[International Review of Cell and Molecular Biology] Volume 309 || Microtubule Plus-End Tracking...

CHAPTER TWO

Microtubule Plus-End TrackingProteins and Their Roles in CellDivisionJorge G. Ferreira*,†, Ana L. Pereira*, Helder Maiato*,†,1*Chromosome Instability & Dynamics Laboratory, Instituto de Biologia Molecular e Celular, Universityof Porto, Porto, Portugal†Cell Division Unit, Department of Experimental Biology, University of Porto, Porto, Portugal1Corresponding author: e-mail address: [email protected]

Contents

1. Introduction 602. Microtubules in Cell Division 64

2.1 Mitotic entry 642.2 Prometaphase–metaphase transition 652.3 Metaphase 662.4 Metaphase–anaphase transition 672.5 Mitotic exit and cytokinesis 67

3. Families of Microtubule Plus-End-Tracking Proteins (þTIPs) 693.1 CLIP family 693.2 EB family 733.3 CLASP family 773.4 APC family 813.5 Motor proteins 823.6 Lis1 843.7 Kinesin-13 family 853.8 TOG family 873.9 Other þTIPs 88

4. Recognition of Microtubule Plus Ends by þTIPs 894.1 Recognizing the microtubule plus end 894.2 Copolymerization 914.3 Diffusion versus motor-based transport 924.4 Hitchhiking 934.5 Turnover at microtubule plus end 95

5. þTIPs in Mitosis 975.1 þTIPs in mitotic spindle organization and positioning 975.2 þTIPs at mitotic centrosome 1025.3 þTIPs at kinetochore 1045.4 þTIPs regulation during mitosis 1095.5 þTIPs in mitotic exit and cytokinesis 113

International Review of Cell and Molecular Biology, Volume 309 # 2014 Elsevier Inc.ISSN 1937-6448 All rights reserved.http://dx.doi.org/10.1016/B978-0-12-800255-1.00002-8

59

http://dx.doi.org/10.1016/B978-0-12-800255-1.00002-8

6. Concluding Remarks 116Acknowledgments 117References 117

Abstract

Microtubules are cellular components that are required for a variety of essential pro-cesses such as cell motility, mitosis, and intracellular transport. This is possible becauseof the inherent dynamic properties of microtubules. Many of these properties are tightlyregulated by a number of microtubule plus-end-binding proteins or þTIPs. These pro-teins recognize the distal end of microtubules and are thus in the right context to con-trol microtubule dynamics. In this review, we address how microtubule dynamics areregulated by different þTIP families, focusing on how functionally diverse þTIPs spa-tially and temporally regulate microtubule dynamics during animal cell division.

1. INTRODUCTION

Division of one cell into two genetically identical daughter cells occurs

through two coordinated processes known as mitosis (division of the

nucleus) and cytokinesis (division of the cytoplasm). In order to do so, cells

have to assemble a dynamic array of MTs known as the mitotic spindle.

Differences in MT dynamic behavior are observed in vivo and can occur

via two distinct mechanisms. One involves the addition and loss of tubulin

subunits at the same end of MTs—a mechanism known as dynamic instabil-

ity (Mitchison and Kirschner, 1984; Sammak and Borisy, 1988; Schulze and

Kirschner, 1988). The other occurs through gain of tubulin at the plus

ends of MTs and loss of tubulin at the minus ends of MTs—a mechanism

known as treadmilling (Margolis and Wilson, 1978; Rodionov and

Borisy, 1997).

Dynamic instability is driven mainly by GTP hydrolysis (Hyman et al.,

1992). Tubulin subunits are incorporated into a protofilament when bound

to GTP (Fig. 2.1). After incorporation, GTP hydrolysis occurs very rapidly

in the b-tubulin subunit (Desai and Mitchison, 1997). This means that the

MT lattice is enriched in GDP-tubulin. As a consequence, MT plus ends are

less stable and tend to adopt a curved conformation, favoring depolymeri-

zation (Desai and Mitchison, 1997; Melki et al., 1989). Given this, how is it

then possible for MTs to stabilize and polymerize? Hydrolysis of GTP is

favored by the addition of new heterodimers and therefore does not occur

60 Jorge G. Ferreira et al.

in the last subunit added to the protofilament but in the one before last. For

this reason, it was proposed that MTs have a GTP b-tubulin cap that would

be sufficient to stabilize them (Mitchison and Kirschner, 1984). The exact

size of the GTP cap is still unclear and many studies have reached different

conclusions, with values ranging from 40 GTP subunits (Voter et al., 1991)

to a single GTP subunit on each protofilament (Caplow and Shanks, 1996;

Drechsel and Kirschner, 1994).

Four parameters are currently used to define dynamic instability: growth

velocity, shrinking velocity, rescue frequency, and catastrophe frequency

(Walker et al., 1988). MT growth velocity depends on soluble tubulin con-

centration and the rate of association of GTP-tubulin to the MT. On the

other hand, shrinking velocity is independent of tubulin concentration

but depends on the dissociation rate of GDP-tubulin. Therefore, increasing

tubulin concentration can increase growth rate which, in turn, leads to a

Polymerization

Growing microtubule

Shrinking microtubule

Catastrophe Rescue

Depolymerization

GDP

GTP

Figure 2.1 Microtubule dynamic instability. GTP-bound tubulin assembles at themicro-tubule plus end creating a stable GTP cap that prevents microtubules fromdepolymerizing. When GTP hydrolysis occurs, the microtubule becomes unstable anddepolymerizes by the outward curving of individual protofilaments, which leads to fur-ther destabilization of themicrotubule structure. When GDP is substituted for GTP in thedisassembled tubulin subunits, the cycle can begin again.

61+TIPs in Cell Division

decrease in MT shortening rate. Catastrophe/rescue frequency is defined as

the number of catastrophes/rescues undergone during the total growth time

of an MT, respectively. These dynamic parameters can be easily visualized

using kymographic tools (Pereira and Maiato, 2010), which provide a visual

representation of the MT plus end over time (Fig. 2.2).

Recent in vitro systems have made possible to recreate physiologic

dynamic instability with minimal components such as MTs, an MT stabi-

lizer, and an MT destabilizer (Kinoshita et al., 2001; Li et al., 2012;

Zanic et al., 2013). Typical growth velocities of MTs in vitro are around

1–5 mm/min, but these values can be much higher in living cells. MT short-

ening velocities are in the order of 10–50 mm/min and are normally 10 times

higher than growth velocities. Hydrolysis of GTP, which occurs at the MT

plus ends, also plays a crucial role in the transition between MT growth and

shrinkage (Hyman et al., 1992). In fact, GTP hydrolysis causes tubulin to

adopt a curved conformation, ultimately leading to destabilization of the lat-

tice (Melki et al., 1989). Because these GDP-tubulin subunits are not

allowed to completely curve while in the lattice, energy released from

hydrolysis is stored as mechanical strain within an MT (Caplow et al.,

1994). This means that when catastrophe events occur, protofilaments adopt

an outward curvature, leading to rapid depolymerization of an MT

(Fig. 2.1).

MT treadmilling was first proposed when it was observed that

isolated bovine brain tubulin continuously incorporated intoMTs at a con-

stant rate, while the MT length remained constant (Margolis and Wilson,

1978). This mechanism implies that (1) there has to be a unidirectional flow

of tubulin subunits with incorporation at the plus end and dissociation

at the minus end and (2) the rate of tubulin association has to be similar

to the rate of tubulin dissociation. The treadmilling model implies that

this mechanism could be bidirectional, depending on the available tubulin

concentration at each given moment. In fact, fluorescence speckle micro-

scopy techniques demonstrated a lack of polarity in treadmilling (Grego

et al., 2001).

It is now widely known that MT behavior is modulated by a number of

MT-associated proteins (MAPs), which can influence dynamic instability

parameters and consequently impact on mitotic progression and fidelity.

Many of these MAPs share the ability to recognize only the distal part of

a polymerizingMT, known as theMT plus end. For this reason, theseMAPs

are currently known as MT plus-end-tracking proteins (þTIPs)

(Akhmanova and Steinmetz, 2008; Schuyler and Pellman, 2001). In this

review, we will cover a range of topics related to the role and regulation


of þTIPs in animal cell division which include (1) how þTIPs can specif-

ically recognize and bind to the plus ends of MTs, (2) howþTIPs are able to

modify MT behavior, and finally, (3) how different þTIPs interact with

each other to coordinate entry, progression, and exit from mitosis.

Space

Tim

e

Growth

Shrinkage

Figure 2.2 Typical kymograph (plot of distance vs. time) with changes in microtubulelength and transitions over time. Kymograph obtained from HeLa cell-expressing GFP-tubulin. Vertical scale bar is 10 s; horizontal scale bar 5 mm. Microtubules will normallyswitch stochastically between growth and shrinkage. Highlighted is one growth phaseand a subsequent shrinkage phase (dashed white lines). A rescue event corresponds toa transition from shrinkage to growth and a catastrophe corresponds to a transitionfrom growth to shrinkage.


2. MICROTUBULES IN CELL DIVISION

2.1. Mitotic entryCell-cycle progression is accompanied by changes in MT dynamics at very

specific stages. In fact, there is an increase in MT dynamics which occurs

concomitantly with NEB that could be important for spindle morphogen-

esis (Piehl and Cassimeris, 2003; Zhai et al., 1996). These changes in MT

dynamics are accompanied by a decrease in tubulin polymer (Zhai and

Borisy, 1994; Zhai et al., 1996), leading to the hypothesis that

MT-stabilizing proteins would have to be inactivated upon mitotic entry

(Cassimeris, 1999). This was supported by the fact that addition of cyclins

or activated CDK1 toXenopus extracts was sufficient to induce a mitotic-like

catastrophe rate ofMTs (Belmont et al., 1990;Murray and Kirschner, 1989).

Curiously, inactivation of CDK1 upon anaphase onset was shown to require

intact MTs (Andreassen and Margolis, 1994) and inhibition of CDK1 pro-

motes MT growth (Moutinho-Pereira et al., 2009; Skoufias et al., 2007).

Why is it necessary for a cell to alter MT dynamics dramatically upon

mitotic entry? During the initial stages of mitosis, spindle poles nucleate

MTs that spatially search for kinetochores. This “search and capture” model

proposed that MTs randomly probe the entire cell volume until they contact

the kinetochore (Kirschner and Mitchison, 1986). However, it became

obvious, based on experimental and theoretical evidence, that this model

alone could not account for the typical mitotic timing (Magidson et al.,

2011; Paul et al., 2009; Wollman et al., 2005). In fact, it was demonstrated

that both the distribution of chromosomes in prometaphase and their move-

ments and rotations significantly reduce spindle assembly time without

compromising mitotic fidelity (Magidson et al., 2011; Paul et al., 2009).

Curiously, assembly or disassembly of MTs can also generate force with-

out direct contribution of motor proteins (Dogterom and Yurke, 1997;

Koshland et al., 1988), and these are sufficient to move subcellular structures

such as chromosomes and organelles, or assist in mitotic spindle positioning

(Dogterom et al., 2005; Inoue and Salmon, 1995;Mogilner andOster, 2003;

Tolic-Norrelykke, 2008). Accordingly, MT polymerization can generate

pushing forces. Addition of tubulin subunits to the MT plus end will induce

its compression when MT hits an object, and this leads to a movement of

MT in the opposite direction, unless MT is attached to some structure

(Dogterom and Yurke, 1997; Holy et al., 1997). These forces can only


be exerted over very short distances because MTs have a tendency to buckle

when they grow too long (Dogterom and Yurke, 1997; Dogterom et al.,

2005). Because of this, in vivo evidence for MT pushing forces has been lim-

ited. However, it has been shown that MTs can contribute to the polar ejec-

tion forces that push chromosome arms away from the pole, thereby

contributing for chromosome alignment (Brouhard and Hunt, 2005;

Inoue and Salmon, 1995; Rieder and Salmon, 1994).

2.2. Prometaphase–metaphase transitionWhen an MT comes into contact with a kinetochore, it becomes stabilized

(Hayden et al., 1990), leading to a poleward movement of the chromosome,

which is dynein dependent (Echeverri et al., 1996; Rieder and Alexander,

1990; Yang et al., 2007). Afterward, CENP-E-mediated forces at the kinet-

ochore move the chromosome to the metaphase plate (Kapoor et al., 2006;

Wood et al., 1997). These traction forces are coordinated with polar ejection

forces, which act on chromosome arms and are driven by chromokinesins

and MT polymerization (Brouhard and Hunt, 2005; Cane et al., 2013;

Ke et al., 2009; Rieder and Salmon, 1994; Yajima et al., 2003). Altogether,

these forces facilitate chromosome alignment at the metaphase plate and help

stabilize kinetochore–MT attachments. However, in the initial stages of

mitosis, most kinetochores can become attached in an incorrect way as mon-

otelic (i.e., only one kinetochore attached), syntelic (i.e., both kinetochores

attached and oriented to the same spindle pole), or merotelic (i.e., one kinet-

ochore attached and oriented to both spindle poles). These need to be

corrected so that kinetochores become amphitelically attached (i.e., each

kinetochore attached to MTs oriented to a single spindle pole). The mech-

anisms involved in kinetochore–MT error correction have been extensively

studied and include the destabilization of kinetochore–MTs (k-fibers) by

Aurora-B-mediated activity (Biggins and Walczak, 2003; Cimini et al.,

2003, 2006; Kline-Smith and Walczak, 2004; Lampson et al., 2004; Liu

et al., 2009a; Loncarek et al., 2007; Magidson et al., 2011). Interestingly,

increasing kinetochore tension, such as happens when chromosomes

become bioriented, induces a spatial separation of Aurora-B from its kinet-

ochore substrates, leading to stabilization of k-fibers (Liu et al., 2009a).

In addition to the biochemical signals generated at the kinetochore, the

dynamic state of MTs is also important for mitotic fidelity. Accordingly, it

has been shown that the temporal regulation of MT dynamics during early

mitosis is essential for genomic stability (Bakhoum et al., 2009a,b).


This regulation depends on the activity of kinesin-13 members Kif2B

and MCAK, which control MT turnover in prometaphase and metaphase,

respectively. Interestingly, it was shown that interaction of Kif2B with

CLASP1 during prometaphase promotes k-fiber turnover, whereas in

metaphase, CLASP1 associates with Astrin to promote k-fiber stability

(Manning et al., 2010). In agreement, increasing the stability of k-fibers pre-

maturely in prometaphase resulted in chromosome missegregation

(Bakhoum et al., 2009a).

Taken together, these results demonstrate that, during prometaphase,

k-fiber stability is reduced so that erroneous attachments can be efficiently

corrected. As cells progress to metaphase and chromosomes become bio-

riented, there is an increase in k-fiber stability which is essential for spindle

assembly checkpoint (SAC) satisfaction. In fact, the SAC constantly moni-

tors for unattached kinetochores so that the mitotic progression is delayed

until all kinetochores are stably attached to k-fibers (Rieder and Maiato,

2004; Rieder et al., 1995).

2.3. MetaphaseUpon establishment of the metaphase spindle, its length and shape appear

relatively stable. However, the spindle itself is quite heterogeneous and

dynamic. Experiments demonstrated that spindle MT turnover was mainly

derived from the high dynamic instability of nonkinetochore–MTs (Buster

et al., 2007; Gorbsky et al., 1990; Salmon et al., 1984; Zhai et al., 1995).

Similar measurements made in kinetochore–MTs showed that, although still

capable of turnover, they do so at much lower rates relative to non-

kinetochore–MTs (�10�) (Zhai et al., 1995). Interestingly, there is a strik-

ing reduction of MT turnover rates and MT flux at anaphase onset,

suggesting that kinetochore–MT attachment is stabilized at this stage

(Gorbsky and Borisy, 1989; Zhai et al., 1995). This further demonstrates that

MTs can also change their dynamic behavior during different stages of

mitosis.

In addition to dynamic instability, a second mechanism also ensures

proper spindle dynamics in metaphase, which is known asMT poleward flux

(Mitchison, 1989). This is a highly conserved feature of the mitotic spindle

in higher eukaryotes and is associated with the incorporation ofMT subunits

at the MT plus ends and disassembly of subunits at the MT minus ends

(Mitchison et al., 1986). Current models proposed to explain MT flux take

into account the following premises: active incorporation of tubulin sub-

units at the kinetochore, disassembly of tubulin subunits at the centrosome


(Buster et al., 2007; Kwok and Kapoor, 2007; Mitchison, 1989), as well

as sliding of MTs through the action of plus-end-directed motors

(Brust-Mascher and Scholey, 2002; Matos et al., 2009; Pereira and

Maiato, 2012). The net result is the stabilization of the spindle size while

maintaining the structure highly dynamic.

2.4. Metaphase–anaphase transitionDifferentMT populations have distinct dynamic properties in the metaphase

spindle. Nonkinetochore–MTs have a higher turnover when compared to

k-fibers (Cassimeris et al., 1990; Mitchison et al., 1986; Saxton et al., 1984).

These differences will be reflected on MTs as cells enter anaphase. In fact,

turnover of k-fiber was shown to decrease as cells enter anaphase by as much

as fivefold when compared to the same population ofMTs in metaphase cells

(Gorbsky and Borisy, 1989; Zhai et al., 1995), whereas turnover of non-

kinetochore fibers is not affected during the transition from metaphase to

anaphase (Zhai et al., 1995).

Shortening of k-fibers during anaphase should occur either by activeMT

depolymerization at the pole region (known as the “Traction Fiber” model)

(Buster et al., 2007;Matos et al., 2009;Waters et al., 1996) orbydisassemblyof

MTs at the kinetochore (known as the “Pacman” model) (Cassimeris and

Salmon, 1991; Gorbsky et al., 1987, 1988; Maiato, 2010; Nicklas, 1989).

MT depolymerization per se is sufficient to drive chromosome movement

in vitro (Coue et al., 1991; Koshland et al., 1988) and for generating force

(Grishchuk et al., 2005). This was first demonstrated in vitro when it was

shown that depolymerizingMTs alone could generate sufficient pulling force

to move chromosomes without the contribution of motors (Koshland et al.,

1988). Subsequent reports demonstrated that, in an in vitro system, chromo-

somes were being pulled at about 30 mm/min in an ATP-independent man-

ner and, thus,were relying only onMTdepolymerization (Coue et al., 1991).

However, there is evidence that this process might also require the assistance

ofmotor proteins tomove chromosomes (Desai andMitchison, 1997;Maiato

and Lince-Faria, 2010; Pfarr et al., 1990).

2.5. Mitotic exit and cytokinesisMTs are also necessary for changes in cell shape and size during anaphase and

telophase. Upon anaphase onset, depolymerization of spindle MTs has to be

compensated by an increase in astral MT polymerization/elongation

(Morrison and Askham, 2001; Strickland et al., 2005b). Elongation of astral


MTs is necessary for their interaction with the cell cortex and definition of

the cytokinetic furrow, but apparently is not essential for anaphase progres-

sion itself, as the cytokinetic furrow can still be formed even in the absence of

astral MTs (Rankin and Wordeman, 2010; Strickland et al., 2005a,b;

Sullivan and Huffaker, 1992).

MT reorganization during mitotic exit is strictly associated with the inac-

tivation of the mitotic kinase CDK1, which triggers the formation of ana-

phase MTs and the midbody (Wheatley et al., 1997). A similar phenomenon

was also observed in Drosophila S2 cells and shown to involve acentriolar

MT-organizing centers (aMTOCs). These aMTOCs were able to nucleate

MTs de novo upon CDK1 inhibition at anaphase onset (Moutinho-Pereira

et al., 2009), and this was dependent on the activity of Msps/XMAP215

and KLP10A/kinesin-13. This reorganization also depends on the precise

regulation of MT dynamics and allows daughter cells to adhere simulta-

neously to the substrate (Ferreira et al., 2013).

Cytokinesis relies on MTs in several ways. First, definition of the cleav-

age plane is specifically determined by astral MTs (and not spindle MTs) as

furrowing still occurs in the presence of asters without any intervening spin-

dle (Rieder et al., 1997). However, successful completion of cleavage does

require interaction of midzone MT bundles with the cell cortex (Wheatley

and Wang, 1996). Moreover, if anaphase astral MT formation is suppressed

by interfering with the þTIP EB1 or with dynactin, cytokinesis is delayed

(Strickland et al., 2005b), which supports the necessity of MT interaction

with the cortex to define cleavage plane localization (Bement et al., 2005;

Strickland et al., 2005a). At this stage, regulation of MT dynamics seems

to be dispensable, as contact of MTs with the cortex is sufficient to trigger

the process. In contrast with earlier stages of cytokinesis, MTs are essential

for completion of the process (Savoian et al., 1999). MTs that establish the

midbody are acetylated, highly stable (Margolis et al., 1990), and resistant to

nocodazole treatment (Foe and von Dassow, 2008; Piperno et al., 1987).

Nevertheless, some midbody MTs are still able to exhibit a highly dynamic

behavior as can be seen by live imaging of MT plus ends with fluorescent-

tagged EB proteins, which show comets moving in and out of the midbody

(Rosa et al., 2006). Thus, it is not surprising that g-tubulin was found in themidbody during late cytokinesis (Julian et al., 1993), suggesting active MT

nucleation. Notably, g-tubulin interacts with the Augmin complex during

anaphase, and this is required for MT nucleation in the central spindle and

successful cytokinesis (Uehara et al., 2009). Final disassembly of the midbody


requires that MTs are cut, which is accomplished by a mechanism that

involves the MT-severing enzyme spastin (Guizetti et al., 2011).

3. FAMILIES OF MICROTUBULE PLUS-END-TRACKINGPROTEINS (+TIPs)

Many proteins have the ability to associate with MTs. Among these, a

large number of MAPs specifically recognize the terminal portion of MT

(Table 2.1). These are collectively known as MT plus-end-tracking proteins

or þTIPs (Akhmanova and Steinmetz, 2008; Schuyler and Pellman, 2001).

When theseþTIPs are labeled with a fluorescent tag, they appear as comets

in the MT tip, moving throughout the cell as MT grows and disappearing

when MT shrinks (Howard and Hyman, 2003). In this section, we will

cover the structural features and function of the main families of þTIPs.

3.1. CLIP familyThe first descriptionof tip-tracking behavior came fromworkofKreis and col-

laborators, who demonstrated that cytoplasmic linker protein (CLIP) 170 was

able to specifically associate with the plus end of polymerizing MTs

(Diamantopoulos et al., 1999; Perez et al., 1999). The CLIP family of proteins

is comprisedof twomembers inmammalians:CLIP170 andCLIP115.The lat-

ter is a brain-specific CLIP that shares functional similarities with CLIP170

(DeZeeuwet al., 1997).These proteins have a characteristicCAP-Gly domain

(Fig. 2.3) which is necessary for interaction with tubulin and EB1 (Weisbrich

et al., 2007). TheseCAP-Gly domains are surrounded bybasic, serine-rich res-

idues that assist in the binding to MTs (Hoogenraad et al., 2000). In order to

perform its function, CLIP170 needs to form a parallel homodimer. Each

monomer is composed of an N-terminal domain required for MT binding

(with two CAP-Gly domains per monomer), a central coiled-coil domain

required for dimerization, and a C-terminal metal-binding domain (with

two zinc fingers per monomer; Fig. 2.3) (Gupta et al., 2009; Pierre et al.,

1994; Scheel et al., 1999). Both the CAP-Gly domains at the N-terminus

and the zinc fingers at the C-terminus are thought to play an important role

in the autoregulation of CLIP170 (Hayashi et al., 2007; Lansbergen et al.,

2004). In accordance, it was shown that they can interact with each other, cre-

ating a doughnut-shapedmolecule that no longer interacts withMTs. In addi-

tion, these autoinhibitory interactions use the same binding determinants as

CLIP170’s intermolecular interactions with p150glued, suggesting that


Table 2.1 Main þTIP families and main functions during cell divisionþTIP Homologues Interacting þTIPs Main mitotic functions References

EB1 family (EB1,

EB2, EB3)

Mal3 (Sp)

Bim1 (Sc)

AtEB1 (At)

Most þTIPs (with

SxIP and CAP-Gly

domains)

Nucleation of astral microtubules;

loading of þTIPs to plus ends;

spindle positioning; cytokinesis

completion; postmitotic cell

adhesion

Tirnauer and Bierer (2000),

Toyoshima and Nishida (2007),

Rogers et al. (2002), Stout et al.

(2011), Ferreira et al. (2013)

CLIP family

(CLIP170,

CLIP115)

Tip1 (Sp)

Bim1 (Sc)

CLIP190

(Dm)

EB family

CLASPs p150glued

Cytoplasmic dynein

Microtubule rescue and

stabilization; targeting of dynein to

plus ends; required for mitotic

progression; microtubule

interaction with cell cortex and

kinetochores

Arnal et al. (2004), Dujardin et al.

(1998), Wieland et al. (2004),

Tanenbaum et al. (2006)

APC family (APC,

APC2/APC-L)

Kar9 (Sc)

APC1/2

(Dm)

APR-1 (Ce)

EB family Microtubule stabilization;

regulation of kinetochore–

microtubule interaction;

chromosome segregation; spindle

positioning

Kaplan et al. (2001), Fodde et al.

(2001), Green et al. (2005),

McCartney et al. (2001), Zhang

et al. (2007a)

CLASP family

(CLASP1,

CLASP2)

Peg1 (Sp)

Stu1 (Sc)

MAST/Orbit

(Dm)

Cls-2 (Ce)

CLASP (At)

EB family

CLIP170

CLIP115

Kinesin-7

Spindle microtubule dynamics;

mitotic spindle organization and

assembly; spindle pole integrity;

kinetochore–microtubule

attachment; cytokinesis completion

Mimori-Kiyosue et al. (2006),

Logarinho et al. (2012), Lemos

et al. (2000), Maiato et al. (2005),

Maiato et al. (2003a), Pereira et al.

(2006), Maffini et al. (2009)

Motor proteins

(kinesin-7, kinesin-

14, dynein)

Kinesin-7

Tea2 (Sp)

Klp2 (Sc)

Kinesin-14

Ncd (Dm)

KLP2 (Sp)

Kar3 (Sc)

Cytoplasmic

Dynein

EB family

Dynein

CLASPs;

EB family;

Dynactin

(p150glued)

LIS1

Spindle formation; chromosome

congression; microtubule plus-end

elongation; interpolar microtubule

sliding; metaphase chromosome

alignment; spindle pole focusing;

spindle positioning

Kapoor et al. (2006), Wood et al.

(1997), Kapitein et al. (2005),

Cooke et al. (1997), Sardar et al.

(2010), Goshima et al. (2005),

Kiyomitsu and Cheeseman (2012),

Maffini et al. (2009), O’Connell

and Wang (2000)

Kinesin-13 family

(Kif2C/MCAK)

XKCM1 (Xl)

KLP10A

(Dm)

AtKinesin-13

(At)

EB family

CLIP170

APC

Microtubule depolymerization;

spindle assembly; kinetochore–

microtubule turnover; error

correction

Ganem andCompton (2004), Ems-

McClung et al. (2007), Moore and

Wordeman (2004), Wordeman

et al. (2007), Bakhoum et al.

(2009b), Ganem et al. (2005)

TOG family (ch-

TOG)

XMAP215

(Xl)

Dis1

Alp14(Sp)

Stu2 (Sc)

Msps (Dm)

ZYG-9 (Ce)

EB family

Dynein

Microtubule stabilization; spindle

pole organization; centrosome

integrity; spindle assembly;

protecting kinetochore fiber

disassembly

Gergely et al. (2003), Cassimeris

and Morabito (2004), Barr and

Gergely (2008), Booth et al. (2011)

At, Arabidopsis thaliana; Sc, Saccharomyces cerevisiae; Sp, Schizosaccharomyces pombe; Dm, Drosophila melanogaster; An, Aspergillus nidulans; Ce, Caenorhabditis elegans; Xl,Xenopus laevis; Hs, Homo sapiens.

regulation of MT binding by þTIPs occurs through direct competition

between homologous binding interfaces (Hayashi et al., 2007).

CLIP proteins were described to impact on MT dynamics either directly

or by recruiting a rescue factor (Komarova et al., 2002). Although CLIP115

1 444 735

TOG/TOG-like domain Helical region Basic/Ser

CLASP-like protein

Helical region Helical region

1538

1 14 116 185

CH domain Coiled-coil EBH domain EEY/F

268255

EB-like protein

Zn fingerBas/Ser

1 78 120 232 274 350 1353

Bas/Ser CAP-Gly Bas/Ser CAP-Gly Coiled-coil Zn finger EEY/F

1438

CLIP-like protein

Basic/Ser

1 248 767

Coiled-coil Helical region

2843

APC protein

Coiled-coil

Armadillo repeats

Basic/Ser

453

Coiled-coil

1 181 1868

Dynein heavy chain protein

Helical region Coiled-coil Helical region Coiled-coilAAA ATPase AAA ATPase

444 1171 3189 3553 4646

1 255 658

Kinesin domainHelical region Basic/Ser Coiled-coil

725618518

Kinesin-like protein

1 159

TOG domain Basic/Ser Helical region

1399 2032

TOG domain TOG domain TOG domain TOG domain

ch-TOG protein

1 39

WD40 repeatLisH Coiled-coil

41085 96

Lis1 protein

Figure 2.3 Structural diagram of the main þTIP families. Cartoon depicting relevantdomains in the main þTIP families. Bas/Ser-basic and proline/serine-rich sequenceregions; CAP/Gly, cytoskeleton-associated protein/glycine-rich domain; Zn finger, zincfinger; CH, calponin homology domain; TOG, tumor overexpressed gene domain;EBH, end binding homology domain; LisH, Lis1-homology motif.


lacks the C-terminal domain of CLIP170, they share a similar N-terminal

domain, which means they could regulate MT dynamics in a similar fashion

(Hoogenraad et al., 2000; Komarova et al., 2002). In fact, both in vivo and

in vitro studies using the N-terminus of CLIP170 demonstrated that this pro-

tein acts by preventing catastrophes or promoting rescue events (Arnal et al.,

2004; Komarova et al., 2002). The exact mechanism of CLIP170-mediated

MT rescue is still unclear but may involve stabilization of the curved

protofilaments by the N-terminus of CLIP170 or coassembly of CLIP170

with tubulin oligomers into MTs (Arnal et al., 2004; Diamantopoulos

et al., 1999). Interestingly, although CLIPs track only growing MT plus

ends, they also influence the behavior of depolymerizingMTs. This is a puz-

zling observation and indicates that CLIPs function is not totally understood.

3.2. EB familyEnd binding (EB) proteins are part of a highly conserved family which, in

mammalians, comprises three members encoded from three different genes:

EB1, EB2 (RP1), and EB3 (EB3F) (Su and Qi, 2001). EB1 was the first

member identified in a yeast two-hybrid screen as an interactor of the

C-terminus of the adenomatous polyposis coli (APC) tumor suppressor pro-

tein (Su et al., 1995). Both EB1 and EB3 seem to be ubiquitously expressed,

whereas EB2 expression is restricted to only certain cell types/tissues (Su and

Qi, 2001). Normally, EB1 is expressed in higher levels when compared to

other EBs. However, EB3 is also highly expressed in specific cell types. EB3

was originally reported in neurons, where it was shown to interact with a

brain-specific form of APC (APC2), but it is also highly abundant in muscle

cells (Nakagawa et al., 2000; Straube and Merdes, 2007).

EBs are relatively small, elongated proteins (around 32 kDa) with con-

served structural features (Fig. 2.3). All members have at the N-terminal

region an MT-binding portion containing a calponin homology (CH)

domain with a highly conserved fold (Akhmanova and Steinmetz, 2008).

The structural basis for EB1 binding to MTs has already been described

(Hayashi and Ikura, 2003; Slep and Vale, 2007). It was shown that this

CH domain is both required and sufficient for binding to MT plus ends

(Hayashi and Ikura, 2003; Komarova et al., 2009). The C-terminal portion

of EB1, on the other hand, contains a coiled-coil region which is necessary

for EB dimerization (Su and Qi, 2001). This is essential not only because

they need two CH domains to interact with MTs but also to form the func-

tional C-terminal domain (Buey et al., 2011; Honnappa et al., 2005).


Recently, it was demonstrated that EB1 and EB3 prefer heterodimerization

to EB1/EB1 or EB3/EB3 homodimers (De Groot et al., 2009), while EB2

does not show preferential association with any other EB member. This

chain exchange between EBs can be suppressed by specific EB interaction

partners, which indicates an extra layer of regulation of EB function

(De Groot et al., 2009).

The coiled-coil region partially overlaps the end binding homology

(EBH) domain, which was shown to be required for efficient interaction

with EB-binding partners (Akhmanova and Steinmetz, 2008; Bjelic et al.,

2011). Solving the C-terminal structure of EB1 (EB1c) by X-ray crystallog-

raphy demonstrated that the coiled-coil terminates in a 4-helix bundle with

a hydrophobic cavity (Honnappa et al., 2005; Slep et al., 2005). In addition,

EB1c has an EEY/F motif that is very similar to the one found in a-tubulinand CLIP170 (Komarova et al., 2005; Mishima et al., 2007;Weisbrich et al.,

2007) and might be important to help in the regulation of EB1/CLIP170/

tubulin association (Bieling et al., 2008;Mishima et al., 2007). Both EB1 and

EB3 have very similar structures, which are highlighted by the fact that they

share some functional similarity (Komarova et al., 2005, 2009). On the other

hand, EB2 appears to have fewer similarities with the other two family

members. Not only the interaction partners are substantially different

between this and other EBs, but also EB2 does not promote persistent

MT growth or restore CLIP association to the MT plus ends (De Groot

et al., 2009; Komarova et al., 2005, 2009). In fact, EB2 does not interact

to the same extent with MCAK, APC, or CLIP170 (Bu and Su, 2003;

Komarova et al., 2005; Lee et al., 2008). This can be explained by the fact

that the C-terminal domain of EB2 is significantly different from EB1 and

EB3, with fewer acidic residues. Furthermore, EB2 has a longer N-terminal

region, containing approximately 40 amino acids in excess when compared

to EB1 and EB3 (Komarova et al., 2009). Interestingly, this difference in the

N-terminal domain is clustered around the sequence SRHD in the CH

domain, which is essential for MT binding and can explain the differences

observed between EB2 and the other family members in this aspect

(Komarova et al., 2009).

EB proteins are associated with MT plus ends in both interphase and

mitotic cells (Fig. 2.4; Berrueta et al., 1998; Mimori-Kiyosue et al.,

2000b; Morrison et al., 1998). The first report regarding the possible role

of EB proteins inMT dynamics came from the observation that, when over-

expressed, these proteins induced the formation of acetylated MT bundles

that were resistant to nocodazole treatment (Bu and Su, 2001). In addition,


their ability to tip-track MTs led to the possibility that they might be

involved in MT dynamics regulation, particularly in promoting MT growth

(Nakamura et al., 2001; Tirnauer and Bierer, 2000). This was confirmed in

many independent studies using not only different model organisms such as

budding and fission yeast, Drosophila, and human cells, but also in vitro sys-

tems (Beinhauer et al., 1997; Coquelle et al., 2009; Komarova et al., 2009;

Nakamura et al., 2001; Rogers et al., 2002; Tirnauer et al., 1999).

The overall picture that has emerged confirms the role of EB proteins in

the regulation of MT dynamics, but their precise effect is still not fully

understood. In mouse fibroblasts, EB1 depletion leads to an increase in

MT pausing and a decrease in MT growth time (Kita et al., 2006). In addi-

tion, EB1 was also shown to induce MT stabilization by interacting with

mDia and APC (Wen et al., 2004) and to localize to stable Glu-MTs. In

these conditions, knockdown of EB1 leads to the appearance of more

dynamic MTs, as demonstrated by the concomitant decrease in Glu-

MTs. EB3 also interferes with MT dynamics. In fact, it was shown in myo-

blasts that EB3 depletion induced MT overgrowth near the cell cortex and a

significant decrease in MT shrinkage rate (Straube and Merdes, 2007). EB1

and EB3 also promote persistent growth of internal MTs by suppressingMT

catastrophes (Komarova et al., 2009).

The impact of EB proteins on interphase MT dynamics may also involve

their interaction with otherþTIPs. In fact, differences in the expression and

regulation of several þTIPs in different cell types may be responsible for

the observed differences in specific MT populations (Ligon et al., 2003).

A B

Figure 2.4 Localization of EB1 during (A) interphase and (B) mitosis. Immunolocalizationof EB1 (green) and a-tubulin (red) in fixed cells using specific antibodies. EB proteinsassociate with the growing ends of microtubules throughout the cell cycle. In addition,EB1 also associates with the centrosome. Scale bars, 5 mm.


In addition, data derived from in vitro assays demonstrated that EB1 can act

cooperatively with other þTIPs such as CLIP170 in the regulation of MT

dynamics. Accordingly, it was demonstrated that both EB1 and CLIP170

can synergize to modulate MT dynamics, possibly by modifying the

MT-stabilizing cap (Lopus et al., 2012). Moreover, EB association with

CLASPs was also reported to affect MT dynamics at the cell cortex by

increasing MT rescue events (Mimori-Kiyosue et al., 2005). Interestingly,

EBs can also associate with and load MT depolymerizers such as MCAK

to the MT plus ends (Montenegro Gouveia et al., 2010; Moore et al.,

2005). This interaction is important for the localization ofMCAK to the plus

ends but also to enhance its catastrophe-inducing activity. Thus, by allowing

the accumulation of polymerizers and depolymerizers at the MT plus end,

EB proteins facilitate the rapid switching between MT growth and

shortening.

Interestingly, modulation of MT dynamics by EB proteins can also be

regulated by phosphorylation. In budding yeast, the single EB-like protein

was described to be phosphorylated by Ipl1p/Aurora-B and this is important

to regulate the association of EB to spindle MTs (Zimniak et al., 2009).

Moreover, a mutation in the fission yeast EB-like protein was sufficient

to increase MT binding, leading to their stabilization (Iimori et al., 2012).

In humans, less is known about the phosphoregulation of EB proteins.

Recent work demonstrated that EB3 is phosphorylated by Aurora kinases

on S176 during mitosis (Ban et al., 2009). This Aurora-mediated EB3

phosphorylation leads to a significant increase in MT growth, allowing

stabilization of the midbody (Ferreira et al., 2013). In this context, dephos-

phorylation of EB3 restricts cortical MT growth, allowing proper daughter

cell adhesion to the substrate. Inversely, phosphorylation of EB3 on S162 by

the Src-PLCg2 signaling pathway was shown to block MT growth, leading

to adherens junction stabilization in interphase cells (Komarova et al., 2012).

Taken together, these data demonstrate that EB protein association to the

MT plus ends can be regulated by phosphorylation, although it is still unclear

how different phosphorylation events integrate to control EB function, thus

regulating MT dynamics in different tissues.

Besides its plus end localization, EB proteins were also shown to bind

other subcellular structures either directly (centrosome) or indirectly

(F-actin and membranes). In fact, EB1 is a functional component of centro-

somes and binds to this structure independently of MTs through its

C-terminal domain (Louie et al., 2004). Curiously, the C-terminal domain

of EB1 is also required for the recruitment of g-tubulin to centrosomes and


anchoring of MTs to this structure (Askham et al., 2002). Moreover, EB1

was shown to interact with the centrosomal protein FOP (Yan et al.,

2006). This interaction is essential for recruitment of EB1 to the centrosome

and its later association with CAP350, forming an MT-anchoring complex.

EB3 also localizes to the centrosome (Ban et al., 2009; Ferreira et al., 2013).

EBs can also interact indirectly with actin filaments or membrane structures.

EB1was shown to interactwith the spectraplakinACF7/MACF1, providing

a link between MT and actin cytoskeletons (Kodama et al., 2003). Overall,

EB proteins act, either directly or through interaction with a partner, as

mediators of cellular functions by regulating MT dynamics. More detailed

data on the physiological relevance of EB proteins still await the develop-

ment of mammalian knockout models.

3.3. CLASP familyCLIP-associating proteins (CLASPs) are highly conserved þTIPs involved

in the regulation and organization of cellular MT dynamics, motility, and

cell division. The CLASP protein family was first identified in a genetic

screen for mitotic mutants in Drosophila and was named as multiple asters

(MAST)/Orbit (Inoue et al., 2000; Lemos et al., 2000). In mammals, there

are two paralog genes encoding for CLASP1 and CLASP2 proteins, which

were found in a yeast two-hybrid screen as interacting proteins with

CLIP115 and CLIP170 (Akhmanova et al., 2001). While CLASP1 is more

ubiquitously expressed, CLASP2 is predominantly expressed in the brain

and reproductive organs (Akhmanova et al., 2001), as well as in the hema-

topoietic organs in mice (Drabek et al., 2012). All the data collected in dif-

ferent model organisms suggest a functional role of CLASPs starting at

embryogenesis (Inoue et al., 2000; Lemos et al., 2000; Park et al., 2012).

Both clasp1 and clasp2 genes can undergo alternative splicing events, origi-

nating several isoforms. So far, only one biologically active isoform has been

found for CLASP1, known as CLASP1a (�170 kDa). On the other hand,

three isoforms have been described for CLASP2, namely, CLASP2a(�170 kDa), CLASP2b (�140 kDa), and CLASP2g (�140 kDa), which

result from alternative splicing events (Akhmanova et al., 2001).

CLASPs display a conserved structure, sharing approximately 77%

sequence homology (Akhmanova et al., 2001), and contain two short

Ser-x-Ile-Pro (SxIP) polypeptide motifs embedded in an extensive central

sequence region enriched with positively charged serine and proline residues

(Fig. 2.4), which is highly conserved across species (Honnappa et al., 2009;


Kumar et al., 2012; Mimori-Kiyosue et al., 2005). These motifs have been

shown to be essential for the interaction with the C-terminal domain of EB

proteins and are important for the plus-end-tracking activity of CLASPs

(Honnappa et al., 2009). The recurrence of the SxIPmotif found in CLASPs

also enables the intermolecular cooperation between them to significantly

improve the efficiency of MT tip-tracking (Honnappa et al., 2009).

Until recently, the general perception regarding the N-terminal domain

of CLASPs was that it contained only one TOG domain, accompanied by a

variable number of TOG-like regions that hold a weak sequence homology

to those found in proteins of the Dis1/TOG family (Lemos et al., 2000).

However, recent evidence gathered from X-ray crystallography determined

that human CLASP1 has, at least, two legitimate TOG domains: TOG1 and

TOG2 (previously classified as cryptic TOG-like 2) (Leano et al., 2013). In

yeast, the TOG domains of CLASP are capable of binding directly to soluble

tubulin dimers, but not to dimers that are already incorporated in the MT

lattice (Al-Bassam and Chang, 2011; Al-Bassam et al., 2010). The detailed

mechanism behind the interaction of TOG domains with soluble tubulin

is yet to be fully understood, but important new data may have shed light

on the precise mechanism that controls association of human CLASP1

with MTs. Accordingly, CLASP1 TOG2 domain has a distinctive bent

conformation, which is hypothesized to be a good fit to bind to the curved

conformation of tubulin dimers on depolymerizing MTs. This leads to their

stabilization, possibly leading to a rescue event (Leano et al., 2013). How-

ever, the authors suggest that this conformational variation in TOG2 may

only occur upon lattice binding. This particular domain also seems to be

important for the establishment of a CLASP-mediated bipolar spindle

(Leano et al., 2013).

Interestingly, the N-terminal region is different between the CLASP2

isoforms. Notably, the previously described CLASP2 TOG domain only

exists in the longer alpha isoform, while being absent from the shorter

isoforms (Akhmanova et al., 2001). In CLASP2b, it is replaced by a short

N-terminal palmitoylation motif, which gives CLASP2b the ability to

anchor membranes. On the other hand, CLASP2g contains the inconspic-

uous peptide—MAMGDD—in this region.

The central region of CLASPs contains six HEAT repeats embedded

between the TOG domains. These repeats were suggested to be involved

in intracellular transport, MT dynamics, and chromosome segregation,

but their exact function is still unknown (Neuwald and Hirano, 2000;

Tournebize et al., 2000). Within this central region, the SxIP motif and


the innermost TOG domain of CLASP1 are responsible for its binding to

PRC1, an MT-bundler protein of the central spindle (Patel et al., 2012).

Of interest, CLASP2a association with actin stress fibers occurs through this

middle serine–arginine (SR)-rich motif and the N-terminal Dis1/TOG

domain in an MT-independent way, providing a direct cross-link between

MTs and the actin cytoskeleton, which is important for cell morphogenesis

(Tsvetkov et al., 2007).

TheC-terminal domain of CLASPs participates in the interactionwith the

Golgi apparatus by binding with the trans-Golgi network protein GCC185,

an interaction that contributes to an asymmetry of the MT array nucleated at

the Golgi (Efimov et al., 2007). The coiled-coil domain present in this region

is also important for the binding of CLASPs to interacting partner proteins,

such as CLIP170, CENP-E, and Plk1 (Akhmanova et al., 2001; Hannak

and Heald, 2006; Maffini et al., 2009; Maia et al., 2012), as well as kinesin-

10/Kid, a chromokinesin which is involved in chromosome congression by

generating polar ejection forces (Antonio et al., 2000; Levesque and

Compton, 2001; Patel et al., 2012; Wandke et al., 2012). The interaction

of CLIP170 with the C-terminal domain of CLASPs has been shown to

enhance CLASPs plus-end association (Mimori-Kiyosue et al., 2005). How-

ever, this CLIP170 interacting region does not seem to be required forCLASP

plus-end tracking or lattice binding (Wittmann andWaterman-Storer, 2005).

Finally, the C-terminal region also seems to be implicated in the

homodimerization of CLASPs (Al-Bassam et al., 2010; Patel et al., 2012).

In interphase,CLASPs canbe found associatedwith theplus endsof grow-

ing MTs, centrosomes, and perinuclear region, consistent with Golgi appa-

ratus localization (Akhmanova et al., 2001; Efimov et al., 2007). They were

also shown to be required for the stabilization of MTs at the leading edge of

motile fibroblasts (Akhmanova et al., 2001). It was demonstrated that

CLASP2 is necessary for the establishment of a stable, polarized MT array

in mouse embryonic fibroblasts, promoting persistent directional motility

in these cells (Drabek et al., 2006). Depletion of both CLASPs by RNAi

resulted in a decrease in the levels of acetylated tubulin, which was accompa-

nied by a reduction inMTdensity (Mimori-Kiyosue et al., 2005). This led to

the hypothesis that, when bound to the plus ends of MTs, CLASPs are

required for rescue events by reducing the number of long depolymerization

episodes (Akhmanova et al., 2001; Al-Bassam et al., 2010; Mimori-Kiyosue

et al., 2005; Sousa et al., 2007).Additional evidence further demonstrated that

CLASPs also increase MT longevity by promoting MT “pausing,” and con-

sequently their stability, without affecting overall MT polymerization rate.


The accumulation of CLASPs at MT plus ends and their

MT-stabilization capacity are also important for the interaction of MTs with

the cell cortex through a complex with LL5b, a process that also seems to

require the spectraplakin MACF/ACF7 (Drabek et al., 2006; Lansbergen

et al., 2006). Notably, it was recently demonstrated, using a mouse knockout

model for Clasp2, that this protein is important for cell attachment and

proper organization of the MT network in hematopoietic stem cells

(Drabek et al., 2012). In this way, CLASP2 is an important player in the

homing and maintenance of hematopoietic stem cells in vivo.

The mechanisms by which CLASPs are able to interact with MTs are

only now being unraveled. It is now known that association of CLASPs

to MTs can be regulated by posttranslational modifications, such as phos-

phorylations. It was demonstrated that the interaction of CLASPs with

MTs is negatively regulated by glycogen synthase kinase (GSK)-3b, a

downstream target of phosphoinositide PI3-kinase (Akhmanova et al.,

2001). Initial observations implied a major increase in CLASP2 signal at dis-

tal MT ends upon GSK-3b inhibition in 3T3 fibroblasts (Akhmanova et al.,

2001). On the other hand, overexpression of a constitutively active GSK-3bform severely prevented CLASP2 localization to MT plus ends and strongly

disrupted CLASP2 MT lattice binding (Akhmanova et al., 2001; Wittmann

and Waterman-Storer, 2005), reinforcing the requirement of GSK-3bkinase activity for CLASP2 association to different subsets of MTs. On

the contrary, inhibition of the kinase stimulated ectopic MT lattice associ-

ation in the cell body. Based on these data, it was proposed that the

MT-binding domain of CLASP2 comprises different functions: it is required

for high affinity binding of CLASP2 to the MT lattice in the lamella, as well

as plus-end tracking. Later experiments identified the GSK-3b phosphory-

lation sites in the MT-binding domain that are involved in the transition

between plus-end tracking and lattice binding (Kumar et al., 2009), con-

firming that CLASP2 is spatially regulated in cells. The fact that these phos-

phorylations by GSK-3b affect the association between CLASP2 and EB1

may explain the alterations observed in CLASP2 tip-tracking ability

(Kumar et al., 2012). Importantly, a priming site phosphorylation of

GSK-3b by CDKs is necessary for GSK3b-mediated CLASP2 phosphory-

lation. Similarly, location and regulation of CLASPs in specific structures

during mitosis seems to be controlled through the phosphorylation activity

of CDK1 and Plk1 (Kumar et al., 2012; Maia et al., 2012), which will be

discussed in more detail in Section 5.4. Finally, the latest results obtained

with a Clasp2 knockout mouse model confirmed the importance of GSK3b


in the regulation of CLASP2 activity, especially at the neuromuscular junc-

tions (Schmidt et al., 2012).

3.4. APC familyThe APC protein is a large protein of approximately 300 kDa encoded by

theApc gene (Smith et al., 1993). In mammalians, an additional form of APC

can be found, which is a product of theAPCL/APC2 gene (Nakagawa et al.,

1998). In structural terms, APC is composed of several domains (Fig. 2.3).

Close to the N-terminus, there is an oligomerization domain and an arma-

dillo repeat domain (ARD). In the middle of the protein, there are

b-catenin-binding motifs, Axin-binding motifs, and also a mutation cluster

region. In addition, there is a KKKK stretch, which is postulated as a putative

nuclear localization signal. In the C-terminus, there is an MT-binding

domain and an EB1-binding domain (Bienz, 2002). The interaction of

EB1 with APC was first mapped to a small region in the C-terminus of

APC which comprises amino acids 2559–2843 (Su et al., 1995). Subsequent

work narrowed this region to the last 170 amino acids of APC (Askham

et al., 2000), and finally, the interaction was attributed to a basic, serine-rich

sequence in the C-terminus of APC named APCp1 (Honnappa et al., 2005).

More specifically, interaction of APC with EB1 depends on the SxIP motif

(Ile2805 and Pro2806) of APC (Honnappa et al., 2005, 2009). Interestingly,

mutations within this region are sufficient to abolish EB1 interaction and

also the ability of APC to tip-track.

APC is involved in the regulation of MT function. In fact, APC directly

associates with MTs and promotes their polymerization and stabilization

in vitro (Munemitsu et al., 1994; Nakamura et al., 2001; Zumbrunn et al.,

2001). As was mentioned earlier, interaction of APC with EB1 seems to

be important for its ability to track MT plus ends (Mimori-Kiyosue et al.,

2000a). However, this might not be the only mechanism that APC uses

to localize to growing MT ends, as APC association to MTs can occur even

in the absence of EB1 (Kita et al., 2006). Moreover, APC can also accumu-

late at the MT plus ends by interacting with Kif3A/Kif3B (Jimbo et al.,

2002). Nevertheless, it seems that APC is mainly loaded onto plus ends

by hitchhiking on EB1 (Honnappa et al., 2009). This interaction is impor-

tant because it was shown that it can help regulate MT stability and promote

cell migration (Wen et al., 2004), although another study with mouse

embryonic fibroblasts derived from mice carrying a truncated Apc allele

demonstrated that the APC–EB1 interaction is not essential for MT


stabilization (Drabek et al., 2006). APC is also involved in the interaction

between MT and actin cytoskeletons (Moseley et al., 2007) and in the

regulation of cell polarity (Etienne-Manneville and Hall, 2003).

3.5. Motor proteinsMany organisms have a set of motor proteins that can travel along MTs

toward their plus- or the minus ends. Recently, many of these motor pro-

teins have also been identified asþTIPs (Wu et al., 2006). These include the

plus-end directed, kinesin-7 family member, CENP-E (Cooke et al., 1997;

Sardar et al., 2010) as well as kinesin-5 Eg5 (Jiang et al., 2012) and theminus-

end-directed dynein (Kobayashi and Murayama, 2009). In this section, we

will focus on the functional relevance and mechanisms involved in motor

protein accumulation at MT plus ends.

3.5.1 KinesinsMost kinesins show plus-end-directedmotility. Kinesins have ATPase activ-

ity, generate movement through the motor domain (Vale and Fletterick,

1997), and are classified according to its position within the proteins

(Miki et al., 2005). These structural features led to the separation of kinesins

into 15 different families (Hirokawa et al., 2009). In addition to the motor

domain, all kinesins have one or more coiled-coil domains. Depending on

the kinesin family, they can also have a CAP-Gly domain, a pleckstrin

homology (PH) domain, a Phox homology (PX) domain, and WD40

repeats (Hirokawa et al., 2009). Any kinesin that does not have a dis-

tinguishing feature falls into the orphan kinesin group (Fig. 2.3; Miki

et al., 2005). So far, kinesins have been involved in many cellular functions

such as organization of the interphase MT cytoskeleton, axonal transport,

organelle movement, and mitosis.

Some kinesin-like proteins have already been described to tip-track

MTs. CENP-E was described to localize to the plus ends of MTs, where

it promotes their elongation, possibly by stabilizing a straight-end conforma-

tion, which favors tubulin addition to the plus end (Sardar et al., 2010). In

theory, all plus-end-directed motors could concentrate on MT plus ends

due to their function, but most of them do not. This probably happens

because they have to interact with otherþTIPs or, in alternative, must show

some specificity for the MT plus end to do so (Bieling et al., 2007; Busch

et al., 2004). In fact, the yeast kinesin Tea2 needs to interact with Mal3

(the EB-like homologue) to track MT plus ends and to stimulate its ATPase

activity (Bieling et al., 2007; Browning and Hackney, 2005; Busch and


Brunner, 2004). The same behavior was described for kinesin-5 Eg5, which

interacts with EB proteins through a classical SxIP motif (Jiang et al., 2012)

and for kinesin-8 Kif18B which does not contain a canonical SxIP motif but

has similar sequences (Stout et al., 2011).

3.5.2 DyneinDynein is a large macromolecular complex with a molecular weight of

approximately 1.2 MDa. It is composed of heavy intermediate, light inter-

mediate, and light chains. The heavy chains contain the motor domains with

six AAA ATPase domains and an MT-binding stalk (Fig. 2.3; Oiwa and

Sakakibara, 2005). Dynein is a minus-end-directed motor that uses ATP

hydrolysis to power its movement and requires interaction with the dynactin

complex. One of the subunits of the dynactin complex is p150glued. This

protein is a þTIP that has a CAP-Gly domain and two coiled-coil regions

which are required for dimerization and interaction with the dynein inter-

mediate chain (King et al., 2003). Early reports of dynein accumulation on

MT plus ends came from work with the filamentous fungus Aspergillus. In

this organism, dynein exhibits plus-end-directed movement at velocities

similar to MT polymerization rates, which suggests that dynein is associated

to, and moving with, the polymerizing ends of MTs (Xiang et al., 2000).

Subsequent reports described the accumulation of both dynein and NUDF

(the homologue of Lis1) at MT plus ends in a comet-like structure (Zhang

et al., 2003). In the same system, dynein and dynactin required each other for

plus-end accumulation but NUDF specifically required dynein to tip-track.

After arriving at the plus ends, dynein also exhibits some retrograde move-

ment and this movement is also MT dependent (Xiang et al., 2000). The

interaction of dynein with LIS1 is important for dynein-mediated retrograde

transport because it allows the release of the dynein–dynactin complex from

CLIP170-decorated MT plus ends (Lansbergen et al., 2004). In vitro work

estimated that the dynein comet consists of approximately 55 dyneinmotors.

About half of the motors show a slow turnover and are actively kept at the

plus ends by a retention mechanism that requires interaction with dynactin

and EB1 (Schuster et al., 2011). Therefore, dynein retention at the plus ends

involves a combination of both stochastic accumulation and active retention

to allow formation of the dynein comet and ensure capturing of organelles

by minus-end-directed motors (Schuster et al., 2011).

During mitosis, dynein localizes at the cell cortex (Kiyomitsu and

Cheeseman, 2012; O’Connell and Wang, 2000). In yeast, it was proposed

that dynein offloads directly from the MT plus ends to the cell cortex by an


active MT-mediated delivery (Markus and Lee, 2011), a process which

requires the neck region of dynein. Longer neck regions allow enhanced

off-loading without affecting motor activity, while shorter necks block

delivery to the cortex. This led to the proposal that a conformational change

in dynein could be regulated by a masking/unmasking event that controls

dynein off-loading from MTs. Moreover, the N-terminal tail domain is

essential for targeting dynein to cortical receptor sites, whereas the

C-terminal domain is required for plus-end targeting in a Bik1/CLIP170-

and Pac1/LIS1-dependent manner (Markus et al., 2009). Curiously, expres-

sion of the motor domain alone blocks the MT plus-end accumulation of

dynein, and this can be rescued by overexpression of LIS1.

Additional dynein functions include centrosome separation and nuclear

translocation (Tsai et al., 2007). Dynein and Lis1 appear to generate tension

between the nucleus and the centrosome (Tanaka et al., 2004) and also at the

interface between MT tips and the cell cortex (Dujardin et al., 2003). Inter-

estingly, the role of dynein in nuclear movement appears to be conserved in

different cell types. Both dynein and kinesin seem to be required for the

bidirectional movement of the nucleus by interacting with the nuclear pore

complex. Interaction of dynein or kinesin-1 with Bicaudal D2 is essential for

nuclear and centrosomal position during mitotic entry (Splinter et al., 2010).

This may also involve the interaction of dynein/dynactin with a CENP-F–

NudE/EL–Nup133 complex (Bolhy et al., 2011).

3.6. Lis1Lissencephaly 1 (Lis1) proteins were first described as the result of a mutation

that leads to severe defects in brain development in humans (Dobyns et al.,

1993; Vallee et al., 2001). So far, many orthologs have been identified from

yeast (Geiser et al., 1997) to Caenorhabditis elegans (Dawe et al., 2001) and

Drosophila (Sheffield et al., 2000). Sequences from all orthologs are highly

conserved, suggesting a functional conservation. In structural terms, Lis1

proteins have three distinct regions (Fig. 2.3). The N-terminal region is

called LIS1-homology motif (LisH), which ranges between residues 1–39

and has been recently recognized as an ubiquitous motif, found in another

114 eukaryotic proteins (Emes and Ponting, 2001; Kim et al., 2004). The

region between amino acids 40–85 is predicted to be a coiled-coil region

which, together with the LisH domain, is involved in dimerization (Tai

et al., 2002). Near the C-terminal region, there are seven WD40 repeats

which range from amino acids 96–410 containing a b propeller domain,


which are important for lateral interactions with other proteins (Tarricone

et al., 2004).

Although it can act as a þTIP, Lis1 seems to target MTs by WD40-

mediated binding to CLIP170, dynein, and dynactin, rather than binding

the plus ends directly (Coquelle et al., 2002; Tai et al., 2002). The interac-

tion of Lis1 with CLIP170 is positively regulated by phosphorylation

(Coquelle et al., 2002). During mitosis Lis1 is recruited to the cell cortex

and kinetochores in a dynein/dynactin-dependent manner (Coquelle

et al., 2002; Faulkner et al., 2000). The C-terminal WD40 repeat region

of Lis1 seems to be sufficient for kinetochore targeting (Tai et al., 2002).

When overexpressed, Lis1 induces a displacement of CLIP170 from the

kinetochores but also interferes with spindle orientation and mitotic pro-

gression (Faulkner et al., 2000; Tai et al., 2002; Vallee et al., 2001).

3.7. Kinesin-13 familyMembers of the kinesin-13 family were named so because of the position of

the motor domain in the middle of the protein. The first 12 families (from

kinesin-1 to -12) have the motor domain close to the N-terminal region,

and kinesin-14 has the motor domain in the C-terminal region (Lawrence

et al., 2004; Miki et al., 2005). These kinesin-13 members were also initially

named M kinesin family (for “Middle Type Motor”) or KinI family (for

“InternalTypeMotor”).Within thekinesin-13 family, there are two subfam-

ilies: the ubiquitous KIF24 subfamily and the mammalian-specific KIF2 sub-

family. This last subfamily is comprised of three members: Kif2A, Kif2B, and

Kif2C/MCAK. All members of the family have an N-terminal globular

domain, followed by a positively charged neck upstream of the centrally

located catalytic core, and a C-terminal dimerization domain (Fig. 2.3;

Ogawa et al., 2004;Wordeman, 2005). TheKIF24 subfamily has the catalytic

core close to the N-terminal region, whereas the KIF2 subfamily has the cat-

alytic core closer to the center of the molecule (Miki et al., 2005). Interest-

ingly, it was demonstrated that MCAK requires dimerization through the

coiled-coil domain in the C-terminal region, and this has a role in regulating

the ATPase activity of the protein (Ems-McClung et al., 2007).

Members of the kinesin-13 family have been implicated in vesicle trans-

port (Noda et al., 1995) and, more importantly, in MT depolymerization

(Desai et al., 1999; Manning et al., 2007; Mennella et al., 2005; Walczak,

2003). Upon binding to the MT end, they induce a conformational change

in its structure that leads to a catastrophe event (Desai et al., 1999). The

MT-destabilizing properties of kinesin-13 members are unique because they


use ATP hydrolysis to induce depolymerization of MTs from both ends,

instead of using it to walk along MTs (Desai et al., 1999; Helenius et al.,

2006; Hunter et al., 2003). The best studiedmember of the family isMCAK.

This þTIP was shown to target the plus ends of MTs and, once there,

14 MCAK dimers form an ATP-hydrolyzing complex that processively

depolymerizes MTs (Hunter et al., 2003). For this reason, MCAK was

described as a major MT remodeler by preventing MT aging and inducing

random catastrophes (Gardner et al., 2012).

One puzzling observation comes from the fact that these proteins, while

having potent MT depolymerization activity, are still able to accumulate in

the plus ends of MTs (Moore et al., 2005). This suggests that the

MT-depolymerizing activity must be inhibited or controlled at this location

and raises the question of how does MCAK reach the MT tip. Microscopy

studies using single molecules demonstrated that MCAK rapidly moves

along the MT lattice in a random walk (Helenius et al., 2006). Contrary

to its requirement for the MT-depolymerizing effect, this diffusion does

not require ATP hydrolysis and is more rapid than direct binding to the plus

end from solution (Helenius et al., 2006). In addition to this, MCAK also

associates with EB proteins. In fact, MCAK associates with the

C-terminal region of both EB1 and EB3 and colocalizes with EB1 at MT

plus ends (Lee et al., 2008; Montenegro Gouveia et al., 2010). This raises

the possibility that MCAK could also use an EB-hitchhiking mechanism

to bind MT plus ends, in addition to lattice diffusion. These were proposed

as complementary mechanisms that would allow MCAK to remain associ-

ated with MT even after EB displacement from the plus end. Recent work

demonstrated that MCAK contains an SxIP motif near its C-terminal

domain that is crucial for associating with EB1 (Honnappa et al., 2009). This

property of MCAK seems to be conserved with other kinesin-13 proteins in

Drosophila such as KLP10A, which associates with EB1 and is necessary for

KLP10A targeting to MT plus ends (Mennella et al., 2005). The association

of MCAK with MTs can also be regulated in a posttranslational manner.

Indeed, Aurora-B was shown to phosphorylate MCAK and this is crucial

for its function (Andrews et al., 2004; Lan et al., 2004). In addition, most

of these phosphorylation sites seem to cluster in a region close to the SxIP

motif, which alters the ability of MCAK to interact with EB1 and tip-track

(Honnappa et al., 2009; Moore et al., 2005). Curiously, the other family

members Kif2A and Kif2B do not accumulate at MT plus ends, and this

is explained by the fact that they do not have an SxIP motif.


3.8. TOG familyThe tumor overexpressed gene (TOG) proteins belong to a highly con-

served family involved in MT dynamics regulation (Al-Bassam and

Chang, 2011; Slep, 2009). The founding member of this family, XMAP215,

was identified in Xenopus as a protein that promotes rapid MT growth (Gard

and Kirschner, 1987). Many orthologues have been described, including

ch-TOG in humans (Charrasse et al., 1998). In structural terms, these pro-

teins are characterized by the presence of a variable number of TOG

domains near the N-terminal region (Fig. 2.3). These domains have approx-

imately 200 amino acids and comprise between 2–5 units, depending on the

organism (Ohkura et al., 2001). Interestingly, each of these domains contains

several HEAT repeats, which are thought to mediate protein–protein inter-

actions (Cassimeris et al., 2001). The human ch-TOG contains five TOG

domains near theN-terminus, regions with sequences rich in serine, glycine,

and lysine (SK-rich domains) and a conserved C-terminal nonrepeat domain

(Al-Bassam and Chang, 2011). Interestingly, CLASPs also have TOG

domains and SR-rich regions, which provide a structural link between

the possible functions of both classes of proteins (Lemos et al., 2000; Slep,

2010). Detailed studies revealed that the N-terminal domain contains

an MT-stabilizing region, whereas the C-terminal domain is necessary for

centrosome and MT targeting (Popov et al., 2001).

TOG proteins not only localize to MT plus ends but can also bind the

MT lattice and soluble tubulin. They have an intrinsic ability to promote

MT elongation from both ends although they do so more efficiently on

the plus ends (Gard and Kirschner, 1987; Vasquez et al., 1994). In vitro stud-

ies with recombinant XMAP215 confirmed that these molecules can asso-

ciate directly to MT plus ends, stimulating their growth (Brouhard et al.,

2008; Kinoshita et al., 2001). These studies further demonstrated that

XMAP215 transiently binds the MT plus end and adds 25 tubulin dimers

to MT before dissociating (Brouhard et al., 2008). The initial hypothesis

for XMAP215 action involved the binding and recruitment of tubulin olig-

omers to MT ends (Cassimeris et al., 2001). However, later it became clear

that TOG proteins can only bind one tubulin dimer at a time (Al-Bassam

et al., 2006; Brouhard et al., 2008). Curiously, in Xenopus egg extracts,

the N-terminal region is able to stimulate MT growth at the plus ends by

inhibiting catastrophes, while the C-terminal region suppresses MT growth

by promoting catastrophes (Popov et al., 2001). Additional studies in differ-

ent systems further confirmed the role of TOG proteins in MT growth and


stabilization (Charrasse et al., 1998; Dionne et al., 2000; Tournebize et al.,

2000). ch-TOG has been shown to promote MT assembly both in solution

and from nucleation centers (Charrasse et al., 1998), and to be essential for

the formation of taxol-induced asters in human mitotic extracts (Dionne

et al., 2000). In vivo, these proteins increase MT growth and knockdown

of the proteins is associated with short interphase MTs, reduced growth

rates, and increased catastrophes and pauses (Brittle and Ohkura, 2005;

Cullen et al., 1999; Tournebize et al., 2000; Wang and Huffaker, 1997).

It was proposed that the stabilizing effect of these proteins might be due

to their interaction with MT-destabilizing proteins. In fact, XMAP215

seems to stabilize MTs by opposing the action of destabilizers such as

XKCM1 (the Xenopus homologue of MCAK).

3.9. Other þTIPsIt is well known that many þTIPs require interaction with EB proteins in

order to localize to the MT plus end (Akhmanova and Steinmetz, 2008).

The discovery, that conserved SxIP motifs are sufficient to target these pro-

teins to the plus ends (Honnappa et al., 2009), has allowed for the screening

and identification of an ever increasing number ofþTIPs (Jiang et al., 2012).

Examples of some of theseþTIPs include the stromal interactionmolecule 1,

which exhibits EB1-dependent tip-tracking behavior (Grigoriev et al.,

2008) and is involved in ER remodeling. Similarly, navigators were

described to associate withMT plus ends and to be important for cytoskeletal

reorganization (Martinez-Lopez et al., 2005; van Haren et al., 2009). In

addition, þTIPs which are involved in MT organization such as tastin

and DDA3 also have SxIP motifs (Jiang et al., 2012; Zhang et al., 2013).

Both tastin and DDA3, unlike the majority of other þTIPs, also have the

ability to track depolymerizing MTs. Surprisingly, among the new SxIP-

containing proteins, there were also membrane-associated þTIPs such as

AMER2/FAM123A, which was originally described as an APC-binding

protein (Grohmann et al., 2007), and kinases such as TTBK1 and TTBK2,

which are involved in the phosphorylation of MT-associated tau (Houlden

et al., 2007; Sato et al., 2006).

On the other hand, there areþTIPs that do not seem to interact directly

with EB proteins but are able to tip-track nonetheless. Two of such þTIPs

are Astrin and Kinastrin. Astrin was originally identified as a mitotic,

MAP (Mack and Compton, 2001). Recently, it was shown that Astrin

can bind to MT plus ends by associating with its interactor Kinastrin


(Dunsch et al., 2011). Once at the plus ends, this Astrin/Kinastrin complex

can induce MT polymerization, possibly by stabilizing the plus ends.

4. RECOGNITION OF MICROTUBULE PLUSENDS BY +TIPs

Accumulation at the MT plus ends is what defines a þTIP. Although

their localization can be confined to a small terminal region of the MT, they

employ different mechanisms to recognize and move along MTs. This sec-

tion will focus on howþTIPs recognize the plus end and how they are able

to move along the MT lattice. Tip-tracking behavior implies that þTIPs

must either have the ability to directly bind tubulin orMTs or, in alternative,

be recruited indirectly through binding to other factors. The fact that many

different classes of proteins can exhibit tip-tracking (Table 2.1) led to the

proposal of four models to account for this behavior: end binding, copoly-

merization, directed transport, and hitchhiking (Fig. 2.5). Curiously, it

seems that the same þTIP can exhibit different behaviors depending on

the conditions or organism: for example, in mammalian cells, APC can

be loaded to the plus ends in an EB1-dependent manner (Slep et al.,

2005), can tip-track autonomously (Kita et al., 2006), or can do so by asso-

ciating with kinesin-2 (Jimbo et al., 2002). On the other hand, loading of

CLIP170 to plus ends can be mediated by motors in yeast (Busch et al.,

2004; Carvalho et al., 2004; Maekawa and Schiebel, 2004), whereas in

mammalian cells, it involves direct binding and treadmilling on MT plus

ends (Perez et al., 1999).

4.1. Recognizing the microtubule plus endHow is it that some þTIPs such as EB proteins are able to directly associate

to the growing end ofMTs? This question is of great importance because EB

proteins are responsible for loading the majority of other þTIPs, including

CLIPs, CLASP, and APC (Lansbergen and Akhmanova, 2006), and can

influence drastic changes in MT dynamics. This means that they must rec-

ognize specific features on plus ends that are different from the lattice

(Fig. 2.5). The first obvious hypothesis is the GTP cap itself. Recently, it

was reported that introducing GTPgS (a slowly hydrolysable form of GTP)

on plus ends mimicked the EB-binding site (Maurer et al., 2011). This is in

line with the finding that EB1 can recognize the nucleotide state of tubulin

independently of its location. Under these conditions, EB1 recognizes the

GMPCPP MT lattice as opposed to the GDP lattice (Zanic et al., 2009).


A different study revealed that EBs can recognize the nucleotide state of the

plus end and this is crucial both for EB binding and for stabilizing a structural

cap that protects MT from depolymerization (Maurer et al., 2012).

Although it is tempting to assume that the nucleotide state of tubulin alone

is sufficient to determine plus-end binding, there is evidence that argues

against such a simple model. In fact, the GTP cap size is thought to be very

small when compared to the region decorated by the EB comet. Typical

comets can vary between 0.5 and 3 mm in length, depending on the growth

rate but not þTIP concentration (Bieling et al., 2007). This means that

comets have to encompass several hundreds or thousands of tubulin sub-

units, which is much bigger than the presumable GTP cap size (Caplow

and Shanks, 1996; Seetapun et al., 2012; Walker et al., 1991). It should

be noted, however, that recent studies propose the existence of longer

GTP caps that exhibit dynamic behavior and could partly account for this

discrepancy (Schek et al., 2007).

Additionally, it was suggested that EB1, instead of binding the

protofilaments themselves (Maurer et al., 2012), could bind to tubulin while

still in the sheet conformation (Vitre et al., 2008). Thismeans that EB1would

promote sheet closure and bind to theMT seam instead of the protofilaments

(Vitre et al., 2008). In fact,Mal3, the EB1homologue,was reported to act as a

Kinesin-mediatedtransport

Lateral diffusion

Direct recognition of plus-end-specific structure

Copolymerizationwith tubulin

Hitchhiking

Plus-end-directed kinesin

Autonomous +TIP

+TIP with partner-binding plus-end tracking

Tubulin dimer

Figure 2.5 Mechanisms of plus-end recognition by þTIPs. þTIPs can arrive at the plusend by lateral diffusion along the microtubule lattice or diffusion from the cytoplasm. Inalternative, they can be transported by kinesins or associate with the growing end ofmicrotubules by attaching to another þTIP (hitchhiking). Some þTIPs can recognizespecial structural features of the plus ends of microtubules, or they may copolymerizewith tubulin dimers or oligomers. Adapted with permission fromMacmillan Publishers Ltd:Nature Reviews Molecular Cell Biology (A Akhmanova and MO Steinmetz; Tracking theends: a dynamic protein network controls the fate of microtubule tips), copyright (2008).


molecular zipper by binding to the seam and leading to changes inMT struc-

ture (desGeorges et al., 2008; Sandblad et al., 2006).Overall these results indi-

cate that the nucleotide state of tubulin plays an important role in plus-end

binding, but there may be additional mechanisms that contribute to þTIP-

MT association. An alternative explanation for specific EB association to the

plus end could depend on the electrostatic interactions between the

C-terminal portion of EB1 and theMT lattice (Buey et al., 2011). In this case,

long-range electrostatic repulsive interactions between the C-terminus of

EB1 and theMT latticemay be able to drive accumulation of EBs on growing

MT ends. In fact, replacing the negatively charged C-terminal portion for a

neutral coiled-coil increased the dwell time of EB1 onMTwithout affecting

interaction with the plus end. Other possible mechanisms may involve the

posttranslational modification of EB proteins themselves. Actually, recent

reports demonstrated that phosphorylation of EB proteins might have an

important role in their association to the plus end. One study described a

mutation on the linker region of Mal3 that is sufficient to reduce the affinity

of the protein forMTs (Iimori et al., 2012), while another demonstrated that

phosphorylation of Bim1p by Aurora/Ipl1p was sufficient to remove Bim1p

from static and dynamic MTs (Zimniak et al., 2009). Curiously, the same

study indicates that both dimerization of Bim1p and the presence of the linker

domain are required for efficient tip-tracking.

4.2. CopolymerizationIn addition to recognizing MT plus ends, someþTIPs such as CLIP170 also

have the ability to directly bind tubulin subunits (Fig. 2.5) (Arnal et al., 2004;

Folker et al., 2005). This suggests that, in order to tip-track, these proteins

copolymerize with tubulin into MT and then quickly dissociate from the

“older” part of MTs as it grows (Akhmanova and Hoogenraad, 2005).

Moreover, these þTIPs must have a higher affinity for free GTP-tubulin

subunits than the GTP or GDP polymer itself. Association of CLIP170 with

free GTP-tubulin subunits is thought to occur through its CAP-Gly domain

which is able to bind directly the EEY/F motif on the C-terminal a-tubulintail (Mishima et al., 2007). Interestingly, the CAP-Gly domain of CLIP170

also interacts with EB1 and explains how it recognizes a composite binding

site on MTs plus ends composed of EB1 (including its C-terminal tyrosine)

and tyrosinated a-tubulin (Bieling et al., 2008; Mishima et al., 2007). Taken

together, these results provide a model for copolymerization of CLIP170

with tubulin, but they do not explain how CLIP170 dissociates from the

growing MT.


While copolymerization seems to explain CLIP170 behavior, it falls

short of explaining general þTIP behavior. First, other þTIPs such as

EB1 do not use the copolymerizationmechanism. In fact, EB1 seems to bind

the tubulin polymer but not the individual subunits (Gache et al., 2005).

Moreover, using reconstituted in vitro systems, it was possible to demonstrate

that the yeast EB-like protein Mal3 did not bind tubulin subunits and that

accumulation of Mal3 on plus ends was independent of tubulin concentra-

tion (Bieling et al., 2007). Furthermore, in vitro systems were able to recreate

plus-end tracking without the presence of exogenous enzymes, which

means that tip-tracking is independent of GTP or GDP and argues against

its role in MT recognition (Bieling et al., 2007, 2008; Maurer et al., 2011).

Recent experiments using FRAP demonstrated that þTIPs associate

very transiently with the plus end of MTs (Dragestein et al., 2008;

Wittmann and Waterman-Storer, 2005). Interestingly, turnover measure-

ments of CLIP170 and EB3 demonstrated that they show diffusion at both

the plus and the minus ends of MTs, which is inconsistent with the copo-

lymerization model (Dragestein et al., 2008). Taken together, these results

argue against the role of copolymerization as the major contributor to plus-

end tracking.

4.3. Diffusion versus motor-based transportAccumulation of þTIPs does not necessarily involve direct binding to the

plus end in all situations. SometimesþTIPs will bind to the lattice and move

toward the plus end ofMTswhere they accumulate. To do so, these proteins

use two different mechanisms: diffusion and motor-based transport

(Fig. 2.5).

Diffusional motility is defined as a one-dimensional walk along the MT

lattice driven solely by thermal energy (Cooper and Wordeman, 2009).

Simple diffusion of molecules along an MT is a simple, low-energy mech-

anism that also has the advantage of allowing bidirectional movement. This

mechanism is represented by the same mathematical equations that define

Brownian motion although diffusion coefficients tend to be smaller (Ali

et al., 2007; Gestaut et al., 2008; Helenius et al., 2006). The first observations

of single-molecule diffusional motility on MTs were performed using non-

processive kinesin motors (Inoue et al., 2001; Okada and Hirokawa, 1999).

While kinesin motor proteins usually “walk” along MTs using ATP hydro-

lysis, they can sometimes show a “biased diffusion.” This has already been

demonstrated for a number of kinesins which include KIF1A, CENP-E,


Ncd, and Eg5 (Furuta and Toyoshima, 2008; Kim et al., 2008; Kwok et al.,

2006; Okada and Hirokawa, 1999). It has been proposed that this type of

motor protein motility occurs when the motor domain is not so tightly

bound to the MT. In accordance, in experiments where ADP is added

instead of ATP, these proteins exhibit a pure diffusional movement presum-

ably because of weaker binding to MT (Kwok et al., 2006; Okada and

Hirokawa, 1999). This type of diffusion appears to occur ubiquitously

and provides some advantages over motor-based movement. First, it makes

the system more flexible by allowing unbiased binding of proteins at both

MT ends. Interestingly, many MAPs (such as MCAK) that require localiza-

tion at both the plus and minus end ofMTs also use this mechanism (Oguchi

et al., 2011). Second, by making weaker attachments to the lattice, in theory

it could allow these proteins to overcome obstacles that may exist along MT

by jumping between protofilaments in a side-step manner (Wang et al.,

1995). Third, diffusion does not require ATP consumption to move pro-

teins. Finally, over short distances (<1 mm), diffusional motility is faster than

directed motility which may allow quicker delivery of molecules to the plus

ends (Cooper andWordeman, 2009). However, because it is a random pro-

cess, it is very ineffective over longer distances.

Contrary to unbiased diffusional motility, molecules can exhibit a direc-

tional movement on MTs that is dependent on motor proteins. This mech-

anism requires the action of kinesin motor proteins which can contribute to

plus end accumulation. However, kinesin action alone is not sufficient to

induce the formation of a comet, as theremust be some retentionmechanism

that allows the þTIP to remain associated with the plus end (Galjart and

Perez, 2003). In yeast, CLIP170 homologues Tip1 and Bik1 are transported

to the plus end by the action of kinesins Tea2 and Kip2, respectively (Busch

et al., 2004; Carvalho et al., 2004). Although not so common in mammals,

motor-mediated transport can also be observed for APC, which requires

kinesin-2 for plus-end accumulation (Jimbo et al., 2002). This accumulation

occurs if the motor transport velocity is higher than MT polymerization/

depolymerization velocity (Busch et al., 2004; Carvalho et al., 2004). Inter-

estingly,þTIPs that are transported bymotors can also track depolymerizing

MTs in a mechanism known as backtracking (Carvalho et al., 2004).

4.4. HitchhikingLoading þTIPs via a motor-based transport requires that they hitchhike on

motor proteins. However, mostþTIPs accumulate at plus ends indirectly by


hitchhiking on other þTIPs (Lansbergen and Akhmanova, 2006). This

implies that a core component of þTIPs must exist that is able to associate

to MTs independently of any other factor (Fig. 2.5). EB proteins and also

XMAP215/ch-TOG can perform this function because it has been shown

that they can autonomously track MT plus ends (Al-Bassam and Chang,

2011; Bieling et al., 2007, 2008; Brouhard et al., 2008; Gard and

Kirschner, 1987). Furthermore, in the case of EB proteins, they are known

to interact with most other plus-end-associated proteins (Akhmanova and

Steinmetz, 2008; Jiang et al., 2012).

The interaction of EBs with other þTIPs occurs through the EB

C-terminal domain, also known as EBH domain (Bu and Su, 2003).

Although all these EB-interacting proteins share the ability to tip-track,

there is no apparent functional similarity between them, raising the question

of whether there is any common feature that accounts for their behavior. It is

now known that these EB partner proteins rely on the SxIP motif, which is

embedded within basic and proline/serine-rich sequence regions, to bind to

EB proteins and hitchhike onMTs. The first report on the role of an Ile-Pro

peptide in þTIP interaction came from structural work on the EB1–APC

interaction (Honnappa et al., 2005). The authors demonstrated that this Ile-

Pro peptide was part of the APC region that bound EB1 and that mutating it

was sufficient to impair the interaction. Further studies revealed that many

þTIPs such as MCAK, CLASPs, APC, and ACF7/MACF1 have a similar

SxIP motif (Honnappa et al., 2009; Jiang et al., 2012). In addition, it was also

demonstrated that this SxIPmotif is sufficient to load theseþTIPs to theMT

plus ends through interaction with EB1. Two interesting observations were

derived from this study: it is possible to abolish the interaction between these

þTIPs and EB1 and, as a consequence, inhibit tip-tracking by simply mutat-

ing the SxIP motif (for instance, by substituting the Ile-Pro with Asp); it also

became possible to “transform” a protein into a þTIP by introducing the

SxIP motif in its amino acid sequence. Taken together, these findings allow

the establishment of a generalMT tip localization signal and create a unifying

mechanism for plus-end targeting.

In addition to SxIP motifs, proteins can also use the CAP-Gly domain to

interact with the plus ends of MTs. Proteins that have a CAP-Gly domain

were the first to be identified that exhibit tip-tracking behavior (Perez et al.,

1999), and these include CLIP170, CLIP115, and p150glued, among others

(Steinmetz and Akhmanova, 2008). The CAP-Gly domain is highly con-

served in eukaryotes, can exist in either single or multiple copies, and is

involved in the regulation of protein interactions and formation of protein


networks (Akhmanova and Steinmetz, 2008; Galjart, 2005). Structural data

derived from X-ray crystallographic analyses of these domains demonstrated

that the conserved motif GKNDG is essential for interaction with the

EEY/F motif of a-tubulin, EB1, and CLIP170 (Steinmetz and

Akhmanova, 2008; Weisbrich et al., 2007). CAP-Gly proteins are unable

to bind tubulin dimers that lack the C-terminal tyrosine of EEY/F (Peris

et al., 2006), and moreover, mutations in the Lys-Asn of the GKNDG or

the EEY/F motives are sufficient to abolish this interaction (Steinmetz

and Akhmanova, 2008; Weisbrich et al., 2007). These interactions with

CAP-Gly proteins have dissociation constants in the micromolar range,

which is similar to what is observed for the interaction of the C-terminal

of EB1with an APCC-terminal peptide and indicates they are very dynamic

(Honnappa et al., 2005, 2006; Mishima et al., 2007; Weisbrich et al., 2007).

Conceptually, the hitchhiking mechanism implies that þTIPs that use

this mechanism are not able to interact efficiently with tubulin or MTs,

but this is not always the case. Notably, proteins that contain CAP-Gly

domains can efficiently associate with tubulin (Dixit et al., 2009; Folker

et al., 2005; Mishima et al., 2007). In addition, although some proteins with

SxIP domains such as RhoGEF2 and melanophilin do not bind tubulin

directly, many others such as MCAK and CLASPs are able to do so

(Al-Bassam et al., 2010; Helenius et al., 2006; Rogers et al., 2004; Wu

et al., 2005). Although direct binding to tubulin or MTs circumvents the

necessity for hitchhiking, it seems that plus-end accumulation mainly

depends on the hitchhiking mechanism. In fact, CLIP170 requires EB1

to tip-track (Dixit et al., 2009) and both EB1 and EB3 enhance the binding

of CLIPs to the MT plus ends (Komarova et al., 2005). Interestingly,

hitchhiking seems to be necessary for the loading but not dissociation of

þTIPs. These results were based on in vitro observations that CLIP170

(which hitchhikes on EB1) remains associated with MT longer than EB1

itself (Dixit et al., 2009). This probably happens because CLIP170, besides

binding to EB1, is able to bind directly to the C-terminal tails of tubulin

(Mishima et al., 2007).

4.5. Turnover at microtubule plus endThe balance betweenMT association–dissociation must be tightly regulated

so thatþTIPs remain confined to the plus end. This is clearly observedwhen

þTIPs are overexpressed and label the entire MT lattice (Schwartz et al.,

1997; Tirnauer and Bierer, 2000). Dissociation of þTIPs from MT may

involve changes in the MT lattice (such as GTP hydrolysis) or a structural


change in the þTIP itself (Akhmanova and Hoogenraad, 2005). Another

convenient mechanism would be þTIP phosphorylation. In fact, CLIP170

association to MT is negatively regulated by phosphorylation (Rickard and

Kreis, 1991). So far, several kinases have been described to affect CLIP170

phosphorylation, including mTOR (Choi et al., 2002), Plk1 and CK2

(Li et al., 2010), and AMPK (Nakano et al., 2010). Surprisingly, inhibition

of mTOR decreases the binding of CLIP170 to MTs, whereas inhibition of

AMPK increases the binding of CLIP170 toMTs. These data emphasize that

multiple layers of regulation must exist that control association of CLIP170

to MT. Experiments with the EB-like protein in yeast cells have also

suggested phosphorylation as a possible mechanism for EB binding to

MTs in a cell-cycle-dependent manner (Iimori et al., 2012; Zimniak

et al., 2009). However, there is no evidence so far for a phosphoregulatory

mechanism that specifically controls association/dissociation of individual

molecules to the MT plus ends.

While the mechanisms that regulate þTIP association to MT are still

elusive, considerable progress has been made in defining how these proteins

turnover at the plus end. The first model proposed that þTIPs bind only

once to the MT plus end and then dissociate when the MT lattice becomes

“mature” (Carvalho et al., 2003; Galjart, 2005). This process is called

treadmilling because of the similarities with the behavior of tubulin subunits

within MT (Fig. 2.6). While it seems that þTIPs are moving along as MT

Microtubulegrowth

Rapid exchange betweenplus-end and cytoplasm

Association

Dissociation

Association+TIP treadmillingalong microtubule(1A) (1B)

(2)

Figure 2.6 Mechanisms of þTIP dissociation from the microtubule. A þTIP associateswith the microtubule (1A) and remains attached to the structure until the plus-end isconverted into a regular lattice and then dissociates (1B). A new þTIP can then bindto the new microtubule plus-end (2). This mechanism is known as treadmilling. Onthe other hand, þTIPs may exchange rapidly with the cytoplasmic pool at their bindingsites in the plus end, while these sites decay exponentially over time. This mechanism isknown as rapid exchange. Adapted with permission fromMacmillan Publishers Ltd: NatureReviews Molecular Cell Biology (A Akhmanova and MO Steinmetz; Tracking the ends: adynamic protein network controls the fate of microtubule tips), copyright (2008).


grows, they are, in fact, stationary and it is the addition of new þTIPs and

tubulin at the plus end that creates this optical illusion (Carvalho et al., 2003).

The treadmilling model also implies that fluorescence decay observed in the

comet’s tail is due to the dissociation of þTIPs as the MT matures. Initial

approaches using fluorescent speckle microscopy techniques proposed that

EB1 and CLIP170 probably used treadmilling onMT plus ends (Perez et al.,

1999; Tirnauer et al., 2002b; Waterman-Storer et al., 1998). However,

recent studies demonstrated that this is not the case. Based on single-

molecule studies and FRAP experiments on MT plus ends (Bieling et al.,

2007; Dragestein et al., 2008; Kumar et al., 2009), a model of fast exchange

was proposed (Fig. 2.6). This is supported by the observation that the MT

decoration time is much longer than the dwell time of single molecules of

Mal3 (Bieling et al., 2007), meaning that Mal3 molecules must continuously

turnover on the plus end. In addition, it was demonstrated that individual

þTIPs are very dynamic and can repeatedly bind to the same plus end with

low affinity. Accordingly, both CLIP170 and EB3 molecules exhibit rapid

turnover behavior on plus ends (Dragestein et al., 2008). This turnover

means that several molecules can attach to the same binding site on MT

and continuously exchange with the cytoplasmic pool, as was shown for

EB3 (Dragestein et al., 2008). Further studies confirmed that EB1 also

exhibited the same behavior (Dixit et al., 2009). Taken together, this means

that þTIP turnover is much higher than binding site turnover, which is in

disagreement with the treadmilling model. As a consequence, these exper-

iments show that accumulation of þTIPs in a comet-like structure depends

on the exponential decay of EB-binding sites in the MT. EB proteins bind

and dissociate very rapidly, which creates a large number of binding sites for

other þTIPs. This is in agreement with studies that demonstrate a necessity

of CLIP170 to bind simultaneously to EB1 and tubulin composite sites

(Bieling et al., 2008). In addition, other þTIPs also show a slower dissoci-

ation rate from theMT, when compared to EB proteins (Bieling et al., 2007;

Dragestein et al., 2008), which further supports the role of EBs in facilitating

the binding of þTIPs to MTs.

5. +TIPs IN MITOSIS

5.1. +TIPs in mitotic spindle organization and positioningThe transition from interphase to mitosis involves a dramatic reorganization

of the MT cytoskeleton. This is accompanied by an increase in MT dynam-

ics and an abrupt decrease inMT polymer level which tightly correlates with

NEB (Zhai et al., 1996). Moreover, mitotic MTs show increased


catastrophe frequencies and spend less time in the “paused” state (Belmont

et al., 1990; Rusan et al., 2001). Many of these changes appear to be con-

trolled by phosphorylation-dependent regulatory mechanisms. Accord-

ingly, CDK1 was shown to play a role in remodeling the MT

cytoskeleton, as adding active CDK1 to Xenopus extracts increases MT

dynamics to mitosis-like levels (Verde et al., 1990). Furthermore, CDK1

induces the depolymerization of interphase MTs when injected into mam-

malian cells and also leads to the destabilization ofMTs when added tomam-

malian cell lysates (Lamb et al., 1990; Lieuvin et al., 1994). Furthermore,

protein phosphatases PP1 and PP2A have been shown to differentially reg-

ulate MT dynamics (Tournebize et al., 1997). While PP1 is required for

transitions into and out of mitosis, PP2A is required to maintain a steady-

state spindle length by controlling the level of catastrophes.

Many different classes of þTIPs have been involved in mitotic spindle

organization, some of which are regulated by phosphorylation. Notably,

CLIP170 is necessary for establishment of spindle bipolarity by interacting

with dynein (Tanenbaum et al., 2008). Interestingly, CLIP170 association

to the MTs is regulated by phosphorylation (Choi et al., 2002; Rickard

and Kreis, 1991), although it is not known whether this has an impact on

spindle organization.

EB proteins have also been implicated in spindle organization. Both

immunofluorescence analyses and live imaging using GFP tagging showed

that EB1 is able to localize to the growing ends of MTs throughout mitosis

(Berrueta et al., 1998; Morrison et al., 1998; Piehl and Cassimeris, 2003).

More in-depth observations demonstrated that EB1 can target to kineto-

chores with attached growingMTs (Tirnauer et al., 2002a). The first reports

indicated that depletion of EB1 in Drosophila by RNAi leads to the forma-

tion of short spindles and short astral MTs (Rogers et al., 2002), but in mam-

malian cells, depletion of EB1 by RNAi does not seem to interfere with

spindle assembly (Bruning-Richardson et al., 2012; Draviam et al., 2006;

Ferreira et al., 2013). In Xenopus egg extracts, EB1 was involved in spindle

organization and chromosome segregation by interacting with XMAP215

(Kronja et al., 2009). In addition, EB1 is also involved in astral MT nucle-

ation/stabilization possibly by interacting with Kif18B (Stout et al., 2011;

Toyoshima and Nishida, 2007). Interestingly, the EB1-interactor APC is

hyperphosphorylated during mitosis, which suggests that its binding to

MTs is regulated by phosphorylation (Bhattacharjee et al., 1996). Moreover,

depletion of APC has been shown to compromise the formation of spindles

in Xenopus extracts (Dikovskaya et al., 2004), although direct evidence for


the role of APC in spindle assembly in animal somatic cells is still lacking.

APC is also involved in mitotic spindle positioning. When a mutated dom-

inant form of APC is expressed in cells or when APC is disrupted, the spindle

is displaced from the cell center (Draviam et al., 2006; Green and Kaplan,

2003). This was also observed when EB1, a known APC interactor, is

depleted from cells, and this correlates with a marked loss of astral MTs

(Draviam et al., 2006; Ferreira et al., 2013; Green et al., 2005;

Toyoshima and Nishida, 2007). Therefore, by stabilizing astral MTs, EB1

helps orient the spindle parallel to the cell-substrate and provides an addi-

tional link between the spindle and the cell cortex. This may be accom-

plished through the interaction between EB1 and the motor protein

Kif18B, a plus-end-directed kinesin that can modulate MT dynamics and

has been shown to regulate astral MT length in early mitosis (Stout et al.,

2011). Recently, EB1 was also shown to be required for spindle symmetry.

Upon injection of specific antibodies or a dominant-negative form of EB1 in

mitotic cells, the resulting daughter cells displayed unequal MT content, and

this correlated with an asymmetric spindle pole movement (Bruning-

Richardson et al., 2012). However, in this study, there was no significant

displacement of the spindle from the cell center.

Several studies have confirmed that both the localization pattern of

CLASPs and their role in mitotic spindle organization are conserved

between species (Inoue et al., 2000; Lemos et al., 2000; Maiato et al.,

2003a; Mimori-Kiyosue et al., 2006; Pereira et al., 2006). Studies in mam-

malian cells revealed that simultaneous depletion of both CLASPs resulted in

an increased mitotic index and a plethora of mitotic abnormalities including

misaligned chromosomes, shorter or collapsed bipolar spindles, as well as

multipolar and disorganized spindles (Logarinho et al., 2012; Maiato

et al., 2003a; Mimori-Kiyosue et al., 2006; Pereira et al., 2006). Further-

more, studies with cells derived from Clasp2 knockout mice demonstrated

that the absence of this protein per se results in a significant number of mitotic

abnormalities, enhancing the susceptibility for aneuploidy and chromosomal

instability (Pereira et al., 2006). These cells exhibit numerous spindle defects

that can be partially rescued by ectopic expression of CLASP1. Curiously,

when individual CLASPs are depleted from human cells by RNAi, mitotic

progression does not seem to be affected (Mimori-Kiyosue et al., 2005). This

is in agreement with the observation that removal of one of the CLASP par-

alogues does not affect localization of the other (Pereira et al., 2006). Taken

together, these observations suggest that CLASPs play, at least, partially

redundant roles in mitosis.


However, some specific mitotic roles can still be assigned to individual

CLASPs. In fact, association of CLASP1 (but not CLASP2) with the tau-

related protein MAP4 was described as important for maintaining spindle

position and defining the division axis in human cells (Samora et al.,

2011). This association serves two purposes: whereas CLASP1 is required

for astral MT capture at the cortex, MAP4 is necessary to prevent engage-

ment of excess dynein motors creating an equilibrium situation. Impor-

tantly, under these conditions, depletion of MAP4 specifically induces

spindle misorientation relative to the substrate without affecting astral

MT nucleation, suggesting that the presence of astral MTs per se might

not be sufficient for accurate spindle positioning. InC. elegans, CLASPs have

a partial redundant role in spindle positioning and astral MT regulation dur-

ing asymmetric cell division (Espiritu et al., 2012). In this system, simulta-

neous depletion of the CLASP homologue CLS-1 together with CLS-2 or

CLS3 induces displacement of the spindle, together with changes in spindle

length. These cells also have a reduced complement of astral MTs, which

accounts for the positioning phenotype. However, depletion of CLS-2

alone in C. elegans embryos leads to defects in chromosome biorientation

without inducing spindle displacement (Cheeseman et al., 2005). Notably,

under these conditions, chromosome biorientation could be rescued by

inhibiting astral MT pulling forces. Overall, these results strengthen the

importance of CLASPs for proper mitotic progression and promotion of

mitotic fidelity.

TheTOG family of proteins also plays an important role in spindle assem-

bly. In fact, XMAP215 is required for this process inXenopus extracts, and its

immunodepletion results in either absence of spindle formation or very short

spindles (Tournebize et al., 2000). In mammalian cells, ch-TOG seems to be

required for the organization of spindle poles but has only a minor role in the

stabilization of spindle MTs (Gergely et al., 2003). The mechanism of

ch-TOG-mediated MT stabilization is partly regulated by its interaction

with TACC3 (Gergely et al., 2003). In addition, it can also protect kineto-

chore–MTs from depolymerization by MCAK (Barr and Gergely, 2008).

Overall, ch-TOG contributes to spindle bipolarity by increasing MT length

and density, focusing MT minus ends at the spindle poles and maintaining

centrosome integrity (Cassimeris and Morabito, 2004).

Many motor proteins that act asþTIPs also have an essential role in spin-

dle organization. Dynein is a minus-end-directed motor that shows tip-

tracking behavior (Vaughan et al., 1999). Dynein can bind to MTs and

induce their stabilization by tethering the plus ends (Hendricks et al., 2012;


Steuer et al., 1990;Yoshida et al., 1985). Furthermore, cytoplasmic dynein also

localizes to the cell cortex and serves as an anchor for astralMTs (Busson et al.,

1998). This localization led to the hypothesis that dynein could be involved in

spindle positioning. In yeast, mutations in the dynein gene affect the move-

ment of the spindle into the budding daughter cell without affecting spindle

assembly or chromosome segregation (Li et al., 1993). Later work done in

mammalian cells showed that when the shape of epithelial cells is mechani-

cally manipulated during mitosis, the mitotic spindle will always align with

the longer cell axis (O’Connell and Wang, 2000) and this can be blocked

by inhibiting dynein. This led to the hypothesis that longer astral MTs, by

having a higher number of dynein motors, would be able to generate

increased forces on the spindle and align it with the long cell axis. This model

implies that either astral MTs are in contact with the actin cortex along their

entire length or that dyneinmotors can anchor to cytoplasmic complexes and

exert a pulling force on MTs, as was later also suggested for interphase cells

(Brodsky et al., 2007). Interestingly, dynein localization at the cortex seems to

depend on both spindle pole and chromosome-derived signals which affects

cortical force generation (Kiyomitsu and Cheeseman, 2012). Proximity of

spindle poles with the cortex displaces dynein to the opposite pole, which

results in spindle centering. Activity of Plk1 at the spindle poles is necessary

because it regulates the interaction between dynein–dynactin and the cortical

factors NuMA and LGN (Kiyomitsu andCheeseman, 2012). Furthermore, a

chromosome-derivedRanGTPgradient restricts the localization ofNuMA–

LGN to the lateral cortex which enforces the spindle orientation axis.

In addition, by using their minus-end-directedmotion, thesemotors exert

pulling forces that maintain spindle pole separation during mitosis (Laan et al.,

2012; Vaisberg et al., 1993) and transport different cargo to the centrosome

where they help maintain spindle pole integrity (Purohit et al., 1999;

Young et al., 2000). Interestingly, minus-end-directed motors can also bind

toMT ends such as theDrosophila kinesin-14Ncd (Goshima et al., 2005). This

accumulation occurs through interaction with EB1 and is thought to play a

role in the capture and transport of k-fibers along centrosomal MTs and help

to form a tightly focused bipolar spindle (Goshima et al., 2005).

Astrin has also been involved in mitotic progression and spindle assem-

bly. Its association to spindle MTs and kinetochores was shown to depend

on GSK3b-mediated phosphorylation (Cheng et al., 2008), as inhibition of

the kinase impairs Astrin accumulation and spindle formation. Additionally,

depletion of Astrin by RNAi in human cells also leads to the formation of

disordered spindles (Gruber et al., 2002). Overall, these results highlight the


importance of Astrin for the formation of a bipolar spindle, which could be

due to the ability of Astrin to regulate spindle MT dynamics (Dunsch

et al., 2011).

There are other proteins such as kinesin-13 family member MCAK that

also plays a role in spindle assembly. MCAK localizes to spindle poles, cen-

tromeres, kinetochores, plus ends of MTs, and also the cytoplasm during

mitosis (Ems-McClung and Walczak, 2010; Moore et al., 2005;

Wordeman and Mitchison, 1995). During early mitosis, MCAK is required

for bipolar spindle assembly. How does MCAK regulate spindle bipolarity?

It has been reported that knockdown of another kinesin-13 member Kif2A

by RNAi leads to a dramatic increase in the number of monopolar spindles

(Ganem and Compton, 2004). When cells depleted of Kif2A are codepleted

for MCAK or treated with low doses of nocodazole, spindle bipolarity is

restored (Ganem and Compton, 2004). This means that Kif2A and MCAK

must be acting on spindle bipolarity through their ability to regulate MT

dynamics. In fact, in extracts depleted of XKCM1 (the Xenopus homologue

of MCAK), there is a fourfold decrease in catastrophe frequencies, which

leads to the formation of very longMTs and assembly of a monopolar spindle

(Walczak et al., 1996). In addition, the EB1-associated pool of MCAK was

proposed to limit the length of MTs in the assembling mitotic spindle, thus

favoring the formation of robust kinetochore–MT attachments (Domnitz

et al., 2012).

Interestingly, excessive nucleation can also induce defects in spindle

positioning. In fact, when MCAK is depleted from HeLa cells, very long

astral MTs are produced (Rankin and Wordeman, 2010). This same effect

can be accomplished by treating cells with the MT-stabilizing drug taxol. As

a consequence, the spindle shows dramatic rocking inside the cell, which is

dependent on Myosin II (Rankin and Wordeman, 2010). During mitosis,

other kinesins are involved in spindle formation (Haraguchi et al., 2006),

chromosome congression (Kapoor et al., 2006;Wood et al., 1997), and inte-

rpolar MT sliding (Kapitein et al., 2005). Interpolar MT sliding is achieved

by kinesin-5/Eg5, which is a plus-end-directed motor that can also tether

MT plus ends (Jiang et al., 2012; Kapitein et al., 2005).

5.2. +TIPs at mitotic centrosomeIn addition to their tip-tracking ability, many þTIPs are also capable of

binding to or contribute to centrosome function. EB1 was first reported

to localize to centrosomes in Dictyostelium. In this system, EB1 localized


to MT-free isolated centrosomes. Moreover, EB1 is thought to be required

for initiation of spindle MT growth (Rehberg and Graf, 2002). In mamma-

lian cells, EB1 interacts with centrosomes independently ofMTs, through its

C-terminal domain (Louie et al., 2004). In addition, EB1 depletion leads to a

reduction in MT minus-end anchoring and delays MT regrowth from cen-

trosomes. More recently, it was shown that FOP (FGFR1 oncogene part-

ner) interacts with the C-terminal region of CAP350 and forms a

centrosomal complex necessary for MT anchoring. Interestingly, FOP is

required for EB1 centrosomal localization (Yan et al., 2006). This localiza-

tion could also be mediated by an interaction with CDK5RAP2. In fact, in

addition to its centrosomal localization, CDK5RAP2 exhibits a tip-tracking

behavior that depends on EB1 binding, through a basic and Ser-rich motif

(Fong et al., 2009). Moreover, CDK5RAP2 contains a centrosome-

targeting domain that has a high homology to the Motif 2 of Centrosomin

(CM2) and mediates the association with Pericentrin and AKAP450 (Fong

et al., 2009; Wang et al., 2010). Similar to EB1, APC also associates with

centrosomes (Louie et al., 2004). This interaction is mediated by the

N-terminal domain of APC (Louie et al., 2004; Tighe et al., 2001), although

the exact interaction sequence is not known.

The observation that CLASPs can accumulate at the centrosome suggests

a function at this level, but little is known about their role in this structure

(Maiato et al., 2003a; Pereira et al., 2006). In both HeLa and Drosophila S2

cells, following colchicine treatment, CLASPs were found to colocalize

with g-tubulin in an MT-independent manner (Lemos et al., 2000;

Maiato et al., 2003a). Drosophila CLASP hypomorphic mutants displayed

atypical MT morphology that correlated with an abnormal pattern of cen-

trosome separation (Lemos et al., 2000), in spite of the fact that these cen-

trosomes were still capable of MT nucleation. Recently, CLASPs were

shown to be required for spindle pole integrity after bipolarization in

response to traction forces exerted by the motor proteins CENP-E and

Kid during chromosome alignment, by recruiting ninein to the centrosome

(Logarinho et al., 2012). This mechanism explains why suppression of

CLASPs leads to an increase in the number of multipolar spindles.

The TOG family of proteins also plays a relevant role at the centrosome.

InDrosophila embryos, the centrosomal protein D-TACC is required to effi-

ciently recruit ch-TOG/Msps to centrosomes (Lee et al., 2001). The role of

ch-TOG in spindle organization was proposed to occur in multiple ways. In

human somatic cells, ch-TOG is thought to play a major role in organizing

spindle poles and a more minor role in stabilizing spindle MTs via an


interaction with TACC3 (Gergely et al., 2003). ch-TOG seems to be

required for centrosomal MT nucleation or stabilization, as absence of the

protein leads to both diminished assembly and less dynamic MTs (Barr

and Gergely, 2008). In addition, ch-TOG also acts by focusing MT minus

ends at the spindle poles ensuring centrosome integrity (Cassimeris and

Morabito, 2004) but also protects spindle MTs from MCAK activity at

the centrosome, which could lead to multipolar spindles (Holmfeldt

et al., 2004). The joint localization of ch-TOG with MCAK at the centro-

some, and subsequent centrosome stabilization, is regulated by Aurora-A

(De Luca et al., 2008). In fact, depletion of Aurora-A leads to an accumu-

lation of ch-TOG at spindle poles with a concomitant delocalization of

MCAK (De Luca et al., 2008).

Like other þTIPs, Astrin was also shown to localize to spindle poles

(Mack and Compton, 2001). Subsequently, it was reported that targeting

of Astrin to the centrosome during S and G2 phases of the cell cycle requires

its interaction with the centrosomal protein ninein (Cheng et al., 2007).

Interestingly, depleting Astrin by RNAi or inducing its mislocalization leads

to loss of spindle pole integrity and centriole disengagement (Cheng et al.,

2007; Thein et al., 2007).

5.3. +TIPs at kinetochoreManyþTIPs are also involved in the regulation of MT–kinetochore attach-

ments. Initial experiments with CLIP170 described its transient association

with prometaphase kinetochores, even before CLIP170 was shown to tip-

track (Dujardin et al., 1998). CLIP170 colocalizes with dynein and dynactin

at kinetochores and is required for the formation of robust k-fibers (Dujardin

et al., 1998). Subsequent studies determined that CLIP170 is necessary for

mitotic progression (Wieland et al., 2004) and that interfering with CLIP170

expression leads to defects in chromosome congression and a decrease in

the number of kinetochore–MT attachments (Tanenbaum et al., 2006).

However, this does not seem to affect MT dynamics or the stability of

kinetochore–MT attachments. These observations indicate that CLIP170

may help in the formation of kinetochore–MT attachments by mediating

the direct capture of MTs at the kinetochore (Tanenbaum et al., 2006).

Interestingly, this kinetochore–MT attachment mechanism may involve

phosphoregulation of CLIP170. Indeed, it was proposed that CK2-

mediated phosphorylation of CLIP170 is involved in its kinetochore local-

ization (Li et al., 2010). Moreover, Plk1 is necessary to enhance this


association with CK2. Overall, Plk1- and CK2-associated phosphorylations

of CLIP170 are necessary for the timely formation of kinetochore–MT

attachments during mitosis. However, the mitotic defects that were attrib-

uted to loss of CLIP170 in cultured cells were not confirmed in mouse

models of CLIP170 deficiency, which raises the possibility that loss of

CLIP170 per se is not essential for establishing kinetochore–MT attachments

(Akhmanova et al., 2005). Interestingly CLIP190, theDrosophila orthologue

of CLIP170, was also reported to localize to unattached kinetochores and

this was shown to be dynein/dynactin-dependent (Dzhindzhev et al., 2005).

Given the localization of CLASPs at the kinetochore, one can assume a

functional role in this structure. Kinetochore localization of CLASPs relies

on CENP-E, independently of its motor activity (Maffini et al., 2009;

Maiato and Logarinho, 2011). This kinetochore targeting requires the

C-terminal domain of CLASP1 and CLASP2 but is independent of MTs

or CLIP170 association (Maia et al., 2012; Maiato et al., 2003a; Mimori-

Kiyosue et al., 2006).

Evidence to support the critical role of CLASPs in the regulation of spin-

dleMT dynamics in mammalian cells initially surfaced after injection of anti-

CLASP1 antibodies in HeLa cells stably expressing GFP-a-tubulin (Maiato

et al., 2003a,b). In this situation, injection of anti-CLASP1 antibodies sig-

nificantly reduced or suppressed the typical oscillatory dynamic behavior

of k-fibers, resulting in spindle collapse. Additionally, the fact that CLASP1

accumulates at the outer corona region strongly argued for a role in the reg-

ulation of the kinetochore–MT interface. Amore detailed analysis of mature

k-fibers using FRAP revealed that, in cells depleted for Drosophila CLASP,

k-fibers were not able to flux (Maiato et al., 2005). When severed with a

laser, the fraction of the k-fiber that remained attached to the kinetochore

was unable to regrow, contrary to what happens in the wild-type control

cells. This provided conclusive evidence regarding the essential role of

Drosophila CLASP in the incorporation of tubulin subunits at the kineto-

chore level. This also explains why its absence results in progressively short

k-fibers through tubulin depolymerization at the minus end, leading to

bipolar spindle collapse (Maiato et al., 2005). Interestingly, spindle collapse

could be reverted by depleting KLP10A (a Drosophila kinesin-13 MT

depolymerizer), which prevents MT minus-end depolymerization (Buster

et al., 2007; Laycock et al., 2006; Matos et al., 2009).

In mammalian cells, depletion of CLASPs at the kinetochores caused a

considerable decrease of k-fiber poleward flux and turnover rates, increasing

their stability (Maffini et al., 2009; Manning et al., 2010). Thus, the short


spindles detected in mammalian cells upon CLASPs depletion can be related

to their particular function at the MT–kinetochore interface. In this way,

CENP-E-mediated recruitment of CLASPs to kinetochores is critical for

the rapid exchange of attached MTs, contributing to their instability. This

view is consistent with the decreased accumulation of CLASPs at kineto-

chores during anaphase and concomitant with the reduction of k-fiber

dynamics at anaphase onset when compared to prometaphase (Bakhoum

et al., 2009b; Gorbsky and Borisy, 1989; Gorbsky et al., 1987). Therefore,

it was proposed that during early mitosis an increase in kinetochore–MT

turnover would allow the correction of erroneous attachments. This balance

is achieved through a functional interaction between CLASPs and the MT

depolymerizer Kif2B, which localize to kinetochores during early mitosis

(Bakhoum et al., 2009b; Maffini et al., 2009; Manning et al., 2010). How-

ever, as cells go into metaphase, this complex is replaced by a CLASP1–

Astrin complex, which promotes k-fiber stability, chromosome alignment,

and SAC silencing. These different complexes appear to be mutually exclu-

sive, suggesting that their recruitment to kinetochores is sufficient to change

the dynamics of attached MTs. At the transition from metaphase to ana-

phase, CLASP levels at kinetochores are reduced via a dynein-dependent

minus-end-directed removal (Reis et al., 2009). It should be noted that

depletion of CLASPs does not seem to interfere with the targeting of other

proteins that might be involved in kinetochore–MT attachment, such as

CLIP170 or dynein (Maiato et al., 2002, 2003a).

EB1 localizes to the plus ends of polymerizing MTs, suggesting that it

may regulateMT dynamics duringmitosis (Tirnauer et al., 2002a). Although

EB1 was originally identified as an APC-interacting protein, its localization

is independent of APC (Berrueta et al., 1998; Morrison et al., 1998). Curi-

ously, the inverse is not true, as APC localization to the plus ends requires an

interaction with EB1 (Askham et al., 2000; Mimori-Kiyosue et al., 2000b).

Moreover, the interaction between APC and EB1 does not seem to be rel-

evant for EB1 mitotic localization, as immunoprecipitation studies demon-

strated that the EB1–APC interaction does not occur or is not detectable

during mitosis, possibly because of APC hyperphosphorylation (Askham

et al., 2000; Bhattacharjee et al., 1996; Nakamura et al., 2001). In Xenopus

meiotic extracts, both EB1 and APC interact with kinetochore-associated

BubR1 (Zhang et al., 2007a). In this system, BubR1 directly interacts with

APC and this is essential for chromosome positioning in the metaphase plate.

Curiously, earlier reports had already identified interaction of APC with

checkpoint proteins Bub1 and Bub3 at the kinetochore (Kaplan et al.,


2001). This may explain why cells carrying a truncated APC gene (Min)

show defects in chromosome segregation. Furthermore, loss of APC leads

to changes in mitotic progression associated with a decrease in metaphase

interkinetochore tension (Dikovskaya et al., 2007; Draviam et al., 2006).

This was accompanied by a decrease in the association of Bub1 and BubR1

with kinetochores, which suggests that APC may be, directly or indirectly,

involved in the loading of these proteins.

The minus-end-directed motor dynein can also be found at kineto-

chores. This localization is regulated by MT attachment to the kinetochores

but does not depend on tension (King et al., 2000). In fact, dynein binding to

the kinetochore is very sensitive, as “fewer than half the normal number of

kinetochore–MTs leads to the loss of most kinetochore–dynein” (King

et al., 2000). Furthermore, the association of dynein to kinetochores was

reported to depend on Spindly (Barisic et al., 2010; Chan et al., 2009;

Gassmann et al., 2010). A significant pool of kinetochore–dynein is regu-

lated by Plk1-mediated phosphorylation, as inhibiting Plk1 severely affects

dynein localization to the kinetochore without affecting dynactin or Zw10

(Bader et al., 2011). What could be the role of dynein at the kinetochore?

Dynein associates with kinetochores during prometaphase and, as a minus-

end-directed motor, generates a pulling force on MTs. By interfering with

dynein localization at the kinetochore, cells fail to achieve efficient chromo-

some alignment and exhibit problems in MT capture (Li et al., 2007; Yang

et al., 2007). For this reason, kinetochore–dynein was proposed to produce a

poleward force that brings monooriented kinetochores close to the pole,

which facilitates MT capture by the kinetochore and promotes chromosome

congression. Accordingly, depleting or inhibiting kinetochore–dynein pre-

vents the rapid poleward motion of attached kinetochores but does not

interfere with kinetochore fiber formation (Yang et al., 2007). In addition,

dynein also plays a role in stableMT attachment and kinetochore orientation

during metaphase, although its kinetochore levels are reduced at that stage

(Varma et al., 2008; Yang et al., 2007). This effect may be related to the abil-

ity of dynein to remove some kinetochore components during mitosis to

ensure MT stability as was shown for CLASPs (Reis et al., 2009). Interest-

ingly, kinetochore–dynein is also required for normal anaphase chromo-

some movement, but it remains unknown whether this is directly due to

its ATPase activity (Yang et al., 2007).

Kinesin-7/CENP-E is a plus-end-directed motor required for meta-

phase chromosome alignment (Kapoor et al., 2006; Wood et al., 1997).

Although CENP-E is not a conventionalþTIP, due to its plus-end-directed


movement, it appears to accumulate in the distal end of MTs in vitro (Sardar

et al., 2010) and in vivo (Cooke et al., 1997). In addition, CENP-E has crit-

ical roles during mitosis including kinetochore–MT attachment and move-

ment of chromosomes to the metaphase plate (Cooke et al., 1997; Kapoor

et al., 2006; Schaar et al., 1997; Wood et al., 1997).

In prophase, kinesin-13 MCAK localizes to the inner kinetochore and,

during chromosome congression, MCAK specifically associates with the

leading kinetochore (Kline-Smith et al., 2004). Depletion or disruption of

MCAK leads to defects in alignment and segregation of chromosomes

(Kline-Smith et al., 2004; Wordeman et al., 2007; Zhu et al., 2005). These

defects may be the consequence of improper kinetochore–MT attachments

that arise whenMCAK is disrupted and lead to the formation of merotelic or

syntelic attachments. For this reason, MCAK was proposed to play a role in

the prevention and/or correction of kinetochore–MT attachments (Kline-

Smith et al., 2004). Additional work demonstrated that MCAK is required

for the turnover of k-fibers, contributing to the directional switching

between sister centromeres (Rizk et al., 2009; Wordeman et al., 2007).

The pool of MCAK that is associated with EB1 at the MT tips was proposed

to be important for the promotion of stable kinetochore–MT attachments in

an indirect way, by limiting nonkinetochore–MT length (Domnitz et al.,

2012). Overall, MCAK’s role would be to contribute to error correction

either by allowing the release of MTs from the kinetochore or by promoting

MT turnover. This process seems to be regulated by Aurora-B-mediated

phosphorylation of MCAK at serine 196, which inhibits MCAK

MT-depolymerizing activity (Andrews et al., 2004; Lan et al., 2004). Given

their role in controlling k-fiber turnover, it is not surprising that these proteins

can influence chromosome segregation. In fact,MCAKwas first shown to be

important for chromosome segregation, as introduction of a motorless,

dominant-negative version of the protein leads to lagging chromosomes

(Maney et al., 1998). Simultaneous knockdownofMCAKandKif2A induces

an even higher number of lagging chromosomes (Ganem et al., 2005). Inter-

estingly,MCAK activitymay also be regulated by interactionwith ch-TOG.

By forming a complex with TACC3 and clathrin, ch-TOG physically cross-

links k-fibers and reduces MT catastrophes (Barr and Gergely, 2008; Booth

et al., 2011).Moreover, these TACC3/ch-TOG/clathrin k-fiber bridges are

regulated by Aurora-A (Cheeseman et al., 2011).

In addition to its centrosomal localization, Astrin and its interactor

Kinastrin also localize at the kinetochores (Dunsch et al., 2011; Manning

et al., 2010; Schmidt et al., 2010). This localization is negatively regulated


by an Aurora-B-mediated phosphorylation so that Astrin only localizes to

bioriented kinetochores (Manning et al., 2010; Schmidt et al., 2010).

Accordingly, Astrin was shown to help stabilize kinetochore–MT attach-

ments and to promote mitotic progression (Dunsch et al., 2011; Gruber

et al., 2002; Manning et al., 2010; Schmidt et al., 2010). Importantly, the

plus-end localization of Astrin was reported to facilitate chromosome align-

ment (Dunsch et al., 2011). Moreover, depletion of Kinastrin induces the

same mitotic defects observed in Astrin-depleted cells.

5.4. +TIPs regulation during mitosisMany þTIPs exhibit different localization or behavior during mitosis. This

raises the question of how the different þTIPs are regulated in space and

time. CLIP170 was one of the first þTIPs described to be phosphorylated

in vivo at multiple sites (Choi et al., 2002). In this report, the authors iden-

tify an interaction of CLIP170 with FRAP kinase and treatment with

rapamycin interferes with the ability of CLIP170 to associate with MTs.

However, the same report describes several rapamycin-sensitive and -

insensitive phosphorylation sites, indicating there must be other kinases

regulating CLIP170 function. Accordingly, both Plk1 and CK2 have been

recently identified as CLIP170 kinases (Li et al., 2010). The CK2-mediated

phosphorylation is essential for kinetochore targeting of CLIP170 in a

dynactin-dependent manner. In this context, Plk1 seems to act as a priming

kinase, which enhances the ability of CLIP170 to bind to CK2. Expression

of phospho-null mutants of CLIP170 is sufficient to displace the protein

from the kinetochore and induce defects in the formation of k-fibers,

which further highlights the importance of CLIP170 phosphoregulation

(Li et al., 2010). This phosphoregulatory mechanism may also be relevant

to control CLIP170 association to the plus ends by inducing conforma-

tional changes in the protein. CLIP170 switches between two conforma-

tional states that alter its affinity for MTs (Lansbergen et al., 2004). In

its phosphorylated state, CLIP170 shows enhanced binding between the

N- and C-terminal domains and remains in a “closed” conformation

(Lee et al., 2010). This phosphorylated form of CLIP170 has lower affinity

for MTs and does not interact with p150glued. The phospho-null mutant

of CLIP170 is in an “open” conformation and has higher affinity for the

plus ends of MTs and p150glued (Lee et al., 2010). This leads to an auto-

inhibitory mechanism that confers tighter control of CLIP170 association

to the MT.


In a similar fashion, CLASPs localization during mitosis is also regulated

by phosphorylation, both in a direct and in an indirect manner. Astrin and

SKAP both bind MTs directly and are required for CLASP1 kinetochore

localization (Schmidt et al., 2010). However, the Astrin/SKAP complex tar-

gets only to bioriented kinetochores due to an Aurora-B-mediated regula-

tory mechanism. This provides a spatiotemporal control of kinetochore

composition through Aurora-B. In addition to this indirect regulation,

CLASP2 can also be directly phosphorylated during mitosis. Although

CLASP2 shows tip-tracking behavior in interphase, it is not easily detectable

at MT plus ends during metaphase. This raised the possibility that plus-end

localization could be under the regulation of specific mitotic kinases. Indeed,

priming phosphorylations of CLASP2 by CDK1 and subsequent GSK3bphosphorylation are required to induce CLASP2 displacement from the plus

ends of MTs during mitosis (Kumar et al., 2012). Interestingly, this tip-

tracking behavior depends on the interaction of CLASP2 with EB1, as

imposing the phosphorylations induced a disruption in the interaction

between the two proteins, leading to the displacement of CLASP2 from

the plus ends. Conversely, introducing phospho-null mutations on these

specific sites was sufficient to restore EB1 association and binding to the

plus end. More recently, it was shown that during mitosis CLASP2 is

predominantly phosphorylated at its C-terminal domain, close to the

kinetochore-associated and dimerization region (Maia et al., 2012). These

phosphorylations were mediated by CDK1 and Plk1 kinases at different sites

and were specific for CLASP2. Noteworthy, CLASP2 phosphorylation by

CDK1 acts as a priming event for further association of CLASP2 with Plk1.

In accordance, colocalization of CLASP2 and Plk1 was reported in the Golgi

apparatus, centrosomes, kinetochores (in an MT-independent way), spindle

midzone, and midbody. CDK1 phosphorylation of CLASP2 was required

not only to increase Plk1 levels at the kinetochore but was also necessary

to maintain spindle bipolarity. More specifically, CLASP2 phosphorylation

on serine 1234 by CDK1 was shown to stabilize kinetochore–MTs, as

expression of the respective phospho-null mutant of CLASP2 leads to the

formation of monopolar spindles due to k-fiber instability (Maia et al.,

2012). In this way, phosphorylation of CLASP2 by both CDK1 and Plk1

were shown to be important for proper kinetochore–MT attachment and

chromosome alignment.

Given their crucial role in the regulation of MT dynamics, surprisingly

little is known on how EB function is regulated. Much of the recent work

has focused on the budding or fission yeast homologues of EB1. The


budding yeast EB-like protein Bim1p is regulated by multisite phosphory-

lation by the Aurora-B homologue Ipl1p (Zimniak et al., 2009). This

EB-like protein forms a stable complex with Aurora-B, which then phos-

phorylates a serine cluster in the linker region of EB, and this phosphoryla-

tion is sufficient to reduce the affinity of the protein for MTs. On the other

hand, a mutation of the fission yeast EB-like protein Mal3 was reported to

increase the affinity of the protein to MTs (Iimori et al., 2012). When the

glutamine on position 89 in the CH domain was replaced with an arginine,

this EB no longer behaved as a þTIP but associated with the entire MT

lattice. This also prevented EB dissociation from MTs even when it was

not growing, leading to a reduction in the shrinkage rate. How do these

phosphorylations regulate the affinity of EB proteins for MTs? One may

consider that they affect the interaction of EBs with partner proteins such

as CLASP2, and this could impact on the overall affinity of EBs for the plus

end. On the other hand, phosphorylation of EB proteins could introduce

negative charges in the protein which would disrupt the association with

MTs through electrostatic repulsive interactions as was recently proposed

(Buey et al., 2011). Nevertheless, it seems plausible that phosphorylation

of EBs could lead to an overall decrease of interaction with the MT. In fact,

phosphorylation of EB3 was recently reported in human cells. During mito-

sis, EB3 was reported to be phosphorylated by Aurora kinases on serine 176

(Ban et al., 2009). This phosphorylation induces a stabilization of the protein

because it prevents its polyubiquitination and proteasome-mediated degra-

dation. Furthermore, EB3 phosphorylation by Aurora-B during mitosis was

shown to spatially regulate MT dynamics (Ferreira et al., 2013). Such a spa-

tial phosphorylation pattern allows distinct pools of EB3 to fine-tune MT

dynamics at different cellular locations.

Motor proteins are also subject to regulation during mitosis. Dynein was

shown to be phosphorylated during meiosis when added to Xenopus egg

extracts. This was dependent on CDK1 and occurred specifically after incu-

bation with metaphase but not interphase extracts (Dell et al., 2000). At least

partially, dynein phosphorylation could direct its binding to specific partners

or structures. Phosphorylation of dynein intermediate chain favors its asso-

ciation to Zw10 instead of dynactin, and this triggers dynein accumulation at

the kinetochore (Whyte et al., 2008). Interestingly, this association persists

until chromosomes become bioriented, which results in dynein dephos-

phorylation. Dephosphorylated dynein then associates preferentially with

dynactin and exhibits poleward streaming, which removes it from the kinet-

ochore (Whyte et al., 2008). In addition to CDK1, kinetochore–dynein is


also phosphorylated by Plk1 (Bader et al., 2011). Interfering with Plk1

induces mislocalization of dynein without significantly affecting dynactin

or Zw10, and this leads to defects in MT capture at the kinetochore.

MCAK is regulated by phosphorylation through several kinases, of

which Aurora-B seems to be the most significant (Tanenbaum and

Medema, 2011). Addition of phosphates in the N-terminal region of

MCAK, near the SxIP motif, seems to affect the tip-tracking behavior of

MCAK. Namely, Aurora-B-mediated phosphorylation of MCAK was pro-

posed to disrupt its interaction with theþTIPs TIP150 and EB1, decreasing

MCAK affinity for MT plus ends (Honnappa et al., 2009; Jiang et al., 2009;

Moore et al., 2005). Overall, these phosphorylations may contribute to a

decrease in the recruitment of MCAK to the plus ends of MTs, favoring

MT growth. In agreement, in vivo phosphorylation of the neck region of

MCAK varies according to the mitotic stage: it is high in early mitosis

but decreases when chromosomes become aligned and kinetochore–MT

attachments have to be stabilized (Lan et al., 2004). Many phosphorylation

sites regulate MCAK binding to spindle poles, kinetochores, centromeres,

and chromosome arms (Andrews et al., 2004; Lan et al., 2004; Zhang

et al., 2007b, 2008). Moreover, these phosphorylation events seem to affect

specific pools of MCAK. Accordingly, MCAK neck phosphorylation can be

found mainly at the centromere, whereas Aurora-B-mediated phosphoryla-

tion at serine 95 inhibits this localization. Strikingly, phosphorylation at ser-

ine 110 by Aurora-B increases centromere binding (Zhang et al., 2007b).

Taken together, these observations suggest multiple layers of regulation

depending on spatiotemporal constraints. A mitotic-specific phosphoryla-

tion of MCAK by CDK1 has also been reported (Sanhaji et al., 2010). This

modification inhibits MCAK’s MT-depolymerizing activity and can be

reproduced by expressing a phosphomimetic mutant. However, it is not

yet clear what the functional relevance of this modification is. In fact, if

MTs need to be more dynamic during mitosis, why should MCAK activity

be impaired at this stage? It may be that this phosphorylation (such as hap-

pens with Aurora-B-mediated phosphorylations) affects only a small pool of

MCAK, suggesting that there must be a local regulation of MT dynamics

during mitosis (Tanenbaum et al., 2011). In this regard, it is possible that

impairment of MCAK MT-depolymerizing activity on k-fibers alone

would allow them to stabilize, while active MCAK would still be acting

on nonkinetochore–MTs, allowing their faster turnover. Finally, the

C-terminal domain of MCAK, which affects its own MT depolymerase

activity, is phosphorylated by Plk1 (Moore and Wordeman, 2004; Zhang


et al., 2010). Unlike CDK1- and Aurora-B-mediated phosphorylations,

Plk1 phosphorylation promotes the MT-destabilizing activity of MCAK.

Thus, many layers of regulation ensure that the localization and activity

ofþTIPs during mitosis are tightly regulated to allow successful completion

of the process. While phosphorylation emerges as the major regulatory

mechanism to control þTIPs interactions, it remains unclear how all these

can be integrated to form a coherent picture.

5.5. +TIPs in mitotic exit and cytokinesisThe completion of mitosis involves the final separation of sister chromatids

into two daughter cells and partitioning of the cytoplasm. This last step

requires the formation of an actomyosin ring that will constrict MTs in

the midzone region (Fujiwara and Pollard, 1976; Schroeder, 1972, 1973).

Myosin function in the cytokinetic ring requires astral MTs to interact with

the cortex (Foe and von Dassow, 2008), but this does not seem to depend on

the precise regulation of MT dynamics. In fact, both MT stabilization and

increases in MT dynamics are able to induce furrow formation (Strickland

et al., 2005a). Changing of the midzone to midbody correlates with furrow

ingression, and when this is prevented, cells accumulate midzone-like MT

structures (Straight et al., 2003). Most MTs that compose the spindle mid-

body are antiparallel MTs that derive from the spindle midzone (Elad et al.,

2010; Euteneuer and McIntosh, 1980; Mullins and Biesele, 1977). As

opposed to earlier stages of cytokinesis, MTs are essential for completion

of the process (Savoian et al., 1999). Although midbody MTs are relatively

stable (Margolis et al., 1990), they can also show de novo nucleation. This

process may involve g-tubulin, which is also required for successful comple-

tion of cytokinesis (Julian et al., 1993; Shu et al., 1995). Live imaging of MT

plus ends with EB proteins also indicated that some midbody MTs are still

able to exhibit a highly dynamic behavior (Rosa et al., 2006).

Exit from mitosis requires the inactivation of CDK1. This inactivation

induces a reorganization of the MT cytoskeleton that includes increased

astral MT nucleation and midbody formation (Wheatley et al., 1997). In

addition to this more general role in MT organization, yeast CDK1 was

shown to control Aurora kinase by phosphorylating its N-terminal domain

(Zimniak et al., 2012). Interestingly, this phosphorylation blocks association

of Aurora with the yeast EB-like protein until anaphase onset. Association

between Aurora and Bim1p is required for Bim1p phosphorylation on its

linker region (Zimniak et al., 2009). This phosphorylation is necessary for


efficient EB tip-tracking and occurs specifically during anaphase as a means

to ensure normal spindle elongation and disassembly of the spindle midzone.

Therefore, if CDK1 fails to phosphorylate Aurora kinase, this leads to pre-

mature targeting of Aurora kinase to the spindle and constitutive EB phos-

phorylation, resulting in problems duringmitotic exit (Zimniak et al., 2012).

In human cells, EB proteins are tightly associated with midzone and mid-

body MTs (Berrueta et al., 1998; Morrison et al., 1998). Moreover, EB1

and Aurora-B were shown to colocalize in these same structures (Sun

et al., 2008). However, unlike in yeast cells, human EB1 is not a substrate

of Aurora-B but is required to enhance the kinase activity of Aurora-B. It

does so by preventing association of PP2A with Aurora-B and protecting it

from dephosphorylation (Sun et al., 2008). This is in apparent contradiction

with the studies performed in yeast but one must bear in mind that human

cells have more than one EB protein. Accordingly, in human cells, both

Aurora-A and Aurora-B were shown to phosphorylate EB3 during mitosis,

leading to EB3 stabilization (Ban et al., 2009). Interestingly, EB3 seems to be

a target of the recently described Aurora-B-mediated phosphorylation gra-

dient in late mitosis (Ferreira et al., 2013; Fuller et al., 2008). Accordingly,

Aurora-B-mediated phosphorylation of EB3 on serine 176 promotes MT

growth, stabilizing the midbody and allowing completion of cytokinesis

(Ferreira et al., 2013). Importantly, EB3 dephosphorylation near the cell

cortex restricts MT growth, which allows stabilization of focal adhesions

and daughter cell adhesion.

Taken together, the interactions between EB proteins and other mitotic

exit-related proteins highlight the importance of þTIPs in this context.

Accordingly, if formation of astral MTs is suppressed during anaphase by

interfering with either EB1 or dynactin, there is a significant delay in cyto-

kinesis (Strickland et al., 2005b). During anaphase, phosphorylated MCAK

localizes to the spindle midzone, and this is important because it helps reg-

ulate its MT depolymerization activity (Lan et al., 2004). The phosphory-

lation of MCAK is also carried out by Aurora-B (Fuller et al., 2008; Lan

et al., 2004). Recently, a new þTIP termed TIP150 was shown to localize

to the plus ends until anaphase B, in an EB1-dependent manner. TIP150 also

interacts with the MT depolymerase MCAK and appears to assist in the

EB1-mediated recruitment of MCAK to the plus ends (Jiang et al.,

2009). Interestingly, MCAK shares common cellular localizations with

EB1 and Aurora-B. Taken together, this means that, either directly or indi-

rectly, Aurora-B and CDK1 seem to regulate the localization or activity of

many EB1-associated proteins after anaphase onset and until cytokinesis.


Other EB-associated proteins such as APC or CLASPs were also shown

to independently regulate mitotic exit. During anaphase and telophase, both

CLASP1 and CLASP2 accumulate in the spindle midzone and midbody

(Maiato et al., 2003a; Mimori-Kiyosue et al., 2006). Given the localization

of CLASPs in the spindle midzone and midbody, one can predict a role for

these proteins during mitotic exit. In accordance, previous studies have

implicated CLASPs in cytokinesis in C. elegans, Drosophila weak hypo-

morphic mutants, and human cells (Inoue et al., 2004; Mimori-Kiyosue

et al., 2006; Pereira et al., 2006; Skop et al., 2004). More recently, CLASP1

recruitment to the central spindle was shown to be dependent on its asso-

ciation with PRC1 at anaphase B (Liu et al., 2009b). Furthermore, reduc-

tion in CLASP1 levels or interference with the CLASP1–PRC1 interaction

leads to a disorganization of the central spindle, due to a reduction in the

amount of antiparallel MTs, as well as a chromatin-bridge phenotype and

failures in accurate chromosome segregation (Liu et al., 2009b). These

observations suggest a role for the CLASP1–PRC1 complex in antiparallel

MT elongation and central spindle stabilization. Interestingly, PRC1 itself is

directly regulated by phosphorylation by CDK1 and Plk1, which puts it in

the right context for regulating mitotic exit (Hu et al., 2012; Jiang et al.,

1998). Due to the partial redundancy of CLASPs during mitosis, it is not

surprising that cells derived fromCLASP2 knockout mice also present a mild

cytokinetic phenotype (Pereira et al., 2006). The presence of chromatin

bridges in CLASP2-depleted cells can be pointed out as a cause of cytoki-

nesis failure, leading to the generation of polyploid cells with multiple

centrosomes (Pereira et al., 2006).

APC, one of the main interactors of EB1, has been extensively impli-

cated in cytokinesis completion. Reports on APC mutants demonstrated

that these cells become polyploid over time (Fodde et al., 2001; Kaplan

et al., 2001; Tighe et al., 2004), which suggests that APC plays a role in cyto-

kinesis. Although the different APC alleles behave in a distinct manner, it is

believed that they may interfere with anchoring of the mitotic spindle

(Caldwell et al., 2007). In fact, in a C-terminal-truncated mutant of

APC, MTs make fewer contacts with the cell cortex. For this reason, spin-

dles rotate excessively and this leads to cytokinetic failures (Caldwell et al.,

2007). Interestingly, both inMin mice and APC knockout mice, there is an

increase in the number of tetraploid cells, which is a hallmark of cytokinesis

failure (Caldwell et al., 2007; Dikovskaya et al., 2007).

Other non-EB1-associated proteins play important roles in postanaphase

cells. The kinesin CENP-E localizes to the midbody where it uses its


coiled-coil domain, to interact with Skp1 (Liu et al., 2006). These proteins

show an inverse correlation at the midbody, with CENP-E levels decreasing

as Skp1 associates with this structure. In fact, there is a tight spatiotemporal

regulation of CENP-E at the midbody that is essential for completion of

cytokinesis. In this context, the Skp1 interaction may be essential for

CENP-E proteolysis. Dynein may also play a role in cytokinesis, although

the mechanism is not completely clear. Dynein light intermediate chain 1

(LIC1) is concentrated at the midbody during abscission (Horgan et al.,

2010). Moreover, it was recently shown that dynein is necessary for

transport of Rab8-positive vesicles to the midbody, and this is required

for completion of cytokinesis (Kaplan and Reiner, 2011).

In conclusion, þTIPs can impact mitotic exit at many different levels.

They interact with the major kinases regulating transition from mitosis to

G1 such as CDK1, Plk1, and Aurora-B. Moreover, they are prominently

localized to the spindle midzone and midbody which are crucial in the out-

come of mitosis. Although þTIPs have a significant role in the direct reg-

ulation of MT function, there is a network of reciprocal interactions at the

midzone and midbody which is regulated by Aurora-B or CDK1-mediated

phosphorylations and involves many different families of þTIPs.

6. CONCLUDING REMARKS

In this review, we aimed to provide an up-to-date, systematic orga-

nization of MT functions as well as the role that the major þTIP families

play in this context. More specifically, we addressed the significance that

þTIPs have on the regulation of MT dynamics and stability, which makes

them important players during cell division. It is interesting that many of the

þTIPs share either a functional or spatial overlap, highlighting the complex

role that MT plus ends play throughout mitosis. While many of the features

that define a þTIP are already known, it is still unclear how they interact

with each other in specific contexts to regulate mitotic progression. The

identification of the main structural features that regulate association of pro-

teins to the MT plus ends has led to an overwhelming increase in the num-

ber of potential þTIPs. So far, around 800 proteins were defined as

containing SxIP motifs and many have been confirmed to track on MT plus

ends ( Jiang et al., 2012). Strikingly, little is known about the mitotic role of

most of these proteins, providing a fruitful ground for study in the

coming years.


ACKNOWLEDGMENTSAna L. Pereira is supported by fellowship SFRH/BPD/66707/2009 from Fundacao para a

Ciencia e Tecnologia (FCT, Portugal). Work in the laboratory of H. M. is funded by Grants

PTDC/SAU-ONC/112917/2009 from FCT (COMPETE-FEDER), the Human Frontier

Science Program and the 7th framework program Grant PRECISE from the European

Research Council.

REFERENCESAkhmanova, A., Hoogenraad, C.C., 2005. Microtubule plus-end-tracking proteins: mech-

anisms and functions. Curr. Opin. Cell Biol. 17, 47–54.Akhmanova, A., Hoogenraad, C.C., Drabek, K., Stepanova, T., Dortland, B., Verkerk, T.,

Vermeulen, W., Burgering, B.M., De Zeeuw, C.I., Grosveld, F., Galjart, N., 2001.Clasps are CLIP-115 and -170 associating proteins involved in the regional regulationof microtubule dynamics in motile fibroblasts. Cell 104, 923–935.

Akhmanova, A., Mausset-Bonnefont, A.L., van Cappellen, W., Keijzer, N.,Hoogenraad, C.C., Stepanova, T., Drabek, K., van der Wees, J., Mommaas, M.,Onderwater, J., van der Meulen, H., Tanenbaum, M.E., Medema, R.H.,Hoogerbrugge, J., Vreeburg, J., Uringa, E.J., Grootegoed, J.A., Grosveld, F.,Galjart, N., 2005. The microtubule plus-end-tracking protein CLIP-170 associates withthe spermatid manchette and is essential for spermatogenesis. Genes Dev. 19, 2501–2515.

Akhmanova, A., Steinmetz, M.O., 2008. Tracking the ends: a dynamic protein networkcontrols the fate of microtubule tips. Nat. Rev. Mol. Cell Biol. 9, 309–322.

Al-Bassam, J., Chang, F., 2011. Regulation of microtubule dynamics by TOG-domainproteins XMAP215/Dis1 and CLASP. Trends Cell Biol. 21, 604–614.

Al-Bassam, J., Kim, H., Brouhard, G., van Oijen, A., Harrison, S.C., Chang, F., 2010.CLASP promotes microtubule rescue by recruiting tubulin dimers to the microtubule.Dev. Cell 19, 245–258.

Al-Bassam, J., van Breugel, M., Harrison, S.C., Hyman, A., 2006. Stu2p binds tubulin andundergoes an open-to-closed conformational change. J. Cell Biol. 172, 1009–1022.

Ali, M.Y., Krementsova, E.B., Kennedy, G.G., Mahaffy, R., Pollard, T.D., Trybus, K.M.,Warshaw, D.M., 2007. Myosin Va maneuvers through actin intersections and diffusesalong microtubules. Proc. Natl. Acad. Sci. U.S.A. 104, 4332–4336.

Andreassen, P.R., Margolis, R.L., 1994. Microtubule dependency of p34cdc2 inactivationand mitotic exit in mammalian cells. J. Cell Biol. 127, 789–802.

Andrews, P.D., Ovechkina, Y., Morrice, N., Wagenbach, M., Duncan, K., Wordeman, L.,Swedlow, J.R., 2004. Aurora B regulates MCAK at the mitotic centromere. Dev. Cell 6,253–268.

Antonio, C., Ferby, I., Wilhelm, H., Jones, M., Karsenti, E., Nebreda, A.R., Vernos, I.,2000. Xkid, a chromokinesin required for chromosome alignment on the metaphaseplate. Cell 102, 425–435.

Arnal, I., Heichette, C., Diamantopoulos, G.S., Chretien, D., 2004. CLIP-170/tubulin-curved oligomers coassemble at microtubule ends and promote rescues. Curr. Biol.14, 2086–2095.

Askham, J.M., Moncur, P., Markham, A.F., Morrison, E.E., 2000. Regulation and functionof the interaction between the APC tumour suppressor protein and EB1. Oncogene 19,1950–1958.

Askham, J.M., Vaughan, K.T., Goodson, H.V., Morrison, E.E., 2002. Evidence that aninteraction between EB1 and p150(Glued) is required for the formation andmaintenanceof a radial microtubule array anchored at the centrosome.Mol. Biol. Cell 13, 3627–3645.


http://refhub.elsevier.com/B978-0-12-800255-1.00002-8/rf0005









































Bader, J.R., Kasuboski, J.M., Winding, M., Vaughan, P.S., Hinchcliffe, E.H.,Vaughan, K.T., 2011. Polo-like kinase1 is required for recruitment of dynein to kinet-ochores during mitosis. J. Biol. Chem. 286, 20769–20777.

Bakhoum, S.F., Genovese, G., Compton, D.A., 2009a. Deviant kinetochore microtubuledynamics underlie chromosomal instability. Curr. Biol. 19, 1937–1942.

Bakhoum, S.F., Thompson, S.L., Manning, A.L., Compton, D.A., 2009b. Genome stabilityis ensured by temporal control of kinetochore-microtubule dynamics. Nat. Cell Biol. 11,27–35.

Ban, R., Matsuzaki, H., Akashi, T., Sakashita, G., Taniguchi, H., Park, S.Y., Tanaka, H.,Furukawa, K., Urano, T., 2009. Mitotic regulation of the stability of microtubule plus-end tracking protein EB3 by ubiquitin ligase SIAH-1 and Aurora mitotic kinases. J. Biol.Chem. 284, 28367–28381.

Barisic, M., Sohm, B., Mikolcevic, P., Wandke, C., Rauch, V., Ringer, T., Hess, M.,Bonn, G., Geley, S., 2010. Spindly/CCDC99 is required for efficient chromosome con-gression and mitotic checkpoint regulation. Mol. Biol. Cell 21, 1968–1981.

Barr, A.R., Gergely, F., 2008. MCAK-independent functions of ch-Tog/XMAP215 inmicrotubule plus-end dynamics. Mol. Cell. Biol. 28, 7199–7211.

Beinhauer, J.D., Hagan, I.M., Hegemann, J.H., Fleig, U., 1997. Mal3, the fission yeasthomologue of the human APC-interacting protein EB-1 is required for microtubuleintegrity and the maintenance of cell form. J. Cell Biol. 139, 717–728.

Belmont, L.D., Hyman, A.A., Sawin, K.E., Mitchison, T.J., 1990. Real-time visualization ofcell cycle-dependent changes in microtubule dynamics in cytoplasmic extracts. Cell 62,579–589.

Bement, W.M., Benink, H.A., von Dassow, G., 2005. A microtubule-dependent zone ofactive RhoA during cleavage plane specification. J. Cell Biol. 170, 91–101.

Berrueta, L., Kraeft, S.K., Tirnauer, J.S., Schuyler, S.C., Chen, L.B., Hill, D.E., Pellman, D.,Bierer, B.E., 1998. The adenomatous polyposis coli-binding protein EB1 is associatedwith cytoplasmic and spindle microtubules. Proc. Natl. Acad. Sci. U.S.A. 95,10596–10601.

Bhattacharjee, R.N., Hamada, F., Toyoshima, K., Akiyama, T., 1996. The tumor suppressorgene product APC is hyperphosphorylated during the M phase. Biochem. Biophys. Res.Commun. 220, 192–195.

Bieling, P., Kandels-Lewis, S., Telley, I.A., van Dijk, J., Janke, C., Surrey, T., 2008. CLIP-170 tracks growing microtubule ends by dynamically recognizing compositeEB1/tubulin-binding sites. J. Cell Biol. 183, 1223–1233.

Bieling, P., Laan, L., Schek, H., Munteanu, E.L., Sandblad, L., Dogterom, M., Brunner, D.,Surrey, T., 2007. Reconstitution of a microtubule plus-end tracking system in vitro.Nature 450, 1100–1105.

Bienz, M., 2002. The subcellular destinations of APC proteins. Nat. Rev. Mol. Cell Biol. 3,328–338.

Biggins, S., Walczak, C.E., 2003. Captivating capture: how microtubules attach to kineto-chores. Curr. Biol. 13, R449–R460.

Bjelic, S., De Groot, C.O., Scharer, M.A., Jaussi, R., Bargsten, K., Salzmann, M., Frey, D.,Capitani, G., Kammerer, R.A., Steinmetz, M.O., 2011. Interaction of mammalian endbinding proteins with CAP-Gly domains of CLIP-170 and p150(glued). J. Struct. Biol.177, 160–167.

Bolhy, S., Bouhlel, I., Dultz, E., Nayak, T., Zuccolo, M., Gatti, X., Vallee, R., Ellenberg, J.,Doye, V., 2011. A Nup133-dependent NPC-anchored network tethers centrosomes tothe nuclear envelope in prophase. J. Cell Biol. 192, 855–871.

Booth, D.G., Hood, F.E., Prior, I.A., Royle, S.J., 2011. A TACC3/ch-TOG/clathrincomplex stabilises kinetochore fibres by inter-microtubule bridging. EMBO J. 30,906–919.






















































Brittle, A.L., Ohkura, H., 2005. Mini spindles, the XMAP215 homologue, suppresses paus-ing of interphase microtubules in Drosophila. EMBO J. 24, 1387–1396.

Brodsky, I.B., Burakov, A.V., Nadezhdina, E.S., 2007. Microtubules’ interaction with cellcortex is required for their radial organization, but not for centrosome positioning. CellMotil. Cytoskeleton 64, 407–417.

Brouhard, G.J., Hunt, A.J., 2005. Microtubule movements on the arms of mitotic chromo-somes: polar ejection forces quantified in vitro. Proc. Natl. Acad. Sci. U.S.A. 102,13903–13908.

Brouhard, G.J., Stear, J.H., Noetzel, T.L., Al-Bassam, J., Kinoshita, K., Harrison, S.C.,Howard, J., Hyman, A.A., 2008. XMAP215 is a processive microtubule polymerase.Cell 132, 79–88.

Browning, H., Hackney, D.D., 2005. The EB1 homolog Mal3 stimulates the ATPase of thekinesin Tea2 by recruiting it to the microtubule. J. Biol. Chem. 280, 12299–12304.

Bruning-Richardson, A., Langford, K.J., Ruane, P., Lee, T., Askham, J.M., Morrison, E.E.,2012. EB1 is required for spindle symmetry in mammalian mitosis. PLoSOne 6, e28884.

Brust-Mascher, I., Scholey, J.M., 2002. Microtubule flux and sliding in mitotic spindles ofDrosophila embryos. Mol. Biol. Cell 13, 3967–3975.

Bu, W., Su, L.K., 2001. Regulation of microtubule assembly by human EB1 family proteins.Oncogene 20, 3185–3192.

Bu, W., Su, L.K., 2003. Characterization of functional domains of human EB1 family pro-teins. J. Biol. Chem. 278, 49721–49731.

Buey, R.M., Mohan, R., Leslie, K., Walzthoeni, T., Missimer, J.H., Menzel, A., Bjelic, S.,Bargsten, K., Grigoriev, I., Smal, I., Meijering, E., Aebersold, R., Akhmanova, A.,Steinmetz,M.O., 2011. Insights into EB1 structure and the role of its C-terminal domainfor discriminating microtubule tips from the lattice. Mol. Biol. Cell 22, 2912–2923.

Busch, K.E., Brunner, D., 2004. The microtubule plus end-tracking proteins mal3p andtip1p cooperate for cell-end targeting of interphase microtubules. Curr. Biol. 14,548–559.

Busch, K.E., Hayles, J., Nurse, P., Brunner, D., 2004. Tea2p kinesin is involved inspatial microtubule organization by transporting tip1p on microtubules. Dev. Cell 6,831–843.

Busson, S., Dujardin, D., Moreau, A., Dompierre, J., De Mey, J.R., 1998. Dynein anddynactin are localized to astral microtubules and at cortical sites in mitotic epithelial cells.Curr. Biol. 8, 541–544.

Buster, D.W., Zhang, D., Sharp, D.J., 2007. Poleward tubulin flux in spindles: regulation andfunction in mitotic cells. Mol. Biol. Cell 18, 3094–3104.

Caldwell, C.M., Green, R.A., Kaplan, K.B., 2007. APC mutations lead to cytokinetic fail-ures in vitro and tetraploid genotypes in Min mice. J. Cell Biol. 178, 1109–1120.

Cane, S., Ye, A.A., Luks-Morgan, S.J., Maresca, T.J., 2013. Elevated polar ejection forcesstabilize kinetochore-microtubule attachments. J. Cell Biol. 200, 203–218.

Caplow, M., Ruhlen, R.L., Shanks, J., 1994. The free energy for hydrolysis of amicrotubule-bound nucleotide triphosphate is near zero: all of the free energy for hydro-lysis is stored in the microtubule lattice. J. Cell Biol. 127, 779–788.

Caplow, M., Shanks, J., 1996. Evidence that a single monolayer tubulin-GTP cap is bothnecessary and sufficient to stabilize microtubules. Mol. Biol. Cell 7, 663–675.

Carvalho, P., Gupta Jr., M.L., Hoyt, M.A., Pellman, D., 2004. Cell cycle control of kinesin-mediated transport of Bik1 (CLIP-170) regulates microtubule stability and dyneinactivation. Dev. Cell 6, 815–829.

Carvalho, P., Tirnauer, J.S., Pellman, D., 2003. Surfing on microtubule ends. Trends CellBiol. 13, 229–237.

Cassimeris, L., 1999. Accessory protein regulation of microtubule dynamics throughout thecell cycle. Curr. Opin. Cell Biol. 11, 134–141.






















































Cassimeris, L., Gard, D., Tran, P.T., Erickson, H.P., 2001. XMAP215 is a long thin mol-ecule that does not increase microtubule stiffness. J. Cell Sci. 114, 3025–3033.

Cassimeris, L., Morabito, J., 2004. TOGp, the human homolog of XMAP215/Dis1, isrequired for centrosome integrity, spindle pole organization, and bipolar spindle assem-bly. Mol. Biol. Cell 15, 1580–1590.

Cassimeris, L., Rieder, C.L., Rupp, G., Salmon, E.D., 1990. Stability of microtubule attach-ment to metaphase kinetochores in PtK1 cells. J. Cell Sci. 96 (Pt. 1), 9–15.

Cassimeris, L., Salmon, E.D., 1991. Kinetochore microtubules shorten by loss of subunits atthe kinetochores of prometaphase chromosomes. J. Cell Sci. 98 (Pt. 2), 151–158.

Chan, Y.W., Fava, L.L., Uldschmid, A., Schmitz, M.H., Gerlich, D.W., Nigg, E.A.,Santamaria, A., 2009. Mitotic control of kinetochore-associated dynein and spindleorientation by human Spindly. J. Cell Biol. 185, 859–874.

Charrasse, S., Schroeder, M., Gauthier-Rouviere, C., Ango, F., Cassimeris, L., Gard, D.L.,Larroque, C., 1998. The TOGp protein is a new human microtubule-associated proteinhomologous to the Xenopus XMAP215. J. Cell Sci. 111 (Pt. 10), 1371–1383.

Cheeseman, I.M., MacLeod, I., Yates 3rd., J.R., Oegema, K., Desai, A., 2005. The CENP-F-like proteins HCP-1 and HCP-2 target CLASP to kinetochores to mediate chromo-some segregation. Curr. Biol. 15, 771–777.

Cheeseman, L.P., Booth, D.G., Hood, F.E., Prior, I.A., Royle, S.J., 2011. Aurora A kinaseactivity is required for localization of TACC3/ch-TOG/clathrin inter-microtubulebridges. Commun. Integr. Biol. 4, 409–412.

Cheng, T.S., Hsiao, Y.L., Lin, C.C., Hsu, C.M., Chang, M.S., Lee, C.I., Yu, R.C.,Huang, C.Y., Howng, S.L., Hong, Y.R., 2007. hNinein is required for targetingspindle-associated protein Astrin to the centrosome during the S and G2 phases. Exp.Cell Res. 313, 1710–1721.

Cheng, T.S., Hsiao, Y.L., Lin, C.C., Yu, C.T., Hsu, C.M., Chang, M.S., Lee, C.I.,Huang, C.Y., Howng, S.L., Hong, Y.R., 2008. Glycogen synthase kinase 3beta interactswith and phosphorylates the spindle-associated protein astrin. J. Biol. Chem. 283,2454–2464.

Choi, J.H., Bertram, P.G., Drenan, R., Carvalho, J., Zhou, H.H., Zheng, X.F., 2002. TheFKBP12-rapamycin-associated protein (FRAP) is a CLIP-170 kinase. EMBO Rep. 3,988–994.

Cimini, D., Moree, B., Canman, J.C., Salmon, E.D., 2003. Merotelic kinetochore orienta-tion occurs frequently during early mitosis in mammalian tissue cells and error correctionis achieved by two different mechanisms. J. Cell Sci. 116, 4213–4225.

Cimini, D., Wan, X., Hirel, C.B., Salmon, E.D., 2006. Aurora kinase promotes turnover ofkinetochore microtubules to reduce chromosome segregation errors. Curr. Biol. 16,1711–1718.

Cooke, C.A., Schaar, B., Yen, T.J., Earnshaw, W.C., 1997. Localization of CENP-E in thefibrous corona and outer plate of mammalian kinetochores from prometaphase throughanaphase. Chromosoma 106, 446–455.

Cooper, J.R., Wordeman, L., 2009. The diffusive interaction of microtubule bindingproteins. Curr. Opin. Cell Biol. 21, 68–73.

Coquelle, F.M., Caspi, M., Cordelieres, F.P., Dompierre, J.P., Dujardin, D.L., Koifman, C.,Martin, P., Hoogenraad, C.C., Akhmanova, A., Galjart, N., De Mey, J.R., Reiner, O.,2002. LIS1, CLIP-170’s key to the dynein/dynactin pathway. Mol. Cell. Biol. 22,3089–3102.

Coquelle, F.M., Vitre, B., Arnal, I., 2009. Structural basis of EB1 effects on microtubuledynamics. Biochem. Soc. Trans. 37, 997–1001.

Coue, M., Lombillo, V.A., McIntosh, J.R., 1991. Microtubule depolymerization promotesparticle and chromosome movement in vitro. J. Cell Biol. 112, 1165–1175.





















































Cullen, C.F., Deak, P., Glover, D.M., Ohkura, H., 1999. mini spindles: A gene encodinga conserved microtubule-associated protein required for the integrity of the mitoticspindle in Drosophila. J. Cell Biol. 146, 1005–1018.

Dawe, A.L., Caldwell, K.A., Harris, P.M., Morris, N.R., Caldwell, G.A., 2001. Evolution-arily conserved nuclear migration genes required for early embryonic development inCaenorhabditis elegans. Dev. Genes Evol. 211, 434–441.

De Groot, C.O., Jelesarov, I., Damberger, F.F., Bjelic, S., Scharer, M.A., Bhavesh, N.S.,Grigoriev, I., Buey, R.M., Wuthrich, K., Capitani, G., Akhmanova, A.,Steinmetz, M.O., 2009. Molecular insights into mammalian end-binding proteinheterodimerization. J. Biol. Chem. 285, 5802–5814.

De Luca, M., Brunetto, L., Asteriti, I.A., Giubettini, M., Lavia, P., Guarguaglini, G., 2008.Aurora-A and ch-TOG act in a common pathway in control of spindle pole integrity.Oncogene 27, 6539–6549.

De Zeeuw, C.I., Hoogenraad, C.C., Goedknegt, E., Hertzberg, E., Neubauer, A.,Grosveld, F., Galjart, N., 1997. CLIP-115, a novel brain-specific cytoplasmic linkerprotein, mediates the localization of dendritic lamellar bodies. Neuron 19, 1187–1199.

Dell, K.R., Turck, C.W., Vale, R.D., 2000. Mitotic phosphorylation of the dynein lightintermediate chain is mediated by cdc2 kinase. Traffic 1, 38–44.

des Georges, A., Katsuki, M., Drummond, D.R., Osei, M., Cross, R.A., Amos, L.A., 2008.Mal3, the Schizosaccharomyces pombe homolog of EB1, changes the microtubulelattice. Nat. Struct. Mol. Biol. 15, 1102–1108.

Desai, A., Mitchison, T.J., 1997. Microtubule polymerization dynamics. Annu. Rev. CellDev. Biol. 13, 83–117.

Desai, A., Verma, S., Mitchison, T.J., Walczak, C.E., 1999. Kin I kinesins are microtubule-destabilizing enzymes. Cell 96, 69–78.

Diamantopoulos, G.S., Perez, F., Goodson, H.V., Batelier, G., Melki, R., Kreis, T.E.,Rickard, J.E., 1999. Dynamic localization of CLIP-170 to microtubule plus ends iscoupled to microtubule assembly. J. Cell Biol. 144, 99–112.

Dikovskaya, D., Newton, I.P., Nathke, I.S., 2004. The adenomatous polyposis coli protein isrequired for the formation of robust spindles formed in CSFXenopus extracts. Mol. Biol.Cell 15, 2978–2991.

Dikovskaya, D., Schiffmann, D., Newton, I.P., Oakley, A., Kroboth, K., Sansom, O.,Jamieson, T.J., Meniel, V., Clarke, A., Nathke, I.S., 2007. Loss of APC induces poly-ploidy as a result of a combination of defects in mitosis and apoptosis. J. Cell Biol. 176,183–195.

Dionne, M.A., Sanchez, A., Compton, D.A., 2000. ch-TOGp is required for micro-tubule aster formation in a mammalian mitotic extract. J. Biol. Chem. 275,12346–12352.

Dixit, R., Barnett, B., Lazarus, J.E., Tokito, M., Goldman, Y.E., Holzbaur, E.L., 2009.Microtubule plus-end tracking by CLIP-170 requires EB1. Proc. Natl. Acad. Sci.U.S.A. 106, 492–497.

Dobyns, W.B., Reiner, O., Carrozzo, R., Ledbetter, D.H., 1993. Lissencephaly. A humanbrain malformation associated with deletion of the LIS1 gene located at chromosome17p13. JAMA 270, 2838–2842.

Dogterom, M., Kerssemakers, J.W., Romet-Lemonne, G., Janson, M.E., 2005. Force gen-eration by dynamic microtubules. Curr. Opin. Cell Biol. 17, 67–74.

Dogterom, M., Yurke, B., 1997. Measurement of the force-velocity relation for growingmicrotubules. Science 278, 856–860.

Domnitz, S.B., Wagenbach, M., Decarreau, J., Wordeman, L., 2012. MCAK activity atmicrotubule tips regulates spindle microtubule length to promote robust kinetochoreattachment. J. Cell Biol. 197, 231–237.





















































Drabek, K., Gutierrez, L., Vermeij, M., Clapes, T., Patel, S.R., Boisset, J.C., van Haren, J.,Pereira, A.L., Liu, Z., Akinci, U., Nikolic, T., van Ijcken, W., van den Hout, M.,Meinders, M., Melo, C., Sambade, C., Drabek, D., Hendriks, R.W., Philipsen, S.,Mommaas, M., Grosveld, F., Maiato, H., Italiano Jr., J.E., Robin, C., Galjart, N.,2012. The microtubule plus-end tracking protein CLASP2 is required for hematopoiesisand hematopoietic stem cell maintenance. Cell Reports 2, 781–788.

Drabek, K., van Ham, M., Stepanova, T., Draegestein, K., van Horssen, R., Sayas, C.L.,Akhmanova, A., Ten Hagen, T., Smits, R., Fodde, R., Grosveld, F., Galjart, N.,2006. Role of CLASP2 in microtubule stabilization and the regulation of persistentmotility. Curr. Biol. 16, 2259–2264.

Dragestein, K.A., van Cappellen, W.A., van Haren, J., Tsibidis, G.D., Akhmanova, A.,Knoch, T.A., Grosveld, F., Galjart, N., 2008. Dynamic behavior of GFP-CLIP-170reveals fast protein turnover on microtubule plus ends. J. Cell Biol. 180, 729–737.

Draviam, V.M., Shapiro, I., Aldridge, B., Sorger, P.K., 2006. Misorientation and reducedstretching of aligned sister kinetochores promote chromosome missegregation inEB1- or APC-depleted cells. EMBO J. 25, 2814–2827.

Drechsel, D.N., Kirschner, M.W., 1994. The minimum GTP cap required to stabilizemicrotubules. Curr. Biol. 4, 1053–1061.

Dujardin, D., Wacker, U.I., Moreau, A., Schroer, T.A., Rickard, J.E., De Mey, J.R., 1998.Evidence for a role of CLIP-170 in the establishment of metaphase chromosome align-ment. J. Cell Biol. 141, 849–862.

Dujardin, D.L., Barnhart, L.E., Stehman, S.A., Gomes, E.R., Gundersen, G.G.,Vallee, R.B., 2003. A role for cytoplasmic dynein and LIS1 in directed cell movement.J. Cell Biol. 163, 1205–1211.

Dunsch, A.K., Linnane, E., Barr, F.A., Gruneberg, U., 2011. The astrin-kinastrin/SKAPcomplex localizes to microtubule plus ends and facilitates chromosome alignment.J. Cell Biol. 192, 959–968.

Dzhindzhev, N.S., Rogers, S.L., Vale, R.D., Ohkura, H., 2005. Distinct mechanisms governthe localisation of Drosophila CLIP-190 to unattached kinetochores and microtubuleplus-ends. J. Cell Sci. 118, 3781–3790.

Echeverri, C.J., Paschal, B.M., Vaughan, K.T., Vallee, R.B., 1996. Molecular characteriza-tion of the 50-kD subunit of dynactin reveals function for the complex in chromosomealignment and spindle organization during mitosis. J. Cell Biol. 132, 617–633.

Efimov, A., Kharitonov, A., Efimova, N., Loncarek, J., Miller, P.M., Andreyeva, N.,Gleeson, P., Galjart, N., Maia, A.R., McLeod, I.X., Yates 3rd., J.R., Maiato, H.,Khodjakov, A., Akhmanova, A., Kaverina, I., 2007. Asymmetric CLASP-dependentnucleation of noncentrosomal microtubules at the trans-Golgi network. Dev. Cell 12,917–930.

Elad, N., Abramovitch, S., Sabanay, H., Medalia, O., 2010. Microtubule organization in thefinal stages of cytokinesis as revealed by cryo-electron tomography. J. Cell Sci. 124,207–215.

Emes, R.D., Ponting, C.P., 2001. A new sequence motif linking lissencephaly, TreacherCollins and oral-facial-digital type 1 syndromes, microtubule dynamics and cell migra-tion. Hum. Mol. Genet. 10, 2813–2820.

Ems-McClung, S.C., Hertzer, K.M., Zhang, X., Miller, M.W., Walczak, C.E., 2007. Theinterplay of the N- and C-terminal domains of MCAK control microtubule depolymer-ization activity and spindle assembly. Mol. Biol. Cell 18, 282–294.

Ems-McClung, S.C., Walczak, C.E., 2010. Kinesin-13s in mitosis: Key players in thespatial and temporal organization of spindle microtubules. Semin. Cell Dev. Biol. 21,276–282.

Espiritu, E.B., Krueger, L.E., Ye, A., Rose, L.S., 2012. CLASPs function redundantly toregulate astral microtubules in the C. elegans embryo. Dev. Biol. 368, 242–254.






















































Etienne-Manneville, S., Hall, A., 2003. Cdc42 regulates GSK-3beta and adenomatous poly-posis coli to control cell polarity. Nature 421, 753–756.

Euteneuer, U., McIntosh, J.R., 1980. Polarity of midbody and phragmoplast microtubules.J. Cell Biol. 87, 509–515.

Faulkner, N.E., Dujardin, D.L., Tai, C.Y., Vaughan, K.T., O’Connell, C.B., Wang, Y.,Vallee, R.B., 2000. A role for the lissencephaly gene LIS1 in mitosis and cytoplasmicdynein function. Nat. Cell Biol. 2, 784–791.

Ferreira, J.G., Pereira, A.J., Akhmanova, A., Maiato, H., 2013. Aurora B spatially regulatesEB3 phosphorylation to coordinate daughter cell adhesion with cytokinesis. J. Cell Biol.201, 709–724.

Fodde, R., Kuipers, J., Rosenberg, C., Smits, R., Kielman, M., Gaspar, C., van Es, J.H.,Breukel, C., Wiegant, J., Giles, R.H., Clevers, H., 2001. Mutations in the APC tumoursuppressor gene cause chromosomal instability. Nat. Cell Biol. 3, 433–438.

Foe, V.E., von Dassow, G., 2008. Stable and dynamic microtubules coordinately shape themyosin activation zone during cytokinetic furrow formation. J. Cell Biol. 183, 457–470.

Folker, E.S., Baker, B.M., Goodson, H.V., 2005. Interactions between CLIP-170, tubulin,and microtubules: implications for the mechanism of Clip-170 plus-end tracking behav-ior. Mol. Biol. Cell 16, 5373–5384.

Fong, K.W., Hau, S.Y., Kho, Y.S., Jia, Y., He, L., Qi, R.Z., 2009. Interaction ofCDK5RAP2 with EB1 to track growing microtubule tips and to regulate microtubuledynamics. Mol. Biol. Cell 20, 3660–3670.

Fujiwara, K., Pollard, T.D., 1976. Fluorescent antibody localization of myosin in the cyto-plasm, cleavage furrow, and mitotic spindle of human cells. J. Cell Biol. 71, 848–875.

Fuller, B.G., Lampson, M.A., Foley, E.A., Rosasco-Nitcher, S., Le, K.V., Tobelmann, P.,Brautigan, D.L., Stukenberg, P.T., Kapoor, T.M., 2008. Midzone activation of auroraB in anaphase produces an intracellular phosphorylation gradient. Nature 453,1132–1136.

Furuta, K., Toyoshima, Y.Y., 2008. Minus-end-directed motor Ncd exhibits processivemovement that is enhanced by microtubule bundling in vitro. Curr. Biol. 18, 152–157.

Gache, V., Louwagie, M., Garin, J., Caudron, N., Lafanechere, L., Valiron, O., 2005.Identification of proteins binding the native tubulin dimer. Biochem. Biophys. Res.Commun. 327, 35–42.

Galjart, N., 2005. CLIPs and CLASPs and cellular dynamics. Nat. Rev. Mol. Cell Biol. 6,487–498.

Galjart, N., Perez, F., 2003. A plus-end raft to control microtubule dynamics and function.Curr. Opin. Cell Biol. 15, 48–53.

Ganem, N.J., Compton, D.A., 2004. The KinI kinesin Kif2a is required for bipolar spindleassembly through a functional relationship with MCAK. J. Cell Biol. 166, 473–478.

Ganem, N.J., Upton, K., Compton, D.A., 2005. Efficient mitosis in human cells lackingpoleward microtubule flux. Curr. Biol. 15, 1827–1832.

Gard, D.L., Kirschner, M.W., 1987. A microtubule-associated protein from Xenopus eggsthat specifically promotes assembly at the plus-end. J. Cell Biol. 105, 2203–2215.

Gardner, M.K., Zanic, M., Gell, C., Bormuth, V., Howard, J., 2012. Depolymerizingkinesins Kip3 and MCAK shape cellular microtubule architecture by differential controlof catastrophe. Cell 147, 1092–1103.

Gassmann, R., Holland, A.J., Varma, D.,Wan, X., Civril, F., Cleveland, D.W., Oegema, K.,Salmon, E.D., Desai, A., 2010. Removal of Spindly from microtubule-attached kinet-ochores controls spindle checkpoint silencing in human cells. Genes Dev. 24, 957–971.

Geiser, J.R., Schott, E.J., Kingsbury, T.J., Cole, N.B., Totis, L.J., Bhattacharyya, G., He, L.,Hoyt, M.A., 1997. Saccharomyces cerevisiae genes required in the absence of the CIN8-encoded spindle motor act in functionally diverse mitotic pathways. Mol. Biol. Cell 8,1035–1050.






















































Gergely, F., Draviam, V.M., Raff, J.W., 2003. The ch-TOG/XMAP215 protein is essentialfor spindle pole organization in human somatic cells. Genes Dev. 17, 336–341.

Gestaut, D.R., Graczyk, B., Cooper, J., Widlund, P.O., Zelter, A., Wordeman, L.,Asbury, C.L., Davis, T.N., 2008. Phosphoregulation and depolymerization-drivenmovement of the Dam1 complex do not require ring formation. Nat. Cell Biol. 10,407–414.

Gorbsky, G.J., Borisy, G.G., 1989. Microtubules of the kinetochore fiber turn over in meta-phase but not in anaphase. J. Cell Biol. 109, 653–662.

Gorbsky, G.J., Sammak, P.J., Borisy, G.G., 1987. Chromosomes move poleward in anaphasealong stationary microtubules that coordinately disassemble from their kinetochore ends.J. Cell Biol. 104, 9–18.

Gorbsky, G.J., Sammak, P.J., Borisy, G.G., 1988. Microtubule dynamics and chromosomemotion visualized in living anaphase cells. J. Cell Biol. 106, 1185–1192.

Gorbsky, G.J., Simerly, C., Schatten, G., Borisy, G.G., 1990. Microtubules in themetaphase-arrested mouse oocyte turn over rapidly. Proc. Natl. Acad. Sci. U.S.A. 87,6049–6053.

Goshima, G., Nedelec, F., Vale, R.D., 2005. Mechanisms for focusing mitotic spindle polesby minus end-directed motor proteins. J. Cell Biol. 171, 229–240.

Green, R.A., Kaplan, K.B., 2003. Chromosome instability in colorectal tumor cells is asso-ciated with defects in microtubule plus-end attachments caused by a dominant mutationin APC. J. Cell Biol. 163, 949–961.

Green, R.A., Wollman, R., Kaplan, K.B., 2005. APC and EB1 function together in mitosisto regulate spindle dynamics and chromosome alignment. Mol. Biol. Cell 16,4609–4622.

Grego, S., Cantillana, V., Salmon, E.D., 2001. Microtubule treadmilling in vitro investigatedby fluorescence speckle and confocal microscopy. Biophys. J. 81, 66–78.

Grigoriev, I., Gouveia, S.M., van der Vaart, B., Demmers, J., Smyth, J.T., Honnappa, S.,Splinter, D., Steinmetz, M.O., Putney Jr., J.W., Hoogenraad, C.C., Akhmanova, A.,2008. STIM1 is a MT-plus-end-tracking protein involved in remodeling of the ER.Curr. Biol. 18, 177–182.

Grishchuk, E.L., Molodtsov,M.I., Ataullakhanov, F.I., McIntosh, J.R., 2005. Force produc-tion by disassembling microtubules. Nature 438, 384–388.

Grohmann, A., Tanneberger, K., Alzner, A., Schneikert, J., Behrens, J., 2007. AMER1regulates the distribution of the tumor suppressor APC between microtubules and theplasma membrane. J. Cell Sci. 120, 3738–3747.

Gruber, J., Harborth, J., Schnabel, J., Weber, K., Hatzfeld, M., 2002. The mitotic-spindle-associated protein astrin is essential for progression through mitosis. J. Cell Sci. 115,4053–4059.

Guizetti, J., Schermelleh, L., Mantler, J., Maar, S., Poser, I., Leonhardt, H.,Muller-Reichert,T., Gerlich, D.W., 2011. Cortical constriction during abscission involves helices ofESCRT-III-dependent filaments. Science 331, 1616–1620.

Gupta, K.K., Paulson, B.A., Folker, E.S., Charlebois, B., Hunt, A.J., Goodson, H.V., 2009.Minimal plus-end tracking unit of the cytoplasmic linker protein CLIP-170. J. Biol.Chem. 284, 6735–6742.

Hannak, E., Heald, R., 2006. Xorbit/CLASP links dynamic microtubules to chromosomesin the Xenopus meiotic spindle. J. Cell Biol. 172, 19–25.

Haraguchi, K., Hayashi, T., Jimbo, T., Yamamoto, T., Akiyama, T., 2006. Role of thekinesin-2 family protein, KIF3, during mitosis. J. Biol. Chem. 281, 4094–4099.

Hayashi, I., Ikura, M., 2003. Crystal structure of the amino-terminal microtubule-bindingdomain of end-binding protein 1 (EB1). J. Biol. Chem. 278, 36430–36434.

Hayashi, I., Plevin, M.J., Ikura, M., 2007. CLIP170 autoinhibition mimics intermolecularinteractions with p150Glued or EB1. Nat. Struct. Mol. Biol. 14, 980–981.






















































Hayden, J.H., Bowser, S.S., Rieder, C.L., 1990. Kinetochores capture astral microtubulesduring chromosome attachment to the mitotic spindle: direct visualization in live newtlung cells. J. Cell Biol. 111, 1039–1045.

Helenius, J., Brouhard, G., Kalaidzidis, Y., Diez, S., Howard, J., 2006. The depolymerizingkinesin MCAK uses lattice diffusion to rapidly target microtubule ends. Nature 441,115–119.

Hendricks, A.G., Lazarus, J.E., Perlson, E., Gardner, M.K., Odde, D.J., Goldman, Y.E.,Holzbaur, E.L., 2012. Dynein tethers and stabilizes dynamic microtubule plus ends.Curr. Biol. 22, 632–637.

Hirokawa, N., Noda, Y., Tanaka, Y., Niwa, S., 2009. Kinesin superfamily motor proteinsand intracellular transport. Nat. Rev. Mol. Cell Biol. 10, 682–696.

Holmfeldt, P., Stenmark, S., Gullberg, M., 2004. Differential functional interplay of TOGp/XMAP215 and the KinI kinesin MCAK during interphase and mitosis. EMBO J. 23,627–637.

Holy, T.E., Dogterom, M., Yurke, B., Leibler, S., 1997. Assembly and positioning of micro-tubule asters in microfabricated chambers. Proc. Natl. Acad. Sci. U.S.A. 94, 6228–6231.

Honnappa, S., Gouveia, S.M., Weisbrich, A., Damberger, F.F., Bhavesh, N.S., Jawhari, H.,Grigoriev, I., van Rijssel, F.J., Buey, R.M., Lawera, A., Jelesarov, I., Winkler, F.K.,Wuthrich, K., Akhmanova, A., Steinmetz, M.O., 2009. An EB1-binding motif actsas a microtubule tip localization signal. Cell 138, 366–376.

Honnappa, S., John, C.M., Kostrewa, D., Winkler, F.K., Steinmetz, M.O., 2005. Structuralinsights into the EB1-APC interaction. EMBO J. 24, 261–269.

Honnappa, S., Okhrimenko, O., Jaussi, R., Jawhari, H., Jelesarov, I., Winkler, F.K.,Steinmetz, M.O., 2006. Key interaction modes of dynamic þTIP networks. Mol. Cell23, 663–671.

Hoogenraad, C.C., Akhmanova, A., Grosveld, F., De Zeeuw, C.I., Galjart, N., 2000. Func-tional analysis of CLIP-115 and its binding to microtubules. J. Cell Sci. 113 (Pt. 12),2285–2297.

Horgan, C.P., Hanscom, S.R., McCaffrey, M.W., 2010. Dynein LIC1 localizes to themitotic spindle and midbody and LIC2 localizes to spindle poles during cell division. CellBiol. Int. 35, 171–178.

Houlden, H., Johnson, J., Gardner-Thorpe, C., Lashley, T., Hernandez, D., Worth, P.,Singleton, A.B., Hilton, D.A., Holton, J., Revesz, T., Davis, M.B., Giunti, P.,Wood, N.W., 2007. Mutations in TTBK2, encoding a kinase implicated in tau phos-phorylation, segregate with spinocerebellar ataxia type 11. Nat. Genet. 39, 1434–1436.

Howard, J., Hyman, A.A., 2003. Dynamics and mechanics of the microtubule plus end.Nature 422, 753–758.

Hu, C.K., Coughlin, M., Mitchison, T.J., 2012. Midbody assembly and its regulation duringcytokinesis. Mol. Biol. Cell 23, 1024–1034.

Hunter, A.W., Caplow, M., Coy, D.L., Hancock, W.O., Diez, S., Wordeman, L.,Howard, J., 2003. The kinesin-related protein MCAK is a microtubule depolymerasethat forms an ATP-hydrolyzing complex at microtubule ends. Mol. Cell 11, 445–457.

Hyman, A.A., Salser, S., Drechsel, D.N., Unwin, N., Mitchison, T.J., 1992. Role of GTPhydrolysis in microtubule dynamics: information from a slowly hydrolyzable analogue,GMPCPP. Mol. Biol. Cell 3, 1155–1167.

Iimori, M., Ozaki, K., Chikashige, Y., Habu, T., Hiraoka, Y., Maki, T., Hayashi, I.,Obuse, C.,Matsumoto, T., 2012. Amutation of the fission yeast EB1 overcomes negativeregulation by phosphorylation and stabilizes microtubules. Exp. Cell Res. 318, 262–275.

Inoue, S., Salmon, E.D., 1995. Force generation by microtubule assembly/disassembly inmitosis and related movements. Mol. Biol. Cell 6, 1619–1640.

Inoue, Y., Iwane, A.H., Miyai, T., Muto, E., Yanagida, T., 2001. Motility of single one-headed kinesin molecules along microtubules. Biophys. J. 81, 2838–2850.























































Inoue, Y.H., do Carmo Avides, M., Shiraki, M., Deak, P., Yamaguchi, M., Nishimoto, Y.,Matsukage, A., Glover, D.M., 2000. Orbit, a novel microtubule-associated proteinessential for mitosis in Drosophila melanogaster. J. Cell Biol. 149, 153–166.

Inoue, Y.H., Savoian, M.S., Suzuki, T., Mathe, E., Yamamoto, M.T., Glover, D.M., 2004.Mutations in orbit/mast reveal that the central spindle is comprised of two microtubulepopulations, those that initiate cleavage and those that propagate furrow ingression.J. Cell Biol. 166, 49–60.

Jiang, K., Toedt, G., Montenegro Gouveia, S., Davey, N.E., Hua, S., van der Vaart, B.,Grigoriev, I., Larsen, J., Pedersen, L.B., Bezstarosti, K., Lince-Faria, M., Demmers, J.,Steinmetz,M.O.,Gibson,T.J.,Akhmanova,A., 2012.Aproteome-wide screen formam-malian SxIP motif-containing microtubule plus-end tracking proteins. Curr. Biol. 22,1800–1807.

Jiang, K.,Wang, J., Liu, J.,Ward, T.,Wordeman, L., Davidson, A.,Wang, F., Yao, X., 2009.TIP150 interacts with and targets MCAK at the microtubule plus ends. EMBORep. 10,857–865.

Jiang, W., Jimenez, G., Wells, N.J., Hope, T.J., Wahl, G.M., Hunter, T., Fukunaga, R.,1998. PRC1: a human mitotic spindle-associated CDK substrate protein required forcytokinesis. Mol. Cell 2, 877–885.

Jimbo, T., Kawasaki, Y., Koyama, R., Sato, R., Takada, S., Haraguchi, K., Akiyama, T.,2002. Identification of a link between the tumour suppressor APC and the kinesin super-family. Nat. Cell Biol. 4, 323–327.

Julian, M., Tollon, Y., Lajoie-Mazenc, I., Moisand, A., Mazarguil, H., Puget, A.,Wright, M., 1993. Gamma-tubulin participates in the formation of the midbody duringcytokinesis in mammalian cells. J. Cell Sci. 105 (Pt. 1), 145–156.

Kapitein, L.C., Peterman, E.J., Kwok, B.H., Kim, J.H., Kapoor, T.M., Schmidt, C.F., 2005.The bipolar mitotic kinesin Eg5 moves on both microtubules that it crosslinks. Nature435, 114–118.

Kaplan, A., Reiner, O., 2011. Linking cytoplasmic dynein and transport of Rab8 vesicles tothe midbody during cytokinesis by the doublecortin domain-containing 5 protein.J. Cell Sci. 124, 3989–4000.

Kaplan, K.B., Burds, A.A., Swedlow, J.R., Bekir, S.S., Sorger, P.K., Nathke, I.S., 2001.A role for the Adenomatous Polyposis Coli protein in chromosome segregation. Nat.Cell Biol. 3, 429–432.

Kapoor, T.M., Lampson, M.A., Hergert, P., Cameron, L., Cimini, D., Salmon, E.D.,McEwen, B.F., Khodjakov, A., 2006. Chromosomes can congress to themetaphase platebefore biorientation. Science 311, 388–391.

Ke, K., Cheng, J., Hunt, A.J., 2009. The distribution of polar ejection forces determines theamplitude of chromosome directional instability. Curr. Biol. 19, 807–815.

Kim, M.H., Cooper, D.R., Oleksy, A., Devedjiev, Y., Derewenda, U., Reiner, O.,Otlewski, J.,Derewenda,Z.S., 2004.The structure of theN-terminal domainof theprod-uct of the lissencephaly gene Lis1 and its functional implications. Structure 12, 987–998.

Kim, Y., Heuser, J.E.,Waterman, C.M., Cleveland, D.W., 2008. CENP-E combines a slow,processive motor and a flexible coiled coil to produce an essential motile kinetochoretether. J. Cell Biol. 181, 411–419.

King, J.M., Hays, T.S., Nicklas, R.B., 2000. Dynein is a transient kinetochore componentwhose binding is regulated by microtubule attachment, not tension. J. Cell Biol. 151,739–748.

King, S.J., Brown, C.L., Maier, K.C., Quintyne, N.J., Schroer, T.A., 2003. Analysis of thedynein-dynactin interaction in vitro and in vivo. Mol. Biol. Cell 14, 5089–5097.

Kinoshita, K., Arnal, I., Desai, A., Drechsel, D.N., Hyman, A.A., 2001. Reconstitutionof physiological microtubule dynamics using purified components. Science 294,1340–1343.






















































Kirschner, M., Mitchison, T., 1986. Beyond self-assembly: from microtubules to morpho-genesis. Cell 45, 329–342.

Kita, K., Wittmann, T., Nathke, I.S., Waterman-Storer, C.M., 2006. Adenomatous poly-posis coli on microtubule plus ends in cell extensions can promote microtubule netgrowth with or without EB1. Mol. Biol. Cell 17, 2331–2345.

Kiyomitsu, T., Cheeseman, I.M., 2012. Chromosome- and spindle-pole-derived signals gen-erate an intrinsic code for spindle position and orientation. Nat. Cell Biol. 14, 311–317.

Kline-Smith, S.L., Khodjakov, A., Hergert, P., Walczak, C.E., 2004. Depletion of centro-mericMCAK leads to chromosome congression and segregation defects due to improperkinetochore attachments. Mol. Biol. Cell 15, 1146–1159.

Kline-Smith, S.L., Walczak, C.E., 2004. Mitotic spindle assembly and chromosome segre-gation: refocusing on microtubule dynamics. Mol. Cell 15, 317–327.

Kobayashi, T., Murayama, T., 2009. Cell cycle-dependent microtubule-based dynamictransport of cytoplasmic dynein in mammalian cells. PLoS One 4, e7827.

Kodama, A., Karakesisoglou, I., Wong, E., Vaezi, A., Fuchs, E., 2003. ACF7: an essentialintegrator of microtubule dynamics. Cell 115, 343–354.

Komarova, Y., De Groot, C.O., Grigoriev, I., Gouveia, S.M., Munteanu, E.L.,Schober, J.M., Honnappa, S., Buey, R.M., Hoogenraad, C.C., Dogterom, M.,Borisy, G.G., Steinmetz, M.O., Akhmanova, A., 2009. Mammalian end bindingproteins control persistent microtubule growth. J. Cell Biol. 184, 691–706.

Komarova, Y., Lansbergen, G., Galjart, N., Grosveld, F., Borisy, G.G., Akhmanova, A.,2005. EB1 and EB3 control CLIP dissociation from the ends of growing microtubules.Mol. Biol. Cell 16, 5334–5345.

Komarova, Y.A., Akhmanova, A.S., Kojima, S., Galjart, N., Borisy, G.G., 2002. Cytoplas-mic linker proteins promote microtubule rescue in vivo. J. Cell Biol. 159, 589–599.

Komarova, Y.A., Huang, F., Geyer, M., Daneshjou, N., Garcia, A., Idalino, L., Kreutz, B.,Mehta, D., Malik, A.B., 2012. VE-cadherin signaling induces EB3 phosphorylation tosuppress microtubule growth and assemble adherens junctions. Mol. Cell 48, 914–925.

Koshland, D.E., Mitchison, T.J., Kirschner, M.W., 1988. Polewards chromosome move-ment driven by microtubule depolymerization in vitro. Nature 331, 499–504.

Kronja, I., Kruljac-Letunic, A., Caudron-Herger, M., Bieling, P., Karsenti, E., 2009.XMAP215-EB1 interaction is required for proper spindle assembly and chromosomesegregation in Xenopus egg extract. Mol. Biol. Cell 20, 2684–2696.

Kumar, P., Chimenti, M.S., Pemble, H., Schonichen, A., Thompson, O., Jacobson, M.P.,Wittmann, T., 2012. Multisite phosphorylation disrupts arginine-glutamate salt bridgenetworks required for binding of the cytoplasmic linker-associated protein 2(CLASP2) to end-binding protein 1 (EB1). J. Biol. Chem. 287, 17050–17064.

Kumar, P., Lyle, K.S., Gierke, S., Matov, A., Danuser, G., Wittmann, T., 2009. GSK3betaphosphorylation modulates CLASP-microtubule association and lamella microtubuleattachment. J. Cell Biol. 184, 895–908.

Kwok, B.H., Kapitein, L.C., Kim, J.H., Peterman, E.J., Schmidt, C.F., Kapoor, T.M., 2006.Allosteric inhibition of kinesin-5 modulates its processive directional motility. Nat.Chem. Biol. 2, 480–485.

Kwok, B.H., Kapoor, T.M., 2007. Microtubule flux: drivers wanted. Curr. Opin. Cell Biol.19, 36–42.

Laan, L., Pavin, N., Husson, J., Romet-Lemonne, G., van Duijn, M., Lopez, M.P.,Vale, R.D., Julicher, F., Reck-Peterson, S.L., Dogterom, M., 2012. Cortical dyneincontrols microtubule dynamics to generate pulling forces that position microtubuleasters. Cell 148, 502–514.

Lamb, N.J., Fernandez, A., Watrin, A., Labbe, J.C., Cavadore, J.C., 1990. Microinjection ofp34cdc2 kinase induces marked changes in cell shape, cytoskeletal organization, andchromatin structure in mammalian fibroblasts. Cell 60, 151–165.






















































Lampson,M.A.,Renduchitala,K.,Khodjakov,A.,Kapoor,T.M., 2004.Correcting improperchromosome-spindle attachments during cell division. Nat. Cell Biol. 6, 232–237.

Lan, W., Zhang, X., Kline-Smith, S.L., Rosasco, S.E., Barrett-Wilt, G.A., Shabanowitz, J.,Hunt, D.F., Walczak, C.E., Stukenberg, P.T., 2004. Aurora B phosphorylates centro-meric MCAK and regulates its localization and microtubule depolymerization activity.Curr. Biol. 14, 273–286.

Lansbergen, G., Akhmanova, A., 2006. Microtubule plus end: a hub of cellular activities.Traffic 7, 499–507.

Lansbergen, G., Grigoriev, I., Mimori-Kiyosue, Y., Ohtsuka, T., Higa, S., Kitajima, I.,Demmers, J., Galjart, N., Houtsmuller, A.B., Grosveld, F., Akhmanova, A., 2006.CLASPs attachmicrotubule plus ends to the cell cortex through a complex with LL5beta.Dev. Cell 11, 21–32.

Lansbergen, G., Komarova, Y., Modesti, M., Wyman, C., Hoogenraad, C.C.,Goodson, H.V., Lemaitre, R.P., Drechsel, D.N., van Munster, E., Gadella Jr., T.W.,Grosveld, F., Galjart, N., Borisy, G.G., Akhmanova, A., 2004. Conformational changesin CLIP-170 regulate its binding to microtubules and dynactin localization. J. Cell Biol.166, 1003–1014.

Lawrence, C.J., Dawe, R.K., Christie, K.R., Cleveland, D.W., Dawson, S.C., Endow, S.A.,Goldstein, L.S., Goodson, H.V., Hirokawa, N., Howard, J., Malmberg, R.L.,McIntosh, J.R., Miki, H., Mitchison, T.J., Okada, Y., Reddy, A.S., Saxton, W.M.,Schliwa, M., Scholey, J.M., Vale, R.D., Walczak, C.E., Wordeman, L., 2004.A standardized kinesin nomenclature. J. Cell Biol. 167, 19–22.

Laycock, J.E., Savoian, M.S., Glover, D.M., 2006. Antagonistic activities of Klp10A andOrbit regulate spindle length, bipolarity and function in vivo. J. Cell Sci. 119,2354–2361.

Leano, J. B., Rogers, S. L.,Slep, K. C., 2013. A Cryptic TOG Domain with a DistinctArchitecture Underlies CLASP-Dependent Bipolar Spindle Formation. Structure.

Lee, H.S., Komarova, Y.A., Nadezhdina, E.S., Anjum, R., Peloquin, J.G., Schober, J.M.,Danciu, O., van Haren, J., Galjart, N., Gygi, S.P., Akhmanova, A., Borisy, G.G., 2010.Phosphorylation controls autoinhibition of cytoplasmic linker protein-170. Mol. Biol.Cell 21, 2661–2673.

Lee, M.J., Gergely, F., Jeffers, K., Peak-Chew, S.Y., Raff, J.W., 2001. Msps/XMAP215interacts with the centrosomal protein D-TACC to regulate microtubule behaviour.Nat. Cell Biol. 3, 643–649.

Lee, T., Langford, K.J., Askham, J.M., Bruning-Richardson, A., Morrison, E.E., 2008.MCAK associates with EB1. Oncogene 27, 2494–2500.

Lemos, C.L., Sampaio, P., Maiato, H., Costa, M., Omel’yanchuk, L.V., Liberal, V.,Sunkel, C.E., 2000. Mast, a conserved microtubule-associated protein required forbipolar mitotic spindle organization. EMBO J. 19, 3668–3682.

Levesque, A.A., Compton, D.A., 2001. The chromokinesin Kid is necessary for chromo-some arm orientation and oscillation, but not congression, on mitotic spindles. J. CellBiol. 154, 1135–1146.

Li, H., Liu, X.S., Yang, X., Wang, Y., Turner, J.R., Liu, X., 2010. Phosphorylation ofCLIP-170 by Plk1 and CK2 promotes timely formation of kinetochore-microtubuleattachments. EMBO J. 29, 2953–2965.

Li, W., Moriwaki, T., Tani, T., Watanabe, T., Kaibuchi, K., Goshima, G., 2012. Recon-stitution of dynamic microtubules with Drosophila XMAP215, EB1, and Sentin. J. CellBiol. 199, 849–862.

Li, Y., Yu, W., Liang, Y., Zhu, X., 2007. Kinetochore dynein generates a poleward pullingforce to facilitate congression and full chromosome alignment. Cell Res. 17, 701–712.

Li, Y.Y., Yeh, E., Hays, T., Bloom, K., 1993. Disruption of mitotic spindle orientation in ayeast dynein mutant. Proc. Natl. Acad. Sci. U.S.A. 90, 10096–10100.




















































Lieuvin, A., Labbe, J.C., Doree, M., Job, D., 1994. Intrinsic microtubule stability in inter-phase cells. J. Cell Biol. 124, 985–996.

Ligon, L.A., Shelly, S.S., Tokito, M., Holzbaur, E.L., 2003. The microtubule plus-endproteins EB1 and dynactin have differential effects on microtubule polymerization.Mol. Biol. Cell 14, 1405–1417.

Liu, D., Vader, G., Vromans,M.J., Lampson,M.A., Lens, S.M., 2009a. Sensing chromosomebi-orientation by spatial separation of aurora B kinase from kinetochore substrates.Science 323, 1350–1353.

Liu, D., Zhang, N., Du, J., Cai, X., Zhu, M., Jin, C., Dou, Z., Feng, C., Yang, Y., Liu, L.,Takeyasu, K., Xie, W., Yao, X., 2006. Interaction of Skp1 with CENP-E at the mid-body is essential for cytokinesis. Biochem. Biophys. Res. Commun. 345, 394–402.

Liu, J., Wang, Z., Jiang, K., Zhang, L., Zhao, L., Hua, S., Yan, F., Yang, Y., Wang, D.,Fu, C., Ding, X., Guo, Z., Yao, X., 2009b. PRC1 cooperates with CLASP1 to organizecentral spindle plasticity in mitosis. J. Biol. Chem. 284, 23059–23071.

Logarinho, E., Maffini, S., Barisic, M., Marques, A., Toso, A., Meraldi, P., Maiato, H., 2012.CLASPs prevent irreversible multipolarity by ensuring spindle-pole resistance to tractionforces during chromosome alignment. Nat. Cell Biol. 14, 295–303.

Loncarek, J., Kisurina-Evgenieva, O., Vinogradova, T., Hergert, P., La Terra, S.,Kapoor, T.M., Khodjakov, A., 2007. The centromere geometry essential for keepingmitosis error free is controlled by spindle forces. Nature 450, 745–749.

Lopus, M., Manatschal, C., Buey, R. M., Bjelic, S., Miller, H. P., Steinmetz, M. O.,Wilson,L., 2012. Cooperative Stabilization of Microtubule Dynamics by EB1 and CLIP-170Involves Displacement of Stably Bound P(i) at Microtubule Ends. Biochemistry.

Louie, R.K., Bahmanyar, S., Siemers, K.A., Votin, V., Chang, P., Stearns, T., Nelson, W.J.,Barth, A.I., 2004. Adenomatous polyposis coli and EB1 localize in close proximity of themother centriole and EB1 is a functional component of centrosomes. J. Cell Sci. 117,1117–1128.

Mack, G.J., Compton, D.A., 2001. Analysis of mitotic microtubule-associated proteins usingmass spectrometry identifies astrin, a spindle-associated protein. Proc. Natl. Acad. Sci.U.S.A. 98, 14434–14439.

Maekawa, H., Schiebel, E., 2004. CLIP-170 family members: a motor-driven ride to micro-tubule plus ends. Dev. Cell 6, 746–748.

Maffini, S., Maia, A.R., Manning, A.L., Maliga, Z., Pereira, A.L., Junqueira, M.,Shevchenko, A., Hyman, A., Yates 3rd., J.R., Galjart, N., Compton, D.A.,Maiato, H., 2009. Motor-independent targeting of CLASPs to kinetochores byCENP-E promotes microtubule turnover and poleward flux. Curr. Biol. 19,1566–1572.

Magidson, V., O’Connell, C.B., Loncarek, J., Paul, R., Mogilner, A., Khodjakov, A., 2011.The spatial arrangement of chromosomes during prometaphase facilitates spindle assem-bly. Cell 146, 555–567.

Maia, A.R., Garcia, Z., Kabeche, L., Barisic, M., Maffini, S., Macedo-Ribeiro, S.,Cheeseman, I.M., Compton, D.A., Kaverina, I., Maiato, H., 2012. Cdk1 and Plk1mediate a CLASP2 phospho-switch that stabilizes kinetochore-microtubule attach-ments. J. Cell Biol. 199, 285–301.

Maiato, H., 2010. Mitosis: wisdom, knowledge, and information. Cell. Mol. Life Sci. 67,2141–2143.

Maiato, H., Fairley, E.A., Rieder, C.L., Swedlow, J.R., Sunkel, C.E., Earnshaw, W.C.,2003a. Human CLASP1 is an outer kinetochore component that regulates spindlemicrotubule dynamics. Cell 113, 891–904.

Maiato, H., Khodjakov, A., Rieder, C.L., 2005. Drosophila CLASP is required for the incor-poration of microtubule subunits into fluxing kinetochore fibres. Nat. Cell Biol. 7,42–47.



















































Maiato, H., Lince-Faria, M., 2010. The perpetual movements of anaphase. Cell. Mol. LifeSci. 67, 2251–2269.

Maiato, H., Logarinho, E., 2011. Motor-dependent and -independent roles of CENP-E atkinetochores: the cautionary tale of UA62784. Chem. Biol. 18, 679–680.

Maiato, H., Rieder, C.L., Earnshaw, W.C., Sunkel, C.E., 2003b. How do kinetochoresCLASP dynamic microtubules? Cell Cycle 2, 511–514.

Maiato, H., Sampaio, P., Lemos, C.L., Findlay, J., Carmena, M., Earnshaw, W.C.,Sunkel, C.E., 2002. MAST/Orbit has a role in microtubule-kinetochore attachmentand is essential for chromosome alignment and maintenance of spindle bipolarity.J. Cell Biol. 157, 749–760.

Maney,T.,Hunter,A.W.,Wagenbach,M.,Wordeman,L.,1998.Mitoticcentromere-associatedkinesin is important for anaphase chromosome segregation. J. Cell Biol. 142, 787–801.

Manning, A.L., Bakhoum, S.F., Maffini, S., Correia-Melo, C., Maiato, H., Compton, D.A.,2010. CLASP1, astrin and Kif2b form a molecular switch that regulates kinetochore-microtubuledynamics topromotemitotic progression and fidelity. EMBOJ. 29, 3531–3543.

Manning, A.L., Ganem, N.J., Bakhoum, S.F., Wagenbach, M., Wordeman, L.,Compton, D.A., 2007. The kinesin-13 proteins Kif2a, Kif2b, and Kif2c/MCAK havedistinct roles during mitosis in human cells. Mol. Biol. Cell 18, 2970–2979.

Margolis, R.L., Rauch, C.T., Pirollet, F., Job, D., 1990. Specific association of STOP pro-tein with microtubules in vitro and with stable microtubules in mitotic spindles of cul-tured cells. EMBO J. 9, 4095–4102.

Margolis, R.L., Wilson, L., 1978. Opposite end assembly and disassembly of microtubules atsteady state in vitro. Cell 13, 1–8.

Markus, S.M., Lee, W.L., 2011. Regulated offloading of cytoplasmic dynein from microtu-bule plus ends to the cortex. Dev. Cell 20, 639–651.

Markus, S.M., Punch, J.J., Lee, W.L., 2009. Motor- and tail-dependent targeting of dyneinto microtubule plus ends and the cell cortex. Curr. Biol. 19, 196–205.

Martinez-Lopez, M.J., Alcantara, S., Mascaro, C., Perez-Branguli, F., Ruiz-Lozano, P.,Maes, T., Soriano, E., Buesa, C., 2005. Mouse neuron navigator 1, a novel microtubule-associated protein involved in neuronal migration. Mol. Cell. Neurosci. 28, 599–612.

Matos, I., Pereira, A.J., Lince-Faria, M., Cameron, L.A., Salmon, E.D., Maiato, H., 2009.Synchronizing chromosome segregation by flux-dependent force equalization at kinet-ochores. J. Cell Biol. 186, 11–26.

Maurer, S.P., Bieling, P., Cope, J., Hoenger, A., Surrey, T., 2011. GTPgammaS microtu-bules mimic the growing microtubule end structure recognized by end-binding proteins(EBs). Proc. Natl. Acad. Sci. U.S.A. 108, 3988–3993.

Maurer, S.P., Fourniol, F.J., Bohner, G., Moores, C.A., Surrey, T., 2012. EBs recognize anucleotide-dependent structural cap at growing microtubule ends. Cell 149, 371–382.

McCartney, B.M., McEwen, D.G., Grevengoed, E., Maddox, P., Bejsovec, A., Peifer, M.,2001. Drosophila APC2 and Armadillo participate in tethering mitotic spindles to cor-tical actin. Nat. Cell Biol. 3 (10), 933–938.

Melki, R., Carlier, M.F., Pantaloni, D., Timasheff, S.N., 1989. Cold depolymerization ofmicrotubules to double rings: geometric stabilization of assemblies. Biochemistry 28,9143–9152.

Mennella, V., Rogers, G.C., Rogers, S.L., Buster, D.W., Vale, R.D., Sharp, D.J., 2005.Functionally distinct kinesin-13 family members cooperate to regulate microtubuledynamics during interphase. Nat. Cell Biol. 7, 235–245.

Miki, H., Okada, Y., Hirokawa, N., 2005. Analysis of the kinesin superfamily: insights intostructure and function. Trends Cell Biol. 15, 467–476.

Mimori-Kiyosue, Y., Grigoriev, I., Lansbergen, G., Sasaki, H., Matsui, C., Severin, F.,Galjart, N., Grosveld, F., Vorobjev, I., Tsukita, S., Akhmanova, A., 2005. CLASP1





















































and CLASP2 bind to EB1 and regulate microtubule plus-end dynamics at the cell cortex.J. Cell Biol. 168, 141–153.

Mimori-Kiyosue, Y., Grigoriev, I., Sasaki, H., Matsui, C., Akhmanova, A., Tsukita, S.,Vorobjev, I., 2006. Mammalian CLASPs are required for mitotic spindle organizationand kinetochore alignment. Genes Cells 11, 845–857.

Mimori-Kiyosue, Y., Shiina, N., Tsukita, S., 2000a. Adenomatous polyposis coli (APC) pro-tein moves along microtubules and concentrates at their growing ends in epithelial cells.J. Cell Biol. 148, 505–518.

Mimori-Kiyosue, Y., Shiina, N., Tsukita, S., 2000b. The dynamic behavior of theAPC-binding protein EB1 on the distal ends of microtubules. Curr. Biol. 10,865–868.

Mishima, M., Maesaki, R., Kasa, M., Watanabe, T., Fukata, M., Kaibuchi, K.,Hakoshima, T., 2007. Structural basis for tubulin recognition by cytoplasmic linker pro-tein 170 and its autoinhibition. Proc. Natl. Acad. Sci. U.S.A. 104, 10346–10351.

Mitchison, T., Evans, L., Schulze, E., Kirschner, M., 1986. Sites of microtubule assembly anddisassembly in the mitotic spindle. Cell 45, 515–527.

Mitchison, T., Kirschner, M., 1984. Dynamic instability of microtubule growth. Nature 312,237–242.

Mitchison, T.J., 1989. Polewards microtubule flux in the mitotic spindle: evidence fromphotoactivation of fluorescence. J. Cell Biol. 109, 637–652.

Mogilner, A., Oster, G., 2003. Polymer motors: pushing out the front and pulling up theback. Curr. Biol. 13, R721–R733.

Montenegro Gouveia, S., Leslie, K., Kapitein, L.C., Buey, R.M., Grigoriev, I.,Wagenbach, M., Smal, I., Meijering, E., Hoogenraad, C.C., Wordeman, L.,Steinmetz, M.O., Akhmanova, A., 2010. In vitro reconstitution of the functionalinterplay between MCAK and EB3 at microtubule plus ends. Curr. Biol. 20,1717–1722.

Moore, A., Wordeman, L., 2004. The mechanism, function and regulation ofdepolymerizing kinesins during mitosis. Trends Cell Biol. 14, 537–546.

Moore, A.T., Rankin, K.E., von Dassow, G., Peris, L., Wagenbach, M., Ovechkina, Y.,Andrieux, A., Job, D., Wordeman, L., 2005. MCAK associates with the tips of polymer-izing microtubules. J. Cell Biol. 169, 391–397.

Morrison, E.E., Askham, J.M., 2001. EB 1 immunofluorescence reveals an increase in grow-ing astral microtubule length and number during anaphase in NRK-52E cells. Eur. J.Cell Biol. 80, 749–753.

Morrison, E.E., Wardleworth, B.N., Askham, J.M., Markham, A.F., Meredith, D.M., 1998.EB1, a protein which interacts with the APC tumour suppressor, is associated with themicrotubule cytoskeleton throughout the cell cycle. Oncogene 17, 3471–3477.

Moseley, J.B., Bartolini, F., Okada, K., Wen, Y., Gundersen, G.G., Goode, B.L., 2007.Regulated binding of adenomatous polyposis coli protein to actin. J. Biol. Chem.282, 12661–12668.

Moutinho-Pereira, S., Debec, A., Maiato, H., 2009. Microtubule cytoskeleton remodelingby acentriolar microtubule-organizing centers at the entry and exit from mitosis in Dro-sophila somatic cells. Mol. Biol. Cell 20, 2796–2808.

Mullins, J.M., Biesele, J.J., 1977. Terminal phase of cytokinesis in D-98s cells. J. Cell Biol. 73,672–684.

Munemitsu, S., Souza, B., Muller, O., Albert, I., Rubinfeld, B., Polakis, P., 1994. The APCgene product associates with microtubules in vivo and promotes their assembly in vitro.Cancer Res. 54, 3676–3681.

Murray, A.W., Kirschner, M.W., 1989. Cyclin synthesis drives the early embryonic cellcycle. Nature 339, 275–280.





















































Nakagawa, H., Koyama, K., Murata, Y., Morito, M., Akiyama, T., Nakamura, Y., 2000.EB3, a novel member of the EB1 family preferentially expressed in the central nervoussystem, binds to a CNS-specific APC homologue. Oncogene 19, 210–216.

Nakagawa, H., Murata, Y., Koyama, K., Fujiyama, A., Miyoshi, Y., Monden, M.,Akiyama, T., Nakamura, Y., 1998. Identification of a brain-specific APC homologue,APCL, and its interaction with beta-catenin. Cancer Res. 58, 5176–5181.

Nakamura, M., Zhou, X.Z., Lu, K.P., 2001. Critical role for the EB1 and APC interaction inthe regulation of microtubule polymerization. Curr. Biol. 11, 1062–1067.

Nakano, A., Kato, H., Watanabe, T., Min, K.D., Yamazaki, S., Asano, Y., Seguchi, O.,Higo, S., Shintani, Y., Asanuma, H., Asakura, M., Minamino, T., Kaibuchi, K.,Mochizuki, N., Kitakaze, M., Takashima, S., 2010. AMPK controls the speed of micro-tubule polymerization and directional cell migration through CLIP-170 phosphoryla-tion. Nat. Cell Biol. 12, 583–590.

Neuwald, A.F., Hirano, T., 2000. HEAT repeats associated with condensins, cohesins, andother complexes involved in chromosome-related functions. Genome Res. 10,1445–1452.

Nicklas, R.B., 1989. The motor for poleward chromosome movement in anaphase is in ornear the kinetochore. J. Cell Biol. 109, 2245–2255.

Noda, Y., Sato-Yoshitake, R., Kondo, S., Nangaku, M., Hirokawa, N., 1995. KIF2 is anew microtubule-based anterograde motor that transports membranous organelles dis-tinct from those carried by kinesin heavy chain or KIF3A/B. J. Cell Biol. 129,157–167.

O’Connell, C.B., Wang, Y.L., 2000. Mammalian spindle orientation and position respondto changes in cell shape in a dynein-dependent fashion. Mol. Biol. Cell 11,1765–1774.

Ogawa, T., Nitta, R., Okada, Y., Hirokawa, N., 2004. A common mechanism formicrotubule destabilizers-M type kinesins stabilize curling of the protofilament usingthe class-specific neck and loops. Cell 116, 591–602.

Oguchi, Y., Uchimura, S., Ohki, T., Mikhailenko, S.V., Ishiwata, S., 2011. The bidirec-tional depolymerizer MCAK generates force by disassembling both microtubule ends.Nat. Cell Biol. 13, 846–852.

Ohkura, H., Garcia,M.A., Toda, T., 2001. Dis1/TOGuniversal microtubule adaptors—oneMAP for all? J. Cell Sci. 114, 3805–3812.

Oiwa, K., Sakakibara, H., 2005. Recent progress in dynein structure and mechanism. Curr.Opin. Cell Biol. 17, 98–103.

Okada, Y., Hirokawa, N., 1999. A processive single-headed motor: kinesin superfamilyprotein KIF1A. Science 283, 1152–1157.

Park, E.C., Lee, H., Hong, Y., Kim, M.J., Lee, Z.W., Kim, S.I., Kim, S., Kim, G.H.,Han, J.K., 2012. Analysis of the expression of microtubule plus-end tracking proteins(þTIPs) during Xenopus laevis embryogenesis. Gene Expr. Patterns 12, 204–212.

Patel, K., Nogales, E., Heald, R., 2012. Multiple domains of human CLASP contribute tomicrotubule dynamics and organization in vitro and in Xenopus egg extracts. Cytoskel-eton (Hoboken) 69, 155–165.

Paul, R., Wollman, R., Silkworth, W.T., Nardi, I.K., Cimini, D., Mogilner, A., 2009.Computer simulations predict that chromosome movements and rotations acceleratemitotic spindle assembly without compromising accuracy. Proc. Natl. Acad. Sci.U.S.A. 106, 15708–15713.

Pereira, A.J., Maiato, H., 2010. Improved kymography tools and its applications to mitosis.Methods 51, 214–219.

Pereira, A.J., Maiato, H., 2012. Maturation of the kinetochore-microtubule interface and themeaning of metaphase. Chromosome Res. 20, 563–577.






















































Pereira, A.L., Pereira, A.J., Maia, A.R., Drabek, K., Sayas, C.L., Hergert, P.J., Lince-Faria,M., Matos, I., Duque, C., Stepanova, T., Rieder, C.L., Earnshaw, W.C., Galjart, N.,Maiato, H., 2006. Mammalian CLASP1 and CLASP2 cooperate to ensure mitotic fidel-ity by regulating spindle and kinetochore function. Mol. Biol. Cell 17, 4526–4542.

Perez, F., Diamantopoulos, G.S., Stalder, R., Kreis, T.E., 1999. CLIP-170 highlights grow-ing microtubule ends in vivo. Cell 96, 517–527.

Peris, L., Thery,M., Faure, J., Saoudi, Y., Lafanechere, L., Chilton, J.K., Gordon-Weeks, P.,Galjart, N., Bornens, M., Wordeman, L., Wehland, J., Andrieux, A., Job, D., 2006.Tubulin tyrosination is a major factor affecting the recruitment of CAP-Gly proteinsat microtubule plus ends. J. Cell Biol. 174, 839–849.

Pfarr, C.M., Coue, M., Grissom, P.M., Hays, T.S., Porter, M.E., McIntosh, J.R., 1990.Cytoplasmic dynein is localized to kinetochores during mitosis. Nature 345, 263–265.

Piehl, M., Cassimeris, L., 2003. Organization and dynamics of growing microtubule plusends during early mitosis. Mol. Biol. Cell 14, 916–925.

Pierre, P., Pepperkok, R., Kreis, T.E., 1994. Molecular characterization of two functionaldomains of CLIP-170 in vivo. J. Cell Sci. 107 (Pt. 7), 1909–1920.

Piperno, G., LeDizet, M., Chang, X.J., 1987. Microtubules containing acetylated alpha-tubulin in mammalian cells in culture. J. Cell Biol. 104, 289–302.

Popov, A.V., Pozniakovsky, A., Arnal, I., Antony, C., Ashford, A.J., Kinoshita, K.,Tournebize, R., Hyman, A.A., Karsenti, E., 2001. XMAP215 regulates microtubuledynamics through two distinct domains. EMBO J. 20, 397–410.

Purohit, A., Tynan, S.H., Vallee, R., Doxsey, S.J., 1999. Direct interaction of pericentrinwith cytoplasmic dynein light intermediate chain contributes to mitotic spindle organi-zation. J. Cell Biol. 147, 481–492.

Rankin, K.E., Wordeman, L., 2010. Long astral microtubules uncouple mitotic spindlesfrom the cytokinetic furrow. J. Cell Biol. 190, 35–43.

Rehberg, M., Graf, R., 2002. Dictyostelium EB1 is a genuine centrosomal componentrequired for proper spindle formation. Mol. Biol. Cell 13, 2301–2310.

Reis, R., Feijao, T., Gouveia, S., Pereira, A.J., Matos, I., Sampaio, P., Maiato, H.,Sunkel, C.E., 2009. Dynein and mast/orbit/CLASP have antagonistic roles in regulatingkinetochore-microtubule plus-end dynamics. J. Cell Sci. 122, 2543–2553.

Rickard, J.E., Kreis, T.E., 1991. Binding of pp 170 to microtubules is regulated by phosphor-ylation. J. Biol. Chem. 266, 17597–17605.

Rieder, C.L., Alexander, S.P., 1990. Kinetochores are transported poleward along a singleastral microtubule during chromosome attachment to the spindle in newt lung cells.J. Cell Biol. 110, 81–95.

Rieder, C.L., Cole, R.W., Khodjakov, A., Sluder, G., 1995. The checkpoint delaying ana-phase in response to chromosome monoorientation is mediated by an inhibitory signalproduced by unattached kinetochores. J. Cell Biol. 130, 941–948.

Rieder, C.L., Khodjakov, A., Paliulis, L.V., Fortier, T.M., Cole, R.W., Sluder, G., 1997.Mitosis in vertebrate somatic cells with two spindles: implications for the metaphase/ana-phase transition checkpoint and cleavage. Proc. Natl. Acad. Sci. U.S.A. 94, 5107–5112.

Rieder, C.L., Maiato, H., 2004. Stuck in division or passing through: what happens whencells cannot satisfy the spindle assembly checkpoint. Dev. Cell 7, 637–651.

Rieder, C.L., Salmon, E.D., 1994. Motile kinetochores and polar ejection forces dictatechromosome position on the vertebrate mitotic spindle. J. Cell Biol. 124, 223–233.

Rizk, R.S., Bohannon, K.P., Wetzel, L.A., Powers, J., Shaw, S.L., Walczak, C.E., 2009.MCAK and paclitaxel have differential effects on spindle microtubule organizationand dynamics. Mol. Biol. Cell 20, 1639–1651.

Rodionov, V.I., Borisy, G.G., 1997. Microtubule treadmilling in vivo. Science 275,215–218.





















































Rogers, S.L., Rogers, G.C., Sharp, D.J., Vale, R.D., 2002. Drosophila EB1 is important forproper assembly, dynamics, and positioning of the mitotic spindle. J. Cell Biol. 158,873–884.

Rogers, S.L., Wiedemann, U., Hacker, U., Turck, C., Vale, R.D., 2004. Drosophila Rho-GEF2 associates with microtubule plus ends in an EB1-dependent manner. Curr. Biol.14, 1827–1833.

Rosa, J., Canovas, P., Islam, A., Altieri, D.C., Doxsey, S.J., 2006. Survivin modulates micro-tubule dynamics and nucleation throughout the cell cycle. Mol. Biol. Cell 17,1483–1493.

Rusan, N.M., Fagerstrom, C.J., Yvon, A.M., Wadsworth, P., 2001. Cell cycle-dependentchanges in microtubule dynamics in living cells expressing green fluorescent protein-alpha tubulin. Mol. Biol. Cell 12, 971–980.

Salmon, E.D., Leslie, R.J., Saxton, W.M., Karow, M.L., McIntosh, J.R., 1984. Spindlemicrotubule dynamics in sea urchin embryos: analysis using a fluorescein-labeled tubulinand measurements of fluorescence redistribution after laser photobleaching. J. Cell Biol.99, 2165–2174.

Sammak, P.J., Borisy, G.G., 1988. Direct observation of microtubule dynamics in living cells.Nature 332, 724–726.

Samora, C.P., Mogessie, B., Conway, L., Ross, J.L., Straube, A., McAinsh, A.D., 2011.MAP4 and CLASP1 operate as a safety mechanism to maintain a stable spindle positionin mitosis. Nat. Cell Biol. 13, 1040–1050.

Sandblad, L., Busch, K.E., Tittmann, P., Gross, H., Brunner, D., Hoenger, A., 2006. TheSchizosaccharomyces pombe EB1 homolog Mal3p binds and stabilizes the microtubulelattice seam. Cell 127, 1415–1424.

Sanhaji, M., Friel, C.T., Kreis, N.N., Kramer, A., Martin, C., Howard, J., Strebhardt, K.,Yuan, J., 2010. Functional and spatial regulation of mitotic centromere-associatedkinesin by cyclin-dependent kinase 1. Mol. Cell. Biol. 30, 2594–2607.

Sardar, H.S., Luczak, V.G., Lopez, M.M., Lister, B.C., Gilbert, S.P., 2010. Mitotic kinesinCENP-E promotes microtubule plus-end elongation. Curr. Biol. 20, 1648–1653.

Sato, S., Cerny, R.L., Buescher, J.L., Ikezu, T., 2006. Tau-tubulin kinase 1 (TTBK1), aneuron-specific tau kinase candidate, is involved in tau phosphorylation and aggregation.J. Neurochem. 98, 1573–1584.

Savoian, M.S., Earnshaw, W.C., Khodjakov, A., Rieder, C.L., 1999. Cleavage furrowsformed between centrosomes lacking an intervening spindle and chromosomes containmicrotubule bundles, INCENP, and CHO1 but not CENP-E. Mol. Biol. Cell 10,297–311.

Saxton, W.M., Stemple, D.L., Leslie, R.J., Salmon, E.D., Zavortink, M., McIntosh, J.R.,1984. Tubulin dynamics in cultured mammalian cells. J. Cell Biol. 99, 2175–2186.

Schaar, B.T., Chan, G.K., Maddox, P., Salmon, E.D., Yen, T.J., 1997. CENP-E function atkinetochores is essential for chromosome alignment. J. Cell Biol. 139, 1373–1382.

Scheel, J., Pierre, P., Rickard, J.E., Diamantopoulos, G.S., Valetti, C., van der Goot, F.G.,Haner, M., Aebi, U., Kreis, T.E., 1999. Purification and analysis of authentic CLIP-170and recombinant fragments. J. Biol. Chem. 274, 25883–25891.

Schek 3rd., H.T., Gardner, M.K., Cheng, J., Odde, D.J., Hunt, A.J., 2007. Microtubuleassembly dynamics at the nanoscale. Curr. Biol. 17, 1445–1455.

Schmidt, J.C., Kiyomitsu, T., Hori, T., Backer, C.B., Fukagawa, T., Cheeseman, I.M.,2010. Aurora B kinase controls the targeting of the Astrin-SKAP complex to biorientedkinetochores. J. Cell Biol. 191, 269–280.

Schmidt, N., Basu, S., Sladecek, S., Gatti, S., van Haren, J., Treves, S., Pielage, J., Galjart, N.,Brenner, H.R., 2012. Agrin regulates CLASP2-mediated capture of microtubules at theneuromuscular junction synaptic membrane. J. Cell Biol. 198, 421–437.





















































Schroeder, T.E., 1972. The contractile ring. II. Determining its brief existence, volumetricchanges, and vital role in cleaving Arbacia eggs. J. Cell Biol. 53, 419–434.

Schroeder, T.E., 1973. Actin in dividing cells: contractile ring filaments bind heavy mero-myosin. Proc. Natl. Acad. Sci. U.S.A. 70, 1688–1692.

Schulze, E., Kirschner, M., 1988. New features of microtubule behaviour observed in vivo.Nature 334, 356–359.

Schuster, M., Kilaru, S., Ashwin, P., Lin, C., Severs, N.J., Steinberg, G., 2011. Controlledand stochastic retention concentrates dynein at microtubule ends to keep endosomes ontrack. EMBO J. 30, 652–664.

Schuyler, S.C., Pellman, D., 2001. Microtubule “plus-end-tracking proteins”: The end isjust the beginning. Cell 105, 421–424.

Schwartz, K., Richards, K., Botstein, D., 1997. BIM1 encodes a microtubule-binding pro-tein in yeast. Mol. Biol. Cell 8, 2677–2691.

Seetapun, D., Castle, B.T., McIntyre, A.J., Tran, P.T., Odde, D.J., 2012. Estimating themicrotubule GTP cap size in vivo. Curr. Biol. 22, 1681–1687.

Sheffield, P.J., Garrard, S., Caspi, M., Aoki, J., Arai, H., Derewenda, U., Inoue, K., Suter, B.,Reiner, O., Derewenda, Z.S., 2000. Homologs of the alpha- and beta-subunits of mam-malian brain platelet-activating factor acetylhydrolase Ib in the Drosophila melanogastergenome. Proteins 39, 1–8.

Shu, H.B., Li, Z., Palacios, M.J., Li, Q., Joshi, H.C., 1995. A transient association of gamma-tubulin at the midbody is required for the completion of cytokinesis during themammalian cell division. J. Cell Sci. 108 (Pt. 9), 2955–2962.

Skop, A.R., Liu, H., Yates 3rd., J., Meyer, B.J., Heald, R., 2004. Dissection of the mam-malian midbody proteome reveals conserved cytokinesis mechanisms. Science 305,61–66.

Skoufias, D.A., Indorato, R.L., Lacroix, F., Panopoulos, A., Margolis, R.L., 2007. Mitosispersists in the absence of Cdk1 activity when proteolysis or protein phosphatase activity issuppressed. J. Cell Biol. 179, 671–685.

Slep, K.C., 2009. The role of TOG domains in microtubule plus end dynamics. Biochem.Soc. Trans. 37, 1002–1006.

Slep, K.C., 2010. Structural and mechanistic insights into microtubule end-binding proteins.Curr. Opin. Cell Biol. 22, 88–95.

Slep, K.C., Rogers, S.L., Elliott, S.L., Ohkura, H., Kolodziej, P.A., Vale, R.D., 2005. Struc-tural determinants for EB1-mediated recruitment of APC and spectraplakins to themicrotubule plus end. J. Cell Biol. 168, 587–598.

Slep, K.C., Vale, R.D., 2007. Structural basis of microtubule plus end tracking byXMAP215, CLIP-170, and EB1. Mol. Cell 27, 976–991.

Smith, K.J., Johnson, K.A., Bryan, T.M., Hill, D.E., Markowitz, S., Willson, J.K.,Paraskeva, C., Petersen, G.M., Hamilton, S.R., Vogelstein, B., et al., 1993. TheAPC gene product in normal and tumor cells. Proc. Natl. Acad. Sci. U.S.A. 90,2846–2850.

Sousa, A., Reis, R., Sampaio, P., Sunkel, C.E., 2007. The Drosophila CLASP homologue,Mast/Orbit regulates the dynamic behaviour of interphase microtubules by promotingthe pause state. Cell Motil. Cytoskeleton 64, 605–620.

Splinter, D., Tanenbaum, M.E., Lindqvist, A., Jaarsma, D., Flotho, A., Yu, K.L.,Grigoriev, I., Engelsma, D., Haasdijk, E.D., Keijzer, N., Demmers, J., Fornerod, M.,Melchior, F., Hoogenraad, C.C., Medema, R.H., Akhmanova, A., 2010. Bicaudal D2,dynein, and kinesin-1 associate with nuclear pore complexes and regulate centrosomeand nuclear positioning during mitotic entry. PLoS Biol. 8, e1000350.

Steinmetz, M.O., Akhmanova, A., 2008. Capturing protein tails by CAP-Gly domains.Trends Biochem. Sci. 33, 535–545.





















































Steuer, E.R., Wordeman, L., Schroer, T.A., Sheetz, M.P., 1990. Localization of cytoplasmicdynein to mitotic spindles and kinetochores. Nature 345, 266–268.

Stout, J.R., Yount, A.L., Powers, J.A., Leblanc, C., Ems-McClung, S.C., Walczak, C.E.,2011. Kif18B interacts with EB1 and controls astral microtubule length during mitosis.Mol. Biol. Cell 22, 3070–3080.

Straight, A.F., Cheung, A., Limouze, J., Chen, I., Westwood, N.J., Sellers, J.R.,Mitchison, T.J., 2003. Dissecting temporal and spatial control of cytokinesis with amyosin II Inhibitor. Science 299, 1743–1747.

Straube, A., Merdes, A., 2007. EB3 regulates microtubule dynamics at the cell cortex and isrequired for myoblast elongation and fusion. Curr. Biol. 17, 1318–1325.

Strickland, L.I., Donnelly, E.J., Burgess, D.R., 2005a. Induction of cytokinesis is indepen-dent of precisely regulated microtubule dynamics. Mol. Biol. Cell 16, 4485–4494.

Strickland, L.I., Wen, Y., Gundersen, G.G., Burgess, D.R., 2005b. Interaction between EB1and p150glued is required for anaphase astral microtubule elongation and stimulation ofcytokinesis. Curr. Biol. 15, 2249–2255.

Su, L.K., Burrell, M., Hill, D.E., Gyuris, J., Brent, R., Wiltshire, R., Trent, J.,Vogelstein, B., Kinzler, K.W., 1995. APC binds to the novel protein EB1. CancerRes. 55, 2972–2977.

Su, L.K., Qi, Y., 2001. Characterization of human MAPRE genes and their proteins.Genomics 71, 142–149.

Sullivan, D.S., Huffaker, T.C., 1992. Astral microtubules are not required for anaphase B inSaccharomyces cerevisiae. J. Cell Biol. 119, 379–388.

Sun, L., Gao, J., Dong, X., Liu, M., Li, D., Shi, X., Dong, J.T., Lu, X., Liu, C., Zhou, J.,2008. EB1 promotes Aurora-B kinase activity through blocking its inactivation byprotein phosphatase 2A. Proc. Natl. Acad. Sci. U.S.A. 105, 7153–7158.

Tai, C.Y., Dujardin, D.L., Faulkner, N.E., Vallee, R.B., 2002. Role of dynein, dynactin, andCLIP-170 interactions in LIS1 kinetochore function. J. Cell Biol. 156, 959–968.

Tanaka, T., Serneo, F.F., Higgins, C., Gambello, M.J., Wynshaw-Boris, A., Gleeson, J.G.,2004. Lis1 and doublecortin function with dynein to mediate coupling of the nucleus tothe centrosome in neuronal migration. J. Cell Biol. 165, 709–721.

Tanenbaum, M.E., Galjart, N., van Vugt, M.A., Medema, R.H., 2006. CLIP-170 facilitatesthe formation of kinetochore-microtubule attachments. EMBO J. 25, 45–57.

Tanenbaum, M.E., Macurek, L., Galjart, N., Medema, R.H., 2008. Dynein, Lis1 and CLIP-170 counteract Eg5-dependent centrosome separation during bipolar spindle assembly.EMBO J. 27, 3235–3245.

Tanenbaum,M.E., Medema, R.H., 2011. Localized Aurora B activity spatially controls non-kinetochore microtubules during spindle assembly. Chromosoma 120, 599–607.

Tanenbaum, M.E., Medema, R.H., Akhmanova, A., 2011. Regulation of localization andactivity of the microtubule depolymerase MCAK. Bioarchitecture 1, 80–87.

Tarricone, C., Perrina, F., Monzani, S., Massimiliano, L., Kim, M.H., Derewenda, Z.S.,Knapp, S., Tsai, L.H., Musacchio, A., 2004. Coupling PAF signaling to dynein regula-tion: structure of LIS1 in complex with PAF-acetylhydrolase. Neuron 44, 809–821.

Thein, K.H., Kleylein-Sohn, J., Nigg, E.A., Gruneberg, U., 2007. Astrin is required for themaintenance of sister chromatid cohesion and centrosome integrity. J. Cell Biol. 178,345–354.

Tighe, A., Johnson, V.L., Albertella, M., Taylor, S.S., 2001. Aneuploid colon cancer cellshave a robust spindle checkpoint. EMBO Rep. 2, 609–614.

Tighe, A., Johnson, V.L., Taylor, S.S., 2004. Truncating APC mutations have dominanteffects on proliferation, spindle checkpoint control, survival and chromosome stability.J. Cell Sci. 117, 6339–6353.

Tirnauer, J.S., Bierer, B.E., 2000. EB1 proteins regulate microtubule dynamics, cell polarity,and chromosome stability. J. Cell Biol. 149, 761–766.






















































Tirnauer, J.S., Canman, J.C., Salmon, E.D., Mitchison, T.J., 2002a. EB1 targets to kineto-chores with attached, polymerizing microtubules. Mol. Biol. Cell 13, 4308–4316.

Tirnauer, J.S., Grego, S., Salmon, E.D., Mitchison, T.J., 2002b. EB1-microtubule interac-tions in Xenopus egg extracts: role of EB1 in microtubule stabilization and mechanismsof targeting to microtubules. Mol. Biol. Cell 13, 3614–3626.

Tirnauer, J.S., O’Toole, E., Berrueta, L., Bierer, B.E., Pellman, D., 1999. Yeast Bim1p pro-motes the G1-specific dynamics of microtubules. J. Cell Biol. 145, 993–1007.

Tolic-Norrelykke, I.M., 2008. Push-me-pull-you: how microtubules organize the cell inte-rior. Eur. Biophys. J. 37, 1271–1278.

Tournebize, R., Andersen, S.S., Verde, F., Doree, M., Karsenti, E., Hyman, A.A., 1997.Distinct roles of PP1 and PP2A-like phosphatases in control of microtubule dynamicsduring mitosis. EMBO J. 16, 5537–5549.

Tournebize, R., Popov, A., Kinoshita, K., Ashford, A.J., Rybina, S., Pozniakovsky, A.,Mayer, T.U., Walczak, C.E., Karsenti, E., Hyman, A.A., 2000. Control of microtubuledynamics by the antagonistic activities of XMAP215 and XKCM1 in Xenopus eggextracts. Nat. Cell Biol. 2, 13–19.

Toyoshima, F., Nishida, E., 2007. Integrin-mediated adhesion orients the spindle parallel tothe substratum in an EB1- and myosin X-dependent manner. EMBO J. 26, 1487–1498.

Tsai, J.W., Bremner, K.H., Vallee, R.B., 2007. Dual subcellular roles for LIS1 and dynein inradial neuronal migration in live brain tissue. Nat. Neurosci. 10, 970–979.

Tsvetkov, A.S., Samsonov, A., Akhmanova, A., Galjart, N., Popov, S.V., 2007.Microtubule-binding proteins CLASP1 and CLASP2 interact with actin filaments. CellMotil. Cytoskeleton 64, 519–530.

Uehara, R., Nozawa, R.S., Tomioka, A., Petry, S., Vale, R.D., Obuse, C., Goshima, G.,2009. The augmin complex plays a critical role in spindle microtubule generation formitotic progression and cytokinesis in human cells. Proc. Natl. Acad. Sci. U.S.A.106, 6998–7003.

Vaisberg, E.A., Koonce, M.P., McIntosh, J.R., 1993. Cytoplasmic dynein plays a role inmammalian mitotic spindle formation. J. Cell Biol. 123, 849–858.

Vale, R.D., Fletterick, R.J., 1997. The design plan of kinesin motors. Annu. Rev. Cell Dev.Biol. 13, 745–777.

Vallee, R.B., Tai, C., Faulkner, N.E., 2001. LIS1: cellular function of a disease-causing gene.Trends Cell Biol. 11, 155–160.

van Haren, J., Draegestein, K., Keijzer, N., Abrahams, J.P., Grosveld, F., Peeters, P.J.,Moechars, D., Galjart, N., 2009. Mammalian navigators are microtubule plus-end track-ing proteins that can reorganize the cytoskeleton to induce neurite-like extensions. CellMotil. Cytoskeleton 66, 824–838.

Varma, D., Monzo, P., Stehman, S.A., Vallee, R.B., 2008. Direct role of dynein motor instable kinetochore-microtubule attachment, orientation, and alignment. J. Cell Biol.182, 1045–1054.

Vasquez, R.J., Gard, D.L., Cassimeris, L., 1994. XMAP from Xenopus eggs promotes rapidplus end assembly of microtubules and rapid microtubule polymer turnover. J. Cell Biol.127, 985–993.

Vaughan, K.T., Tynan, S.H., Faulkner, N.E., Echeverri, C.J., Vallee, R.B., 1999.Colocalization of cytoplasmic dynein with dynactin and CLIP-170 at microtubule distalends. J. Cell Sci. 112 (Pt. 10), 1437–1447.

Verde, F., Labbe, J.C., Doree, M., Karsenti, E., 1990. Regulation of microtubuledynamics by cdc2 protein kinase in cell-free extracts of Xenopus eggs. Nature 343,233–238.

Vitre, B., Coquelle, F.M., Heichette, C., Garnier, C., Chretien, D., Arnal, I., 2008. EB1regulates microtubule dynamics and tubulin sheet closure in vitro. Nat. Cell Biol. 10,415–421.






















































Voter, W.A., O’Brien, E.T., Erickson, H.P., 1991. Dilution-induced disassembly of micro-tubules: relation to dynamic instability and the GTP cap. Cell Motil. Cytoskeleton 18,55–62.

Walczak, C.E., 2003. The Kin I kinesins are microtubule end-stimulated ATPases. Mol. Cell11, 286–288.

Walczak, C.E., Mitchison, T.J., Desai, A., 1996. XKCM1: a Xenopus kinesin-related pro-tein that regulates microtubule dynamics during mitotic spindle assembly. Cell 84,37–47.

Walker, R.A., O’Brien, E.T., Pryer, N.K., Soboeiro, M.F., Voter, W.A., Erickson, H.P.,Salmon, E.D., 1988. Dynamic instability of individual microtubules analyzed byvideo light microscopy: rate constants and transition frequencies. J. Cell Biol. 107,1437–1448.

Walker, R.A., Pryer, N.K., Salmon, E.D., 1991. Dilution of individual microtubulesobserved in real time in vitro: evidence that cap size is small and independent of elon-gation rate. J. Cell Biol. 114, 73–81.

Wandke, C., Barisic, M., Sigl, R., Rauch, V., Wolf, F., Amaro, A.C., Tan, C.H.,Pereira, A.J., Kutay, U., Maiato, H., Meraldi, P., Geley, S., 2012. Human chro-mokinesins promote chromosome congression and spindle microtubule dynamics duringmitosis. J. Cell Biol. 198, 847–863.

Wang, P.J., Huffaker, T.C., 1997. Stu2p: a microtubule-binding protein that is an essentialcomponent of the yeast spindle pole body. J. Cell Biol. 139, 1271–1280.

Wang, Z., Khan, S., Sheetz, M.P., 1995. Single cytoplasmic dynein molecule movements:characterization and comparison with kinesin. Biophys. J. 69, 2011–2023.

Wang, Z., Wu, T., Shi, L., Zhang, L., Zheng, W., Qu, J.Y., Niu, R., Qi, R.Z., 2010. Con-servedmotif of CDK5RAP2mediates its localization to centrosomes and the Golgi com-plex. J. Biol. Chem. 285, 22658–22665.

Waterman-Storer, C.M., Desai, A., Bulinski, J.C., Salmon, E.D., 1998. Fluorescent specklemicroscopy, a method to visualize the dynamics of protein assemblies in living cells.Curr. Biol. 8, 1227–1230.

Waters, J.C., Mitchison, T.J., Rieder, C.L., Salmon, E.D., 1996. The kinetochore micro-tubule minus-end disassembly associated with poleward flux produces a force that cando work. Mol. Biol. Cell 7, 1547–1558.

Weisbrich, A., Honnappa, S., Jaussi, R., Okhrimenko, O., Frey, D., Jelesarov, I.,Akhmanova, A., Steinmetz, M.O., 2007. Structure-function relationship of CAP-Glydomains. Nat. Struct. Mol. Biol. 14, 959–967.

Wen, Y., Eng, C.H., Schmoranzer, J., Cabrera-Poch, N., Morris, E.J., Chen, M.,Wallar, B.J., Alberts, A.S., Gundersen, G.G., 2004. EB1 and APC bind to mDia tostabilize microtubules downstream of Rho and promote cell migration. Nat. Cell Biol.6, 820–830.

Wheatley, S.P., Hinchcliffe, E.H., Glotzer, M., Hyman, A.A., Sluder, G., Wang, Y., 1997.CDK1 inactivation regulates anaphase spindle dynamics and cytokinesis in vivo. J. CellBiol. 138, 385–393.

Wheatley, S.P., Wang, Y., 1996. Midzone microtubule bundles are continuously requiredfor cytokinesis in cultured epithelial cells. J. Cell Biol. 135, 981–989.

Whyte, J., Bader, J.R., Tauhata, S.B., Raycroft, M., Hornick, J., Pfister, K.K., Lane, W.S.,Chan, G.K., Hinchcliffe, E.H., Vaughan, P.S., Vaughan, K.T., 2008. Phosphorylationregulates targeting of cytoplasmic dynein to kinetochores during mitosis. J. Cell Biol.183, 819–834.

Wieland, G., Orthaus, S., Ohndorf, S., Diekmann, S., Hemmerich, P., 2004. Functionalcomplementation of human centromere protein A (CENP-A) by Cse4p from Saccha-romyces cerevisiae. Mol. Cell. Biol. 24, 6620–6630.





















































Wittmann, T., Waterman-Storer, C.M., 2005. Spatial regulation of CLASP affinity formicrotubules by Rac1 and GSK3beta in migrating epithelial cells. J. Cell Biol. 169,929–939.

Wollman, R., Cytrynbaum, E.N., Jones, J.T., Meyer, T., Scholey, J.M., Mogilner, A., 2005.Efficient chromosome capture requires a bias in the ‘search-and-capture’ process duringmitotic-spindle assembly. Curr. Biol. 15, 828–832.

Wood, K.W., Sakowicz, R., Goldstein, L.S., Cleveland, D.W., 1997. CENP-E is a plus end-directed kinetochore motor required for metaphase chromosome alignment. Cell 91,357–366.

Wordeman, L., 2005. Microtubule-depolymerizing kinesins. Curr. Opin. Cell Biol. 17,82–88.

Wordeman, L., Mitchison, T.J., 1995. Identification and partial characterization of mitoticcentromere-associated kinesin, a kinesin-related protein that associates with centromeresduring mitosis. J. Cell Biol. 128, 95–104.

Wordeman, L., Wagenbach, M., von Dassow, G., 2007. MCAK facilitates chromosomemovement by promoting kinetochore microtubule turnover. J. Cell Biol. 179, 869–879.

Wu, X., Xiang, X., Hammer 3rd., J.A., 2006. Motor proteins at the microtubule plus-end.Trends Cell Biol. 16, 135–143.

Wu, X.S., Tsan, G.L., Hammer 3rd., J.A., 2005. Melanophilin and myosin Va track themicrotubule plus end on EB1. J. Cell Biol. 171, 201–207.

Xiang, X., Han, G., Winkelmann, D.A., Zuo, W., Morris, N.R., 2000. Dynamics ofcytoplasmic dynein in living cells and the effect of a mutation in the dynactin complexactin-related protein Arp1. Curr. Biol. 10, 603–606.

Yajima, J., Edamatsu, M., Watai-Nishii, J., Tokai-Nishizumi, N., Yamamoto, T.,Toyoshima, Y.Y., 2003. The human chromokinesin Kid is a plus end-directedmicrotubule-based motor. EMBO J. 22, 1067–1074.

Yan, X., Habedanck, R., Nigg, E.A., 2006. A complex of two centrosomal proteins,CAP350 and FOP, cooperates with EB1 in microtubule anchoring. Mol. Biol. Cell17, 634–644.

Yang, Z., Tulu, U.S., Wadsworth, P., Rieder, C.L., 2007. Kinetochore dynein is requiredfor chromosome motion and congression independent of the spindle checkpoint. Curr.Biol. 17, 973–980.

Yoshida, T., Ito, A., Izutsu, K., 1985. Association of anti-dynein-1 cross-reactive antigenwith the mitotic spindle of mammalian cells. Cell Struct. Funct. 10, 245–258.

Young, A., Dictenberg, J.B., Purohit, A., Tuft, R., Doxsey, S.J., 2000. Cytoplasmic dynein-mediated assembly of pericentrin and gamma tubulin onto centrosomes. Mol. Biol. Cell11, 2047–2056.

Zanic, M., Stear, J.H., Hyman, A.A., Howard, J., 2009. EB1 recognizes the nucleotide stateof tubulin in the microtubule lattice. PLoS One 4, e7585.

Zanic, M., Widlund, P.O., Hyman, A.A., Howard, J., 2013. Synergy between XMAP215and EB1 increases microtubule growth rates to physiological levels. Nat. Cell Biol. 15,688–693.

Zhai, Y., Borisy, G.G., 1994. Quantitative determination of the proportion of microtubulepolymer present during themitosis-interphase transition. J. Cell Sci. 107 (Pt. 4), 881–890.

Zhai, Y., Kronebusch, P.J., Borisy, G.G., 1995. Kinetochore–microtubule dynamics and themetaphase-anaphase transition. J. Cell Biol. 131, 721–734.

Zhai, Y., Kronebusch, P.J., Simon, P.M., Borisy, G.G., 1996. Microtubule dynamics at theG2/M transition: abrupt breakdown of cytoplasmic microtubules at nuclear envelopebreakdown and implications for spindle morphogenesis. J. Cell Biol. 135, 201–214.

Zhang, J., Ahmad, S., Mao, Y., 2007a. BubR1 and APC/EB1 cooperate to maintainmetaphase chromosome alignment. J. Cell Biol. 178, 773–784.





















































Zhang, J., Li, S., Fischer, R., Xiang, X., 2003. Accumulation of cytoplasmic dynein anddynactin at microtubule plus ends in Aspergillus nidulans is kinesin dependent. Mol.Biol. Cell 14, 1479–1488.

Zhang, L., Shao, H., Huang, Y., Yan, F., Chu, Y., Hou, H., Zhu, M., Fu, C.,Aikhionbare, F., Fang, G., Ding, X., Yao, X., 2010. PLK1 phosphorylates mitoticcentromere-associated kinesin and promotes its depolymerase activity. J. Biol. Chem.286, 3033–3046.

Zhang, L., Shao, H., Zhu, T., Xia, P., Wang, Z., Liu, L., Yan, M., Hill, D.L., Fang, G.,Chen, Z., Wang, D., Yao, X., 2013. DDA3 associates with microtubule plus endsand orchestrates microtubule dynamics and directional cell migration. Sci. Rep. 3, 1681.

Zhang, X., Ems-McClung, S.C., Walczak, C.E., 2008. Aurora A phosphorylates MCAK tocontrol ran-dependent spindle bipolarity. Mol. Biol. Cell 19, 2752–2765.

Zhang, X., Lan, W., Ems-McClung, S.C., Stukenberg, P.T., Walczak, C.E., 2007b. AuroraB phosphorylates multiple sites on mitotic centromere-associated kinesin to spatially andtemporally regulate its function. Mol. Biol. Cell 18, 3264–3276.

Zhu, C., Zhao, J., Bibikova, M., Leverson, J.D., Bossy-Wetzel, E., Fan, J.B.,Abraham, R.T., Jiang, W., 2005. Functional analysis of human microtubule-basedmotor proteins, the kinesins and dyneins, in mitosis/cytokinesis using RNA interference.Mol. Biol. Cell 16, 3187–3199.

Zimniak, T., Fitz, V., Zhou, H., Lampert, F., Opravil, S., Mechtler, K., Stolt-Bergner, P.,Westermann, S., 2012. Spatiotemporal regulation of ipl1/aurora activity by direct cdk1phosphorylation. Curr. Biol. 22, 787–793.

Zimniak, T., Stengl, K., Mechtler, K., Westermann, S., 2009. Phosphoregulation of thebudding yeast EB1 homologue Bim1p by Aurora/Ipl1p. J. Cell Biol. 186, 379–391.

Zumbrunn, J., Kinoshita, K., Hyman, A.A., Nathke, I.S., 2001. Binding of the adenomatouspolyposis coli protein to microtubules increases microtubule stability and is regulated byGSK3 beta phosphorylation. Curr. Biol. 11, 44–49.





























[International Review of Cell and Molecular Biology] Volume 309 || Microtubule Plus-End Tracking...

Documents

Transcript of [International Review of Cell and Molecular Biology] Volume 309 || Microtubule Plus-End Tracking...