UC San Francisco Electronic Theses and Dissertations

UCSF UC San Francisco Electronic Theses and Dissertations

Title Biochemical characterization of microtubule minus-end binding proteins

Permalink https://escholarship.org/uc/item/4wk8m4kh

Author Hendershott, Melissa

Publication Date 2014

Peer reviewed|Thesis/dissertation

eScholarship.org Powered by the California Digital Library University of California Biochemical characterization of microtubule minus-end binding proteins

Melissa Hendershott

DISSERTATION

Submitted in partial satisfaction of the requirements for the degree of

DOCTOR OF PHILOSOPHY

CELL BIOLOGY

in the

GRADUATE DIVISION

of the

Melissa Clayton Hendershott

ii Abstract

The microtubule (MT) cytoskeleton plays an essential role in mitosis, intracellular transport, cell shape, and cell migration. The assembly and disassembly of MTs, which can occur through the addition or loss of subunits at either the plus- or minus-ends of the polymer, is essential for MTs to carry out their biological functions. In Chapter 1, I describe my work on a recently described family of MT minus-end binding proteins called CAMSAP/Patronin/Nezha that act on MT ends to regulate their dynamics. Patronin, the single member of this family in Drosophila, was previously shown to stabilize MT minus-ends against depolymerization in vitro and in vivo. Here, I show that all three mammalian CAMSAP family members also bind specifically to MT minus-ends and protect them against kinesin-13-induced depolymerization. However, these proteins differ in their abilities to suppress tubulin addition at minus-ends and to dissociate from MTs. CAMSAP1 does not interfere with polymerization and tracks along growing minus-ends. CAMSAP2 and CAMSAP 3 decrease the rate of tubulin incorporation and remain bound, thereby creating stretches of decorated MT minus- ends. Using truncation analysis, I find that somewhat different minimal domains of CAMSAP and Patronin are involved in minus-end localization. However, I find that in both cases, a highly conserved C-terminal domain and a more variable central domain cooperate to suppress minus-end dynamics in vitro and that both regions are required to stabilize minus-ends in Drosophila S2 cells.

These results show that members of the CAMSAP/Patronin family all localize to and protect minus- ends but have evolved distinct eﬀects on MT dynamics. In Chapter 2, I describe work characterizing a set of inhibitors of a kinesin-12 family member called Kif15, which has a role in spindle organization during mitosis. Kif15 has been shown in previous work to cooperate with the tetrameric kinesin

Eg5 to facilitate bipolar spindle formation; we were interested in the possible application of these molecules to spindle disruption in hopes that the combination of Eg5 and Kif15 inhibition might be clinically eﬃcacious. I show that these small molecules have activity against Kif15 through an uncompetitive mechanism and that Kif15 inhibition with these molecules aﬀects spindle organization and cell division in vivo.

iii Acknowledgements

I would like to thank Ron for being a wonderful and supportive mentor, and for creating a welcoming and intellectually engaging environment in the lab. I would like to thank Sarah Goodwin, without whom I probably would not have joined the lab- Sarah taught me everything I know about Patronin, and made me feel welcome from day one. Sabine Petry was extremely helpful in giving me a crash course in biochemistry and helping me set up the dynamic assay that was the foundation of my project. Nico was absolutely instrumental in my project being a success; I learned an immense amount about microscopy through Nico and am so grateful to have had access to the amazing knowledge Nico possesses. My bay mates were wonderful science partners and friends. I would like to thank Walter for being an amazingly thoughtful and kind friend and Susana for being patient when I asked her tons of annoying questions, as well as generally being a kind and generous friend.

I would like to thank Erik for putting up with me in Woods Hole, and also for sharing in my lab triumphs and low moments. And for pureeing the brains. I would like to thank Minhaj for helping me in many ways- generously donating reagents and providing lots of advice about protein puriﬁcation and in vitro assays. I want to thank Marvin for all the help with the Kif15 project- having access to all the reagents and getting to dive into experiments made the project really fun.

I would like to thank Nan for keeping everything running in the lab-we really would all fall apart without Nan. I would like to thank Phoebe for all the help over the years; Phoebe does so much for the lab, and with amazing grace and cheerfulness. And overall, I would like to thank everyone in the Vale lab, past and present, for being an amazing group of people. I can’t imagine ever finding another environment where people are so scientifically driven and smart, but also genuinely caring, respectful and kind. I couldn’t have found a better place to do my PhD. I would like to thank my friends and family for reminding me that failures in lab do not equate to failing at life. And finally,

I would like to thank Forrest, who supported me in every way I could have asked.

Portions of this work (largely the content of Chapter 1) have been previously published in M.

C. Hendershott and R. D. Vale. Regulation of microtubule minus-end dynamics by CAMSAPs and

Patronin. Proc Natl Acad Sci U S A. vol. 111, pp. 5860-5, Apr 2014.

iv Contents

Abstract ...... iii

Acknowledgements ...... iv

1 Chapter 1 1

1.1 Introduction ...... 1

1.2 Results ...... 8

1.3 Methods ...... 19

1.4 Conclusion ...... 25

2 Chapter 2 30

2.1 Introduction ...... 30

2.2 Results ...... 32

2.3 Methods ...... 40

2.4 Conclusions ...... 41

3 Chapter 3 43

3.1 Final Thoughts ...... 43

v List of Figures

1.1 CAMSAPs have a conserved function of binding MT minus-ends ...... 9

1.2 CAMSAPs diﬀerentially aﬀect tubulin subunit addition at MT minus-ends ...... 11

1.3 CAMSAP domains involved in minus-end MT binding and regulation of dynamics . 13

1.4 All CAMSAP CKK domains exhibit similar MT binding behavior ...... 14

1.5 CAMSAPs protect MT minus-ends from depolymerization by MCAK ...... 15

1.6 Domains required for Drosophila Patronin minus-end binding, suppression of growth

in vitro and MT stability in S2 cells ...... 17

1.7 A sub-domain of the Patronin CC speciﬁcally binds MT minus-ends ...... 18

1.8 CAMSAPs and Patronin cannot bind to yeast microtubules ...... 20

2.1 Schematic of PK/LDH ATPase Reaction ...... 33

2.2 Suppression of ATPase activity of the Kif15 motor domain by addition of Kif15 inhibitors 34

2.3 Measurement of Kif15 ATPase in the presence of varying MT or ATP concentrations 35

2.4 Inhibition of Kif15 leads to spindle collapse ...... 37

2.5 Ispinesib-resistant cell lines exhibit diﬀerent behavior in the presence of a Kif15 inhibitor 39

vi Chapter 1

Chapter 1

1.1 Introduction

A cell’s purpose is to divide and replicate itself; all life comes from life (leaving aside the issue of the first cell). However, this simple goal, to divide and produce more cells, belies the complex and intricate activities required to divide hundreds or thousands of times without error. Any individual cell is like a tiny city unto itself, with thousands of protein ”parts” executing hundreds of jobs within the cell. Cells must produce energy, create new cellular material and replicate their genetic information; orchestrating all these tasks is extremely complex. Over the time scales of evolution, cells have grown from simple, single-celled bacteria, which are small and can grow and divide in the absence of other cells, to eukaryotic cells, which are hundreds of times larger than bacteria, and can form assemblies that create tissues, organs and ultimately entire organisms. Increases in size and cellular complexity, however, led to the requirement for structural elements. Like the steel beams of a building, large cells require structural support to maintain their shape and structural integrity; additionally, the large size of eukaryotic cells means that simple diffusion is too slow to efficiently transport materials across the cell. And the additional genomic complexity means that chromosomes must be allocated precisely, to ensure that genetic material is equally distributed; large chromosomes must be actively pulled into the newly formed cells, rather than relying on chance to obtain genetic material, as bacteria do for plasmid DNA. Among the many molecules that cells use to carry out these tasks, one of the central players are molecular polymers known as microtubules.

Microtubules (MTs) are composed of dimers of α- and β-tubulin that assemble head-to-tail, an arrangement that creates a polar protoﬁlament. Protoﬁlaments assemble laterally to form the

1 canonical 13-protoﬁlament MT structure, with β-tubulin exposed at fast-growing plus-ends, and α- tubulin exposed at slow-growing minus-ends (1). MTs derive the energy for their growth from GTP; both α- and β-tubulin bind GTP. However, because the GTP bound to α-tubulin is tightly wedged at the α-β interface, only the GTP bound to β-tubulin can be hydrolyzed. As a MT polymerizes, the β-tubulin-bound GTP is hydrolyzed, creating a MT lattice populated by GDP-bound tubulin, with a growing tip populated by un-hydrolyzed GTP-bound tubulin (2). This composition leads to MTs exhibiting an intriguing property termed ”dynamic instability,” whereby the polymer can abruptly switch between episodes of net growth and net shrinkage (3). Dynamic instability takes place because the hydrolysis of GTP to GDP in the polymer lattice slightly changes the curvature of the tubulin dimers(4). Because GDP-bound tubulin has a slightly greater curvature than GTP- bound tubulin, once the GTP has been hydrolyzed, the GDP-bound tubulin dimers strain against the lattice (5). This strain leads to stochastic destabilization of the entire microtubule, triggering a depolymerization event, or a catastrophe. These catastrophes will either fully depolymerize the MT, or can terminate in ”rescue” events, in which a microtubule can regrow. The rates of growth and shrinkage as well as the frequency of transitions between these two states are regulated by numerous microtubule-associated proteins, many of which bind to the ends of the polymer (6; 7).

Methods for studying MT in vitro have dramatically improved our understanding of MT behavior and the functions of microtubule associated proteins (MAPs). Tubulin can be puriﬁed from a variety of sources and given an energy source (GTP), can be polymerized in vitro. These MTs can be stabilized by the MT binding drug taxol, or by polymerization in the presence of a non-hydrolyzable

GTP analogue (8). MTs have been dynamically observed for decades; early strategies used DIC imaging of MT growth from structures such as isolated axonemes from sea urchin sperm (9); more modern methods use total internal reﬂection microscopy (TIRF) and ﬂuorescently labeled tubulin.

These in vitro systems and labeling strategies have allowed detailed observations of MT dynamics and interactions with binding proteins. For example, in vitro work has enabled precise characterization of the network of plus-end tracking proteins regulated by EB1 (10; 11; 12; 13). This has allowed detailed mechanistic studies of microtubule binding and tip-tracking proteins, which frequently help elucidate observations made in vivo. For example, some proteins are known to promote MT growth in vivo; these proteins can be directly measured as increasing the MT growth rate in vitro. Other microtubule associated proteins, such as MAPs, regulate microtubule stability; many of these proteins are up- regulated in neurons, presumably to help stabilize the long microtubule bundles that animate the long processes (axons and dendrites) formed by neurons. These proteins have been shown to stabilize

MT in vitro as well; these experiments have also revealed subtle diﬀerences in their functions that

2 are not readily apparent from in vivo study.

Significant progress has been made in understanding the behavior of MT plus-ends; in vivo, MT plus ends are highly dynamic, constantly shrinking and growing at the periphery of the cell. These dynamics can be visualized in vivo by labeling either the tubulin itself with a fluorescent protein, or by visualizing the tips of the MT themselves using proteins that localize specifically at growing

MT ends. The dynamics of MT plus-ends in vivo are regulated by a well-characterized network of plus-end tracking proteins (+TIPs) (14). End-binding (EB) proteins are thought to recognize the plus tip of MTs by speciﬁcally binding to the GTP-cap and can aﬀect the dynamics of plus- ends both by intrinsically altering the structure of the MT end (15; 16; 17), as well as recruiting other interacting proteins (18). Among the many proteins interacting with EBs, TOG-domain containing proteins, such as XMAP215, promote MT growth and have been suggested to act as MT

”polymerases” (19; 20). Conversely, kinesin-13s (e.g. MCAK from hamster) increase instability of

MT ends, leading to increased catastrophe frequency (21; 22). Thus regulation of these and other

+TIPs can dramatically aﬀect the stability and turnover of the microtubule network (23; 24).

In comparison to the well-characterized +TIPS, much less is known about regulation of MT minus-ends. In many cells, minus-ends in vivo are anchored at the centrosome by the γ-tubulin ring complex (γ-TuRC). The γ-TuRC is the only complex of proteins known to bind directly at the exposed α-tubulin end of a microtubule (25). Each protoﬁlament of the microtubule is capped by a

γ-tubulin molecule; the γ-tubulin ring complex is made up of assemblies of GCP proteins that form the canonical cone shape that is characteristic of the γ-TuRC (26; 27). In cells, γ-TuRC cone shapes can be visualized by EM on the ends of microtubules, although not all microtubules are capped (28).

Although γ-TuRC is thought to be the only nucleator of MTs, other proteins are involved in localizing

γ-tubulin and regulating its function. Ninein is often associated with MT minus-ends and likely plays a role in localizing γ-TuRCs to sites throughout the cell. A protein complex known as augmin was shown in our lab to localize γ-tubulin throughout the mitotic spindle, promoting non-centrosomal

MT nucleation during cell division (29; 30). Many membrane-associated proteins localize γ-tubulin to membrane-bound structures throughout the cell; for example, Ori-McKenney recently showed that the Drosophila homologue of AKAP450 (31) is involved in localizing γ-tubulin to nucleation sites located at Golgi outposts in branched neuronal architectures (32). However, while microtubules are nucleated by γ-tubulin, it remains an unresolved question of whether all microtubules are capped at their minus-ends at all times. Indeed, microtubules frequently undergo rearrangements during the cell cycle that require release of microtubules from their site of nucleation; for example, during mitosis, large numbers of microtubules are released from the centrosome to assemble the mitotic

3 spindle. However, whether these microtubules are released with their minus-ends capped remains unclear. Additionally, interphase cells also are observed to release microtubules from the centrosome

(33); the state of the minus-ends of these microtubules is also not known.

One reason that MT minus-ends are somewhat difficult to study is that, in contrast to plus-ends, where many end-binding proteins have been identified that can be used to mark their presence, very few minus-end binding proteins have been identified. γ-tubulin was until recently the only known protein to directly bind to MT minus-ends. Thus, difficulty in labeling and visualizing minus-ends in vivo has complicated detailed study of their behavior. However, when minus-ends that are not connected to the centrosome have been observed, either due to breakage or transport away from the centrosome, they appear to be highly stable and neither grow nor shrink, in contrast to the behavior of minus-ends composed of pure tubulin. Newly created minus-ends formed by breakage or severing with a laser tend to neither grow nor shrink, while the newly created plus-ends tend to rapidly depolymerize (34; 35; 33; 36; 37). It is unclear how this stability of minus-ends is mediated.

While it is possible that newly formed minus-ends become capped by γ-tubulin immediately upon their generation, this behavior has not actually been visualized.

While some cells organize their MTs around a MT organizing center (MTOC), many cells, such as epithelial cells, Drosophila S2 cells, skeletal muscle cells, neurons and plant cells have non-centrosomal MT arrays. Many epithelial cells have MT arrays that are arranged along the apical-basal axis of the cell, where minus ends are oriented toward the apical side of the cell (38).

Drosophila cells tend to have a less organized microtubule array, in which MT emanate throughout the cytoplasm, rather than being gathered around a centrosome. In skeletal muscle, which is made of large, multinucleated cells known as myotubes, MTs are arrayed in long, linear arrays that are not central to a MTOC; in fact, as myotubes mature, centrosomes are lost entirely (39; 40). Neurons have one of the most unique microtubule arrangements, as MTs are essential for animating the long processes that characterize neurons. MTs in these processes are also essential to provide tracks for transport of materials throughout the neuron (41; 42). Microtubules have stereotypic arrangements in neurons; axons have MT arrays with plus-ends oriented away from the cell body, while dendrites, which tend to be shorter than axons, have MTs of mixed polarity (43; 44; 45). These microtubules are not anchored to the centrosome and are most likely localized by release and transport from the cell body. It is unclear how the stability of these microtubule populations are mediated, or how they are regulated. Similarly, many of the MTs in the spindle also are not directly connected to centrosomes (46), although as mentioned above, it is known that microtubule nucleation in the spindle is mediated by the augmin complex (47). However, whether these MT remain associated

4 with γ-tubulin throughout cell division is unknown. Thus, although free minus ends are present in some abundance in many biological systems, very little is known about their regulation, or proteins that bind to them. There are many open questions related to these minus ends; for example, how is stability of the ends maintained? Are there diﬀerential regulatory mechanisms for diﬀerent types of minus-ends (ie. minus-ends that are free in the spindle, versus minus-ends that are located in neuronal process)? Where are minus ends localized in cells with non-centrosomal microtubule arrays?

Because of the paucity of markers for minus-ends, their visualization and study remains a challenge in cell biology.

In addition to proteins that bind and regulate MTs at their ends, there are many proteins that are involved in binding MTs along their lengths. These proteins include bundling and cross-linking proteins, such as PRC-1, which bundles antiparallel microtubules in mitosis (48; 49), microtubule- associated proteins (MAPs), such as tau, which binds MTs along their length and helps stabilize them (50; 51) and motor proteins, which bind to and use energy from ATP hydrolysis to actually walk along microtubules (52; 53). Many microtubule-based motors have been identiﬁed. The two major families are kinesin and dynenin; kinesins are canonically plus-end directed motors, while dynein moves the opposite direction and walks towards MT minus-ends (52). These two motors, aside from moving in opposite directions, have very diﬀerent structures and regulatory mechanisms.

Kinesins are most often dimers; they possess two motor domains, which are akin to ”feet” and use a long stalk, or linker, to dimerize and coordinate the activity of the two motor heads (54; 55; 56).

There are approximately 40-50 known mammalian kinesins which perform diverse functions (52).

Conventional kinesin, or kinesin-1, is largely neuronal and is highly processive; kinesin-1 is important for cargo transport (52). Other kinesins, such as the tetrameric kinesin Eg5, slide and bundle anti-parallel microtubules, a function that is largely important during mitosis (57; 58). Still other kinesins are involved in chromosome dynamics during mitosis. A family of kinesins known as kinesin-

13s, or MCAK, are involved in microtubule depolymerization, most likely by a mechanism that involves bending tubulin dimers in a way that creates strain in the polymer, leading to catastrophic strain on the MT lattice (59; 60; 22). These diverse functions are achieved through the enormous diversity of kinesin family members. In contrast, dynein, which also plays a huge number of roles in cells, is diﬀerentially regulated largely through the activities of regulatory subunits (52). Dynein exists as a large, megadalton multi-protein assembly. Although the heavy chain of yeast dynein is known to walk processively without additional components, no such activity has been observed for mammalian dynein. Instead, many regulatory proteins bind to the dynein heavy chain and regulate its processivity and attachments to cargo. For example, the light-intermediate chain of dynein

5 (LIC) is thought to mediate binding to cargo, while new work by Rick McKenney in our lab has also demonstrated the important of another complex, the dynactin complex (another megadalton protein assembly), for processive dynein motility (61). Both dynein and kinesin are essential for cargo transport throughout the cell, and knockdown of many kinesin family members, or of dynein regulators, can have deleterious eﬀects on the cell.

Study of the complex regulation of microtubules has revealed many fundamental aspects of cell biology; one context in which MTs have been extensively studied is cell division. Previous work in our lab focusing on spindle assembly identiﬁed the Drosophila protein Patronin (from the Latin patronus) in a whole genome RNAi screen for mitotic spindle formation (62) (originally named ssp4). In this screen, each gene in the Drosophila genome was systematically knocked down by

RNAi. Automated imaging was used to image hundreds of spindles and automatically characterize their properties (these classiﬁcations were manually conﬁrmed as well). One phenotype that was observed was a short spindle phenotype; one of the short spindle phenotype genes, ssp4, was studied further by Sarah Goodwin. Knock-down of ssp4 in interphase cells showed treadmilling MT traveling through the cytoplasm. These small, ”severed” pieces of MT led to the original hypothesis that ssp4’s function was to regulate MT severing by severing proteins such as katanin. However,

Goodwin’s work showed that this protein binds to and protects the microtubule minus-end against depolymerization by kinesin-13 both in vitro and in vivo (63). Patronin is also important for maintaining MT structures in Drosophila cells. In addition to Patronin RNAi causing small fragments of MTs to appear in the cytoplasm, there is a global decrease in MT density in the cells, likely due to increased depolymerization of the MT cytoskeleton by kinesin-13. This result suggests that

Patronin is important for the regulation and maintenance of MT organization in Drosophila cells.

Purified Patronin was incubated with stabilized MT (either by taxol or GMPCPP stabilization) and was shown to bind specifically and strongly at MT minus-ends. This minus-end binding activity additionally allows the protection of minus-ends from depolymerization; Goodwin showed that in vitro, MTs incubated with Patronin and kinesin-13 are depolymerized only from their unprotected plus-ends. Minus-ends, capped by Patronin, are protected from the activity of kinesin-13. Further work in flies has shown that Patronin antagonizes kinesin-13 during mitosis and that regulation of

Patronin’s activity facilitates spindle elongation in anaphase B (64).

In mammals, the vertebrate homologues of Patronin, named CAMSAP/Nezha, have also been shown to associate with MT minus ends. Bioinformatic analyses suggest that this protein family

ﬁrst evolved in metazoans; invertebrates possess a single gene (e.g. Patronin in ﬂies, PTRN-1 in

C. elegans), while vertebrates possess three CAMSAP genes (CAMSAP1, CAMSAP2 and CAM-

6 SAP3/Nezha) (1.1A) (65; 66). Takeichi and co-workers (65; 67) have shown that one of the mammalian homologues, Nezha, anchors minus-ends of MTs in cell-cell adherens junctions. This work also demonstrated the speciﬁc in vitro binding of Nezha to MT minus ends. The protein family was also identiﬁed through a spectrin binding activity and named CAMSAP (CAlModulin-regulated,

Spectrin-Associated Protein) (66; 68). This work also identiﬁed the MT binding activity of the protein family (although not the minus-end speciﬁcity) and localized the MT binding region to the

C-terminal domain of the protein, which they termed the CKK domain. Recent work has demonstrated that CAMSAP2 and 3 bind MT minus-ends of mammalian cells and that depletion of these proteins by RNAi causes a reduction in microtubule numbers and a change in the MT organization in epithelial cells (69; 70). These studies also show that both CAMSAP2 and CAMSAP3 can be found at the minus-ends of the same MTs, and that they form somewhat elongated caps, rather than small puncta. Additionally, recent studies also have shown that the C. elegans homolog of Pa- tronin/CAMSAP, PTRN-1, stabilizes MTs in neurons and promotes neurite and synapse stability

(71; 72).

While these studies collectively demonstrate that CAMSAPs and Patronin stabilize MT minus- ends in vivo, many questions remain as to how these proteins aﬀect minus-end dynamics, whether all vertebrate CAMSAPs possess minus-end regulatory activity, and how minus-end recognition is achieved. For my thesis work, I wanted to understand this family of proteins in more mechanistic detail than had previously been appreciated. Since work in our lab focused on the Drosophila homologue, I wanted to compare the activity of Drosophila Patronin with the mammalian homologues.

Additionally, in Drosophila, only one CAMSAP homologue is present; therefore, I wanted to understand how the activity of Drosophila Patronin might differ from the activities of the CAMSAPs in mammals. Finally, I wanted to gain a deeper understanding of differences between the mammalian homologues; what distinct activities might the three CAMSAP homologues possess to differentiate themselves from each other? Through this work, I showed that all vertebrate CAMSAP family members bind to MT minus-ends in vitro. However, I found two separable activities for full length or truncated subdomains of CAMSAPs and Patronin at minus-ends: an ability to track along growing

MT minus-ends and an ability to suppress tubulin incorporation at minus-ends. The latter activity is correlated with MT minus-ends stabilization in Drosophila S2 cells. These results indicate that minus-end binding and regulation is universal for all CAMSAP members but that this activity is tunable in ways that might be exploited by cells to regulate MT organization. My hope is that these studies will provide insight into the biochemical functions of these proteins that can help guide future studies that will help elucidate the distinct roles that these proteins play, as well as guide

7 studies into whether these proteins create or regulate distinct sub-populations of MT minus-ends.

1.2 Results

All Vertebrate CAMSAP family members bind microtubule minus-ends in vitro

Previous studies have shown that CAMSAP2 and CAMSAP3 bind to MT minus-ends in vivo (65; 66).

To determine whether these proteins alone are suﬃcient for this activity and whether this property is also true for CAMSAP1 (Figure 1.1A), I puriﬁed full-length mammalian CAMSAP1 (human),

CAMSAP2 (human) and CAMSAP3 (mouse (Nezha), chosen because of the prior in vivo work performed on this protein (65)) with an N-terminal GFP and performed in vitro binding experiments.

The GFP-CAMSAPs (10-12 nM) were incubated with ﬂuorescently-labeled, GMPCPP-stabilized

MTs, which were then bound to and moved along a glass surface coated with either kinesin or dynein motors. In the kinesin gliding assay, the minus-end of the MT is leading in the direction of motion, while the direction of gliding is the opposite with dynein. These gliding assays allowed us to distinguish the polarity of end binding and also determine whether the proteins were stably bound to the end as the microtubule translocated across the surface. For all three GFP-tagged CAMSAPs, a bright fluorescent spot was observed at the minus-ends of the majority of gliding MTs, and, in rarer cases (<10%), was observed at both ends (Figure 1.1B,C). Some MTs also had GFP spots along their length, but these fluorescent spots tended not to be stably bound as the MT moved along the surface and were also more sensitive to salt-induced dissociation. The minus-end-bound GFP punctae were ∼2-4-fold brighter than GFP-CAMSAP fluorescent spots on the glass (Figure 1.4A).

This result suggests that one or a few CAMSAP molecules are suﬃcient to bind at the minus-end of a stabilized MT. These data suggest that that binding of CAMSAPs to minus-ends diﬀers from

γ-tubulin, which, as part of a γ-tubulin ring complex, caps each of the 13 protofilaments on the minus-end (73). In summary, I find that all vertebrate CAMSAPs can specifically recognize the minus-ends of non-dynamic MTs.

CAMSAP proteins diﬀerentially regulate dynamic MT minus-ends

Previous in vitro work with Patronin and CAMSAP3, as well as the work described above, were performed with stabilized MT (63; 65; 66). However, in vivo, MTs are dynamic polymers, and therefore, I wanted to determine if CAMSAP proteins aﬀected the incorporation of tubulin into growing MTs in vitro. For this experiment, a GMPCPP-stabilized ”seed” MT with incorporated

Alexa647-tubulin was adhered onto the cover glass surface. Then tubulin monomers were added,

8 A. B. MT GFP-CAMSAP Kinesin Motor Protein

Hs CAMSAP2 CAMSAP2

0 sec

CAMSAP3

20 sec

CAMSAP1 35 sec C. Patronin, Fruitfly 100 ptrn-1, C. elegans 80 LEGEND Trailing MT End Ray-finned Fish Coelacanth 60 Leading MT End Reptiles and Birds Drosophila 40 Binding at Both Mammals C. elegans MT Ends 20 Unbound MT Xenopus Marsupials Percent of MT Bound Percent of MT x10 branch length CAMSAP1 CAMSAP2 CAMSAP2 CAMSAP3 CAMSAP3 Kinesin Dynein Kinesin Dynein Kinesin

Figure 1.1: A. Most vertebrate species possess 3 CAMSAP/Patronin homologues, while Drosophila and C. elegans each have only one. B. Motor gliding assay for determining the end of the MT bound by CAMSAPs; in a kinesin gliding assay, the minus-end of the GMPCPP-stabilized MT is the leading; polarity is reversed in a dynein gliding assay. Kinesin assay for GFP-CAMSAP2 is shown. Arrows show GFP-CAMSAP2 on MT minus-ends. Bar, 5 µm. Movement of a MT-CAMSAP2 complex over time (rectangle inset, bar, 2 µm). C. Quantiﬁcation of number of MT with GFP- CAMSAPs on their ends. Results compiled from two independent experiments (>100 MT scored per assay). a subset of which were labeled with Alexa561-tubulin to discern new subunit addition to the MT ends; GFP-CAMSAP was added into the mixture with the tubulin monomers (Figure 1.2A). Triple color images, to visualize MT seeds, new tubulin addition, and GFP-CAMSAPs, were acquired every 2 sec for 3-4 min, by TIRF microscopy; this acquisition time allowed accurate measurements of plus- and minus-end growth of control MT with minimal photobleaching. In the assay buﬀer

(80 mM PIPES, 2 mM MgCl2, 1 mM EGTA, 60 mM KCl) and the concentrations of tubulin (20-

25 µm) used in this assay, the majority (>80%) of MTs grew from both ends (Figure 1.2B,D,

E); catastrophes (rapid depolymerization) were occasionally observed, but rare. Plus-ends could be diﬀerentiated from minus-ends by their two-fold faster growth rates (Figure 1.2G). Interestingly, when GFP-CAMSAP1 was added, ﬂuorescent molecules were observed tracking at the tip of growing

MT minus-ends (Figure 1.2D) and did not aﬀect the rate of tubulin incorporation at the minus- end (Figure 1.2G); plus-ends were not aﬀected (Figure 1.2G). In contrast, upon the addition of

GFP-CAMSAP2 and GFP-CAMSAP3, most MT minus-ends (∼70% and 80% respectively) did not

9 display growth at the limit of detection (∼0.5 µm growth) over the ∼4 min time of image acquisition

(Figure 1.2C,D,E). However, the ﬂuorescence intensity of the GFP-CAMSAP at the minus-ends in these dynamic assays with monomeric tubulin (Figure 1.2C) was brighter than that observed on minus-ends of GMPCPP-stabilized MTs (Figure 1.1B), suggesting that perhaps some minus-end tubulin addition occurred and deposited CAMSAPs on the MT. Neither CAMSAP2 nor CAMSAP3 aﬀected the rate of plus-end growth (Fig. 1.2G).

The above experiments showed that CAMSAP2 and CAMSAP3 dramatically suppress microtubule minus-end growth, while CAMSAP1 has little eﬀect despite its presence at the MT minus-end.

To determine the extent of growth suppression more accurately for CAMSAP2 and CAMSAP3, I performed longer term imaging experiments (Figure 1.2F). Over the course of 20 min, discernible tubulin incorporation at the minus-end could be observed for the majority of MTs that had a

GFP-CAMPSAP2 or GFP-CAMPSAP3 at the end. GFP-CAMSAP2 coated the entire stretch of new tubulin incorporation, suggesting that, once bound to the minus-end, it can remain associated with the MT lattice after new tubulins have been added on (Figure 1.2F). For GFP-CAMSAP3 the decoration appeared more discontinuous, suggesting that occasionally tubulin could grow past a

GFP-CAMSAP3 cap and then become recapped with GFP-CAMSAP3 (Figure 1.2F). These longer time-lapse experiments enabled measurements of minus-end polymerization rates and the results show that CAMSAP2 and CAMSAP3 reduce minus-end growth by 86% and 93%, respectively, compared with control MTs (Figure 1.2G).

In summary, CAMSAP2 and CAMSAP3 both strongly suppress minus-end growth, with CAM-

SAP3 being somewhat more potent, but do not affect plus-end growth. In contrast, CAMSAP1 tracks along the growing tips of minus-ends without significantly affecting the polymerization rate.

These diﬀerences in the dynamic assay suggest that the three CAMSAP proteins, despite their shared ability to bind selectively to MT minus-ends, regulate minus-end dynamics in distinct ways.

Cooperation between CAMSAP domains enhances minus-end localization and suppression of MT minus-end dynamics

The three vertebrate CAMSAP proteins and the single Patronin protein from invertebrates all share a similar overall domain structure of an N-terminal calponin-homology (CH) domain (which has actin binding activity in some proteins (74)), a central domain of three coiled-coils with a predicted poorly structured region between coils 2 and 3 (herein referred to as the ”CC” domain), and a C- terminal conserved globular domain known as the CKK domain (66). Previous studies have shown that Patronin’s CH domain is diﬀusely localized when expressed in cells and does not target to any

10 A. GFP-CAMSAP MT Seed Dynamic MT Anti-biotin antibody B. C.

D. Control CAMSAP2 E. 100

80 Time

60 Distance CAMSAP1 CAMSAP3 40

20 Percent Dynamic Minus Ends

Control CAMSAP1 CAMSAP2 CAMSAP3 F. CAMSAP2 CAMSAP3 G. 1.8 Plus Ends Minus Ends 1.4 0 min. 1.0

0.6 MT Growth Rate ( μ m/min) MT

20 min. 0.2

Control CAMSAP1 CAMSAP2 CAMSAP3

Figure 1.2: A. GMPCPP seed MTs (with Alexa-647 tubulin and biotin tubulin) were attached to an anti-biotin antibody-coated coverslip (see Methods). Unpolymerized tubulin (23 µm) was added to the chamber with Alexa561-labeled tubulin (2 µm) and allowed to polymerize at room temperature. B. Control field of polymerizing MT. GMCPP seeds shown in blue and newly polymerized tubulin shown in red. Image was background corrected in Fiji. Image shown ∼3-4 min after reaction set-up. Bar, 5 µm. C.Dynamic assay performed with addition of 10-12 nM GFP-CAMSAP3 (green spot at MT minus-ends) shows little growth after 3-4 min of polymerization. Bar, 5 µm. D. Representative kymographs showing polymerization of control MT and MTs bound by GFP-CAMSAPs. Photo- bleaching of CAMSAP2 was partially corrected using Fiji plugin. Minus-ends oriented to left. Bar: 1 µm horizontal and 20 sec vertical. E. Quantification of MT minus-ends showing visible growth in <4 min rapid acquisition window (control: N=4, CAMSAP1: N=2, CAMSAP2: N=3, CAMSAP3: N=4, where N is number of experimental days, n=59-215 MTs total). Minus-ends were scored as dynamic if they underwent growth of >0.5 µm over the course of imaging (200 sec). Mean and SEM are shown for averages of each experimental day. Only MTs with GFP at the minus-end were scored. F. Long-term (20 min) imaging of MT seeds (blue), newly polymerized tubulin (red) and GFP-CAMSAP2 and 3 (green). Minus-ends oriented to left. Scale bar: 1 µm. G. Quantification of MT plus- and minus-end growth rates; short-term imaging (<4 min) was measured for plus- and minus-end growth rates; long-term imaging was used for CAMSAP2 and CAMSAP3 minus-ends. Number of independent experiments on separate days: plus ends: Control: N=6; CAMSAP1: N=2; CAMSAP2: N=5; CAMSAP3: N=5. Minus-ends: Control: N=6; CAMSAP1: N=2; CAMSAP2: N=2; CAMSAP3: N=2, for 19-77 MT per day. Mean and SEM of independent days of measurement shown.

11 speciﬁc structure (63). The CKK domain has been shown to bind to MT both in vivo (63) and in vitro (66) but minus-end-localization was not carefully examined in these studies.

To understand the roles of each domain in minus-end binding, I expressed and puriﬁed three truncations of CAMSAP3: the CKK domain, the coiled-coil (CC) domain and the CC+CKK domains

(Figure1.3). I ﬁrst incubated these GFP-tagged, puriﬁed proteins with GMPCPP-stabilized MTs and performed kinesin gliding assays, as described for the full-length CAMSAPs. I found that the

CKK domain of CAMSAP3 (CKK3) bound to MTs along their entire lengths, although MTs showed a higher concentration of GFP-CKK3 at their minus-ends (Figure 1.3B). However, this association was very sensitive to salt; virtually all of the GFP-CKK dissociated completely from the MTs when

60 mM KCl was added to the buﬀer (Figure 1.3B). Similar results were also obtained for GFP-CKK domains from CAMSAP1 and CAMSAP2 1.4B. In contrast, the CC domain from CAMSAP3 did not bind to MTs (Figure 1.3B). However, the CC+CKK domain from CAMSAP3 (Figure 1.3B) and

CAMSAP2 (1.4C) bound to MT minus-ends at 60 mM KCl and few molecules were found along the length of the microtubule. These data indicate that the CKK domain is responsible for MT localization of mammalian CAMSAP proteins, but that the CC domain enhances the strength and speciﬁcity of minus-end binding at higher, more physiological ionic strengths.

I next tested the CC+CKK domains from CAMSAP2 and 3, which exhibited specific minus-end localization in our standard assay buffer, for their effects on MT dynamics in the short time-lapse assay (Figure 1.3C, D). The CC+CKK domains from both CAMSAP2 and 3 suppressed MT minus- end growth (Figure 1.3C,D). Thus, the CH domain is not needed for either minus-end localization or suppression of growth.

CAMSAP proteins protect stabilized MT from depolymerization by kinesin-13

Previous work has shown that Patronin protects MT minus-ends from depolymerization by MCAK, a kinesin-13 protein that causes subunit loss and destabilizes microtubule ends (63). I next in- vestigated whether the three mammalian CAMSAPs are able to protect microtubule minus-ends against kinesin-13-induced depolymerization. In this assay, GMPCPP-stabilized, ﬂuorescent MTs with incorporated biotinylated tubulin were adhered to streptavidin-coated coverslips; kinesin-13

(MCAK; 6 nM) was added, and the depolymerization of MT ends was followed by TIRF microscopy

(Figure 1.5A). For control MTs without CAMSAPs, 93% of the MTs were depolymerized from both ends, and 7% resisted depolymerization at both ends; depolymerization from just one end was never observed (Figure 1.5B,C). In contrast, the majority of MT depolymerized from just one end in the presence of CAMSAP3 (74%) CAMSAP2 (63%) and CAMSAP1 (54%) (Figure 1.5B,C). These

12 A. CH CC1 CC2 CC3 CKK

B. CAMSAP3 Domain Analysis GFP GFP GFP GFP

CKK- 0 mM KCl CKK-60 mM KCl CC-0 mM KCl CC+CKK-60 mM KCl GFP 647-MT Merge GFP 0.4 C. D. CAMSAP2 CC+CKK CAMSAP3 CC+CKK 0.3 0.2 ( μ m/min) Time 0.1

Distance Control CAMSAP2 CAMSAP3 Minus End Grwoth Rate CC+CKK CC+CKK

Figure 1.3: A. CAMSAP domain structure; predicted CC regions shown as cylinders. B. The CKK domain of CAMSAP3 (20 nM) binds along the length of microtubules with an accumulation at MT minus-ends, as shown by TIRF microscopy. CKK binding is abolished in the presence of 60 mM KCl. CAMSAP3 CC does not exhibit MT binding, but the CC+CKK domain shows speciﬁc MT minus-end binding in the presence of 60 mM KCl. Bar, 2 µm. C. Representative kymographs of MTs capped by the CAMSAP2 and CAMSAP3 CC+CKK domains. For both CAMSAPs, the CC+CKK domains recapitulate minus-end capping to the same extent as the full-length proteins. Scale bar: 1 µm horizontal, 20 sec vertical. D. Quantiﬁcation of minus-end growth rates determined by long-term (20 min) imaging of MT dynamics (control growth rate from Fig. 2, for CAMSAP2 and 3 CC+CKK, N=2, 14-32 MTs per day). Mean and SEM of each experimental day are shown. Only MTs with GFP initially at the minus-end were analyzed.

13 ncnrs otemmainCKdmis h a K oande o idM,ee n0 in even MT, bind not all does almost domain KCl, CKK mM 2 Pat Left). 60 the bar: arrow, (Scale domains, of (white CKK (Right). minus-ends presence MT KCl mammalian the to mM the binds In 2 to domain MT CC+CKK contrast bar: the CAMSAP2 Scale columns). In for the preference columns). third 3), fourth slight and (Fig. and a CC+CKK (second (ﬁrst with for lost concentrations, KCl plotted is nM binding and mM 10-12 microtubule CAMSAP2 surface 0 and at glass CAMSAP1 length in the the MT 3), on minus-end, to (Figure bound CAMSAP3 bind as molecules to domains compared Similar nearby was CKK B. ﬁve minus-end of member. microtubule family intensity a CAMSAP average at each the bound to GFP-CAMSAP of ratio intensity a The A. 1.4: Figure Count 15 25 35 5 A. 1 C. B. Merge 647-MT GFP Merge 647-MT GFP 2 CAMSAP2 CC+CKK Intensity Ratio CAMSAP1 CKK- 0mMKCl 60 mMKCl CAMSAP1 3 5 4 µm.) 6 Count CKK-60 mMKCl 10 2 30 40 0 CAMSAP1 0 mMKCl Pat CKK 6 5 4 3 2 1 Intensity Ratio CAMSAP2 14 CKK- 0mMKCl CAMSAP2

Count 1 20 30 40 50 60 0 .C sfrteCAMSAP3 the for As C. µm. CKK-60 mMKCl 6 5 4 3 2 1 CAMSAP2 Intensity Ratio CAMSAP3 Anti-biotin A. MCAK MT CAMSAP C. Antibody Depolymerization at One End 100 Depolymerization from Both Ends No Depolymerization 80

B. Control CAMSAP3 60 40 Percent of MTs 20 Time

Distance Control CAMSAP1 CAMSAP2 CAMSAP3

Figure 1.5: A. Schematic of assay. B. Representative kymographs of control MT depolymerizing from both ends (left), and MT protected in the presence of CAMSAP3 (right). In the low-salt conditions needed for MCAK depolymerization, CAMSAPs tend to bind along MT length; hence, only the MT imaging channel is shown. Horizontal bar 1 µm; vertical bar 60 sec. C. Quantiﬁcation of microtubules exhibiting kinesin-13-induced end-wise depolymerization behaviors. Number of independent experiments on separate days: control, N=3; CAMSAP1: N=2; CAMSAP2, N=2; CAMSAP3, N=2, for 17-142 MTs per day. Mean and SEM are shown for the diﬀerent of experimental days. results suggest that the function of protecting MTs from kinesin-13 mediated depolymerization is conserved throughout all members of the family.

Suppression of microtubule minus-end dynamics is essential for the cell biological activity of the Drosophila CAMSAP homologue, Patronin

Drosophila has a single CAMSAP homologue, Patronin, which is most closely aligned with CAM-

SAP2 and CAMSAP3 (Figure 1.1A). The presence of a single gene, coupled with a high eﬃciency of

RNAi-mediated protein knockdown, makes Drosophila S2 cells a good system for studying the roles of these minus-end regulatory proteins. Prior RNAi knockdown of Patronin was shown to decrease the number of interphase MTs and cause the appearance of short, treadmilling MT fragments (63).

The latter phenotype is easily scored by time-lapse microscopy and provides a clear in vivo assay for Patronin function.

I first conducted a functional analysis of purified Patronin domains (Figure 1.6A), similar to that described for the mammalian CAMSAP3. As was true for CAMSAP3, I found that the CC+CKK domain bound tightly and specifically to MT minus-ends (Figure 1.6B). Interestingly, and in contrast to what I found for CAMSAP3, the Patronin GFP-CKK domain (10 nM) did not exhibit observable binding to MT in a TIRF assay, even without addition of KCl (Figure 1.4A). Also in contrast to the mammalian CAMSAPs, GFP-CC domain alone bound highly specifically to MT minus-ends

(Figure 1.6B). Through truncation analysis, I identiﬁed an ∼70 kDa fragment in the C-terminal

15 half of the CC domain (a.a. 868-1457) that is necessary and suﬃcient for binding selectively to

MT minus-ends (Figure 1.7A,B, construct 4). This region eliminates the ﬁrst two predicted coiled coils and contains a predicted unstructured region plus the C-terminal coiled coil; the C-terminal coiled-coil (a.a. 1288-1457) alone, however, was insuﬃcient to confer MT minus-end binding (Figure

1.7A).

To assess how these domains aﬀect MT dynamics, I performed dynamic MT growth assays, as previously described for the CAMSAP proteins. Surprisingly, I found that the Patronin CC domain tracked along growing MT minus-ends in vitro, but did not change the rate of minus-end growth (Figure 1.6B,C), similar to the behavior observed for CAMSAP1. In contrast, the Patronin

CC+CKK domain bound to minus-ends and suppressed tubulin incorporation by ∼90% compared with minus-ends of control MTs (Figure 1.6C). The GFP-CC+CKK domain tracked along the tip of these very slow-growing minus-ends, but did not coat the region of new growth like CAMSAP2

(Figure 1.6C). These results show that the Patronin CC domain can bind to MT minus-ends, but must work in coordination with the CKK domain to suppress minus-end dynamics.

The Patronin domain analysis uncovered a construct that binds MT minus-ends but does not suppress growth (Pat CC) and a construct that binds and suppresses minus-end dynamics (Pat

CC+CKK). I then performed rescue experiments with these constructs, as well as full length (FL)

Patronin, to determine which domains are necessary to rescue the interphase Patronin RNAi phenotype. Endogenous Patronin was knocked down using dsRNAs against the 3’ and 5’ UTRs, so that rescue experiments could be performed with Patronin constructs lacking these UTR sequences.

Similar to our prior results (63), the Patronin knock-down with UTR sequences led to decreased mass of the interphase MT network and caused the appearance of small, treadmilling MT fragments in the cytoplasm (Figure 1.6D,E). Transfection of the FL and the CC+CKK constructions led to full rescue (no treadmilling, short MTs) (Figure 1.6D,E). In contrast, the CC domain, which localizes to minus-ends but does not suppress minus-end dynamics, did not rescue the knock-down phenotype

(Figure 1.6D,E). These results demonstrate that suppression of MT minus-end dynamics by the

CC+CKK domain is essential for Patronin’s function in stabilizing MT minus-ends in vivo.

CAMSAP and Patronin proteins cannot bind yeast microtubules, but are unaﬀected by subtilisin treatment

For both the CAMSAP family members and Patronin, I was able to identify minimal regions of the protein necessary for MT minus-end recognition. However, I also wanted to try to gather information about features of tubulin that might be important for specifying minus-end recognition. Recently,

16 A. B.

CH C1CC2C CC3 CKK Time

GFP Pat CC GFP Pat CC+CKK Pat CC

Pat CC+CKK Distance C. 0.6 D. 0 s 0 s Drosophila S2 Rescue 100

0.4 80 40 s 25 s 60

0.2 40 100 s 45 s 20 Minus End Growth Rate (mm/min) % Cells with >5 Free MT E1 E2 E1 E2E1 E2 E1 E2 Control Pat CC Pat CC+CKK no Pat FLPat CC CC+ CKK E. Patronin RNAi +FL-Patronin +Patronin CC +Patronin CC+CKK

Figure 1.6: A. Schematic of Patronin domain structure. The Patronin CC domain and CC+CKK domains bind to MT minus-ends (white arrows) in a kinesin gliding assay. Bar, 2 µm. B. Represen- tative kymographs of dynamic MTs capped by Patronin GFP-CC and GFP-CC+CKK. Horizontal bar, 1 µm; vertical bar, 20 sec. C. Quantitation of minus-end growth rates in the presence of Pa- tronin truncations. Short-term images were scored for Pat CC; long-term images scored for Patronin CC+CKK. For control, N=6; Pat CC, N=1; Pat CC+CKK, N=2, n=10-39 MTs per day; mean and SD shown for each experimental day. Inset: MT growth (red) from stabilized seeds (blue) after 15 min in presence of Pat CC+CKK (green). Scale bar: 1 µm. D. Quantitation of Drosophila rescue experiments. Experiment was performed in duplicate (n=10-12 cells scored on each day); results from each day are shown as separate bars. E. Representative cells from rescue experiments; insets: zoom of cell edge. Yellow arrows in insets mark representative MT fragments used for scoring. Scale bar: full cell, 10 µm, inset, 2.5 µm.

17 534-1457 Crgo htcudbn oM;ti osrc otie infiatprino predicted CC a 10 minimal of smallest GFP-CC868-1457 the bar: portion Scale the with and significant was arrows. labeled microtubules a white Minus-ends 868-1457) labeled with contained GFP-labeled acid Alexa-561 marked construct (amino 4); are with this construct 4 assay 776-1457, (GFP-CC868-1457, MT; Construct Gliding 3: domain to B. 638-1457, 1288-1457. Numbered bind bind 2: region. length). could 6: not 534-1457, unstructured the did and that 1: along that 534-868 region or those regions: minus-end CC and 5: acid the green amino 868-1457, to in following bind shown 4: the to are failed to assay constructs specifi- correspond gliding (these predictions bound constructs red kinesin structure that in a on constructs shown in based GFP-CC are minus-ends made schematic). were MT prediction CC to CC Patronin cally under the denoted of are truncations (truncations Indicated A. 1.7: Figure 534-868 A. Likelihood of Coiled-coil 0.2 0.4 0.6 0.8 1.0 638-1457

776-1457 Patronin Coiled-coilDomain Analysis 868-1457 Does notbindMT Binds MT MinusEnds 1288-1457 MinusEnds µm. 18 6 5 4 3 2 1 B.

Construct 4 Kinesin Gliding Assay Luke Rice has shown that pure tubulin isotopes can be puriﬁed from yeast using a His tag (75);

Minhaj Sirrajuddin has pioneered the use of this technique in our lab (76). Using this purified tubulin, I wanted to compare the binding of CAMSAPs and Patronin to MTs polymerized from yeast tubulin. To this end, I purified tubulin from yeast, using a small amount of labeled porcine tubulin to fluorescently label the tubulin, polymerized MT in vitro and used these microtubules in a gliding assay to assess binding of CAMSAP and Patronin truncations. Interestingly, I found that none of the full-length CAMSAPs, nor the Patronin coiled-coil domain could bind to the yeast microtubules (Figure 1.8A,B,C). I also tested whether binding was affected by the presence of C- terminal tubulin tails, which regulate binding of many MAPs. For the Patronin CC, subtilisin treatment, which cleaves off the C-terminal tail, had no effect on minus-end recognition, suggesting that Patronin (and likely the CAMSAPs) are not recognizing a feature of the C-terminal tails, but rather are recognizing an exposed feature on the α-tubulin that is absent in yeast tubulin. I also tested the CAMSAP3 CKK domain for MT binding; since this protein binds the length of MTs, I wondered whether it would be able to recognize highly conserved features on the surface of the yeast

MT. Interestingly, I saw much less binding of the CKK domain to the yeast MT than to porcine

MT (Figure 1.8D). This suggests that even on the MT surface, there are unique features that are absent in yeast tubulin that the CKK domain of the CAMSAPs recognizes.

1.3 Methods

Homology Tree Generation

Homology trees were generated using the ENSEMBL tree-generating tool (http://www.ensembl.org/ info/genome/compara/homology/method.html) (77; 78).

Plasmid construction and protein expression

For protein expression, CKK domains were cloned into the NotI and KpnI sites of pET28a-GFP plasmid using Gibson cloning (New England Biolabs) and expressed in BL-21 A1 E. coli (Life Tech- nologies). Brieﬂy, overnight starter cultures were diluted 1:100, grown to log phase and induced using

1 mM IPTG for 6-8 hr at 20C. Cells were harvested and lysed using an Emulsiflex, and proteins were purified using NiNTA beads according to manufacturer’s instructions (Qiagen, Inc); following purification, proteins were dialyzed into 25 mM HEPES, pH 7.4, 150 mM KCl, 1 mM TCEP and

10%glycerol. For FL-CAMSAP, Patronin, CAMSAP CC and CC+CKK proteins, coding sequences of the genes were cloned into a pFasBac HTA+EGFP vector and expressed in Sf9 cells. EGFP

19 Merge MTs GFP A. Porcine MT Subtilisin Porcine MT Yeast MT Patronin Coiled-coil

B. FL-CAMSAP2

C. FL-CAMSAP3

D. CAMSAP3 CKK

Figure 1.8: A. Binding of Patronin CC to porcine, subtilisin-treated and yeast MT. B and C. FL-CAMSAP2 and 3 bind to minus-ends of porcine MT, but not to yeast microtubules. D. The CAMSAP3 CKK domain weakly binds to yeast MT, but at a much lower aﬃnity than to porcine MT. Minus-ends labeled with GFP-CAMSAPs/Patronin CC are marked with white arrows. Scale bar: 2 µm.

20 was cloned into the pFasBac vector between BamHI and SacI using conventional cloning; coding sequences of CAMSAP genes and truncations were cloned between SacI and XhoI sites using Gibson cloning. Truncation constructs encode the following amino acids: Pat CC: 534-1457, Pat CC+CKK:

534-1689, Pat CKK :1546-1689, CAMSAP2 CC: 707-1303, CAMSAP2 CC+CKK: 707-1484, CAM-

SAP2 CKK 1342-1484, CAMSAP3 CC: 578-950 CAMSAP3 CC+CKK: 578-1239, CAMSAP3: CKK

1062-1239. Plasmids were transformed into DH10Bac cells (Life Technologies) and bacmids were prepared according to Bac-to-Bac Baculovirus Expression System manual (Life Technologies). Bacmids were transfected into Sf9 cells and virus was harvested according to manufacturer’s instructions. Sf9 cells were used for protein expression and were cultured in suspension in Sf-900 II SFM (Invitrogen) supplemented with antibiotic/antimycotic. Expression was completed for 60-72 hr for constructs with cells at a starting density of 2x106 cells/ml. After collection of infected cells, proteins were puriﬁed following cell lysis in 1% Triton X-100, using NiNTA beads as for CKK proteins.

Protein Preparation

Pig brain tubulin was prepared according to Castoldi and Popov (79). MTs were prepared for each day of imaging by polymerizing ∼6 µm unlabeled tubulin in the presence of 1 mM GMPCPP (Jena

Biosciences). Labeled tubulin (either Alexa-561, Cy5 or biotinylated, as speciﬁed in main text) was mixed into the reaction at a ratio of approximately 1:10 and polymerized in a ﬁnal volume of 20 µl for 30-60 min at 37 C.

The kinesin motor domain, K420, was puriﬁed with an N-terminal 6XHis tag. The dynein motor domain, VY268, GST dimerized, and puriﬁed using a ZZ-tag, as in (80).

Microtubule seeds were formed by incubating 6 µm tubulin, plus Alex-647 and biotinylated tubulin at a ratio of 1 to 10 with unlabeled tubulin, with 1 mM GMPCPP (Jena Biosciences) at

37C for 1 hr.

For yeast MT preparation, yeast were transformed with a plasmid expressing recombinant α- and β-tubulin; cultures were grown in media containing 2% glucose, amino acids and yeast nitrogen base. After approximately 48 hours of growth, raﬃnose was added to the media (to relieve repression of of the galactose promoter); expression was induced with 2% galactose for 20-24 hours. Cells were ground in liquid nitrogen after harvesting and incubated in buﬀer I (50mM PIPES pH6.8, 5mM

MgCl2) with Ni-NTA beads; tubulin was eluted with 50mM PIPES pH6.8, 5mM MgCl2, 500mM

NaCl, 30mM, Imidazole, 500uM ATP. A Q-column was loaded with eluate and fractions collected with varying salt concentrations from the column (Buﬀer A: 25mM PIPES pH6.8, 2mM MgCl2,

1mM EGTA and Buﬀer B: 25mM PIPES pH6.8, 2mM MgCl2, 1mM EGTA, 1M NaCl). Tubulin

21 was concentrated using ammonium sulfate precipitation and polymerized with 1:10 labeled tubulin

(Cy5), 2 mM GTP, 5 µM epothilone-B in BRB80 at 30C overnight. Gliding assays were performed as described for porcine tubulin in In vitro assays section.

Image Analysis: Quantiﬁcation of Minus-End Intensities

Images were background-corrected where indicated using the Fiji Background Subtraction plug-in.

Similarly, where indicated, photobleaching correction was done using the Fiji Bleach Correction plug-in.

To determine the approximate numbers of CAMSAPs bound to minus-end of GMPCPP-stabilized microtubules, I used Fiji (81) to select particles on the surface of the coverslip, and particles on MT minus-ends. I used MatLab code to determine the integrated intensity of the ﬁve closest GFP spots on the glass coverslip to each minus-end-bound GFP-CAMSAP spot and calculated the ratio of these intensities.

Patronin RNAi experiments in Drosophila S2 cells

Drosophila S2 cell lines were cultured in Schneider’s media supplemented with 10% FBS (Invitrogen) and antibiotic/antimycotic mix. Cell lines stably expressing GFP-tubulin and mCherry-Patronin constructs were generated as previously described (63). GFP-tubulin was expressed under a consti- tutive actin promoter; mCherry-Patronin genes were expressed under an inducible metallothionine promoter. For RNAi experiments, dsRNA against the 3’ and 5’ UTRs of Patronin was synthesized by in vitro transcription with the Ambion T7 Megascript kit using S2 cell genomic DNA template and primers amplifying 5’ and 3’ UTR sequences (5’ F: TAATACGACTCACTATAGGGCACAT-

GAAAATTTGTAAG;

5’ R: TAATACGACTCACTATAGGGGACTCCGGCTCTCCGACGCCCGCC;

3’ F: TAATACGACTCACTATAGGGGGAAATGAAATCGTGTATGGGCCG;

3’ R: TAATACGACTCACTATAGGGGTGTACATCCGCTGGCTCTCAC)

Cells were subjected to two rounds of RNAi over the course of 8 days. Cells were prepared for imaging by inducing expression of mCh-Patronin truncation with 100 µm Cu2+ overnight and plating onto concanavalin A for 1 hr prior to imaging. For rescue experiments, cells expressing both GFP- tubulin and the mCh-Patronin construct were selected for time-lapse imaging. Cells were imaged on a Zeiss spinning disc confocal microscope with a 100 1.45 NA objective, using an Andor EM

CCD camera at 5-10 sec intervals. Cells were scored for the Patronin phenotype by three blinded observers for the presence of >5 treadmilling MT fragments at the cell edge (scores were identical

22 for all but ﬁve cells).

In vitro assays

Single molecule assays were imaged using a Nikon Ti microscope (1.49 N.A., 100x objective) by total internal reflection microscopy (TIRF); images were acquired with an Andor EM-CCD camera and MicroManager software (82). Flow chambers (∼8 µl) were created using double-stick tape and silanized coverslips, using a procedure modified from Gell et. al. (83). Briefly, coverslips were washed by sonication in 1 M NaOH, rinsed, dried and plasma cleaned, silanized for 1 hr. using trichloroethylene plus 0.1% dimethyl-dichlorosilane (DDS), washed in three consecutive baths with methanol and sonication, and then dried. Pig brain tubulin was prepared according to Castoldi and

Popov (79). MTs were prepared for each day of imaging by polymerizing ∼6 µm unlabeled tubulin in the presence of 1 mM GMPCPP (Jena Biosciences). Labeled tubulin (either Alexa-561, Cy5 or biotinylated, as speciﬁed in main text) was mixed into the reaction at a ratio of approximately 1:10 and polymerized in a ﬁnal volume of 20 µl for 30-60 min at 37 C.

For motor-driven gliding assays, a ﬂow chamber (described above) was ﬁrst incubated with 0.2 mg/ml of anti-His antibody (5 min), blocked with 1 mg/ml kappa-casein (5 min), followed by ∼100-

200 pM of a truncated, dimeric human kinesin (K420, puriﬁed with an N-terminal 6XHis tag). For dynein gliding assays, the motor domain of dynein (VY268, GST dimerized, puriﬁed using ZZ-tag

(80)), was incubated in a ﬂow cell with acid-base washed coverslips, followed by blocking with 1 mg/ml kappa-casein and addition of motility mix. Motility assay buﬀer (80 mM PIPES, 2 mM

MgCl2, 1 mM EGTA, with 60 mM KCl, as speciﬁed in text) plus CAMSAP proteins (at 10-12 nM) with MT, 4 µm ATP and 2 mM PCA/PCD/Trolox (to reduce photobleaching, (84)) was added to the chamber and images were acquired at 2, 5 or 10 sec intervals for 20-50 frames.

For measuring dynamic tubulin addition to MT seeds, I employed a modiﬁcation of the assay developed in Gell et al. (36). Microtubule seeds were formed by incubating 6 µm tubulin, plus Alex-

647 and biotinylated tubulin at a ratio of 1 to 10 with unlabeled tubulin, with 1 mM GMPCPP (Jena

Biosciences) at 37C for 1 hr. Flow chambers were incubated (5 min each) sequentially with 0.2 mg/ml anti-biotin antibody, followed by blocking with 1 mg/ml kappa-casein, and biotinylated MT seeds.

The mix for measuring MT dynamics was then added, which contained 80 mM PIPES (pH 6.8), 2 mM MgCl2, 1 mM EGTA, 25 µm unlabeled tubulin, 0.33 mg/ml Cy5 or Alexa561-labeled tubulin

(as speciﬁed in text), 60 mM KCl, 1 mg/ml kappa-casein, 0.1% methyl cellulose, 2-4 mM GTP and 2 mM PCA/PCD/Trolox. CAMSAP and Patronin GFP-tagged proteins were added at a ﬁnal concentration of 10-20 nM. Imaging was performed at room temperature. Under these conditions,

23 microtubules grew from both ends and catastrophes were rare. Images were acquired at 2 sec intervals for 100-200 sec; generally; 2-3 fields were imaged per flow chamber. For experiments examining longer time periods, flow chambers were constructed identically to short-term experiments, but were sealed using valap, and images were acquired at 1 or 5 min intervals over 20 min.

Kinesin-13 Depolymerization Assays

For kinesin-13 assays, ﬂow chambers were constructed as described for polymerization assays (sequential incubation with 0.2 mg/ml anti-biotin antibody, 1 mg/ml kappa-casein, and biotinylated,

Alexa-647 labeled MT seeds, polymerized as described above). Depolymerization mix containing

6 nM MCAK (85), 80 mM PIPES (pH 6.8), 2 mM MgCl2, 1 mM EGTA, 2 mM ATP, 1 mg/ml kappa-casein and Trolox/PCA/PCD was ﬂowed into the chamber for imaging. Imaging was done in

TIRF at 10 sec intervals for 10-20 min.

Image Analysis

For MT polymerization and depolymerization experiments, kymographs were made using the Reslice function in Fiji (81) and used to measure growth at microtubule ends. MTs were selected for scoring in which both ends of the MT could be clearly resolved (ie. not blocked by other MTs or out of the TIRF field). Plus-ends were differentiated from minus-ends by the difference in growth rates.

Minus-ends were scored as dynamic if they underwent growth of more than ∼200-300 nm (2-3 pixels) over the course of imaging (200 sec per ﬁeld). Rates of growth for both long and short-term imaging experiments were measured over a period for which the MT could be clearly resolved (usually greater than half of the imaging period) by making a rectangle where the corners marked the beginning and end of growth; the height of the rectangle corresponded to time elapsed, and the width corresponded to the growth distance. For depolymerization assays with kinesin-13, similar criteria were used for choosing MT; MT were then assessed for depolymerization at each end over the course of imaging.

To determine the approximate numbers of CAMSAPs bound to minus-end of GMPCPP-stabilized microtubules, I used Fiji (41) to select particles on the surface of the coverslip, and particles on MT minus-ends. I used MatLab code to determine the integrated intensity of the ﬁve closest GFP spots on the glass coverslip to each minus-end-bound GFP-CAMSAP spot and calculated the ratio of these intensities.

24 1.4 Conclusion

This work, in combination with several other recent studies, provides support for the model that metazoan CAMSAPs/Patronins act as stabilizing proteins for MT minus-ends. Previous work has shown that Drosophila Patronin stabilizes minus-ends against depolymerization by kinesin-13 (63).

Here, I show that this kinesin-13 protection activity extends to the mammalian CAMSAPs and that Patronin, CAMSAP2 and 3 also strongly suppress minus-end growth. While my paper was in submission, Jiang et al. (86) published a paper showing that CAMSAP2 and 3 decrease minus-end polymerization in vitro and protect minus-ends from depolymerization after a laser-induced cut of

MTs in vitro and in vivo. They also showed that a stretch of MT coated with CAMSAP2 is protected against depolymerization traveling towards it from the plus end following a catastrophe.

In addition to these in vitro studies, the MT minus-end protection activity of CAMSAPs/-

Patronin has been shown to be important for maintaining the normal array of non-centrosomal microtubules (63; 69; 86) and the loss of this activity impacts cell migration (86), neurite morphology (71; 72) and mitosis (62; 64). Interestingly, while Patronin plays an important role in mitosis in flies, Jiang et al. did not find mitotic defects of CAMSAP2 knockdown (86). While this result may reveal a difference in the biological function of Drosophila versus mammalian proteins, it is possible that the knockdown of all three CAMSAPs may be needed to uncover a mitotic phenotype.

CAMSAPs also have been reported to interact biochemically with spectrins (an activity from which the CAMSAPs derive their name) and overexpression studies have suggested that this interaction may be important in vivo (68). However, the current evidence supporting a physiological connec- tion of this protein family with spectrin is less clear than data supporting their role in regulating

MT minus-ends. Thus, it may be warranted to develop a new unifying nomenclature for both the invertebrate and vertebrate proteins reﬂecting their MT function in the future.

Despite their common activity in binding MT minus-ends, the three mammalian CAMSAPs exhibit different properties with respect to their effectiveness as a ”cap” for growing MT minus- ends. In my work and that of Jiang et al. (86), we show that CAMSAP1 tracks at the tips of growing minus-ends, while CAMSAP2 and CAMSAP3 suppress minus-end growth, with CAMSAP3 being more robust in its activity. Thus, all of the CAMSAPs can remain bound to the MT end and still allow the addition of new tubulins to varying extents. The different CAMSAPs/Patronin also exhibit different off-rates from the MT lattice (this work and Jiang et al. (86)). CAMSAP1 and Patronin appear to bind at the very tip of the minus-end, and thus, in a similar manner to

+TIP tracking proteins, must dissociate after allowing tubulin incorporation. CAMSAP2, however,

25 appears to remain tightly bound to the microtubule lattice after tubulin incorporation, creating an elongated stretch of bound molecules. CAMSAP3 may show an intermediate dissociation activity.

The mechanism of minus-end recognition (where α-tubulin is exposed) by CAMSAP/Patronin is unclear, although several results suggest a complex binding mechanism. The CAMSAPs and

Patronin bind to the ends of non-dynamic, GMPCPP-stabilized MT in sub-stoichiometric (<13) numbers, suggesting that they can recognize some feature that is unique to the exposed α-tubulin surface at the minus-end. Our data from experiments done with yeast microtubules suggests that this feature is also unique to a structure found in mammalian and eukaryotic tubulins that is absent in yeast tubulins; this may suggest amino acids that are unique to minus-ends. While I did pick a small subset of amino acids (4) that are distinct in yeast tubulin to mutate to the human amino acids, these mutations did not restore binding of Patronin/CAMSAP to the MT minus-ends; thus, further mutagenesis will be required to pinpoint the site of binding at the minus-end. In a dynamic MT assay, tubulin addition results in increasing numbers of CAMSAPs being deposited near the minus end. Since not all of these CAMSAPs can bind to the limited number of exposed α-tubulins at the very minus-end, they must be retained by other means. The lattice aﬃnity of individual CAMSAPs on stabilized MTs appears to be relatively weak (Figure 1.1). However, once deposited at the exposed minus-end by tubulin subunit addition, adjacent CAMSAPs might interact cooperatively with one another to be retained on the newly polymerized MT lattice near the minus-end.

I also used truncation experiments to identify a minimal domain involved in minus-end binding and emerged with somewhat diﬀerent results for the mammalian CAMSAP proteins and invertebrate

Patronin. For CAMSAP, the CKK domain binds to MTs weakly and shows preferential minus- end localization (results here and Jiang et al. (86)), but the Patronin CKK domain does not bind MTs under the same conditions. Conversely, the Patronin CC domain shows clear minus- end binding, while the CAMSAP3 CC domain shows no detectable MT binding activity in the microscopy assay. However, for both CAMSAPs and Patronin, a combination of the CKK+CC produces robust minus-end recognition and suppression of minus-end dynamics in vitro (results here and Jiang et al. (86) and in vivo (Figure1.6)). It seems plausible that mammalian CAMSAPs and invertebrate Patronin have two domains that facilitate minus-end binding; the evolution of binding strengths of these domains and their cooperativity in the intact protein may account for the distinct minus-end behavior of invertebrate Patronin and mammalian CAMSAPs, and between the mammalian CAMSAPs. Consistent with this idea, a region in the CC domain that is important for CAMSAP function and conserved among vertebrate CAMSAPs is not conserved in Drosophila

Patronin (86). Interestingly, the N-terminal calponin-homology domain, which is conserved among

26 all invertebrate and vertebrate CAMSAPS, does not appear to be necessary for MT minus-end regulation and perhaps is involved in a yet-unidentiﬁed function of these proteins.

Our work, as well as the recent studies by Jiang et. al. (86) and Tanaka et al. (69) suggest that

MT minus-ends may be subject to more regulation than previously thought. The vertebrate and invertebrate CAMSAPs all appear to strongly protect minus-ends against depolymerization caused by MT depolymerases or breaks. However, the CAMSAPs and Patronin permit tubulin subunit addition at the minus-end at very different rates. These differences could lead to a tunable system for regulation of minus-end dynamics in cells, allowing for normal growth when bound by CAMSAP1, and slow growth when bound by CAMSAP2 or CAMSAP3. I also find somewhat different efficiencies of protection against kinesin-13-mediated depolymerization by the CAMSAPs. The different off-rates of CAMSAPs from MTs also may have a physiological role, as elegantly demonstrated by Jiang et al. (86) who showed that long, decorated stretches of CAMSAP2 at minus-ends can create stable seeds that can undergo repeated rounds of growth after plus-end catastrophes. The interplay of

CAMSAPs within a single cell may play an important role in regulating the MT network. For example, cooperation between CAMSAPs has been documented in epithelial cells by Tanaka et al.

(86) who observed that microtubule minus-ends are decorated with similar sized, short stretches of CAMSAP2 and CAMSAP3, but found that the length of minus-end decoration by CAMSAP2 increased considerably after CAMSAP3 depletion by RNAi. This result suggests that the size of these minus-end caps may be governed by the relative ratios of these CAMSAPs in the cell.

A number of interesting questions remain about the role of CAMSAP family members. One is how these proteins recognize and stabilize a microtubule minus-end. The features recognized by

CAMSAPs that are specific to the minus-end are unknown. Although one clue comes from our work with yeast tubulin, which suggests that a non-conserved minus-end exposed region is recognized by the CAMSAPs, the high conservation of tubulin between yeast and other organisms means that this feature may be very subtle. It will be useful to determine whether a ”humanized” yeast tubulin, in which surface exposed non-conserved residues have been mutated to the mammalian amino acids, will facilitate CAMSAP minus-end binding. Our results also suggest that Patronin binds MT minus-ends in a way that is fundamentally different from that of γ-tubulin. γ-tubulin caps all protofilaments on a MT end through the γ-tubulin ring complex (γ-TuRC), and completely blocks growth of the MT minus-end. In contrast, CAMSAP proteins individually recognize minus ends, and additionally do not block addition of dimers, permitting slow growth. This data raises a number of questions about how CAMSAPs recognize and bind to minus-ends. For example, how much space does an individual

CAMSAP occupy on a minus-end? How many protoﬁlaments does a bound CAMSAP ”obscure”?

27 How are CAMSAPs retained on a polymerizing microtubule? What differences between the Patronin coiled-coil and the CAMSAP’s coiled-coils allow the Patronin CC to recognize a minus-end on its own? What structural features of the the different CAMSAPs give them different properties for altering MT minus-end growth rates? These questions are all compelling and interesting and will likely require structural approaches to elucidate their answers. These studies are complicated by the structures involved; it seems likely that the CAMSAPs bind to MT minus ends, rather than tubulin dimers, requiring perhaps EM images of CAMSAPs bound at a minus-end. However, locating MT ends on an EM grid to image restricts the sample size it is possible to attain. Therefore, it is currently unclear what structural methods might be tractable for this problem. Further mutagenesis, of both yeast tubulin, and CAMSAP proteins, may help guide future studies. For example, domain swap experiments, such as moving the CKK domain of CAMSAP1 to CAMSAP3 and determining how the swap alters the protein’s behavior, could further our understanding of the specific function of each domain.

Another element of the structural puzzle is understanding how exactly CAMSAPs protect MT from depolyermerization by kinesin-13 proteins. Kinesin-13’s are thought to promote depolymerization by bending back tubulin dimers, leading to induction of catastrophe as lateral contacts are lost between protoﬁlaments (22). Thus, there are several possible mechanisms by which CAMSAPs might protect minus-ends. They could simply block the bending of the tubulin dimers by kinesin-

13, preventing the loss of lateral protoﬁlament contact. Alternatively, they could prevent binding of kinesin-13 at the minus ends (although not along the length of MT- using mCh-tagged kinesin-13, I can visualize binding of the kinesin-13 along the MT length). Or, perhaps the CKK domain (which seems to be required for minus-end protection, at least for Patronin) actually inhibits or blocks the activity of kinesin-13 proteins at the minus-end of a MT. I have no evidence favoring any of these mechanisms; more detailed work could help illuminate this process.

Finally, the role of CAMSAPs in cells remains a rich and unexplored area of study. Jiang et. al. elegantly showed that CAMSAP2 is recruited to severed MT sites in vivo, suggesting that CAMSAP2 is involved in binding to and stabilizing newly formed minus-ends. However, much remains unknown about the function of CAMSAPs in vivo. For example, minus-ends have long been thought to be stable. Jiang et. al., however, showed that minus ends in vivo are dynamic (albeit exhibiting very slow growth), suggesting this accepted dogma is likely untrue. Plus end dynamics have important functions in cell division, cell migration and axonal growth; however, the role of minus-end dynamics is unexplored. The fact that different CAMSAPs have differing effects on minus-ends suggests that perhaps minus-ends are differentially regulated. CAMSAP1 allows growth from the minus end;

28 under what circumstances might this be beneficial? How might the expression of CAMSAP family members and minus-end dynamics be different in different cell types? For example, CAMSAP1 is up-regulated in neurons; perhaps it plays an important role in axonal or dendritic growth. It is possible that different CAMSAPs have activity localized to particular regions of a neuron. The role of CAMSAPs in mitosis is also not understood. The presence of multiple family members in a cell has made knockdown studies difficult, as it is hard to know if loss of one family member can be rescued by the presence of a different CAMSAP. Thus, it is possible that CAMSAPs play a role in mammalian mitosis; it has been shown that Patronin is important for anaphase onset in Drosophila, so it seems likely that these proteins could be important in mammals as well. Finally, I found no role for the calponin-homology domain in MT recognition and growth suppression. However, this domain is conserved through all CAMSAP family members, suggesting it likely plays a functional role in vivo.

What this function might be remains completely unknown. During my rotation, I completed pull- down experiments with Patronin and many of the hits were actin-associated proteins; additionally, calponin-homology domains are often implicated in actin binding. These results raise the suggestive

(although at this point completely speculative) possibility that CAMSAPs/Patronin may have a role in linking the MT and actin cytoskeletons. Alternatively, the CH domain may be involved in other protein-protein interactions; for example, CAMSAP3 has been shown to anchor microtubules at adherens junctions. Perhaps the CH domain is involved in localizing CAMSAP3 at these junctions.

Many open questions remain related to the function of the CAMSAP proteins; hopefully, future studies will reveal how different cell types (e.g. epithelial cells, neurons, fibroblasts) utilize and regulate the different activities of CAMSAP proteins to control non-centrosomal MTs during cell division, migration and differentiation.

29 Chapter 2

Chapter 2

2.1 Introduction

One of the essential functions of the microtubule cytoskeleton is to faithfully segregate genetic material during cell division. To achieve the goal of appropriating the proper amount of genetic material to each cell, there are several steps that must take place, and many controls exist to ensure that all chromosomes are segregated correctly. First, each sister chromatid must become attached to a MT. This task is accomplished by a search and capture mechanism (87), whereby MTs grow and shrink at their plus-ends, probing space until they encounter a chromosome kinetochore, the location of MT attachment. When a kinetochore is encountered, an initial attachment is made laterally along the MT by dynein and a complex known as the RZZ complex (88; 89). This lateral attachment is weak, and is therefore converted to a more stable end-on attachment, mediated in eukaryotes by the

Dam1 complex and a protein known Hec1 (90; 91). These attachments are capable of sensing force across the kinetochore; when all kinetochores are under tension, the spindle checkpoint is released, allowing anaphase, and the segregation of the chromosomes, to begin (92).

The processes described above all require extensive regulation to maintain spindle bipolarity and organization. Many proteins are involved in this process. A protein complex known as augmin binds along the length of MTs in the spindle, anchoring gamma-TuRCs and promoting MT nucleation within the spindle (47). Spindle bipolarity is maintained through a network of cross-linking proteins that perform dual functions of sliding and bundling MTs. PRC1 is a static MT cross linker, while motor proteins such as Eg5 both bundle and slide MT (49; 58). Additionally, at the poles, MT are constantly being disassembled by the activity of depolymerizing proteins such as kinesin-13 family

30 MT depolymerases (93), while at the plus ends, which are oriented toward the center of the spindle,

MT growth is promoted by TOG family proteins, among others (94).

Microtubule motors are deeply involved in process of cell division. There are two major classes of microtubule based motors: kinesins and dynein. Both motor families walk on microtubule tracks.

Kinesins are most often plus-end directed (although there are exceptions), while dynein exclusively walks to the MT minus-end (95). There are approximately 40 kinesins in mammals, and many more in plants; in contrast, dynein exists in just two forms, cytoplasmic and ciliary (96; 52). Dynein forms a large complex, providing many layers of regulation and specialization. Kinesins achieve their specialization by existing in many diﬀerent sub-types. Both classes of motors are extremely important during cell division. Dynein is involved in assembly of the spindle (97; 98), pulling chromosomes toward the spindle poles for chromosome segregation, as well as in the initial capture of chromosomes (99). Kinesins also play diverse roles. Kinesin-13 family proteins are specialized proteins that depolymerize MTs by inducing catastrophes from MT ends. This activity is important for maintaining MT dynamics within the spindle, as well as modulating spindle length (19). Chromokinesins are a class of kinesins that interact with chromosomes and help facilitate chromosome condensa- tion and alignment along the metaphase plate (100). The kinesin-5 family, which includes Eg5, are tetrameric motor proteins, which possess splayed heads that are able to slide anti-parallel microtubules (101; 102). This activity is important for formation of the bipolar spindle, and also facilitates the separation of the spindle at anaphase (103; 104; 105).

To maintain spindle bipolarity, it is essential to have cross linking proteins opposing the tendency of the spindle MT to splay out and become monopolar; in early spindle formation, Eg5 is essential for performing this function. In the absence of Eg5, either by RNAi knockdown or the addition of Eg5 inhibitors such as monastral, spindle formation is perturbed and spindle bipolarity is lost, leading to the formation of monopolar spindles (104). These spindles are not competent for chromosome segregation and lead to cell death after failure in mitosis (106; 107). Because treatment of cells with

Eg5 inhibitors lead to cell death, there was extensive interest in their use as therapeutic agents, and a number of pharmaceutical companies developed potent and highly speciﬁc inhibitors of Eg5.

However, many of these therapeutics failed in clinical trials because of a lack of eﬃcacy; cancers treated with the Eg5 inhibitors frequently develop resistance to the drugs.

Work done by Tanenbaum et. al. pointed to a possible the cause of this resistance (108). By systematically knocking down all kinesins known to be involved in mitosis, Tanenbaum showed that a kinesin-12 family member, Kif15, was able to compensate for the loss of Eg5 activity. Kif15 is thought to have a similar structure to Eg5 (102). In cells that were resistant to Eg5 inhibition and

31 formed bipolar spindles in the presence of Eg5 inhibition, Kif15 knock down led to the formation of monopolar spindles. Further experiments showed that in wild-type cells, Eg5 is necessary only for bipolar spindle formation; once a bipolar spindle has formed, it can be maintained in the absence of Eg5 activity (108). However, Kif15 is required for the maintenance of bipolarity; loss of both

Eg5 and Kif15 leads to spindle collapse. This experiment provides an assay for testing eﬃcacy of possible Kif15 inhibitors.

In the studies described in this chapter, I tested Kif15 inhibitors developed by Cytokinetics for eﬃcacy at inhibiting Kif15 activity by two methods; in vitro ATPase assays with the Kif15 motor domain and in vivo assays for spindle collapse. I ﬁnd that of the three inhibitors tested, all three possess activity against Kif15, both in vitro and in vivo. Two of the inhibitors (herein referred to as

”GSK03” and ”CK57”) are the most active; ”GSK01” is the least active both assays. Additionally,

I selected two cell lines, starting with wild-type HeLa and U20S cells, with an ispinesib-like molecule

(although the molecule used in these studies is a derivative of ispinesib, a patented compound which we could not obtain for use in the lab, I will use the name ispinesib throughout this text since they have very similar structures and activity), a potent inhibitor of Eg5 which is used in the clinic. Once these cell lines were resistant to 5-10 nM ispineisb, I tested them to see if the acquisition of Eg5 resistance would lead them to have increased sensitivity to Kif15 inhibition.

2.2 Results

Addition of Kif15 inhibitors reduces ATPase rate of Kif15 motor domain

Our first goal upon receipt of the drugs was to confirm that they could inhibit ATPase activity of purified Kif15 motor domains. Therefore, I performed ATPase assays using a PK/LDH (pyruvate kinase/lactate dehydrogenase) assay. This assay measures a decrease in absorbance of NADH over time; through enzyme activities shown in Figure 2.1, generation of free Pi is coupled to a reduction in NADH and absorbance at 340 nm. I employed this assay to determine if the addition of the

Cytokinetics inhibitors would slow the motor domain’s ATPase activity.

To purify the Kif15 motor domain, we cloned the motor domain (with no coiled-coil linker) into a pET-28a vector; because this protein is somewhat unstable, a solubility tag (GB1) was fused in-frame with the protein to increase stability of the motor domain. Puriﬁed protein was added to the assay mixture and absorbance at 340 nm was measured every 5 seconds for 7 minutes. In the absence of microtubules, the motor domain of molecular motors that walk on microtubules is

32 Figure 2.1: Pyruvate kinase catalyzes the reaction of PEP to pyruvate using phosphate generated by the ATPase activity of the enzyme being studied, which the lactate dyhydrogenase enzyme then converts to LDH. This process leads to the reduction of NADH to NAD+; by measuring the decrease in absorbance of NADH at 340 nm, the rate of phosphate production of the motor protein under study can be indirectly measured. Image from the Department of Cell Biology at Duke University. inactive; therefore, a reaction done in the absence of microtubules was used as a baseline for complete motor inhibition.

The results shown in Figure 2.2A show sample reaction traces for the uninhibited Kif15 motor domain (green), the no-MT control reaction (red), and the motor domain in the presence of the three inhibitors (shades of yellow). In the absence of MT, very little motor activity can be measured. In the presence of each of the three inhibitors, a moderate reduction in motor ATPase activity is observed.

Figure 2.2B shows the fold inactivation for each inhibitor over the native ATPase activity of the

Kif15 motor domain. As a control, I included the motor Eg5, for which we have a potent inhibitor

(a close analogue of the clinical drug Ispinesib). For all inhibitors tested, I saw between a 2-4 fold reduction in motor activity. This result was somewhat surprising; I expected a full (approximately

8-10 fold) inhibition of motor activity compared to the native activity. The reasons for this relatively weak inhibition remain unclear.

Kif15 inhibitors have an uncompetitive mechanism of action against Kif15

In addition to confirming that the Kif15 inhibitors had activity against purified Kif15 protein, I also wanted to obtain information about how these inhibitors were affecting Kif15 activity. Because kinesins have activity only in the presence of MTs and hydrolyze ATP to generate force for movement, kinesins are considered to act on two substrates: MTs and ATP. Therefore, binding to either of these substrates could compete for binding to the inhibitors. This type of inhibition would suggest that at high concentrations of the substrate, it would compete for binding to the binding pocket, reducing the potency of the inhibitor. Alternatively, the inhibitors could act by an uncompetitive mechanism; here, we would expect that even at high substrate concentrations the inhibitor would continue to have the same effect on Kif15 activity. To differentiate between these two mechanisms, I performed

33 A. B. 1.5 6.0

1.0 4.0

No MT, Kif15 GSK03 0.5 CK57 Fold Inhibition Fold ATPase Rate (a.u.) ATPase 2.0 GSK01 Kif15

20 60 100 140 180 Eg5/ Kif15/ Kif15/ Kif15/ Time (seconds) Isp. GSK01 GSK03 CK57

Figure 2.2: A. Example traces of the activity of the Kif15 motor domain, shown by a decrease in absorbance at 340 nm. Green shows the rate of activity of the uninhibited motor domain; red shows a control in which no microtubules have been added. Yellow traces show activity in the presence of the three Cytokinetics inhibitors, as indicated. B. Fold inhibition of activity of the motor domain compared to uninhibited motor activity.

ATPase assays with a range of MT and ATP concentrations and measured ATPase activity using the same PK-LDH reaction described above; this time, I utilized a plate reader format to facilitate using a panel of concentrations.

Figure 2.3A,B shows the curves for the Kif15 ATPase activity with increasing MT and ATP concentrations; as expected, activity increases logarithmically with increasing concentrations, and plateaus. Similar activity is seen in the presence of inhibitors. To determine the mechanism of inhibition, I then used this data to construct a Lineweaver-Burk plot (109). If the fit lines for the double reciprocal plot of enzyme activity vs. substrate concentration for uninhibited versus inhibited reactions cross at the x-axis, inhibition is considered competitive; if the fit lines intersect at the y- axis, then non-competitive inhibition is being observed. Points of intersection in between those two axes are considered ”mixed” inhibition. Based on Lineweaver-Burk plots of the varying MT and ATP concentrations, it is clear that the fit lines for the inhibited and uninhibited reactions are not crossing at the x-axis; while the fit lines are not crossing at y-axis either, they are closer to intersecting at the y-axis. This data is suggestive that the inhibitors function through a non-competitive mechanism for both their MT substrate, as well as ATP.

34 Control (no inhibitor) CK57 A. B. 10 14 GSK01 GSK03 12 8 10 6 8

4 6 4 ATPase Rate (a.u.) ATPase 2 Rate (a.u.) ATPase 2

504 0302010 1.00.80.60.40.2 [MT] (mM) [ATP] (mM) C. D. 0.4 0.6 1/ATPase Rate (a.u.) 1/ATPase 1/ATPase Rate (a.u.) 1/ATPase

-2.5 -1.5 -0.5 1.5 1.5 -6.0 -4.0 -2.0 6.04 .02.0 1/[MT] (μM) 1/[ATP] (mM)

Figure 2.3: A. Measurements of ATPase rate were made using varying concentrations of MT, from 0 to 100 µM, in the presence of absence of 10µM of each inhibitor, as indicated. Measurements were made using a plate reader; the absolute value of the ATPase rate is shown with standard error for n=4 replicates. B. As done in A, measurements were made of Kif15 ATPase activity in the presence of varying concentrations of ATP, ranging from 0 to 1 mM. Standard error of n=3 replicates shown. To determine the mechanism of inactivation, data shown in A and B were plotted in a Lineweaver-Burk plot (109). The inverse of the ATPase rate was plotted against the inverse of substrate concentration; (C) shows this plot for varying MT concentrations, (D) for varying ATP concentrations. Fit lines are shown in black.

35 Simultaneous inhibition of Kif15 and Eg5 in pre-formed spindles leads to spindle collapse

I next wanted to test whether the addition of Kif15 inhibitors could disrupt spindle maintenance in vivo. Because Eg5 can compensate for loss of Kif15 activity in spindle formation and maintenance, no eﬀect of adding the Kif15 inhibitors had been previously observed. To circumvent this problem, and isolate the activity of Kif15 from that of Eg5 in wild-type cells, I performed the spindle collapse assay developed by Tanenbaum et. al. (108).

In this assay, I used a proteasome inhibitor (MG132) to arrest cells in metaphase, after formation of a bipolar spindle. Once the spindle is formed, the activity of either Kif15 or Eg5 is suﬃcient to maintain spindle bipolarity (Eg5 is dominant for bipolar spindle formation). Thus, inhibiting just

Kif15 or Eg5 has no eﬀect on spindle bipolarity (Figure 2.4B,C). Indeed, I saw no eﬀect on spindle bipolarity after the addition of only either ispinesib, or any of the individual Kif15 inhibitors (Figure

2.4C, data not shown). However, with a combination of 5µ M ispinesib plus 10µ M CK57 and GSK03, spindle collapse was observed in approximately 40% of spindles. This percentage was slightly lower, approximately 20%, for GSK01 which I found to be a less potent inhibitor of Kif15 (Figure 2.4B).

I additionally determined the concentration dependence of each drug on spindle collapse (Figure

Figure 2.4C). I found that for all the Kif15 inhibitors, 20µ M concentrations of the inhibitors produced the most potent response. For GSK03 and CK57, almost 100% of the spindles collapsed in the presence of 20µ M drug. For GSK01, even a 20µ M drug concentration led to only approximately

30% collapsed spindles. These results indicate that the Kif15 inhibitors are active against Kif15 in vivo, although high drug doses are required to observe spindle collapse.

The ultimate goal of the development of the Kif15 inhibitors would ideally be their application in a clinical setting, where the hope would be that the combination of Kif15 and Eg5 inhibition would prevent resistance to Eg5 inhibition, since Kif15 seems to be the dominant motor that compensates for loss of Eg5 activity (108). Therefore, we sought to test whether cell lines that had been selected to grow in the presence of ispinesib, and therefore presumably in the absence of Eg5 activity, would be more sensitive to Kif15 inhibition. Both the HeLa and U20S cell lines did not show a spindle morphology phenotype in the presence of Kif15 inhibition prior to selection with ispinesib (data not shown). After selecting cells for approximately 30 passages in increasing concentrations of ispinesib,

I used these cell lines to look at the eﬀect of Kif15 inhibition on ispinesib resistant cells. I tested several conditions using both HeLa and U20S cells, as well as diﬀerent concentrations of the Kif15 inhibitor (I used GSK03 for all these experiments) and ispinesib. To automate the analysis of these

36 MG132+ 5 μM isp.+ A. B. MG132 20 μM GSK03 60%

40%

20% Tubulin DAPI Tubulin 10 μm Percent Monopolar Spindles Percent C. MG isp. CK57 GSK01 GSK03 100% +ispenisib 80%

60%

40%

20% Percent Monopolar Spindles Percent

MG MG+ MG+ MG+ CK57 GSK01GSK03 CK57 GSK01 GSK03 +ispenisib

Figure 2.4: A. Images of bipolar and collapsed, monopolar spindles; bipolar spindle shown in the presence of 5µ M MG132; collapsed spindle shown with 5µ M MG132, 5µ M ispinesib and 20µ M GSK03. B. Graph showing percent monopolar spindles from 3 independent experiments in the presence of 5µ M inpinesib plus 10µ M of the indicated inhibitor. For ispinesib alone, results and standard deviation shown for 2 independent experiments with 5 µ M ispinesib. C. Concentration dependence of spindle collapse. For single drug experiments, 10µ M Kif15 inhibitor (indicated) was incubated with cells; for Eg5 and Kif15 inhibition, 5µ MG132, 5µ M ispinesib and 5, 10 or 20 µ M Kif15 inhibitor was incubated with cells.

37 images, I used the TrackMate plugin in Fiji; since cells round up during mitosis, I was able to use

TrackMate to detect primarily mitotic cells with fairly good fidelity (as is always the case with automated image analysis, the tracking was not perfect- however, given the large number of data points, I feel that these imperfections are fairly minor). Tracks were generated for each mitotic cell detected; when a cell actually completed a division and flattened, the track often underwent a split prior to disappearing; by plotting these events in a different color than all other tracks, I was able to identify bona fide cell divisions. To determine the effect of drug combinations on the cells, I used the track data to make histograms of the frequency of track persistence; cells that divide successfully should have, on average, short tracks which split. In contrast, cells that are arrested in mitosis should have long tracks that do not split.

In figure 12, for each experimental condition on two experimental days, I have plotted the frequencies of track lengths (green) and the frequencies of split tracks (red). Figure 2.5A shows an example event that is plotted in red; these are cell divisions in which a mitotic cell is tracked and splits, representing a verified successful mitotic event. Figure 2.5B shows a set of events that would be plotted in green; these are mitotic cells that persist for many frames in the movie. While the data are not perfect, there are several take-aways that can be readily appreciated. One immediate feature of the data is that the HeLa and U20S cells behave very differently from each other in this experiment (Figure 2.5C,D). The HeLa cells were more resistant to ispinesib (in the presence of

5-10 nM ispinesib, most track lengths are short, and many splitting events are observed) and also more sensitive to Kif15 inhibition; in the presence of both GSK03 alone, and GSK03+ispinesib, at all concentrations, longer track lengths and fewer splits are observed (Figure 2.5C). In contrast, the

U20S cells are likely only partially ispinesib resistant; fewer track splits are observed, as well as fewer cell division events being detected overall. Additionally, the U20S cells were much less sensitive to

Kif15 inhibition. There were fewer cell divisions in the GSK03 or GSK03/ispinesib condition overall compared to ispinesib or DMSO treated cells, but those cells that did divide largely were able to do so successfully, as evidenced by the short track lengths (Figure 2.5D). Overall, these results suggest that cells may acquire ispinesib resistance through different mechanisms, and that the mechanism of resistance may affect the effect of Kif15 inhibition on those cell lines.

38 A. HeLa Cells- “Split path” B. HeLa Cells- No split

0 2 3 5 8 0 4 9 23

C. Day 1

no drug 10 nM isp. 10 μM 20 μM 10 nM isp GSK03 GSK03 20 μM GSK03 HeLa Day 2

no drug DMSO 5 nM isp. 10 μM 20 μM 40 μM 5 nM isp 5 nM isp 5 nM isp GSK03 GSK03 GSK03 10 μM 20 μM 40 μM D. GSK03 GSK03 GSK03 Day 1

no drug 10 μM isp. 10 μM 20 μM 10 nM isp, GSK03 GSK03 20 μM

U20S GSK03 Day 2

no drug DMSO 5 nM isp. 10 μM 20 μM 40 μM 5 nM isp 5 nM isp 5 nM isp GSK03 GSK03 GSK03 10 μM 20 μM 40 μM GSK03 GSK03 GSK03

Figure 2.5: A. Example of a successful cell division; a single mitotic cell is tracked for 2 frames, followed by a track splitting event, cell ﬂattening and loss of the track. B. Example of arrested mitotic cells being tracked over many sequential frames. C. Results of tracking HeLa cells in two independent experiments with drug concentrations and combinations as indicated. Green frequencies show all tracking events; red frequencies show tracks with splits; histograms have 5 frame bins (x- axis); each frequency hashmark represents 20 events (y-axis). D. Results of tracking HeLa cells in two independent experiments with drug concentrations and combinations as indicated; histograms arranged as in D. 39 2.3 Methods

Protein Puriﬁcation and ATPase Assays

Kif15-GB1-His was cloned into the pET-28a expression vector; protein was expressed in BL-21A1 cells overnight at 18C. Cells were lysed using an Emulsiflex and protein was purified using Ni-NTA beads (Qiagen). ATPase assays were done with 250 nM Kif15. Briefly, PK/LDH reactions were prepared by mixing 20µM taxol, 65 µM microtubules (polyermized in BRB80 plus 1 mM GTP and

20 µ M taxol at 37C for 1 hour), 2 mM PEP, 50 µM NADH, 2µl PK/LDH enzyme mix and 1 mM

ATP, rapidly mixing, and reading absorbance at 340 nm every 5 seconds for 7 min.

To determine the mechanism of action of Kif15 inhibitors, plate reader assays were completed essentially as described; a master mix with components listed above was mixed and pipetted into a

Corning 96-well half-volume plate. Readings were taken every 15 seconds in the plate reader at 340 nm for 15 minutes.

Cell Culture and Cell-based Assays

U20S and HeLa cells were cultured in DMEM plus 10% FBS and pen/strep/glutamine (Invitrogen).

Spindle collapse assays were performed by plating cells on glass-bottom plates overnight, followed by drug treatment the following day. Cells were treated with 5 µ M MG132 for 30 minutes, followed by one hour treatment with ispinesib or Kif15 inhibitors, at concentrations indicated in the text. Cells were ﬁxed with 3.3% formaldehyde for 5 minutes, washed with PBS, permeabilized with 1% Triton

(in PBS) and stained with α-DM1α antibody (to stain tubulin, 1:10,000) for one hour, then stained with α-mouse FITC-conjugated secondary antibody for 30-60 min, followed by a brief (5-10 min) incubation with 1 µ g/ml DAPI (to stain DNA). Cells were imaged by epiﬂuorescence microscopy using a Nikon microscope (aka superscope).

For resistance assays, U20S and HeLa cells were cultured as described above, with the addition of increasing concentrations of ispinesib (starting at 0.1 nM ispinesib and working up to 5-10 nM ispinesib over the course of approximately 30 passages). For overnight spindle imaging, cells were plated onto glass-bottom dishes (Brooks Life Science Systems, MatriPlate, 96-well glass bottom micro well plates) without selection drug overnight. Cells were transferred into imaging media (In- vitrogen DMEM/F-12, HEPES, no phenol red) plus 10% FBS plus drugs at concentrations indicated and imaged at 15 minute intervals overnight (12-15 hours) using transmitted light. For each condition tested, I imaged 10 ﬁelds of the well. Images were analyzed using TrackMate (81), a Fiji plugin.

Brieﬂy, I implemented the TrackMate code in Python to run on a batch of images; TrackMate’s

40 algorithm detects round objects, which correspond (mostly) to dividing cells. Data about the track length and the number of splits in a tracked cell (a split in a track generally corresponds to a cell division) was obtained from the tracking and was graphed in MatLab; 5-frame intervals were used to bin track lengths, and frequency was plotted on the y-axis. Subsets of tracks containing splits

(bona-ﬁde cell divisions) were plotted as sub-bars within the overall track length bins.

2.4 Conclusions

The results summarized above suggest several major conclusions. First, and most obviously, is that the group of inhibitors tested has activity against the Kif15 motor domain and is capable of recapitulating the ﬁndings of Tanenbaum et. al. (108). Treatment of cells with MG132 to arrest cells in mitosis, followed by inhibition of Eg5 and Kif15, leads to spindle collapse, while inhibition of just one of the spindle motors allows the bipolar spindle to persist. These results are in keeping with

Tanenbaum et. al.’s results using Kif15 RNAi in conjunction with a pharmacological Eg5 inhibitor.

Although I found minor diﬀerences in the abilities of the compounds to produce spindle collapse and inhibit Kif15 ATPase activity, overall they are eﬀective against their targets.

I also did some experiments to attempt to address the mechanism of action that these inhibitors use to attenuate Kif15 activity. By performing ATPase assays with varying concentrations of the motor’s two substrates, ATP and microtubules, I was able to show that the Kif15 inhibitors most likely interfere with the activity of Kif15 through an uncompetitive mechanism; however, I do not know the structural basis of the binding of these inhibitors or how they might interfere with Kif15 activity (it is possible that Cytokinetics has more information about these details).

Finally, in an attempt to determine the eﬀect of Kif15 inhibition on cells that can divide in the

(presumptive) absence of Eg5 activity, I selected cells using ispinesib, a potent Eg5 inhibitor. I used these ispinesib resistant cell lines to test whether cells that had been selected to be able to divide in the absence of Eg5 activity showed an increased dependence on Kif15, and would thus be sensitive to Kif15 inhibition. I was able to select HeLa cells that were able to divide in the presence of 5-10 nM ispinesib; the U20S cells were more difficult to select, and therefore may not have been completely ispinesib resistant. I found that for the HeLa cells, there was marked decrease in successful cell divisions and many arrested cells both in the presence of GSK03 (one of the Kif15 inhibitors) alone, as well as with GSK03 plus ispinesib; the drugs did not seem to have a strong additive effect. In contrast, for the U20S cells, I found that the cells were partially arrested in the presence of ispinesib (suggesting their incomplete resistance), but did not seem to be affected by

41 the inhibition of Kif15. However, although I am not sure how significant this result is, it seemed that there were fewer dividing U20S cells overall in the presence of GSK03; it is unclear if Kif15 inhibition actually prevented cells from entering mitosis, if this was an experimental coincidence or perhaps an off-target effect of the molecule. The combination of these results point to two major facets of the data. One is that there is a clear effect of Kif15 inhibition on the ispinesib resistant

HeLa cells; treatment of these cells with GSK03 led to a marked increase in arrested cells, both in terms of depressing the number of divisions and increasing the track length. The U20S cells behaved somewhat differently, showing only minor effects in the presence of either GSK03 or the combination of GSK03 and ispinesib. Second, the clear differences in behavior between these two cell lines points to the important of testing multiple cell lines before generalizing results; it is clear that there are major differences in the behaviors of these cells. A deeper understanding of these differences between cell lines in their response to ispinesib selection and what it might mean for drug efficacy would be essential before moving forward with further development of these molecules.

42 Chapter 3

Chapter 3

3.1 Final Thoughts

The work described in the preceding chapters, like all scientiﬁc work, uncovers at least as many questions as it addresses. Therefore, many directions for future study may be worth undertaking in the future (although not by me). I believe that the study of CAMSAP proteins is a rich and interesting area that is worth pursuing further. This family of proteins is unique and I believe provides both a tool for visualizing minus-ends of microtubules, and is an intriguing protein in its own right. In the introduction, I mentioned that very little is known about microtubule minus-ends.

I believe one major limitation on their study previously has simply been the diﬃculty in visualizing them. In contrast to microtubule plus ends, which are dynamic, generally extend to the periphery of the cell, facilitating imaging, and have a plurality of bright, easily tagged markers, minus-ends are comparatively very diﬃcult to locate, visualize and track. CAMSAPs have clear applications for this basic purpose; several recent papers (86; 69; 65; 70) have clearly visualized CAMSAPs at MT minus-ends, which are located throughout the cell (and not only clustered around the centrosome).

Additionally, in preliminary work with Marvin Tanenbaum, in our lab, using a construct Marvin calls ”sunGFP”, which labels proteins with 24x GFPs, we can see very bright labeling of individual

CAMSAPs in the cell; we also were able to see clear minus-end labeling of MTs, and saw many minus- ends non-intuitively facing toward the outer periphery of U20S cells. Thus, I think that CAMSAPs are an important new labeling tool to study the localization of MT minus-ends in varied cell types.

In cell types with non-canonical MT arrays, such as neurons, myotubes or polarized epithelial cells, labeling with CAMSAP proteins may help elucidate the MT organization that allows these cells

43 to function. Recent work by Yau et. al (110) has used CAMSAP2 to mark minus-ends of MT in neurons, and shown that the presence of CAMSAP2 helps stabilize MT in neuronal processes.

However, many aspects of MT localization and behaviour remain open areas for exploration.

Another intriguing dimension of CAMSAP biology is the existence of three homologues in most mammals. Current work has not addressed what divergent roles the homologues may play in diﬀerent biological systems, or even clearly established where the homologues are expressed. Work done in the

Allen Brain Atlas project (111) found low expression of CAMSAP1 in mouse brain, but other studies

(112) have shown the presence of CAMSAP1 in neurons. CAMSAP2 appears to be the most highly expressed CAMSAP in neuronal tissues, while CAMSAP3 has lower expression (111). However, expression studies for these proteins outside of large screening contexts have not been established yet. Therefore, simply determining a profile of tissue expression for each of these proteins would be valuable. Additionally, little is known about the regulation of these proteins throughout the cell cycle. Interestingly, Wang et. al. recently showed that Patronin has a role in regulating the timing of anaphase onset in Drosophila (64), suggesting that in Drosophila, Patronin exhibits cell-cycle dependent regulation as well. In general, however, it appears that CAMSAP2 and CAMSAP3 are the most widely expressed homologues of the CAMSAP family; they also behave the most similarly biochemically (86; 113). Given that CAMSAP1 allows growth at the minus-end of the microtubule, it is intriguing to speculate that it may have a specialized role in either a specific tissue or perhaps at specific times during the cell cycle. For example, perhaps in differentiating neurons, it is expressed early in differentiation to allow fine-tuning of the MT architecture, but once the neuronal processes are established, the MT are stabilized by CAMSAP2/3. While these ideas are wild speculation, this line of investigation could lead to interesting information about MT organization and regulation by

CAMSAPs.

Very little is known about the structure of CAMSAP proteins. While we now have significantly more information about the domain architecture of Patronin and CAMSAPS (86; 113), what these domains look like structurally remains unclear. Although a structure of the CAMSAP3 CKK domain has been deposited in the PDB (1UGJ), how this structure might relate to the function of the protein is unknown. There are a number of pieces of information that could be obtained from structural studies. For example, Drosophila Patronin and the mammalian CAMSAPs have variability in their ability to block growth at MT minus-ends. Differences in the crystal structures of these proteins (if solved) might point to functional consequences in their effects on MT minus-ends. More detailed structural analysis might also help point to clues about how these proteins actually recognize and affect minus-ends. Currently, there is no structural insight into how CAMSAPs specifically recognize

44 minus-ends, how they can bind to minus-ends while permitting growth or how they might stabilize minus-ends, both intrinsically and against depolymerization by kinesin-13. Obtaining structural information on CAMSAP proteins alone, while likely to be a challenge, should be fairly straight- forward. However, determining how CAMSAPs actually interact with the minus-end of a MT is likely to be more challenging. It is possible that electron microscopy of MT minus-ends could help reveal some information about CAMSAP binding to minus-ends, although this approach is likely to be technically challenging, given that the end of a microtubule makes up such a small part of the molecule. Thus, obtaining enough images of minus-ends to obtain a meaningful reconstruction might prove impossible. Alternatively, using new techniques developed ﬁrst in Luke Rice’s lab and in our lab (75; 76) that allow production of recombinant tubulin in yeast, it may be possible to infer the binding site of the CAMSAP proteins. My results show that CAMSAPs and Patronin cannot recognize yeast tubulin. By making a humanized yeast tubulin mutant, in which all non-identical surface residues of the tubulin have been mutated to the human amino acids, it may be possible to isolate the patch of exposed α-tubulin at the minus-end that is recognized by the CAMSAPs.

An additional intriguing observation about the binding of CAMSAPs is that for the mammalian proteins, binding at the polymerizing end of a microtubule leads to the deposition of CAMSAPs along the length of the microtubule lattice (86); interestingly, this behavior was not seen for the

Patronin protein, even after long periods of polymerization (113). It is possible that the coiled-coil of the CAMSAP proteins is involved in mediating interactions between adjacent CAMSAPs as they interact with the polymerizing MT minus-end. However, the absence of this behavior for Patronin could mean that structural information about the conﬁguration of the coiled-coils in these proteins might provide information that would help clarify this behavior.

In Chapter 2, I describe work that I undertook brieﬂy in the lab to characterize the function of a set of inhibitors that act against Kif15, a kinesin-15 family motor protein that is known to play a role during mitosis. The rationale for the development of these inhibitors was to improve the eﬃcacy of treatment of cancers with inhibitors of the kinesin-5 family protein Eg5. Eg5 is important for maintaing spindle bipolarity and its inhibition leads to the formation of monopolar spindles during cell division and arrest of cells in mitosis. However, cancers treated with Eg5 inhibitors frequently develop resistant to these drugs. Marvin Tanenbaum showed during his graduate work that Kif15 is a key molecule who over expression can compensate for the loss of Eg5 activity, either through drug inactivation or knock-down by RNAi (108). In parallel to this work, Cytokinetics developed drugs with activity against Kif15; after Marvin’s paper came out, it seemed possible that treatment with

Eg5 and Kif15 inhibitors might help prevent resistance to Eg5 inhibition. While the Kif15 inhibitors

45 clearly have activity against Kif15, I remain skeptical about their efficacy as a clinical entity. First, the molecules that I tested do not, at least in my hands, seem to potently inhibit Kif15 in vitro or in vivo. In the spindle collapse experiments described above, I had to use 20 µM drug concentrations to achieve collapse of approximately 90% of spindles in the assay; this concentration is much higher than I expected for drugs with a reported 30-300 nM binding affinity. However, more broadly, I am skeptical of the concept of this class of drugs. The Eg5 inhibitors work extremely effectively to induce monopolar spindles, but still have low clinical efficacy against human cancers. While this may be partially explained by redundancy between motor proteins, I feel that it is also possible and even likely that the reason that other drugs that inhibit mitosis (i.e. taxanes, which are widely used as a therapeutic) are effective because of their pleiotropic effect on cells. Thus, drugs that are more specific as spindle poisons may actually have reduced clinical efficacy compared to taxanes, although they are more specific. However, I think the Kif15 inhibitors could be useful for studying the roles of mitotic kinesins during cell cycle progression. The activity of Eg5 is thought to be necessary early in mitosis for coalescing microtubules into a bipolar spindle, which Kif15 may play a role in sliding microtubule apart later in mitosis. However, because these activities are fairly acute and happen over the timescale of mitosis, which takes place in only 30-40 minutes, it is difficult to inhibit these activities within this timeframe by RNAi. Drug application, on the other hand, can be done rapidly during mitosis and has an acute effect on the proteins in question. Thus, the combination of Eg5 and Kif15 inhibitors could be effectively used to study what contributions these motors make during different phases of mitosis.

46 Bibliography

[1] T. J. Mitchison, “Localization of an exchangeable gtp binding site at the plus end of micro-

tubules.,” Science, vol. 261, pp. 1044–1047, Aug 1993.

[2] D. N. Drechsel and M. W. Kirschner, “The minimum gtp cap required to stabilize micro-

tubules,” Curr Biol, vol. 4, pp. 1053–61, Dec 1994.

[3] T. Mitchison and M. Kirschner, “Dynamic instability of microtubule growth.,” Nature, vol. 312,

pp. 237–242, Nov 1984.

[4] H.-W. Wang and E. Nogales, “Nucleotide-dependent bending ﬂexibility of tubulin regulates

microtubule assembly,” Nature, vol. 435, pp. 911–915, 06 2005.

[5] E. Nogales, M. Whittaker, R. A. Milligan, and K. H. Downing, “High-resolution model of the

microtubule,” Cell, vol. 96, pp. 79–88, Jan 1999.

[6] P. Bieling, L. Laan, H. Schek, E. L. Munteanu, L. Sandblad, M. Dogterom, D. Brunner,

and T. Surrey, “Reconstitution of a microtubule plus-end tracking system in vitro.,” Nature,

vol. 450, pp. 1100–1105, Dec 2007.

[7] A. Akhmanova and M. O. Steinmetz, “Tracking the ends: a dynamic protein network controls

the fate of microtubule tips.,” Nat Rev Mol Cell Biol, vol. 9, pp. 309–322, Apr 2008.

[8] A. A. Hyman, S. Salser, D. N. Drechsel, N. Unwin, and T. J. Mitchison, “Role of gtp hydrolysis

in microtubule dynamics: information from a slowly hydrolyzable analogue, gmpcpp,” Mol Biol

Cell, vol. 3, pp. 1155–67, Oct 1992.

[9] S. Inou´eand H. Sato, “Cell motility by labile association of molecules. the nature of mitotic

spindle ﬁbers and their role in chromosome movement,” J Gen Physiol, vol. 50, pp. Suppl:259–

92, Jul 1967.

47 [10] P. Bieling, L. Laan, H. Schek, E. L. Munteanu, L. Sandblad, M. Dogterom, D. Brunner,

and T. Surrey, “Reconstitution of a microtubule plus-end tracking system in vitro,” Nature,

vol. 450, pp. 1100–5, Dec 2007.

[11] P. Bieling, S. Kandels-Lewis, I. A. Telley, J. van Dijk, C. Janke, and T. Surrey, “Clip-170

tracks growing microtubule ends by dynamically recognizing composite eb1/tubulin-binding

sites,” J Cell Biol, vol. 183, pp. 1223–33, Dec 2008.

[12] R. Dixit, B. Barnett, J. E. Lazarus, M. Tokito, Y. E. Goldman, and E. L. F. Holzbaur,

“Microtubule plus-end tracking by clip-170 requires eb1,” Proc Natl Acad Sci U S A, vol. 106,

pp. 492–7, Jan 2009.

[13] S. Honnappa, S. M. Gouveia, A. Weisbrich, F. F. Damberger, N. S. Bhavesh, H. Jawhari,

I. Grigoriev, F. J. A. van Rijssel, R. M. Buey, A. Lawera, I. Jelesarov, F. K. Winkler,

K. W¨uthrich, A. Akhmanova, and M. O. Steinmetz, “An eb1-binding motif acts as a mi-

crotubule tip localization signal,” Cell, vol. 138, pp. 366–76, Jul 2009.

[14] S. C. Schuyler and D. Pellman, “Microtubule ”plus-end-tracking proteins”: The end is just

the beginning.,” Cell, vol. 105, pp. 421–424, May 2001.

[15] B. Vitre, F. M. Coquelle, C. Heichette, C. Garnier, D. Chretien, and I. Arnal, “Eb1 regulates

microtubule dynamics and tubulin sheet closure in vitro.,” Nat Cell Biol, vol. 10, pp. 415–421,

Apr 2008.

[16] K. A. Blake-Hodek, L. Cassimeris, and T. C. Huﬀaker, “Regulation of microtubule dynamics

by bim1 and bik1, the budding yeast members of the eb1 and clip-170 families of plus-end

tracking proteins.,” Mol Biol Cell, vol. 21, pp. 2013–2023, Jun 2010.

[17] S. P. Maurer, F. J. Fourniol, G. Bohner, C. A. Moores, and T. Surrey, “Ebs recognize a

nucleotide-dependent structural cap at growing microtubule ends.,” Cell, vol. 149, pp. 371–

382, Apr 2012.

[18] S. Honnappa, S. M. Gouveia, A. Weisbrich, F. F. Damberger, N. S. Bhavesh, H. Jawhari,

I. Grigoriev, F. J. A. van Rijssel, R. M. Buey, A. Lawera, I. Jelesarov, F. K. Winkler,

K. Wuthrich, A. Akhmanova, and M. O. Steinmetz, “An eb1-binding motif acts as a mi-

crotubule tip localization signal.,” Cell, vol. 138, pp. 366–376, Jul 2009.

[19] R. Tournebize, A. Popov, K. Kinoshita, A. J. Ashford, S. Rybina, A. Pozniakovsky, T. U.

Mayer, C. E. Walczak, E. Karsenti, and A. A. Hyman, “Control of microtubule dynamics by

48 the antagonistic activities of xmap215 and xkcm1 in xenopus egg extracts.,” Nat Cell Biol,

vol. 2, pp. 13–19, Jan 2000.

[20] G. J. Brouhard, J. H. Stear, T. L. Noetzel, J. Al-Bassam, K. Kinoshita, S. C. Harrison,

J. Howard, and A. A. Hyman, “Xmap215 is a processive microtubule polymerase.,” Cell,

vol. 132, pp. 79–88, Jan 2008.

[21] A. W. Hunter, M. Caplow, D. L. Coy, W. O. Hancock, S. Diez, L. Wordeman, and

J. Howard, “The kinesin-related protein mcak is a microtubule depolymerase that forms an

atp-hydrolyzing complex at microtubule ends.,” Mol Cell, vol. 11, pp. 445–457, Feb 2003.

[22] C. A. Moores, J. Cooper, M. Wagenbach, Y. Ovechkina, L. Wordeman, and R. A. Milligan,

“The role of the kinesin-13 neck in microtubule depolymerization,” Cell Cycle, vol. 5, pp. 1812–

5, Aug 2006.

[23] R. Heald and E. Nogales, “Microtubule dynamics.,” J Cell Sci, vol. 115, pp. 3–4, Jan 2002.

[24] J. Howard and A. A. Hyman, “Dynamics and mechanics of the microtubule plus end.,” Nature,

vol. 422, pp. 753–758, Apr 2003.

[25] C. Wiese and Y. Zheng, “A new function for the gamma-tubulin ring complex as a microtubule

minus-end cap,” Nat Cell Biol, vol. 2, pp. 358–64, Jun 2000.

[26] M. Moritz, M. B. Braunfeld, V. Gu´enebaut,J. Heuser, and D. A. Agard, “Structure of the

gamma-tubulin ring complex: a template for microtubule nucleation,” Nat Cell Biol, vol. 2,

pp. 365–70, Jun 2000.

[27] Y. Zheng, M. L. Wong, B. Alberts, and T. Mitchison, “Nucleation of microtubule assembly by

a gamma-tubulin-containing ring complex,” Nature, vol. 378, pp. 578–83, Dec 1995.

[28] E. T. O’Toole, K. L. McDonald, J. M¨antler, J. R. McIntosh, A. A. Hyman, and T. M¨uller-

Reichert, “Morphologically distinct microtubule ends in the mitotic centrosome of caenorhab-

ditis elegans,” J Cell Biol, vol. 163, pp. 451–6, Nov 2003.

[29] R. Uehara, R.-s. Nozawa, A. Tomioka, S. Petry, R. D. Vale, C. Obuse, and G. Goshima, “The

augmin complex plays a critical role in spindle microtubule generation for mitotic progression

and cytokinesis in human cells,” Proc Natl Acad Sci U S A, vol. 106, pp. 6998–7003, Apr 2009.

49 [30] S. Petry, A. C. Groen, K. Ishihara, T. J. Mitchison, and R. D. Vale, “Branching microtubule

nucleation in xenopus egg extracts mediated by augmin and tpx2,” Cell, vol. 152, pp. 768–77,

Feb 2013.

[31] S.-i. Kawaguchi and Y. Zheng, “Characterization of a drosophila centrosome protein cp309

that shares homology with kendrin and cg-nap,” Mol Biol Cell, vol. 15, pp. 37–45, Jan 2004.

[32] K. M. Ori-McKenney, L. Y. Jan, and Y.-N. Jan, “Golgi outposts shape dendrite morphology

by functioning as sites of acentrosomal microtubule nucleation in neurons,” Neuron, vol. 76,

pp. 921–30, Dec 2012.

[33] A. M. Yvon and P. Wadsworth, “Non-centrosomal microtubule formation and measurement of

minus end microtubule dynamics in a498 cells.,” J Cell Sci, vol. 110 ( Pt 19), pp. 2391–2401,

Oct 1997.

[34] T. J. Keating, J. G. Peloquin, V. I. Rodionov, D. Momcilovic, and G. G. Borisy, “Microtubule

release from the centrosome.,” Proc Natl Acad Sci U S A, vol. 94, pp. 5078–5083, May 1997.

[35] C. M. Waterman-Storer and E. D. Salmon, “Actomyosin-based retrograde ﬂow of microtubules

in the lamella of migrating epithelial cells inﬂuences microtubule dynamic instability and

turnover and is associated with microtubule breakage and treadmilling.,” J Cell Biol, vol. 139,

pp. 417–434, Oct 1997.

[36] V. Rodionov, E. Nadezhdina, and G. Borisy, “Centrosomal control of microtubule dynamics.,”

Proc Natl Acad Sci U S A, vol. 96, pp. 115–120, Jan 1999.

[37] I. A. Vorobjev, V. I. Rodionov, I. V. Maly, and G. G. Borisy, “Contribution of plus and minus

end pathways to microtubule turnover.,” J Cell Sci, vol. 112 ( Pt 14), pp. 2277–2289, Jul 1999.

[38] R. Bacallao, C. Antony, C. Dotti, E. Karsenti, E. H. Stelzer, and K. Simons, “The subcellular

organization of madin-darby canine kidney cells during the formation of a polarized epithe-

lium,” J Cell Biol, vol. 109, pp. 2817–32, Dec 1989.

[39] J. A. Connolly, B. W. Kiosses, and V. I. Kalnins, “Centrioles are lost as embryonic myoblasts

fuse into myotubes in vitro,” Eur J Cell Biol, vol. 39, pp. 341–5, Jan 1986.

[40] A. M. Tassin, B. Maro, and M. Bornens, “Fate of microtubule-organizing centers during myo-

genesis in vitro,” J Cell Biol, vol. 100, pp. 35–46, Jan 1985.

50 [41] D. Signor and J. M. Scholey, “Microtubule-based transport along axons, dendrites and ax-

onemes,” Essays Biochem, vol. 35, pp. 89–102, 2000.

[42] A. C. Horton and M. D. Ehlers, “Dual modes of endoplasmic reticulum-to-golgi transport in

dendrites revealed by live-cell imaging,” J Neurosci, vol. 23, pp. 6188–99, Jul 2003.

[43] P. W. Baas, J. S. Deitch, M. M. Black, and G. A. Banker, “Polarity orientation of microtubules

in hippocampal neurons: uniformity in the axon and nonuniformity in the dendrite,” Proc Natl

Acad Sci U S A, vol. 85, pp. 8335–9, Nov 1988.

[44] P. W. Baas, M. M. Black, and G. A. Banker, “Changes in microtubule polarity orientation

during the development of hippocampal neurons in culture,” J Cell Biol, vol. 109, pp. 3085–94,

Dec 1989.

[45] S. R. Heidemann, J. M. Landers, and M. A. Hamborg, “Polarity orientation of axonal micro-

tubules,” J Cell Biol, vol. 91, pp. 661–5, Dec 1981.

[46] K. S. Burbank, A. C. Groen, Z. E. Perlman, D. S. Fisher, and T. J. Mitchison, “A new

method reveals microtubule minus ends throughout the meiotic spindle,” J Cell Biol, vol. 175,

pp. 369–75, Nov 2006.

[47] G. Goshima, M. Mayer, N. Zhang, N. Stuurman, and R. D. Vale, “Augmin: a protein complex

required for centrosome-independent microtubule generation within the spindle,” J Cell Biol,

vol. 181, pp. 421–9, May 2008.

[48] P. Bieling, I. A. Telley, and T. Surrey, “A minimal midzone protein module controls formation

and length of antiparallel microtubule overlaps,” Cell, vol. 142, pp. 420–32, Aug 2010.

[49] R. Subramanian, E. M. Wilson-Kubalek, C. P. Arthur, M. J. Bick, E. A. Campbell, S. A.

Darst, R. A. Milligan, and T. M. Kapoor, “Insights into antiparallel microtubule crosslinking

by prc1, a conserved nonmotor microtubule binding protein,” Cell, vol. 142, pp. 433–43, Aug

2010.

[50] Y. Kanai, J. Chen, and N. Hirokawa, “Microtubule bundling by tau proteins in vivo: analysis

of functional domains,” EMBO J, vol. 11, pp. 3953–61, Nov 1992.

[51] D. N. Drechsel, A. A. Hyman, M. H. Cobb, and M. W. Kirschner, “Modulation of the dynamic

instability of tubulin assembly by the microtubule-associated protein tau,” Mol Biol Cell,

vol. 3, pp. 1141–54, Oct 1992.

51 [52] R. D. Vale, “The molecular motor toolbox for intracellular transport,” Cell, vol. 112, pp. 467–

80, Feb 2003.

[53] M. A. Welte, “Bidirectional transport along microtubules,” Curr Biol, vol. 14, pp. R525–37,

Jul 2004.

[54] J. T. Yang, R. A. Laymon, and L. S. Goldstein, “A three-domain structure of kinesin heavy

chain revealed by dna sequence and microtubule binding analyses,” Cell, vol. 56, pp. 879–89,

Mar 1989.

[55] N. Hirokawa, K. K. Pﬁster, H. Yorifuji, M. C. Wagner, S. T. Brady, and G. S. Bloom, “Sub-

molecular domains of bovine brain kinesin identiﬁed by electron microscopy and monoclonal

antibody decoration,” Cell, vol. 56, pp. 867–78, Mar 1989.

[56] G. S. Bloom, M. C. Wagner, K. K. Pﬁster, and S. T. Brady, “Native structure and physical

properties of bovine brain kinesin and identiﬁcation of the atp-binding subunit polypeptide,”

Biochemistry, vol. 27, pp. 3409–16, May 1988.

[57] D. J. Sharp, K. L. McDonald, H. M. Brown, H. J. Matthies, C. Walczak, R. D. Vale, T. J.

Mitchison, and J. M. Scholey, “The bipolar kinesin, klp61f, cross-links microtubules within

interpolar microtubule bundles of drosophila embryonic mitotic spindles,” J Cell Biol, vol. 144,

pp. 125–38, Jan 1999.

[58] L. C. Kapitein, E. J. G. Peterman, B. H. Kwok, J. H. Kim, T. M. Kapoor, and C. F. Schmidt,

“The bipolar mitotic kinesin eg5 moves on both microtubules that it crosslinks,” Nature,

vol. 435, pp. 114–8, May 2005.

[59] L. Wordeman and T. J. Mitchison, “Identiﬁcation and partial characterization of mitotic

centromere-associated kinesin, a kinesin-related protein that associates with centromeres dur-

ing mitosis,” J Cell Biol, vol. 128, pp. 95–104, Jan 1995.

[60] A. W. Hunter, M. Caplow, D. L. Coy, W. O. Hancock, S. Diez, L. Wordeman, and

J. Howard, “The kinesin-related protein mcak is a microtubule depolymerase that forms an

atp-hydrolyzing complex at microtubule ends,” Mol Cell, vol. 11, pp. 445–57, Feb 2003.

[61] R. J. McKenney, W. Huynh, M. E. Tanenbaum, G. Bhabha, and R. D. Vale, “Activation of

cytoplasmic dynein motility by dynactin-cargo adapter complexes,” Science, vol. 345, pp. 337–

41, Jul 2014.

52 [62] G. Goshima, R. Wollman, S. S. Goodwin, N. Zhang, J. M. Scholey, R. D. Vale, and N. Stuur-

man, “Genes required for mitotic spindle assembly in drosophila s2 cells.,” Science, vol. 316,

pp. 417–421, Apr 2007.

[63] S. S. Goodwin and R. D. Vale, “Patronin regulates the microtubule network by protecting

microtubule minus ends.,” Cell, vol. 143, pp. 263–274, Oct 2010.

[64] H. Wang, I. Brust-Mascher, G. Civelekoglu-Scholey, and J. M. Scholey, “Patronin mediates

a switch from kinesin-13-dependent poleward ﬂux to anaphase b spindle elongation.,” J Cell

Biol, vol. 203, pp. 35–46, Oct 2013.

[65] W. Meng, Y. Mushika, T. Ichii, and M. Takeichi, “Anchorage of microtubule minus ends to

adherens junctions regulates epithelial cell-cell contacts.,” Cell, vol. 135, pp. 948–959, Nov

2008.

[66] A. J. Baines, P. A. Bignone, M. D. A. King, A. M. Maggs, P. M. Bennett, J. C. Pinder, and

G. W. Phillips, “The ckk domain (duf1781) binds microtubules and deﬁnes the camsap/ssp4

family of animal proteins.,” Mol Biol Evol, vol. 26, pp. 2005–2014, Sep 2009.

[67] M. N. Shahbazi, D. Megias, C. Epifano, A. Akhmanova, G. G. Gundersen, E. Fuchs, and

M. Perez-Moreno, “Clasp2 interacts with p120-catenin and governs microtubule dynamics at

adherens junctions.,” J Cell Biol, vol. 203, pp. 1043–1061, Dec 2013.

[68] M. D. A. King, G. W. Phillips, P. A. Bignone, N. V. L. Hayes, J. C. Pinder, and A. J.

Baines, “A conserved sequence in calmodulin regulated spectrin-associated protein 1 links its

interaction with spectrin and calmodulin to neurite outgrowth,” Journal of Neurochemistry,

vol. 128, no. 3, pp. 391–402, 2014.

[69] N. Tanaka, W. Meng, S. Nagae, and M. Takeichi, “Nezha/camsap3 and camsap2 cooperate in

epithelial-speciﬁc organization of noncentrosomal microtubules.,” Proc Natl Acad Sci U S A,

vol. 109, pp. 20029–20034, Dec 2012.

[70] S. Nagae, W. Meng, and M. Takeichi, “Non-centrosomal microtubules regulate f-actin orga-

nization through the suppression of gef-h1 activity.,” Genes Cells, vol. 18, pp. 387–396, May

2013.

[71] J. D. Marcette, J. J. Chen, and M. L. Nonet, “The caenorhabditis elegans microtubule minus-

end binding homolog ptrn-1 stabilizes synapses and neurites,” eLife, vol. 3, 2014.

53 [72] C. E. Richardson, K. A. Spilker, J. G. Cueva, J. Perrino, M. B. Goodman, and K. Shen,

“Ptrn-1, a microtubule minus end-binding camsap homolog, promotes microtubule function in

caenorhabditis elegans neurons,” eLife, vol. 3, 2014.

[73] J. M. Kollman, J. K. Polka, A. Zelter, T. N. Davis, and D. A. Agard, “Microtubule nucleating

gamma-tusc assembles structures with 13-fold microtubule-like symmetry.,” Nature, vol. 466,

pp. 879–882, Aug 2010.

[74] M. Gimona, K. Djinovic-Carugo, W. J. Kranewitter, and S. J. Winder, “Functional plasticity

of ch domains.,” FEBS Lett, vol. 513, pp. 98–106, Feb 2002.

[75] V. Johnson, P. Ayaz, P. Huddleston, and L. M. Rice, “Design, overexpression, and puriﬁcation

of polymerization-blocked yeast -tubulin mutants,” Biochemistry, vol. 50, pp. 8636–44, Oct

2011.

[76] M. Sirajuddin, L. M. Rice, and R. D. Vale, “Regulation of microtubule motors by tubulin

isotypes and post-translational modiﬁcations,” Nat Cell Biol, vol. 16, pp. 335–44, Apr 2014.

[77] T. J. P. Hubbard, B. L. Aken, S. Ayling, B. Ballester, K. Beal, E. Bragin, S. Brent, Y. Chen,

P. Clapham, L. Clarke, G. Coates, S. Fairley, S. Fitzgerald, J. Fernandez-Banet, L. Gordon,

S. Graf, S. Haider, M. Hammond, R. Holland, K. Howe, A. Jenkinson, N. Johnson, A. Ka-

hari, D. Keefe, S. Keenan, R. Kinsella, F. Kokocinski, E. Kulesha, D. Lawson, I. Longden,

K. Megy, P. Meidl, B. Overduin, A. Parker, B. Pritchard, D. Rios, M. Schuster, G. Slater,

D. Smedley, W. Spooner, G. Spudich, S. Trevanion, A. Vilella, J. Vogel, S. White, S. Wilder,

A. Zadissa, E. Birney, F. Cunningham, V. Curwen, R. Durbin, X. M. Fernandez-Suarez, J. Her-

rero, A. Kasprzyk, G. Proctor, J. Smith, S. Searle, and P. Flicek, “Ensembl 2009.,” Nucleic

Acids Res, vol. 37, pp. D690–7, Jan 2009.

[78] A. J. Vilella, J. Severin, A. Ureta-Vidal, L. Heng, R. Durbin, and E. Birney, “Ensemblcom-

para genetrees: Complete, duplication-aware phylogenetic trees in vertebrates.,” Genome Res,

vol. 19, pp. 327–335, Feb 2009.

[79] M. Castoldi and A. V. Popov, “Puriﬁcation of brain tubulin through two cycles of

polymerization-depolymerization in a high-molarity buﬀer.,” Protein Expr Purif, vol. 32,

pp. 83–88, Nov 2003.

[80] A. Gennerich and S. L. Reck-Peterson, “Probing the force generation and stepping behavior

of cytoplasmic dynein.,” Methods Mol Biol, vol. 783, pp. 63–80, 2011.

54 [81] J. Schindelin, I. Arganda-Carreras, E. Frise, V. Kaynig, M. Longair, T. Pietzsch, S. Preibisch,

C. Rueden, S. Saalfeld, B. Schmid, J.-Y. Tinevez, D. J. White, V. Hartenstein, K. Eliceiri,

P. Tomancak, and A. Cardona, “Fiji: an open-source platform for biological-image analysis.,”

Nat Methods, vol. 9, pp. 676–682, Jul 2012.

[82] A. Edelstein, N. Amodaj, K. Hoover, R. Vale, and N. Stuurman, “Computer control of mi-

croscopes using micromanager.,” Curr Protoc Mol Biol, vol. Chapter 14, p. Unit14.20, Oct

2010.

[83] C. Gell, V. Bormuth, G. J. Brouhard, D. N. Cohen, S. Diez, C. T. Friel, J. Helenius,

B. Nitzsche, H. Petzold, J. Ribbe, E. Schaﬀer, J. H. Stear, A. Trushko, V. Varga, P. O.

Widlund, M. Zanic, and J. Howard, “Microtubule dynamics reconstituted in vitro and imaged

by single-molecule ﬂuorescence microscopy.,” Methods Cell Biol, vol. 95, pp. 221–245, 2010.

[84] C. E. Aitken, R. A. Marshall, and J. D. Puglisi, “An oxygen scavenging system for improvement

of dye stability in single-molecule ﬂuorescence experiments.,” Biophys J, vol. 94, pp. 1826–1835,

Mar 2008.

[85] L. Peris, M. Wagenbach, L. Lafanechere, J. Brocard, A. T. Moore, F. Kozielski, D. Job,

L. Wordeman, and A. Andrieux, “Motor-dependent microtubule disassembly driven by tubulin

tyrosination.,” J Cell Biol, vol. 185, pp. 1159–1166, Jun 2009.

[86] K. Jiang, S. Hua, R. Mohan, I. Grigoriev, K. W. Yau, Q. Liu, E. A. Katrukha, A. F. M.

Altelaar, A. J. R. Heck, C. C. Hoogenraad, and A. Akhmanova, “Microtubule minus-end

stabilization by polymerization-driven camsap deposition.,” Dev Cell, vol. 28, pp. 295–309,

Feb 2014.

[87] M. Kirschner and T. Mitchison, “Beyond self-assembly: from microtubules to morphogenesis,”

Cell, vol. 45, pp. 329–42, May 1986.

[88] D. A. Starr, R. Saﬀery, Z. Li, A. E. Simpson, K. H. Choo, T. J. Yen, and M. L. Goldberg,

“Hzwint-1, a novel human kinetochore component that interacts with hzw10,” J Cell Sci,

vol. 113 ( Pt 11), pp. 1939–50, Jun 2000.

[89] G. J. P. L. Kops, Y. Kim, B. A. A. Weaver, Y. Mao, I. McLeod, J. R. Yates, 3rd, M. Tagaya,

and D. W. Cleveland, “Zw10 links mitotic checkpoint signaling to the structural kinetochore,”

J Cell Biol, vol. 169, pp. 49–60, Apr 2005.

55 [90] C. Ciferri, J. De Luca, S. Monzani, K. J. Ferrari, D. Ristic, C. Wyman, H. Stark, J. Kilmartin,

E. D. Salmon, and A. Musacchio, “Architecture of the human ndc80-hec1 complex, a critical

constituent of the outer kinetochore,” J Biol Chem, vol. 280, pp. 29088–95, Aug 2005.

[91] R. R. Wei, J. Al-Bassam, and S. C. Harrison, “The ndc80/hec1 complex is a contact point for

kinetochore-microtubule attachment,” Nat Struct Mol Biol, vol. 14, pp. 54–9, Jan 2007.

[92] S. L. Kline-Smith, S. Sandall, and A. Desai, “Kinetochore-spindle microtubule interactions

during mitosis,” Curr Opin Cell Biol, vol. 17, pp. 35–46, Feb 2005.

[93] C. A. Moores and R. A. Milligan, “Lucky 13-microtubule depolymerisation by kinesin-13

motors.,” J Cell Sci, vol. 119, pp. 3905–3913, Oct 2006.

[94] F. Gergely, V. M. Draviam, and J. W. Raﬀ, “The ch-tog/xmap215 protein is essential for

spindle pole organization in human somatic cells,” Genes Dev, vol. 17, pp. 336–41, Feb 2003.

[95] B. M. Paschal, H. S. Shpetner, and R. B. Vallee, “Map 1c is a microtubule-activated atpase

which translocates microtubules in vitro and has dynein-like properties,” J Cell Biol, vol. 105,

pp. 1273–82, Sep 1987.

[96] E. A. Vaisberg, P. M. Grissom, and J. R. McIntosh, “Mammalian cells express three distinct

dynein heavy chains that are localized to diﬀerent cytoplasmic organelles,” J Cell Biol, vol. 133,

pp. 831–42, May 1996.

[97] T. Gaglio, M. A. Dionne, and D. A. Compton, “Mitotic spindle poles are organized by struc-

tural and motor proteins in addition to centrosomes,” J Cell Biol, vol. 138, pp. 1055–66, Sep

1997.

[98] E. A. Vaisberg, M. P. Koonce, and J. R. McIntosh, “Cytoplasmic dynein plays a role in

mammalian mitotic spindle formation,” J Cell Biol, vol. 123, pp. 849–58, Nov 1993.

[99] D. A. Starr, B. C. Williams, T. S. Hays, and M. L. Goldberg, “Zw10 helps recruit dynactin

and dynein to the kinetochore,” J Cell Biol, vol. 142, pp. 763–74, Aug 1998.

[100] J. Takagi, T. Itabashi, K. Suzuki, and S. Ishiwata, “Chromosome position at the spindle

equator is regulated by chromokinesin and a bipolar microtubule array,” Sci Rep, vol. 3,

p. 2808, 2013.

56 [101] J. Turner, R. Anderson, J. Guo, C. Beraud, R. Fletterick, and R. Sakowicz, “Crystal structure

of the mitotic spindle kinesin eg5 reveals a novel conformation of the neck-linker,” J Biol

Chem, vol. 276, pp. 25496–502, Jul 2001.

[102] H. Drechsler, T. McHugh, M. R. Singleton, N. J. Carter, and A. D. McAinsh, “The kinesin-12

kif15 is a processive track-switching tetramer,” Elife (Cambridge), vol. 3, p. e01724, 2014.

[103] A. Blangy, H. A. Lane, P. d’H´erin,M. Harper, M. Kress, and E. A. Nigg, “Phosphorylation

by p34cdc2 regulates spindle association of human eg5, a kinesin-related motor essential for

bipolar spindle formation in vivo,” Cell, vol. 83, pp. 1159–69, Dec 1995.

[104] K. E. Sawin, K. LeGuellec, M. Philippe, and T. J. Mitchison, “Mitotic spindle organization

by a plus-end-directed microtubule motor,” Nature, vol. 359, pp. 540–3, Oct 1992.

[105] K. E. Sawin and T. J. Mitchison, “Mutations in the kinesin-like protein eg5 disrupting local-

ization to the mitotic spindle,” Proc Natl Acad Sci U S A, vol. 92, pp. 4289–93, May 1995.

[106] R. Sakowicz, J. T. Finer, C. Beraud, A. Crompton, E. Lewis, A. Fritsch, Y. Lee, J. Mak,

R. Moody, R. Turincio, J. C. Chabala, P. Gonzales, S. Roth, S. Weitman, and K. W. Wood,

“Antitumor activity of a kinesin inhibitor,” Cancer Res, vol. 64, pp. 3276–80, May 2004.

[107] J. W. Purcell, J. Davis, M. Reddy, S. Martin, K. Samayoa, H. Vo, K. Thomsen, P. Bean,

W. L. Kuo, S. Ziyad, J. Billig, H. S. Feiler, J. W. Gray, K. W. Wood, and S. Cases, “Activity

of the kinesin spindle protein inhibitor ispinesib (sb-715992) in models of breast cancer,” Clin

Cancer Res, vol. 16, pp. 566–76, Jan 2010.

[108] M. E. Tanenbaum, L. Mac˚urek,A. Janssen, E. F. Geers, M. Alvarez-Fern´andez,and R. H.

Medema, “Kif15 cooperates with eg5 to promote bipolar spindle assembly,” Curr Biol, vol. 19,

pp. 1703–11, Nov 2009.

[109] D. B. Hans Lineweaver, “The determination of enzyme dissociation constants,” J. Am. Chem.

Soc., vol. 56, pp. 658–666, March 1934.

[110] K. W. Yau, S. F. B. van Beuningen, I. Cunha-Ferreira, B. M. C. Cloin, E. Y. van Bat-

tum, L. Will, P. Sch¨atzle,R. P. Tas, J. van Krugten, E. A. Katrukha, K. Jiang, P. S. Wulf,

M. Mikhaylova, M. Harterink, R. J. Pasterkamp, A. Akhmanova, L. C. Kapitein, and C. C.

Hoogenraad, “Microtubule minus-end binding protein camsap2 controls axon speciﬁcation and

dendrite development,” Neuron, vol. 82, pp. 1058–73, Jun 2014.

57 [111] C. L. Thompson, L. Ng, V. Menon, S. Martinez, C.-K. Lee, K. Glattfelder, S. M. Sunkin,

A. Henry, C. Lau, C. Dang, R. Garcia-Lopez, A. Martinez-Ferre, A. Pombero, J. L. R. Ruben-

stein, W. B. Wakeman, J. Hohmann, N. Dee, A. J. Sodt, R. Young, K. Smith, T.-N. Nguyen,

J. Kidney, L. Kuan, A. Jeromin, A. Kaykas, J. Miller, D. Page, G. Orta, A. Bernard, Z. Riley,

S. Smith, P. Wohnoutka, M. J. Hawrylycz, L. Puelles, and A. R. Jones, “A high-resolution spa-

tiotemporal atlas of gene expression of the developing mouse brain,” Neuron, vol. 83, pp. 309–

23, Jul 2014.

[112] R. Petryszak, T. Burdett, B. Fiorelli, N. A. Fonseca, M. Gonzalez-Porta, E. Hastings, W. Hu-

ber, S. Jupp, M. Keays, N. Kryvych, J. McMurry, J. C. Marioni, J. Malone, K. Megy, G. Rus-

tici, A. Y. Tang, J. Taubert, E. Williams, O. Mannion, H. E. Parkinson, and A. Brazma,

“Expression atlas update–a database of gene and transcript expression from microarray- and

sequencing-based functional genomics experiments,” Nucleic Acids Res, vol. 42, pp. D926–32,

Jan 2014.

[113] M. C. Hendershott and R. D. Vale, “Regulation of microtubule minus-end dynamics by cam-

saps and patronin,” Proc Natl Acad Sci U S A, vol. 111, pp. 5860–5, Apr 2014.

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