Molecular Dissection of Nde1's Role in Mitosis

Molecular Dissection of Nde1’s Role in Mitosis

Caitlin Lazar Wynne

Submitted in partial fulfillment of the requirements for the degree of Doctor of Philosophy under the Executive Committee of the Graduate School of Arts and Sciences

COLUMBIA UNIVERSITY

2016

ABSTRACT

Molecular dissection of Nde1’s role in mitosis

Caitlin Lazar Wynne

Upon entry into G2 and mitosis (G2/M), dynein dissociates from its interphase cargos and forms mitotic-specific interactions that direct dynein to the nuclear envelope, cell-cortex, kinetochores, and spindle poles to ensure equal segregation of genetic

material to the two daughter cells. Although the need for precise regulation of dynein’s

activity during mitosis is clear, questions remain about the mechanisms that govern the cell-cycle dependent dynein interactions. Frequently dynein cofactors provide platforms for regulating dynein activity either by directing dynein to specific sites of action or by tuning the motor activity of the dynein motor. In particular the dynein cofactor Nde1 may play a key role in defining dynein’s mitotic activity. During interphase, Nde1 is involved in the dynein-dependent processes of Golgi positioning and minus-end directed lysosome transport (Lam et al., 2009; Yi et al., 2011), but as the cell progresses into

G2/M, Nde1 adopts mitotic specific interactions at the nuclear envelope and kinetochores. It is unknown how Nde1’s cell-cycle specific localization is regulated and how, if at all, Nde1 is ultimately able to influence dynein’s recruitment and activity at each of these sites. One candidate is cell-cycle specific phosphorylation of Nde1 by a

G2/mitotic specific kinase, cyclinB/Cdk1 (Alkurayaet al. 2011). To study the potential function of the phosphorylation by Cdk1, we assayed the localization of GFP Cdk1Nde1 phospho-mimetic and phospho-mutant constructs at the NE and kinetochores. We demonstrate Cdk1 phosphorylation of Nde1 is required for Nde1 localization to both the

NE and to the kinetochore, and also the phosphorylation of Nde1 directly activates

physical interactions between Nde1 and its nuclear envelope and the kinetochore-binding

partner, CENP-F. Furthermore, physiological studies of Nde1 phosphorylation constructs

show that over-expression of GFP Nde1 phospho-mutant causes a significant delay in time from NEBD to anaphase onset, specifically demonstrating a late prometaphase/metaphase arrest. Therefore, we conclude Cdk1 phosphorylation of Nde1 not only regulates its localization to the nuclear envelope and kinetochore but also plays an important functional role in Nde1’s mitotic activity in vivo.

In addition to understanding how the cell cycle specific activity of Nde1 is regulated, to fully comprehend how dynein functions during mitosis it is necessary to understand how Nde1 is able to modulate dynein’s activity. Nde1 is typically believed to act as a bridge between dynein and specific cellular cargo by physically interacting both with the cargo and dynein/Lis1 to specify the sites of dynein’s activity. Therefore, to understand how Nde1 functions with Lis1 and dynein during mitosis, we created point mutations in the N-terminal coiled-coil domain that specifically disrupted either the

Nde1-Lis1 interaction or the Nde1-dynein interaction. We find that disrupting the Nde1- dynein interaction has more severe phenotypic effects compared to disrupting the Nde1-

Lis1 interaction: expression of GFP Nde1 del dynein mutant caused a significant delay in anaphase onset while GFP Nde1 del Lis1 only caused a slight increase in cell cycle duration before anaphase onset. Phenotypic analysis suggests that the effects of abolishing the Nde1-dynein interaction on mitotic progression may be due to defects in maintaining kinetochore-microtubule stability during metaphase. Nde1 plays a role in this dynein-dependent mitotic activity through recruitment of a subfraction of dynein to the kinetochore by Nde1’s coiled-coil domain. While the phenotypic effect of removing

the Lis1-Nde1 interaction is less severe than removing the dynein-Nde1 interaction, the interaction between Lis1 and Nde1 plays an important role in Nde1’s mitotic behavior as it is affects Nde1’s localization at the kinetochore, specifically by influencing Nde’1 interaction with its kinetochore recruitment partner, CENP-F.

The entirety of this work demonstrates that Nde1 acts as a link between cellular cargo and dynein behavior as phospho-regulation of Nde1 throughout the cell cycle allows Nde1’s to interact with unique mitotic cargoes and influence the recruitment and activity of dynein at the kinetochore.

Table of Contents

List of Figures and Tables iii

Acknowledgements v

Dedication viii

Chapter 1: Cell-Cycle Regulation of Cytoplasmic Dynein

1. Introduction 1

2. Cytoplasmic Dynein 2

A. Motor Activity 3 Dynein Motor Domain 3 Non-motor domain regulation of dynein motor activity 8 B. Cargo-Binding 10 Light intermediate chains/intermediate chains 10 Dynactin 15 Nde1/Ndel1 16

3. Dynein’s role in G2 and mitosis 18

4. Cell-cycle regulation of dynein and dynein accessory proteins 26

A. Dynein Complex 26 B. Mitotic specific cargo interactions 27 C. Cell-cycle regulated phosphorylation 29

5. Conclusion 35

Chapter 2: Cdk1 phosphorylation of Nde1 activates it interaction with Nuclear Envelope and Kinetochore binding partner, CENP-F and regulates mitotic function

1. Introduction 37 2. Results 39

Cdk1 phosphorylated Nde1 and phospho-mimetic Nde1 localize to known Nde1 mitotic sites 39 Nde1 phosphorylation increases interaction with its kinetochore-binding partner, CENP-F 50 Role of Nde1 phosphorylation in mitosis 54 Role of Nde1 in Anaphase A 61

3. Discussion 65

Role of Nde1 phosphorylation in G2-M Phase Targeting and CENP-F binding 66 Functional Analysis 68 Post-metaphase roles of Nde1 72

Chapter 3: Structure-Function approach to elucidating Nde1’s function during mitosis

1. Introduction 74 2. Results 76

Characterization of the Nde1-Dynein and Nde1-Lis1 interactions 76 Importance of Nde1’s interaction with dynein or Lis1 in mitotic progression 81 Investigation of Metaphase defects caused by lack of Nde1-Dynein interaction 84 Nde1 recruits dynein directly to the kinetochore through its coiled-coil domain 90 Nde1-Lis1 interaction is required for Nde1 localization to the kinetochore 91

3. Discussion 98

Importance of the N-terminal domain of Nde1 for its function with dynein 98 Role of the Nde1-lis1 interaction in mitosis 101

Chapter 4: Conclusion 104

Chapter 5: Experimental Procedures 114

References 126

Appendix I: Additional Work 150

Appendix II: Additional Work 152

LIST OF FIGURES AND TABLES

Chapter 1: Cell-Cycle Regulation of Cytoplasmic Dynein

Figure 1-1. Representation of cytoplasmic dynein subunits and dynein adaptor proteins 3

Figure 1-2. Cytoplasmic dynein motor domain 5

Figure 1-3. Cytoplasmic dynein’s mechanochemical cycle 7

Table 1-1. Summary of dynein cargo interactions 11

Figure 1-4. Cell-Cycle specific dynein recruitment pathways 19

Figure 1-5. Cytoplasmic dynein’s mitotic localization 20

Chapter 2: Cdk1 phosphorylation of Nde1 activates it interaction with Nuclear Envelope and Kinetochore binding partner, CENP-F and regulates mitotic function

Figure 2-1. Examination of endogenous Nde1/Ndel1 localization at the nuclear envelope in G2 and the kinetochore during mitosis 41

Figure 2-2. Localization of Nde1 phospho-mimetic and phospho-mutant constructs at the NE in G2. 43

Figure 2-3. Localization of GFP wt Nde1, phospho-mimetic Nde1 and phospho-mutant Nde1 at kinetochores in mitotic cells 45

Figure 2-4. Endogenous Cdk1 phosphorylated Nde1 localizes to the kinetochore 47

Figure 2-5. Effects of Cdk1 phosphorylation on Nde1 kinetochore localization 49

Figure 2-6. CENP-F siRNA decreases endogenous Nde1/el signal from kinetochores 51

Figure 2-7. Biochemical analysis of Cdk1 effects on the Nde1-CENP-F interaction 53

Figure 2-8. Effects of over expression of GFP Nde1 phosphorylation constructs on mitotic progression 56

Figure 2-9. Rescue of Nde1 siRNA mitotic progression defects with wt, phospho-mimetic or phospho-mutant Nde1 58

iii

Figure 2-10. Role of Nde1 in Anaphase 62

Figure 2-11. Summary of the mitotic localization of Cdk1 phosphorylated Nde1 at the nuclear envelope and kinetochore 65

Chapter 3: Structure-Function approach to elucidating Nde1’s function at the kinetochore

Figure 3-1. Biochemical characterization of Nde1’s interaction with dynein 78

Figure 3-2. Characterization of Nde1 mutant constructs used to study the mitotic role of Nde1’s interaction with Lis1 or dynein 80

Figure 3-3. Effects of over expression of Nde1 mutants on mitotic progression 83

Figure 3-4. Rescue of Nde1 siRNA mitotic progression defects with Nde1 mutant constructs 85

Figure 3-5. Role of Nde1-dynein interaction on formation of stable KT-MT interactions 87

Figure 3-6. Characterization of metaphase defects caused by loss of Nde1 or the Nde1-dynein interaction 89

Figure 3-7. Nde1 recruits dynein to the kinetochore via its N-terminal coiled-coil 91

Figure 3-8. Requirement of Lis1 and Nde1 for localization of each to the kinetochore 93

Figure 3-9. Localization of GFP Nde1 del Lis1 mutant to the kinetochore 95

Figure 3-10. Biochemical analysis of the effect of Lis1 binding mutant on Nde1’s interaction with CENP-F 97

Appendix I

Sup. Figure 1.1. Characterization of WT-MD and Delta CT-MD purifications 151

Appendix II

Sup. Figure 2.1. Single molecule studies of C-terminus Nde1 truncations 153

ACKNOWLEDGMENTS

The past seven years of my graduate school career at Columbia University have been filled with interesting scientific conversations with colleagues in the halls of the

15th floor, discussions of data in Richard’s office and laughter with fellow lab-mates during lunch breaks in the computer room. It has been an important period of personal intellectual growth that has challenged me to become a better scientist, student and person. There are a great number of people that have had a significant role in shaping the positive experience and scientific progress I have experienced during graduate school.

First, I would like to thank my mentor, Richard Vallee for his guidance and support during my graduate school experience. Richard has been an invaluable resource for my progression in my scientific learning. I not only learned to be an independent scientist but how to think critically.

The members of the Vallee lab have fostered a supportive lab environment that allowed for interesting scientific discussions and collaborative work. First, I would specifically like to thank Dan Hu for his friendship throughout my entire six years together in the Vallee Lab. Although our scientific interests are not aligned, he has always taken the time to discuss experiments and ideas that have shaped views on my project. I want to thank Aurelie for her kindness and support during this last year. I would like to thank Tiago for pushing me to try the new spinning confocal microscope, which has revolutionized my imaging and data acquisition during the last year of my

PhD. I want to thank Nupoor for her positive attitude and kindness she brings to lab everyday. Additionally, Alex, Jie, and Dave have all been amazing colleagues that have contributed to the environment of the Vallee lab.

I want to thank the past members of the Vallee lab that had an important role in

the foundation of my graduate school experience. I want to thank Sarah for teaching me

biochemistry and for the years of interesting scientific conversations that shaped the

direction of my project. I want to thank Julian for his example of how to design

experiments and for his friendship and willingness to take 3:00 pm diet coke breaks.

Finally I want to thank Shahrnaz for being a supportive bay-mate during the first five years of graduate school.

I want to thank the Cell Biology program, especially the directors Ron Liem and

Donna Farber for accepting me into the graduate program and for their continued guidance throughout my graduate experience. I want to thank Zaia Sivo for her support since the first day I arrived in New York. I am forever grateful for the time she took to listen to my concerns.

I want to thank my committee, Gregg Gundersen, Yungui Mao and Ulrich Hengst for their continued guidance and support during my committee meetings. Without their advice and expertise, I would not have progressed in my project. Additionally, I would like to thank Professor Chloe Bulinski for joining the thesis defense committee. I want to

thank the entire Mao lab for their continued help and advice in troubleshooting various

experiments.

I want to thank the first friends I met a Columbia, Colleen, Sarah and Jayne, for

their willingness to take crazy photo-booth pictures and for the fun nights that allowed for

an escape from the lab. There are no greater people I would have rather shared my entire

20s with. We have come a long way and it has been an honor to be able to share many

great experiences and life events with you. Also, I want to thank Eileen, Shahrnaz and

Punita for the Antika dinners with nutella pizza and for being helpful resources for what to expect during the final months of graduate school. I am truly grateful for the friendship and support during my entire PhD.

I want to thank Dr. Kevin Vaughan for his mentorship and guidance. I would not be working towards a PhD at Columbia University without his mentorship and friendship.

I would also like to thank all my friends and my family for their love, friendship and understanding during the past six years. I want to thank Pat, Louise and Maureen for their support and encouragement throughout the past seven years. I want to thank Anna for providing pep talks and motivation, sometimes in the form of emojis. I want to thank

Daniel and John for being supportive and understanding brothers. It has been a pleasure to be able to share this experience with you and commiserate together about graduate school. I want to thank my parents, Eileen and George, for teaching me the importance of education, for pushing me to achieve my goal of getting my PhD and for supporting me every step of the way.

And last, I want to give a heartfelt thank you to my husband Matt for his never- ending support and unwavering belief I could accomplish my goal. I could not ask for a more supportive partner during this graduate school journey. From coming to lab with me at 5:00 in the morning to taking the subway home with me late at night, you have gone above and beyond to help me in any way you could.

vii

DEDICATION

For Matt.

Forever grateful for your love, support and understanding

I am only where I am today because of you.

viii

Chapter 1: Cell-Cycle Regulation of Cytoplasmic Dynein

1. Introduction to microtubule motors and their adaptation to cargos

Directed transport of diverse cellular cargo including vesicles, mRNA, virus, organelles and proteins along the microtubule cytoskeleton network requires the regulated action of two classes of microtubule motor proteins: dynein and kinesin.

Dynein is responsible for minus-end directed transport, while kinesin acts as the plus-end directed motor. By targeting these motors to specific cargo and locations, movement throughout the cell can be carried out in a highly precise manner. A notable example of the requirement for highly regulated motor activity is in mitosis when the microtubule network reorganizes to form the mitotic spindle and microtubule motors are required to adopt specialized mitotic interactions and function at unique mitotic structures in order to ensure the correct segregation of genetic material to two daughter cells.

While the activity of both kinesin and dynein is regulated to carry out specific cellular functions, the mechanism by which the regulation is achieved differs. Kinesin activity is mainly regulated by differential isoform usage; there are 15 kinesin families encompassing ~45 different kinesins and individual kinesins have unique cargo interactions to dictates cellular functions (Hirokawa et al., 2009). For example, cell-cycle specific kinesins—Eg5, MCAK or CENP-E—contribute to kinesin’s mitotic specific activity. Unlike kinesin, only one complex, cytoplasmic dynein is responsible for all dynein’s cytoplasmic cellular functions. Instead several adaptor proteins are required for facilitating cargo interactions and influencing dynein’s motor properties. Given the disconnect between the diverse cellular processes dynein carries out and the single dynein complex, a main focus in cell biology is understanding the regulation of dynein and how

the single dynein complex is able to carry out diverse cellular function. The focus of this chapter is an introduction to the cell-cycle regulation of cytoplasmic dynein and how both intra and inter molecular regulation of dynein allows it to adopt unique cargo interactions during G2 and mitosis to target dynein to mitotic specific sites.

2. Cytoplasmic Dynein

Cytoplasmic Dynein is a ~1.6 MDa protein complex comprised of heavy chains, light-intermediate chains (LICs), intermediate chain (ICs) and lights chains (LCs) that all function together to modulate the enzymatic activity of or facilitate a cargo interaction for dynein’s complex and numerous cellular function. Classically, the dynein subunits have been divided into subunits responsible for motor activity, with the motor domain of the heavy chain mainly responsible for force production, and cargo binding by the LCs, LICS and ICS. Because there is only one dynein responsible for the numerous cytoplasmic functions, dynein requires the interaction with additional class of proteins, the dynein adaptor proteins. These are dynactin, Nde1 and Ndel1, Lis1. Dynactin (Dynein-

Activator) is a large 1.2 mDA complex comprised of 11 different proteins forming the complex essential for most dynein cellular functions. Lis1 was first identified as the causal gene for the human cortical developmental disease, lissencephaly, (Reiner et al.,

1999), but later characterized to be a regulator of dynein motor activity. Nde1 and Ndel1 interact with the both Lis1 and dynein to influence dynein’s persistent force production and recruitment to cellular environments (McKenney et al., 2010; Stehman et al., 2007).

It is the orchestration of subunits within the dynein complex as well as binding of accessory proteins that contribute to dynein’s motor activity as well as cargo-binding capability (Figure 1.1)

Figure 1.1. Representation of cytoplasmic dynein subunits and dynein adaptor proteins (modified from Jaarsma and Hoogenrad, 2015) A. The cytoplasmic dynein complex is comprised of the heavy chains, light intermediate chains (LIC1 and LIC2), intermediate chain (IC) and light chains (LC8, TcTex and LC7). Dynein adaptor proteins, dynactin and Nde1/Ndel1, bind to the intermediate chain of dynein to contribute to cytoplasmic dynein’s cellular functions. Lis1 interacts with both Nde1/Ndel1 and the motor domain of dynein. Each subunit of the dynein complex and dynein adaptor proteins are required to modulate dynein’s motor activity or interaction with cellular cargos.

A. Motor Activity

Dynein motor domain and cross-bridge cycle

The two motor domains within the dynein heavy chains generate the force

required for the translocation of cytoplasmic dynein towards the minus end of the

microtubule. Structurally, the motor domain is an asymmetrical planar ring of 6 AAA

domains, AAA1-AAA6, arranged around a central pore. Nucleotide (ATP) binds and is hydrolyzed within the AAA ring. Although AAA1-AAA4 have the ability to bind nucleotide (Koonce et al., 2004), the main site of ATP hydrolysis essential for dynein motility is AAA1 (Reck-Peterson et al., 2004; Kon et al., 2004). The nucleotide binding ability of AAA3 is required to regulate AAA1 nucleotide binding ability (Dewitt et al.,

2015; Nicholas et al., 2015). The final two AAA domains of the motor domain, AAA5 and AAA6, lack nucleotide-binding ability, but add structural rigidity to the ring (Bhabha et al., 2014).

Three appendages emerge from the AAA ring. From AAA4/5 of the AAA ring, the stalk of the motor domain emerges, consisting of a coiled-coil domain made up of two coils named CC1 and CCC2, which span 10-15nm and terminate in the microtubule binding domain (MTBD) (Gee et al. 1997; Gibbons et al., 2005). ATP binding and hydrolysis in AAA1 cause changes in the MTBD’s affinity for microtubule—low affinity when ATP is bound and high affinity in the APO state (Imamula et al., 2007; Redwine et al., 2012). CC1 and CC2 make contact with the MTBD and their orientation relative to each other affects the MTBD affinity for the microtubule (MT) (Kon et al., 2009; Carter et al., 2008).

The second motor domain appendage, the N-terminal linker, lies on the face of the motor domain (Koonce et al., 1998; Burgess et al., 2004; Roberts et al., 2009). The linker is the MD structure responsible for the movement of the dynein motor domain relative to the heavy chain tail to produce the power-stroke and propagate mechanical force for movement of the motor (Burgess 2003); specifically, conformational changes and movement of the linker produce this power-stroke (Burgess et al., 2004; Roberts et

al., 2009; Kon et al., 2011; Carter et al., 2011; Kon et al., 2012; Schmidt et al., 2012

Bhabha et al., 2014). The final appendage of the motor domain, the C-sequence, lies on the surface of the AAA1/5/6/ opposite the linker (Roberts et al., 2009; Kon et al., 2011).

Little is known about the role of this structure, though it has been implicated in the regulation of dynein processivity and force production (Numata et al., 2011; Nicholas et al., 2014 and See Appendix) (See Figure 1.2).

Figure 1.2 Cytoplasmic dynein motor domain A. Schematic diagram crystal structure representation of the dynein motor domain (Modified from Hook and Vallee, 2012). AAA1-AAA6 forms the central ring around a central pore in a clockwise orientation. The stalk emerges from AAA5 and AAA5 and terminates in the MTBD. The crystal structure revealed a novel appendage, the buttress, which extends from AAA5 to make contact with the stalk. The force-producing component of the motor domain, the linker, lies on top of the motor domain and changes its position depending on the nucleotide state of AAA1. In the APO state of the motor domain, the linker is docked on AAA4. Not depicted is the C-terminal domain which is absent from yeast dynein, but in all other species the C-terminal domain on the motor domain opposite the linker. B. 2.8A° Crystal structure of dynein motor domain (From Kon et al., 2012)

The dynein motor domains (MDs) require the binding and hydrolysis of ATP to induce conformation changes in the AAA ring to influence MT binding affinity and a force producing power-stroke, ultimately driving the movement towards the minus-end of

the microtubule. ATP binding to AAA1 closes the ring by rotation of AAA2 and AAA3,

allowing a cascade of changes within the motor domain including the linker adopting a

bent “pre-power stroke” conformation and the MTBD releasing from the microtubule

moving towards the minus-end of the MT. The sliding of the two coiled-coils within the stalk decreases the affinity of MTBD. The long-rang communication between AAA1 and the MTBD, a distance of ~25nm, is an unanswered yet critical question in the dynein field. It is unknown how the registry of the coiled-coil changes in an ATP-binding dependent manner, especially since AAA1 is a significant distance away. It is possible the buttress, a coiled-coil structure emerging from AAA5 and contacts the stalk, and a direct interaction could propagate the ATP signal (Bhabha et al., 2014; Schmidt et al.,

2012; Kon et al., 2009; Carter et al., 2008). Upon ATP hydrolysis, the linker undergoes a movement to its original position and the MTBD makes a low-affinity interaction with the microtubule (Burgess et al., 2003, Roberts et al., 2009; Kon et al., 2005). This interaction induces the release of ADP-Pi and the formation of a strong MTBD-MT interaction to move the dynein complex towards the minus-end of the MT.

Figure 1.3: Conformational changes in the linker influence the power-stroke state of the dynein motor domain. (Modified from Cianfrocco et al 2015; Carter et al. 2016). A. The linker adopts multiple conformations depending on nucleotide state. Upon binding to ATP in AAA1, the linker adopts a bent conformation, positioning itself over AAA3. This is the pre-power stroke state. After ATP hydrolysis and release of –Pi, the linker moves to dock at AAA4/5. This movement, or power stroke is responsible for force production of dynein (Burgess et al., 2003). B. A representation of the mechanochemical cycle of dynein. Conformational changes induced by ATP binding to AAA1 cause reorganization of the linker and subsequent force production during the power- stroke. Concurrently, ATP state of the motor domain influences the microtubule affinity

of the MTBD. Binding of ATP releases dynein from the MT. In the ADP-Pi state, MTBD makes a low affinity contact with tubulin towards the minus end of the microtubule. The release of ADP-Pi increases dynein’s interaction with the microtubule allow for translocation down the microtubule. It is the combination of changes in linker conformation and MTBD affinity that allows for dynein’s force production towards the minus end of the microtubule.

The force production from the linker and the changes in microtubule binding does

not occur in isolation but occurs in each motor domain of the dynein complex. Questions remain whether there is coordination in the two motor domains relative to each other, but single molecule studies have started to examine this question. Relative to each other, the

motor domains are uncoordinated, producing alternating or non-alternating movements.

Additionally, rather than stepping consistently along the microtubules, individual motor domains are able to take both forward and backward directed steps with a variable size ranging from 12-24 nm or 24-34nm measured (Mallik et al., 2004; Reck-Peterson et al.,

2006; Qiu et al., 2012; Dewitt et al., 2012). While additional single molecule studies are required to completely understand the coordination of the motor domains for dynein’s minus-end directed movements, the motor properties of the dynein complex have been measured. Velocity of a single mammalian dynein was reported to range from .6-8 μm

sec-1 and an average run length ranging from .4-.7 μm (Wang et al., 1995; Ori McKenney et al., 2012; McKenney et al., 2010; King and Schoer 2004; Mallik et al., 2005). The linker movement during the mechano-chemical cycle dynein measured force production of ~1pN for mammalian dynein (McKenney et al., 2010, Mallik et al., 2004; Rai et al.,

2013). This translocation and force production that allow dynein to function as the minus-end directed microtubule motor.

Non-dynein motor domain regulation of motor activity

The motor activity of dynein is affected by additional subunits of the dynein

complex as well as dynein adaptor proteins. The heavy chain of dynein consists of not

only the motor domains but also the heavy chain tail that contains the sites of contact for

the light intermediate chains and intermediate chains (Tynan et al., 2000). Mutants in the

N-terminus of the heavy chain decreased inter-head ATP coordination, causing a premature release from the MT (Ori-McKenney et al., 2011). In yeast, deletion of light

chain (LC8) ortholog, Dyn2, reduced dynein’s the run length by decreasing the stability

of the intermediate chain/heavy chain interaction (Rao et al., 2013). These results

highlight the role of the tail and stability of the dynein complex in allosterically

controlling the behavior of the motor domains.

Additionally, extra-molecular interactions with either dynactin or Nde1/Lis1

modulate the dynein motor behavior. Dynactin (Dynein-Activator), the large 1.2 mDA

complex, increased the processivity of dynein by increasing run length (King and Shroer,

2004; Ross et al., 2006). The interaction between dynactin’s p150 domain and dynein

intermediate chain stimulated dynein processivity by decreasing lateral stepping of

dynein (Tripathy et al., 2014; Kardon et al., 2009).

Recently, the dynein field has questioned the motor behavior of tissue purified

mammalian dynein and dynactin in single-molecule studies. Two papers reported fluorescently labeled purified dynein demonstrated diffusive behavior in contrast to the processive movements measured in optical trap experiments (McKenney et al., 2014;

Schlager et al., 2014). However, processive movements seen previously for dynein were

only achieved when the complex of BicD2-N-terminal, dynactin, and dynein was formed.

The dynactin complex and dynein alone did not interact or allow for a processive dynein

motor. In addition to Bicd2-N, there are multiple other proteins that activate dynein

motility with dynactin, including Spindly, FIP3 and HookA (McKenney et al., 2014).

Increasingly it is clear that dynactin requires binding to additional factors to not only

associate with the dynein complex, but also to affect dynein’s processivity. Additional

characterization of dynein’s interaction with either the dynactin complex or the p150

subunit alone could elucidate the differences seen in dynactin’s ability to bind dynein and modulate of dynein motor behavior.

The second group of dynein adaptor proteins that influence dynein motor activity is Nde1/Ndel1 and Lis1. Nde1 recruits Lis1 to the dynein complex to influence the force production of the dynein complex both in vitro and in vivo (McKenney et al., 2010;

Reddy et al., 2016). Formation of a Lis1-Nde1-dynein complex transforms dynein in a persistent force producing motor allowing dynein to spend a longer time at its 1.1pN stall force (McKenney et al., 2010). Lis1 interacts with the motor domain, specifically AAA3, to increase dynein-MT interaction (Huang et al., 2012; Toropova et al. 2014). Even in the presence of high concentrations of ATP, dynein with Lis1 is unable to release

effectively from microtubules and exhibits pausing in motility analysis (McKenney et al.,

2010; Huang et al., 2012; Yamada et al., 2009). Single-molecule experiments suggest

that Lis1 and Nde1 could be required for dynein’s role in high-force cellular functions.

B. Cargo Binding

The dynein heavy chain tail extends from the dynein motor domains and provides

the platform for binding of the dynein subunits required cargo interactions (Taynn et al.,

2000; Habara et al., 1999). The light intermediate chains (LICs) and the intermediate

chain (ICs) within the dynein complex are essential for cargo binding along with dynein

adaptor proteins dynactin, Nde1l and Nde1. The diverse cargo interactions are required for dynein’s the numerous cellular functions.

The LICS were identified as the 50-60 molecular weight proteins in the bovine brain purified dynein complex (Hughes et al., 1995). Higher eukaryotes have two LICs,

LIC1 and LIC2, which are 65% identical and 71% similar (Tynan et al., 2000). Both

LICs have a highly conserved N-terminal P-loop structure (Hughes et al., 1995) that resembles the Ras-like GTP-binding protein family (Schroeder et al., 2014). This Ras-

like domain facilitates LIC’s interaction with the dynein heavy chain while the C-

terminal domain is required for LIC’s interaction with known binding partners (Taynan et

al., 2000; Schroeder et al., 2014).

The interactions between the C-terminal domain of the LICs and a diverse set of proteins are required for targeting of dynein to cellular cargo for specific dynein activities. LIC1 has been shown to be involved in dynein-dependent processes including minus-end directed vesicle transport, Golgi positioning and transport of Golgi to ER vesicles (Palmer 2009). LIC1 interacts with RILP, a lysosomal protein, and is required for the recruitment of dynein to the lysosome (Scherer et al., 2014, Tan et al., 2011). In addition, LIC1 has been shown to bind pericentrin at the centrosome (Tynan et al., 2000),

BICD2 (Splinter et al., 2012), Rab11-FIP3 on the endosomal recycling compartment

(Horgan et al., 2011), Rab4A on early endosome for regulation of membrane recycling

(Bieli et al., 2001) (Table 1.1).

Dynein Dynein Binding partner Specific Cell- Reference Subunit adaptor Cycle Stage protein Light Pericentrin Interphase/Mitosis Tynan et al., Intermediate - 2000 Chain-1 RILP Interphase Scherer et al.,

2014 Tan et al., 2011 BicD2 Interphase/Mitosis Splinter et al., 2010 Rab11-FIP3 Interphase Horgan et al., 2011 Hexon Interphase-Virus Bremner et al., 2009 Rab4A Interphase Bieli et al., 2001 Mad2 Mitosis Sivaram et al., 2009 Par3 Interphase Schmoranzer et Light al., 2009 Intermediate - Rab11-FIP3 Interphase Horgan et al., Chain-2 2010 Golgin160 Interphase Yadav et al., 2012 Herpes Simplex Interphase-Virus Ye et al., 2000 Virus Protein uL Intermediate (34), - Chain Zw10 Mitosis Whyte et al., 2008 Huntingtin Interphase Caviston et al., 2007 Snapin Interphase Cai et al., 2012 Clip-170 Interphase/Mitosis Vaughan et al., 2002; Tai et al., 2002 Zw10 Mitosis Starr et al., 1998; Chan et al., 2000 Spindly Mitosis Griffis et al., 2007 Rab6 Interphase Short et al., Intermediate 2002 Dynactin Chain RILP Interphase Jordens et al., 2001 HookA Interphase Zhang et al., 2014 BICD2 Interphase/G2 Splinter et al., 2012; Hoogenraad et al., 2013 SNX5/6 Interphase Wassmer et al., 2009

FIP3 Interphase Horgan et al., 2010 NuMA Mitosis Merdes et al., 1996 AnkG Interphase Kuijpers et al., 2016 CENP-F Mitosis Vergnolle and Taylor, 2007 Zw10 unknown Stehman et al., 2007 DISC1 mitosis Bradshaw et al., 2004 Gamma tubulin unknown Feng et al., 2000 Intermediate Nde1 and p60 subunit of Interphase/mitosis Toyo-oko et Chain Ndel1 katanin al., 2006 Su48 mitosis Hirohashi et al., 2006a; Hirohashi et al., 2006b dynein light chain unknown McKenney et LC8 al., 2011 14-3-3e unknown Toyo-oka 2003 pericentrin Interphase/mitosis Feng et al., 2000

Table 1.1. Summary of reported cargo interactions with components of the dynein complex or dynein adaptor proteins and their relevance in specific states of the cell-cycle (if known).

The dynein intermediate chain (IC) is the critical hub for binding of dynein accessory proteins, dynactin and Nde1/Ndel1, as well as direct interactions with physiological cargos. The N-terminal domain of IC consists of an N-terminal alpha helical region, followed by a serine-rich unstructured domain, and the dynein light chain binding sites (Harbura et al., 1999; Nayaorodk et al., 2011). It was previously reported that the interaction of Light chains, LC8, TcTex, and Roadblock, with the intermediate chain facilitated binding of cargo to the dynein complex through light chain cargo

interactions (King et al., 1996a; King et al., 1996b; King et al., 1998). Structural studies

of LC8 dispute this cargo-adaptation model since the intermediate chain and light chain

interactions occupy the same binding groove in LC8 (Bensison et al., 2007), preventing

the recruitment of additional dynein cargo through the light chains. Upstream of the light

chain binding sites, the C-terminus of the IC forms two WD-40 domains, required for the

association with the heavy chain (Ma et al., 1999) so the cargo-binding activity of the

intermediate chain is contained within the N-terminal region.

A small class of proteins binds directly to the N-terminus of the intermediate chain including—Huntingtin which binds to facilitate dynein-dependent vesicular transport (Caviston et al., 2007); Snapin, a neuronal snare binding protein that links dynein to endosome-lysosome transport (Cai et al., 2012); and the Hexon subunit of

Adenovirus to transport adenovirus to the nucleus during viral infection (Bremner et al.,

2009) (See Table 1-1). The majority of intermediate chain cargo interactions occur through the interaction with dynactin, Nde1 and Ndel1 (Table 1-1). Both proteins bind to the N-terminal domain of the IC and compete for a binding site in the first 70 amino acids

(McKenney et al., 2011). This competition could be critical to distinguish specific dynein functions. How each protein recruits cargo to the dynein complex, through their

interaction with the intermediate chain, will be discussed in detail below.

The intermediate chain possibly achieves the diversity of cargo interactions

required for all of cytoplasmic dynein’s cellular functions in part because of the number

of IC isoforms and phospho-regulation. There are two isoforms of IC, IC1 and IC2, each

with multiple spliceforms. In total there are 5 IC isoforms and splice variants (IC-1A,

IC-1B, IC-2A, IC-2B and IC-2C) (Vaughan et al., 1995; Pfister et al., 1996). The

expression pattern varies for each isoform, with IC1 predominately found in the brain

while IC-2C exhibits universal expression (for here forward, referring to IC means IC-

2C) (Vaughan et al., 1995; Pfister et al., 1996). Isoform specificity could regulate tissue

specific cargo. Additionally, intermediate chain phosphorylation at S84 and T89

decreases its interaction with dynactin in interphase and mitosis (Vaughan et al., 2002;

Whyte et al., 2008), which could decrease dynein’s association with cargo through a loss

of the dynein-dynactin interaction. Further examples are phosphorylation of the

intermediate chain in axons by GskB and Erk1/2 decreases IC’s interaction with Nde1

and affects dynein’s function axonal transport of TrkA positive endosomes and Rab7

vesicles (Guo et al., 2015; Mitchell et al., 2014). In this manner, the intermediate chains

are a platform regulation of dynein-cargo interactions.

Dynactin

As mentioned previously, dynactin is required dynein’s processivity, but also has

cargo-binding functions. Dynactin is a large complex is comprised of four distinct

structural units: the shoulder, the filament, the barbed end and the pointed end that carry

out a roles in stabilization of the dynactin complex or cargo binding. The filament,

barbed end and pointed-end of dynein provide structural support for the dynactin

complex (Miller et al., 1992; Schafer et al., 1994; Schroer et al., 2004). In addition to a

structural role, the p25 subunit of the pointed end was demonstrated to have an additional

role in cargo binding in endosome transport in Aspergillus (Zhang et al., 2011). The final

structure of the dynactin complex is the shoulder, which is comprised of the p50, p150

and p22/24 subunits (Urnavicius et al 2015). The p50 and p150 subunits are essential for interactions with microtubules, cargo proteins and the dynein complex.

The p150 subunit of interacts with the intermediate chain of dynein (Holzbaur et

al., 1992; Vaughan and Vallee, 1995). It is through this interaction that dynactin provides

the link from cargo to dynein. For example, in mitosis, dynactin interacts with

kinetochore proteins zw10 and Spindly to influence dynein’s kinetochore function

(Griffis et al., 2007; Chan et al., 2000). Plus-end TIP proteins EB1 and Clip170 interact with p150 and target dynein, via dynactin, to the plus-end of the microtubules (Vaughan et al 2002). Additionally, the interaction between dynein and dynactin is essential in specifying dynein’s role in is organelle transport through the interaction of p50 or p150 subunits with interacting proteins embedded in the organelle (Haghnia et al., 2007)

(Table 1.1).

The interaction between dynactin and Bicd2, a protein first identified in mRNA transport in drosophila but implicated dynein’s role in vesicle transport, Golgi positioning and nuclear envelope recruitment of dynein in mammalian cells, has shed light on the complex nature of dynein, dynactin and their relationship to cargo. Studies initiated to determine the interaction between BicD2 and dynactin showed that dynactin requires

BICD2 N-terminal to form a complex with dynein (Splinter et al., 2012). Bicd2 binds both to LIC1 and the p50 subunit of dynactin. Although a direct interaction between dynein and dynactin with p150 and IC had been reported, the formation of the entire dynein-dynactin complex in vivo can only be formed in the presence of additional protein

Bicd2 (Splinter et al., 2012). Further research will determine the role of cargo in stabilizing the dynein-dynactin interaction and the presence of additional cargos, which bind to both LICs and dynactin to perform the similar functions as BicD2.

Nde1 and Ndel1

The second class of dynein adaptor proteins is Nde1 and Ndel1. The paralogues,

Nde1 and Ndel1, share 55 % identity and 73% similarity. Little is known about specific

differences between Nde1 and Ndel1 apart from varying in expression patterns in the

developing neocortex (Feng et al., 2000; Sasaki et al., 2000) as well as knockout

phenotypes: Ndel knockout mice are embryonic lethal (Sasaki et al., 2005), while Nde1

knockout mice survive but have a smaller brain as a result of defects in cortical

development (Feng et al., 2004). It is assumed that Nde1 and Ndel1 have similar

structures, cargo-binding properties and role in dynein functions but extensive

characterization of Nde1 and Ndel1 binding partners and physiological roles will clarify

the role of each paralogue (For this reason Nde1 and Ndel1 will be referred to as

Nde1/el1 when talking about principles that apply to both).

Nde1/Ndel1 a highly elongated have a N-terminal coiled-coiled domain followed

by a C-terminal unstructured region. The C-terminal domain contains multiple

phosphorylation sites shown to be targeted by Aurora A, Erk1/2, Cdk1, Cdk5 or PKA

(Mori et al 2007; Bradshaw et al., 2011; Yan et al., 2003) as well as multiple sites of

interaction for known binding partners. Nde1/el1 interact with CENP-F, DISC1, dynein

light chain LC8, p60 subunit of katanin, 14-3-3e, Su48, Zw10, pericentrin, Ankryn G—to name a few (see review by in Bradshaw 2011) (Vergnolle and Taylor 2007; Bradshaw et al 2004; McKenney et al 2011; Toyo-oko et al 2006; Kuijers et al., 2016; Stehman et al.,

2007) (Table 1.1). These interactions specify Nde1/el1 to function in dynein-dependent process including lysosome transport in cell culture and in neurons, (Yi et al 2011,

Pandey et al 2012), vesicle sorting at the axon-initial segment (Kuijpers et al 2016),

formation of spindle asters in Xenopus mitotic egg extract (Zyłkiewicz et al., 2011), and

Golgi positioning (Lam et al 2010).

Nde1/Ndel1 acts as a dynein adaptor protein because of its ability to bind to

multiple cargos as well as dynein and Lis1. Nde1/Ndel1 binds to Lis1 and the dynein

intermediate chain through the N-terminal coil-coiled domain (Feng et al.,2000,

Derewenda et al.,2007; Zyłkiewicz et al., 2011; Wang et al., 2011) . Early yeast-two hybrid studies demonstrated that Ndel1 bound to the dynein motor domain, but this interaction has not been confirmed biochemically (Sasaki et al., 2000; McKenney et al.,

2010). Biochemically, Nde1/el1 recruits Lis1 to the dynein motor domain, through the formation of a dynein-Lis1-Nde1 complex. Through this interaction, Lis1 is able to influence the motor behavior of cytoplasmic dynein (McKenney et al., 2010). Given the requirement of Nde1 in Lis1 biochemical functions, it was thought that Nde1l was essential to recruit Lis1, as well as dynein, to specific subcellular location at the kinetochore in mitosis and to membranes of organelles in interphase (Stehman et al.,

2007, Vergnolle and Taylor’ 2007; Liang et al., 2007; Lam et al., 2010). Because Nde1 is recruited to specific subcellular localizations through its diverse interactions with binding partners, it plays an essential role in directing dynein activity to specific sites.

Lis1 is thought to have a greater role in regulating dynein’s motor properties than facilitating cargo interactions. While Lis1 interacts with Nde1, the only other protein shown to interact with Lis1 is the microtubule plus-end tip protein Clip-170 (Coquelle et al., 2002).

3. Dynein’s function during G2/M

As cells progress through the cell-cycle, cytoplasmic dynein adopts specific interactions required for dynein’s recruitment to mitotic sites. During interphase, dynein recruitment to the Golgi and lysosomes/late endosomes allow for Golgi positioning and minus-end directed vesicle transport (Vallee et al., 2012)(Figure 1.4). Upon entry into

G2, dynein accumulates at the nuclear envelope followed by localization to the kinetochore (KT), spindle pole, and cell cortex in mitosis (Figure 1.5). At each site, dynein is recruited by site-specific interactions. Below describes how dynein is recruited to mitotic sites to perform its functions in G2 and mitosis.

Figure 1.4. Cytoplasmic dynein adopts cell-cycle specific interactions. (Modified from Vallee et al., 2012) Diagram of dynein’s recruitment to specific cellular sites depending on the stage of the cell cycle. At each localization, dynein adaptor proteins, dynactin and Nde1/el1, bind to site-specific cargo. During interphase, dynein is recruited to the Golgi and lysosome, while in G2/mitosis dynein is recruited to the nuclear envelope and kinetochore.

Figure 1.5. Cytoplasmic dynein localization during G2 and mitosis. Staining of Hela cells with dynein intermediate chain antibody reveals nuclear envelope staining in G2 and prophase, kinetochore localization in prometaphase, spindle localization during metaphase and in anaphase, IC signal is detected at the cell cortex. Throughout G2 and mitosis, dynein is recruited to distinct cellular sites depending on the stage of the cell- cycle. Scale = 5 μm Unpublished data.

During G2, dynein recruitment to the nuclear envelope is dependent on an early

and a late recruitment pathway, which are required for dynein-dependent G2 functions.

Dynein at the NE in G2 and prophase helps prepare the cell for entry into mitosis. In

mammalian cultured cells, nuclear envelope dynein is thought to produce force on the NE

in prophase to form prophase nuclear envelope invaginations indicative of forces on the

nuclear envelope, though the exact mechanism is unknown (Salina et al., 2000).

Additionally, dynein at the NE provides the link between the microtubule network and

the nucleus. Disruption of dynein recruitment factors at the NE, cause defects in

centrosome anchoring as well as centrosome separation in prophase (Robinson et al.,

1999, Bohley et al., 2011, Splinter et al., 2010, Raaijimakers et al., 2012). To perform

these functions, dynein is recruited to NE in early G2 and late G2. During early G2,

dynactin is recruited to the NE by Bicd2, which itself is recruited to by the nuclear pore

protein RANBP2. Through this interactions and Bicd2’s interaction with dynactin, the

entire dynein-dynactin-Lis1 complex is properly localized to the NE in G2 (Splinter et

al., 2012; Baffet et al., 2015; Busson et al., 1998). In late G2/prophase, CENP-F exits the

nucleus and binds to nuclear pore protein, Nup133, in turn recruiting Nde1/Ndel1

(Bohley et al., 2011; Baffet et al., 2015). In mammalian cells, this late pathway has been

implicated in the microtubule anchoring as well as prophase nuclear invagination

functions of dynein (Bohley et al., 2011; Hebbar et al., 2008). The purpose of two

pathways is thought to be to recruit additional dynein required for late G2/prophase functions, for example providing additional forces for nuclear envelope break down.

While disruption of dynein recruitment to the NE does not block entry in mitosis, in neuronal progenitor cells, the sequential recruitment of dynein to the NE is essential interkinetic nuclear migration and translocation of the nucleus to the ventricle for entry into mitosis Any block in the early and late dynein recruitment pathways cause a G2 arrest (Hu et al., 2013; Baffet et al., 2015).

Dynein localizes to the kinetochore in prometaphase, followed by a decrease in kinetochore signal upon MT-attachment. Cytoplasmic dynein is absent at the kinetochores in metaphase (Hoffman et al., 2001; Pfarr et al., 1999; Steuer et al., 1999;

Wordeman et al., 1991; Hoffman et al., 2001, King et al., 2000). At the kinetochore, there are multiple dynein recruitment pathways. The reason for multiple cargo interactions is not clear but might be due to the requirement for separation of specific dynein functions or to ensure dynein’s kinetochore localization by having redundant

recruitment pathways since defects in dynein kinetochore function could have

catastrophic defects. Recruitment and regulation of dynein activity at the kinetochore is

critical for proper mitotic progression. It has been hard to determine the precise

mechanism of dynein in multiple steps of mitosis since there is significant inter-

dependency among the kinetochore proteins, but dynein has been shown to have a role in

chromosome movement towards the metaphase plate (Yang et al., 2007), stabilization of

end-on microtubule attachments, kinetochore orientation (Varma et al., 2007), and

removal of spindle assembly checkpoint proteins to inactivate the spindle assembly

checkpoint (Howell et al. 2001; Wojcik et al., 2001).

The first kinetochore dynein recruitment pathway is through Zw10, a component

of the RZZ complex. The RZZ complex is comprised of three proteins: Zw10, rod, and

Zwilch (Chan et al., 2000; Williams et al., 2003). Zw10 localized to the kinetochore

through the direct interaction with kinetochore protein Zwint but its interactions with Rod

and Zwilch are required for both RZZ formation and their localization to the kinetochore

(Chan et al 2000). Zw10 recruits dynein through interactions with the p50 subunit of dynactin (Starr et al 1998), dynein accessory protein Spindly (Gassman et al., 2008;

Griffis et al., 2007, Chan et al., 2009; Barisc et al., 2010), and by binding directly to the intermediate chain (Whyte et al., 2008). The timing and function of these interactions is not clear; but zw10 recruitment of spindly and dynactin is essential for dynein’s function in spindle assembly checkpoint removal and dynein-dependent removal of spindle assembly checkpoint proteins, Mad1-Mad2 (Griffis et al., 2007, Barisic et al., 2010). The complex of Mad1-Mad2 produces a wait signal that inhibits the activity of the anaphase- promoting complex (APC/C) until proper kinetochore-microtubule attachment. After the

correct attachment, dynein is required in the essential removal of Mad1/2 from the

kinetochore to activate Anaphase-promoting complex (APC/C) to initiate anaphase onset.

(Buffin et al., 2005; reviewed in Salmon and Mussachio 2007). The RZZ complex is the

required the link between Mad1/2 and dynein to allow for proper SAC activation and

timely inactivation when all chromosomes have proper MT attachments (Howell et al.,

2001, Wojack et al., 2001, Kops et al 2005;Yang et al 2007). Knockdown of Zw10 and

Rod in Xenopus or mammalian Hela cells prevents the kinetochore recruitment of Mad2, causing premature inactivation of the SAC and subsequence early anaphase onset (Chan et al., 2000; Basto et al., 2000).

Dynein adaptor proteins, Nde1 and Ndel1, recruit both Lis1 and dynein to the kinetochore through its link with kinetochore binding-partner, CENP-F (Liang et al.,

2007; Stehman et al., 2007; Vergnolle and Taylor, 2007). Less is known about the exact

role of this second dynein recruitment pathway and how it relates to the zw10 dependent

dynein recruitment pathway. Nde1 has been implicated in dynein-mediated removal of

spindle assembly checkpoint proteins since perturbation of Ndel1 decreased the spindle pole accumulation of zw10 in Az-Dog experiments (Yan et al., 2003; Liang et al 2007).

Additionally, Nde1 and Nde1l have been implicated in the MT-KT stabilization role of dynein, though the exact mechanism and role is unknown (Steheman et al., 2007).

Especially since in biochemical experiments, Nde1 interacts with zw10 (Steheman et al.,

2007), further work is required to understand the relationship between the two dynein recruitment pathways at the kinetochore and how exactly each contribute to dynein kinetochore function.

Given that dynein is a microtubule motor, dynein and accessory proteins are

required for proper formation of the mitotic spindle and spindle pole focusing. The

bipolar mitotic spindle is formed in part by the anti-parallel sliding of over-lapping

microtubules. Both dynein and mitotic spindle kinesin, Eg5, provide opposing forces to

allow the separation of the centrosome and proper bipolar spindle formation

(Taunembaum et al., 2008, Raajiimakers et al., Van heesenben et al., 2014, Gaglio et al.,

1996,Vaisberg et al., 1993). Little is known about the dynein is involved in this process.

It is unclear how dynein requires interaction with specific interactors, since dynactin,

Nde1/Nde1l and Lis1 have all been implicated in this dynein-dependent function

(Echeverri et al., 1996; Burkheart et al., 1997; Raajimakers et al 2013).

In addition to formation of bipolar spindle, dynein has been implicated in spindle pole focusing both in cell-free Xenopus egg extract or mammalian culture cells. Cell-free

Xenopus egg extract experiments demonstrate dynein focuses of microtubules into

spindle pole asters and in mammalian cells, for the maintenance of the centrosome-

spindle pole connection (Verde et al., 1991; Heald et al., 1996; Heald et al., 1997). In

mammalian cells, NuMA (Nuclear/mitotic apparatus protein), recruits dynactin and

dynein to the spindle pole where the complex functions in spindle pole focusing (Merdes

et al., 1996; Merdes et al.,2000). Additionally Nde1/Nde1l have been implicated in this

dynein-dependent function. Depletion of Nde1/Nde1l prevented focused spindles in both

Xenpus egg extract and mammalian cell culture (Wang et al., 2011; Zyłkiewicz et al

2011). Consistent with its involvement in aster and spindle pole formation, multiple

reports have demonstrated that Nde1 null mutants don’t have proper spindle formation in

drosophila (Wainman et al., 2009). Questions remain about how Nde1/Nde1l are

recruited to the spindle pole and its function relative to the NuMA-dynein-dynactin complex.

The final site of dynein localization in mitosis is the cell cortex. In mammalian cells, dynein localization to the cell cortex in metaphase and anaphase is required for proper spindle positioning in metaphase and elongation of the spindle in Anaphase B.

Proper distribution of dynein forces on the cell cortex drives the positioning of the spindle required to correctly separate the two daughter cells. The mechanism through which dynein correctly positions the mitotic spindle involves capturing end-on the astral microtubules and providing a balanced pulling force on the spindle (Laan et al., 2012).

Additionally, in anaphase, dynein drives chromosome separation in anaphase B (Kotak et al 2014). In metaphase and anaphase, dynein localizes to the cell cortex at distal ends of the cell, but is absent from cortex at the spindle midzone (Bosson et al., 1999; Kiyomitsu and Cheeseman, 2012; Kotak et al., 2012). This cortical localization is dependent on

NuMA, which facilitates the link between dynein/dynactin and protein complexes of the cell cortex. During metaphase, the NuMA-LGN-Gai complex recruits dynein to the distal ends of the cell. Specifically the N-terminal domain of NuMA mediates the interaction with dynein and dynactin (Kotak et al., 2012; Kiyomitsu and Cheeseman,

2012). In anaphase, NuMA is directly recruited to the cortex by protein 4.1G, independently of the LGN-Gai complex, to provides a direct link to increase levels of

dynein at the cortex (Kiyomitsu and Cheeseman, 2013).

Dynein, through its dynein adaptor proteins dynactin and Nde1/el1, adopt cell-

cycle specific interactions with NuMA, zw10, Spindly, CENP-F, and BicD2. Questions

remain on how these interactions are cell cycle regulated to dictate mitotic specific

localization and functions of dynein. The following section addresses how these G2 and

mitotic specific cargo interactions are formed and regulated.

4. Cell-Cycle regulation of dynein-cargo

As mentioned in the previous section, dynein is recruited to mitotic sites through

cell-cycle dependent cargo interactions. A main outstanding question is how do dynein

and its adaptor proteins release from interphase cargo to adopt G2/M specific

interactions. Recently, it was demonstrated that Cdk1 phosphorylation of dynein in G2

dissociated dynein from interphase cargo (Chung et al., 2016). Given the specialized

functions of dynein in G2 and mitosis, cell-cycle regulation of the dynein complex and its adaptor proteins is essential. How dynein performs its diverse roles in mitosis as well as how does dynein adopt specific interactions is unclear. Discussed below are clues into the regulation of cell-cycle specific cargo interactions.

A. Complex

The first means of regulating dynein’s cell-cycle behavior is different compositions of the dynein complex, specifically the association of either light intermediate chain-1 or light intermediate chain-2. At the molecular level, the LICs form distinct homodimers and compete with each other to bind to the dynein heavy chain since the heavy chain is unable to bind the two LICs simultaneously (Tynan et al., 2000).

Although the LICs have similar sequences and structures (Hughes et al., 1995), the LICS have a distinct set of binding partners (see Table 1.1), which could possibly lead to

specific dynein complexes with unique functions.

In mitosis, both Lic1 and Lic2 localization to the kinetochore (Mahale et al.,

2016), but the spindle localization patterns differ. LIC1 localizes to the mitotic spindle

while LIC2 stains the spindle pole (Horgan et al., 2011). While it has been shown that

Lic1 binds to spindle assembly checkpoint protein, Mad2, identification of mitotic

specific cargo is lacking (Sivaram et al., 2009). Further examination of mitotic

interactors of LIC1 or LIC2 will allow for a better understanding of unique functions of

dynein complexes containing LIC1 or LIC2.

Additionally, differential phosphorylation of LIC1 and LIC2 in mitosis could

contribute to unique mitotic specific interactions. Cdk1 phosphorylation of LIC1 is

required for dynein’s role in inactivation of the spindle assembly checkpoint (Dell et al.,

2000; Niclas et al., 1996; Sivaram et al., 2009). It has not been demonstrated how this

phosphorylation regulates cargo, but previous work on the role of PKA phosphorylation

of LIC1 shows phosphorylation regulates its interaction with RILP to affect lysosome

transport during adenovirus infection (Scherer et al., 2014). Further identification of phosphorylation events, specifically by G2/M kinases, will elucidate the role of mitotic phospho-regulation of LIC1 and LIC2 cargo. This will lead to a better understanding how the presence of LIC1 or LIC2 directs dynein to specific mitotic sites for LIC-unique functions.

B. Association with mitotic specific proteins

The next point discussed is the interaction of dynein and its adaptor proteins, dynactin and Nde1l/el, with proteins only expressed or available for interaction in

G2/mitotsis. Timing of protein expression or physical barriers of protein localization are a means to restrict dynein interactions until the proper time of the cell cycle. By preventing the access to mitotic cargo until the correct stage, dynein is recruited to the

nuclear envelope, kinetochore, spindle pole or cell-cortex only when required for specific

mitotic functions.

Spindly is a novel kinetochore protein involved in dynein kinetochore function

and has no known expression or function interphase. Spindly localizes to the kinetochore

at early prometaphase through an interaction with zw10 and is removed in a

dynein/dynactin dependent manner when cells achieve microtubule attachment and align

at the metaphase plate (Griffis et al., 2007 Chan et al., 2009; Gassman et al., 2008; Griffis

et al., 2007, Chan et al., 2009; Barisc et al., 2010). Mitotic specific expression of Spindly

allows for it to act as a mitotic-specific recruitment factor for dynein at the kinetochore

through interaction with dynactin and zw10 (Gassman et al., 2010, Griffis et al., 2007,

Chan et al., 2009).

Dynein recruitment factors, CENP-F and NuMA, are confined to the nucleus until the proper cell-cycle stage to indirectly recruit dynein to the NE or spindle pole and cell cortex, respectively. During interphase, NuMA is restricted to the nuclear matrix but upon entry into mitosis, NuMA localizes to the spindle pole and cell-cortex (Lydersen et al., 1980). This redistribution to the spindle pole and cell cortex allows for dynactin- dynein recruitment to these mitotic sites (Merdes et al., 1996; Merdes et al.,2000; Kotak et al., 2012; Kiyomitsu and Cheeseman, 2012; Kiyomitsu and Cheeseman, 2013). CENP-

F expression is absent in G1 and S phases but protein levels increases in G2. CENP-F signal is restricted to the nucleus until late G2 when it exits the nucleus (Liao et al.,

1995), then CENP-F binds to the nuclear envelope via nuclear pore protein, Nup133

(Bohley et al., 2011). This late G2/prophase localization of CENP-F to the NE is required to recruit dynein accessory proteins Nde1l and Nde1 which function in the late

G2 dynein recruitment pathway to the NE (Bolhey et al., 2011; Hu et al., 2014) CENP-F

has an additional role at the kinetochore in mitosis (Vergnolle and Taylor, 2007), but the

initial interaction in a cell-cycle specific state is limited by CENP-Fs localization until the proper time of the cell-cycle. While the restriction of NuMA and CENP-F to the nucleus is required to temporally regulate dynein cargo-interactions, it is unknown how the nuclear exit is regulated. It is possible that nuclear envelope breakdown frees NuMA from the nuclear matrix to localize to the spindle pole or that phosphorylation of CENP-F

is in late G2 to allow for active nuclear export. Future studies on cell-cycle regulation of

CENP-F or NuMA will provide insight into the localization pattern of each protein.

C. Phosphorylation-

The final means to regulate cell-cycle dependent cargo interactions is phosphorylation. A group of kinases are only expressed or active in G2 and mitosis, which make them great candidates for cell-cycle regulation of dynein interactions. These mitotic specific kinases include Cdk1-Cyclin B, Plk1, the Aurora family of kinases and the NIMA family (Erich Nigg, 2001). Phosphorylation of proteins in the dynein pathways by these mitotic kinases could allow for precise regulation of interactions specifically in

G2 and mitosis. i. Direct phosphorylation of the intermediate chain of dynein

Dynein intermediate chain undergoes phospho-regulation to determine recruitment to and cargo-interactions at the kinetochore during prometaphase and metaphase. The dynein intermediate chain is phosphorylation at residue T89 by PLk1 during mitosis and a phospho-specific antibody recognizes phosphorylated dynein at the kinetochore in prometaphase (Whyte et al., 2009; Bader et al., 2011). The

phosphorylation state of dynein IC regulates its interaction with mitotic cargo zw10 and

dynactin: Phospho-IC binds to zw10, while the dephosphorylated state of the

intermediate chain binds to dynactin (Whyte et al., 2009). This phosphorylation

dependent cargo-switching is essential for dynein’s role in spindle assembly checkpoint

inactivation. Dephosphorylation of dynein IC at the T89 residue is required for dynein- dependent removal of zw10 and spindle assembly checkpoint proteins (Whyte et al.,

2008).

The intermediate chain of dynein contains a serine rich region in the N-terminal

domain, where T89 is located. It is possible that there are more sites targeted by

additional kinases to dictate additional interactions that specify subpopulations of

cytoplasmic dynein for specific functions.

ii. Cell-Cycle phosphorylation regulation of Dynein recruitment factors

Although specific cargo might not interact directly with dynein, their role in

binding to dynein adaptor proteins is critical for the localization and regulation of

cytoplasmic dynein. Cell-cycle specific phospho regulation of the recruitment of dynein cargo to mitotic sites is critical to determine the timing of dynein and dynein adaptor protein recruitment.

As mentioned previously, dynein recruitment and localization to the nuclear envelope in early G2 is dependent on the RanBP2-BicD2-Dynactin pathway (Doye et al.,

2010; Hut et al., 2014). Biochemically, Cdk1 phosphorylation increases the interaction between BICD2 and RanBP2. In mammalian cultured cells, Cdk1 phosphorylation of

RanBP2 in the early stages of G2 is required for Bicd2 localization to the NE as well as the recruitment of dynactin and dynein. This Cdk1 phospho-regulation of the RanBP2

and BicD2 is not only for the localization of dynein to the NE in early G2 but for its

function in interkinetic nuclear migration in neuronal progenitor cells (Baffet et al.,

2015). Since Cdk1/CyclinB kinase is only active at low levels in G2 and high levels in

mitosis (Hochegger et al., 2008), this interaction is restricted to a specific time of the cell

cycle so the initiation of dynein recruitment to the NE is restricted to early G2.

An additional example of a dynein cargo’s recruitment under cell-cycle control is

the recruitment of zw10 to the kinetochore by Zwint. Zw10 localizes to the kinetochore

through a direct interaction with kinetochore protein Zwint (Kasaboski et al 2011;

Famulski et al., 2008; Kops et al., 2005; Wang et al., 2004; Starr et al., 2000). Inhibition

of Aurora family kinase, Aurora B, or expression of Zwint Auroa B phospho-mutant

construct prevented kinetochore localization of zw10 (Kasaboski et al 2011; Famulski et

al., 2008). Since Aurora B is a kinetochore-specific kinase only active during mitosis,

Zwint phosphorylation is restricted both spatially and temporally. It is this

phosphorylation of Zwint and the subsequent recruitment of zw10 that allows for

Spindly, dynactin and dynein localization to the prometaphase kinetochore.

During metaphase, dynein is recruited to the distal ends of the cell cortex by the

NuMA-LGN-Gai complex. There is an asymmetric localization of dynein at the distal ends, depending on the position of the mitotic spindle; dynein is present on the cortex of the distal end of the cell furthest away from the mitotic spindle. In order to achieve this asymmetric localization, NuMA and dynactin interaction at the cell cortex is regulated by

Plk1 kinase. Plk1 signal from the spindle pole decreases the interaction between NuMa and dynein/dynactin to prevent recruitment of dynein to the cortex only when the spindle pole is in closer proximity to one distal end of the cell. Plk1 is a negative regulator of

dynein localization to the cell cortex, allowing for the asymmetric recruitment depending on the position of the mitotic spindle (Kotak et al., 2012; Kiyomitsu and Cheeseman,

2012).

It was mentioned previously that NuMA functions with dynein and dynactin both at the spindle pole and at that cell cortex. The switch between NuMA’s localization and function in metaphase and anaphase is regulated by the Cdk1 phosphorylation state of

NuMA. During metaphase, Cdk1 phosphorylated NuMA localizes to the spindle pole, but upon entry into anaphase, when the levels of Cdk1 drop leading to a decrease in the phosphorylation levels of NuMA, Cdk1 dephosphorylated NuMA localizes to the cell cortex (Kotak et al., 2013). In this manner, the Cdk1 phosphorylation state of NuMA temporally regulates its localization to either the spindle pole or the cell cortex allowing for dynactin and dynein recruitment to each location at different stages of the cell cycle. iii Cell-Cycle phosphorylation regulation of Dynein accessory proteins

Previous sections introduce the dynein accessory protein dynactin, Nde1/el1 and lis1 and address their role in modulation of dynein motor activity and dynein’s cargo binding. Although each has been implicated in dynein’s function in G2/M, the knowledge of their cell-cycle regulation is lacking. Little is known about how direct phosphorylation dictates their activity at and localization to mitotic sites.

Dynactin

The p150 subunit of dynactin is a target for phosphorylation in M phase, though many of the specific sites or kinases involved are not known (Huang et al., 1999). There have been two reports of mitotic phosphorylation of p150. In Drosophila, Aurora A phosphorylation of p150 is required for proper spindle formation potentially through the

regulation of p150’s microtubule binding activity. Expression of a p150 phospho-mutant

demonstrated strong spindle localization that was absent in the wt or phospho-mimetic

condition (Rome et al., 2010). In mammalian cells, Plk1 kinase activity regulated

dynactin’s localization to the nuclear envelope in prophase to aid in dynein’s role in

nuclear envelope breakdown. Again the p150 subunit, specifically amino acid 179, of

dynactin is the target for the Plk1 kinase (Li et al., 2010). While both are examples of

how mitotic specific kinases influence dynactin’s behavior through phosphorylation of

p150, the mechanism is not known. Information from interphase PKA phosphorylation

of p150 suggests that phosphorylation in the N-terminal domain of p150 decreases the interaction between the microtubule binding domain of p150 and microtubules (Vaughan et al. 2000). The role of this lower microtubule affinity is not clear. Additionally, little is known about the role of phosphorylation in cargo or dynein interactions. Since the p50 subunit of dynactin is also involved in cargo-binding, knowledge of the mitotic phosphorylation of this subunit is required to gain a better understanding of cell-cycle regulation of dynactin interactions.

Lis1

In its early characterization, Lis1 was identified as a phospho-protein (Sapir et al.,

1999), but no information is known about specific sites, kinases responsible for

phosphorylation or functional relevance of phosphorylation. A significant study of Lis1 phosphorylation is required to start to understand the role of lis1 phosphorylation in regulation of its G2/M function. Given its role in modulation of dynein’s motor activity rather than as a recruitment factor of dynein cargo, phosphorylation could be a means to regulate Lis1 ability to bind to Nde1/el1 or dynein.

Nde1/Ndel1

Nde1/Nde1 are great candidates for phospho-regulation of dynein activity.

Xenopus Nde1l (MP43) was originally identified in a screen of mitotic phospho proteins

(Stukenberg 1997), and both Nde1/Ndel1 are phosphorylated in M phase (Yan et al.,

2003). Within the C-terminal unstructured region of Nde1/Ndel1 there are multiple

phosphorylation sites shown to be targets of Aurora A, Erk1/2, Cdk1, Cdk5, PKA kinases

(Mori et al 2007; Bradshaw et al., 2011; Yan et al., 2003).

Specifically focusing on the phosphorylation of Nde1 by mitotic kinases Cdk1

and Aurora A, much of the studies have focused on phospho-regulation of Nde1/Ndel1

activity and function at the centrosome. Phosphorylation of Nde1l by Aurora A

implicated in Nde1l’s localization to the centrosome in prophase to influence centrosomal protein TACC3 and Aurora A for the maturation and separation of centrosomes in prophase (Mori et al., 2007). Additionally, Cdk1 phosphorylation of Nde1 decreases

Nde1 recruitment to centrosome recruitment factor su48 (Hiroheshi et al., 2006). The dephosphorylation of Cdk1 phosphorylated Nde1/el1 is essential not only for recruitment of Nde1, but regulation of spindle orientation in neuronal progenitor cells (Xie et al.,

2012). Although Aurora A phosphorylation of Ndel1 or dephosphorylated Cdk1 Nde1 regulate their localization to the kinetochore, little is known about the mechanism of

Nde1/el1 action or the effect on dynein activity. Additionally, the requirement for each phosphorylation in regulating specific Nde1/el1 centrosome functions is not clear.

In addition to the centrosome, Nde1/el1 localize to the nuclear envelope in G2 and the kinetochore during mitosis (Bohley et al., 2010; Stehman et al., 2007; Yang et al.,

2007). Questions remain about whether mitotic phosphorylation by Aurora A or Cdk1

influences Nde1 recruitment to these mitotic sites. Since Aurora A localizes to the

spindle pole during G2/M (Mori et al., 2007), Cdk1 phosphorylation is the better

candidate for regulation of Nde1 localization to the G2 nuclear envelope and kinetochore.

Small-molecule inhibition of Cdk1 prevented of Nde1/el1 localization at the nuclear envelope in G2. Given the role of Cdk1 phosphorylation of Ran-BP2 in recruitment of

Bicd2 to the NE, it is possible that Cdk1 could also be directly influencing the recruitment of Nde1/Nde1l in the late recruitment pathway. The role of Cdk1 phosphorylation of Nde1 in its recruitment to the NE and kinetochore is investigated in

Chapter 2.

While phosphorylation of Nde1/el1 has the potential to regulate its localization to specific mitotic sites through regulation of cargo interactions, it also has been implicated in modulating the direct interactions with Lis1 and dynein (Yan et al., 2003, Pandey and

Smith., 2011; Hebbar et al., 2008) to influence their cellular functions.

5. Conclusion-

The precise coordination and regulation of dynein activity during mitosis is required for faithful segregation of chromosomes. While the dynein motor domains orchestrate the force-production and translocation ability of the dynein complex, it is the

LICs, ICs and the interaction with dynein adaptors Nde1/Ndel1 and dynactin that dictate cargo interactions. During G2 and mitosis, dynein adaptor proteins, dynactin and

Nde1/Nde1l adopt cell-cycle specific interactions. These interactions direct dynein to the

NE in G2, and to the kinetochore, spindle pole and cell cortex in mitosis. An outstanding question is how exactly these interactions are regulated in a cell cycle specific manner.

Dynein complex composition, association with mitotic specific proteins and

phosphorylation by mitotic kinases are potential means for cargo regulation though additional information is required. Understanding how dynein is targeted to sites in mitosis will provide a more detailed understanding of dynein’s mechanism during G2 and mitosis. The work in this thesis aims to clarify the role of Cdk1 phosphorylation of Nde1 in the regulation and Nde1’s role in dynein-dependent functions in G2 and mitosis.

Chapter 2: Cdk1 phosphorylation of Nde1 activates its interaction

with NE and kinetochore binding partner, CENP-F, and regulates

mitotic function

Introduction

Cytoplasmic dynein is a minus-end directed microtubule motor protein involved in a broad range of cellular functions throughout the cell cycle. However, mechanisms for cell cycle-dependent dynein cargo recruitment and activation remain poorly understood. Dynein gradually accumulates at the nuclear envelope (NE) during G2 and then appears at kinetochores after nuclear envelope breakdown. We have recently found that the initial stages of dynein NE recruitment are activated by Cdk1 phosphorylation of the nucleoporin RanBP2, which, in turn, recruits dynein through its accessory protein

BicD2 (Splinter et al., 2010; Baffet et al., 2014). Curiously, Cdk1 had an additional role in the late G2 NE dynein recruitment pathway; in this case in controlling nuclear export of CENP-F, which binds to another nucleoporin, Nup133 (Baffet et al., 2014; Bohly et al

2011). CENP-F, then in turn, recruits dynein regulatory factors Nde1 and Ndel1 to the

NE (Bohly et al., 2011), though control of this and of subsequent mitotic behavior for each of these proteins remains largely unexplored.

Nde1 and Ndel1 are closely-related genes. Their protein products, referred to by these names or NudE and NudEL, (54% identical and 73% similar) consist of highly elongated N-terminal coiled-coil domains, which contain dynein and LIS1 binding sites

(Derewenda et al., 2007; Zyłkiewicz et al., 2011; Wang and Zheng, 2011; Feng et al.,

2000; Sasaki et al., 2000) (Figure 1A). We have found from in vitro studies that, NdE1 recruits LIS1 and dynein into a supercomplex, which enhances the dynein-microtubule

(MT) interaction under load; resulting in increased dynein force production in vivo and in

vitro (McKenney et al., 2010; Reddy et al., 2016). We have also determined using RNAi

rescue analysis in developing rat neocortex that Nde1 and Ndel1 are almost

interchangeable functionally, though Nde1 alone is essential for mitotic entry in the

neural progenitor cells (Doobin et al., 2016).

Nde1 and Ndel1 interact through their C-terminal disordered regions (see Figure

1A) with diverse subcellular proteins, including, in addition to CENP-F, the dynein light chain LC8, 14-3-3e, DISC1, su48, MCRS1/p78, and Ank-G (Vergnolle and Taylor, 2007;

Stehman et al., 2007, Toyo-oka et al., 2003, Ozeki et al., 2003; Hirohashi et al., 2006b;

Hirohashi et al., 2006a; Kuijpers et al., 2016). Knockdown of CENP-F, in particular,

causes marked reduction in the concentration of Nde1 and Ndel1 at the G2 NE and

mitotic kinetochores (Bohly et al., 2011;Vergnolle and Taylor, 2007). The mechanisms

responsible for cycle-dependent recruitment of Nde1 and Ndel1 to their G2-M cargo sites

and the physiological importance of these interactions remain important outstanding

questions.

Nde1 and Ndel1 contain phosphorylation sites for Cdk1, Cdk5, Aurora A and

Erk1/2 kinases (Yan et al., 2003; Alkuraya et al., 2011; Mori et al., 2007). Cdk5

phosphorylation of Ndel1 was found to affect lysosome motility in axons though reports

dispute the phosphorylation state required to cause an effect (Pandey and Smith, 2011;

Klinman and Holzbaur, 2015). In vitro Nde1-dynein and Nde1-LIS1 binding interactions were found to be affected (Hebbar et al., 2009; Zyłkiewicz et al., 2011) However, because phosphorylation sites are concentrated within the C-terminal Nde1 and Ndel1 domains, we have chosen to investigate a role in regulating cargo binding. Given the

contribution of Cdk1 in dynein NE recruitment, we have chosen to test the effects of this

protein kinase on Nde1 behavior.

The current study was initiated to understand the role of Cdk1-mediated Nde1

phosphorylation during G2-M phase of the cell cycle. We find that phosphorylation

clearly stimulates the Nde1-CENP-F interaction in vitro, and is required for Nde1

targeting both to the NE and kinetochores. Of considerable interest, phospho-Nde1

persists at kinetochores well beyond the metaphase/anaphase transition and much of

anaphase, similar to CENP-F. These results, together with physiological analysis of

Cdk1-phosphorylated Nde1 provide the first strong evidence that Nde1 phosphorylation

regulates Nde1/Ndel1 cargo binding, and suggests a novel Nde1 role in mitotic

progression.

Results

Cdk1 phosphorylated Nde1 and phospho-mimetic Nde1 localize to known Nde1 mitotic

sites

Anti-Nde1/el1 antibody revealed NE and kinetochore staining in Hela cells

(Stehman et al., 2007; Baffet et al., 2015). Staining coincided with that for CENP-F at

kinetochores even before nuclear envelope breakdown (Figure 2.1B). Nde1/el1 signal

persists at kinetochores through mid-anaphase, highly unusual behavior for a dynein-

related protein, but strikingly similar to that for CENP-F (Figure 2.1B). Unfortunately,

GFP wt Nde1 is not detected at the kinetochore during live-imaging preventing the measurement of kinetochore flux to understand the behavior of kinetochore Nde1.

Analysis was limited to quantitative analysis of kinetochore fluorescent intensity for the

Nde1/el1 antibody, which revealed an ~50% decrease upon chromosome alignment

(Figure 2.1C). Hela cells were treated with either nocodazole or Taxol to determine the importance of microtubule attachment or tension on Nde1/el1 kinetochore signal intensity. Measurement of Nde1/el1 signal intensity in nocodazole treated cells revealed a ~2 fold increase in fluorescent intensity compared to cells treated with Taxol (Figure

2.1D) which suggests MT-attachment contributes to the decrease of Nde1/el1 kinetochore signal upon chromosome alignment.

Figure 2.1. Examination of endogenous Nde1/Ndel1 localization at the nuclear envelope in G2 and the kinetochore during mitosis. A. Schematic diagram of Nde1 labeled with important phosphorylation sites (T215, T243, T246) and interacting regions of known binding partners. Ankyrin-G (Ank-G) and su48 have interact with the C-

terminal domain of Nde1 though a smaller domain has not been mapped. B. Endogenous Nde1/el1 and CENP-F co-localize at the nuclear envelope in G2 and the kinetochore throughout mitosis. The CENP-F and Nde1/el1 kinetochore signal is present prior to nuclear envelope breakdown and persists into anaphase. Scale = 5μm C. Levels of endogenous Nde1l/Ndel1 kinetochore signal was measured relative to ACA (ACA: anti- centromere antibody) signal in cells with unaligned or aligned chromosomes. The average ratio of Nde1/ACA is represented +/- SEM. Student’s T-Test was performed on 3 replicates of ~25 kinetochores/ cell for 15 cells ***p<.001 The relative Nde1/el1 kinetochore signal drops by ~50% from cells with unaligned chromosomes to cells with aligned chromosomes. Scale=5μm D. Hela cells were treated for one hour with 5μM Nocodazole, 10μM Taxol or 10μM MG132, to determine if MT attachment or MT- Kinetochore tension contributed to the decrease in kinetochore Nde1/el1 kinetochore signal in aligned chromosomes. Relative Nde1/Ndel1 signal levels were quantified relative to ACA signal. Average ratio of Nde1/ACA is represented +/- SEM. Student’s T-Test was performed *p<.05. Nde1/el1 kinetochore signal in cells treated with nocodazole was twice that of cells treated with MG132 or Taxol.

As a means to examine the role of Nde1 Cdk1 phosphorylation during G2 and mitosis, we monitored the distribution of GFP-tagged wt Nde1 as well as triple Cdk1 phospho-mimetic (Nde1-215E, 243E,246E) and phospho-mutant Nde1 constructs (Nde1-

215A, 243A,246A). The mutations were at amino acids T215, T243 and T246, which

have been shown to be phosphorylated in vivo and/or predicted by consensus site analysis

to be Cdk1 substrates (Alkuraya et al., 2011; Dephoure et al., 2008) Given Nde1’s

localization to the NE in G2 and prophase, we assayed for the localization of GFP Nde1

constructs for co-localization with CENP-F at the NE (Figure 2.2A). Both Nde1 WT and

Nde1 phospho-mimetic co-localized with CENP-F at the G2 NE, though levels of Nde1

phospho-mutant were reduced. Investigation of the effects of expression of wt Nde1,

Nde1 phospho-mimetic or phospho-mutant on progression through G2 reveals Nde1 phospho-mutant expressing cells an increased ratio in G2+/total GFP+ cells in

comparison to GFP Nde1 phospho-mimetic. This suggests a possible delay in G2 due to

decreased localization of phospho-mutant at the nuclear envelope.

Figure 2.2 Localization of Nde1 phospho-mimetic and phospho-mutant constructs at the NE in G2. A. Hela cells were transfected with GFP WT Nde1, GFP Nde1 phospho-mimetic (Nde1 215E, 243E, 246E) and GFP Nde1 phospho-mutant (Nde1 215A, 243A,246A) and examined for localization at the G2 NE as determined by presence of CENP-F at the NE. Anti-GFP signal co-localized with CENP-F at the NE in cells expressing wt Nde1 and Nde1 phospho-mimetic, though a weak signal was detected in cells expressing Nde1 phospho-mutant. Scale=10μm B. Quantification of the ratio of

G2+ GFP cells in Hela cells transfected with GFP wt Nde1, GFP Nde1 215E, 243E, 246E or GFP Nde1 215A, 243A,246A. The ratio of G2 cells was increased in the GFP Nde1 phospho-mutant condition in comparison to both wt Nde1 or GFP phospho-mimetic.

Since endogenous Nde1/el1 localizes to the kinetochore from prophase and persists into anaphase, we assayed the localization of GFP Nde1 constructs at the kinetochore in all stages of mitosis. GFP wt Nde1 displayed a kinetochore localization pattern consistent with endogenous Nde1/el1 (Figure 2.3A). We found that GFP Nde1 phospho-mimetic localizes to kinetochores from or soon before NEB and persists at these sites though anaphase. Surprisingly, GFP Nde1 phospho-mutant also showed kinetochore localization in prometaphase, though staining appeared weaker than for phospho-mimetic Nde1. More dramatically, GFP-Nde1 phospho-mutant was undetectable from kinetochores of aligned chromosomes in metaphase and during anaphase (Figure 2.3 B,C).

Figure 2.3. Localization of GFP wt Nde1, phospho-mimetic Nde1 and phospho- mutant Nde1 at kinetochores in mitotic cells. A-C. Hela cells were transfected with GFP wt Nde1, phospho-mimetic or phospho-mutant Nde1. In wt Nde1 and phospho-mimetic Nde1 expressing cells, anti-GFP antibody detected signal at kinetochores in prometaphase, metaphase and anaphase cells. GFP wt Nde1 and phospho-mimetic Nde1 co-localize with endogenous CENP-F at the kinetochore from prometaphase into anaphase. In contrast, GFP phospho-mutant Nde1 localizes to kinetochores in prometaphase, but is absent from metaphase and anaphase kinetochores. Scale= 5μm

As a means to confirm the kinetochore localization pattern of GFP Nde1 phospho- mimetic, we performed immunocytochemistry using an antibody raised against a Nde1 peptide corresponding to phosphorylated at residue T246 (Alkuraya et al., 2011; and see

Methods). This antibody recognized kinetochores in prometaphase through into anaphase

(Figure 2.4A). Measurement of average kinetochore fluorescence intensity again

revealed a decrease upon chromosome alignment (Figure 2.4B). Additionally, phospho

Nde1 antibody co-localized with CENP-F at kinetochores (Figure 2.4C).

Figure 2.4. Endogenous Cdk1 phosphorylated Nde1 localizes to the kinetochore. A. Hela cells were stained with p246, a phospho-specific antibody recognizing amino acid 246, which is exclusively phosphorylated by Cdk1(Alkuraya et al., 2012). Phospho Nde1 signal is detected on kinetochores in prometaphase and persists on anaphase kinetochores. B. Quantification of kinetochore p246 signal intensity relative to ACA levels reveals a drop in antibody signal on aligned chromosomes in comparison to unaligned chromosomes. Mean +/- SEM is represented. C. Endogenous Cdk1 phosphorylated Nde1, determined by pT246 phospho-antibody, co-localizes with CENP- F at the kinetochore of 2 μM nocodazole treated cells. Scale = 5μm

In addition to differential temporal localization of Cdk1 phospho-mimetic and

Cdk1 phospho-mutant Nde1, phosphorylation could regulate association with the

kinetochore. We measured the average intensity of GFP Nde1 phospho-mimetic or

phospho-mutant at kinetochores of nocodazole treated cells. Calculations of average

GFP intensity relative to centromere marker ACA intensity reveal Nde1 phospho-mutant has a decreased intensity at the kinetochore in comparison to phospho-mimetic (Figure

2.5A,B). CENP-F kinetochore signal intensity did not differ between conditions (Figure

2.5C). This suggests that Nde1 phosphorylation state could regulate interaction with kinetochore binding protein, influencing the targeting of Nde1 to the kinetochore.

Figure 2.5. Effects of Cdk1 phosphorylation on Nde1 kinetochore localization. A. Hela cells were transfected with cDNAs encoding GFP wt , phospho-mimetic, and phospho-mutant Nde1 and examined for localization to the kinetochore in nocodazole- treated cells. All three versions of the expressed Nde1 clearly localized to kinetochore, though the intensity for the GFP Nde1 phospho-mutant appeared to be decreased. The intensity of the CENP-F signal appeared to be independent of Nde1 phosphorylation state. Bar = 5 μm. B. Quantification of average GFP-Ndel intensity relative to average ACA immunofluorescence signal. Mean +/- SEM of three independent experiments is represented. Anova statistical analysis was performed and a significant difference among

the three conditions was determined (p-value =.0033). Student’s T-test performed between columns indicates a significant difference in the levels of GFP Nde1 depending on phosphorylation state. The difference between WT and the phospho-mimetic Nde1 signal was not was a significant.

Nde1 phosphorylation increases interaction with its kinetochore-binding partner,

CENP-F

Nde1 localization to the kinetochore is dependent on CENP-F, specifically through the direct interaction between C-terminal Nde1 and CENPF (Vergnolle and

Taylor 2007; Liang et al., 2007, Bohley et al 2011). Though CENP-F knockdown does not completely abolish endogenous Nde1/Ndel1 from the kinetochore, endogenous

Nde1/Ndel1 signal is absent from metaphase and anaphase kinetochores (Figure 2.6 A-

C). Because our results show that Nde1 Cdk1 phospho-mimetic increases Nde1 at the

NE and kinetochore in comparison to Nde1 phospho-mutant, we tested whether phosphorylation affects the Nde1 interaction with CENP-F in vitro. In view of the size of

CENP-F we used a bacterially-expressed GST-tagged CENP-F fragment containing its

Nde1-Binding Domain (NBD) (Figure 2.7A, B). GST-CENPF (NDB) was incubated with Hela cell lysate expressing wt Nde1, phospho-mimetic Nde1 or phospho-mutant

Nde1. Each fragment bound to the CENP-F fragment, though levels of GFP phospho- mimetic Nde1 bound were increased by ~2 fold (Figure 2.7C). As an alternative approach, bacterially-expressed full-length Nde1 was phosphorylated in vitro with recombinant CyclinB/Cdk1 then incubated with GST-CENP-F NBD. We observed an

~2.5 fold increase in levels of Nde1 pulled down by GST-CENP-F NBD in the phosphorylated condition (Figure 2.7D). Together these experiments suggest that Cdk1 phosphorylation of Nde1 activates binding to the NE and kinetochore through CENP-F.

Figure 2-6. CENP-F siRNA decreases endogenous Nde1/el signal from kinetochores. A. Hela cells were treated with 2μM Control siRNA or 2μM CENP-F siRNA for 48 hours and endogenous Nde1/el1 kinetochore signal was determined in 5μM nocodazole treated cells. Depletion of endogenous CENP-F signal in CENP-F knockdown indicates effectiveness of CENP-F siRNA. B. Quantification of endogenous

Nde1/el1 fluorescence signal intensity relative to ACA for control siRNA or CENP-F siRNA conditions reveals a decrease in Nde1/el1 signal in CENP-F siRNA condition. Average ratio of Nde1/el1/ACA is represented +/- SEM. Student’s T-Test was performed on 3 replicates of ~25 kinetochores/ cell for ~20 cells and a significance of <.0001 was calculated. There is a significant decrease in endogenous Nde1/el1 kinetochore intensity in cells treated with CENP-F siRNA. C. Staining of endogenous Nde1/el1 in mitotic cells treated either with 2μM control siRNA or 2μM CENP-F siRNA reveals Nde1/el1 signal is lost from kinetochores in metaphase and anaphase cells but Nde1/el1 is present on prometaphase kinetochores.

Figure 2-7. Biochemical analysis of Cdk1 effects on the Nde1-CENP-F interaction. A. Schematic diagram of the CENP-F fragment used for biochemical analysis (NBD: Nde1 binding domain, amino acids 2122-2297) B. GST pull-down of Nde1 by GST CENP-F NBD fragment. C. GST CENP-F NBD pull down of GPF Nde1 constructs from Hela cells expressing wt Nde1, Nde1 phospho-mimetic or Nde1 phospho-mutant.

Quantification of GFP Nde1 in GST CENP-F NBD pellet demonstrates an increased amount of GFP Nde1 phospho-mimetic. D. GST pull down of CENP-F NBD with HA- Nde1 following phosphorylation of Nde1 using recombinant in the presence or absence of Cdk1/CyclinB. Levels of Nde1 in the CENP-F NBD pull-downs were quantified and revealed increased amount of Cdk1/Cyclin B phosphorylated Nde1 in the GST CENP-F NBD pellet. ** p<.01.

Role of Nde1 phosphorylation in mitosis

Our results indicate that Nde1 phosphorylation enhances its recruitment to the NE and kinetochore generally through an increased interaction with kinetochore binding partner CENP-F. We sought further to determine whether phosphorylation regulates

Nde1 mitotic activity. Live-cell imaging of Hela H2b-Cherry cells was performed to analyze the effect of over-expression of Nde1 phosphorylation constructs on mitotic progression (Figure 2.8A-E). Cells expressing GFP Nde1 phospho-mutant (215A, 243A,

246A), spent an average 2X longer from NEBD to alignment as well as time spent in metaphase (average unaligned=67.14 min; average aligned=53.91min). The delay in the unaligned state was often due to lack of compact chromosome alignment at the metaphase plate. The Nde1 phospho-mimetic (215E, 243E, 246E) expressing cells displayed a slight increase in the average time spent at metaphase (aligned- ~27min), though not significantly different from wt Nde1 or GFP conditions. Monitoring of H2B

Cherry cells expressing the Nde1 phosphorylation constructs demonstrates the dominant negative effect of Nde1 phospho-mutant causing defects in chromosome alignment and metaphase arrest.

To further investigate the requirement of Nde1 Cdk1 phosphorylation constructs, rescue experiments were performed. Reduction of Nde1 by siRNA decreased endogenous levels of Nde1 from Hela cells (Figure 2.9G) Nde1 Cdk1 phosphorylation constructs could rescue Nde1 knockdown mitotic progression phenotype. Cells were

treated for 48 hours with Nde1 siRNA followed by the addition of GFP constructs 24

hours before imaging. Nde1 RNAi caused a significant delay in anaphase onset, as has

been observed using Nde1 siRNA, treatment with function-blocking AB or GFP Nde1 dominant negative contructs (Raaijimakers, vergnolle, stehman). Surprisingly, Nde1 siRNA rescue with wt Nde1, Nde1 phospho-mimetic (215E, 243E, 246E) and Nde1 phospho-mutant (215A, 243A, 246A) all decreased the time in mitosis to similar extents.

There was no significance difference significant difference in the average time from nuclear envelope breakdown until anaphase onset between the Nde1 phosphorylation conditions. There is a ~15min increase in mitotic duration when Nde1 siRNA is rescued with Nde1 phospho-mutant construct (215A, 243A, 246A) in comparison to wt Nde1 and

Nde1 (215E, 243E, 246E). Specifically there was a subtle increase in the unaligned and aligned; NUMBERS (Figure 2.9 D, E, F). This slight delay in anaphase onset could indicate a subtle role for the phosphorylated population of Nde1 in prometaphase and metaphase Nde1 functions but both Nde1 phospho-mutant and Nde1 phospho-mimetic

have the ability to rescue Nde1 mitotic progression defects to the same levels as WT

Nde1 (Figure 2.9 E,F).

Figure 2.8. Effects of over expression of GFP Nde1 phospho constructs on mitotic progression. A-D. H2B cherry Hela cells were transfected with either GFP vector or GFP Nde1 constructs for 24 hours and then imaged for the occurrence of mitotic events. The time from NEBD to alignment (unaligned) was measured as well as the time spent in

metaphase prior to anaphase onset (aligned). Each mitotic event for GFP, wt Nde1, phospho-mimetic (Nde1 215E, 243E, 246E) or phospho-mutant (Nde1 215A, 243A, 246A) expressing cells were graphed. ** denotes cells that did not enter anaphase throughout the duration of the movie. Scale = 10μm E. Quantification of the average time spent from NEBD to anaphase onset. Cells expressing Nde1 phospho-mutant on average spent a longer time from NEBD to anaphase in comparison to GFP, wt Nde1 or phospho-mimetic Nde1. Summary of the time in unaligned vs. aligned: Nde1 215A, 243A, 246A-67.72 min, 53.59 min; Nde1 215E, 243E, 246E-32.91 min, 27.70 min; wt Nde1-34.50 min, 13.68 min; GFP-35.31 min, 12.15 min) Representative images of Hela H2B cherry cells expressing GFP, GFP wt Nde1, GFP Nde1 phospho-mimetic or GFP Nde1 phospho-mutant. Image panels represent time point 9 minutes apart. Scale =10µm A.

Figure 2.9. Rescue of Nde1 siRNA mitotic progression defects with wt, phospho-mimetic or phospho-mutant Nde1. A. Live cell imaging of H2B cherry Hela cells treated with control siRNA or Nde1 siRNA. Representative images of control siRNA or Nde1 siRNA treated cells during mitosis. Scale = 10μm B. Graphical representation of time spent from NEBD to alignment (unaligned) and time spent in metaphase (aligned) for either control siRNA treated cells or Nde1 siRNA treated cells. Each bar represents a single

mitotic event. ** denotes cells that did not enter anaphase throughout the duration of the movie. C-D. Images of mitotic events of H2B cherry Hela cells treated for 48 hours with Nde1 siRNA then transiently transfected with wt Nde1, Nde1 phospho-mimetic (Nde1 215E, 243E, 246E) or Nde1 phospho-mutant (Nde1 215A, 243A, 246A). Quantification of time spent from NEBD to alignment (unaligned) and time spent in metaphase (aligned) for each condition. Each bar represents one mitotic event E. Summary of the total time from NEBD to anaphase onset for each condition. Scale = 10μm A Non-parametric Mann-Whitney test was run to compare the means between rescue constructs. There is no significant difference between the conditions. F. Summary of the time spent with chromosomes unaligned or aligned for control siRNA treated cells, Nde1 siRNA treated cells, or Nd1 siRNA cells transiently transfected with either wt Nde1, phospho-mimetic or phospho-mutant Nde1. G. Endogenous Nde1 protein levels in Hela cell lysate either expressing control siRNA or Nde1 siRNA. Cells were harvested after 48 hours.

Role of Nde1 in Anaphase A

Unique to Nde1/el relative to cytoplasmic dynein mitotic localization is the persistence of endogenous and Cdk1 phosphorylated Nde1 on kinetochores in anaphase. Since dynein

has been reported to decrease the velocity of chromosomes in anaphase A by ~30%

(Howell et al. 2000), we tested whether Nde1 has a similar role in anaphase. H2B Cherry

Hela cells were treated with control siRNA or Nde1 siRNA for 48 hours before imaging.

Aligned cells were imaged, following treatment with Cdk1 inhibitor RO-3306 to

stimulate anaphase onset. Measurement of maximum chromosome mass distance in the first 2.5 minutes of anaphase (anaphase A) was used to determine the chromosome velocity in each condition. Cells treated with control siRNA had an average anaphase A velocity of ~1.5μm/min, which is similar to previous reports (Reider and Salmon; 1998)

while cells treated with Nde1 siRNA exhibited an average anaphase A velocity of .998

μm/min; a decrease of ~25% (Figure 2-10 A, B, E).

Figure 2.10 Role of Nde1 in Anaphase A-D. H2B Cherry Hela cells were treated with control siRNA or Nde1 siRNA for 48 hours prior to imaging. Aligned cells were located prior to addition of 9uM RO-3306, which was used to simulate anaphase onset. One minute post drug application, movies were acquired every 30 seconds and chromosome movements in anaphase were monitored. To determine the rate of chromosome separation in anaphase A, maximal inter-chromosome distance was measure the first 2.5 minutes. A-B. Representative trace of chromosome behavior in control siRNA and Nde1 siRNA treated cells. Average anaphase A velocity of control siRNA was 1.556 μm/min (n=25 cells) while Nde1 siRNA average velocity was .998 μm/min (n=34 cells). Scale = 10μM C. Representative images of anaphase movies of Nde1

siRNA treated cells rescued with GFP wt Nde1, phospho-mimetic Nde1 or phospho- mutant Nde1. Scale = 10μM D. Average anaphase velocities for Nde1 siRNA treated cells rescued with wt Nde1 (1.67 μm/min, n=7), phospho-mimetic Nde1(1.121 μm/min, n=7), or phospho-mutant Nde1 (1.47μm/min n=7). E. Comparison of average Anaphase A velocities.

To test the role of Nde1 phospho-mimetic and phospho-mutant on anaphase A behavior, we expressed GFP wt, phospho-mimitic or phospho-mutant Nde1 in cells treated with Nde1 siRNA. GFP wt Nde1 and phospho-mutant Nde1 expressing cells had average Anaphase A velocities of 1.67 μm/min and 1.47 μm/min, respectively (Figure 2-

10 C,D, E). These values are comparable to that measured for control siRNA cells, indicating that wt Nde1 and phospho-mutant Nde1 rescue Nde1 siRNA anaphase A defects. In comparison, Nde1 siRNA cells expressing Nde1 phospho-mimetic had an average anaphase A velocity of 1.121 μm/min. (Figure 2-10 C,D, E). GFP Nde1 phospho-mimetic did not rescue Nde1 siRNA anaphase A velocity defects.

Figure 2.11. Summary of the nuclear envelope and kinetochore localization of Cdk1 phosphorylated Nde1. Cdk1 phosphorylated Nde1 (Nde1***) localizes to the nuclear envelope in late G2 and prophase, while dephosphorylate Nde1 (Nde1) has a weaker association. During mitosis, Cdk1 phosphorylated Nde1 binds to the kinetochore in prometaphase, metaphase and anaphase. In contrast, dephosphorylated Nde1 only is detected at the prometaphase kinetochore.

Discussion

Nde1 is a known target of Cdk1 kinase activity, but the effects of phosphorylation on its subcellular localization and mitotic behavior have remained largely unexplored.

We find that Cdk1 phosphorylation targets Nde1 to the late G2 NE and to mitotic kinetochores from prophase through mid-anaphase (Figure 2.11). This behavior appears to be mediated by a Cdk1-enhanced interaction between Nde1 and CENP-F. Phenotypic studies reveal Nde1 phospho-mutant (Nde1 215A, 243A, 246A) has a dominant negative effect, causing cells to exhibit a late prometaphase or metaphase arrest. RNAi rescue

experiments show that both phosphorylated as well as dephosphorylated Nde1 function

during mitosis, and suggest that Cdk1 phosphorylation regulates Nde1 behavior

throughout late G2 and mitosis.

Role of Nde1 phosphorylation in G2-M Phase Targeting and CENP-F binding

We find that Cdk1 phosphorylation state of Nde1 dictates Nde1’s NE and kinetochore localization in G2/M. Phospho-mimetic and wt Nde1 localized to the G2 NE

and mitotic kinetochores, in the latter case apparently from before NEBD to anaphase.

Use of an Nde1 phosphorylation site-specific antibody (Fig. 1) clearly supported the

association of Cdk1 phosphorylated Nde1 with the kinetochore. In contrast, the Cdk1

phospho-mutant Nde1 showed weak labeling at the NE and a kinetochore signal on

prometaphase kinetochores but absent at metaphase and anaphase. This pattern is

consistent with decreased kinetochore association upon end-on MT attachment like other

dynein-related proteins. The differential localization of phospho-mimetic and phospho-

mutant Nde1 is the first report of Cdk1 phospho-regulation of localization of dynein

adaptor proteins in mitosis.

A key finding of the current study is that Cdk1 phosphorylation enhances the

interaction of Nde1 with CENP-F. Both in vivo-expressed phospho-mimetic Nde1 and in

vitro Cdk1-Cyclin B1 phosphorylated Nde1 were observed to interact more strongly with

a recombinant CENP-F fragment in comparison to in-vivo expressed phospho-mutant

Nde1 or non-phosphorylated Nde1. This is the first evidence for protein kinase-mediated

activation of Nde1 or Ndel1 cargo binding and its regulation of sub-cellular targeting.

Given the reported role of Cdk5 phosphorylation in Ndel1 or Nde1 function in axonal

transport (Pandey et al., 2010; Klinman and Holzbaur, 2015), phosphorylation could be regulating the interaction between Nde1 and Ndel1 and vesicular factors as well.

Additionally, in vivo, phospho-mimetic Nde1 showed stronger signal at the mitotic kinetochore relative to Nde1 phospho-mutant, presumably through an increased interaction with CENP-F. It is possible that over-expression of the phospho-mutant Nde1 artificially increases its levels at kinetochores. A favored model is that since CENP-F siRNA does not completely abolish endogenous Nde1/Ndel1 from the kinetochore, an additional Nde1 kinetochore recruitment factor, potentially zw10 (Stehman et al., 2007), could be responsible for anchoring phospho-mutant to the kinetochore. Phosphorylation state of Nde1 could dictate Nde1 to specific mitotic cargo.

Endogenous Nde1/Ndel1 have a unique kinetochore pattern of the dynein-related proteins. Along with CENP-F, Nde1/Ndel1 remain at the kinetochore into anaphase. In addition, Nde1/Ndel1 signal declines ~50% upon chromosome alignment. To confirm the decrease in kinetochore Nde1 from prometaphase to metaphase, a GFP wt Nde1 stable cell line could allow for the visualization of GFP-wt Nde1 on kinetochores. FRAP recovery experiments of GFP wt-Nde1 turnover at unaligned or aligned kinetochores, would allow for quantitative insight into Nde1 kinetochore dynamics.

Cdk1 phosphorylated Nde1, monitored by both over-expression of phospho- mimetic or phospho-specific AB, reveal a strikingly similar kinetochore pattern. Cdk1 phosphorylated Nde1 persists at the anaphase kinetochore, even when Cdk1 levels have dropped. Additionally, we found that phosphorylated Nde1, monitored using the phospho-specific antibody, showed a ~50% decline in signal upon chromosome

alignment. This suggests that the Cdk1 phosphorylated form of the protein is

predominant at kinetochores both prior to chromosome alignment and into anaphase.

The decrease in p-Nde1 at the aligned chromosomes may be a simple reflection of

the reduced levels of its recruitment protein. We find that CENP-F kinetochores levels

drop by ~half upon chromosome alignment (Data not shown). Cytoplasmic dynein, at

least some of its regulators, and spindle assembly checkpoint (SAC) proteins are

substantially depleted upon chromosome alignment and presumably end-on MT

attachment. It seems reasonable that the partial loss of both phospo-Nde1 and CENP-F might be mediated by dynein-mediated poleward transport mechanism. It is important to

note, CENP-F and Nde1/Ndel1 do not have the classic streaming/spindle localization

pattern like dynactin or Lis1. An alternative possibility is that Nde1/Ndel1 and CENP-F

release from the kinetochore and remain at the precursor to the spindle mid-zone for post-

metaphase functions. Further research is required to explore the role of non-kinetochore

bound Nde1 and CENP-F.

How Cdk1-phosphorylated Nde1 can persist at anaphase kinetochores despite the

well-established drop in Cdk1 activity at anaphase onset is unknown. An appealing

possibility involves a potential role for member proteins of the 14-3-3 family, which

protect Ndel1’s Cdk 1/5 sites from dephosphorylation (Toyo-oka et al., 2003). Nde1 has been reported to interact with 14-3-3e, in a phospho-dependent manner (Kimura et al.,

2014; Lu and Prehoda, 2013). The interaction between Nde1 and 14-3-3e could allow for a pool of Nde1 to resist kinetochore-associated phosphatase activity and ultimately allow for the persistence of Nde1 at the kinetochore into anaphase.

Functional Analysis

Phenotypic analysis of Nde1 siRNA and Cdk1 phospho constructs implicates

phosphorylation of Nde1, not only in regulating Nde1 localization to the kinetochore, but

also Nde1’s mitotic function.

Examination of cells expressing Nde1 phosphorylation construct reveals an increase in G2+ cells in the Nde1 phospho-mutant condition. This result is consistent

with a previous report implicating a single Cdk1 phospho-mutant Nde1 construct (Nde1

246A) in increasing the percentage of G2 cells (Alkuraya et al., 2011). Since defects in

dynein and dynein accessory proteins do not cause a block in mitotic entry in mammalian

culture cells (Salinas et al., 2000), the increase in ratio of cells in G2 could reflect a delay

in cell cycle progression. One possibility is that Cdk1 phosphorylation of Nde1 aids in

NE dynein functions during G2/prophase, since Ndel1 Cdk1 and Cdk5 phosphorylation

was demonstrated to have a role in the formation of prophase nuclear envelope

invaginations (Hebbar et al., 2008). Further examination of defects in prophase

centrosome separation or centrosome-nucleus anchoring is required to elucidate the role

of Cdk1 phosphorylation of Nde1 in Nde1 and dynein G2/prophase functions.

Over-expression of Nde1 phosphorylation constructs reveals Nde1 phospho-

mutant causes a delay in anaphase due to a prolongment in late prometaphase and a

metaphase arrest. This could be caused by either defects in formation of stable end-on

MT attachments or stabilization of KT-MT attachments. In multiple cells, chromosomes did not form a tight alignment at the metaphase plate and often exhibited more exaggerated chromosome movements upon alignment, suggesting a possible lack of KT-

MT tension and stabilization. For mechanistic insight into the chromosome behavior of

Nde1 phospho-mutant expressing cells, oscillations of kinetochore pairs in CENPA-YFP

expressing Hela cells could be quantified. Increased oscillations of kinetochore pairs in

the phospho-mutant would suggest phosho-Nde1 is required for dynein’s role in KT-MT stabilization at the metaphase kinetochore. In contrast, Cdk1 phosphorylated Nde1 does not play a role in the SAC inactivation of dynein, since Nde1 phospho-mutant expressing cells do not enter anaphase with incomplete MT attachments since no lagging chromosomes in anaphase were detected.

GFP phospho-mutant Nde1 signal at the kinetochore is decreased in comparison to GFP wt Nde1 or GFP phospho-mimetic Nde1. How this decreased kinetochore localization is affecting the over-expression phenotype of phospho-mutant Nde1 is

unknown. One possibility is that Nde1 phospho-mutant is sequestering an essential

binding partner independent of CENP-F. Proteomics analysis of differential interactions

between phospho-mimetic and phospho-mutant Nde1 will provide clues into unique

interactions. Potential candidates are zw10, as well as Lis1 and dynein. Alternatively,

expression of phospho-mutant Nde1 could displace of endogenous Nde1, presumably both Cdk1 phosphorylated Nde1 and dephosphorylated Nde1. The defects observed in

expression studies would be a result of the lack of phosphorylated Nde1 at the

kinetochore and clearly indicate a role for Cdk1 phosphorylated Nde1 at the kinetochore.

Testing for endogenous phosphorylated Nde1 localization at the kinetochore in Nde1

phospho-mutant cells would provide clues whether the over-expression of Nde1 phospho-

mutant is displacing endogenous Nde1.

A subtle delay in anaphase onset was observed in Nde1 phospho-mimetic cells,

specifically an increase in the average time in metaphase in comparison to GFP or GFP

wt Nde1 conditions. This suggests the possibility for a role for Nde1 phospho-mutant in

Nde1’s mitotic function, though further characterization will be required to determine the

exact role of phospho-mutant Nde1.

Nde1 knockdown caused a delay in anaphase onset, consistent with prior reports

(Vergnolle and Taylor, 2007; Raaijmakers et al., 2013). Cells exhibited both a

prolongment in chromosome alignment and time spent aligned at the metaphase plate

consistent with previously reported roles in dynein-dependent functions of MT

attachment and stabilization (Stehman et al., 2007). Surprisingly, wt, phospho-mimetic

and phospho-mutant Nde1 all rescued mitotic progression delays, albeit incompletely,

perhaps reflecting incomplete rescue. This suggests that both phospho-mimetic and

phospho-mutant have the activity required to rescue Nde1 mitotic defects.

Rescue with wt Nde1 does not decrease the time in mitosis to control siRNA treated levels, potentially a reflection of incomplete rescue since over-expression of wt

Nde1 does not cause delayed mitotic progression phenotype. In addition to the kinetochore, Nde1 has roles at the centrosome and spindle pole (Yan et al., 2003;

Hirohashi et al., 2006b). Nde1 siRNA could cause a disruption of the mitotic spindle not completely rescued by expression of wt Nde1 or Nde1 phospho-constructs. This could be reflected in the out-lying time points in each condition. Given multiple functions of

Nde1 in mitosis at the kinetochore and centrosome, it is important to remember rescue experiments reflect Nde1 phospho-mutant’s and phospho-mimetic’s role at both of these sites and defects are not just indicative of the kinetochore fraction of Nde1.

The physiological requirement for the CENP-F-phospho-Nde1 interaction at kinetochores throughout mitosis may involve the fidelity of MT capture and stability of the MT-KT interaction. CENP-F depletion causes chromosome misalignment,

suggesting a role in efficient and correctly orientated MT attachments potentially through

its MT binding ability (Holt et al., 2005; Feng et al., 2006; Yang et al., 2005; Volkov et

al., 2015). As proper MT attachment is reduced in Nde1 knockdown cells and potentially

in phospho-mutant Nde1 over-expression, we reason that phospho-Nde1 may contribute with CENPF in this function. The persistence of CENP-F and phospho-Nde1 perhaps

Nde1 serves in a fail-safe role, available even upon anaphase onset to recruit cytoplasmic dynein back to kinetochores that might have lost their attachment. Residual Nde1 at kinetochores might acquire dynein without delay to rescue attachment errors. Further work will be required to test these important possibilities.

Post-Metaphase roles of Nde1

To begin to clarify the role of Nde1 in post-metaphase functions, we examined the effect of Nde1 siRNA on anaphase A velocity since disruption of dynein and dynein adaptor protein zw10 caused defects in anaphase A velocity. Nde1 siRNA caused a decrease in anaphase A velocity. Unlike Nde1 siRNA treated H2B rescued with Nde1 siRNA with GFP wt Nde1 or GFP Nde1 phospho-mutant, cells rescued with GFP Nde1 phospho-mimetic had a slower average anaphase A velocity. This would suggest that the kinetochore localization of Cdk1 phosphorylated Nde1 is not essential for its function in anaphase A. The mechanism of dynein’s role in anaphase A is unknown. It is possible that additional sub-fractions of Nde1 localized to the centrosome and spindle pole are contributing to Nde1’s anaphase function by influencing forces on the mitotic spindle.

GFP Nde1 phospho-mimetic has decreased localization to the centrosome and spindle pole in comparison to GFP Nde1 phospho-mutant (Yan et al., 2003;Hirohashi et al.,

2006b). Although Nde1 phospho-mimetic is present on the kinetochore, the decreased

localization at the centrosome and spindle pole could be contributing to inability to rescue Nde1 siRNA anaphase A velocity defects while GFP Nde1 phospho-mutant is able to rescue the defects.

The function of the CENP-F-Nde1 interaction in the later stages of mitosis is also uncertain, though CENP-F and Nde1/Ndel1 can be detected at the central spindle in late anaphase and telophase cells. Nde1’s localization to the mid-zone and function in post- metaphase roles has not been reported or investigated. CENP-F c.elegans homolog

HCP1/2 has been reported to have a role in central spindle initiation (Maton et al., 2015), though a similar role in mammalian cells will require further research to clarify as well as implication of Nde1 in this role. As mentioned previously, the maintenance of CENP-F- phospho Nde1 at the kinetochore in anaphase might be to reserve a sub fraction of Nde1 at the kinetochore for residual MT-attachment functions. Further experiments will clarify this role of the Cdk1 phosphorylated Nde1 and determine whether Cdk1 dephosphorylated Nde1 has a non-centrosome localization pattern that could contribute to a central spindle function of Nde1.

Chapter 3: Structure-function dissection of Nde1’s role in mitosis

Introduction

The previous chapter explored how Nde1 phosphorylation by Cdk1 increased

Nde1’s interaction with its nuclear envelope (NE) and a kinetochore recruitment factor,

CENP-F. The importance of phosphorylation in controlling Nde1’s recruitment to mitotic sites was demonstrated as a Nde1 phospho-mimetic construct specifically localized to the NE in G2 and to the mitotic kinetochore from prophase into anaphase.

However, the regulation of Nde1’s localization at the NE and kinetochore is ultimately only important as far as it allows the cell to modulate the activity of cytoplasmic dynein at these sites in a cell cycle specific fashion.

The precise coordination of cytoplasmic dynein’s activity at the kinetochore during mitosis affects several crucial steps necessary for proper chromosome segregation including chromosome congression (Yang et al., 2007), stabilization of kinetochore- microtubule attachments during metaphase (Varma et al., 2009), and removal of the spindle assembly checkpoint (SAC) proteins (Howell et al., 2000). In order to accomplish these diverse functions during mitosis, dynein requires interactions with multiple cofactors including Nde1/el1, dynactin, Lis1 and the kinetochore proteins Zw10 and Spindly (Stehman et al., 2007; Gassman et al., 2010; Starr et al., 2000; Faulkner et al., 2000). For some of these interactions, the molecular consequences are well known— for example Zw10 and Spindly interact with dynein to contribute to dynein’s role in the removal of the spindle assembly checkpoint protein Mad2 upon MT attachment (Chan et al., 2000; Buffin et al., 2005)—but questions remain about the exact mechanism of

Nde1’s interaction with dynein and its roles in specific mitotic functions.

Inhibition of Nde1, both by siRNA or Nde1/Ndel1 function blocking AB, has

demonstrated Nde1’s involvement in multiple dynein-dependent mitotic functions including SAC inactivation through removal of Zw10, chromosome alignment at metaphase, and formation of stable kinetochore-MT attachments (Stehman et al., 2007;

Vergonolle et al., 2007; Liang et al., 2007). Nde1 depletion might produce these

phenotypes though multiple different physical interactions as both in vitro and in vivo

studies show Nde1 can play a role in modulating dynein’s activity through direct

interaction with dynein intermediate chain as well as the recruitment of Lis1.

Additionally, in vitro, Nde1 interacts with both dynein and Lis1 via its N-terminal coiled-

coil domain and this tri-protein complex of Nde1-Lis1-Dynein transforms dynein into a persistent force-producing motor, and in vivo this complex is required for high force production of dynein in lipid droplet motility (McKenney et al., 2010; Reddy et al.,

2016).

To address the role of the direct interactions between Nde1 and Lis1/dynein, we

created point mutations in Nde1 to target either Nde1’s Lis1 interaction or its dynein

interaction. Through molecular dissection of Nde1’s interactions with dynein and Lis1

we uncover the distinct functions for each interaction. Phenotypic studies of Nde1

mutants reveal that the Nde1-del dynein mutant causes severe defects in mitotic

progression while Nde1-del Lis1 mutant causes only a slight increase in the time from

NEBD to anaphase onset. Biochemical assays show Nde1 directly recruits dynein to the

kinetochore through its N-terminal coiled-coil domain and this direct interaction is required for Nde1’s role in stabilization of MT-KT at metaphase. While interestingly,

Lis1 binding to Nde1 influences Nde1’s localization to the kinetochore through Nde1’s

interaction with CENP-F. By genetically separating the two main interactions of Nde1

we demonstrate how each is required for unique aspects of Nde1’s function.

Results

Characterization of the Nde1-Dynein and Nde1-Lis1 interactions

Construction of Nde1 mutants specifically targeting either the dynein or Lis1

interactions requires precise knowledge of the sites of interaction within the Nde1

molecule.

Previously, disruption of the Nde1-dynein interaction was accomplished by a

deletion in the C-terminal domain of Nde1, specifically amino acids 257-291, since this

region was identified as the dynein-interacting region in yeast-two hybrid experiments

(Sasaki et al., 2000). Discovery of an additional dynein-binding domain within Nde1’s

N-terminal coiled-coil domain complicates the presence of a single dynein-binding site

(Zyłkiewicz et al., 2011; Wang et al., 2011). We performed biochemical pull-down experiments with recombinant Nde1 fragments and point mutations to clarify the role of

Nde1’s N-terminal or C-terminal domains in Nde1’s dynein-binding ability. A series of

Nde1 truncations were created to target the previously reported C-terminal dynein- binding domain (amino acid 252-291); Nde1 1-291 maintains the entire c-terminal dynein-binding domain, while Nde1 fragments 1-248, 1-218 and 1-191 completely lack the C-terminal dynein binding site. To test of the role of the Nde1 fragments on Nde1’s dynein binding activity, we performed HA immuno-precipitations of recombinant HA tagged Nde1 in the presence or recombinant dynein intermediate chain fragment (IC 1-

250) or purified dynein complex (Figure 3.1A-C). Western blot analysis reveals each HA

Nde1 construct bound to both dynein intermediate chain and purified dynein complex.

These observations suggest the C-terminal domain of Nde1 is not required for Nde1’s

interaction with dynein.

To target the N-terminal dynein-binding domain and assess its role in Nde1’s dynein binding activity, we mutated two amino acids in the coiled-coil region, amino acid

T48 and amino acid T51 (Zyłkiewicz et al., 2011). GST tagged wt Nde1 or Nde1 47.51A was incubated with bovine brain purified dynein. Wt Nde1 pulled down dynein, while

Nde1 47.51A did interact with the purified dynein complex (Figure 3.1D). The HA immuno-precipitation experiment suggests the N-terminal dynein-binding domain, specifically amino acids T47 and T51, is essential for Nde1’s interaction with dynein.

To further confirm of the role of the N-terminal of Nde1 in its interaction with dynein, we tested ability to form a tri-protein complex with CENP-F, Nde1 and dynein

IC. Nde1 interacts with CENP-F via is C-terminal unstructured region, which overlaps the predicted C-terminal dynein binding domain (Vergnolle and Taylor, 2007). Any IC pulled down by Nde1 bound to CENP-F should be via the N-terminal coiled-coil domain since the C-terminal domain is occupied by CENP-F. GST-pull down experiments were performed to test this hypothesis. GST-CENPF fragment containing only the Nde1 was incubated with Nde1, washed, and then incubated with dynein IC fragment. Intermediate chain only pulled down with GST-CENP-F in the presence of Nde1 (Figure 3.1E). The formation of a triple complex between Nde1, CENP-F and IC suggests that Nde1 can interact with both IC and CENP-F simultaneously and is able to recruit dynein when bound to CENP-F.

Figure 3.1 Biochemical characterization of Nde1’s interaction with dynein. A. Schematic diagram of Nde1 domains. The N-terminal coiled-coil domain contains a dynein-binding site and a lis1-binding site. The C-terminal unstructured domain contains multiple phosphorylation sites (*), around amino acid 219-246. The C-terminus contains LC8 bind site and a previously identified C-terminal dynein-binding site. A summary of the results of HA immuno-precipitations where HA-Nde1 truncations were incubated with either recombinant intermediate chain fragment (IC 1-250) or bovine brain purified dynein. Each of the C-terminal truncations was able to interact with both dynein complex and recombinant IC. B. HA-tagged Nde1 constructs were recruited to protein A beads. HA- Nde1 constructs recruited to monoclonal HA conjugated protein A beads were incubated with bacterially purified N-terminal IC fragment (1-250) or bovine brain purified dynein complex. C. Recombinant Nde1 or Nde1 del dynein (Dynein 47.51AA) were recruited to protein A beads via HA antibody and incubated with bovine purified dynein complex. HA-Nde1 del dynein did not interact with purified dynein complex. D. A GST pull-down experiment with GST CENP-F NBD pull down of dynein intermediate chain in the absence or presence of Nde1. Dynein IC only pulls down with CENP-F NBD in the presence of Nde1.

Biochemical experiments validated the N-terminal dynein-binding domain as the main interacting site between Nde1 and dynein. A mammalian expression construct, named GFP Nde1 del dynein, containing point mutations at amino acid 47 and 51 to

abolish the N-terminal dynein-binding domain, was created to study in vivo the affect of

this mutation (Figure 3.2A). Interestingly, immuno-precipitation of GFP Nde1 constructs, never detected an interaction with dynein. Alternatively, to confirm the lack of dynein-binding activity in the Nde1 del dynein construct, we expressed GFP- intermediate chain and tested for an interaction with recombinant Nde1, either GST-WT

Nde1 or GST-Nde1 del dynein (47.51AA). The levels of dynein that interacted with

Nde1 decreased in the Nde1 del Dynein condition (Figure 3.2 C).

Figure 3.2 Characterization of Nde1 mutant constructs used to study the mitotic role of Nde1’s interaction with Lis1 or dynein. A. Diagram representations of the Nde1 mutants used in this study, specifically Nde1 del dynein, which has two alanine mutations at amino acid 47 and 51, and Nde1 del Lis1, which as three alanine mutations—119A, 127A,129A. B. GFP Immuno-precipitation of GFP, GFP wt Nde1, GFP del dynein or GFP del Lis1 reveals a loss of pull-down of endogenous Lis1 in the GFP alone and GFP Nde1 del Lis1 condition. C. GST pull down of dynein intermediate chain from cells

expressing GFP-IC2C with either GST wt Nde1 or GST Nde1 47.51AA. GFP IC2C only pulls down with GST wt Nde1.

To specifically abolish the interaction between Nde1 and Lis1, three amino acids

(119,127,129) were mutated in the third coiled-coil region of Nde1, specifically due to their reported role in decreasing the biochemical interaction with Lis1 (Derewenda et al.,

2008). Confirmation of proof-of-principle was achieved through anti-GFP immuno-

precipitation with Hela cell lysate expressing either GFP WT, GFP del Lis1, or GFP del

Dynein. In both GFP wt Nde1 and GFP Nde1 del dynein conditions, an interaction with

Lis1 was detected. An interaction with Lis1 was not detected in the GFP-Nde1 Del Lis1 condition (Figure 3.2B). The Nde1 mutants were confirmed to disrupt their respective interactions and validated as tools to dissect the role of Nde1’s direct interaction with either dynein or Lis1.

Importance of Nde1’s interaction with dynein or Lis1 in mitotic progression

To determine how Nde1’s interaction with either Lis1 or dynein influenced

Nde1’s role during mitosis, we performed live-cell imaging experiments of H2B cherry

Hela cells expressing GFP Nde1 constructs to assay defects in mitotic progression.

Mammalian expression constructs, GFP, GFP wt Nde1, GFP Nde1 del dynein or GFP del

Lis1, were transfected into cells stable-expressing H2B cherry and monitored for the time spent from nuclear envelope breakdown to anaphase onset. Expression of GFP Nde1 del dynein significantly increased the time in mitosis, with an average time measured to be

189 min. Cells exhibited both a delay chromosome alignment and a metaphase arrest

(Figure 3.3A-E). There was a slight delay in mitotic progression seen in cells expressing

Nde1 del Lis1. The average time from NEBD to anaphase onset increased from 48.18 min in wt Nde1 expressing cells to 81.34 min. There was a slight increase in the time

spent in alignment, consistent with reported role for Lis1 in mitosis (Faulkner et al.,

2000). Over-expression studies of Nde1 mutants shows expression of GFP Nde1 del dynein causes significant mitotic progression defects suggesting an essential role for this interaction.

Figure 3.3 Effects of over expression of Nde1 mutants on mitotic progression. A- D. H2B cherry Hela cells were transfected with either GFP vector or GFP Nde1 mutant constructs for 24 hours and then imaged for the occurrence of mitotic events. The time

from NEBD to alignment (unaligned) was measured as well as the time spent in metaphase prior to anaphase onset (aligned). Each mitotic event for GFP, wt Nde1, Nde1 del dynein or Nde1 del Lis1 expressing cells was graphed. Note the scale in the Nde1 del Dynein condition is in two segments to accommodate long time points measured. E. Quantification of the average time spent from NEBD to anaphase onset, separate by time from NEBD to alignment (unaligned) and time spent aligned until anaphase onset (aligned) for each condition. Nde1 del dynein-expressing cells showed an increase a large increase in average time from NEBD to anaphase onset in comparison to GFP, wt Nde1 or Nde1 del lis1 conditions. Nde1 del LIs1-expressing cells show an increase in average time (NUMBERS), though not as severe as the Nde1 del dynein condition.

To further confirm the role of Nde1 dynein-binding and lis1-binding mutants in mitosis, Nde1 rescue experiments were performed (Figure 3.4). As discussed in the previous chapter, Nde1 siRNA causes a significant increase in the time from NEBD to anaphase onset. Rescue of Nde1 siRNA treated cells with GFP Nde1 del. Lis1 slightly increased the average time until anaphase onset (86.45 min), but the difference was not significantly differently from Nde1 siRNA rescued with GFP wt Nde1 (73.03 min). In contrast, GFP Nde1 del. dynein did not rescue Nde1 siRNA defects and had a potent inhibition of mitotic progression. Cells remained arrested at metaphase for a significant period of time and multiple cells were unable to enter anaphase during the course of imaging (Figure 3.4). Chromosomes of cells arrested in metaphase often displayed loss tight alignment at the metaphase plate. Interestingly, the average time spent in mitosis for Nde1 siRNA cells rescued with Nde1 del dynein was significantly greater than Nde1 siRNA alone, 283.5 min compared to 123 min respectively. Consistent with the over- expression studies, cells expressing Nde1 del dynein had significant defects in mitotic progression while there was a lesser contribution of Nde1l Lis1 to Nde1’s role in mitotic progression.

Investigation of Metaphase defects caused by lack of Nde1-Dynein interaction

Figure 3.4 Rescue of Nde1 siRNA mitotic progression defects with Nde1 mutant constructs. A. H2B Cherry cells were treated with Nde1 siRNA for 24 hours prior to transfection with Nde1 rescue constructs. 18 hours later cells were imaged for ~12 hours. Time from nuclear envelope breakdown to anaphase onset was calculated. >18 cells were analyzed for each condition. Representative images of Nde1 siRNA treated Hela H2B cherry cells expressing either GFP Nde1 del Dynein (∆dynein ) or GFP Nde1 del Lis1 (∆Lis1) . Image panels represent time point 9 minutes apart. Scale =10µm. B. Quantification of the time from NEBD to anaphase onset control siRNA, Nde1 siRNA alone or Nde1 siRNA with GFP rescue constructs. Dynein-binding mutant is unable to rescue Nde1 siRNA mitotic progression defects while Lis1-binding mutant is able to rescue to levels comparable to WT Nde1. Cells that are boxed in the Nde1 siRNA +

Nde1 del Dynein condition never entered anaphase during the entire length of the movie. C. Quantification of the average time spent from NEBD to anaphase onset, separated by time from NEBD to alignment (unaligned) and time spent aligned until anaphase onset (aligned). Nde1 siRNA cells rescued with Nde1 del dynein on average spent a significantly longer time until anaphase onset, specifically a large portion of the time was in the unaligned state.

To determine the defects caused by a lack of Nde1-dynein interaction leading to significant metaphase arrest, we investigated the role of Nde1 in dynein-dependent processes at metaphase. Function blocking Nde1/Ndel1 antibody was reported to cause a decrease in kinetochore-microtubule (KT-MT) stabilization (Stehman et al., 2007), leading us to test the effects of Nde1 siRNA rescue with either wt Nde1 or GFP Nde1 del dynein on the ability to form stable KT-MT attachments. Cells were exposed to 0° degrees for 10 min to destabilize dynamic microtubules. Cold-stabilization experiments, demonstrated that rescue with Nde1 del dynein construct did not grossly effect the KT-

MT stabilization at the kinetochore (Figure 3.5). Both Nde1 siRNA treated Hela cells expressing wt Nde1 or Nde1 del dynein formed stable KT-MT interactions, indicating the metaphase arrest in Nde1 del dynein cells is not due to a global loss of KT-MT interactions. It is possible that single kinetochores have defects in stable microtubule formation but increased resolution is required to detect lack of KT-MT interactions on an individual scale.

Figure 3.5 Role of Nde1-dynein interaction in the formation of stable KT-MT attachments. Nde1 siRNA treated Hela cells rescued with GFP wt Nde1 or GFP Nde1 del dynein, were exposed to 0°C for 15 min prior to fixation. Immuno-histochemistry of alpha tubulin detects the presence of microtubules that were insensitive to cold treatment. Rescue with wt Nde1 or Nde1 del dynein does not prevent the formation of stable kinetochore-microtubule interactions on cell-wide scale. Scale = 5μm

We performed live-cell imaging on CENPA-YFP expressing cells to visualize the details of metaphase chromosome dynamics. Cells were treated with 10μM MG132 for 1 hour and followed by observation of kinetochore pair oscillations in either control siRNA, Nde1 siRNA or Nde1 siRNA treated rescued with wt Nde1 or Nde1 del dynein.

The number of cells exhibiting mis-alignment defects, as judged by escape the metaphase plate or significant movement towards the poles, was quantified. We observed that the lack of Nde1-dynein interaction did not rescue Nde1 siRNA mis-alignment defects and increased the number of cells with kinetochore alignment/maintenance defects

(Figure3.6A). On an individual kinetochore level, measurement the inter-kinetochore distance is an indication of the tension generation on the kinetochore by opposing pulling forces of mitotic spindle transmitted through stable/correct MT-KT attachments. We

measured the inter-kinetochore distance of aligned cell arrested in MG132 for 1 hour.

The distance in between CENPA+ sister kinetochore pairs was decreased in Nde1 siRNA

treated cells, but this decreased was rescued by expression of WT Nde1 (Figure 3.6B).

Nde1 siRNA treated cells expressing Nde1+del dynein construct did not rescue the inter-

kinetochore distance defects since there was a significant difference between wt Nde1

and Nde1 del dynein conditions. The average kinetochore distances for mock treated,

Nde1 siRNA, Nde1 siRNA+ wt Nde1 or Nde1 siRNA + Nde1 del dynein were 1.29 μm,

1.15μm, 1.26μm or 1.21μm, respectively. Nde1 and the Nde1-dyenin interaction are

required for normal tension generation at the kinetochore.

Dynein forces on the spindle oppose mitotic kinesin Eg5 for proper spindle length

and mitotic spindle tension. Raaijmakers et al., 2013 reported that Nde1 was required to

maintain spindle length in metaphase providing an additional mitotic role for Nde1 that

could be disrupted by the loss of the Nde1-dynein interaction (Raaijmakers et al., 2013).

To determine Nde1 mutant’s role in this process, we measured the length of the mitotic spindle in control siRNA, Nde1 siRNA or Nde1 siRNA cells rescued with wt Nde1 or

Nde1 del dynein. Measurements of spindle pole-spindle pole distance reveals Nde1 del

Dynein does not rescue spindle length defects seen in Nde1 siRNA (Figure 3.6C).

Figure 3.6 Characterization of metaphase defects caused by loss of Nde1 or the Nde1- dynein interaction. A. Hela CENP-A YFP cells were treated with Nde1 siRNA for 24hours followed by a 24-hour expression of mCherry wt Nde1 or mCherry Nde1 del dynein to not conflict with the YFP CENP-A signal. Cells were treated with 10μM MG132 to prevent anaphase onset and dynamics of sister kinetochore pairs were observed for a period of 15 min with frames acquired every 15 secs. Cells were scored for the presence of mis-aligned kinetochore pairs during the course of the movie. Quantification represents the ratio of cells with mis-aligned kinetochore/ total cells counted. Data was pooled from three different independent experiments. Statistics and error bars are absent since this is an absolute average of all the numbers pooled for each condition. Scale = 5μm B. The distance between YFP CENPA kinetochore pairs was measured mock treated, Nde1 siRNA or Nde1 siRNA rescued with wt Nde1 or Nde1 del dynein. CENP-A YFP cells were treated with 10μM MG132 for 30 min prior to measurement. Student’s T-test performed shows a significant difference between the

inter-kinetochore distances between cells rescued with wt Nde1e and Nde1 del dynein as well a Nde1 siRNA with either mock treated or wt Nde1 rescued cells. * p<.05, ***p<.001 C. Hela cells mock treated, Nde1 siRNA or Nde1 siRNA rescued with wt Nde1 or Nde1 del dynein were fixed and stained for endogenous alpha tubulin to stain the mitotic spindle for measurement of spindle length. The length of the spindle was determined as the distance between the spindle poles. Measurements revealed cells in both Nde1 siRNA and Nde1 siRNA + Nde1 del dynein conditions had increased spindle lengths.

Nde1 recruits dynein directly to the kinetochore through its coiled-coil domain

The phenotype in GFP Nde1 del Dynein expressing cells could be due defects in the

recruitment of dynein to the KT by Nde1, but the role of Nde1 in recruitment of dynein to

the kinetochore is controversial. A Nde1/Ndel1 function blocking antibody abolished

dynein signal on unaligned kinetochores, while Nde1 siRNA or Ndel1 siRNA did not

decrease the levels of GFP-Dynein Heavy Chain (DHC) at kinetochores in nocodazole treated cells (Raajimakers et al., 2013; Stehman et al., 2007). Rescue of Nde1 siRNA with GFP Nde1 del dynein construct would allow for direct read-out of the role Nde1 and

Nde1’s direct interaction with dynein in recruitment of dynein to the kinetochore.

Calculation of the intensity of dynein signal relative to ACA signal on kinetochores in mock treated or Nde1 siRNA treated cells, reveals Nde1 knockdown decreased dynein levels by ~80%. Expression of GFP wt Nde1 in Nde1 siRNA treated cells increased dynein levels measured on kinetochores, though dynein levels were slightly decreased in comparison to control cells, potentially due to incomplete rescue. Nde1 siRNA cells rescued with GFP Nde1 del Dynein did not display increased levels of dynein on the kinetochore. This rescue experiment suggests Nde1 directly recruits dynein to the kinetochore via Nde1’s N-terminal dynein-binding domain.

Figure 3.7. Nde1 recruits dynein to the kinetochore via its N-terminal coiled coil. A. Hela cells were treated with 2uM Nde1 siRNA for 48 hours then stained for endogenous Nde1/el1 or dynein intermediate chain. Signal intensity of Nde1/Ndel1 or dynein antibody was decreased in the siRNA condition. Hela cells treated with 2um siRNA were rescued with either GFP WT Nde1 or GFP Nde1 del Dynein for 24 hours. Dynein kinetochore signal intensity was quantified for GFP expressing cells. Scale = 5μm B. Quantification of kinetochore dynein levels in comparison to an internal standard, ACA intensity was performed. Relative (to Mock treated) average dynein/ACA levels were plotted +/-SEM. ~30 kinetochores/cell for >15 cells over three independent experiments. Paired student’s T-test was performed to analyze significance between conditions. * p<.05, **p<.01. The measurements reveal GFP Nde1 del dynein does not rescue the decrease in dynein localization in Nde1 siRNA treated cells. Although expression of wt Nde1 does not increase dynein signal intensity to mock treated levels, there is a significant difference between wt Nde1 and Nde1 del dynein.

Nde1-Lis1 interaction is required for Nde1 localization to the kinetochore

Phenotypic studies demonstrate that GFP Nde1 del Lis1 did not causes significant defects

in Nde1’s role in mitotic progression which is surprising since Nde1/Nde1l are thought to be the recruitment factor for Lis1 in dynein-dependent processes especially at the kinetochore (Stehman et al., 2007; Liang et al., 2007). To investigate the role of Nde1 in recruitment of Lis1 to the kinetochore, we treated cells with Nde1 siRNA and measured the signal intensity of Lis1 at kinetochores in nocodazole treated Hela cells.

Unfortunately due antibody species conflicts and fixation methods required for detection of Lis1 at kinetochore, rescue experiments with GFP Nde1 del Lis1 could not be performed. Nde1 siRNA decreased Lis1 signal by 40%, demonstrating Nde1 does not solely recruit Lis1 to the kinetochore by Nde1 but additional factors including Nde1l could contribute to Lis1 localization at the kinetochore (Figure 3.8).

Figure 3.8. Requirement of Lis1 and Nde1 for localization of each to the kinetochore. B. Hela cells were treated with Nde1 siRNA for 48 hours, followed by staining for endogenous Lis1 in nocodazole treated cells. The representative image demonstrates a decrease in endogenous Lis1 signal at the kinetochore in comparison to control siRNA. Scale=5μm B. Endogenous Lis1 intensity relative to ACA intensity was quantified from

~25 kinetochores of >30 cells from three independent experiments. Mean relative intensity +/- SEM. Students T-test was performed. **p<.01. Cells treated with Nde1 siRNA showed a significant decrease in Lis1 kinetochore intensity relative to control siRNA treated cells. C. Hela cells were treated with Lis1 siRNA for 96 hours. The representative image demonstrates a decrease in endogenous Nde1/Ndel1 signal at the kinetochore of nocodazole treated cells in comparison to control siRNA. Scale=5μm D. Endogenous Nde1 intensity relative to ACA intensity was quantified from ~25 kinetochores of >30 cells from three independent experiments. Mean relative intensity +/- SEM. Students T-test was performed. **p<.01. Cells treated with Lis1 siRNA showed a significant decrease in Nde1/el1 kinetochore intensity relative to control siRNA treated cells. *** p<.001 E. Western blot detection of endogenous levels of Lis1 from Hela cell lysate of control siRNA or Lis1 siRNA treated cells reveals a 60% decrease in protein levels in the siRNA condition.

Recently, Lis1 has been shown to be involved in localization of Bicd2, dynactin

and dynein to the plus-end of the MT (Splinter et al., 2012). It is unknown whether this

is a general function of lis1 or specific to the plus-end microtubule localization of dynein.

Given the interaction between Nde1 and Lis1 we tested the role of Lis1 in Nde1 localization at the kinetochore. Quantification of Nde1/Ndel1 kinetochore signal in control or Lis1 siRNA treated cells revealed Lis1 siRNA decreased the levels of

Nde1/Ndel1 by ~42% of control levels (Figure 3.8B). This demonstrates Lis1 has a role in Ndel/Ndel1’s localization at the kinetochore.

To determine if the effect of Lis1 on Nde1’s kinetochore localization is indirect or due to the direct interaction of Nde1 and Lis1, we quantified the levels of GFP signal at the kinetochore of nocodazole treated Hela cells expressing either GFP wt Nde1, GFP

Nde1 del dynein or GFP Nde1 del Lis1. Measurement of GFP intensity relative to anti- centromere protein (ACA) revealed GFP Nde1 del Lis1 had a decreased intensity at the kinetochore relative to GFP wt Nde1 or GFP del Dynein (Figure 3.9). Importantly, other mutants in the N-terminal coiled-coil do not decrease GPP Nde1 localization to the kinetochore; no significant difference between the GFP intensity in wt Nde1 and Nde1

del dynein was detected. This suggests that the loss of the direct interaction between

Nde1 and Lis1 decreases Nde1 localization to the kinetochore.

Figure 3.9. Localization of GFP Nde1 del Lis1 mutant to the kinetochore. A. GFP WT Nde1, GFP Nde1 del Dynein, GFP Nde1 del Lis1 localization to the kinetochore in nocodazole treated cells. Intensity of GFP signal, determined by anti-GFP staining, is decreased in cells expressing GFP Nde1 del Lis1. B. Quantification of average GFP

intensity relative to average ACA intensity. Mean +/- SEM of three independent experiments is represented. Student’s T-test performed shows a significant difference between the relative GFP intensities between WT Nde1 and Nde1 del Lis1. Expression of mutant GFP Nde1 del dynein does not decrease GFP Nde1 signal localization at the kinetochore. * p<.05, ** p<.01 Scale=5μm

CENP-F is the recruitment factor for Nde1 at the kinetochore (Vergnolle and

Taylor, 2007). In chapter 2, we demonstrated that phosphorylation state of Nde1 affects the binding of Nde1 to CENP-F. Since Lis1 siRNA decreased the levels of endogenous

Nde1 at the kinetochore, we wanted to test the role of Lis1-Nde1 interaction on Nde1 binding to CENPF. We aimed to determine if Lis1’s interaction with Nde1 influenced

Nde1’s interaction with CENP-F. An additional mutant construct, Nde1 cdk1 phospho- mimetic lacking lis1-binding site (Nde1 EEE, del Lis1) was created (Figure 3.11A).

GST-CENP-F (NBD) was incubated with Hela cell lysate expressing either GFP Nde1 phospho-mimetic (Nde1 EEE), GFP Nde1 phospho-mimetic with lis1 binding site mutated (EEE, del LIs1 3mt) or GFP Nde1 del Lis1 in Hela cells (Figure 3.10B). GST pull down experiments revealed CENP-F NBD pulled down less Nde1 when the Lis1 binding domain was mutated. GFP Nde1 phospho-mimetic lacking Lis1 (Nde1 EEE, del

Lis1) binding displayed decreased levels of interaction with CENPF and Nde1 del Lis1 show complete abolishment of an Nde1 interaction with CENPF. To ensure Lis1 did not mediate the interaction between Nde1 and CENPF, we tested the ability of Lis1 to bind to

CENPF. Baculovirus purified Lis1 did not interact with recombinant CENP-NBD

(Figure 3.10C). The direct interaction of Lis1 with Nde1 affected Nde1’s interaction with CENP-F through an unknown mechanism.

Figure 3.10. Biochemical analysis of the effect of Lis1 binding mutant on Nde1’s interaction with CENP-F. A. Description of Nde1 constructs used to examine the requirement of Lis1-Nde1 binding for an interaction with CENP-F: Cdk1 phospho- mimetic (Nde1EEE), Cdk1 phospho-mimetic lacking Lis1 binding (Nde1 EEE, del Lis1) and WT Nde1 del Lis1. B-C. Pull down of over-expressed Nde1 mutant constructs from cell lysate with bacterially expressed recombinant CENP-F fragment (NBD). Quantification of relative GFP-Nde1 signal from 3 independent experiments reveals Nde1 lacking Lis1 binding ability decreases its interaction with CENP-F. Decreased amount of GFP-Nde1 EEE, del Lis1 interacts with GST-CENP-F NDB, but the

interaction is not completely abolished. D. GST-Pull down with recombinant CENP-F NBD and baculovirus purified Lis1 reveals no interaction between the two proteins.

Discussion

The purpose of this study was to analyze specific Nde1 mutations to determine the physiological role of Nde1’s interaction with Lis1 or dynein. Ultimately, this leads to a greater understanding of the role and mechanism of Nde1 function in mitosis. Both the interaction between Nde1 and Lis1 or Nde1 and dynein are required for Nde1’s role in mitotic progression, though the severity of defects with the loss of the Nde1-dynein interaction is significantly more severe. Nde1 recruits dynein to the kinetochore to allow for stabilization of the KT-MT interaction. Alternatively, Nde1 and Lis1 have a cooperative interaction to influence the recruitment of each to the kinetochore. Lis1 indirectly affects Nde1’s interaction with kinetochore recruitment partner, CENP-F.

Mutational analysis demonstrates that the roles of the interactions between Nde1 and dynein or lis1 are unique and contribute to specific, unique physiological functions.

Importance of the N-terminal domain of Nde1 for its function with dynein

This work confirms the importance of Nde1’s N-terminal coiled-coil domain in its interaction with cytoplasmic dynein and provides a physiological relevance for this interaction. Previous identification of the N-terminal dynein-binding domain discussed the possibility of a second dynein binding site within the C-terminal domain of Nde1

(Wang et al. 2011; Zyłkiewicz et al., 2011). In vitro experiments ruled out the presence of a second binding site since recombinant Nde1 45.51AA did not pull down dynein complex. While amino acids 47 and 51 are the major sites of interaction, adjacent amino acids could contribute to the Nde1-dynein interaction by having a lower affinity for dynein, which is not detected in biochemical experiments. But it is the n-terminal

domain of Nde1 that is the site of interaction, leading to changes in orientation of the

Nde1 molecule for how Nde1 interacts with dynein.

Both over-expression of GFP Nde1 del dynein and rescue of Nde1 siRNA with

GFP Nde1 del dynein caused a prolongment of alignment and a significant metaphase arrest. The alignment phenotype was a late prometaphase defect suggesting the initial

MT-KT was not inhibited but late chromosome congression was disrupted by the lack of the Nde1-dynein interaction. Chromosomes in cells arrested in metaphase exhibited alignment instability, a behavior characterized by periods of a loss of alignment or a loose alignment at the metaphase plate before initiation of anaphase onset. This behavior could be a result of a decrease in the stability of the kinetochore-microtubule interaction or loss of end-on KT-MT attachments.

The rescue phenotype for cells expressing Nde1 del dynein was significantly more severe than Nde1 siRNA alone. This indicates that not only is the Nde1-dynein interaction required for mitotic functions of Nde1, but also GFP Nde1 del dynein could be causing dominant negative effects leading to disruption of the mitotic spindle pole organization. Ndel1, along with Lis1, are required for spindle aster formation in Xenopus cell-free system (Zyłkiewicz et al., 2011). Sequestration of Lis1 by GFP Nde1 del dynein could be causing significant spindle defects, preventing normal progression in anaphase while this mis-localization or sequestration of Lis1 does not occur in Nde1 siRNA conditions.

The metaphase arrest observed in Nde1 del dynein over-expression or rescue experiments was the result of defects in proper kinetochore-microtubule interactions preventing spindle assembly checkpoint inactivation. The lack of Nde1-dyenin

interaction did not cause gross defects in end-on microtubule stabilization, but led to

subtle defects in kinetochore-microtubule stability and tension generation. Nde1-dynein

interaction is not required for the initial lateral kinetochore-microtubule attachment, since defects were observed in late prometaphase and metaphase. Given Nde1’s role with dynein and lis1 in persistent force production, the metaphase kinetochore could a physiological site requiring this high force-production. Maintenance of stable end-on

kinetochore-microtubule interactions as well as tension generation across the sister kinetochore pairs could require dynein’s force production. Loss of dynein required for this metaphase function could cause the observed defects. The increase in the ratio of kinetochore pairs with mis-alignments defects in Nde1 del dynein conditions could

represent an inequality of forces between the sister kinetochores or loss of stable end-on

attachments at a single kinetochore. Additionally, decreased forces along the mitotic

spindle result in both a lack of tension generation across stable KT-MT interactions and

an increase in spindle length.

Nde1 has been described as a dynein recruitment factor. Rescue experiments

confirm that Nde1 recruits dynein to the kinetochore, and it is specifically due to dynein’s

interaction with Nde1’s coiled-coil domain. This is the first report of the N-terminal

dynein-binding domain within Nde1 acting as the site of dynein recruitment. Recently

the role of Nde1 in recruitment of dynein to the kinetochore has been disputed

(Raaijmakers et al., 2013). It is possible that cell type or experimental design could be

the reason for differing results. In vitro pull-down experiments confirm the recruitment

ability of Nde1. Biochemical experiments demonstrate Nde1 is able to interact with both

CENP-F via Nde1’s C-terminal domain and dynein through it N-terminal coiled-coil

100

domain. It has been assumed from siRNA experiments that CENP-F recruits dynein to nuclear envelope or kinetochore via Nde1, but this work provides biochemical evidence of the ability to for a tri-protein complex with CENP-F-Nde1 and dynein. The role for

Nde1 in recruitment of dynein to specific cargo has now been shown both physiologically and biochemically.

Role of the Nde1-lis1 interaction in mitosis

A surprising result of this study was the apparent non-essential nature of for the

Nde1-Lis1 interaction during mitosis: Over-expression of Nde1-del Lis1 only showed a slight delay in time to alignment and Nde1 siRNA treated cells rescued with GFP Nde1 del Lis1 did not have a significant difference compared to cells treated with GFP wt Nde1 in time to anaphase onset. Further examination of Nde1’s mitotic functions will clarify if the Nde1-lis1 interaction is required for Nde1’s role in spindle assembly checkpoint protein removal or kinetochore-microtubule attachment, even though there is only a subtle defect.

Previous studies of Lis1’s role in mitosis demonstrated that knockdown of Lis1 or inhibition with a function-blocking antibody caused a chromosome alignment phenotype

resulting in a significant increase in the time spent in mitosis (Raaijmakers et al., 2013;

Faulkner et al., 2000). This defect in mitotic progression was not detected in over-

expression of or rescue with GFP Nde1 del Lis1. Phenotypic studies of the requirement

for the Nde-Lis1 interaction demonstrate the direct interaction is not essential for Nde1

mitotic functions or contribute to all of Lis1’s activity in mitosis. Closer examination of

the defects observed in Nde1 del Lis1 phenotypic studies are required to understand

which Lis1 mitotic activities require its interaction with Nde1.

101

Consistent with what has been published on the role of Ndel1 in recruitment of

Lis1 to the kinetochore, Nde1 siRNA decreased endogenous Lis1 signal from

kinetochores, though Lis1 only dropped by ~50%. This reflects either incomplete Nde1 knockdown or a role for additional Lis1 recruitment factors at the kinetochore.

Increasing evidence of non-Nde1/el1 recruitment of Lis1 to the kinetochore. At the NE in G2, Lis1 signal is present prior to Nde1/el1 NE localization and is recruited to the NE with BicD2, dynactin in the early dynein recruitment pathway (Baffet et al. 2015;

Splinter et al., 2010). Additionally, disruption of dynactin localization by p50 over- expression causes mis-localization of Lis1 at the kinetochore (Tai et al., 2002). This data, along with results from this section provide further support for a non-Nde1 dependent recruitment pathway of Lis1 calling into question previous models of Nde1/el1 function as the sole recruitment factor for Lis1.

Surprisingly, interaction with Lis1 is required for Nde1 localization at the kinetochore. Lis1 siRNA decreased endogenous Nde1/el1 at the kinetochore while expression of GFP Nde1 del. Lis1 mutant displayed reduced GFP intensity at the kinetochore in comparison to GFP wt Nde1. This is the first report of Lis1 and its interaction with Nde1 in influencing Nde1 localization.

CENP-F is Nde1’s recruitment factor at the NE and kinetochore. Biochemical analysis reveals that Nde1 interaction with Lis1 is required for Nde1’s binding to CENP-

F, since less Nde1 del Lis1 GFP pull downs with GST-CENP-F. Lis1 acts directly on

Nde1 to influence Nde1’s behavior, since Lis1 does not bind to GST-CENP-F. Nde1 binding to Lis1 could influence the conformation of Nde1 to allow for a more stable interaction with CENP-F. A role for Lis1 in stabilization of interactions between dynein-

102

related proteins has been reported in stabilizing the dynein-dynactin-mRNA cargo interaction (Dix et al., 2008). Further investigation is required to clarify the mechanism of action between Lis1 and Nde1 to influence Nde1’s binding with CENP-F. One way to test the role of the Lis1-Nde1-CENP-F complex is to perform GST-CENP-F pull down experiments with recombinant Nde1 and Lis1. If the incubation of Lis1 with Nde1 increases the amount of Nde1 pulled down by GST-CENP-F NBD, this would suggest the binding of Lis1 to Nde11 affects Nde1’s interaction with CENP-F. Additionally, potentially this is a universal mechanism for Lis1, Nde1 and Nde1’s C-terminal interacting proteins in other physiological scenarios.

Questions remain about the physiological relevance of stabilization of the CENP-

F-Nde1-Lis1 interaction. Reported in Chapter 2, endogenous Nde1/el1 kinetochore signal drops ~50% upon chromosome alignment. Given the role of Lis1 in stabilization of an Nde1 fraction at the kinetochore, Lis1 could be influencing this kinetochore pattern of Nde1. Loss of Lis1 upon MT-attachment could lead to the decrease of Nde1/Ndel1 from the kinetochore, through a decreased interaction with CENP-F. To test this hypothesis, quantification of endogenous Nde1/el1 signal at the kinetochore of Lis1 siRNA treated cells. Similar Nde1/el1 antibody signal intensities between kinetochores of unaligned and aligned chromosomes would suggest a role for Lis1 in the MT- attachment dependent drop of Nde1/el1 signal in mitosis.

103

Chapter 4: Discussion

The entirety of this work aims to understand the regulation and function of Nde1

in mitosis by targeted structure-function experiments. We start by testing how

phosphorylation state of Nde1 affects its localization during G2/M and find that Nde1

phosphorylated by Cdk1 activates its interaction with CENP-F for localization to the NE

and kinetochore. Phenotypic studies examining the role of Cdk1 phospho-mimetic and

phospho-mutant Nde1 on Nde1’s mitotic function demonstrate both phospho-mimetic and phospho-mutant Nde1 rescue mitotic progression defects though over-expression of

GFP Nde1 phospho-mutant causes a late prometaphase/metaphase arrest.

Because localization of Nde1 potentially specifies a site for recruitment of Lis1 and dynein, we next investigated the roles of the interaction between Nde1 and Lis1 or

Nde1 and dynein. We report that Lis1-Nde1 interaction is required to localize Nde1 to the kinetochore through affecting Nde1’s interaction with CENP-F. Additionally, the

Nde1-dynein interaction is critical for Nde1’s function in metaphase, specifically in the stabilization of the kinetochore-microtubule interaction, by recruiting a fraction of dynein to the kinetochore.

By using a structure-function approach to understanding the mechanism of

Nde1‘s action during mitosis, targeting both phospho-regulation and specific interactions, we can more clearly understand the role of Nde1 as a dynein accessory protein, especially how Nde1 is required in G2/M for the complexity of dynein physiological functions.

Nde1 Phosphorylation

While the work presented in this thesis describes the role of Cdk1

phosphorylation in regulation the localization of Nde1 at mitotic sites during G2/M

104

through an increased interaction with recruitment factor, CENP-F, this is only the

beginning of the understanding of the importance of phosphorylation of Nde1. Discussed

below are broader implications of Cdk1 phosphorylation on Nde1 cargo-binding activity

as well as general principles about the role of phospho-regulation of Nde1 activities.

Phosphorylation of Nde1 could dictate cargo interactions and regulate Nde1’s

scaffold properties. We present a model for Cdk1 activation of the interaction between

Nde1 and CENP-F, which allows for the recruitment of Cdk1 phosphorylated Nde1 to the

NE and kinetochore, from prophase to anaphase. While Cdk1 phosphorylation of Nde1

specifies it to the NE and kinetochore, it is possible that the same phosphorylation

decreases the interaction between Nde1 and an interphase binding-partner. It has been

demonstrated that dephosphorylation regulates the Nde1-Su48 interaction at the

centrosome (Hirohashi et al., 2006b). With Nde1 involved in lysosome transport and

Golgi positioning in interphase (Lam et al., 2010; Liang et al., 2004), Cdk1

phosphorylation of the C-terminal domain of Nde1 could release Nde1 from these cargo

allowing for the precise orchestration of Nde1 cell-cycle specific functions. Additionally,

since phosphorylation of Nde1 is involved in Nde1’s lysosome function, (Pandey et al.,

2011; Kliman and Holzbaur et al, 2015), phosphorylation be a general mechanism to

regulate the timing and location of Nde1’s interactions.

This work specifically focused on the role of Cdk1 phospho-regulation of during

G2/M. While Cdk1 is a critical kinase in the G2/M transition, a phospho-proteome study

showed that there were approximately 6 (Dehoure et al., 2008) G2/M phosphorylation sites identified on Nde1. The identified or predicted Cdk1 kinase sites account for only 2

fractions, while the identity and role of additional mitotic phosphorylation sites are

105

unknown. Cdk1 has been known to be a priming phosphorylation, for example Cdk1

phosphorylation of MT tip-end binding protein CLIP-170, is phosphorylated by Cdk1 which in term allowed for a subsequent PLK1 phosphorylation. Given Nde1 is phosphorylated in G2, Cdk1 phosphorylation could allow for priming of additional phosphorylations required for Nde1 kinetochore functions. Additionally, differential phosphorylation of Nde1 could dictate subfractions of Nde1 at the centrosome or the spindle pole. Aurora A phosphorylation of Ndel1 localizes it to the spindle pole, a similar phosphorylation in Nde1 could target Nde1 to non-kinetochore mitotic sites. To further understand the role of phosphorylation in regulating Nde1 localization to additional sites or Nde1 activity at the kinetochore, it is critical to identify the additional kinases responsible for phosphorylation of Nde1.

One main unanswered question in the study of Nde1 is the similarities between

Nde1 and Ndel1 cellular functions. Differential phosphorylation of Nde1 and Ndel1 possibly distinguishes paralogue function in mitosis. The only reported phenotypic

differences between Nde1 and Ndel1 have been in their roles in mitosis, specifically in

mammalian cells and radial glial progenitor cells of the developing neocortex. Nde1, not

Ndel1, is required for mitotic entry in radial glial progenitor cells since Nde1 knockdown

blocks cells in G2. Additionally, rescue of Nde1 knockdown with Nde1l did not rescue

Nde1 G2 arrest, suggesting that Nde1l lacks a Nde1-specific activity required for its function in mitotic entry (Doobin et al., 2016). In mammalian cultured cells, knockdown of Nde1 or Ndel1 caused different phenotypes in time spent in mitosis. Nde1 siRNA cells demonstrated a significant delay in anaphase onset, consistent with reports of this thesis. In contrast, Nde1l siRNA treated cells did not delay mitotic progression, although

106

a few cells demonstrated lagging chromosomes indicative of incomplete KT-MT

attachments and a weak spindle assembly checkpoint (Vergnolle and Taylor. 2007).

Differential phosphorylation of Nde1 or Nde1l could be responsible for unique mitotic behaviors and requirement of a single paralogue in mitosis. Aurora A has been shown to phosphorylate Nde1l, but it has yet to be proven to phosphorylate Nde1, which lacks an

Aurora A consensus site. A complete study of Nde1 and Ndel1 phosphorylation in mitosis is required to get a better clue into the possible role of phosphorylation in distinguishing Nde1 and Ndel1 functions.

Generally, the importance of Nde1 phosphorylation lies in its role in the regulation of Nde1’s interaction with dynein and Lis1 to influence dynein-dependent functions. Phosphorylation has been reported to regulate the interaction of Ndel1 with dynein and/or Lis1 (Zyłkiewicz et al., 2011, Hebbar et al., 2008). Cdk1/5 phosphorylation increases Ndel1’s interaction with Lis1 (Yan et al., 2003; Hebbar et al.,

2008; Pandey and Smith., 2011). However there are disputing reports on how phosphorylation alters the interactions with dynein; one demonstrates Cdk1/Ckd5 phosphorylation of Nde1l decreases the interaction between Nde1 and dynein while another reports Nde1 phospho-mutant (246A) decreases association with dynein. This is a critical lack of understanding of the role of Cdk1 phosphorylation that requires additional experiments to clarify. Comparison of the rescue phenotypes of Nde1 phosphorylation constructs and Nde1 mutants can provide insight into whether phosphorylation is pheno-copying the lack of Lis1 or dynein interaction. In the rescue experiments, phospho-mimetic or phospho-mutant Nde1 has a dramatically less severe rescue phenotype in comparison to the complete loss of dynein-Nde1 interaction.

107

Phosphorylation state of Nde1 could be modulating, not abolishing, the interaction with

dynein, allowing for rescue of Nde1 mitotic progression defects with both phospho-

mimetic and phospho-mutant.

A main finding of this work is the role of Cdk1 phosphorylation of Nde1 in the activation of the Nde1-CENP-F interaction. The Cdk1 phosphorylation sites in Nde1 are adjacent to the CENP-F binding region within the C-terminus of the Nde1 molecule.

Additionally, Cdk1/5 phosphorylation has been shown to affect the interaction between

Nde1 and Lis1 and Nde1-Dynein, both of which bind Nde1 in the N-terminal coiled- coiled domain (Zyłkiewicz et al., 2011, Feng et al., 2000). Given the long-range affects on the interaction with binding-partners in the N-terminal domain and a lack of complete over-lap with the CENP-F binding region, an existing hypothesis is that phosphorylation causes long-range changes within the molecules. Cross-linking experiments with Nde1 detected interactions between the C-terminal domain of Nde1 and the Nde1 N-terminal coiled-coil (Soares et al., 2012). Coopertivity between N-terminal and C-terminal potentially allows for multiple conformations of the Nde1 molecule, one elongated and the other folded on itself. Cdk1 phosphorylation of the C-terminal could influence the open verses closed conformation of the molecule. To test this hypothesis, cross-linking experiments in the presence or absence of phosphorylated Nde1 would determine if interactions between the C-terminal and N-terminal domains are dependent on the phosphorylation state of Nde1. Additionally, multiple conformations of Nde1 could have a role in how Nde1 interacts with its cargo, dynein and Lis1. Further experiments will determine if conformation states of Nde1 have functional relevance. One hypothesis is that Nde1 is the tension/strain-sensing molecule for the dynein pathway and the

108

conformation of Nde1 transmits this information to dynein and Lis1. Upon experiencing

strain, Nde1 could adopt the open conformation allowing for increased recruitment of

Lis1 for persistent force production. Conformational regulation of Nde1 is an interesting

hypothesis that will allow for better understanding of the role of Nde1 in dynein-

dependent functions.

Nde1’s Role in Mitosis:

The functions of dynein at the kinetochore include chromosome congression

(Yang et al., 2007), stabilization of kinetochore-microtubule attachments during metaphase (Varma et al., 2009), and removal of the spindle assembly checkpoint (SAC) proteins (Howell et al., 2000). The recruitment of dynein to the kinetochore by Zw10 is required for dynein’s role in spindle assembly checkpoint inactivation through removal of spindle assembly checkpoint proteins but Nde1 has yet to be implicated in a specific mitotic function of dynein. Nde1 siRNA causes a slight defect in chromosome congression (Chapter 2 and Chapter 3), indicating a subtle role in dynein’s role in lateral microtubule attachment or initial MT attachment but the main defect observed in Nde1

siRNA, rescue with Nde1 mutants or over-expression of Nde1 GFP mutant constructs

was a late prometaphase/metaphase arrest suggesting Nde1 is involved in the KT-MT stabilization function of dynein or maintenance of end-on kinetochore microtubule attachments.

Both Nde1 siRNA, Nde1 del dynein or disruption of dynein function in metaphase demonstrated increased oscillations or escapes of kinetochore pairs from the

metaphase plate when analyzing the dynamics of CENP-A YFP kinetochores; although

the defect was significantly more severe in disruption of cytoplasmic dynein (Varma et

109

al., 2008). This could indicate an imbalance or altering of forces between the kinetochore pairs with one kinetochore experiencing stronger pulling forces that the other.

The mitotic spindle exerts significant forces on chromosomes. Measurement of forces on anaphase chromosomes has been measure to be 700 pN, with individual forces on a microtubule K-fiber to be around 10-15pN (Nicklas, 1983; Alexander and Rieder,

1991) Dynein and Nde1 could be required to counter this force production. Laser ablation of kinetochore microtubules of aligned chromosomes caused a rapid poleward movement of the chromosomes (Rieder and Salmon, 1994). While this represents a complete abolishment of MT interactions at metaphase, loss of dynein and Nde1 could result in subtle, incomplete loss of MT attachments caused by an imbalance of forces. To test this hypothesis, Nde1 manipulated cells, either with siRNA or expression of mutants, could be monitored for the loss of microtubule attachment in cells arrested at metaphase.

Mad2+ kinetochores would indicate loss of microtubule attachment. If Nde1 is involved in balancing forces and maintenance of KT-MT interactions, the number of Mad2+ kinetochores could increase or change over a period of time in metaphase-arrested cells indicating microtubule attachments are formed then lost.

Therefore, the role of Nde1 and dynein could be to assist additional protein complexes required for MT-KT interaction, specifically HEC1/Ndc80 complex, to counter-balance the forces on the kinetochores to allow for maintenance of KT-MT interaction at metaphase.

The one caveat is the requirement for the complex of Nde1-Lis1-dynein transforming dynein to a persistent force-producing motor (McKenney et al., 2010).

Phenotypic studies of Nde1 lacking the Lis1 interaction did not produce a significant

110

defect, though the behavior of kinetochore pairs was not measured. It is possible that

Lis1 is still able to interact with dynein independently of Nde1 to influence the motor

behavior of dynein.

Broader implications for the role of Nde1 in mitosis-

Mechanistic insight into the role of Nde1 in mitosis in mammalian cell culture

cells can be applied to understanding the role of Nde1 in mitotic events in neuronal

progenitor cells. Consequences of Nde1funtion in mitosis in radial glial progenitor cells

have broader implications in the development of the cortex.

Both knock-out of Nde1 in mice and homozygous mutations in human patients

cause significant cortical development phenotypes (Feng and Walsh, 2004; Alkuraya et al., 2011). The Nde1 null mouse exhibited a microcephalic brain, characterized by a decreased size of the cortex. The neuronal progenitor population of cells is depleted causing a decreased number of neurons in the later layers of the developing cortex (Feng and Walsh, 2004). In human patients, homozygous mutations in Nde1, a frameshift rendering the mutated protein unstable and functionally inactive, cause severe microlissencephly. This is characterized by a significant reduction in the circumference of the cortex and cortical layering malformations (Alkuraya et al., 2011). The microlissencephly seen in knock-out or knock-down Nde1 conditions is caused by multiple cell cycle blocks, one of which is entry into mitosis (Doobin et al., 2016). In contrast, Nde1 siRNA in cultured cells do not demonstrate G2 block seen in radial glial progenitor cells. The recruitment of additional dynein in the late G2 pathway, of which

CENP-F and Nde1 are a part of, could be essential in progenitor cells to increase dynein force production required to translocate the nucleus towards the crowded ventricle for

111

mitotic entry. As described in my work, Nde1 siRNA causes defects in mitotic

progression due to a prolongment of prometaphase and metaphase. While the phenotypes

are not exactly the same between Nde1 in mammalian cell culture studies and radial glial

progenitor cells, both cell types require Nde1 for the proper progression through mitosis.

Given the importance of Cdk1 phosphorylation of Nde1 behavior in mammalian

cultured cells, it is probably this phosphorylation event is also regulating Nde1 behavior

in the radial glial progenitor cells. Over-expression of GFP Nde1 phospho-mutant led to

an increase in the percentage of G2+cells suggesting the Cdk1 phosphorylation of Nde1

plays a role in Nde1’s role in mitotic entry. Therefore in neuronal progenitor cells, Cdk1

phosphorylation of Nde1 could be required for entry into mitosis. Rescue of Nde1

knockdown with Nde1 with either phospho-mimetic or phospho-mutant could test this

hypothesis. If Nde1 phospho-mimetic rescues the G2+ block seen in Nde1 siRNA while

Nde1 phospho-mutant does not, this would suggest that Cdk1 phosphorylation of Nde1 is

required for Nde1’s role in mitotic entry. The increase of the CENP-F-Nde1 interaction

by Cdk1 phosphorylation of Nde1 leading to Nde1 localization at the NE could be

required to recruit additional cytoplasmic dynein for late G2 functions.

Conclusion

The importance of Nde1 has been over-shadowed by the role of Lis1 in dynein-

dependent functions. It has been shown that Nde1 recruits Lis1 to the dynein complex in single-molecule studies to allow for persistent force generation by dynein, through increased MT interaction. Given the interaction between Lis1 and Nde1, Nde1/Ndel1 has been described as recruitment factor for Lis1 to subsequently bind dynein both in biochemical and physiological scenarios. The recent reports of Nde1 independent

112

functions of Lis1, specifically localization of Lis1 to the NE with dynactin and Bicd2

(Baffet et al., 2016), or Lis1 influencing the localization of dynactin and Bicd2 at the

plus-end of microtubules (Splinter et al., 2012), highlight the need to understand the role of Nde1, not just as a recruitment factor for Lis1. Nde1 is the dynein adaptor protein whose mechanism is still not completely known.

113

Chapter 5: Experimental Procedures

DNA Constructs

All Nde1 constructs were cloned from a mouse Nde1 template (gene ID: 67203)

PCR amplification was performed using KOD Hotstart (Millipore) kit following the specified protocol. Point mutagenesis was accomplished using Agilent Quick-change II kit and performed according to their protocol.

Table 1: Description of Constructs used in this study

Construct Vector Insert Cloning Notes Ha-Nde1 pGex 6p-1 Full Length Mouse Cloned by SJW Nde1 Ha Nde1 1-291 His pGex 6p-1 Nde1 aa 1-291 Cloned into pGex using BamH1, EcoR1 Ha Nde1 1-277 His pGex 6p-1 Nde1 aa 1-277 Cloned into pGex using BamH1, EcoR1 Ha Nde1 1-248 pGex 6p-1 Nde1 aa 1-248 Cloned into pGex His using BamH1, EcoR1 Nde1 1-191 pGex 6p-1 Nde1 aa 1-191 Cloned by SJW Nde1 10-191 pGex 6p-1 Nde1 10-191 Cloned by SK Nde1 10-191 pGex 6p-1 Nde1 10-191 E4747A 47.51AA 47.51AA E5151A Nde1 47.51AA pGex 6p-1 Full Length Nde1 E4747A 47.51AA E5151A GST-NudE 3mt pGex 6p-1 Nde1 119A, 126A, E119119A 130A R126126A R130130A CenpF D6 pGex 6p-1 Human CenpF aa Insert was 2122-2297 purchased from Genescript Cloned into pGex6p1 with BamH1 and Xho1 Restriction sites HA Nde1 45E pGex 6p-1 Nde1 243E,246E T243243E T246246E HA Nde1 245E pGex 6p-1 Nde1 T243243E 219E, 243E, 246E T246246E

114

S219219E HA Nde1 245A pGex 6p-1 Nde1 219A, 243A, T243243E 246A T246246E S219219E GFP Nde1 WT pIC113 Full Length Nde1 From SK/SS (pEGFP-TEV-S) GFP Nde1 47.51AA pIC113 Full Length Nde1 E4747A (pEGFP-TEV-S) 47.51AA E5151A GFP Nde1 3mt pIC113 Nde1 119A, 126A, E119119A (pEGFP-TEV-S) 130A R126126A R130130A GFP Nde1 245A pIC113 Nde1 219A, 243A, T243243A (pEGFP-TEV-S) 246A T246246A S219219A GFP Nde1 245E pIC113 Nde1 219E, 243E, T243243E (pEGFP-TEV-S) 246E T246246E S219219E mCherry Nde1 mCherry-C1 Full Length Nde1 From SK/DD mCherry Nde1 mCherry-C1 Full Length Nde1 E4747A 47.51AA 47.51AA E5151A mCherry Nde1 mCherry-C1 Nde1 219A, 243A, T243243A 245A 246A T246246A S219219A mCherry Nde1 mCherry-C1 Nde1 219E, 243E, T243243E 245E 246E T246246E S219219E GFP Nde1 245A, pIC113 Nde1 219A, 243A, T243243A 47.51AA (pEGFP-TEV-S) 246A, 47A, 51A T246246A S219219A E4747A E5151A GFP Nde1 3mt, pIC113 Nde1 119A, 126A, E119119A 245E (pEGFP-TEV-S) 130A, 219E, 243E, R126126A 246E R130130A T243243E T246246E S219219E

115

Table 2: Primers used in cloning Nde1 245E E#2: (F)GGCTCTGTACCGTCTGAGCCAGTAGCTCACCG (R)CGGTGAGCTACTGGCTCAGACGGTACAGAGCC

E#45: (F)CCACTAGTGGGGAGCCACTCGAACCTGCAGCCCGG (R)CCGGGCTGCAGGTTCGAGTGGCTCCCCACTAGTGG

Nde1 245A A#2 (F)GGCTCTGTACCGTCTGCTCCAGTAGCTCACCGAGG (R)CCTCGGTGAGCTACTGGAGCAGACGGTACAGAGCC A#4,5: (F)GCTCCACTAGTGGGGCGCCACTCGCACCTGCAGCCCGG (R)CCGGGCTGCAGGTGCGAGTGGCGCCCCACTAGTGGAGC

Nde1 47.51AA (F)GCCGAGAATACGCAGCTGAATTGGCGGCTCAGCTGC and Nde1 (R)GCAGCTGAGCCGCCAATTCAGCTGCGTATTCTCGGC 10-191 47.51AA

Nde1 126A,130A: 119A,126A, (F)CCAATGATGACCTGGAAGCAGCCAAAGCAGCCACAATCATG 130A (3mt) TCCC (R)GGGACATGATTGTGGCTGCTTTGGCTGCTTCCAGGTCATCAT TGG

119A: (F)GAAATACATTAGGGAACTGGCACAAGCCAATGATGACCTGG AAAGAGCC (R)GGCTCTTTCCAGGTCATCATTGGCTTGTGCCAGTTCCCTAATG TATTTC

Nde1 1-248 (R)GGGACGCCACTCACACCTGCAGGGCATCATCATCATCATCAT CATTAGGAATTCGCG Nde1 1-277 (R)CCAAGCTAGCATCATGCAGGAACTTCGGGCATCATCATCATC ATCATCATTAGGAATTCGCG Nde1 1-291 (R) GGACAAGTGGGCCAGCCTCAGGGCATCATCATCATCATCATCAT TAGGAATTCGCG

116

Mammalian Cell Culture

HelaM (gift from Vicki Allen- Lam et al 2010) , H2B-RFP Hela (gift from X),

Hela, and Hela YFP-CenpA (gift from Mao lab) were the cell lines used in this study.

Cells were cultured in DMEM supplemented with 10% HI FBS (Gibco) and 1:10,000 Pen

Strep (Gibco). H2B-RFP Hela cells were imaged overnight in C02-independent media

(Gibco) supplemented with 1mM Glutamate and 10% FBS.

Plasmids, SiRNA and Drug Treatments

Mammalian expression Plasmids

Mammalian Cell Culture expression constructs used in this study are: GFP-C1,

GFP-Nde1 WT, GFP-Nde1 47.51AA, GFP-Nde1 119A, 128A, 130A (3mt), GFP-Nde1

245E, GFP-Nde1 245A, mCherry Nde1 WT, mCherry Nde1 47.51AA. Transient

transfections were performed using Effectene (Qiagen) and cells were fixed or imaged

~24 hours post transfection.

siRNA

Protein knockdown was achieved using Dharmacon smartpool constructs: Human

NudE smartpool siRNA (Dharmacon M-020625-00-0010), Human Lis1 smartpool siRNA (Dharmacon,GE), Human CenpF smartpool siRNA (Dharmacon GE ). ZW10 siRNA (Dharmacon GE) target sequence was AAGGGUGAGGUGUGCAAUAUG

(Varma et al. 2006). Control cells were treated with control siRNA (Dharmacon GE).

2uM siRNA was transfected into Hela cells using Hiperfect reagent (Qiagen). Cells were treated with Nde1 or CenpF siRNA for 48 hours before fixation or live-cell imaging. Lis1 and Zw10 siRNA treatment of cells required two rounds of siRNA and replating.

Twenty-four hours post-transfection, cells were replated onto glass coverslips. After 36

117

hours, a second round of siRNA was applied to the cells. Coverslips were fixed/lysed 96

hours post transfection. Rescue of Nde1 was performed using mouse Nde1 cDNA,

insensitive to human siRNA constructs. Rescue constructs, GFP Nde1 and GFP Nde1

mutants, were transfected according to Effectene (Qiagen) protocol 24 hours prior to

fixing/live-cell imaging.

Drug treatment

Cells were treated with nocodozal overnight to accumulate mitotic cells or for 1 hour to disassemble microtubule network at concentrations of .1uM or 5uM respectively.

Cells were treated with 10uM MG132 for 1 hour, 9-27uM R0-3306 (Sigma) for 1hour or

10uM Taxol for 30 min.

Immunoflourescent microscopy

Cells were plated on glass coverslips for fixed image analysis or glass bottom dishes (MatTek Corp) for live-cell imaging. Cells were fixed in cold methanol for 7-10 min at -20° or with 3-4% paraformaldehyde in PHEM buffer ph 6.9 (60mM Pipes, 25mM

Hepes, 10mM EGTA, 2mM MgCl2) for 20 min at 37°. PFA fixed cells were subjected to pre-extraction, .1% triton in PHEM for 15 sec and washed prior to fixation. PFA-fixed cells were permibilized for 2 min with .1% triton in PBS. Coverslips were blocked with

.5% NDS in PBS for 1 hour at RT, primary antibodies were added for 1 hour at 37° or overnight at 4°, washed and secondary antibodies (Cy2, Cy3 or Cy5) were applied for 1 hour at RT. Secondary antibodies were used at ratio of 1:300. DAPI was added to the second wash post-secondary antibody. Experiments using phospho-Nde1 antibody were performed with TBS. Coverslips were mounted and imaged using Olympus Laser

Scanning Confocal or Olympus Spinning Disc Confocal.

118

Microscopy

Fixed images were acquired on a IX83 Andor Revolution XD Spinning Disk

Confocal System with a 60x silicone oil objective (NA 1.30) or 20X air objective (NA .7)

and a 2x magnifier coupled to an iXon Ultra 888 EMCCD Camera. Z-stacks of at most

.5um were taken for each image. Live-cell movies were taken with spinning disc confocal

using Andor software and the laser intensity did not exceed 6% for live-cell imaging.

Images for movies of mitotic progression were taken every 3 min for a time-course of 12-

16 hours and up to 20 fields were imaged. Movies of kinetochore behavior using Hela

CenpA-YFP cell-line were acquired every 15/sec for 20 min. Up to 5 cells and 4 z-planes were imaged each time point. On the spinning disc, a 20X was used for mitotic progression movies and 60X was used for imaging kinetochore dynamics. Anaphase experiment was performed using H2B-RFP hela cells. Cells were captured in the aligned state, then 9uM RO 3306 was added to the media. One minute post treatment, images were taken every 30 seconds for 30 min to analyze anaphase chromosome movement.

Analysis of images was performed with ImageJ software.

Quantification and Statistical Analysis

ImageJ software was used to analyze imaging data. Quantification of relative kinetochore intensity was calculated using the average maximum fluorescence of a kinetochore relative to the average maximum ACA intensity:

(Average Max Intensity-Average background intensity)/(Average Max ACA intensity-

Average ACA background intensity) Between 25-50 kinetochore measurements were

taken per cell while 15-20 measurements of the background intensity were polled to give

the average background intensity. At least 30 cells (N=3) were analyzed. Anaphase

119

chromosome movement/velocity was performed by measurement of maximum inter-

chromosome distance. All conditions were graphed relative to the WT or control

condition. Exposure, Laser intensity and EM Gain were consisted between all conditions

in each experiment.

Statistical analysis was carried out using Excel (Microsoft) or Prism (Graph Pad)

software. Significance was determined by students unpaired t-test. *=p<.05, **=p<.01,

***=p< .001. Data represented as mean and SEM of at least three independent replicates.

Protein Purification

Dynein Purification

Bovine dynein was purified according to the protocol described in Paschal 1991.

Briefly, fresh calf brains were collected and stored on ice until flask frozen with liquid

nitrogen. Brains were rapidly thawed and ~40g of brains was homogenized with a

blaneder and mechanical dounce in brain buffer(50mM Pipes, 50mM Hepes ph 7.0, 2mM

MgSO4 (MgCl2), 1mM EDTA supplemented with protease inhibitor (sigma) and 1mM

DTT). Lysate was spun at 12,000 rpm for thirty minutes, followed by a high speed,

60,000 rpm, spin for 1 hour at 4° to prepare calf brain lysate. To polymerize

microtubules, 30uM Taxol was added to the high-speed spin supernatant and incubated at

37° for 15min. 8% sucrose of a volume of 1/3 the supernatant is under-laid slowly to the bottom of the taxol-treated supernatant to collect the microtubule pellet. The microtubule pellet is collected after a 1 hour 14,000 rpm spin and resuspended in brain buffer with

10uM taxol. The solution is then incubated at 37° for 10min followed with a 28,000 rpm spin at 20°. The microtubule pellet is then resuspended in brain buffer supplemented with

120

10uM Taxol, 5mM GTP and 5mM MgSO4 to remove contaminating proteins that have higher affinity for GTP (kinesin, dynamin). To polymerize and collect the microtubules, the resuspended pellet was incubated at 37° for 10 min, followed with a 35,000 rpm spin.

The microtubule pellet is then resuspended in brain buffer supplemented with 10mM

ATP and 5mM MgS04, incubated at 37° for 10 min, followed by a 63,000 rpm spin at

20°. The supernatant was collected. ATP release supernatant was applied to 5-20% Tris-

KCL sucrose gradient and centrifuged overnight. Fractions were collected, analyzed by

SDS-PAGE gel stained with commassie and fractions containing cytoplasmic dynein were aliquoted and flash frozen with liquid nitrogen for long-term storage.

Bacterial Expression

pGex NudE constructs and pGex CenpF DNA was transformed into BL-21

CodonPlus RIPL competent cells (Agilent Technologies) for bacterial expression.

Bacteria cultures were supplemented with ampicillian and incubated shaking at 37°.

When cultures reached exponential growth phase (measured absorbance .600 at 600nm)

1mM IPTG was added. Cultures were then incubated for 4 hours at 20° then bacterial pellet was collected. Purification of GST-tagged protein from bacterial lysate was performed with agarose glutathione beads (USB) in lysis buffer (PBS supplemented with

2mM DTT and 1:100 protease inhibitor). Beads were washed extensively before purified

GST-tagged protein was either cleaved off glutathione beads (USB) by precision protease

(GE Biosciences) or eluted with 10mM reduced glutathione. Buffer exchange, using GE

NAP-10 columns, was performed to have the final purified protein in storage buffer buffer (1mM EDTA, 150 mM NaCl, 50mM Tris ph 7.0, 10% glycerol, 1mM DTT) or DB buffer (35mM Pipes, 5mM MgSO4, 1mM EGTA, .5mM EDTA, 1mM DTT). Protein

121

concentration was measured and quality of protein was determined by SDS-PAGE gel

stained with coomassie dye solution. Aliquots of purified protein were flash frozen with

liquid nitrogen and stored at -80°.

Baculovirus Expression

Lis1 was purified according to protocol detailed in Mckenney et al 2009. Sf9 insect cells (Invitrogen) were cultured with SF-900 II (Gibco) serum free media at 27°.

Cells were infected with Lis1 baculovirus for 48 hours before washing in cold PBS and

harvesting the cells in Ni+2 purification buffer (50 mM NaP04 ph 7.0, 300mM NaCl,

10mM Immidozale, and 10%glycerol). Lysate was homogenized using a glass dounce

followed by 2 min sonication on ice. His-tagged Lis1 was purified with Ni beads

(Qiagen). Purified protein was eluted with His purification buffer and 300mM

Immidozale. GE NP-10 columns were used to buffer exchange purified protein into a

final storage buffer (1mM EDTA, 150 mM NaCl, Tris ph 7.0, 10% glycerol, and 1mM

DTT). Protein concentration was measured and quality of protein was determined by

SDS-PAGE gel stained with coomassie dye solution. Aliquots of purified protein were

flash frozen with liquid nitrogen and stored at -80°.

Rat GST-MD and GST-MD deltaCT were purified according to the methods

described in Nicholas et al 2014. Briefly, Sf9 insect cells cultured in serum-free media

were infected with rat GST WT-MD or Delta CT-MD-expressing baculovirus. Cells were

harvested 48 hours post-infection, then mechanically lysed with DEB buffer (100mM

Pipes pH 7.2, 2mM MgSO4, 2mM EGTA, 50mM NaCl, 1M Glycerol) on ice with a glass

dounce. High-speed spin supernatant was applied to GST-spin trap columns (GE) for 30

min at 4°. Columns were washed with DEB buffer before addition of elution buffer

122

(GE). Columns were incubated for 30min, then the flow through was collected. To remove contaminating free GST, the purified protein product was applied to a hand- packed sepherose (GE) 1 ml column. 1000ul volume of DEB buffer was added to the column and ~150ul fractions were collected. Fraction content was analyzed by Tris-

Glycine SDS-PAGE gel (Invitrogen – Life Sciences) stained with commassie blue. GST- free fractions of purified GST-MD constructs were aliquoted and flash frozen with liquid nitrogen then stored in the -80°.

In Vitro Kinase assay

CyclinB/Cdk1 protein kinase (Millipore) was used to perform in vitro ckd1 kinase assays. 250ng CyclinB/Cdk1 was incubated with 5-10ug NudE, 200uM ATP, 5X PK

(NEB) buffer in a total volume of 50ul. Reaction was incubated at 30° for one hour.

Phosphorylated NudE was then used in pull-down experiments. Phosphorylation of Nde1 was monitored by western blot with pT MAPK/CDK1 antibody.

Immunoprecipitation/Pull-down experiments

Mammalian cell lysates were prepared in RIPA ph 7.4 (100mM NaCl, 1mM

EGTA, 50mM Tris, 1% NP-10, 1mM DTT, and 1:100 protease inhibitor (Sigma)).

Extracts were lysed on ice followed by centrifugation to remove membranous fraction.

500-750 ug of cell lysate was incubated with 5 ug antibody or added to GST-bound recombinant protein. Recombinant protein interaction studies were performed in buffer A

(50mM Hepes ph 7.4, 150 mM NaCl, 1% NP-40, 1mM DTT). For GST pull-down experiments, eluted GST constructs were incubated with glutathione agarose beads

(USB) in buffer A for 1 hour prior to incubation, washed to remove unbound GST protein, then incubated with interactor proteins for 1 hour in a total volume of 300ul.

123

Experiments with proteins recruited to the beads via HA- or His- antibody were performed with equal molar AB to protein. AB and protein complexes were allowed to form for 1 hour at 4° before washing and addition of interactor. Incubation with interactor protein occurred for 1 hour at 4°. Beads were washed four times before resuspended in

50ul of buffer A and addition of 6X sample buffer. Samples were analyzed by SDS-Page gel and western blot.

Western Blot

Western Blot analysis was performed with a PVDF methanol-activated membrane

(Millipore). The membrane was blocked with 5% milk in PBS for one hour at room temperature. Primary antibodies were diluted in 5ml of 5% milk in PBS and incubated with membrane for 1 hour at RT or overnight at 4°, then washed 3 times with PBS supplemented with 5% tween. Secondary antibodies (Alexa Fluor-680nm or -800nm conjugated) were added to 5ml of 5% milk in PBS at a dilution of 1:10000 and incubated at room temperature for 1 hour. The membrane was developed using the Odessey

Imaging system (Licor). Analysis of the membrane was performed using ImageJ software.

Sucrose Gradient Ultracentrifugation

Sucrose gradient analysis of purified GST motor domain constructs was performed with a 1.25ml gradient of layered sucrose consisting of four 325ul layers of

5%, 10%, 15%, and 20% sucrose in DEB buffer (100mM Pipes pH 7.2, 2mM MgSO4,

2mM EGTA, 50mM NaCl supplemented with 1mM DTT). The gradient was allowed to linearize overnight at 4°. The sucrose gradient was loaded with 5ug of WT-MD in 100ul

124

DEB buffer, and then centrifuged at 54,000 rpm for 3 hour at 4°. The gradients were

fractionated and analyzed by SDS-PAGE and western blot.

Antibodies

Antibodies used in this study are listed in the table below. WB designates concentrations used in western blots and IP shows dilution in immuno-fluorescent and western blot experiments.

Table 3: Antibodies used in the study Antibody Species Company Dilution:WB Dilution:IF Catalog # Crest/ACA Human Antibodies 1:400 15-235 Incorporated GFP Rabbit Invitrogen 1:2500 1:100 A11122 GFP Chicken Millipore 1:1000 1:100 16901 CenpF Mouse BD 1:1000 1:100 610768 (Mitosin) Bioscience YL ½ Rat Abcam 1:500 ab6160 Alpha Mouse Abcam 1:5000 1:300 ab7291 Tubulin Lis1 Rabbit Santa Cruz 1:2500 1:100 SC-15319 Dynein IC Mouse 1:5000 1:100 Gift from Kevin (74.1) pfister Lis1 Mouse Sigma 1:5000 L7391 Dynein HC Rabbit 1:5000 (8) NudE/L Rabbit 1:2500 1:100 (month 11) NudE Mouse Abnova 1:5000 p246 Rabbit Gift from Y. 1:1000 1:100 Alkurya 2011 Feng GST Mouse Santa Cruz 1:1000 SC-53909 pT Mouse Cell 1:1000 MAPK/CDK1 Signaling substrate AB Zw10 (AP2) rabbit 1:1000 1:100 Clip170 (F-3) Mouse Santa Cruz 1:100 SC-28325 Mad1 (9B10) Mouse Santa Cruz 1:50 SC-82746 14-3-3 Mouse Epsilon HA.11 Mouse Covance (IP) HA Rabbit Sigma H6908 (IP)

125

References

Alexander, S. P., & Rieder, C. L. (1991). Chromosome motion during attachment to the vertebrate spindle: initial saltatory-like behavior of chromosomes and quantitative analysis of force production by nascent kinetochore fibers. The Journal of Cell Biology, 113(4), 805–815.

Alkuraya, F. S., Cai, X., Emery, C., Mochida, G. H., Al-Dosari, M. S., Felie, J. M., et al. (2011). Human mutations in NDE1 cause extreme microcephaly with lissencephaly [corrected]. American Journal of Human Genetics, 88(5), 536–547.

Bader, J. R., Kasuboski, J. M., Winding, M., Vaughan, P. S., Hinchcliffe, E. H., & Vaughan, K. T. (2011). Polo-like kinase1 is required for recruitment of dynein to kinetochores during mitosis. The Journal of Biological Chemistry, 286(23), 20769– 20777.

Baffet, A. D., Hu, D. J., & Vallee, R. B. (2015). Cdk1 Activates Pre-mitotic Nuclear Envelope Dynein Recruitment and Apical Nuclear Migration in Neural Stem Cells. Developmental Cell, 33(6), 703–716.

Bakircioglu, M., Carvalho, O. P., Khurshid, M., Cox, J. J., Tuysuz, B., Barak, T., et al. (2011). The essential role of centrosomal NDE1 in human cerebral cortex neurogenesis. American Journal of Human Genetics, 88(5), 523–535.

Barbar, E., Kleinman, B., Imhoff, D., Li, M., Hays, T. S., & Hare, M. (2001). Dimerization and folding of LC8, a highly conserved light chain of cytoplasmic dynein. Biochemistry, 40(6), 1596–1605.

Barisic, M., & Geley, S. (2011). Spindly switch controls anaphase: spindly and RZZ functions in chromosome attachment and mitotic checkpoint control. Cell Cycle (Georgetown, Tex.), 10(3), 449–456.

Barisic, M., & Maiato, H. (2015). Dynein prevents erroneous kinetochore-microtubule attachments in mitosis. Cell Cycle (Georgetown, Tex.), 14(21), 3356–3361.

Barisic, M., Sohm, B., Mikolcevic, P., Wandke, C., Rauch, V., Ringer, T., et al. (2010). Spindly/CCDC99 is required for efficient chromosome congression and mitotic checkpoint regulation. Molecular Biology of the Cell, 21(12), 1968–1981.

Basto, R., Gomes, R., & Karess, R. E. (2000). Rough deal and Zw10 are required for the metaphase checkpoint in Drosophila. Nature Cell Biology, 2(12), 939–943.

Basto, R., Scaerou, F., Mische, S., Wojcik, E., Lefebvre, C., Gomes, R., et al. (2004). In vivo dynamics of the rough deal checkpoint protein during Drosophila mitosis. Current Biology : CB, 14(1), 56–61.

126

Benison, G., Karplus, P. A., & Barbar, E. (2007). Structure and dynamics of LC8 complexes with KXTQT-motif peptides: swallow and dynein intermediate chain compete for a common site. Journal of Molecular Biology, 371(2), 457–468.

Blasier, K. R., Humsi, M. K., Ha, J., Ross, M. W., Smiley, W. R., Inamdar, N. A., et al. (2014). Live cell imaging reveals differential modifications to cytoplasmic dynein properties by phospho- and dephosphomimic mutations of the intermediate chain 2C S84. Journal of Neuroscience Research, 92(9), 1143–1154.

Bolhy, S., Bouhlel, I., Dultz, E., Nayak, T., Zuccolo, M., Gatti, X., et al. (2011). A Nup133-dependent NPC-anchored network tethers centrosomes to the nuclear envelope in prophase. The Journal of Cell Biology, 192(5), 855–871.

Bradshaw, N. J., Hennah, W., & Soares, D. C. (2013). NDE1 and NDEL1: twin neurodevelopmental proteins with similar 'nature' but different “nurture.” Biomolecular Concepts, 4(5), 447–464.

Bradshaw, N. J., Soares, D. C., Carlyle, B. C., Ogawa, F., Davidson-Smith, H., Christie, S., et al. (2011). PKA phosphorylation of NDE1 is DISC1/PDE4 dependent and modulates its interaction with LIS1 and NDEL1. The Journal of Neuroscience : the Official Journal of the Society for Neuroscience, 31(24), 9043–9054.

Bremner, K. H., Scherer, J., Yi, J., Vershinin, M., Gross, S. P., & Vallee, R. B. (2009). Adenovirus transport via direct interaction of cytoplasmic dynein with the viral capsid hexon subunit. Cell Host & Microbe, 6(6), 523–535.

Buffin, E., Lefebvre, C., Huang, J., Gagou, M. E., & Karess, R. E. (2005). Recruitment of Mad2 to the kinetochore requires the Rod/Zw10 complex. Current Biology : CB, 15(9), 856–861.

Bullock, S. L., Nicol, A., Gross, S. P., & Zicha, D. (2006). Guidance of bidirectional motor complexes by mRNA cargoes through control of dynein number and activity. Current Biology : CB, 16(14), 1447–1452.

Burgess, S. A., Walker, M. L., Sakakibara, H., Knight, P. J., & Oiwa, K. (2003). Dynein structure and power stroke. Nature, 421(6924), 715–718.

Burgess, S. A., Walker, M. L., Sakakibara, H., Oiwa, K., & Knight, P. J. (2004). The structure of dynein-c by negative stain electron microscopy. Journal of Structural Biology, 146(1-2), 205–216. doi:10.1016/j.jsb.2003.10.005

Busson, S., Dujardin, D., Moreau, A., Dompierre, J., & De Mey, J. R. (1998). Dynein and dynactin are localized to astral microtubules and at cortical sites in mitotic epithelial cells. Current Biology : CB, 8(9), 541–544.

Cai, Q., Lu, L., Tian, J.-H., Zhu, Y.-B., Qiao, H., & Sheng, Z.-H. (2010). Snapin- regulated late endosomal transport is critical for efficient autophagy-lysosomal function in neurons. Neuron, 68(1), 73–86.

127

Caldas, G. V., Lynch, T. R., Anderson, R., Afreen, S., Varma, D., & DeLuca, J. G. (2015). The RZZ complex requires the N-terminus of KNL1 to mediate optimal Mad1 kinetochore localization in human cells. Open Biology, 5(11), 150160.

Carter, A. P., Cho, C., Jin, L., & Vale, R. D. (2011). Crystal structure of the dynein motor domain. Science (New York, N.Y.), 331(6021), 1159–1165.

Carter, A. P., Diamant, A. G., & Urnavicius, L. (2016). How dynein and dynactin transport cargos: a structural perspective. Current Opinion in Structural Biology, 37, 62– 70.

Carter, A. P., Garbarino, J. E., Wilson-Kubalek, E. M., Shipley, W. E., Cho, C., Milligan, R. A., et al. (2008). Structure and functional role of dynein's microtubule-binding domain. Science (New York, N.Y.), 322(5908), 1691–1695.

Caviston, J. P., Ross, J. L., Antony, S. M., Tokito, M., & Holzbaur, E. L. F. (2007). Huntingtin facilitates dynein/dynactin-mediated vesicle transport. Proceedings of the National Academy of Sciences of the United States of America, 104(24), 10045–10050.

Chan, G. K., Jablonski, S. A., Starr, D. A., Goldberg, M. L., & Yen, T. J. (2000). Human Zw10 and ROD are mitotic checkpoint proteins that bind to kinetochores. Nature Cell Biology, 2(12), 944–947.

Chan, Y. W., Fava, L. L., Uldschmid, A., Schmitz, M. H. A., Gerlich, D. W., Nigg, E. A., & Santamaria, A. (2009). Mitotic control of kinetochore-associated dynein and spindle orientation by human Spindly. The Journal of Cell Biology, 185(5), 859–874.

Chowdhury, S., Ketcham, S. A., Schroer, T. A., & Lander, G. C. (2015). Structural organization of the dynein-dynactin complex bound to microtubules. Nature Structural & Molecular Biology, 22(4), 345–347.

Chung, J. Y.-M., Steen, J. A., & Schwarz, T. L. (2016). Phosphorylation-Induced Motor Shedding Is Required at Mitosis for Proper Distribution and Passive Inheritance of Mitochondria. Cell Reports, 16(8), 2142–2155.

Civril, F., Wehenkel, A., Giorgi, F. M., Santaguida, S., Di Fonzo, A., Grigorean, G., et al. (2010). Structural analysis of the RZZ complex reveals common ancestry with multisubunit vesicle tethering machinery. Structure (London, England : 1993), 18(5), 616–626.

Collins, E. S., Balchand, S. K., Faraci, J. L., Wadsworth, P., & Lee, W.-L. (2012). Cell cycle-regulated cortical dynein/dynactin promotes symmetric cell division by differential pole motion in anaphase. Molecular Biology of the Cell, 23(17), 3380–3390.

Coquelle, F. M., Caspi, M., Cordelières, F. P., Dompierre, J. P., Dujardin, D. L., Koifman, C., et al. (2002). LIS1, CLIP-170's key to the dynein/dynactin pathway. Molecular and Cellular Biology, 22(9), 3089–3102.

128

Dell, K. R., Turck, C. W., & Vale, R. D. (2000). Mitotic phosphorylation of the dynein light intermediate chain is mediated by cdc2 kinase. Traffic (Copenhagen, Denmark), 1(1), 38–44.

Derewenda, U., Tarricone, C., Choi, W.-C., Cooper, D. R., Lukasik, S., Perrina, F., et al. (2007). The structure of the coiled-coil domain of Ndel1 and the basis of its interaction with Lis1, the causal protein of Miller-Dieker lissencephaly. Structure (London, England : 1993), 15(11), 1467–1481.

Dewitt, M. A., Chang, A. Y., Combs, P. A., & Yildiz, A. (2012). Cytoplasmic dynein moves through uncoordinated stepping of the AAA+ ring domains. Science (New York, N.Y.), 335(6065), 221–225.

Dewitt, M. A., Cypranowska, C. A., Cleary, F. B., Belyy, V., & Yildiz, A. (2015). The AAA3 domain of cytoplasmic dynein acts as a switch to facilitate microtubule release. Nature Structural & Molecular Biology, 22(1), 73–80.

Dix, C. I., Soundararajan, H. C., Dzhindzhev, N. S., Begum, F., Suter, B., Ohkura, H., et al. (2013). Lissencephaly-1 promotes the recruitment of dynein and dynactin to transported mRNAs. The Journal of Cell Biology, 202(3), 479–494.

Doobin, D. J., Kemal, S., Dantas, T. J., & Vallee, R. B. (2016). Severe NDE1-mediated microcephaly results from neural progenitor cell cycle arrests at multiple specific stages. Nature Communications, 7, 12551.

Drerup, C. M., Ahlgren, S. C., & Morris, J. A. (2007). Expression profiles of ndel1a and ndel1b, two orthologs of the NudE-Like gene, in the zebrafish. Gene Expression Patterns : GEP, 7(6), 672–679.

Echeverri, C. J., Paschal, B. M., Vaughan, K. T., & Vallee, R. B. (1996). Molecular characterization of the 50-kD subunit of dynactin reveals function for the complex in chromosome alignment and spindle organization during mitosis. The Journal of Cell Biology, 132(4), 617–633.

Eckley, D. M., Gill, S. R., Melkonian, K. A., Bingham, J. B., Goodson, H. V., Heuser, J. E., & Schroer, T. A. (1999). Analysis of dynactin subcomplexes reveals a novel actin- related protein associated with the arp1 minifilament pointed end. The Journal of Cell Biology, 147(2), 307–320.

Engelender, S., Sharp, A. H., Colomer, V., Tokito, M. K., Lanahan, A., Worley, P., et al. (1997). Huntingtin-associated protein 1 (HAP1) interacts with the p150Glued subunit of dynactin. Human Molecular Genetics, 6(13), 2205–2212.

Famulski, J. K., & Chan, G. K. (2007). Aurora B kinase-dependent recruitment of hZW10 and hROD to tensionless kinetochores. Current Biology : CB, 17(24), 2143– 2149.

129

Famulski, J. K., Vos, L., Sun, X., & Chan, G. (2008). Stable hZW10 kinetochore residency, mediated by hZwint-1 interaction, is essential for the mitotic checkpoint. The Journal of Cell Biology, 180(3), 507–520.

Faulkner, N. E., Dujardin, D. L., Tai, C. Y., Vaughan, K. T., O'Connell, C. B., Wang, Y., & Vallee, R. B. (2000). A role for the lissencephaly gene LIS1 in mitosis and cytoplasmic dynein function. Nature Cell Biology, 2(11), 784–791.

Feng, Y., & Walsh, C. A. (2001). Protein-protein interactions, cytoskeletal regulation and neuronal migration. Nature Reviews. Neuroscience, 2(6), 408–416.

Feng, Y., & Walsh, C. A. (2004). Mitotic spindle regulation by Nde1 controls cerebral cortical size. Neuron, 44(2), 279–293.

Feng, Y., Olson, E. C., Stukenberg, P. T., Flanagan, L. A., Kirschner, M. W., & Walsh, C. A. (2000). LIS1 regulates CNS lamination by interacting with mNudE, a central component of the centrosome. Neuron, 28(3), 665–679.

Feng, J., Huang, H., & Yen, T. J. (2006). CENP-F is a novel microtubule-binding protein that is essential for kinetochore attachments and affects the duration of the mitotic checkpoint delay. Chromosoma, 115(4), 320–329.

Gaglio, T., Dionne, M. A., & Compton, D. A. (1997). Mitotic spindle poles are organized by structural and motor proteins in addition to centrosomes. The Journal of Cell Biology, 138(5), 1055–1066.

Gaglio, T., Saredi, A., Bingham, J. B., Hasbani, M. J., Gill, S. R., Schroer, T. A., & Compton, D. A. (1996). Opposing motor activities are required for the organization of the mammalian mitotic spindle pole. The Journal of Cell Biology, 135(2), 399–414.

Gao, F. J., Hebbar, S., Gao, X. A., Alexander, M., Pandey, J. P., Walla, M. D., et al. (2015). GSK-3β phosphorylation of cytoplasmic dynein reduces Ndel1 binding to intermediate chains and alters dynein motility. Traffic (Copenhagen, Denmark), n/a–n/a.

Gassmann, R., Essex, A., Hu, J.-S., Maddox, P. S., Motegi, F., Sugimoto, A., et al. (2008). A new mechanism controlling kinetochore-microtubule interactions revealed by comparison of two dynein-targeting components: SPDL-1 and the Rod/Zwilch/Zw10 complex. Genes & Development, 22(17), 2385–2399.

Gassmann, R., Holland, A. J., Varma, D., Wan, X., Civril, F., Cleveland, D. W., et al. (2010). Removal of Spindly from microtubule-attached kinetochores controls spindle checkpoint silencing in human cells. Genes & Development, 24(9), 957–971.

Gee, M. A., Heuser, J. E., & Vallee, R. B. (1997). An extended microtubule-binding structure within the dynein motor domain. Nature, 390(6660), 636–639.

Gennerich, A., Carter, A. P., Reck-Peterson, S. L., & Vale, R. D. (2007). Force-induced bidirectional stepping of cytoplasmic dynein. Cell, 131(5), 952–965.

130

Gibbons, I. R., Garbarino, J. E., Tan, C. E., Reck-Peterson, S. L., Vale, R. D., & Carter, A. P. (2005). The affinity of the dynein microtubule-binding domain is modulated by the conformation of its coiled-coil stalk. The Journal of Biological Chemistry, 280(25), 23960–23965.

Gill, S. R., Schroer, T. A., Szilak, I., Steuer, E. R., Sheetz, M. P., & Cleveland, D. W. (1991). Dynactin, a conserved, ubiquitously expressed component of an activator of vesicle motility mediated by cytoplasmic dynein. The Journal of Cell Biology, 115(6), 1639–1650.

Gönczy, P., Pichler, S., Kirkham, M., & Hyman, A. A. (1999). Cytoplasmic dynein is required for distinct aspects of MTOC positioning, including centrosome separation, in the one cell stage Caenorhabditis elegans embryo. The Journal of Cell Biology, 147(1), 135–150.

Griffis, E. R., Stuurman, N., & Vale, R. D. (2007). Spindly, a novel protein essential for silencing the spindle assembly checkpoint, recruits dynein to the kinetochore. The Journal of Cell Biology, 177(6), 1005–1015.

Grill, S. W., Howard, J., Schäffer, E., Stelzer, E. H. K., & Hyman, A. A. (2003). The distribution of active force generators controls mitotic spindle position. Science (New York, N.Y.), 301(5632), 518–521. .

Haghnia, M., Cavalli, V., Shah, S. B., Schimmelpfeng, K., Brusch, R., Yang, G., et al. (2007). Dynactin is required for coordinated bidirectional motility, but not for dynein membrane attachment. Molecular Biology of the Cell, 18(6), 2081–2089.

Heald, R., Tournebize, R., Blank, T., Sandaltzopoulos, R., Becker, P., Hyman, A., & Karsenti, E. (1996). Self-organization of microtubules into bipolar spindles around artificial chromosomes in Xenopus egg extracts. Nature, 382(6590), 420–425.

Heald, R., Tournebize, R., Habermann, A., Karsenti, E., & Hyman, A. (1997). Spindle assembly in Xenopus egg extracts: respective roles of centrosomes and microtubule self- organization. The Journal of Cell Biology, 138(3), 615–628.

Hebbar, S., Mesngon, M. T., Guillotte, A. M., Desai, B., Ayala, R., & Smith, D. S. (2008). Lis1 and Ndel1 influence the timing of nuclear envelope breakdown in neural stem cells. The Journal of Cell Biology, 182(6), 1063–1071.

Hirokawa, N., Noda, Y., Tanaka, Y., & Niwa, S. (2009). Kinesin superfamily motor proteins and intracellular transport. Nature Reviews. Molecular Cell Biology, 10(10), 682–696.

Hirohashi, Y., Wang, Q., Liu, Q., Du, X., Zhang, H., Sato, N., & Greene, M. I. (2006). p78/MCRS1 forms a complex with centrosomal protein Nde1 and is essential for cell viability. Oncogene, 25(35), 4937–4946.

131

Hirohashi, Y., Wang, Q., Liu, Q., Li, B., Du, X., Zhang, H., et al. (2006). Centrosomal proteins Nde1 and Su48 form a complex regulated by phosphorylation. Oncogene, 25(45), 6048–6055.

Hochegger, H., Takeda, S., & Hunt, T. (2008). Cyclin-dependent kinases and cell-cycle transitions: does one fit all? Nature Reviews. Molecular Cell Biology, 9(11), 910–916.

Hoffman, D. B., Pearson, C. G., Yen, T. J., Howell, B. J., & Salmon, E. D. (2001). Microtubule-dependent changes in assembly of microtubule motor proteins and mitotic spindle checkpoint proteins at PtK1 kinetochores. Molecular Biology of the Cell, 12(7), 1995–2009.

Holt, S. V., Vergnolle, M. A. S., Hussein, D., Wozniak, M. J., Allan, V. J., & Taylor, S. S. (2005). Silencing Cenp-F weakens centromeric cohesion, prevents chromosome alignment and activates the spindle checkpoint. Journal of Cell Science, 118(Pt 20), 4889–4900.

Holzbaur, E. L., Hammarback, J. A., Paschal, B. M., Kravit, N. G., Pfister, K. K., & Vallee, R. B. (1992). Homology of a 150K cytoplasmic dynein-associated polypeptide with the Drosophila gene Glued. Nature, 360(6405), 695–695.

Hoogenraad, C. C., Akhmanova, A., Howell, S. A., Dortland, B. R., De Zeeuw, C. I., Willemsen, R., et al. (2001). Mammalian Golgi-associated Bicaudal-D2 functions in the dynein-dynactin pathway by interacting with these complexes. The EMBO Journal, 20(15), 4041–4054.

Hoogenraad, C. C., Wulf, P., Schiefermeier, N., Stepanova, T., Galjart, N., Small, J. V., et al. (2003). Bicaudal D induces selective dynein-mediated microtubule minus end- directed transport. The EMBO Journal, 22(22), 6004–6015.

Höök, P., & Vallee, R. (2012). Dynein dynamics. Nature Structural & Molecular Biology, 19(5), 467–469.

Horgan, C. P., Hanscom, S. R., & McCaffrey, M. W. (2011). Dynein LIC1 localizes to the mitotic spindle and midbody and LIC2 localizes to spindle poles during cell division. Cell Biology International, 35(2), 171–178.

Horgan, C. P., Hanscom, S. R., Jolly, R. S., Futter, C. E., & McCaffrey, M. W. (2010). Rab11-FIP3 binds dynein light intermediate chain 2 and its overexpression fragments the Golgi complex. Biochemical and Biophysical Research Communications, 394(2), 387– 392.

Horgan, C. P., Hanscom, S. R., Jolly, R. S., Futter, C. E., & McCaffrey, M. W. (2010). Rab11-FIP3 links the Rab11 GTPase and cytoplasmic dynein to mediate transport to the endosomal-recycling compartment. Journal of Cell Science, 123(Pt 2), 181–191.

Howell, B. J., McEwen, B. F., Canman, J. C., Hoffman, D. B., Farrar, E. M., Rieder, C. L., & Salmon, E. D. (2001). Cytoplasmic dynein/dynactin drives kinetochore protein

132

transport to the spindle poles and has a role in mitotic spindle checkpoint inactivation. The Journal of Cell Biology, 155(7), 1159–1172.

Hu, D. J.-K., Baffet, A. D., Nayak, T., Akhmanova, A., Doye, V., & Vallee, R. B. (2013). Dynein recruitment to nuclear pores activates apical nuclear migration and mitotic entry in brain progenitor cells. Cell, 154(6), 1300–1313.

Huang, C. Y., Chang, C. P., Huang, C. L., & Ferrell, J. E. (1999). M phase phosphorylation of cytoplasmic dynein intermediate chain and p150(Glued). The Journal of Biological Chemistry, 274(20), 14262–14269.

Huang, J., Roberts, A. J., Leschziner, A. E., & Reck-Peterson, S. L. (2012). Lis1 acts as a “clutch” between the ATPase and microtubule-binding domains of the dynein motor. Cell, 150(5), 975–986.

Hughes, S. M., Vaughan, K. T., Herskovits, J. S., & Vallee, R. B. (1995). Molecular analysis of a cytoplasmic dynein light intermediate chain reveals homology to a family of ATPases. Journal of Cell Science, 108 ( Pt 1), 17–24.

Hussein, D., & Taylor, S. S. (2002). Farnesylation of Cenp-F is required for G2/M progression and degradation after mitosis. Journal of Cell Science, 115(Pt 17), 3403– 3414.

Ikeda, M., Hikita, T., Taya, S., Uraguchi-Asaki, J., Toyo-oka, K., Wynshaw-Boris, A., et al. (2008). Identification of YWHAE, a gene encoding 14-3-3epsilon, as a possible susceptibility gene for schizophrenia. Human Molecular Genetics, 17(20), 3212–3222.

Imamula, K., Kon, T., Ohkura, R., & Sutoh, K. (2007). The coordination of cyclic microtubule association/dissociation and tail swing of cytoplasmic dynein. Proceedings of the National Academy of Sciences of the United States of America, 104(41), 16134– 16139.

Inoue, M., Arasaki, K., Ueda, A., Aoki, T., & Tagaya, M. (2008). N-terminal region of ZW10 serves not only as a determinant for localization but also as a link with dynein function. Genes to Cells : Devoted to Molecular & Cellular Mechanisms, 13(8), 905–914.

Jaarsma, D., & Hoogenraad, C. C. (2015). Cytoplasmic dynein and its regulatory proteins in Golgi pathology in nervous system disorders. Frontiers in Neuroscience, 9, 397.

Jordens, I., Fernandez-Borja, M., Marsman, M., Dusseljee, S., Janssen, L., Calafat, J., et al. (2001). The Rab7 effector protein RILP controls lysosomal transport by inducing the recruitment of dynein-dynactin motors. Current Biology : CB, 11(21), 1680–1685.

Kardon, J. R., & Vale, R. D. (2009). Regulators of the cytoplasmic dynein motor. Nature Reviews. Molecular Cell Biology, 10(12), 854–865.

Kardon, J. R., Reck-Peterson, S. L., & Vale, R. D. (2009). Regulation of the processivity and intracellular localization of Saccharomyces cerevisiae dynein by dynactin.

133

Proceedings of the National Academy of Sciences of the United States of America, 106(14), 5669–5674.

Karess, R. (2005). Rod-Zw10-Zwilch: a key player in the spindle checkpoint. Trends in Cell Biology, 15(7), 386–392.

Karki, S., & Holzbaur, E. L. (1999). Cytoplasmic dynein and dynactin in cell division and intracellular transport. Current Opinion in Cell Biology, 11(1), 45–53.

Kasuboski, J. M., Bader, J. R., Vaughan, P. S., Tauhata, S. B. F., Winding, M., Morrissey, M. A., et al. (2011). Zwint-1 is a novel Aurora B substrate required for the assembly of a dynein-binding platform on kinetochores. Molecular Biology of the Cell, 22(18), 3318–3330.

Kim, M. H., Cooper, D. R., Oleksy, A., Devedjiev, Y., Derewenda, U., Reiner, O., et al. (2004). The structure of the N-terminal domain of the product of the lissencephaly gene Lis1 and its functional implications. Structure (London, England : 1993), 12(6), 987–998.

King, J. M., Hays, T. S., & Nicklas, R. B. (2000). Dynein is a transient kinetochore component whose binding is regulated by microtubule attachment, not tension. The Journal of Cell Biology, 151(4), 739–748.

King, S. J., & Schroer, T. A. (2000). Dynactin increases the processivity of the cytoplasmic dynein motor. Nature Cell Biology, 2(1), 20–24.

King, S. J., Bonilla, M., Rodgers, M. E., & Schroer, T. A. (2002). Subunit organization in cytoplasmic dynein subcomplexes. Protein Science : a Publication of the Protein Society, 11(5), 1239–1250.

King, S. J., Brown, C. L., Maier, K. C., Quintyne, N. J., & Schroer, T. A. (2003). Analysis of the dynein-dynactin interaction in vitro and in vivo. Molecular Biology of the Cell, 14(12), 5089–5097.

King, S. M. (2000). AAA domains and organization of the dynein motor unit. Journal of Cell Science, 113 ( Pt 14), 2521–2526.

King, S. M., Barbarese, E., Dillman, J. F., Patel-King, R. S., Carson, J. H., & Pfister, K. K. (1996). Brain cytoplasmic and flagellar outer arm dyneins share a highly conserved Mr 8,000 light chain. The Journal of Biological Chemistry, 271(32), 19358–19366.

Kiyomitsu, T., & Cheeseman, I. M. (2012). Chromosome- and spindle-pole-derived signals generate an intrinsic code for spindle position and orientation. Nature Cell Biology, 14(3), 311–317.

Kiyomitsu, T., & Cheeseman, I. M. (2013). Cortical dynein and asymmetric membrane elongation coordinately position the spindle in anaphase. Cell, 154(2), 391–402.

134

Klinman, E., & Holzbaur, E. L. F. (2015). Stress-Induced CDK5 Activation Disrupts Axonal Transport via Lis1/Ndel1/Dynein. Cell Reports, 12(3), 462–473.

Kon, T., Nishiura, M., Ohkura, R., Toyoshima, Y. Y., & Sutoh, K. (2004). Distinct functions of nucleotide-binding/hydrolysis sites in the four AAA modules of cytoplasmic dynein. Biochemistry, 43(35), 11266–11274.

Kon, T., Oyama, T., Shimo-Kon, R., Imamula, K., Shima, T., Sutoh, K., & Kurisu, G. (2012). The 2.8 Å crystal structure of the dynein motor domain. Nature, 484(7394), 345– 350.

Koonce, M. P. (1997). Identification of a microtubule-binding domain in a cytoplasmic dynein heavy chain. The Journal of Biological Chemistry, 272(32), 19714–19718.

Koonce, M. P., & Tikhonenko, I. (2000). Functional elements within the dynein microtubule-binding domain. Molecular Biology of the Cell, 11(2), 523–529.

Kops, G. J. P. L., Kim, Y., Weaver, B. A. A., Mao, Y., McLeod, I., Yates, J. R., et al. (2005). ZW10 links mitotic checkpoint signaling to the structural kinetochore. The Journal of Cell Biology, 169(1), 49–60.

Kotak, S., & Gönczy, P. (2014). NuMA phosphorylation dictates dynein-dependent spindle positioning. Cell Cycle (Georgetown, Tex.), 13(2), 177–178.

Kotak, S., Busso, C., & Gönczy, P. (2012). Cortical dynein is critical for proper spindle positioning in human cells. The Journal of Cell Biology, 199(1), 97–110.

Kotak, S., Busso, C., & Gönczy, P. (2013). NuMA phosphorylation by CDK1 couples mitotic progression with cortical dynein function. The EMBO Journal, 32(18), 2517– 2529.

Kuijpers, M., van de Willige, D., Freal, A., Chazeau, A., Franker, M. A., Hofenk, J., et al. (2016). Dynein Regulator NDEL1 Controls Polarized Cargo Transport at the Axon Initial Segment. Neuron, 89(3), 461–471.

Kumar, S., Zhou, Y., & Plamann, M. (2001). Dynactin-membrane interaction is regulated by the C-terminal domains of p150(Glued). EMBO Reports, 2(10), 939–944.

Laan, L., Pavin, N., Husson, J., Romet-Lemonne, G., van Duijn, M., López, M. P., et al. (2012). Cortical dynein controls microtubule dynamics to generate pulling forces that position microtubule asters. Cell, 148(3), 502–514.

Lam, C., Vergnolle, M. A. S., Thorpe, L., Woodman, P. G., & Allan, V. J. (2010). Functional interplay between LIS1, NDE1 and NDEL1 in dynein-dependent organelle positioning. Journal of Cell Science, 123(Pt 2), 202–212.

Levy, J. R., & Holzbaur, E. L. F. (2006). Cytoplasmic dynein/dynactin function and dysfunction in motor neurons. International Journal of Developmental Neuroscience : the

135

Official Journal of the International Society for Developmental Neuroscience, 24(2-3), 103–111.

Li, H., Liu, X. S., Yang, X., Song, B., Wang, Y., & Liu, X. (2010). Polo-like kinase 1 phosphorylation of p150Glued facilitates nuclear envelope breakdown during prophase. Proceedings of the National Academy of Sciences of the United States of America, 107(33), 14633–14638.

Li, S. H., Gutekunst, C. A., Hersch, S. M., & Li, X. J. (1998). Interaction of huntingtin- associated protein with dynactin P150Glued. The Journal of Neuroscience : the Official Journal of the Society for Neuroscience, 18(4), 1261–1269.

Li, Y., Yu, W., Liang, Y., & Zhu, X. (2007). Kinetochore dynein generates a poleward pulling force to facilitate congression and full chromosome alignment. Cell Research, 17(8), 701–712.

Liang, Y., Yu, W., Li, Y., Yang, Z., Yan, X., Huang, Q., & Zhu, X. (2004). Nudel functions in membrane traffic mainly through association with Lis1 and cytoplasmic dynein. The Journal of Cell Biology, 164(4), 557–566.

Liang, Y., Yu, W., Li, Y., Yu, L., Zhang, Q., Wang, F., et al. (2007). Nudel modulates kinetochore association and function of cytoplasmic dynein in M phase. Molecular Biology of the Cell, 18(7), 2656–2666.

Liao, H., Winkfein, R. J., Mack, G., Rattner, J. B., & Yen, T. J. (1995). CENP-F is a protein of the nuclear matrix that assembles onto kinetochores at late G2 and is rapidly degraded after mitosis. The Journal of Cell Biology, 130(3), 507–518.

Lo, K. W.-H., Kan, H.-M., & Pfister, K. K. (2006). Identification of a novel region of the cytoplasmic Dynein intermediate chain important for dimerization in the absence of the light chains. The Journal of Biological Chemistry, 281(14), 9552–9559.

Lu, M. S., & Prehoda, K. E. (2013). A NudE/14-3-3 pathway coordinates dynein and the kinesin Khc73 to position the mitotic spindle. Developmental Cell, 26(4), 369–380.

Lydersen, B. K., & Pettijohn, D. E. (1980). Human-specific nuclear protein that associates with the polar region of the mitotic apparatus: distribution in a human/hamster hybrid cell. Cell, 22(2 Pt 2), 489–499.

Mahale, S. P., Sharma, A., & Mylavarapu, S. V. S. (2016). Dynein Light Intermediate Chain 2 Facilitates the Metaphase to Anaphase Transition by Inactivating the Spindle Assembly Checkpoint. PloS One, 11(7), e0159646.

Mallik, R., Carter, B. C., Lex, S. A., King, S. J., & Gross, S. P. (2004). Cytoplasmic dynein functions as a gear in response to load. Nature, 427(6975), 649–652.

136

Mallik, R., Petrov, D., Lex, S. A., King, S. J., & Gross, S. P. (2005). Building complexity: an in vitro study of cytoplasmic dynein with in vivo implications. Current Biology : CB, 15(23), 2075–2085.

McKenney, R. J., Huynh, W., Tanenbaum, M. E., Bhabha, G., & Vale, R. D. (2014). Activation of cytoplasmic dynein motility by dynactin-cargo adapter complexes. Science (New York, N.Y.), 345(6194), 337–341.

McKenney, R. J., Vershinin, M., Kunwar, A., Vallee, R. B., & Gross, S. P. (2010). LIS1 and NudE induce a persistent dynein force-producing state. Cell, 141(2), 304–314.

McKenney, R. J., Weil, S. J., Scherer, J., & Vallee, R. B. (2011). Mutually exclusive cytoplasmic dynein regulation by NudE-Lis1 and dynactin. The Journal of Biological Chemistry, 286(45), 39615–39622.

Merdes, A., Heald, R., Samejima, K., Earnshaw, W. C., & Cleveland, D. W. (2000). Formation of spindle poles by dynein/dynactin-dependent transport of NuMA. The Journal of Cell Biology, 149(4), 851–862.

Merdes, A., Ramyar, K., Vechio, J. D., & Cleveland, D. W. (1996). A complex of NuMA and cytoplasmic dynein is essential for mitotic spindle assembly. Cell, 87(3), 447–458.

Mische, S., He, Y., Ma, L., Li, M., Serr, M., & Hays, T. S. (2008). Dynein light intermediate chain: an essential subunit that contributes to spindle checkpoint inactivation. Molecular Biology of the Cell, 19(11), 4918–4929.

Mitchell, D. J., Blasier, K. R., Jeffery, E. D., Ross, M. W., Pullikuth, A. K., Suo, D., et al. (2012). Trk activation of the ERK1/2 kinase pathway stimulates intermediate chain phosphorylation and recruits cytoplasmic dynein to signaling endosomes for retrograde axonal transport. The Journal of Neuroscience : the Official Journal of the Society for Neuroscience, 32(44), 15495–15510.

Moon, H. M., Youn, Y. H., Pemble, H., Yingling, J., Wittmann, T., & Wynshaw-Boris, A. (2014). LIS1 controls mitosis and mitotic spindle organization via the LIS1-NDEL1- dynein complex. Human Molecular Genetics, 23(2), 449–466.

Mori, D., Yano, Y., Toyo-oka, K., Yoshida, N., Yamada, M., Muramatsu, M., et al. (2007). NDEL1 phosphorylation by Aurora-A kinase is essential for centrosomal maturation, separation, and TACC3 recruitment. Molecular and Cellular Biology, 27(1), 352–367.

Musacchio, A., & Salmon, E. D. (2007). The spindle-assembly checkpoint in space and time. Nature Reviews. Molecular Cell Biology, 8(5), 379–393.

Nicholas, M. P., Berger, F., Rao, L., Brenner, S., Cho, C., & Gennerich, A. (2015). Cytoplasmic dynein regulates its attachment to microtubules via nucleotide state- switched mechanosensing at multiple AAA domains. Proceedings of the National Academy of Sciences of the United States of America, 112(20), 6371–6376.

137

Nicholas, M. P., Höök, P., Brenner, S., Wynne, C. L., Vallee, R. B., & Gennerich, A. (2015). Control of cytoplasmic dynein force production and processivity by its C- terminal domain. Nature Communications, 6, 6206.

Nicklas, R. B. (1983). Measurements of the force produced by the mitotic spindle in anaphase. The Journal of Cell Biology, 97(2), 542–548.

Niethammer, M., Smith, D. S., Ayala, R., Peng, J., Ko, J., Lee, M. S., et al. (2000). NUDEL is a novel Cdk5 substrate that associates with LIS1 and cytoplasmic dynein. Neuron, 28(3), 697–711.

Nigg, E. A. (2001). Mitotic kinases as regulators of cell division and its checkpoints. Nature Reviews. Molecular Cell Biology, 2(1), 21–32.

Nishikawa, Y., Inatomi, M., Iwasaki, H., & Kurisu, G. (2015). Structural Change in the Dynein Stalk Region Associated with Two Different Affinities for the Microtubule. Journal of Molecular Biology.

Nishikawa, Y., Oyama, T., Kamiya, N., Kon, T., Toyoshima, Y. Y., Nakamura, H., & Kurisu, G. (2014). Structure of the entire stalk region of the Dynein motor domain. Journal of Molecular Biology, 426(19), 3232–3245.

Numata, N., Shima, T., Ohkura, R., Kon, T., & Sutoh, K. (2011). C-sequence of the Dictyostelium cytoplasmic dynein participates in processivity modulation. FEBS Letters, 585(8), 1185–1190.

Nurminsky, D. I., Nurminskaya, M. V., Benevolenskaya, E. V., Shevelyov, Y. Y., Hartl, D. L., & Gvozdev, V. A. (1998). Cytoplasmic dynein intermediate-chain isoforms with different targeting properties created by tissue-specific alternative splicing. Molecular and Cellular Biology, 18(11), 6816–6825.

Nyarko, A., & Barbar, E. (2011). Light chain-dependent self-association of dynein intermediate chain. The Journal of Biological Chemistry, 286(2), 1556–1566.

Nyarko, A., Hare, M., Makokha, M., & Barbar, E. (2003). Interactions of LC8 with N- terminal segments of the intermediate chain of cytoplasmic dynein. TheScientificWorldJournal, 3, 647–654.

Nyarko, A., Song, Y., & Barbar, E. (2012). Intrinsic disorder in dynein intermediate chain modulates its interactions with NudE and dynactin. The Journal of Biological Chemistry, 287(30), 24884–24893.

Ogawa, K. (1991). Four ATP-binding sites in the midregion of the beta heavy chain of dynein. Nature, 352(6336), 643–645.

Ori-McKenney, K. M., Xu, J., Gross, S. P., & Vallee, R. B. (2010). A cytoplasmic dynein tail mutation impairs motor processivity. Nature Cell Biology, 12(12), 1228–1234.

138

Pandey, J. P., & Smith, D. S. (2011). A Cdk5-dependent switch regulates Lis1/Ndel1/dynein-driven organelle transport in adult axons. The Journal of Neuroscience : the Official Journal of the Society for Neuroscience, 31(47), 17207– 17219.

Paschal, B. M., Mikami, A., Pfister, K. K., & Vallee, R. B. (1992). Homology of the 74- kD cytoplasmic dynein subunit with a flagellar dynein polypeptide suggests an intracellular targeting function. The Journal of Cell Biology, 118(5), 1133–1143.

Paschal, B. M., Shpetner, H. S., & Vallee, R. B. (1987). MAP 1C is a microtubule- activated ATPase which translocates microtubules in vitro and has dynein-like properties. The Journal of Cell Biology, 105(3), 1273–1282.

Pecreaux, J., Röper, J.-C., Kruse, K., Jülicher, F., Hyman, A. A., Grill, S. W., & Howard, J. (2006). Spindle oscillations during asymmetric cell division require a threshold number of active cortical force generators. Current Biology : CB, 16(21), 2111–2122.

Pei, Z., Lang, B., Fragoso, Y. D., Shearer, K. D., Zhao, L., Mccaffery, P. J. A., et al. (2014). The expression and roles of Nde1 and Ndel1 in the adult mammalian central nervous system. Neuroscience, 271, 119–136.

Pfister, K. K. (2015). Distinct functional roles of cytoplasmic dynein defined by the intermediate chain isoforms. Experimental Cell Research, 334(1), 54–60.

Pfister, K. K., Benashski, S. E., Dillman, J. F., Patel-King, R. S., & King, S. M. (1998). Identification and molecular characterization of the p24 dynactin light chain. Cell Motility and the Cytoskeleton, 41(2), 154–167.

Pfister, K. K., Salata, M. W., Dillman, J. F., Torre, E., & Lye, R. J. (1996). Identification and developmental regulation of a neuron-specific subunit of cytoplasmic dynein. Molecular Biology of the Cell, 7(2), 331–343.

Pfister, K. K., Salata, M. W., Dillman, J. F., Vaughan, K. T., Vallee, R. B., Torre, E., & Lye, R. J. (1996). Differential expression and phosphorylation of the 74-kDa intermediate chains of cytoplasmic dynein in cultured neurons and glia. The Journal of Biological Chemistry, 271(3), 1687–1694.

Purohit, A., Tynan, S. H., Vallee, R., & Doxsey, S. J. (1999). Direct interaction of pericentrin with cytoplasmic dynein light intermediate chain contributes to mitotic spindle organization. The Journal of Cell Biology, 147(3), 481–492.

Qiu, W., Derr, N. D., Goodman, B. S., Villa, E., Wu, D., Shih, W., & Reck-Peterson, S. L. (2012). Dynein achieves processive motion using both stochastic and coordinated stepping. Nature Structural & Molecular Biology, 19(2), 193–200.

Raaijmakers, J. A., & Medema, R. H. (2014). Function and regulation of dynein in mitotic chromosome segregation. Chromosoma, 123(5), 407–422.

139

Raaijmakers, J. A., Tanenbaum, M. E., & Medema, R. H. (2013). Systematic dissection of dynein regulators in mitosis. The Journal of Cell Biology, 201(2), 201–215.

Raaijmakers, J. A., van Heesbeen, R. G. H. P., Meaders, J. L., Geers, E. F., Fernandez- Garcia, B., Medema, R. H., & Tanenbaum, M. E. (2012). Nuclear envelope-associated dynein drives prophase centrosome separation and enables Eg5-independent bipolar spindle formation. The EMBO Journal, 31(21), 4179–4190.

Rai, A. K., Rai, A., Ramaiya, A. J., Jha, R., & Mallik, R. (2013). Molecular adaptations allow dynein to generate large collective forces inside cells. Cell, 152(1-2), 172–182.

Rao, L., Romes, E. M., Nicholas, M. P., Brenner, S., Tripathy, A., Gennerich, A., & Slep, K. C. (2013). The yeast dynein Dyn2-Pac11 complex is a dynein dimerization/processivity factor: structural and single-molecule characterization. Molecular Biology of the Cell, 24(15), 2362–2377.

Rapali, P., Szenes, Á., Radnai, L., Bakos, A., Pál, G., & Nyitray, L. (2011). DYNLL/LC8: a light chain subunit of the dynein motor complex and beyond. The FEBS Journal, 278(17), 2980–2996.

Reck-Peterson, S. L., & Vale, R. D. (2004). Molecular dissection of the roles of nucleotide binding and hydrolysis in dynein's AAA domains in Saccharomyces cerevisiae. Proceedings of the National Academy of Sciences of the United States of America, 101(6), 1491–1495.

Reck-Peterson, S. L., Yildiz, A., Carter, A. P., Gennerich, A., Zhang, N., & Vale, R. D. (2006). Single-molecule analysis of dynein processivity and stepping behavior. Cell, 126(2), 335–348.

Reddy, B. J. N., Mattson, M., Wynne, C. L., Vadpey, O., Durra, A., Chapman, D., et al. (2016). Load-induced enhancement of Dynein force production by LIS1-NudE in vivo and in vitro. Nature Communications, 7, 12259.

Redwine, W. B., Hernández-López, R., Zou, S., Huang, J., Reck-Peterson, S. L., & Leschziner, A. E. (2012). Structural basis for microtubule binding and release by dynein. Science (New York, N.Y.), 337(6101), 1532–1536.

Rieder, C. L., & Salmon, E. D. (1994). Motile kinetochores and polar ejection forces dictate chromosome position on the vertebrate mitotic spindle. The Journal of Cell Biology, 124(3), 223–233.

Roberts, A. J., Malkova, B., Walker, M. L., Sakakibara, H., Numata, N., Kon, T., et al. (2012). ATP-driven remodeling of the linker domain in the dynein motor. Structure (London, England : 1993), 20(10), 1670–1680.

Roberts, A. J., Numata, N., Walker, M. L., Kato, Y. S., Malkova, B., Kon, T., et al. (2009). AAA+ Ring and linker swing mechanism in the dynein motor. Cell, 136(3), 485– 495.

140

Rodríguez-Crespo, I., Yélamos, B., Roncal, F., Albar, J. P., Ortiz de Montellano, P. R., & Gavilanes, F. (2001). Identification of novel cellular proteins that bind to the LC8 dynein light chain using a pepscan technique. FEBS Letters, 503(2-3), 135–141.

Ross, J. L., Wallace, K., Shuman, H., Goldman, Y. E., & Holzbaur, E. L. F. (2006). Processive bidirectional motion of dynein-dynactin complexes in vitro. Nature Cell Biology, 8(6), 562–570.

Rusan, N. M., Tulu, U. S., Fagerstrom, C., & Wadsworth, P. (2002). Reorganization of the microtubule array in prophase/prometaphase requires cytoplasmic dynein-dependent microtubule transport. The Journal of Cell Biology, 158(6), 997–1003.

Salina, D., Bodoor, K., Eckley, D. M., Schroer, T. A., Rattner, J. B., & Burke, B. (2002). Cytoplasmic dynein as a facilitator of nuclear envelope breakdown. Cell, 108(1), 97–107.

Samsó, M., & Koonce, M. P. (2004). 25 Angstrom resolution structure of a cytoplasmic dynein motor reveals a seven-member planar ring. Journal of Molecular Biology, 340(5), 1059–1072.

Sapir, T., Cahana, A., Seger, R., Nekhai, S., & Reiner, O. (1999). LIS1 is a microtubule- associated phosphoprotein. European Journal of Biochemistry / FEBS, 265(1), 181–188.

Sasaki, S., Mori, D., Toyo-oka, K., Chen, A., Garrett-Beal, L., Muramatsu, M., et al. (2005). Complete loss of Ndel1 results in neuronal migration defects and early embryonic lethality. Molecular and Cellular Biology, 25(17), 7812–7827.

Sasaki, S., Shionoya, A., Ishida, M., Gambello, M. J., Yingling, J., Wynshaw-Boris, A., & Hirotsune, S. (2000). A LIS1/NUDEL/cytoplasmic dynein heavy chain complex in the developing and adult nervous system. Neuron, 28(3), 681–696.

Savoian, M. S., Goldberg, M. L., & Rieder, C. L. (2000). The rate of poleward chromosome motion is attenuated in Drosophila zw10 and rod mutants. Nature Cell Biology, 2(12), 948–952.

Scaërou, F., Aguilera, I., Saunders, R., Kane, N., Blottière, L., & Karess, R. (1999). The rough deal protein is a new kinetochore component required for accurate chromosome segregation in Drosophila. Journal of Cell Science, 112 ( Pt 21), 3757–3768.

Scaërou, F., Starr, D. A., Piano, F., Papoulas, O., Karess, R. E., & Goldberg, M. L. (2001). The ZW10 and Rough Deal checkpoint proteins function together in a large, evolutionarily conserved complex targeted to the kinetochore. Journal of Cell Science, 114(Pt 17), 3103–3114.

Scherer, J., Yi, J., & Vallee, R. B. (2014). PKA-dependent dynein switching from lysosomes to adenovirus: a novel form of host-virus competition. The Journal of Cell Biology, 205(2), 163–177.

141

Schlager, M. A., Hoang, H. T., Urnavicius, L., Bullock, S. L., & Carter, A. P. (2014). In vitro reconstitution of a highly processive recombinant human dynein complex. The EMBO Journal, 33(17), 1855–1868.

Schmidt, H., Gleave, E. S., & Carter, A. P. (2012). Insights into dynein motor domain function from a 3.3-Å crystal structure. Nature Structural & Molecular Biology, 19(5), 492–7– S1.

Schmidt, H., Zalyte, R., Urnavicius, L., & Carter, A. P. (2015). Structure of human cytoplasmic dynein-2 primed for its power stroke. Nature, 518(7539), 435–438.

Schmoranzer, J., Fawcett, J. P., Segura, M., Tan, S., Vallee, R. B., Pawson, T., & Gundersen, G. G. (2009). Par3 and dynein associate to regulate local microtubule dynamics and centrosome orientation during migration. Current Biology : CB, 19(13), 1065–1074.

Schroeder, C. M., Ostrem, J. M. L., Hertz, N. T., & Vale, R. D. (2014). A Ras-like domain in the light intermediate chain bridges the dynein motor to a cargo-binding region. eLife, 3, e03351.

Schroer, T. A. (2004). Dynactin. Annual Review of Cell and Developmental Biology, 20(1), 759–779.

Schroer, T. A., Steuer, E. R., & Sheetz, M. P. (1989). Cytoplasmic dynein is a minus end- directed motor for membranous organelles. Cell, 56(6), 937–946.

Schwamborn, J. C., & Knoblich, J. A. (2008). LIS1 and spindle orientation in neuroepithelial cells. Cell Stem Cell, 2(3), 193–194.

Seldin, L., Poulson, N. D., Foote, H. P., & Lechler, T. (2013). NuMA localization, stability, and function in spindle orientation involve 4.1 and Cdk1 interactions. Molecular Biology of the Cell, 24(23), 3651–3662.

Sharp, D. J., Rogers, G. C., & Scholey, J. M. (2000). Cytoplasmic dynein is required for poleward chromosome movement during mitosis in Drosophila embryos. Nature Cell Biology, 2(12), 922–930.

Sheetz, M. P., Steuer, E. R., & Schroer, T. A. (1989). The mechanism and regulation of fast axonal transport. Trends in Neurosciences, 12(11), 474–478.

Sheffield, P. J., McMullen, T. W., Li, J., Ho, Y. S., Garrard, S. M., Derewenda, U., & Derewenda, Z. S. (2001). Preparation and crystal structure of the recombinant alpha(1)/alpha(2) catalytic heterodimer of bovine brain platelet-activating factor acetylhydrolase Ib. Protein Engineering, 14(7), 513–519.

Short, B., Preisinger, C., Schaletzky, J., Kopajtich, R., & Barr, F. A. (2002). The Rab6 GTPase regulates recruitment of the dynactin complex to Golgi membranes. Current Biology : CB, 12(20), 1792–1795.

142

Shpetner, H. S., Paschal, B. M., & Vallee, R. B. (1988). Characterization of the microtubule-activated ATPase of brain cytoplasmic dynein (MAP 1C). The Journal of Cell Biology, 107(3), 1001–1009.

Shu, T., Ayala, R., Nguyen, M.-D., Xie, Z., Gleeson, J. G., & Tsai, L.-H. (2004). Ndel1 operates in a common pathway with LIS1 and cytoplasmic dynein to regulate cortical neuronal positioning. Neuron, 44(2), 263–277.

Silva, P. M. A., Reis, R. M., Bolanos-Garcia, V. M., Florindo, C., Tavares, Á. A., & Bousbaa, H. (2014). Dynein-dependent transport of spindle assembly checkpoint proteins off kinetochores toward spindle poles. FEBS Letters, 588(17), 3265–3273.

Silvanovich, A., Li, M.-G., Serr, M., Mische, S., & Hays, T. S. (2003). The third P-loop domain in cytoplasmic dynein heavy chain is essential for dynein motor function and ATP-sensitive microtubule binding. Molecular Biology of the Cell, 14(4), 1355–1365.

Sivaram, M. V. S., Wadzinski, T. L., Redick, S. D., Manna, T., & Doxsey, S. J. (2009). Dynein light intermediate chain 1 is required for progress through the spindle assembly checkpoint. The EMBO Journal, 28(7), 902–914.

Smith, D. S., Niethammer, M., Ayala, R., Zhou, Y., Gambello, M. J., Wynshaw-Boris, A., & Tsai, L. H. (2000). Regulation of cytoplasmic dynein behaviour and microtubule organization by mammalian Lis1. Nature Cell Biology, 2(11), 767–775.

Soares, D. C., Bradshaw, N. J., Zou, J., Kennaway, C. K., Hamilton, R. S., Chen, Z. A., et al. (2012). The mitosis and neurodevelopment proteins NDE1 and NDEL1 form dimers, tetramers, and polymers with a folded back structure in solution. The Journal of Biological Chemistry, 287(39), 32381–32393.

Splinter, D., Razafsky, D. S., Schlager, M. A., Serra-Marques, A., Grigoriev, I., Demmers, J., et al. (2012). BICD2, dynactin, and LIS1 cooperate in regulating dynein recruitment to cellular structures. Molecular Biology of the Cell, 23(21), 4226–4241.

Splinter, D., Tanenbaum, M. E., Lindqvist, A., Jaarsma, D., Flotho, A., Yu, K. L., et al. (2010). Bicaudal D2, dynein, and kinesin-1 associate with nuclear pore complexes and regulate centrosome and nuclear positioning during mitotic entry. PLoS Biology, 8(4), e1000350.

Starr, D. A., Saffery, R., Li, Z., Simpson, A. E., Choo, K. H., Yen, T. J., & Goldberg, M. L. (2000). HZwint-1, a novel human kinetochore component that interacts with HZW10. Journal of Cell Science, 113 ( Pt 11), 1939–1950.

Starr, D. A., Williams, B. C., Hays, T. S., & Goldberg, M. L. (1998). ZW10 helps recruit dynactin and dynein to the kinetochore. The Journal of Cell Biology, 142(3), 763–774.

Steffen, W., Hodgkinson, J. L., & Wiche, G. (1996). Immunogold localisation of the intermediate chain within the protein complex of cytoplasmic dynein. Journal of Structural Biology, 117(3), 227–235.

143

Steffen, W., Karki, S., Vaughan, K. T., Vallee, R. B., Holzbaur, E. L., Weiss, D. G., & Kuznetsov, S. A. (1997). The involvement of the intermediate chain of cytoplasmic dynein in binding the motor complex to membranous organelles of Xenopus oocytes. Molecular Biology of the Cell, 8(10), 2077–2088.

Stehman, S. A., Chen, Y., McKenney, R. J., & Vallee, R. B. (2007). NudE and NudEL are required for mitotic progression and are involved in dynein recruitment to kinetochores. The Journal of Cell Biology, 178(4), 583–594.

Steuer, E. R., Wordeman, L., Schroer, T. A., & Sheetz, M. P. (1990). Localization of cytoplasmic dynein to mitotic spindles and kinetochores. Nature, 345(6272), 266–268.

Stukenberg, P. T., Lustig, K. D., McGarry, T. J., King, R. W., Kuang, J., & Kirschner, M. W. (1997). Systematic identification of mitotic phosphoproteins. Current Biology : CB, 7(5), 338–348.

Susalka, S. J., & Pfister, K. K. (2000). Cytoplasmic dynein subunit heterogeneity: implications for axonal transport. Journal of Neurocytology, 29(11-12), 819–829.

Sweeney, K. J., Prokscha, A., & Eichele, G. (2001). NudE-L, a novel Lis1-interacting protein, belongs to a family of vertebrate coiled-coil proteins. Mechanisms of Development, 101(1-2), 21–33.

Tai, C.-Y., Dujardin, D. L., Faulkner, N. E., & Vallee, R. B. (2002). Role of dynein, dynactin, and CLIP-170 interactions in LIS1 kinetochore function. The Journal of Cell Biology, 156(6), 959–968.

Takahashi, Y., Edamatsu, M., & Toyoshima, Y. Y. (2004). Multiple ATP-hydrolyzing sites that potentially function in cytoplasmic dynein. Proceedings of the National Academy of Sciences of the United States of America, 101(35), 12865–12869.

Tan, S. C., Scherer, J., & Vallee, R. B. (2011). Recruitment of dynein to late endosomes and lysosomes through light intermediate chains. Molecular Biology of the Cell, 22(4), 467–477.

Tanenbaum, M. E., Macůrek, L., Galjart, N., & Medema, R. H. (2008). Dynein, Lis1 and CLIP-170 counteract Eg5-dependent centrosome separation during bipolar spindle assembly. The EMBO Journal, 27(24), 3235–3245.

Toba, S., Watanabe, T. M., Yamaguchi-Okimoto, L., Toyoshima, Y. Y., & Higuchi, H. (2006). Overlapping hand-over-hand mechanism of single molecular motility of cytoplasmic dynein. Proceedings of the National Academy of Sciences of the United States of America, 103(15), 5741–5745.

Torisawa, T., Ichikawa, M., Furuta, A., Saito, K., Oiwa, K., Kojima, H., et al. (2014). Autoinhibition and cooperative activation mechanisms of cytoplasmic dynein. Nature Cell Biology, 16(11), 1118–1124.

144

Torisawa, T., Nakayama, A., Furuta, K., Yamada, M., Hirotsune, S., & Toyoshima, Y. Y. (2011). Functional dissection of LIS1 and NDEL1 towards understanding the molecular mechanisms of cytoplasmic dynein regulation. The Journal of Biological Chemistry, 286(3), 1959–1965.

Toropova, K., Zou, S., Roberts, A. J., Redwine, W. B., Goodman, B. S., Reck-Peterson, S. L., & Leschziner, A. E. (2014). Lis1 regulates dynein by sterically blocking its mechanochemical cycle. eLife, 3.

Toyo-oka, K., Sasaki, S., Yano, Y., Mori, D., Kobayashi, T., Toyoshima, Y. Y., et al. (2005). Recruitment of katanin p60 by phosphorylated NDEL1, an LIS1 interacting protein, is essential for mitotic cell division and neuronal migration. Human Molecular Genetics, 14(21), 3113–3128. doi:10.1093/hmg/ddi339

Tripathy, S. K., Weil, S. J., Chen, C., Anand, P., Vallee, R. B., & Gross, S. P. (2014). Autoregulatory mechanism for dynactin control of processive and diffusive dynein transport. Nature Cell Biology, 16(12), 1192–1201.

Tsai, J.-W., Bremner, K. H., & Vallee, R. B. (2007). Dual subcellular roles for LIS1 and dynein in radial neuronal migration in live brain tissue. Nature Neuroscience, 10(8), 970– 979.

Tsai, J.-W., Chen, Y., Kriegstein, A. R., & Vallee, R. B. (2005). LIS1 RNA interference blocks neural stem cell division, morphogenesis, and motility at multiple stages. The Journal of Cell Biology, 170(6), 935–945.

Tynan, S. H., Gee, M. A., & Vallee, R. B. (2000). Distinct but overlapping sites within the cytoplasmic dynein heavy chain for dimerization and for intermediate chain and light intermediate chain binding. The Journal of Biological Chemistry, 275(42), 32769–32774.

Tynan, S. H., Purohit, A., Doxsey, S. J., & Vallee, R. B. (2000). Light intermediate chain 1 defines a functional subfraction of cytoplasmic dynein which binds to pericentrin. The Journal of Biological Chemistry, 275(42), 32763–32768.

Urnavicius, L., Zhang, K., Diamant, A. G., Motz, C., Schlager, M. A., Yu, M., et al. (2015). The structure of the dynactin complex and its interaction with dynein. Science (New York, N.Y.), 347(6229), 1441–1446.

Vallee, R. B., Varma, D., & Dujardin, D. L. (2006). ZW10 function in mitotic checkpoint control, dynein targeting and membrane trafficking: is dynein the unifying theme? Cell Cycle (Georgetown, Tex.), 5(21), 2447–2451.

Vallee, R. B., McKenney, R. J., & Ori-McKenney, K. M. (2012). Multiple modes of cytoplasmic dynein regulation. Nature Cell Biology, 14(3), 224–230. van Heesbeen, R. G. H. P., Raaijmakers, J. A., Tanenbaum, M. E., & Medema, R. H. (2013). Nuclear envelope-associated dynein cooperates with Eg5 to drive prophase centrosome separation. Communicative & Integrative Biology, 6(3), e23841.

145

van Heesbeen, R. G. H. P., Tanenbaum, M. E., & Medema, R. H. (2014). Balanced activity of three mitotic motors is required for bipolar spindle assembly and chromosome segregation. Cell Reports, 8(4), 948–956.

Varma, D., Dawn, A., Ghosh-Roy, A., Weil, S. J., Ori-McKenney, K. M., Zhao, Y., et al. (2010). Development and application of in vivo molecular traps reveals that dynein light chain occupancy differentially affects dynein-mediated processes. Proceedings of the National Academy of Sciences of the United States of America, 107(8), 3493–3498.

Varma, D., Dujardin, D. L., Stehman, S. A., & Vallee, R. B. (2006). Role of the kinetochore/cell cycle checkpoint protein ZW10 in interphase cytoplasmic dynein function. The Journal of Cell Biology, 172(5), 655–662. doi:10.1083/jcb.200510120

Varma, D., Monzo, P., Stehman, S. A., & Vallee, R. B. (2008). Direct role of dynein motor in stable kinetochore-microtubule attachment, orientation, and alignment. The Journal of Cell Biology, 182(6), 1045–1054.

Varma, D., Wan, X., Cheerambathur, D., Gassmann, R., Suzuki, A., Lawrimore, J., et al. (2013). Spindle assembly checkpoint proteins are positioned close to core microtubule attachment sites at kinetochores. The Journal of Cell Biology, 202(5), 735–746.

Vaughan, K. T., & Vallee, R. B. (1995). Cytoplasmic dynein binds dynactin through a direct interaction between the intermediate chains and p150Glued. The Journal of Cell Biology, 131(6 Pt 1), 1507–1516.

Vaughan, K. T., Tynan, S. H., Faulkner, N. E., Echeverri, C. J., & Vallee, R. B. (1999). Colocalization of cytoplasmic dynein with dynactin and CLIP-170 at microtubule distal ends. Journal of Cell Science, 112 ( Pt 10), 1437–1447.

Vaughan, P. S., Leszyk, J. D., & Vaughan, K. T. (2001). Cytoplasmic dynein intermediate chain phosphorylation regulates binding to dynactin. The Journal of Biological Chemistry, 276(28), 26171–26179.

Vaughan, P. S., Miura, P., Henderson, M., Byrne, B., & Vaughan, K. T. (2002). A role for regulated binding of p150(Glued) to microtubule plus ends in organelle transport. The Journal of Cell Biology, 158(2), 305–319.

Vergnolle, M. A. S., & Taylor, S. S. (2007). Cenp-F links kinetochores to Ndel1/Nde1/Lis1/dynein microtubule motor complexes. Current Biology : CB, 17(13), 1173–1179.

Volkov, V. A., Grissom, P. M., Arzhanik, V. K., Zaytsev, A. V., Renganathan, K., McClure-Begley, T., et al. (2015). Centromere protein F includes two sites that couple efficiently to depolymerizing microtubules. The Journal of Cell Biology, 209(6), 813– 828.

146

Vos, L. J., Famulski, J. K., & Chan, G. K. T. (2011). hZwint-1 bridges the inner and outer kinetochore: identification of the kinetochore localization domain and the hZw10- interaction domain. The Biochemical Journal, 436(1), 157–168.

Wainman, A., Creque, J., Williams, B., Williams, E. V., Bonaccorsi, S., Gatti, M., & Goldberg, M. L. (2009). Roles of the Drosophila NudE protein in kinetochore function and centrosome migration. Journal of Cell Science, 122(Pt 11), 1747–1758.

Wainman, A., Giansanti, M. G., Goldberg, M. L., & Gatti, M. (2012). The Drosophila RZZ complex - roles in membrane trafficking and cytokinesis. Journal of Cell Science, 125(Pt 17), 4014–4025.

Wang, H., Hu, X., Ding, X., Dou, Z., Yang, Z., Shaw, A. W., et al. (2004). Human Zwint-1 specifies localization of Zeste White 10 to kinetochores and is essential for mitotic checkpoint signaling. The Journal of Biological Chemistry, 279(52), 54590– 54598.

Wang, S., & Zheng, Y. (2011). Identification of a novel dynein binding domain in nudel essential for spindle pole organization in Xenopus egg extract. The Journal of Biological Chemistry, 286(1), 587–593.

Wang, S., Ketcham, S. A., Schön, A., Goodman, B., Wang, Y., Yates, J., et al. (2013). Nudel/NudE and Lis1 promote dynein and dynactin interaction in the context of spindle morphogenesis. Molecular Biology of the Cell, 24(22), 3522–3533.

Waterman-Storer, C. M., Karki, S. B., Kuznetsov, S. A., Tabb, J. S., Weiss, D. G., Langford, G. M., & Holzbaur, E. L. (1997). The interaction between cytoplasmic dynein and dynactin is required for fast axonal transport. Proceedings of the National Academy of Sciences of the United States of America, 94(22), 12180–12185.

Waterman-Storer, C. M., Karki, S., & Holzbaur, E. L. (1995). The p150Glued component of the dynactin complex binds to both microtubules and the actin-related protein centractin (Arp-1). Proceedings of the National Academy of Sciences of the United States of America, 92(5), 1634–1638.

Whyte, J., Bader, J. R., Tauhata, S. B. F., Raycroft, M., Hornick, J., Pfister, K. K., et al. (2008). Phosphorylation regulates targeting of cytoplasmic dynein to kinetochores during mitosis. The Journal of Cell Biology, 183(5), 819–834.

Williams, B. C., Gatti, M., & Goldberg, M. L. (1996). Bipolar spindle attachments affect redistributions of ZW10, a Drosophila centromere/kinetochore component required for accurate chromosome segregation. The Journal of Cell Biology, 134(5), 1127–1140.

Williams, B. C., Li, Z., Liu, S., Williams, E. V., Leung, G., Yen, T. J., & Goldberg, M. L. (2003). Zwilch, a new component of the ZW10/ROD complex required for kinetochore functions. Molecular Biology of the Cell, 14(4), 1379–1391

147

Williams, J. C., Roulhac, P. L., Roy, A. G., Vallee, R. B., Fitzgerald, M. C., & Hendrickson, W. A. (2007). Structural and thermodynamic characterization of a cytoplasmic dynein light chain-intermediate chain complex. Proceedings of the National Academy of Sciences of the United States of America, 104(24), 10028–10033.

Wilson, M. J., Salata, M. W., Susalka, S. J., & Pfister, K. K. (2001). Light chains of mammalian cytoplasmic dynein: identification and characterization of a family of LC8 light chains. Cell Motility and the Cytoskeleton, 49(4), 229–240. doi:10.1002/cm.1036

Wojcik, E., Basto, R., Serr, M., Scaërou, F., Karess, R., & Hays, T. (2001). Kinetochore dynein: its dynamics and role in the transport of the Rough deal checkpoint protein. Nature Cell Biology, 3(11), 1001–1007.

Woodard, G. E., Huang, N.-N., Cho, H., Miki, T., Tall, G. G., & Kehrl, J. H. (2010). Ric- 8A and Gi alpha recruit LGN, NuMA, and dynein to the cell cortex to help orient the mitotic spindle. Molecular and Cellular Biology, 30(14), 3519–3530.

Wynshaw-Boris, A., & Gambello, M. J. (2001). LIS1 and dynein motor function in neuronal migration and development. Genes & Development, 15(6), 639–651.

Xie, Y., Jüschke, C., Esk, C., Hirotsune, S., & Knoblich, J. A. (2013). The phosphatase PP4c controls spindle orientation to maintain proliferative symmetric divisions in the developing neocortex. Neuron, 79(2), 254–265.

Yadav, S., Puthenveedu, M. A., & Linstedt, A. D. (2012). Golgin160 recruits the dynein motor to position the Golgi apparatus. Developmental Cell, 23(1), 153–165.

Yamada, M., Hirotsune, S., & Wynshaw-Boris, A. (2010). The essential role of LIS1, NDEL1 and Aurora-A in polarity formation and microtubule organization during neurogensis. Cell Adhesion & Migration, 4(2), 180–184.

Yamada, M., Toba, S., Yoshida, Y., Haratani, K., Mori, D., Yano, Y., et al. (2008). LIS1 and NDEL1 coordinate the plus-end-directed transport of cytoplasmic dynein. The EMBO Journal, 27(19), 2471–2483.

Yamamoto, T. G., Watanabe, S., Essex, A., & Kitagawa, R. (2008). SPDL-1 functions as a kinetochore receptor for MDF-1 in Caenorhabditis elegans. The Journal of Cell Biology, 183(2), 187–194.

Yan, X., Li, F., Liang, Y., Shen, Y., Zhao, X., Huang, Q., & Zhu, X. (2003). Human Nudel and NudE as regulators of cytoplasmic dynein in poleward protein transport along the mitotic spindle. Molecular and Cellular Biology, 23(4), 1239–1250.

Yang, Z., Guo, J., Chen, Q., Ding, C., Du, J., & Zhu, X. (2005). Silencing mitosin induces misaligned chromosomes, premature chromosome decondensation before anaphase onset, and mitotic cell death. Molecular and Cellular Biology, 25(10), 4062– 4074.

148

Yang, Z., Tulu, U. S., Wadsworth, P., & Rieder, C. L. (2007). Kinetochore dynein is required for chromosome motion and congression independent of the spindle checkpoint. Current Biology : CB, 17(11), 973–980.

Ye, G. J., Vaughan, K. T., Vallee, R. B., & Roizman, B. (2000). The herpes simplex virus 1 U(L)34 protein interacts with a cytoplasmic dynein intermediate chain and targets nuclear membrane. Journal of Virology, 74(3), 1355–1363.

Yi, J. Y., Ori-McKenney, K. M., McKenney, R. J., Vershinin, M., Gross, S. P., & Vallee, R. B. (2011). High-resolution imaging reveals indirect coordination of opposite motors and a role for LIS1 in high-load axonal transport. The Journal of Cell Biology, 195(2), 193–201.

Yoder, J. H., & Han, M. (2001). Cytoplasmic dynein light intermediate chain is required for discrete aspects of mitosis in Caenorhabditis elegans. Molecular Biology of the Cell, 12(10), 2921–2933.

Zhang, G., Lischetti, T., Hayward, D. G., & Nilsson, J. (2015). Distinct domains in Bub1 localize RZZ and BubR1 to kinetochores to regulate the checkpoint. Nature Communications, 6, 7162.

Zhang, J., Li, S., Musa, S., Zhou, H., & Xiang, X. (2009). Dynein light intermediate chain in Aspergillus nidulans is essential for the interaction between heavy and intermediate chains. The Journal of Biological Chemistry, 284(50), 34760–34768.

Zhang, J., Qiu, R., Arst, H. N., Peñalva, M. A., & Xiang, X. (2014). HookA is a novel dynein-early endosome linker critical for cargo movement in vivo. The Journal of Cell Biology, 204(6), 1009–1026. doi:10.1083/jcb.201308009

Zhang, Q., Wang, F., Cao, J., Shen, Y., Huang, Q., Bao, L., & Zhu, X. (2009). Nudel promotes axonal lysosome clearance and endo-lysosome formation via dynein-mediated transport. Traffic (Copenhagen, Denmark), 10(9), 1337–1349.

Zyłkiewicz, E., Kijańska, M., Choi, W.-C., Derewenda, U., Derewenda, Z. S., & Stukenberg, P. T. (2011). The N-terminal coiled-coil of Ndel1 is a regulated scaffold that recruits LIS1 to dynein. The Journal of Cell Biology, 192(3), 433–445.

149

Appendix I: Additional Work

This study investigated the role of the C-terminal domain of mammalian cytoplasmic heavy chain in force production and processivity. Yeast dynein and mammalian dynein differ in their single molecule behaviors. One example is the difference in stall force between yeast and mammalian dynein. Yeast dynein produces

7pN force events (Gennerich et al 2007), while purified rat dynein has been reported to have a stall for of approximately 1 pN (McKenney et al 2010). The question remained how the two species of dynein differ to give starkly different single-molecule behaviors.

Yeast dynein lacks 32 kDA region at the C-terminus of mammalian dynein (Citation). To study the role of this C-terminal domain, two recombinant GST rat motor domain constructs were created, one WT and lacking the c-terminal region (Delta CT). Previous analysis of recombinant mammalian motor domains had been restricted to monomeric motor domain, engineering a functional GST rat motor domain allowed for analysis of dimeric motor domains, mimicking a more physiological relevant form of dynein motor domain. In force production studies, WT-MD gave 1 pN events similar to purified rat or bovine dynein. Surprisingly the removal of the C-terminal domain increased the MD force production to ~6pN, a more yeast-like dynein. The results demonstrate the importance of this additional domain in regulation of motor function of dynein motor domain.

My role in this study was to purify baculovirus expressed motor-domain constructs, both WT-MD and DeltaCT-MD. Analysis of purification products revealed a

150

lower molecular weight fragment, which corresponds to free GST (Sup. Fig1A). I developed a hand-packed SEC column method to remove the free GST from the MD constructs (Sup Fig1B). Additionally, I confirmed the quality of the purified MD constructs by sucrose gradient centrifugation. Each construct ran as a single peak

indicating lack of aggregation (Sup Fig1C).

Supplemental Figure 1. Characterization of WT-MD and Delta CT-MD purifications. A. Representative commassie stained gel of WT-MD and Delta CT-MD protein from GST purification. B. Commassie stained gel of GST-WT and GST-Delta CT preparations applied to a hand-packed Superose size exclusion column. Contaminating free GST eluted in a later faction than WT-MD or Delta CT-MD providing samples of GST-free MD. C. Sucrose density gradient centrifugation of purified WT and Delta CT. Western blot analysis of GST motor domain constructs shows single symmetric peak of MD for both WT and Delta CT indicating lack of aggregation.

151

Appendix II: Additional Work

*I purified bacterially expressed Nde1, bovine brain purified cytoplasmic dynein and Baculovirus purified Lis1 for in vitro lipid droplet experiments to study Nde1 and Lis1’s role in in vivo force production.

Single-molecules study of Nde1 mediated inhibition of dynein bead binding activity

Biochemical studies revealed that the N-terminal domain of Nde1 mediates the

interaction with the intermediate chain of dynein. In collaboration with the Gross Lab at

UC Irvine (Especially post-doc Babu Reddy) we performed single-molecule studies on

the effects of Nde1 truncations on dynein bead-binding ability. The experiment was

carried out in an optical trap experimental set up. Dynein and Nde1 were non-

specifically recruited to latex beads and the ability for dynein to bind to the MT and

produce a force event was monitored. McKenney et al. 2009 found Nde1 alone is

inhibitory to dynein bead binding in single molecule studies. Using the C-terminal

truncations we tested the ratio of Dynein: Nde1 required to completely inhibit dynein-

bead binding activity. Wt Nde1 incubated with dynein led to 0% bead binding at a ratio of 1:100, while Nde1 10-191 required 1:1000 to inhibit dynein’s binding activity. The shortest coiled-coil (10-165) fragment took 7,000 times the amount of dynein to completely inhibit dynein’s binding to the microtubule. Nde1 N-terminal coiled-coil has

the ability to inhibit dynein’s bead binding ability suggesting it is the direct interaction

between Nde1 and dynein that changes dynein’s motor activity rather than a physical

steric interference by Nde1. The increase in Nde1 concentration required to inhibit

dynein could be due to changes in Nde1’s affinity to dynein.

152

Supplemental Figure 2. Single molecule studies of C-terminus Nde1 truncations. A. Schematic of experimental design. Dynein and Nde1 fragments are incubated together prior to recruitment, non-specifically, to latex beads. B. Quantification of the ratio of Nde1:Dynein required to completely inhibit dynein’s bead-binding ability. C-terminal truncations required increased amount to completely inhibit dynein’s bead-binding ability.

153