Cargo Specific Regulation of Cytoplasmic Dynein by Effector Proteins

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Publicly Accessible Penn Dissertations

2018

Cargo Specific Regulation Of Cytoplasmic Dynein By Effector Proteins

Mara Olenick University of Pennsylvania, [email protected]

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Recommended Citation Olenick, Mara, "Cargo Specific Regulation Of Cytoplasmic Dynein By Effector Proteins" (2018). Publicly Accessible Penn Dissertations. 3167. https://repository.upenn.edu/edissertations/3167

This paper is posted at ScholarlyCommons. https://repository.upenn.edu/edissertations/3167 For more information, please contact [email protected]. Cargo Specific Regulation Of Cytoplasmic Dynein By Effector Proteins

Abstract Axonal transport is vital for the development and survival of neurons. The transport of cargo and organelles from the axon to the cell body is driven almost completely by the molecular motor, cytoplasmic dynein. Yet, it remains unclear how dynein is spatially and temporally regulated given the variety of cargo that must be properly localized to maintain cellular function. Previous work has suggested that adaptor proteins provide a mechanism for cargo-specific egulationr of motors. During my thesis work, I have investigated the role of mammalian Hook proteins, Hook1 and Hook3, as potential motor adaptors. Using optogenetic and single molecule assays, I found that Hook proteins interact with both dynein and dynactin, to effectively activate dynein motility, inducing longer run lengths and higher velocities than the previously characterized dynein activator, BICD2. In addition, I found that complex formation requires the N-terminal domain of Hook proteins, which resembles the calponin-homology domain of EB proteins yet cannot bind directly to microtubules. In collaborative studies, we found the Hook domain directly interacts with a helix of the dynein light intermediate chain and this interaction is important for Hook-induced processive motility of dynein. In my final project, I found that Hook1 mediates the transport of TrkB-BDNF signaling endosomes in primary hippocampal neurons. Using live cell microscopy and microfluidic devices, Hook1 depletion resulted in a significant decrease in the flux and processivity of BDNF-Qdots along the mid-axon, an effect specific for Hook1 but not Hook3. ogetherT , my work suggests that dynein effectors like Hook proteins can differentially regulate dynein to allow for organelle-specific tuning of the motor for precise intracellular trafficking.

Degree Type Dissertation

Degree Name Doctor of Philosophy (PhD)

Graduate Group Biochemistry & Molecular Biophysics

First Advisor Erika L. Holzbaur

Subject Categories Biochemistry | Biophysics | Cell Biology

This dissertation is available at ScholarlyCommons: https://repository.upenn.edu/edissertations/3167 CARGO SPECIFIC REGULATION OF CYTOPLASMIC DYNEIN BY EFFECTOR PROTEINS

Mara Alizabeth Olenick

A DISSERTATION

Biochemistry and Molecular Biophysics

Presented to the Faculties of the University of Pennsylvania

Partial Fulfillment of the Requirements for the

Degree of Doctor of Philosophy

2018

Supervisor of Dissertation

______

Erika L. F. Holzbaur, PhD, Professor of Physiology

Graduate Group Chairperson

______

Kim A. Sharp, PhD, Associate Professor of Biochemistry and Biophysics

Dissertation Committee

E. Michael Ostap, PhD, Professor of Physiology

Roberto Dominguez, PhD, Professor of Physiology

Michael A. Lampson, PhD, Associate Professor of Biology

Erfei Bi, PhD, Professor of Cell and Developmental Biology

CARGO SPECIFIC REGULATION OF CYTOPLASMIC DYNEIN BY EFFECTOR PROTEINS

2018

Mara Alizabeth Olenick

ACKNOWLEDGMENT

Getting a PhD can be a very long journey, full of ups and downs but having a good support system can make it an amazing ride. First, I would like to thank my mentor, Erika

Holzbaur for always being positive and encouraging throughout my thesis. Without her encouragement and support along the way, I wouldn’t have the confidence in myself as a scientist that I have today. I would also like to thank my committee members who always pushed me to think more analytically and be more rigorous with my science. Special thanks to Roberto

Dominquez for being a great collaborator throughout my thesis. His scientific advice and structural expertise was critical to my thesis work.

I want to thank the Biochemistry and Molecular Biophysics Graduate Group facility and staff for being supportive and helpful, especially during the early years when you are just trying to get use to the graduate school life style. A big thank you to the Pennsylvania Muscle Institute, a community with such collaborative spirit and excitement for science. The many years of PMI journal club have taught me so much about critical thinking and the art of science, which no class could ever achieve.

I would not be writing this thesis without the tremendous support of the Holzbaur lab members of past and present. First, the biggest thank you to Mariko Tokito, our lab manager for being the cloning master and keeping our lab running with the upmost precision. I would also like to thank Karen Wallace Jahn for being the best lab veterinarian and providing the wit and humor that keeps all of us grounded in the real world. These two women provide such experience and expertise that is invaluable to the Holzbaur lab. Over the years, I have had the pleasure of working with the most talented post-docs and graduate students, who provided a variety of expertise and experience to learn from. Special thank you to Meredith Wilson, Swathi Ayloo,

Sandra Maday and Amy Ghiretti for being my original lab lunch crew and helping me through the first half of my graduate school career. Their support and friendship were vital during those early years. Another big thank you to Pallavi Gopal, Chantell Evans, and Andrea Stavoe for being my second lab lunch crew and helping me make it to the graduate school finishing line. Our afternoon iii crossword breaks kept me sane when the science was driving me insane. I would like to thank everyone in the Holzbaur lab, past and present, for making lab a great place to grow as a scientist.

Of course, I could not have survived the many years of graduate school without the support from my family and friends. I would like to thank all the wonderful friends that I have gained in graduate school over the years. A special thanks to my fellow 2013 BMB incoming class members, also known as BAMB. Not only did we make it through those first two years of class together, but we gained friendship that will last a lifetime. I would also like to thank my family members who got me away from work every now and then to keep me grounded. A special thank you to my parents, Jodie and William Olenick for always believing in me and telling me to shoot for the stars. Finally, I would like to thank my partner in crime and light of my life, Kevin

Walker for always being supportive and loving even when I’m stressed and cranky.

Thank you to everyone who encouraged me to keep going!

iv ABSTRACT

CARGO SPECIFIC REGULATION OF CYTOPLASMIC DYNEIN BY EFFECTOR PROTEINS

Mara A. Olenick

Dr. Erika L.F. Holzbaur

Axonal transport is vital for the development and survival of neurons. The transport of cargo and organelles from the axon to the cell body is driven almost completely by the molecular motor, cytoplasmic dynein. Yet, it remains unclear how dynein is spatially and temporally regulated given the variety of cargo that must be properly localized to maintain cellular function. Previous work has suggested that adaptor proteins provide a mechanism for cargo-specific regulation of motors. During my thesis work, I have investigated the role of mammalian Hook proteins, Hook1 and

Hook3, as potential motor adaptors. Using optogenetic and single molecule assays, I found that Hook proteins interact with both dynein and dynactin, to effectively activate dynein motility, inducing longer run lengths and higher velocities than the previously characterized dynein activator, BICD2. In addition, I found that complex formation requires the N-terminal domain of Hook proteins, which resembles the calponin- homology domain of EB proteins yet cannot bind directly to microtubules. In collaborative studies, we found the Hook domain directly interacts with a helix of the dynein light intermediate chain and this interaction is important for Hook-induced processive motility of dynein. In my final project, I found that Hook1 mediates the transport of TrkB-BDNF signaling endosomes in primary hippocampal neurons. Using live cell microscopy and microfluidic devices, Hook1 depletion resulted in a significant decrease in the flux and processivity of BDNF-Qdots along the mid-axon, an effect

v specific for Hook1 but not Hook3. Together, my work suggests that dynein effectors like

Hook proteins can differentially regulate dynein to allow for organelle-specific tuning of the motor for precise intracellular trafficking.

TABLE OF CONTENTS

ACKNOWLEDGMENT ...... III

ABSTRACT ...... V

LIST OF FIGURES ...... IX

CHAPTER 1: INTRODUCTION ...... 1

I. Intracellular Transport ...... 1

II. Microtubules ...... 3

III. Cytoplasmic Dynein ...... 7

IV. Dynein Effector Proteins ...... 13

V. Hook Proteins ...... 24

CHAPTER 2: HOOK ADAPTORS INDUCE UNIDIRECTIONAL PROCESSIVE MOTILITY BY ENHANCING THE DYNEIN-DYNACTIN INTERACTION ...... 28

I. Summary ...... 29

II. Introduction ...... 30

III. Results ...... 33

IV. Discussion ...... 51

V. Materials and Methods ...... 56

CHAPTER 3: A CONSERVED INTERACTION OF THE DYNEIN LIGHT INTERMEDIATE CHAIN WITH DYNEIN-DYNACTIN EFFECTORS NECESSARY FOR PROCESSIVITY ...... 62

I. Summary ...... 63

II. Introduction ...... 64

III. Results ...... 66

IV. Discussion ...... 87

V. Materials and Methods ...... 93

CHAPTER 4: DYNEIN ACTIVATOR HOOK1 IS REQUIRED FOR TRAFFICKING OF BDNF- SIGNALING ENDOSOMES IN NEURONS ...... 101

vii

I. Summary ...... 102

II. Introduction ...... 103

III. Results ...... 107

IV. Discussion ...... 126

V. Materials and Methods ...... 130

CHAPTER 5: DISCUSSION AND FUTURE DIRECTIONS ...... 135

BIBLIOGRAPHY ...... 145

viii

LIST OF FIGURES

Figure 1.1 Microtubule dynamics and organization ...... 6

Figure 1.2 Dynein-Dynactin schematic ...... 12

Figure 1.3 Model of dynein activation ...... 16

Figure 1.4 Domain structure of dynein effectors ...... 21

Figure 2.1 Hook proteins redistribute peroxisomes to perinuclear region in an optogenetic assay ...... 34

Figure 2.2 Hook proteins differentially redistribute peroxisomes to the MTOC ...... 36

Figure 2.3 Mammalian Hook proteins interact with dynein-dynactin complex ...... 38

Figure 2.4 Hook proteins bind microtubules indirectly ...... 40

Figure 2.5 Hook proteins lack conserved regions for MT binding ...... 42

Figure 2.6 Pull-Down (PD) of Halo-Hook1 with dynein-dynactin complex requires N- terminal region ...... 44

Figure 2.7 Hook proteins display high velocities and long run lengths ...... 46

Figure 2.8 C-terminally truncated Hook proteins display similar motility to full-length ...... 49

Figure 3.1 Hook interacts with LIC1 via the N-terminal Hook domain ...... 69

Figure 3.S1 Domain architecture of Hook1 ...... 71

Figure 3.2 The conserved Helix-1 within the LIC1 effector-binding domain binds the Hook domain...... 74

Figure 3.S2 Alignment of the Effector-Binding Domain of LIC sequences ...... 76

Figure 3.3 Crystal structure of the Hook domain in complex with LIC1 Helix-1...... 79

Figure 3.4 LIC1 Helix-1 mediates the interaction with CC1-Box-containing effectors ...... 82

Figure 3.5 The Helix-1-effector interaction is important for processive motility in vitro and in cells ...... 85

Figure 3.6 Model for cargo transport by dynein-dynactin-effector complexes ...... 91

Figure 3.S3 Table of primers used in this study ...... 100

Figure 4.1 Hook1 co-migrates with Rab5- and Rab7-endosomes ...... 108

Figure 4.S1 Hook1 Knockdown does not significantly change Rab5 or Rab7 motility ...... 111

Figure 4.2 Hook1 KD reduces TrkB-BDNF signaling endosome motility ...... 113

Figure 4.3 Hook1 requires interaction with LIC1 for signaling endosome motility ...... 115

Figure 4.4 Hook1 localizes to distal axon, while other dynein effectors are enriched in other compartments ...... 117

Figure 4.5 Hook1 KD reduces flux of BDNF from distal axon ...... 120

Figure 4.6 Impaired signaling endosome motility reduces BDNF endocytosis ...... 122

Figure 4.7 Loss of Hook1 leads to loss of downstream signaling, measured by pCREB levels ...... 124

Figure 5.1 Co-IP experiments with other dynein adaptors ...... 139

Figure 5.2 Optogenetic recruitment assay with other dynein adaptors ...... 140

Figure 5.3 Summary of results for dynein effectors and adaptors ...... 141

CHAPTER 1: Introduction

I. Intracellular Transport

Spatial and temporal organization of macromolecules and organelles is vital to cellular functions and is driven by a process called intracellular transport. Transport in cells was first observed while studying the growth of peripheral nerves. In 1948, Wiess and Hiscoe described the slow flow, 1mm per day, of axoplasm material from cut or constricted nerves, which was contradictory to the prevailing view that nerve fibers were static threads to conduct nerve impulses (Dahlstrom, 2010; Weiss and Hiscoe, 1948).

Subsequent radiolabeling experiments from Samuels et al. in 1951 described the slow transport of materials in the sciatic nerve over the course of several days (Samuels et al., 1951). After the discovery of this slow transport, Lubinska and her group described fast accumulation of acetylcholinesterase in sciatic nerve from rats (Lubinska et al.,

1964). Later, Lasek would repeat radiolabeling experiments from Samuels’ group to observe the sciatic nerve after a few hours and get rates of transport that were several hundreds of mm per day, in addition to observing retrograde transport (Lasek, 1967).

Collectively, these studies laid the foundation for what would later be known as axonal transport.

At the same time as the discovery of axonal transport, intracellular transport was beginning to be described for the secretory pathway in pancreatic exocrine cells. In the

1960s, pulse chase experiments were used to show the transport of secretory proteins in pancreatic exocrine cells and tissue slices from guinea pigs (Caro and Palade, 1964;

Jamieson and Palade, 1967). Using this method, they observed transport of synthesized proteins from the ER to the Golgi via small vesicles. They also found that this transport was energy dependent since transport could be halted by respiratory inhibitors like 1

Antimycin A (Jamieson and Palade, 1968). Following this work, cell-free systems showed rapid, energy-dependent transport of membranes through the Golgi complex

(Fries and Rothman, 1980).

While these studies provided insights into intracellular transport, the mechanism behind this transport remained to be revealed. Early work in nerve fibers showed that colchicine, a drug that disrupts microtubules, induced the blockage of movement of acetylcholinesterase, implicating the cytoskeleton in intracellular transport (Kreutzberg,

1969), which had already been described in the formation of the mitotic spindle and in flagella (Ledbetter and Porter, 1964; Phillips, 1966; Roth and Daniels, 1962).

Subsequent work using pharmacological inhibition showed the importance of microtubules in transport of secretory proteins in liver and thyroid (Le Marchand et al.,

1973; Williams and Wolff, 1972). Studies in the 1980s would then bring insight into the machines for this microtubule-based transport, the molecular motors, kinesin and dynein

(Allen et al., 1985; Brady, 1985; Paschal and Vallee, 1987; Vale et al., 1985). The next sections will describe in depth the insights gained over the years into microtubules and the machinery that drives minus-end directed microtubule transport.

II. Microtubules

Microtubules are dynamic polymers of the protein tubulin, which contribute to a variety of cellular functions including acting as roads for intracellular transport, driving chromosome separation for cellular division, and maintaining cellular structures including cilia and flagella. Microtubules are made from heterodimers of α- and β-tubulin (Bryan and Wilson, 1971; Feit et al., 1971), which organize head-to-tail to give polarity to the structure, with α-tubulin exposed on the minus-end and β-tubulin exposed at the plus- end (Amos and Klug, 1974; Bergen and Borisy, 1980; Crepeau et al., 1977). Tubulin dimers polymerize longitudinally into protofilaments, which associate laterally (usually 13 protofilaments) to form the hollow tubes of microtubules that are about 25nm in diameter

(Tilney et al., 1973).

Microtubule polymerization is powered by the nucleotide guanosine triphosphate

(GTP) (Olmsted and Borisy, 1975; Weisenberg, 1972). Tubulin dimers are added at a faster rate than GTP hydrolysis, creating what is known as the GTP cap (Mitchison and

Kirschner, 1984). The GTP cap at the end of a growing microtubule allows for continued polymerization, but hydrolysis to GDP makes the microtubule lattice more prone to shrinking, also known as catastrophe (Carlier and Pantaloni, 1981; Carlier et al., 1984).

In fact, hydrolysis of GTP on β-tubulin induces conformational changes in α-tubulin which causes global lattice rearrangements and lattice strain making GDP-microtubules less stable (Alushin et al., 2014; Mandelkow et al., 1991). Polymerization can occur on either end of the microtubule but occurs with very different dynamics (Allen and Borisy,

1974; Kristofferson et al., 1986). The microtubule minus end is very stable with slow growth and very little shrinking, while the microtubule plus end is very dynamic with rapid growth and multiple catastrophe events. Microtubule dynamics can be regulated by

3 different proteins including microtubule associated proteins (MAPs) and molecular motors, which can act to stabilize or increase dynamics.

The C-terminal tails of tubulin are highly charged, acidic, unstructured regions that project from the outer surface of the microtubule lattice. They are critical for interactions with proteins on the surface of microtubules. In addition, the tails can be post-translationally modified (PTM) in a variety of ways including detyrosination, polyglutamylation, and polyglycylation, which can modulate protein interactions. For example, proteins with basic microtubule binding domains like CAP-Gly domains, such as the p150 subunit of dynactin (see next section), interact with the tail tyrosine specifically (Honnappa et al., 2006; Peris et al., 2006), while the plus-end microtubule motor kinesin-1 prefers detyrosinated microtubules (Konishi and Setou, 2009; Liao and

Gundersen, 1998). PTMs can also be indicators of microtubule dynamics; for instance, detyrosination is generally found on long lived stable microtubules, such as those found in the neuronal axons (Cambray-Deakin and Burgoyne, 1987; Robson and Burgoyne,

1989). The microtubule PTM field has had many advances including the discovery of tubulin-modifying enzymes and the creation of new tools, which continue to further insights in this area (reviewed in Janke, 2014).

Microtubule organization is important for cellular maintenance, mitosis, and creating polarized cells. In general, microtubules are organized with the minus ends anchored at a microtubule organization center (MTOC) and the plus ends orientated out.

For proliferative cells, the MTOC is at the centrosome near the nucleus and center of the cell during interphase or when not dividing. Another tubulin isoform, γ-tubulin is important for microtubule nucleation at the MTOC, where γ-tubulin forms a ring of similar size to microtubule diameter (reviewed in Kollman et al., 2011). During mitosis, microtubules are reoriented around spindle poles to help with separation of 4 chromosomes and cellular division (Brinkley et al., 1975; Inoué and Sato, 1967; Inoué et al., 1975). In specialized cells, microtubule organization can be important for creating cellular polarity. In mammalian neurons, microtubules are organized in a uniform polarized array in the axon with plus-ends oriented toward the axon terminal (Burton and

Paige, 1981; Heidemann et al., 1981), while the short and highly branched dendrites have mixed microtubule organization (Baas et al., 1988). In other tissues like intestine and muscle, differentiated cells have non-centrosomal microtubule arrays to create polarity needed for function (reviewed in Dyachuk et al., 2016; Sanchez and Feldman,

2017).

The polar nature of microtubules allows for directional transport by the molecular motors dynein and kinesin. The kinesin family of proteins are mainly plus-end directed motors with a few exceptions, while cytoplasmic dynein is a minus-end directed motor

(discussed in the next section). There are up to 45 different kinesin genes in mammals, which can be classified into 15 families based on phylogenetic analyses (reviwed in

Hirokawa and Tanaka, 2015). In contrast, there is only one major form of cytoplasmic dynein that is responsible for the majority of minus-end directed microtubule transport and is described further in the next section. The coordination and regulation of these microtubule motors for efficient transport and proper function is still a major area of ongoing research, including this thesis.

Figure 1.1 Microtubule dynamics and organization

A) Microtubule with dynamic GTP plus-end. B) Organization of microtubules in fibroblast cells with minus ends at the microtubule organization center (MTOC) and plus-ends oriented towards the periphery. C) Organization of microtubules in mammalian neurons with uniform polarity in axons and mixed polarity in dendrites.

III. Cytoplasmic Dynein

In the 1960s, the first microtubule motor was discovered in the cilia of

Tetrahymena (Gibbons, 1963; Gibbons and Rowe, 1965). They named this protein dynein, from the Greek dyne meaning force, due to its ability to slide microtubules in the cilia using ATP (Gibbons and Rowe, 1965). Since this discovery, sixteen genes have been found in humans to encode dynein heavy chains, including fourteen that function within the axoneme and one involved in retrograde intraflagellar transport (Wickstead and Gull, 2007; Yagi, 2009). The remaining gene encodes cytoplasmic dynein which drives ATP-dependent minus-end directed microtubule transport, opposite of conventional kinesins (Lye et al., 1987; Paschal and Vallee, 1987; Paschal et al., 1987).

Cytoplasmic dynein is responsible for the majority of minus-end directed transport in cells, with a few minus-end directed kinesins contributing to this motility. In animals, cytoplasmic dynein is essential due primarily to its vital role in cellular division and development (Gepner et al., 1996; Robinson et al., 1999; Vaisberg et al., 1993).

Cytoplasmic dynein (hence forth referred to as dynein) is a 1.4 MDa motor complex consisting of dimerized heavy chains (DHC, ~530 kDa), each containing a N- terminal tail, six AAA+ domains to form a motor ring and a flexible stalk with the microtubule binding domain. N-terminal tail of dynein allows for homodimerization of the heavy chains, along with binding sites for other non-catalytic subunits, including two intermediate chains (DIC, ~74 kDa) and two light intermediate chains (LIC, ~33-59 kDa).

Smaller light chains (LC, ~10-14kDa), including LC7/Roadblock, LC8, and TCTEX1 bind to the intermediate chains. These non-catalytic subunits provide sites of attachment and regulation for specific dynein functions. In vertebrates, the intermediate and light chains are encoded by two genes and expression patterns differ in various cell types, likely due to specific cellular demands for dynein (Pfister et al., 2006). 7

Unlike kinesins, dynein is a member of the AAA+ (ATPases Associated with various cellular Activities) family of proteins, which have ring-like hexamers of ATPase domains that can function in a variety of ways to unfold proteins, disassemble aggregates, and produce mechanical force (Miller and Enemark, 2016). Unlike other

AAA+ proteins, the motor domain of dynein is formed by a single polypeptide to form the hexamer of AAA+ domains, a linker domain and a microtubule binding domain. The first four AAA+ domains, AAA1-AAA4 are able to bind ATP, while AAA5 and AAA6 serve as structural units for the stalk and buttress of the microtubule binding domain. While AAA1 is the primary site of ATP hydrolysis, the nucleotide state of the other AAA domains like

AAA3 is important to coordinate microtubule binding and potentially regulate different motility properties of the motor (DeWitt et al., 2015; Nicholas et al., 2015a; Takahide Kon et al., 2004). The microtubule binding domain of dynein is at the end of a 15 nm antiparallel coiled-coil stalk that projects from the AAA4-AAA5 domain (Burgess et al.,

2003; Gee et al., 1997; Kon et al., 2011).

Dynein motility is coupled to ATP-induced conformational changes in the AAA+ ring, with the bending and straightening of the linker region. ATP binds at AAA1 to induce a weak microtubule-binding state and release of the microtubule binding domain

(MTBD) from the microtubule. When ATP is present at AAA1, the linker region also undocks and moves away from the motor ring (Cho and Vale, 2012; Roberts et al., 2012;

Schmidt et al., 2014). The hydrolysis of ATP on the dissociated motor head then causes a primed conformation to allow for rebinding to the microtubule. The forward step is coupled to the release of phosphate and ADP to reset the cycle (Holzbaur and Johnson,

1989b, 1989a; Imamula et al., 2007). Unlike the classic power-stroke mechanism, the working stroke of dynein induces interhead strain to rotate the rings through bending of

8 the stalk and hinging at the microtubule binding domain, instead of tilting the MTBD stalk as previously thought (Lippert et al., 2017).

Single molecule studies have provided insight into the motility of cytoplasmic dynein, in addition to finding differences between dynein from different species.

Mammalian dynein without the presence of interacting partners has been found in a state of autoinhibition, called the phi state, which has low microtubule binding affinity, or as a flexible dimer with very low processivity (Zhang et al., 2017). This is in contrast to yeast dynein, which displays slow processive motility and does not require tight coupling of the two motor domains (DeWitt et al., 2012; Qiu et al., 2012; Reck-Peterson et al.,

2006). Furthermore, mammalian dynein can produce up to 1.1 pN of force, while yeast dynein has a stall force of 7pN (Gennerich et al., 2007; Mallik et al., 2004; Schroeder et al., 2010). Dynein also takes many sideways or backwards steps along the microtubule lattice (Reck-Peterson et al. 2006; Ross et al. 2006) and has a variable stepping pattern, with predominantly an 8 nm step size with a maximum step size of 32 nm, which is in contrast to the uniform 8 nm step size of kinesin-1 (DeWitt et al., 2012; Mallik et al.,

2004; Qiu et al., 2012; Reck-Peterson et al., 2006). Due to the flexible nature of dynein, it has been shown to avoid obstacles better than kinesins (Dixit et al., 2008).

Dynein functions in many different roles in the cell including nuclear positioning, organelle and virus transport, chromosome dynamics, mitotic spindle orientation, axonogenesis, and cell migration (reviewed in Reck-Peterson et al., 2018; Roberts et al.,

2013). Loss of dynein is lethal for many organisms including mice and Drosophila melanogaster (Gepner et al., 1996; Harada et al., 1998). In addition, mutations in dynein lead to neurological diseases like spinal muscular atrophy with lower extremity predominance (SMALED), Charcot–Marie–Tooth (CMT) disease, and malformations of cortical development (MCD), including lissencephaly, pachygyria and polymicrogyria 9

(Fiorillo et al., 2014; Lipka et al., 2013; Poirier et al., 2013; Tsurusaki et al., 2012;

Weedon et al., 2011). Since dynein function is vital for proper development, uncovering the regulatory mechanisms of dynein motility and function has been an imperative area of research.

Dynactin

Dynein has been found to work in complex with dynactin, a 1 MDa, 23 subunit complex which is needed for the majority of dynein’s transport functions. Dynactin was found as an “activating” component that co-purified with 20S dynein and was needed for in vitro vesicle motility (Gill et al., 1991; Schroer and Sheetz, 1991). With many years of work and advancements in cryo-EM technology, the components and structure of dynactin have now been elucidated. Dynactin is comprised of an actin-like Arp1 filament with a large shoulder of alpha-helices at one end (Schroer, 2004). The Arp1 filament contains eight Arp1 molecules, one beta-actin and one Arp11 protein. Arp11 interacts with p25, p27, p62 to form the pointed-end of dynactin. The other end of dynactin is capped by an actin capping protein, CapZαβ, which forms the barbed-end of the filament. The shoulder of dynactin sits on the Arp1 filament near the barbed-end. It is comprised of two copies of p24, four copies of p50 (dynamtin), and two copies of p150Glued (Chowdhury et al., 2015; Urnavicius et al., 2015).

The p150Glued subunit is the largest subunit of dynactin and projects out of the shoulder as a coiled-coil with a N-terminal CAP-Gly domain, which binds microtubules

(Waterman-Storer et al., 1995). In early work, p150Glued was found to interact with the dynein intermediate chain (Karki and Holzbaur, 1995; Vaughan and Vallee, 1995). This subunit is very flexible and likely can fold back on its self to dock against the dynactin filament as seen in a fraction of cryo-EM particles (Urnavicius et al., 2015). The 10 microtubule binding of p150Glued is required to recruit dynein to microtubules and enhance dynein processivity (Ayloo et al., 2014; Culver–Hanlon et al., 2006; King and

Schroer, 2000). In addition, mutations in p150Glued are associated with neurodegenerative disorders such as slowly progressing lower MND (motor neuron disease) and Parkinson-like Perry syndrome (Farrer et al., 2009; Puls et al., 2003;

Stockmann et al., 2013). In the brain, there is also an alternative splice form of p150, p135 which lacks the CAP-Gly domain (Tokito et al., 1996). Motor complexes with p135 seem to have reduced frequency of motility events in vitro (McKenney et al., 2014), but it is still unclear in vivo how p135 affects dynein functions compared to p150.

Dynactin is required for most of dynein’s functions. Recent cryo-EM studies have shown that dynactin reorients the dynein dimer for proper recruitment and positions the motor heads for proper motility along microtubules (Zhang et al., 2017). Furthermore, dynactin is vital to retrograde axonal transport of organelles (Haghnia et al., 2007;

Waterman-Storer et al., 1997). The pointed-end of dynactin has been suggested to play a role in facilitating cargo interaction (Yeh et al., 2012; Zhang et al., 2011), but more recent work has shown that dynactin interacts with dynein via dynein effectors which help link dynein to cargo (see next section for more details).

Figure 1.2 Dynein-Dynactin schematic

Schematics of cytoplasmic dynein and dynactin. Abbreviations: NDD=N-terminal dimerization domain, MTBD=microtubule binding domain, Robl=roadblock, LIC=Light intermediate chain, DIC=dynein intermediate chain.

IV. Dynein Effector Proteins

Adaptor and effectors proteins are required to link cargo to the dynein-dynactin motor complex (Fu and Holzbaur, 2014; Kardon and Vale, 2009). In addition to linking dynein to cargo, some of these proteins can modulate motility properties of dynein, either to enhance or inhibit movement, and some can act as motility switches by interacting with kinesins as well as dynein. This section will highlight the current insights into dynein effectors proteins.

The Bicaudal D (BICD) proteins were the first family of proteins described to link dynein motors to cargo. BICD was initially identified in Drosophila where mutations in

BICD cause abnormal development of a double abdomen or bicaudal (meaning ‘two- tailed’) phenotype (Mohler and Wieschaus, 1986). Further analysis of these flies showed that BICD was vital in mRNA transport and nuclei positioning, suggesting a link to microtubule-mediated transport (Bullock and Ish-Horowicz, 2001; Mach and Lehmann,

1997; Suter and Steward, 1991; Swan and Suter, 1996; Swan et al., 1999; Wharton and

Struhl, 1989). In mammals, there are two BICD orthologs, BICD1 and BICD2 as well as two related proteins, BICDR-1 and BICDR-2. The BICD proteins contain long stretches of coiled-coil domains which interact with dynein and dynactin, with the C-terminal region bending back on itself to produce an autoinhibited state (Liu et al., 2013; Terawaki et al.,

2015; Urnavicius et al., 2015; Wharton and Struhl, 1989). The C-terminal region of all

BICD proteins, except BICDR-2, can be released from this inhibition via cargo binding to the small GTPase Rab6 (Hoogenraad et al., 2001; Matanis et al., 2002; Schlager et al.,

2010; Short et al., 2002). The BICD family has been the most well-characterized family of dynein effectors and has been the starting point for many discoveries into the functions and mechanisms of dynein effectors.

Activation of Dynein

Recently, a set of coiled coil effector proteins including BICD2, Hook3, Spindly and NINL have been shown to enhance the dynein-dynactin interaction and induce superprocessive motility (McKenney et al., 2014; Redwine et al., 2017; Schlager et al.,

2014; Schroeder and Vale, 2016; Splinter et al., 2012). BICD2, the best characterized of these activators, has been shown to increase the affinity of dynein-dynactin interaction through coiled-coil contacts along the Arp1 filament that forms the core of dynactin

(Chowdhury et al., 2015; Urnavicius et al., 2015). BICD2 also interacts with the N- terminal tail of the dynein heavy chain (Chowdhury et al., 2015; Urnavicius et al., 2015) and the dynein light intermediate chain 1 (LIC1) (Lee et al., 2018; Schroeder et al.,

2014), leading to a stabilization of the dynein-dynactin-effector complex. Some dynein effectors including BICDR-1 can recruit two dynein dimers to a single dynactin, which further enhances the force and velocity of the motor complex (Grotjahn et al., 2018;

Schlager et al., 2014b; Urnavicius et al., 2018).

Spindly is another dynein activator which plays a role in mitosis by silencing a mitotic checkpoint after proper spindle assembly (Barisic and Geley, 2011; Griffis et al.,

2007; McKenney et al., 2014). Spindly recruits dynein to kinetochores which induces the movement of chromosomes to the poles (Chan et al., 2009; Gassmann et al., 2008;

Griffis et al., 2007). A recent study on Spindly also found new features of dynein activators, the CC1 box and the Spindly motif (Gama et al., 2017). Using sequence analysis and biochemistry, the CC1 box was found to be important for interaction with dynein while the Spindly motif was found to be important for interaction with dynactin. In addition, sequence alignments with other dynein adaptors showed the presence of these motifs in other proteins including the BICD family, HAP1, and TRAKs (Gama et al.,

2017). The CC1 box is a segment of coiled-coil with two conserved alanines. These two 14 alanines were of interest due to previous studies on BICD2, where mutations of these alanines to valines cause loss of interaction with dynein-dynactin in vitro (Schlager et al.,

2014b) and a loss-of-function phenotype in Drosophila melanogaster (Oh et al., 2000).

Similar alanine to valine mutations in Spindly also resulted in loss of dynein interaction.

The Spindly motif is located after the CC1 box and a stretch of coiled coil and has a sequence of L(F or A)XE for most of dynein effectors. This region of Spindly was found to interact with the pointed end of dynactin. The authors chose to mutate the phenylalanine to an alanine to show a loss of interaction with dynactin but unfortunately, the phenylalanine is not conserved in other dynein effectors and is actually an alanine in most cases (Gama et al., 2017). Further work is still needed to determine the importance of this region in other dynein effectors.

Rab11-FIP3 was also found to activate the motility of dynein. It is part of the

Rab11-FIP family of proteins which are mainly known to regulate the trafficking of recycling endosomes via a conserved Rab11 GTPase binding domain (reviewed in

Horgan and McCaffrey, 2009; Jing and Prekeris, 2009). Class I FIPs (Rab11-FIP1, 2, 5) have N-terminal phospholipid-binding C2-domains, while class II FIPs (Rab11-FIP3 and

-FIP4) possess N-terminal EF-hand domains in addition to longer coiled-coil regions near the C-terminus. Rab11-FIP3 and Rab11-FIP4 seem to have more specialized roles in endosomal trafficking then class I FIPs (Jing and Prekeris, 2009). Rab11-FIP3 plays an important role in the cell cycle dependent trafficking of recycling endosomes to the cleavage furrow but does not affect recycling of proteins in interphase (Horgan et al.,

2010; Inoue et al., 2008; Simon et al., 2008; Wilson et al., 2005). The other class II FIP,

Rab11-FIP4, is highly enriched in the brain but little is known of its function in endosomal trafficking (Jing and Prekeris, 2009). So far, only Rab11-FIP3 has been identified to

15 interact with dynein, despite the high similarities between FIP3 and FIP4 (Horgan et al.,

2010; McKenney et al., 2014).

Figure 1.3 Model of dynein activation

Dynein is found autoinhibited or in open states of flexible conformations. Following the addition of dynactin and a dynein activator, the motor domains of dynein reorganizes for proper microtubule attachment and display processive motility (Zhang et al., 2017).

Adaptors

The following proteins have not been shown to activate dynein motility in vitro like the activators described above but do share similar coiled coil structure and might be identified as activators in future work. These adaptors have mostly been identified through protein interactions, genetic manipulation, and live cell experiments.

One of these identified dynein adaptors is HAP1 (Huntingtin-associated protein

1). HAP1 interacts with huntingtin (htt), which is best known for its causative role in

Huntington’s disease where huntingtin is mutated with an expanded polyglutamine insertion. Huntingtin is a very large protein that has been shown to interact with a variety of proteins but notably interacts with the intermediate chain of dynein (Caviston et al.,

2007). HAP1 can interact with p150Glued (Engelender et al., 1997; Li et al., 1998) as well as kinesin heavy chain and light chain (McGuire et al., 2006; Twelvetrees et al., 2010).

Huntingtin and HAP1 have long been associated with intracellular transport (Block-

Galarza et al., 1997). Huntingtin has been linked to axonal transport of autophagosomes

(Wong and Holzbaur, 2014), synaptic vesicles (Gunawardena et al., 2003; Weiss and

Littleton, 2016), amyloid precursor protein(APP)-positive vesicles, and brain derived neurotrophic factor (BDNF)-positive vesicles (Colin et al., 2008; Her and Goldstein,

2008). HAP1 has been shown to be required for htt-mediated transport and it has been suggested that htt/HAP1 together act as a platform for both dynein and kinesin attachment to vesicles. Due to the large size of htt, it has been difficult to dissect the mechanism of htt and HAP1-mediated dynein motility in vitro assays.

Milton/TRAK proteins are a family of proteins suggested to act as motor adaptors for mitochondria. The Milton/TRAK family have a N-terminal coiled-coil region that has high homology to the HAP1 domain and the C-terminal region interacts with

Mitochondrial Rho GTPase (Miro). In Drosophila, Milton in complex with Miro interacts with kinesin-1 to deliver mitochondria to neuronal synapses (Glater et al., 2006; Stowers et al., 2002). The mammalian homologs of Milton, TRAK1 and TRAK2, have been linked to dynein and kinesin motility and are required for mitochondria distribution in a variety of cell types including neurons (reviewed in Melkov and Abdu, 2018). TRAK1 binds dynein- dynactin and kinesin-1, while TRAK2 predominately interacts with dynein-dynactin (van

Spronsen et al., 2013). Interestingly in neurons, TRAK1 is mainly localized in the axon, while TRAK2 is localized to the dendrites (Loss and Stephenson, 2015; van Spronsen et al., 2013), which could reflect the dependence of each compartment for specialized motors.

Another adaptor, RILP (Rab7-interacting lysosomal protein) has been suggested to link dynein to Rab7 vesicles, including late endosomes and lysosomes. RILP was initially found to interact with the small GTPase Rab7 through yeast two-hybrid screens 17

(Cantalupo et al., 2001). RILP was then suggested to link Rab7 vesicles with dynein through co-localization experiments (Jordens et al., 2001). Later, biochemical studies showed a stepwise process of dynein recruitment by RILP, where RILP and oxysterol- binding protein–related protein 1L (ORP1L) form a complex with Rab7 and then RILP can interact with p150Glued of dynactin, which in turn recruits dynein to the vesicle

(Johansson et al., 2007). This stepwise recruitment suggests dynein association with vesicles can be regulated by cholesterol levels, which are sensed by ORP1L. Consistent with this mechanism, cholesterol-rich lysosomes are transported to the nucleus, but low cholesterol vesicles remain at the cell periphery due to conformational changes in

ORPL1, which leads to dissociation of p150Glued (van der Kant et al., 2013; Rocha et al.,

2009). In addition, RILP has been shown to self-interact, likely as a homodimer, similar to other dynein effectors like BICD2 (Colucci et al., 2005; Wu et al., 2005).

JIP (c-Jun N-terminal kinase (JNK)-interacting proteins) proteins have also been identified as motor adaptors. There are four mammalian JIPs, JIP1-4, which are highly expressed in the brain (Dickens et al., 1997; Kelkar et al., 2000, 2005; Yasuda et al.,

1999). While each JIP protein contains a JNK-binding domain near the N-terminus, JIP1 and JIP2 are structurally distinct from JIP3 and JIP4. JIP1/2 each contain a Src homology-3 domain (SH3) and a phosphotryosine binding domain (PTB) near the C- terminus (Dickens et al., 1997; Yasuda et al., 1999), while JIP3/4 each contain N- terminal coiled coil regions, a leucine zipper domain, and a C-terminal transmembrane region (Kelkar et al., 2000, 2005). JIP proteins interact with multiple kinases of the JNK pathway and p38 MAPK pathway, as positive or negative regulators (Whitmarsh, 2006).

These signaling pathways are involved in a variety of cellular processes including growth, differentiation, and apoptosis. In addition to signaling factors, JIPs have been shown to interact with microtubule motors. All of the JIPs have been found to interact 18 with kinesin-1 (Bowman et al., 2000; Montagnac et al., 2009; Verhey et al., 2001). JIP1,

JIP3, and JIP4 have also been implicated in minus-end motility via dynein-dynactin interaction. JIP1 has been found to coordinate dynein and kinesin motility for APP- positive vesicles, with phosphorylation used to switch from kinesin to dynein motility (Fu and Holzbaur, 2013). JIP4 was shown to transport recycling endosomes during cytokinesis via its interaction with kinesin-1 and dynactin, with ARF6 binding as the regulatory switch for JIP4 interaction with motors (Montagnac et al., 2009). In

Drosophila, JIP3 (aka Sunday Driver) was found to associate with dynein-dynactin during the transport of axonal injury signals (Cavalli et al., 2005). Further work suggested that JIP3 interacts with endosomes for retrograde transport and a small anterograde vesicle population, potentially for axonal growth (Abe et al., 2009; Watt et al., 2015). A recent study showed that JIP3 knockout mouse neurons have lysosome accumulation in axons and impaired maturation, which was previously seen in zebrafish and C. elegans (Drerup and Nechiporuk, 2013; Edwards et al., 2013; Gowrishankar et al., 2017).

Regulatory mechanisms

There are several ways that dynein effector proteins can be regulated for proper spatiotemporal intracellular transport. Some of these adaptor proteins including JIP1 and huntingtin have phosphorylation sites that regulate interaction with molecular motors

(Colin et al., 2008; Fu and Holzbaur, 2013). Other interacting proteins can also act as regulators of dynein effector proteins. For example, Miro is a Ca+2 sensor which alters interaction with Milton and kinesin-1 upon Ca+2 binding, reducing mitochondria motility

(MacAskill et al., 2009; Wang and Schwarz, 2009). Some dynein effectors have also been suggested to be in an autoinhibitory state when not bound to cargo, blocking motor 19 interaction unless properly bound to cargo. For example, the last coiled coil region of

BICD2 folds back on itself but binding its cargo protein Rab6 releases this autoinhibition, unmasking its dynein-dynactin interface (Terawaki et al., 2015). Differential expression of these effectors in tissues and during development is another way to control transport by dynein. For instance, BICD1 has been shown to be highly expressed early in development and decreases during neurite growth (Schlager et al., 2010). Other regulatory mechanisms remain to be elucidated.

Figure 1.4 Domain structure of dynein effectors

Figure legend at the top right. Gray lines indicate identified dynein-dynactin interaction region. Brown lines indicate identified kinesin interaction region. RH=RILP-homology

Other regulators of dynein

Another important regulator of dynein is Lis1. LIS1 was first described as the mutated gene in type 1 lissencephaly, a neurodevelopmental disease (Reiner et al.,

1993). Deletions or duplications of LIS1 cause serious brain development defects. Lis1 plays a role in neuronal migration and proliferation (reviewed in Reiner and Sapir, 2013).

Lis1 interacts with Nudel and NudE, which also have roles in neurodevelopment

(reviewed in Bradshaw et al., 2013). Lis1 and Nudel/NudE were found to interact with dynein during screens for nuclear migration defects in filamentous fungi (Efimov and

Morris, 2000; Minke et al., 1999; Xiang et al., 1995), later confirmed to be conserved in other systems, including Drosophila (Liu et al., 1999, 2000) and mammals (Faulkner et al., 2000; Sasaki et al., 2000; Smith et al., 2000). In polarized cells like neurons, Lis1 has been linked to the motility of several different cargos including mitochondria, endosomes, peroxisomes, and lysosomes, and suggested to be important for motor loading at microtubule plus-end (Egan et al., 2012; Lenz et al., 2006; Moughamian et al.,

2013; Shao et al., 2013; Yi et al., 2011; Zhang et al., 2010). Lis1 is also involved in positioning of centrosomes, nuclei (Gambello et al., 2003; Tsai et al., 2007; Xiang et al.,

1995), and spindle poles (Moon et al., 2014; Yingling et al., 2008).

Lis1 has a N-terminal dimerization domain, a coiled-coil region, a disordered loop, and a β-propeller domain with seven WD repeats (Kim et al., 2004; Tarricone et al.,

2004). The β-propeller domain of Lis1 interacts with the dynein motor domain, at AAA3 and AAA4 (Huang et al., 2012; Toropova et al., 2014). Biochemical experiments suggest that Lis1 increases dynein’s affinity for microtubules and slows down dynein’s velocity in motility assays in vitro (Huang et al., 2012; McKenney et al., 2010; Toropova et al., 2014;

Yamada et al., 2008). Lis1 does not prevent ATP hydrolysis but has been suggested to uncouple ATP hydrolysis from MT binding/release (Huang et al., 2012). Recent work has 22 challenged the idea that Lis1 plays an inhibitory role in dynein motility. For yeast dynein,

Lis1 is now proposed to induce weak or tight microtubule-binding of dynein, depending on the nucleotide state of AAA3 and the number of Lis1 β-propeller domains bound to the motor domain (DeSantis et al., 2017). In recent studies on mammalian dynein with dynactin and BICD2, Lis1 was found to actually increase the frequency and velocity of dynein motility, in a concentration-dependent manner (Baumbach et al., 2017; Gutierrez et al., 2017). Future structural work is needed to better understand the complex of Lis1 with dynein-dynactin-BICD2.

Nudel/NudE are dimeric, coiled-coil proteins that interact with DIC and LC8 via their N-termini and Lis1 with their C-termini (Stehman et al., 2007; Wang and Zheng,

2011; Zyłkiewicz et al., 2011). Nudel/NudE has been suggested to tether Lis1 to dynein and be a regulatory module due to their phosphorylation by Cdk5 (Hebbar et al., 2008;

Klinman and Holzbaur, 2015; Pandey and Smith, 2011). Lis1 in complex with NudE is suggested to induce a high load-bearing state in dynein. Optical trapping experiments showed that Lis1 and NudE allow dynein to have a prolonged force-producing state

(McKenney et al., 2010). Lis1 and NudE/Nudel also enhanced force production of dynein on lipid droplets in cells in a p150Glued independent manner (Reddy et al., 2016).

V. Hook Proteins

With the growing evidence for dynein adaptors, the goal of my thesis work was to uncover new dynein adaptors and understand their role in the regulation of dynein motility. My thesis work focuses on the Hook protein family, due to several studies in other model systems, like Drosophila and filamentous fungi, which suggested roles for

Hook proteins in intracellular transport.

Hook proteins were first investigated in a screen for genes involved in cell- specific ligand endocytosis in Drosophila (Krämer and Phistry, 1996). Hook mutant flies have abnormal bristle morphology which makes them look like a “hook”, in addition to eye degeneration (Krämer and Phistry, 1996; Mohr, 1927). Genetic analysis showed that mutations in the hook gene inhibit endocytosis, as seen by reduced accumulation of transmembrane and soluble ligands into multivesicular bodies, and increased the accumulation of multilamellar late endosomes in the photoreceptor cells of Drosophila eyes (Krämer and Phistry, 1996, 1999; Sunio et al., 1999). The Drosophila Hook protein was also localized to vesicular structures, potentially late endosomes marked by Rab7

(Krämer and Phistry, 1996; Szatmári et al., 2014).

In general, Hook proteins are characterized by three conserved regions: a globular N-terminal putative microtubule binding domain, a central coiled-coil domain, and a divergent, predicted unstructured C-terminal domain thought to mediate cargo binding (Walenta et al., 2001). The N-terminal region of Hook proteins was originally described to interact with microtubules based on homology to the well characterized microtubule binding domains, calponin-homology domains, and initial microtubule pelleting assays with cell lysates (Walenta et al., 2001). The central coiled-coil region was suggested to induce homodimerization by immunoprecipitation and sedimentation assays (Sevrioukov et al., 1999; Walenta et al., 2001; Xu et al., 2008). The C-terminal 24 region is the most divergent region among species and isoforms but has been suggested to link Hook proteins to their intended cargo based on the use of truncated constructs (Bielska et al., 2014; Walenta et al., 2001; Xu et al., 2008; Zhang et al.,

2014).

Hook proteins (HookA or Hok1), were first characterized as potential dynein adaptors in filamentous fungi (Bielska et al., 2014; Zhang et al., 2014). In Aspergillus nidulans, HookA was described as an adaptor on early endosomes regulating dynein, while Hok1 in Ustilago maydis was shown to coordinate dynein and kinesin-3 motors during early endosome transport (Bielska et al., 2014; Zhang et al., 2014). In both fungal

Hook proteins, the N-terminus is required for interaction with dynein and the C-terminus allows for attachment to cargo via interaction with Fused Toes (FTS) and FTS and Hook-

Interacting Protein (FHIP) (Bielska et al., 2014; Yao et al., 2014; Zhang et al., 2014).

These interactions were determined by immunoprecipitation experiments from cell lysates, so direct protein-protein interactions remained to be determined by future studies.

In mammalian cells, three highly conserved Hook proteins are expressed: Hook1,

Hook2, and Hook3. Each Hook protein has been linked to different cellular functions.

Hook1 has been implicated in spermiogenesis as mutations in Hook1 cause abnormal spermatozoon head shape (Mendoza-Lujambio et al., 2002). Hook2 has been linked to centrosomal function and homeostasis (Guthrie et al., 2009; Moynihan et al., 2009;

Szebenyi et al., 2007). Hook3 was originally described as a Golgi-associated protein due to its localization at the Golgi in cell lines and the disruption of Golgi structure when

Hook3 is overexpressed (Walenta et al., 2001). In addition to the aforementioned work,

Hook1 and Hook3 have been implicated in a variety of endosomal trafficking pathways, although there is still no clear consensus on the specific roles of each isoform (Luiro et 25 al., 2004; Maldonado-Báez et al., 2013; Xu et al., 2008). In HeLa cells, Hook proteins were found to interact with members of the HOPS complex and to be important for timely trafficking of EGF through endosomal compartments marked by EEA1, CD63 and

LAMP1, but this study simultaneously knocked down all three Hook isoforms, making it difficult to determine their individual roles (Xu et al., 2008). A recent study in hippocampal neurons suggested that Hook1 and Hook3 are involved in Rab5 retrograde motility in the axon (Guo et al., 2016). Hook1 and Hook3 have been suggested to attach to cargo through C-terminal interactions with Fused Toes (FTS) and FTS-Hook

Interacting (FHIP) proteins, similar to fungal studies (Guo et al., 2016; Xu et al., 2008;

Yao et al., 2014).

Other studies have looked at individual Hook proteins. Using HeLa cells, Hook1 was found to interact with CD147 to facilitate sorting into Rab22-positive recycling tubules (Maldonado-Báez et al., 2013). In COS-1 cells, Hook1 was found to interact with

Rab7, Rab9, and Rab11 using immunoprecipitation (Luiro et al., 2004). The variety of results seen in these studies is likely due to the fluid nature of endosomal pathways and differential cellular demands on these pathways. Studies on Hook3 have implicated that it interactions with a variety of proteins including scavenger receptor A, Pericentriolar

Material 1 (PCM1), IIGP, and Salmonella SpiC (Ge et al., 2010; Kaiser et al., 2004; Sano et al., 2007; Shotland et al., 2003). It remains to be seen if these interactions are reproducible in other systems and studies.

Throughout my thesis, I studied the role of mammalian Hook proteins as dynein adaptors using biochemistry, in vitro motility assays, and live cell microscopy. In the first half of my thesis, I worked to determine how Hook proteins effect dynein motility and which domains are important for motor interaction. In addition, I collaborated with the

Dominguez lab to better understand which dynein components interact with Hook 26 proteins and effect processivity. In the second half of my thesis, I studied the role of

Hook proteins in axonal transport in primary neurons.

CHAPTER 2: Hook Adaptors Induce Unidirectional Processive Motility by

Enhancing the Dynein-Dynactin Interaction

This chapter is adapted from:

Olenick, M. A., Tokito, M., Boczkowska, M., Dominguez, R., Holzbaur, E.L.F. (2016)

Hook adaptors induce unidirectional processive motility by enhancing the dynein- dynactin interaction. Journal of Biological Chemistry 291, 18239-18251.

We gratefully acknowledge Karen Wallace for technical assistance, Michael Woody and

Jeffery Nirschl for assistance with custom image analysis programs, and Chanat

Aonbangkhen for dimerizer reagent.

I. Summary

Cytoplasmic dynein drives the majority of minus-end directed vesicular and organelle motility in the cell. Yet, it remains unclear how dynein is spatially and temporally regulated given the variety of cargo that must be properly localized to maintain cellular function. Recent work has suggested that adaptor proteins provide a mechanism for cargo-specific regulation of motors. Of particular interest, studies in fungal systems have implicated Hook proteins in the regulation of microtubule motors.

Here, we investigate the role of mammalian Hook proteins, Hook1 and Hook3, as potential motor adaptors. We used optogenetic approaches to specifically recruit Hook proteins to organelles and observed rapid transport of peroxisomes to the perinuclear region of the cell. This rapid and efficient translocation of peroxisomes to microtubule minus ends indicates that mammalian Hook proteins activate dynein rather than kinesin motors. Biochemical studies indicate that Hook proteins interact with both dynein and dynactin, stabilizing the formation of a supramolecular complex. Complex formation requires the N-terminal domain of Hook proteins, which resembles the calponin- homology domain of EB proteins yet cannot bind directly to microtubules. Single molecule motility assays using TIRF microscopy indicate that both Hook1 and Hook3 effectively activate cytoplasmic dynein, inducing longer run lengths and higher velocities than the previously characterized dynein activator BICD2. Together, these results suggest that dynein adaptors can differentially regulate dynein to allow for organelle- specific tuning of the motor for precise intracellular trafficking.

II. Introduction

Microtubules provide a polarized highway to facilitate transport of organelles and vesicles throughout the cell. The minus ends of microtubules are usually nucleated near the cell center with the plus ends oriented outwards, toward the cell periphery. This polarity ensures that microtubule motors drive motility in a specific direction; kinesin motors generally drive plus-end motility while minus-end traffic is primarily driven by cytoplasmic dynein. Regulation of these opposing motors is vital for cell survival, particularly in specialized cells like neurons that require efficient transport over long distances (Maday et al., 2014). However, it remains unclear how microtubule motors are spatially and temporally regulated to control the intracellular trafficking of specific cargo.

As a single major form of cytoplasmic dynein drives the transport of a wide array of cargos, including endosomes, RNA granules, and mitochondria (van Niekerk et al.,

2007; van Spronsen et al., 2013; Xiang et al., 2015), it is likely that the transport properties of dynein are modulated by the binding of cargo-specific adaptor molecules.

A number of dynein regulatory and adaptor proteins have been identified to date, including dynactin, Lis1, BICD2, and, more recently, Hook proteins. The first major regulator to be identified was dynactin, a large multi-subunit protein complex required for most functions of dynein within the cell. Dynactin forms a co-complex with dynein

(Chowdhury et al., 2015; Karki and Holzbaur, 1995; Urnavicius et al., 2015; Vaughan and Vallee, 1995) that enhances the initial recruitment of dynein to the microtubule

(Ayloo et al., 2014; Moughamian and Holzbaur, 2012) and mediates association of dynein with some intracellular cargos (Holleran et al., 2001; Muresan et al., 2001; Yeh et al., 2012; Zhang et al., 2011). A second major dynein regulator, Lis1, binds to the dynein motor domain and blocks the required linker swing in the mechanochemical cycle for dynein; thus Lis1 binding induces a non-motile state of dynein that binds tightly to the 30 microtubule (Toropova et al., 2014). In contrast to the inhibitory effect of Lis1 on dynein motility, the dynein adaptor BICD2 has been shown to induce superprocessive motility of dynein, potentially through enhanced stability of the dynein-dynactin complex

(McKenney et al., 2014; Schlager et al., 2014b). Yet another mechanism that adaptors can use to regulate transport is coordination of different motors on the same cargo. For instance, JIP1 acts as a switch between dynein- and kinesin-1-mediated transport depending on its phosphorylation state (Fu and Holzbaur, 2013). Given the wide variety of cargo that must be properly localized within eukaryotic cells, it is likely that many additional adaptors and their underlying regulatory mechanisms remain to be identified and characterized.

Here we focus on another family of potential dynein adaptors, Hook proteins.

Genetic screens in fungal model systems have provided evidence that Hook proteins are required for early endosome trafficking. In general, Hook proteins are characterized by three conserved regions: a globular N-terminal putative microtubule binding domain, a central coiled coil domain, and a variable and predicted unstructured C-terminal domain thought to mediate cargo binding (Walenta et al., 2001). In Aspergillus nidulans, HookA was described as an adaptor on early endosomes regulating dynein, while Hok1 in

Ustilago maydis was shown to coordinate dynein and kinesin-3 motors during early endosome transport (Bielska et al., 2014; Zhang et al., 2014). In both fungal Hook proteins, the C-terminus attaches to cargo through interaction with the proteins, Fused

Toes (FTS) and FTS and Hook-Interacting Protein (FHIP) (Bielska et al., 2014; Yao et al., 2014; Zhang et al., 2014).

These studies led us to ask if such functions of Hook proteins were conserved in mammalian systems. There are three Hook isoforms expressed in humans: Hook1,

Hook2, and Hook3. Each isoform has been associated with a different cargo. Hook3 31 localizes to the Golgi (Walenta et al., 2001), Hook2 is recruited to centrosomes

(Szebenyi et al., 2007), and Hook1 is implicated in endosomal transport (Luiro et al.,

2004; Maldonado-Báez et al., 2013; Xu et al., 2008). To explore adaptors with roles in cargo transport, we focused on Hook1 and Hook3 in our studies.

We used complementary optogenetic and single molecule approaches to establish mammalian Hook proteins as motor adaptors enhancing unidirectional minus end-directed motility driven by dynein. We show that both Hook1 and Hook3 enhance the formation of a dynein-dynactin complex. The formation of this complex requires the

N-terminal globular domain of Hook proteins. Contrary to previous suggestions, this domain does not bind to microtubules directly. In single molecule assays, we find that both Hook1 and Hook3 induce highly processive dynein motility, resulting in both longer run lengths and faster velocities than the previously characterized dynein activator,

BICD2. Together these results support a model in which organelle-specific adaptors differentially regulate dynein motor function within the cell.

III. Results

Differential regulation of dynein-mediated cargo transport by Hook proteins

To assess the role of different adaptors on cargo transport within the cell, we used a light-induced dimerization system to observe changes in cargo motility after recruitment of different adaptors and regulators. In this system, we use the dimerizer cTMP-Htag, a small molecule made of a HaloTag (Htag) ligand linked to a photocaged trimethoprim (TMP). This molecule heterodimerizes HaloTag proteins (Halo) and

Escherichia coli DHFR (eDHFR)-tagged proteins. In our experiments, dimerization between a Halo-tagged cargo and a DHFR-tagged adaptor/motor is induced using

405nm light to cleave photocaged cTMP-Htag (Ballister et al., 2015) (Fig. 2.1A). We used peroxisomes as a model organelle since they are not very motile under endogenous conditions and are uniformly distributed throughout the cell (Smith and

Aitchison, 2013), making them ideal to observe changes in motility.

In live cell experiments in HeLa cells, recruitment of either Hook1 or Hook3 to peroxisomes through light-induced dimerization resulted in a pronounced redistribution of peroxisomes toward the perinuclear region (Fig. 2.1B,C). The organelle redistribution induced by either of the Hook proteins was similar to that observed upon recruitment of the known dynein activator BICD2, suggesting Hook proteins also act as dynein adaptors (Fig. 2.1B,C). In contrast, recruitment of the dynein-binding protein p150Glued was not sufficient to induce robust global redistribution of peroxisomes in this assay. We used K560 (a constitutively active construct of kinesin-1), as a control for kinesin motility and observed robust motility to the periphery of the cells (Fig. 2.1B,C). For additional controls, we also imaged cells expressing these constructs in the absence of dimerizer

33 or in the absence of photoactivation and saw no effects on peroxisome location or motility (data not shown).

Figure 2.1 Hook proteins redistribute peroxisomes to perinuclear region in an optogenetic assay

A) Schematic of inducible dimerization assay and corresponding constructs. B) Using a photoactivatable dimerization system (cTMP-Htag dimerizer) (Ballister et al., 2014), motors/adaptors (-mCh-DHFR tagged) were recruited to peroxisomes (PEX3-Halo-GFP

34 labeled) by 405 nm light and the resulting motility was observed by live cell confocal microscopy. Scale bars=10 μm. Arrows indicate peroxisome clustering after recruitment.

C) Overlay of pre- and post-dimerization images of peroxisomes.

To quantify the redistribution phenotype of each adaptor, we measured the distance of each peroxisome from the microtubule organization center (MTOC) in cells that were fixed 45 minutes after addition of an uncaged TMP-Htag dimerizer and then stained with a gamma-tubulin antibody to visualize the MTOC. Analysis of cells from 3 independent repeats showed that Hook1, Hook3, and BICD2 each induced a significant concentration of peroxisomes near the MTOC in contrast to either p150Glued or K560 (Fig.

2.2). Direct comparison of the distributions shows that Hook1 induced the tightest clustering of peroxisomes near the MTOC, while recruitment of either Hook3 or BICD2 induced similar distributions (Fig. 2.2). Of note, both Hook3 and BICD2 have been linked to Golgi transport (Hoogenraad et al., 2001; Walenta et al., 2001), while Hook1 has been linked to endosomal transport (Maldonado-Báez et al., 2013). While p150Glued was efficiently recruited to peroxisomes in this assay (Fig. 2.2), this recruitment was not sufficient to induce marked peroxisome motility or redistribution, consistent with the idea that dynactin alone is insufficient to induce superprocessive motility. Together, these observations suggest that like BICD2, both Hook1 and Hook3 can activate dynein motility, and that differences among these activators may tune dynein activity to regulate cargo-specific transport.

Figure 2.2 Hook proteins differentially redistribute peroxisomes to the MTOC

A) Dimerization assay in fixed cells stained with gamma-tubulin antibody. Images are maximum projections of confocal z-stacks. Scale bars=10μm. White arrows point to gamma-tubulin stained MTOC and yellow arrows point to peroxisome clusters. Cell outlines were determined from corresponding X-mCH-DHFR images (not shown). B)

Distribution of peroxisomes from MTOC measured in fixed time point dimerization assay 36

(analyzed using Cell Profiler (Carpenter et al., 2006)). The endosomal-linked adaptor

Hook1 tightly clusters peroxisomes to the MTOC compared to Hook3 and BICD2. (Cells analyzed per condition: K560: n=36, p150Glued: n=19, BICD2: n=26, Hk1: n=23, Hk3: n=32) Error bar are S.E.M based on number of cells.

Mammalian Hook proteins interact with dynein-dynactin

To characterize the interactions of mammalian Hook proteins with dynein and dynactin, we performed immunoprecipitation experiments using endogenous and expressed Hook proteins. Using mouse brain lysates, we immunoprecipitated endogenous dynein and dynactin with monoclonal antibodies to the dynein intermediate chain (DIC) and the p150Glued subunit of dynactin, respectively. Co-immunoprecipitation of endogenous Hook1 was seen with the anti-p150Glued antibody but not with the anti-DIC antibody (Fig. 2.3A). This DIC antibody is known to disrupt the interaction of dynein with dynactin (Karki and Holzbaur, 1995), as confirmed by a decreased amount of p150Glued in the DIC IP lane (Fig. 2.3A). These results suggest that Hook proteins either interact with dynactin directly or with the full dynein-dynactin complex, rather than interacting solely with dynein.

Next, we performed IP experiments using Halo-tagged Hook proteins expressed in Cos7 cells. Again, using the p150Glued antibody, we observed co-immunoprecipitation of expressed full length human Hook1 and Hook3 with the dynein-dynactin complex.

With the expression of Hook proteins, we also observed an increase in the co- precipitation of dynein by the anti-p150Glued antibody, as compared to a control experiment in which only Halo-tag was expressed (Fig. 2.3B). We quantified the ratio of

DIC:p150Glued in the immunoprecipitates from each condition, and observed that expression of Hook1, Hook3, or BICD2 each induced enhanced association of dynein 37 with dynactin compared to control IPs (Fig. 2.3C). This observation is consistent with previous studies suggesting that BICD2 enhances dynein-dynactin complex stability

(McKenney et al., 2014; Splinter et al., 2012). Here we find that both Hook proteins were also able to enhance the stability of the dynein-dynactin complex.

Figure 2.3 Mammalian Hook proteins interact with dynein-dynactin complex

A) Western blot showing immunoprecipitation (IP) of endogenous p150Glued (subunit of dynactin) and DIC (Dynein Intermediate Chain) from mouse brain lysates, with anti-myc used as a mouse IgG (MS IgG) control. IP of p150Glued shows interaction with Hook1, while disruption of dynein-dynactin complex in IP with anti-DIC shows loss of this interaction. n=3. B) Western blots showing IP of endogenous p150Glued from Cos7 cells expressing Halo-Hook1, -Hook3, -BICD2 (1-572). HaloTag expressed as a negative control. C) Graph of DIC to p150Glued IP ratio from experiments in B, n=4. The ratio of

DIC to p150Glued IP ratio for control condition (HaloTag only) was normalized to 1 and all other conditions shown as a fold change from the control. Error bars are S.E.M.

The N-terminal domain of Hook proteins does not bind microtubules but is important for interaction with the dynein-dynactin complex

Since previous work suggested that the N-terminal globular domain of Hook proteins binds microtubules (Walenta et al., 2001), we asked whether this domain was necessary for the motor adaptor function of Hook proteins. First, we assessed the ability of Hook proteins to bind microtubules in cell lysates. Using a HA-tagged Hook1 construct expressed in Cos7 cells, we observed pelleting of HA-Hook1 with taxol-stabilized microtubules (Fig. 2.4A,B). However, since this assay utilized cell lysates, the apparent interaction of Hook1 with microtubules could be indirect. To test if the N-terminal region of Hook proteins can bind microtubules directly, we purified recombinant proteins spanning the N-terminal domain of Hook1 for use in microtubule pelleting assays. We tested binding with taxol- or GMPCPP-stabilized microtubules, since they mimic different nucleotide states of microtubules and induce different tubulin conformations, which can affect binding of microtubule associated proteins (Alushin et al., 2014; Yajima et al.,

2012). Purified Hook1 (1-443aa) showed no observable pelleting with either taxol- or

GMPCPP-stabilized microtubules, suggesting this protein has little or no affinity for microtubules (Fig. 2.4C). Next, we tested a recombinant Hook3 N-terminal protein fused to a coiled-coil GCN4 leucine zipper to induce efficient dimerization. Again, we observed no co-pelleting of the purified protein with microtubules, in contrast to a construct of p150Glued that binds to microtubules directly through its N-terminal CAP-Gly domain

(Waterman-Storer et al., 1995) (Fig. 2.4C).

Figure 2.4 Hook proteins bind microtubules indirectly

Microtubule (MT) binding assays were performed using (A) cell lysates from HA-Hook1 transfected Cos7 cells and (C) recombinant purified Hook1 dimer (1-443aa) and Hook3

(1-210aa-GCN4) (1 μM). MT binding assays were performed by mixing equal amounts of protein to increasing amounts of taxol- or GMPCPP- stabilized MTs. Supernatant (S) and pellets (P) were analyzed by SDS-PAGE gels and western blotting, with HA-tag and p150Glued antibodies as noted. Gels in C are Coomassie-stained SDS-PAGE gels. Hook1 from cell lysates co-sediments with MTs, but purified Hook1 and Hook3 constructs do not pellet with MTs, suggesting indirect binding. Endogenous and purified p150Glued (1-

210aa-Htg construct) were used as controls. GMPCPP-stabilized MT binding assay gels not shown. B) Graph of binding assay quantification in A. Error bars are S.E.M. (Cell lysates expt. n=5, purified expt. n=2-3)

Additionally, we used sequence analysis and structure prediction to compare the calponin-homology domain at the N-terminus of Hook isoforms to that of the well- characterized microtubule-binding proteins, EB1 and EB3. Although the overall calponin- homology fold is well-conserved in Hook proteins, the specific regions implicated in microtubule binding according to an EM reconstruction of EB3 on microtubules (Zhang et al., 2015) are very different in Hook proteins as compared to EBs (Fig. 2.5A,B).

Additionally, EB1 residues implicated in microtubule association by mutagenesis studies, including H18, K66, and L67 (Slep and Vale, 2007), are not conserved in Hook isoforms.

In our alignment of Hook and EB sequences based off secondary structure conservation, the corresponding residues in Hook proteins are the same or very similar to the mutations, H18E, K66E, and L67D, that cause a loss of microtubule binding in EB1, further suggesting that Hook proteins do not bind directly to microtubules. However, since we observed co-pelleting of Hook proteins expressed in cell lysates with microtubules, there is likely an indirect interaction mediated by the binding of Hook proteins to dynein-dynactin.

Figure 2.5 Hook proteins lack conserved regions for MT binding

A) Sequence alignment based on secondary structure for Hook1 and EB3. Coloring based off BLOSUM62 score. Magenta boxes indicate MT interaction regions in EB3

(residues within 6Å of tubulin surface in PDB: 3JAK structure (Zhang et al., 2015)).

Arrows indicate residues that ablate microtubule association in EB1 if mutated and are not conserved in Hook proteins (Slep and Vale, 2007). B) Comparison of N-terminal mouse Hook1 (PDB: 1WIX) and EB3 (PDB: 3JAK (Zhang et al., 2015)) structures with predicted microtubule interactions sites highlighted in magenta. Numbers correspond to boxed regions in the alignment (A).

Structural studies of the dynein-dynactin-BICD2 complex indicate that a key aspect in the interaction is the extended coiled-coil domain of BICD2 that threads through a groove along the Arp1 filament (Chowdhury et al., 2015; Urnavicius et al.,

2015). While there are similar extended coiled-coil domains in Hook proteins, the high sequence conservation of the N-terminal calponin-homology domain among Hook isoforms made us question whether this region was also important for the interaction with dynein-dynactin. Using mouse brain lysates, we preformed pull-down experiments with several Hook1 constructs, including the full-length protein, a construct lacking the N- terminus (171-728aa) and a construct truncated at the C-terminus (1-554aa) (Fig. 2.6A).

Cos7 cells expressing Halo-tagged Hook1 constructs were lysed and bound to Halo-link resin. Mouse brain lysates were then mixed with the resins as an abundant source of dynein-dynactin. The resulting pull-downs with full-length and C-terminally-truncated

Hook (1-554aa) showed interaction with dynein and dynactin components, while the construct lacking the N-terminal region showed little or no interaction with dynein- dynactin (Fig. 2.6B,C), consistent with work on other Hook homologs (Malone et al.,

2003; Zhang et al., 2014). This result suggests that the N-terminal region of Hook proteins is necessary for the interaction with dynein-dynactin, potentially providing further contacts in addition to the coiled-coil region to modulate motor activity.

Figure 2.6 Pull-Down (PD) of Halo-Hook1 with dynein-dynactin complex requires

N-terminal region

A) Diagram of conserved domains and predicted coiled-coil regions in Hook1.

MTBD=putative microtubule binding domain, CBD=cargo binding domain. B) Western blot showing PD of Halo-Hook1 constructs with endogenous dynein-dynactin from mouse brain lysates (DHC=Dynein Heavy Chain, DIC=Dynein Intermediate Chain). PD of Hook1 full length (FL) and Hook1 (1-554aa) constructs show interaction with dynein- dynactin, while PD of Hook1 (171-728aa (171-E)) shows loss of interaction with dynein-

44 dynactin complex. NT=non-transfected control. n=3. C) Graphs of DIC or p150Glued to

Hook1 (Hk) ratio from experiments in B, n=3. Error bars are S.E.M.

Hook proteins induce highly processive runs with enhanced velocities

To characterize the functional effects of Hook adaptors on dynein, we utilized an in vitro single molecule approach, using Total Internal Reflection Fluorescence (TIRF) microscopy of cell extracts to characterize dynein-dynactin motility (Ayloo et al., 2014).

We expressed Halo-tagged Hook constructs in HeLa cells and labeled cells with TMR- labeled HaloTag ligand prior to generation of cell lysates. We immobilized taxol- stabilized microtubules to the coverslips of flow chambers using antibodies against β- tubulin. Cell lysates were diluted into motility buffer containing 10 mM Mg-ATP, taxol,

BSA, casein and an oxygen scavenging system and then flowed into the chamber to be imaged.

Using polarity marked microtubules, Hook proteins were seen moving in a unidirectional manner to the minus ends of microtubules, as expected for dynein- mediated motility (Fig. 2.7A). The motility of Hook proteins could be inhibited using siRNA against the dynein heavy chain to knock-down endogenous dynein, confirming that the motility seen is dynein-mediated (Fig. 2.7B,C). Particles were tracked with Fiji

TrackMate (Schindelin et al., 2012) to measure run lengths and velocity. The resulting data were analyzed with a custom Maximum Likelihood Estimation (MLE) modeling program in Matlab (Woody et al., 2016). Velocities were fit to single or double Gaussians as noted and run lengths were fit to single exponential decay curves (Fig. 2.7D,E).

Figure 2.7 Hook proteins display high velocities and long run lengths

A) Example time series of particles moving to the minus end of microtubule (polarity marked for Hk1 and Hk3, Plus end in green). Scale Bar=2μm. B) Maximum projections 46 of Halo-Hook1 (FL) expressed in cells with Mock or DHC siRNA conditions and imaged in TIRF assay. Scale Bar=5μm. C) Western blot of Mock and DHC siRNA knockdown

(KD) lysates used for TIRF assays. D) Track displacement and velocity distributions for particles tracked with ImageJ plugin, TrackMate. Data was fitted with a custom

Maximum Likelihood Estimation (MLE) program (Woody et al., 2016) and plotted as

Probability Density Functions (PDFs) with 95% CI bootstrapping. (BICD2 n=242, Hk1 n=90, Hk3 n=84). E) Table of motility parameters based on fits from data in D. F)

Percent of events with mean velocity greater than 1 μm/sec. G) Per track standard deviation of instantaneous velocity. Data plotted as boxplot with Tukey whiskers.

Overall, full-length Hook1 and Hook3 proteins induced motility with higher velocities and longer run lengths compared to the active BICD2 construct 1-572 (Fig.

2.7D). More than 40% of the Hook-dependent motility events exhibited mean velocities greater than 1 micron/sec (Fig. 2.7F), as compared to a much lower percentage of high- velocity events observed with BICD2. Unlike the distribution of velocities observed for

BICD2, which were adequately fit with a single Gaussian, the distributions for both

Hook1 and Hook3 showed a distinct shoulder at higher velocities, and were best fit to a double Gaussian function (Fig. 2.7D) (Woody et al., 2016). A similar complex distribution is evident in initial data on velocities of dynein-dynactin-Hook3 particles from McKenney et al (McKenney et al., 2014). We also noted that within an individual run, Hook-positive particles showed more pronounced variations in instantaneous velocity than was observed for BICD2-postive particles (Fig. 2.7G).

BICD2 is known to be autoinhibited, with truncation of the C-terminus required for robust activation of dynein in vitro (Hoogenraad et al., 2001; McKenney et al., 2014;

Terawaki et al., 2015). In contrast, for both the optogenetic assay described above and 47 these single molecule approaches, full-length constructs of both Hook1 and Hook3 were active in our assay. However, we wondered whether truncating the C-terminal cargo- binding domain would result in further activation, or perhaps reduce the variations in instantaneous velocities observed within runs of full-length Hook1 or Hook3. However, we found truncated constructs of Hook1 and Hook3 lacking the C-terminal domains

(Hook1 1-554aa and Hook3 1-552aa) moved at velocities very similar to those observed with the full-length proteins and displayed similar run lengths (Fig. 2.8B). Again, more than 40% of motility events exhibited mean velocities greater than 1 micron/sec and displayed increased standard deviation in velocities within individual tracks, similar to the full-length proteins (Fig. 2.8C,D). Thus, the observed variations in velocity during a single run are not likely to be due to transient folding of the Hook proteins into an autoinhibited conformation. We also tested several truncated coiled-coil constructs of

Hook1 and Hook3, lacking both the N-terminal and C-terminal domains, but observed little to no motility with these constructs (Fig. 2.8A). Together, these observations suggest that the interaction of Hook proteins with dynein-dynactin is not solely mediated by the central coiled-coil regions, but instead is likely to involve additional contacts with the N-terminal domain. Based on these observations, we suggest that an extended interaction interface involving both the N-terminal domain and the extended coiled-coil domains of Hook proteins may be necessary to induce the rapid velocities and longer run lengths we observed.

Figure 2.8 C-terminally truncated Hook proteins display similar motility to full- length

A) Diagram of conserved domains and predicted coiled-coil regions in Hook1, truncated constructs below with their corresponding motility in TIRF assays (+ =motility, - =no observable motility). MTBD=putative microtubule binding domain, CBD=cargo binding domain. B) Track displacement and velocity distributions for particles tracked with

ImageJ plugin, TrackMate. Data was fitted with a custom Maximum Likelihood 49

Estimation (MLE) program (Woody et al., 2016) and plotted as Probability Density

Functions (PDFs) with 95% CI bootstrapping. (Hk1 (1-554aa) n=107, Hk3 (1-552aa) n=156). C) Percent of events with mean velocity greater than 1 μm/sec. D) Per track standard deviation of instantaneous velocity. Data plotted as a boxplot with Tukey whiskers. BICD2 data repeated from Figure 2.7 for comparison.

IV. Discussion

Hook proteins have been implicated in the regulation of organelle transport in both fungal model systems and mammalian cells (Bielska et al., 2014; Maldonado-Báez et al., 2013; Xu et al., 2008; Zhang et al., 2014). Here we used optogenetic and single molecule approaches to examine the role of mammalian Hook proteins as motor adaptors. We find that mammalian Hook1 and Hook3 proteins enhance dynein-mediated motility. While fungal Hok1 was suggested to function as a bidirectional adaptor (Bielska et al., 2014), we did not find evidence that either Hook1 or Hook3 acts in this way. In our induced recruitment assay, targeting Hook proteins to peroxisomes induced rapid motility toward the perinuclear region, leading to organelle accumulation near the MTOC.

These observations indicate activation of unidirectional, minus end-directed transport, which would not be expected for a bidirectional adaptor. However, it is possible that in other systems Hook proteins also promote kinesin-dependent motility. HeLa cells express 32 kinesins (Maliga et al., 2013), but may not express the specific isoform that interacts with Hooks 1 or 3; alternatively, productive interactions with kinesin may require a specific regulatory environment not fully reconstituted in our optogenetic recruitment assays. Based on current data, we propose that mammalian Hook proteins are unidirectional, dynein-specific adaptors.

Our observations that both Hook1 and Hook3 robustly activate dynein- dependent motility led us to examine the mechanism underlying this process. We found that Hook proteins interact with dynein-dynactin, as the dynein-dynactin-Hook1 complex was efficiently precipitated by antibodies to dynactin. Further, overexpression of Hook1 enhanced the dynein-dynactin interaction. In contrast to the robust co-precipitation of

Hook1 with dynein and dynactin that we observed with an anti-dynactin antibody, we found that the co-precipitation of the complex was disrupted when a dynein intermediate 51 chain antibody was used. This anti-DIC antibody is known to sterically block the binding of dynein to dynactin (Karki and Holzbaur, 1995). Thus, one interpretation of our observation is that Hook proteins interact directly with subunits of the dynactin complex.

Another possibility is that Hook proteins effectively bind to an assembled dynein- dynactin complex. Alternatively, the DIC antibody might block the region of dynein that is necessary for Hook interaction. Interestingly, BICD2 was immunoprecipitated with the

DIC antibody in other studies (Splinter et al., 2012), suggesting the Hook proteins might have more extensive interactions with dynein, contacts not observed for the dynein- dynactin-BICD2 complex (Chowdhury et al., 2015; Urnavicius et al., 2015). Further structural work is needed to determine the specific interaction sites within the dynein- dynactin-Hook complex.

In our induced recruitment assay and TIRF motility assay, we found that Hook proteins enhance dynein-mediated motility, increasing both velocities and run lengths.

Structural studies have suggested several ways that adaptors might modulate dynein to make it more processive. Given the apparent flexibility of the two dynein heads within the dimeric motor complex, it has been suggested that binding of dynactin locks the dynein motor heads into a more favorable conformation for motility (Chowdhury et al.,

2015). In the absence of other factors, the two heads of the dynein dimer on EM grids display a variety of distances from each other, but in the dynein-dynactin-BICD2 complex the motor heads are locked into a more rigid orientation, potentially allowing for more efficient stepping of the heads along the microtubule (Chowdhury et al., 2015). As

Hook proteins enhance the dynein-dynactin interaction (Fig. 2.3), the binding of either

Hook1 or Hook3 might induce this confined dynein conformation and thus enhance processivity. Furthermore, it has been suggested that the C-terminal tail of the dynein

52 motor causes autoinhibition of the motor (Nicholas et al., 2015b; Torisawa et al., 2014); the binding of Hook proteins to the motor complex might relieve this autoinhibition.

In our TIRF assay, Hook proteins display higher velocities and run lengths, even compared to the previously characterized activator BICD2. Within the BICD family, there are also differences in effects on dynein-mediated velocity. For example, BICD-related protein 1 (BICDR-1) was shown to increase the velocity of Rab6 vesicles almost two-fold more than BICD2 (Schlager et al., 2014b). Additionally, BICD-related proteins have an

N-terminal region before the start of the coiled-coil region that is not seen in other BICD proteins. It is possible that this extra N-terminal region plays a role in enhancing velocity analogous to the enhanced motility observed in our analysis of Hook proteins, which we postulate may be due to additional contacts with dynein or dynactin mediated by the N- terminal domain. However, sequence comparisons of the N-terminal domains of Hooks 1 and 3 with those of BICDR-1 do not reveal significant homology, so the specific mechanisms involved may not be analogous.

Our studies with purified proteins from recombinant constructs of Hook1 and

Hook3 indicate that the previously described microtubule binding domain of these Hook proteins does not directly interact with microtubules despite relatively high secondary structure conservation with other calponin-homology domains proteins, such as EB1 and

EB3. Calponin-homology domains are typically found in actin-binding proteins and signaling proteins but have also been found in microtubule binding proteins. Our results indicate that the N-terminal calponin-homology domain of Hook proteins is important for interactions with the dynein-dynactin complex, and not with microtubules. Since the dynactin filament contains actin-related proteins and one β–actin subunit (Urnavicius et al., 2015), it is possible that the calponin-homology domain of Hook interacts with one or more of these dynactin subunits. Alternatively, several studies have reported specialized 53 roles for individual dynactin subunits in tailoring specific cargo transport and could be potential interaction sites for the N-terminal region of Hook proteins. The pointed end of dynactin p25/p27 has been shown to be vital for proper endosomal transport by dynein

(Xiang et al., 2015; Yeh et al., 2012; Zhang et al., 2011). Since fungal Hook proteins have been linked to endosomal transport, it is possible that the N-terminus of Hook proteins interacts with p25/p27, but this would require the coiled coil of Hook proteins to be oriented in a manner opposite to that of BICD2 along the dynactin filament. If, as it is more likely, the orientation of Hook proteins is the same as that of BICD2, the N-terminal calponin-homology domain would be positioned near CapZαβ at the barbed end of the dynactin filament or in close proximity to the flexible subunit of the shoulder, p150Glued.

An interaction with p150Glued could suggest a mechanism for induction of processive motility induced by the binding of Hook, since previous work has suggested that p150Glued can act as a brake for dynein via the ATP-insensitive binding of the CAP-Gly domain to the microtubule (Ayloo et al., 2014). It is possible that the Hook interaction with p150Glued could “release” this brake to allow long processive dynein runs.

Although BICD2 is known to be tightly regulated by autoinhibition, we did not find evidence for autoinhibition of Hook proteins. Full-length constructs of either Hook1 or

Hook3 were more effective than C-terminally-deleted constructs in induced recruitment assays (data not shown), which is not the case for BICD2. It was reported that in an analogous dimerization assay, full-length BICD2 had a very mild effect on organelle redistribution compared to the C-terminally truncated BICD2 construct (Hoogenraad et al., 2003), which is why most studies use a truncated, constitutively active construct. In our TIRF assay, we did not observe any motility with full-length BICD2, while we did with full-length Hook proteins. Thus, Hook proteins may be regulated by additional factors in the cell, instead of by autoregulation. 54

The more divergent C-terminal regions of Hook proteins likely provide specificity for binding to particular cargoes to regulate the utility of Hook adaptors in transport.

Fused Toes (FTS) and FTS-Hook Interacting Protein (FHIP) have been suggested to link

Hook proteins to early endosomes in Aspergillus nidulans, while in mammalian systems

FTS and FHIP are suggested to link Hook proteins to homotypic vesicular protein sorting

(HOPS) complex for endosomal clustering (Xu et al., 2008; Yao et al., 2014). However,

FTS and FHIP seem to bind promiscuously to all three mammalian Hook homologs.

Other studies on Hook proteins have identified some potential candidates for specific interactions. One study found that Hook1 can specifically interact with clathrin- independent endocytosis (CIE) cargo proteins for recycling tubules from early endosomes but not other CIE cargo proteins (Maldonado-Báez et al., 2013). Hook3 has been linked to scavenger receptor A to participate in the endocytotic turnover of the receptor (Sano et al., 2007), while Hook2 has been suggested to interact with centriolin/CEP110 to maintain centrosomal structure (Szebenyi et al., 2007). These unique protein interactions through the C-terminus of Hook proteins might provide enough specificity to regulate these adaptors to modulate motors for particular functions.

Overall, our study provides evidence that mammalian Hook proteins act as dynein adaptors to modulate dynein-mediated cargo transport. It remains to be determined how Hook proteins play a role in intracellular trafficking in more specialized cells like neurons. A recent study linked Hook proteins to Alzheimer’s disease, showing decreased levels of Hook proteins in diseased brains and localized these proteins to the pathological hallmarks of AD, tau aggregates and amyloid plaques (Herrmann et al.,

2015). Future work is needed to better understand the role of these adaptors in intracellular trafficking under both normal conditions and in disease states like

Alzheimer’s. 55

V. Materials and Methods

Reagents

Halo-Hook constructs were generated from human Hook1 sequence (Uniprot code: Q9UJC3) and human Hook3 sequence (Uniprot code: Q86VS8) using the

HaloTag from the pHTN Halo tag CMV-neo vector (Promega). An HA-Hook1 construct in pCMV-HA vector was also generated. Full length mouse BICD2 in pEGFP vector

(GeneBank accession number AJ250106) was a gift from A. Akhmanova and used to generate a truncated construct spanning residues 1-572 fused to HaloTag and cloned into pcDNA3.1. For recruitment assays, a PEX3-GFP-Halo construct was generated including the N-terminal 42 amino acids of the human PEX3 gene for peroxisome targeting (Kapitein et al., 2010). BICD2-mCherry-eDHFR includes residues 1-572 of mouse BICD2, K560-mCherry-eDHFR includes residues 1-560 of human Kinesin-1 heavy chain, p150-mCherry-eDHFR includes full length human p150Glued (DCTN1 sequence, GeneBank accession number NM_004082). Hook-mCherry-eDHFR constructs were either full length or truncated human constructs (Hook1 1-554aa, Hook3

1-552aa), as noted.

Primary antibodies used for western blots include: p150Glued (610474, 1:5,000) from BD transduction, DIC (MAB1618, 1:1000) from Millipore, DHC (R-325, 1:250) from

Santa Cruz, HaloTag (G928A, 1:1000) from Promega, Hook1 (EPR10103(B), 1:500) from AbCam, HA (16B12, 1:1000) from Covance. For IF staining, gamma-tubulin antibody (GTU-88, 1:1000) from Sigma and secondary alexa-fluorophore 633 conjugated antibody from ThermoFisher (A21052, 1:200) were used. All HRP- conjugated secondary antibodies were from Jackson ImmunoResearch Laboratories (IB

1:5000).

For brain lysates, mice (Mus musculus) that were wild type and homozygous knock-in DIC-eGFP-3x-FLAG were used. All animal protocols are approved by the

Institutional Animal Care and Use Committee (IACUC) at the University of Pennsylvania.

Both male and female (4-10 months old) were used.

For RNA interference knockdown of dynein, short interfering RNA (siRNA) duplex from Dharmacon against human dynein heavy chain (GenBank accession number

NM_001376: 5’-GAGAGGAGGUUAUGUUUAAUU-3’) was used at 50 nM.

Cell culture and transfections

Cos7 cells and HeLa cells were cultured in DMEM with 2 mM glutaMAX and 10% fetal bovine serum. Cell were transiently transfected using Fugene6 (Roche) and cells were harvested 18-20 hours post transfection. For RNAi transfection in knockdown experiments, Lipofectamine RNAiMax (Invitrogen) was used for transfection of siRNA duplexes, with 40-48 hours transfection for optimal knockdown.

Immunoprecipitation and Pull-Down Assays

For immunoprecipitation experiments, Protein-G Dynabeads (Promega) beads were incubated with specific antibody for 10 minutes, prior to the addition of lysates and then incubated with lysates for 15 minutes at room temperature. For endogenous dynein-dynactin IPs, mouse brains were homogenized in PHEM buffer (50 mM PIPES,

50 mM HEPES, 1 mM EGTA, 2 mM MgSO4) with 0.5% triton x-100 and protease inhibitors (1 mM PMSF, 0.01 mg/mL TAME, 0.01 mg/mL leupeptin and 0.001 mg/mL pepistatin-A). and then clarified at 38,400x g at 4˚C for 15 minutes. For p150Glued IP,

Cos7 cells expressing Hook or BICD2 constructs were lysed in 30 mM HEPES, 50 mM

NaCl, 1 mM EGTA, 2 mM MgSO4, pH 7.4 with 1 mM DTT, 0.5% triton X-100 and protease inhibitors. Cell lysates were clarified with a 17,000x g centrifugation before use.

For pull-down assays, HaloLink resin (Promega) was prepped by three washes with lysis buffer. Then lysates with Halo-tagged proteins were incubated with resin for 1 hour at 4˚C to attach protein to the resins, followed by a second 1-hour incubation with mouse brain lysates at 4˚C. Cos7 cells expressing Halo-Hook constructs and mouse brains were both lysed in PHEM buffer and prepped as described above for IP experiments. Blots were visualized using enhanced chemiluminescence (SuperSignal

West Pico Chemiluminescent Substrate, Thermo Scientific) with the G:Box and

GeneSys digital imaging system (Syngene). Densitometry was performed with Fiji (NIH).

Microtubule Pelleting Assays

Unlabeled tubulin was polymerized at 5 mg/mL in BRB80 (80 mM PIPES, 1 mM

EGTA, 1 mM MgCl2, pH 6.8) with either 1 mM GTP stabilized with 20 µM Taxol or just 1 mM GMPCPP. Increasing concentrations of microtubules were incubated at 37°C for 20 minutes with equal concentration of purified protein or cell lysate. Then samples were centrifuged at 38,400x g at 25°C for 20 minutes. The supernatant and the pellet were then separated, denatured, and analyzed by SDS-PAGE. For cell lysate experiments,

Cos7 cells transfected with HA-Hook1 (for 18-20 hrs) were lysed in BRB80 buffer with

0.5% triton x-100 and protease inhibitors (as described above) and clarified with two centrifugation steps (at 17000x g and 32000g).

For purified protein experiments, human Hook1 (Uniprot code: Q9UJC3) and

Hook3 (Uniprot code: Q86VS8) were obtained from Open Biosystems. Hook1 fragment

1-443 was amplified by PCR and the N-terminal TEV protease cleavage site was added with a forward primer. Hook1 was cloned between Not1 and Sal1 sites of a modified vector pMAL-c2x (NEB), in which a hexahistidine affinity purification tag was added N- terminal to MBP and a Sac1 site after MBP residue Asn-367 was replaced with a NotI site. Hook3 fragment 1-230 was fused in register to 28 a.a. of GCN4 58

(MKQLEDKVEELLSKNYHLENEVARLKKL) by overlapping primers, and the fusion construct was cloned as above. The proteins were expressed in BL21(DE3) cells

(Invitrogen), grown in Terrific Broth medium at 37°C until the OD600 reached a value of

1.8–2.0. Expression was induced with addition of 0.5 mM isopropylthio-β-D-galactoside and carried out for 16 hours at 18°C. Cells were harvested by centrifugation, resuspended in 50 mM Tris-HCl, pH 8.0, 500 mM NaCl, 5 mM imidazole and 1 mM phenylmethylsulfonyl fluoride, and lysed using a microfluidizer (Microfluidics, Westwood,

MA). The proteins were first purified through a Ni-NTA affinity column (Qiagen) using standard protocol, followed by size exclusion purification on Superdex 200 HL 26/600 gel filtration column (GE Healthcare) in 20 mM HEPES, pH 7.5, 200 mM NaCl, 1mM

EDTA, 1 mM dithiothreitol (DTT). MBP was cleaved with TEV protease and removed by additional size exclusion purification on the same column.

Inducible Recruitment Assay

HeLa cells were transiently cotransfected with PEX3-GFP-Halo and an adaptor/motor construct (BICD2-, K560-, Hook1-, Hook3-, p150Glued -mCherry-eDHFR) for 18-22 hours. For live cell experiments, cells were plated on glass-bottom plates (from

World Precision Instruments) and the caged dimerizer cTMP-Halo was added 30 minutes prior imaging. The dimerizer cTMP-Htag was dissolved in DMSO at 10 mM and stored in amber plastic microcentrifuge tubes at -80 °C. Dimerizer was diluted in media to a final working concentration of 10 μM. Imaging media was composed of Phenol Red- free DMEM with 25 mM HEPES (Gibco), 10% FBS and 2 mM glutaMAX. Live cell imaging was performed on a spinning disk confocal (UltraVIEW VoX; PerkinElmer) with a 405 nm Ultraview Photokinesis (Perkin Elmer) unit on an inverted microscope (Eclipse

Ti; Nikon) using an Apochromat 100x, 1.49 NA oil immersion objective (Nikon) in an environmental chamber at 37°C. Images were acquired at one frame every 2 seconds 59 using a C9100-50 EMCCD camera (Hamamatsu) controlled by Volocity software (Perkin

Elmer). For whole cell photoactivation, the Photokinesis module was set at 20% laser power for 20 cycles.

For fixed recruitment assays, uncaged TMP-Htag dimerizer was added for 45 minutes to HeLa cells 18-20 hours post-transfection. Cells were fixed with ice-cold methanol with 1 mM EGTA. Fixed cells were then stained for gamma-tubulin with primary and secondary antibodies and mounted on glass coverslips with ProLong Gold anti-fade reagent (Invitrogen). Images were taken with a spinning disk UltraVIEW VoX confocal with 100x objective (as described above) and Z-stacks were taken to encompass the whole depth of each cell.

Single Molecule Motility Assay

Motility assays are performed in flow chambers each made of a glass slide and a silanized (PlusOne Repel Silane, GE Healthcare) coverslip, held together by double sided adhesive tape and forming 15 uL volume chambers with vacuum grease. Each of the following solutions was incubated for 5 minutes before washout. First, a 1:40 dilution of monoclonal anti-β-tubulin antibody (T5201; Sigma) was incubated followed by two incubations with 5% pluronic F-127 (Sigma) for blocking the coverslips. Labeled (labeling ratio of 1:40, HiLyte 488 or 647, Cytoskeleton) taxol-stabilized microtubules are then flowed into the chamber and immobilized on β-tubulin antibodies. Finally, diluted cell lysates are flowed in with assay buffer containing 10 mM Mg-ATP, 0.3 mg/mL bovine serum albumin, 0.3 mg/mL casein, 10 mM DTT and an oxygen scavenging system.

For cell lysate prep, HeLa cells 18-20 hours post-transfection were incubated with Halo ligand TMR (Promega), using manufacturer’s guidelines. Cells were lysed in

40 mM HEPES, 1 mM EDTA, 120 mM NaCl, 0.1% triton X-100, 1 mM Mg-ATP, pH 7.4, supplemented with protease inhibitors (as described above). Lysates were clarified with 60 a 17,000x g centrifugation. Before adding to the imaging chamber, the cell lysate extract is diluted in P12 (12 mM PIPES, 1 mM EGTA, 2 mM MgCl2, 20 μM taxol, pH 6.8). Cells were lysed in 100 μL lysis buffer per 70-80% confluent 10 cm plates and then diluted

1:200 for labelled lysate with non-transfected lysate for a total of 1:50 lysate dilution for imaging. All movies were acquired at room temperature at 4 frames per second using

Nikon TIRF system (Perkin Elmer) on an inverted Ti microscope with a 100x objective and an ImageEM C9100-13 camera (Hamamatsu Photonics) controlled by Volocity software.

Image Analysis

For TIRF assays, particle tracking was performed using the TrackMate plugin in

Fiji (Schindelin et al., 2012). Particles runs were tracked if the start and end of the run were seen over the course of the movie. Particles on microtubule bundles were excluded from analysis. Only processive segments of runs were used for velocity and run length measurements. A custom Maximum Estimation Likelihood Matlab program (38) was used to fit velocity and run length data with PDF fits. For fixed recruitment assay,

CellProfiler was used to measure the distance of peroxisomes to the microtubule organization center (Carpenter et al., 2006). In this program, the MTOC was manually identified, while both peroxisomes and the cell outline were identified by the program.

Measured distances of peroxisomes from the MTOC were normalized by dividing by the longest diameter of the cell and multiplying by 100. Normalized distances were plotted as an averaged distribution with error bars representing S.E.M.

CHAPTER 3: A Conserved Interaction of the Dynein Light Intermediate Chain with Dynein-Dynactin Effectors Necessary for Processivity

This chapter is adapted from:

Lee I.G., Olenick, M.A., Boczkowska, M, Franzini-Armstrong, C., Holzbaur, E.L.F., Dominguez, R.

(2018) A conserved interaction of the dynein light intermediate chain with dynein-dynactin effectors necessary for processivity. Nature Communications 9(1): 986.

I. Summary

Cytoplasmic dynein is the major minus end-directed microtubule-based motor in cells. Dynein processivity and cargo selectivity depend on cargo-specific effectors that, while generally unrelated, share the ability to interact with dynein and dynactin to form processive dynein-dynactin-effector complexes. How this is achieved is poorly understood. Here, we identify a conserved region of the dynein Light Intermediate

Chain-1 (LIC1) that mediates interactions with unrelated dynein-dynactin effectors.

Quantitative binding studies map these interactions to a conserved helix within LIC1 and to N-terminal fragments of Hook1, Hook3, BICD2, and Spindly. A structure of the LIC1 helix bound to the N-terminal Hook domain reveals a conformational change that creates a hydrophobic cleft for binding of the LIC1 helix. The LIC1 helix competitively inhibits processive dynein-dynactin-effector motility in vitro, whereas structure-inspired mutations in this helix impair lysosomal positioning in cells. The results reveal a conserved mechanism of effector interaction with dynein-dynactin necessary for processive motility.

II. Introduction

Cytoplasmic dynein 1 (dynein) is the major minus-end-directed microtubule- based motor in eukaryotic cells. It is responsible for the transport of very diverse cargoes from the periphery to the center of the cell, including lysosomes, mitochondria, and autophagosomes (Barlan and Gelfand, 2017; Bonifacino and Neefjes, 2017; Carter et al., 2016; Hoogenraad and Akhmanova, 2016). Recent work has shown that both cargo specificity and processivity depend on the interaction of dynein with its general adaptor, the dynactin complex, and a series of cargo-specific effectors, including BICD2

(McKenney et al., 2014; Olenick et al., 2016; Schlager et al., 2014a; Splinter et al.,

2012), Hook1/3 (McKenney et al., 2014; Olenick et al., 2016; Schroeder and Vale,

2016), Spindly (McKenney et al., 2014), FIP3 (McKenney et al., 2014), and NIN/NINL

(Redwine et al., 2017). These proteins are generally unrelated at the sequence level, but they all contain large portions of predicted coiled-coil structure and share the ability to interact with both dynein and dynactin to activate processive motility (McKenney et al.,

2014; Olenick et al., 2016; Redwine et al., 2017; Schlager et al., 2014a; Schroeder et al.,

2014). It remains unclear, however, whether each effector has evolved these functions independently or whether they share common structural-functional features and similar interactions with dynein and dynactin. Here, we show that a conserved amphipathic helix within the unstructured C-terminal region of the dynein Light Intermediate Chain 1 (LIC1) interacts with diverse dynein-dynactin effectors. The interactions were quantitatively characterized using purified proteins and isothermal titration calorimetry (ITC). A crystal structure of the LIC1 helix in complex with the N-terminal Hook domain of Hook3 reveals a conformational change within the Hook domain that gives rise to a hydrophobic cleft where the LIC1 helix binds. Supporting the importance of the LIC1-effector interaction, we found that the LIC1 helix competitively inhibits the processive motility of dynein- 64 dynactin in complex with either Hook3 or BICD2 in single-molecule assays using total internal reflection fluorescence (TIRF) microscopy. Finally, in cellular assays, mutating the LIC1 helix leads to defective dynein-driven positioning of lysosomes. Together, the results reveal the existence of a conserved mechanism of interaction between functionally unrelated dynein-dynactin effectors and the dynein LIC1, which is required for processive dynein-driven transport.

III. Results

Hook interacts with the dynein LIC1 via the Hook domain

The dynein LICs, comprising two closely related isoforms (LIC1 and LIC2), consist of two domains – an N-terminal GTPase-like domain that interacts with the dynein heavy chain (Schroeder et al., 2014) and a less conserved and predicted unstructured C-terminal region, referred to here as the effector-binding domain. Using pull-down studies, it had been previously shown that the LIC1 effector-binding domain interacts with several dynein-dynactin effectors, including Hook3, FIP3, BICD2, and

Spindly (Gama et al., 2017; Schroeder and Vale, 2016; Schroeder et al., 2014). On the other hand, a group of dynein-binding proteins, including BICD2, Spindly, HAP1, and

TRAK share a coiled coil segment, termed the CC1-Box, that has been directly implicated in LIC1 binding (Gama et al., 2017). Here, we set out to specifically map and quantitatively characterize the interactions of LIC1 with several dynein-dynactin effectors, including Hook1, Hook3, BICD2, and Spindly.

Hook1 and Hook3 are known dynein effectors that function in endosomal transport (Luiro et al., 2004; Maldonado-Báez et al., 2013; McKenney et al., 2014;

Olenick et al., 2016; Schroeder and Vale, 2016; Xu et al., 2008). We expressed truncated constructs of human Hook1 and Hook3 in E. coli, whereas full-length Hook1 was expressed in insect cells (Fig. 3.1A,B). Because Hook proteins contain several regions of predicted coiled coil (CC1–4) (Fig. 3.1A), we first analyzed whether these constructs were dimeric or monomeric using light scattering. A construct corresponding to the N-terminal Hook domain (Hook111–166) was monomeric, whereas construct

Hook111–443, extending to the end of CC2 was dimeric (Fig. 3.1C). In contrast, Hook111–

238, comprising only the globular Hook domain and CC1 region, was in equilibrium between dimers and monomers, as indicated by an experimentally-measured mass of 66

39.7 kDa, i.e. intermediate between those of the dimer and the monomer. Full-length human LIC1 was expressed as a fusion protein with MBP (maltose binding protein) to increase its solubility and was also found to be monomeric by light scattering (Fig. 3.1C).

Using ITC, MBP-LIC1FL bound Hook111–443 with low micromolar affinity (KD = 8.1

µM) and ~1:1 stoichiometry, i.e. two LIC1 molecules per Hook1 dimer (Fig. 3.1D). Note that this ITC titration was performed at 30°C, instead of 20°C for most titrations performed here, because the amount of heat given off by this reaction was too small to allow for reliable fitting of the thermodynamic parameters. Consistent with the light scattering results, the titration of Hook111–238 into buffer produced a significant endothermic reaction, which was interpreted as indicative of dimer dissociation, with a

KD of 2.1 µM (Fig. 3.1E). This conclusion was confirmed by analysis of Hook11–239GCN4, a dimeric construct stabilized through the addition of a GCN4 leucine zipper at the C- terminus, whose titration into buffer did not produce any significant heat change (Fig.

3.1E). Hook11–239GCN4 bound MBP-LIC1FL with a KD of 12.9 µM and ~1:1 stoichiometry

(Fig. 3.1F), which is very similar to what was observed with Hook111–443 (Fig. 3.1D) despite the fact that the titration was inverted by placing Hook11–239GCN4 in the syringe and MBP-LIC1FL in the cell. The monomeric construct Hook111-166 also bound MBP-

LIC1FL with similar affinity (KD = 12.7 µM) and 1:1 stoichiometry. Together these results show that: a) the LIC1-binding site is fully contained within the conserved N-terminal

Hook domain, b) each Hook dimer interacts with two LIC1s, and c) the CC1 region of

Hook forms an unstable coiled coil, which on its own cannot support stable Hook dimerization.

To gain further insights into the overall structure of Hook and the disposition of the Hook domain with respect to the coiled coil segments, we used rotary shadowing electron microscopy to visualize full-length Hook1 (Fig. 3.1H). Hook1 had a kinesin-like 67 appearance, with most particles displaying two well-separated globular domains at one end, connected through a short neck-like region to a long thin rod, which was often interrupted by a pronounced kink, followed by a shorter thin rod. These features were interpreted to correspond to the N-terminal Hook domain, the unstable CC1 region, CC2, the central so-called Spindly motif (Gama et al., 2017), and CC3, respectively (Fig. 3.1I).

The smaller C-terminal cargo-binding domain (CBD) was only occasionally visualized as a defined structural feature (Fig. 3.1I). This assignment of domains is consistent with the length of the segment extending from the end of the neck region to the central kink, whose mean length of ~31 nm approximately corresponds to the predicted dimensions of CC2 (~27 nm) (Fig. 3.1J), as estimated from the structures of other coiled coil proteins. The dimensions of the remaining smaller domains cannot be accurately measured at this resolution. The assignment of domains is also consistent with structural predictions and sequence conservation analyses, showing a series of coiled coil segments (CC1, CC2, CC3, and CC4) interspersed with three globular regions (Hook domain, Spindly motif, and CBD), connected by short, unstructured loops of lower sequence conservation (Fig. 3.S1). The variability of the kink angle between CC2 and

CC3 suggests that the regions N- and C-terminal to the Spindly motif move relatively independently of each other, i.e. the Spindly motif appears to function as a ‘hinge’.

Finally, the fact that the two globular Hook domains appear well separated from each other in most of the particles visualized is consistent with the two helices that form the

CC1 segment (neck) not forming a stable coiled coil, as also suggested by the light scattering (Fig. 3.1C) and ITC (Fig. 3.1E) results.

Figure 3.1 Hook interacts with LIC1 via the N-terminal Hook domain

Figure 3.1 Hook interacts with LIC1 via the N-terminal Hook domain.

A) Domain organization of Hook1 and constructs used in this study (CC, coiled-coil;

CBD, cargo-binding domain). B) SDS-PAGE (4–12%) showing several of the proteins used in this study. C) SEC-MALS analysis of Hook1 constructs and MBP-LIC1FL (color coded as indicated). The molar mass determined from light scattering (right y-axis) and the UV absorption at 280 nm (left y-axis) are plotted as a function of the elution volume.

The theoretical masses are given in parenthesis. D-G) ITC titrations of LIC1 and Hook1 constructs as indicated. Listed with each titration are the concentrations of the protein in the syringe and in the cell, as well as the temperature of the experiment and parameters of the fit (stoichiometry, N; dissociation constant, KD). Errors correspond to the s.d. of the fits. Open symbols correspond to titrations into buffer (except part e, where both titrations are into buffer). H) Representative rotary shadowing EM image of Hook1FL.

Scale bar, 100 nm. White squares indicate individual Hook1FL molecules highlighted in the zooms shown on the right. Scale bar, 50 nm. I) Close-up view of a representative

Hook1FL molecule shown alongside a cartoon representation of the Hook1 domains based on the rotary shadowing EM, secondary structure and sequence conservation analyses (see Fig. 3.S1). Scale bar, 50 nm. J) Length distribution of the region spanning from the end of the neck to the kink. Bin size, 5 nm, n = 33.

Figure 3.S1 Domain architecture of Hook1

The domain architecture of human Hook1 was analyzed using several bioinformatics approaches, including: sequence conservation analysis with the program Scorecons

(Valdar, 2002), secondary structure prediction with the program Jpred4 (Drozdetskiy et al., 2015), hydrophobic cluster analysis with the program HCA (Gaboriaud et al., 1987), and coiled coil prediction with the program Coils (Lupas et al., 1991). Sequence conservation scores were calculated from an alignment of 96 Hook sequences from different species and isoforms. The resulting per-residue scores were then plotted against the human Hook1 sequence, i.e. residue insertions in other sequences

(compared to human Hook1) are not shown. The secondary structure and coiled coil predictions suggest that Hook consists solely of -helices, since not a single -strand was predicted.

A helix in LIC1 C-terminal region binds the Hook domain

The C-terminal effector-binding domain of LIC1 (human LIC1 residues 390–523) has been shown to interact with dynein-dynactin effectors, including Hook3, BICD2, and

Spindly (Gama et al., 2017; Schroeder and Vale, 2016; Schroeder et al., 2014).

However, it was unknown whether different effectors bound to the same or different regions on the LIC1 C-terminus, and these interactions were characterized by qualitative rather than quantitative analyses. Here, we set out to map the specific region of the LIC1

C-terminus implicated in interactions with Hook proteins and other effectors (see below) and quantitatively characterize the interactions. Most of the LIC1 effector-binding domain is predicted to be unstructured and, unlike the GTPase-like domain, it is not highly conserved among species (Fig. 3.S2). However, sequence analysis reveals two regions of relatively high conservation that coincide with predicted α-helical segments, which we named Helix-1 (human LIC1 residues 440–456) and Helix-2 (residues 493–502) (Fig.

3.2A and Fig. 3.S2). To test whether these conserved helical segments participate in the interaction with the Hook domain, we generated two C-terminally truncated LIC1 constructs, MBP-LIC11–461, which removes the region C-terminal to Helix-1, and MBP-

LIC11–437, which additionally removes Helix-1 (Fig. 3.2A). MBP-LIC11–437 failed to bind the Hook domain by ITC (Fig. 3.2C), whereas MBP-LIC11–461 bound the Hook domain

(Fig. 3.2B) with nearly the same affinity (KD = 10.1 µM) as MBP-LIC1FL (KD = 12.7 µM)

(Fig. 3.1G). These results suggested that the binding site is contained within Helix-1.

Consistent with this conclusion, the Hook domain failed to bind to construct MBP-

LIC1F447A,F448A, in which two strictly conserved phenylalanine residues in the middle of

Helix-1 were simultaneously mutated to alanine (Fig. 3.2D).

To further test the role of Helix-1 in Hook binding, we expressed a 26-a.a. peptide (LIC1433–458), extending several amino acids N- and C-terminally to the predicted 72 helical segment to ensure proper folding of Helix-1. The Hook domain of Hook1 bound to

LIC1433–458 with nearly the same affinity (KD = 15.7 µM) as to MBP-LIC1FL (Figs. 3.1G and 3.2E). The Hook domain of Hook3 (human Hook3 residues 1–160) also bound

MBP-LIC1FL and Helix-1 with similar affinities (Fig. 3.2F,G), and these affinities were comparable to those observed with the Hook domain of Hook1 (Figs. 3.1G and 3.2E).

Together, these results map the LIC1-Hook interaction to the conserved Helix-1 within the effector-binding domain of LIC1 and the N-terminal Hook domain of both Hook1 and

Hook3. Furthermore, the conserved hydrophobic residues F447 and F448 within Helix-1 likely form part of the binding interface.

Figure 3.2 The conserved Helix-1 within the LIC1 effector-binding domain binds the Hook domain

A) Alignment of LIC sequences from different species and isoforms around the predicted Helix-1 within the C-terminal effector-binding domain (top) and domain diagram of human LIC1 showing the constructs used in this study (bottom). The name of each sequence includes the organism of origin and UniProt accession code. Yellow and orange backgrounds indicate 70% and 100% sequence conservation, respectively. Red stars highlight residues F447 and F448 that were mutated to alanine. The predicted

Helix-1 and Helix-2, coinciding with regions of higher sequence conservation (see Fig.

3.S2), are highlighted in the domain diagram, and Helix-1 is also depicted above the sequence alignment. The region corresponding to the Helix-1 (LIC1433–458) peptide is contoured red. B-G) ITC titrations of Hook111–166 and Hook31–160 into LIC1 constructs (as indicated). Listed with each titration are the concentrations of the protein in the syringe and in the cell, as well as the temperature of the experiment and parameters of the fit

(stoichiometry, N; dissociation constant, KD). Errors correspond to the s.d. of the fits.

Open symbols correspond to control titrations into buffer.

Figure 3.S2 Alignment of the Effector-Binding Domain of LIC sequences

A) Domain diagram of human LIC1, showing the location of the predicted Helix-1 and

Helix-2 within the C-terminal Effector-Binding Domain (EBD). B) Sequence alignment of the EBD of LIC sequences from different species and isoforms. The name of each sequence includes the organism of origin. Yellow and orange backgrounds indicate 70% and 100% sequence conservation, respectively. The predicted Helix-1 and Helix-2

(depicted above the sequence alignment) coincide with regions of higher sequence

76 conservation. The region corresponding to the Helix 1 (LIC1433-458) peptide is contoured red.

Structure of a complex of the Hook domain and the LIC1 helix

To further understand the mechanism of interaction between Hook and LIC1, we determined the crystal structure of human Hook31–160 in complex with human LIC1 Helix-

1 at 1.5 Å resolution (Fig. 3.3A–C and 3.S3). The electron density is well defined for

Hook3 residues 10–160 and LIC1 residues 441–454 (Fig. 3.3B). The first nine amino acids of Hook3 and residues 433–440 and 455–458 of Helix-1 were disordered and are thus unlikely to participate in the interaction. As previously reported (Schroeder and

Vale, 2016), the Hook domain displays a canonical 7-helix calponin homology (CH)-like fold, featuring an additional helix at the C-terminus, termed helix α8. Generally, the structure superimposes well with that of the unbound Hook domain determined previously (Schroeder and Vale, 2016), with an root-mean-square deviation of 1.4 Å for

136 equivalent Cα atoms (Fig. 3.3D). However, the Hook domain-specific helix α8, which in the unbound structure is fully extended and interacts in anti-parallel fashion with the same helix from a symmetry-related molecule in the crystal, is broken into two helices

(α8a and α8b) in the current structure (Fig. 3.3D), giving rise to a V-shaped hydrophobic cleft that constitutes the binding site for LIC1 Helix-1 (Fig. 3.3C). As predicted, the visualized portion of Helix-1 is folded as an amphipathic α-helix, with its hydrophobic surface facing the hydrophobic cleft of the Hook domain (Fig. 3.3C). All the highly conserved, hydrophobic amino acids of the LIC1 effector-binding domain are directly inserted into the hydrophobic cleft of the Hook domain, including L444, F447, F448. and

L451, explaining why the mutant MBP-LIC1F447A,F448A failed to bind the Hook domain (Fig.

3.2D). The binding interface also coincides with the most highly conserved surface of the

Hook domain (Fig. 3.3E).

The Hook domain also interacts with a second LIC1 Helix-1 from a neighboring complex in the crystal lattice (Fig. 3.3E). This interaction presents the less conserved, hydrophilic surface of Helix-1 to a less conserved surface on the Hook domain, which a priori is inconsistent with a native interaction. Yet, to rule out this interaction, we generated two Hook domain mutants: A138D, testing the presumed crystal packing contact, and M140D, testing the anticipated native binding site (Fig. 3.3F). The Hook31–

160A138D mutant bound MBP-LIC1FL with the same affinity as wild type Hook31–160

(compare Figs. 3.2F and 3.3G), whereas the Hook31–160M140D mutant failed to bind

MBP-LIC1FL (Fig. 3.3H), confirming that the native binding site of Helix-1 is located at the interface between α8a and α8b and conferring functional significance to the conformational change that splits helix α8 into two helices. Indeed, even in the presence of Helix-1, we obtained a second crystal form showing the reported extended conformation of helix α8 (Schroeder and Vale, 2016), but the LIC1 peptide was not bound in these crystals. To further test the importance of the conformational change in helix α8 for LIC1 binding, we generated a truncated construct, Hook31–143, lacking the

α8b portion of helix α8, i.e. the region that bends back to form the V-shaped cleft (Fig.

3.3D). Hook31–143 failed to bind MBP-LIC1FL (Fig. 3.3I). Collectively, these results confirm that the extended helix α8 of the Hook domain, which distinguishes this domain from the canonical CH fold, undergoes a conformational change to produce a conserved, hydrophobic cleft for binding of the conserved LIC1 Helix-1.

Figure 3.3 Crystal structure of the Hook domain in complex with LIC1 Helix-1

A) Ribbon and surface representation of the structure of Hook31–160 (magenta) in complex with Helix-1 (LIC1433–458, blue). The side chains of Helix-1 are shown using a sticks representation, colored by atom type. B) Close-up view of the Helix-1 binding site, showing the 2Fo-Fc electron density map (blue mesh) at 1.5 Å resolution, contoured at

1σ around an all-atom representation of Helix-1. C) Close-up view of the Helix-1 binding site, showing the residues at the hydrophobic contact interface. D) Superimposition of the structure of the Hook domain from the Helix-1-bound complex (magenta) and unbound structure (grey) (Schroeder and Vale, 2016). A conformational change in the C- terminal helix α8, which distinguishes this domain from the CH domain, leads to the formation of two helices (α8a and α8b) that constitute the binding site for Helix-1. E)

Sequence conservation of the Hook domain (see also Fig. 3.S1) mapped onto the surface of the structure and colored from low to high conservation using a red to green gradient. In the crystal lattice, the Hook domain contacts a second Helix-1 from a neighboring complex (light blue). F) Surface representation of the Hook domain

(magenta), showing in yellow the two amino acids mutated (A138D and M140D) to test the functional relevance of the two Helix-1 interactions. G-I) ITC titrations of the indicated

Hook31–160 mutants into MBP-LIC1FL. Experimental conditions and fitting parameters are listed. Errors correspond to the s.d. of the fits. Open symbols correspond to titrations into buffer.

The LIC1 helix binds diverse dynein-dynactin effectors

Next, we asked whether LIC1 Helix-1 was also implicated in interactions with other dynein-dynactin effectors that are generally structurally and functionally unrelated to each other. As mentioned above, a recent study found that a group of dynein-dynactin 80 effectors share a region termed the CC1-Box that was implicated in LIC1 binding through pull-down and mutagenesis studies (Gama et al., 2017).

To test whether LIC1 Helix-1 also mediates the interaction with CC1-Box- containing effectors, we expressed N-terminal fragments of two effectors: BICD21–98 and

Spindly1–142 (Fig. 3.4A,B). These constructs extend N- and C-terminally from the CC1-

Box to include the first predicted coiled coil segment of each protein and ensure proper dimerization. Note that when bound to dynein-dynactin, the entire N-terminal region of these two proteins appear to form uninterrupted coiled coil structures (Zhang et al.,

2017), as depicted in Fig. 3.4A. Indeed, as verified by light scattering (Fig. 3.4C), both

BICD21–98 and Spindly1–142 form stable coiled coil dimers, with experimentally-determined masses approximately double those calculated from sequence. By ITC, the titrations of

LIC1433–458 into BICD21–98 (Fig. 3.4D) and Spindly1–142 (Fig. 3.4E) fitted best to two- binding-site isotherms. The affinities of the two binding sites were similar to each other, and they were also similar for the two effectors (with KDs ranging from 1.5 to 7.6 µM).

Curiously, however, despite sharing a similar CC1-Box and displaying similar affinities for LIC1 Helix-1, the titrations into BICD21–98 and Spindly1–142 had different overall appearances (Fig. 3.4D,E). For the Spindly1–142 titration, in particular, the two binding sites have very close affinities and are probably saturated at the same time, but the first site has a mild endothermic character whereas the second site has a strong exothermic character, which masks the endothermic signal of the first part of the titration, explaining the peculiar shape of this reaction. Likely, LIC1 binding produces different types of conformational changes in these two proteins, which other than the CC1-Box share no apparent sequence similarity. These results confirm that LIC1 Helix-1 constitutes a common binding site for unrelated dynein-dynactin effectors, including CC1-Box- containing effectors (BICD, Spindly) and Hook-family effectors. 81

Figure 3.4 LIC1 Helix-1 mediates the interaction with CC1-Box-containing effectors

A) Domain organization of human BICD2 and Spindly and constructs used in this study.

B) Sequence alignment of the CC1-Box region (left) of several proteins that link dynein to different cargoes and perform different functions (Chan et al., 2009; Hoogenraad and

Akhmanova, 2016; van Spronsen et al., 2013; Wong and Holzbaur, 2014) (right). Yellow and orange backgrounds indicate 70% and 100% sequence conservation, respectively.

C) SEC-MALS analysis of BICD21–98 (black) and Spindly1–142 (grey). The molar masses

82 determined from light scattering (right y-axis) and the UV absorption at 280 nm (left y- axis) are plotted as a function of the elution volume. The theoretical masses are given in parenthesis. D-E) ITC titrations of Helix-1 (LIC1433–458) into BICD21-98 and Spindly1–142, respectively. The experimental conditions and fitting parameters are listed for each titration. Errors correspond to the s.d. of the fits. Open symbols correspond to control titrations into buffer.

The LIC1 helix/effector interaction is crucial for motility

To test the functional significance of the LIC1 Helix-1 interaction with dynein- dynactin effectors, we utilized an in vitro single molecule approach to track the movement of dynein-dynactin-effector complexes obtained from cell extracts using TIRF microscopy (Olenick et al., 2016). Lysates of HeLa cells expressing Halo-tagged

Hook31–552 labeled with TMR-HaloTag ligand were flowed into a chamber containing

Taxol-stabilized microtubules immobilized on coverslips. The dynein-driven motility of single molecules was then monitored both in the absence or presence of increasing concentrations of Helix-1 or Helix-1F447A,F448A, a peptide carrying two mutations found to inhibit binding of full-length LIC1 to the Hook domain (Fig. 3.2D). Consistent with previous reports (McKenney et al., 2014; Olenick et al., 2016), in the absence of Helix-1 we observed robust motility of Halo-Hook31–552-positive complexes along microtubules, characterized by long run lengths and high velocities (Fig. 3.5A). In contrast, we observed a marked inhibition of processive motility with the addition of Helix-1, with nearly complete inhibition at Helix-1 concentrations of 100 µM or higher, whereas the addition of Helix-1F447A,F448A did not inhibit motility (Fig. 3.5A). Similar results were observed in experiments that tracked the movement of dynein-dynactin-BICD2 complexes obtained from cell extracts expressing Halo-BICD21–572 labeled with TMR- 83

HaloTag ligand (Fig. 3.5B). In this case, however, higher concentrations of Helix-1 (>

200 µM) were required for full inhibition, which is not entirely unexpected for in trans competition of an intramolecular interaction.

To assess whether the LIC1-effector interaction contributes to organelle motility in cells, we analyzed the distribution of lysosomes in HeLa cells expressing GFP,

LIC1WT-GFP or the mutant LIC1F447A,F448A-GFP that does not interact with Hook1 (Fig.

3.2D). Importantly, this mutation is predicted to also block the interaction of LIC1 with other effectors, since we found that Helix-1 is involved in interactions with several effectors (Figs. 3.2E,G and 4D,E). Lysosomes are well-characterized cargoes of dynein, which drives perinuclear clustering of lysosomes near microtubule minus ends

(Johansson et al., 2007; Jordens et al., 2001), and LIC1 is known to be required for this activity (Tan et al., 2011). Compared to the expression of GFP alone, the expression of

LIC1WT-GFP did not significantly change the distribution of lysosomes, visualized by anti-

LAMP1 staining of cells fixed 18–22 h after transfection. In contrast, the expression of the LIC1F447A,F448A-GFP mutant resulted in an abnormal localization of lysosomes (Fig.

3.5C). In these cells, lysosomes appeared dispersed throughout the cytoplasm and did not show the characteristic perinuclear clustering seen in control cells (Fig. 3.5C). In a blind analysis, LIC1F447A,F448A-GFP-expressing cells displayed a significantly higher percentage of abnormally positioned lysosomes compared to cells expressing GFP or

LIC1WT-GFP (Fig. 3.5D). Together, these results show that the LIC1-effector interaction mediated by Helix-1, specifically residues F447 and F448, is absolutely required for processive dynein-based motility in vitro and in cells.

Figure 3.5 The Helix-1-effector interaction is important for processive motility in vitro and in cells

A,B) Time series and kymographs (1 min) of Halo-Hook31–552 and Halo-BICD21–572 runs on microtubules (magenta) in the absence (control) or the presence of Helix-1 or Helix-

1F447A,F448A peptides (as indicated) analyzed by TIRF microscopy. Arrows indicate a motile particle. Max = maximum projection. Scale bar = 5 µm. Quantifications (right) 85 show that the number of motile events declines with increasing Helix-1 concentrations, but not Helix-1F447A,F448A. The statistical significance of the measurements was determined using a one-way ANOVA test, analyzing N = 6–21 videos and a minimum of

3 individual cell lysates per condition (n.s., non-significant; *, p ≤ 0.05; **, p ≤ 0.01; ***, p

< 0.001). Error bars correspond to the s.e.m. C) Representative images of LAMP1 staining of fixed HeLa cells expressing GFP, LIC1WT-GFP or LIC1F447A,448A-GFP. Note that the LAMP1 puncta become more dispersed with the expression of LIC1F447A,448A-

GFP but not LIC1WT-GFP. Cell perimeters are outlined white. Scale bar = 10 µm. D)

Percentage of cells with abnormal LAMP1 staining from fixed HeLa cells expressing

GFP, LIC1WT-GFP and LIC1F447A,448A-GFP. The statistical significance of the measurements was determined using a one-way ANOVA test, analyzing N = 148 (GFP),

N = 77 (LIC1WT-GFP) and N = 48 (LIC1F447A,448A-GFP) cells from three independent repeats (n.s., non-significant; *, p ≤ 0.05; **, p ≤ 0.01). Error bars correspond to the s.e.m.

IV. Discussion

Cytoplasmic dynein is responsible for most cellular activities requiring microtubule minus end-directed motility. However, in isolation, dynein is not very processive (McKenney et al., 2014; Trokter et al., 2012). It is now recognized that dynein’s functional diversity, including cargo-specificity and processivity, depends on its interaction with the general adaptor dynactin, regulated by an ever-expanding family of dynein-dynactin effector proteins, including BICD2 (McKenney et al., 2014; Schlager et al., 2014a; Splinter et al., 2012), Hook1/3 (McKenney et al., 2014; Olenick et al., 2016;

Schroeder and Vale, 2016), Spindly (McKenney et al., 2014), FIP3 (McKenney et al.,

2014), and NIN/NINL (Redwine et al., 2017). These proteins have been distinctly called adaptors (McKenney et al., 2014; Schroeder and Vale, 2016) or regulators (Kardon and

Vale, 2009). We have used here the more general term “effectors” because they do both

– they help bring together dynein and dynactin and recruit specific cargoes, which are typical adaptor functions, but they also activate dynein processivity, thus playing a regulatory role. We have a limited understanding of how dynein-dynactin effectors exert these diverse functions, and the lack of recognizable sequence similarity or a common dynein-dynactin-binding motif among all of them has limited our ability to establish general structural-functional correlations. Most of the effectors, however, appear to have cargo-specific binding domains toward their C-termini (Bielska et al., 2014; Gama et al.,

2017; Hoogenraad et al., 2001; Horgan et al., 2010; Sano et al., 2007; Xu et al., 2008;

Zhang et al., 2014). Another structural feature shared by all the known effectors is the presence of long regions of coiled coil. Cryo-EM (Chowdhury et al., 2015; Grotjahn et al.,

2018; Urnavicius et al., 2015, 2018), biochemical (Karki and Holzbaur, 1995; Morgan et al., 2011; Siglin et al., 2013; Vaughan and Vallee, 1995), and proteomics (Redwine et al., 2017) studies have shown that there are at least three major points of contact among 87 dynein, dynactin, and the effectors, and each of these interactions could in principle contribute toward the overall affinity of their ternary complexes, as well as the adaptor and regulatory activities of each effector. First, dynein and dynactin interact with each other with micromolar affinity (KD = ~3 µM) via a direct interaction between the dynein intermediate chain and the p150Glued subunit of dynactin (Karki and Holzbaur, 1995;

Morgan et al., 2011; Siglin et al., 2013; Vaughan and Vallee, 1995), as well as through the tail domain of the heavy chain (Chowdhury et al., 2015; Grotjahn et al., 2018;

Urnavicius et al., 2015, 2018). Second, a long coiled coil segment in each effector appears to intercalate at the interface between dynein and dynactin, running along the dynactin surface with the N- terminal end directed toward the barbed end of the actin-like dynactin filament (Chowdhury et al., 2015; Grotjahn et al., 2018; Urnavicius et al., 2015,

2018). This interaction appears to modulate the affinity between dynein and dynactin, as well as the number of dynein molecules that are recruited onto the dynactin scaffold

(Grotjahn et al., 2018; Urnavicius et al., 2018).

Evidence for the third type of interaction, involving dynein’s LIC1 subunit and N- terminal sequences in various effectors, has so far been limited to pull-down (Gama et al., 2017; Schroeder and Vale, 2016; Schroeder et al., 2014) and proteomics (Redwine et al., 2017) studies. Here, we have mapped this interaction to a conserved helix (Helix-

1) within the otherwise unstructured and poorly conserved C-terminal region of LIC1. We have further shown that Helix-1 mediates the interaction with structurally and functionally unrelated effectors and that the interactions typically have low micromolar affinities. We were also able to visualize the structural basis of this interaction at high-resolution for the

Hook subfamily of effectors (comprising three isoforms, Hook1–3). Finally, we demonstrated that this interaction enhances the processive motility of dynein in vitro and that disruption of the LIC1-effector interface affects organelle transport in cells. 88

Somewhat analogous to our findings, an interaction between the light and intermediate chains of yeast dynein has been implicated in dimerization and processive motility (Rao et al., 2013).

Curiously, the LIC1-effector interaction involves a conserved motif on the LIC1 side of the interface but different surfaces on the effector side. For Hook-family effectors, the interaction involves the N-terminal Hook domain, which has a globular fold related to the CH domain. However, it is the extended helix α8 of the Hook domain (absent in the

CH fold) that mediates the interaction by forming a V-shaped hydrophobic cleft after splitting into two helices. In BICD2 and Spindly, the interaction involves the CC1-Box, which forms part of a longer coiled coil segment (Fig. 3.4A). Conceivably, the two helices of the coiled coil could separate, partially exposing the hydrophobic core of the coiled coil to create symmetric binding sites for LIC1 Helix-1 on both sides of the coiled coil.

This would give rise to a binding site that is different in sequence but possibly structurally similar to that of the Hook domain.

The effectors analyzed here bind LIC1 with 1:1 stoichiometry, or rather with 2:2 stoichiometry since all the effectors identified to date form dimers. In this way, each effector could in principle tether two LIC1 subunits from a single dynein dimer or from two different dynein dimers bound simultaneously to the dynactin complex. In the case of

Hook-family effectors, the two LIC1-binding sites are physically separated from one another, as suggested by our rotary shadowing EM analysis (Fig. 3.1H,I), whereas in

CC1-Box-containing effectors, the two binding sites occur on the same coiled coil, i.e. adjacent to each other. Such structural differences, as well as differences in the affinities of the LIC1-effector interactions, may play a modulatory role, by forming dynein- dynactin-effector complexes of different affinities and characterized by different run lengths. In this regard, it is important to note that the dynein-dynactin-Hook3 complex 89 displays a bimodal velocity distribution and faster velocities (Olenick et al., 2016) than the dynactin-BICD2 complex characterized by a single velocity distribution (McKenney et al., 2014; Schlager et al., 2014a).

The activation of dynein processivity proceeds through a conformational change from an auto-inhibited so-called “phi-particle” state to a “parallel-heads” state capable of binding microtubules upon complex formation with dynactin-effectors (Zhang et al.,

2017). The auto-inhibited state is stabilized by inter-heavy chain interactions, including near the LIC1 subunit. Because the LIC1-effector interaction appears to “pull” on the

LIC1 subunit and with it on the dynein heavy chain, we propose that it may help reposition the dynein heads for optimal interaction with microtubules by breaking the interaction between heavy chains (Fig. 3.6), and thus this interaction would be specifically important for the activation of dynein processivity. It remains to be demonstrated whether the LIC1-effector interaction is engaged at all times and whether it is absolutely required for dynein-dynactin complex formation, which could primarily depend on the other two interactions mentioned above (i.e. direct dynein-dynactin contacts and contacts mediated by an N-terminal coiled-coil segment of the effectors).

Figure 3.6 Model for cargo transport by dynein-dynactin-effector complexes

Dynein is a 1.4 MDa homodimeric complex of two heavy chains, which each bind smaller subunits, including the intermediate chain, light intermediate chain, and three light chains. Dynactin is a ~1.0 MDa complex of more than 20 proteins, including an actin filament-like core of actin-related protein 1 (Arp1) subunits, capped at both ends by several subunits, and a “shoulder” domain from which emerges the largest subunit, p150Glued, which projects ~50 nm and can bind microtubules directly to help initiate a processive run. Although dynein and dynactin bind directly to each other via the dynein intermediate chain and the dynactin p150Glued subunit (Karki and Holzbaur, 1995;

Morgan et al., 2011; Siglin et al., 2013; Vaughan and Vallee, 1995), they form a stable processive complex only in the presence of effector proteins, including Hook1/3, BICD2,

Spindly, FIP3, and NIN/NINL (McKenney et al., 2014; Olenick et al., 2016; Redwine et al., 2017; Schlager et al., 2014b; Schroeder and Vale, 2016; Splinter et al., 2012). These effectors are unrelated to each other in sequence and recruit different cargoes. Here we have demonstrated that independent of these differences, they all appear to interact with

91 the same region of the dynein LIC1 subunit, which we named Helix-1. The interaction involves the Hook domain in Hook-family effectors (left) or a coiled coil segment in CC1-

Box-containing effectors such as BICD and Spindly (right). We have proposed here that the LIC1-effector interaction may help stabilize the “parallel-heads” conformation thought to be necessary for dynein processivity (Zhang et al., 2017).

V. Materials and Methods

Proteins

The cDNA encoding for human Hook1 (UniProt: Q9UJC3-1) and Hook3 (UniProt:

Q86VS8-1) were purchased from Open Biosystems (Huntsville, AL). Constructs

Hook111–166, Hook111–238 and Hook111–443 were cloned between BamHI and SalI sites of vector pMAL-c2x (NEB, Ipswich, MA). All the primers used in cloning are listed in

Supplementary Table 1. Construct Hook11–239GCN4 was obtained by adding a 28-a.a.

GCN4 sequence (MKQLEDKVEELLSKNYHLENEVARLKKL) by overlapping primers, while respecting the coiled coil heptad register. The fusion construct was cloned as above. Constructs Hook31–143 and Hook31–160 were cloned between NotI and SalI sites of a modified pMAL-c2x (NEB) vector in which the Sac1 site after MBP residue N367 was replaced with a NotI site. Point mutations A138D and M140D in Hook31–160 were introduced using the QuikChange site-directed mutagenesis kit (Agilent Technologies,

Wilmington, DE). All the proteins were expressed in E. coli BL21 (DE3) cells (Invitrogen,

Carlsbad, CA), grown in Terrific Broth medium at 37°C until the OD600 reached a value of

1.5–2, followed by 16 h at 19°C in the presence of 0.25 mM isopropyl-β-D- thiogalactoside. Cells were collected by centrifugation, re-suspended in 20 mM Tris pH

7.0, 100 mM NaCl, 4 mM benzamidine hydrochloride, 1 mM PMSF and 1 mM DTT and lysed using a Microfluidizer large-scale homogenizer (Microfluidics, Newton, MA). All the proteins were purified through an amylose affinity column according to the manufacturer’s protocol (NEB). The MBP tag was removed by incubation with TEV protease overnight at 4°C. All the proteins were additionally purified by gel filtration on a

SD200HL 26/60 column (GE Healthcare, Little Chalfont, UK) in 20 mM Tris pH 7.0, 100 mM NaCl, 1 mM DTT.

The cDNA encoding for human LIC1 (UniProt: Q9Y6G9-1) was a generous gift from Ronald Vale (UCSF). Constructs LIC1FL, LIC11–461 and LIC11–437 were amplified by

PCR and cloned between the BamHI and SalI sites of a modified vector pMAL-c2x that adds a C-terminal Strep-tag to the target protein. Point mutations F447A and F448A in

LIC1FL were introduced using the QuikChange site-directed mutagenesis kit. Proteins were expressed and purified as described above, with one exception; after amylose affinity purification, the proteins were loaded onto a StrepTactin Sepharose column (IBA

Lifesciences, Göttingen, Germany) and eluted after extensive washing with 3 mM desthiobiotin, 20 mM Tris pH 7.0, 100 mM NaCl, 1 mM DTT. To obtain the Helix-1 peptides, the cDNA encoding for LIC1433–458 (Fig. 3.2A) was cloned between the SapI and SalI sites of vector pTYB11 (NEB). Point mutations F447A and F448A were introduced using the QuikChange site-directed mutagenesis kit to obtain the mutant peptide Helix-1F447A,F448A. Proteins were expressed as above, and purified on a chitin affinity column according to the manufacturer’s protocol (NEB), followed by auto- cleavage of the intein tag induced by incubation with 50 mM DTT overnight at 4°C. The cleaved peptides were additionally purified on a Symmetry300 C18 reverse-phase column (Waters, Milford, MA) using an acetonitrile gradient of 0–90% (v/v) and 0.1%

(v/v) trifluoroacetic acid.

The cDNA encoding for full-length human Hook1 was codon optimized for expression in Sf9 cells and synthesized (Genscript Biotech, Piscataway, NJ). The gene was cloned between SalI and XbaI sites of a modified vector pFastBac1, which adds a

V5 epitope tag at the N-terminus and a Strep-tag at the C-terminal of the target protein.

The protein was expressed in Sf9 cells using the Bac-to-Bac baculovirus expression system according to the manufacturer’s protocol (Invitrogen). Cells were collected by centrifugation, re-suspended in lysis buffer (10 mM Na2HPO4 pH 7.4, 100 mM NaCl, 1 94 mM PMSF, 4 mM Benzamidine, 1 mM DTT, and 5% glycerol (v/v)) with addition of a protease inhibitor cocktail (Roche, Basel, Switzerland). Cells were lysed by addition of

0.5% (v/v) Triton X-100 through three cycles of freeze-thaw on ice and centrifuged 20 min at 20,000g. Lysates were loaded onto a StrepTactin Sepharose column, and after washing extensively with lysis buffer, Hook1FL was eluted with the addition of 3 mM desthiobiotin. The protein was additionally purified through a Superose 6 gel filtration column (GE Healthcare), equilibrated with 20 mM Tris pH 7.5, 100 mM NaCl, 1 mM DTT.

The cDNAs encoding for human BICD21–98 (UniProt: Q8TD16-1) and Spindly1–142

(UniProt: Q96EA4-1) were synthesized with codon optimization for E. coli expression

(Genscript Biotech). The BICD21–98 gene was cloned between the BamHI and SalI sites of vector pMAL-c2x. The protein was expressed and purified as described above. The

Spindly1–142 gene was cloned between the BamHI and SalI sites of vector pCold1

(TAKARA BIO, Kusatsu, Japan). The protein was expressed in BL21 (DE3) cells as described above. The protein was purified through a Ni-NTA affinity column in 50 mM Tris pH 8.0, 500 mM NaCl, 4 mM benzamidine hydrochloride, 1 mM PMSF and eluted with 250 mM Imidazole. The His6-tag was removed by overnight incubation with

TEV protease at 4°C, and the protein was additionally purified through a HiLoad 16/600

Superdex 75 pg column (GE Healthcare) in 20 mM Tris pH 7.0, 100 mM NaCl, and 1 mM DTT.

Multi-Angle Light Scattering

Samples were separated by size exclusion chromatography on a Superose 6

10/300 GL column (GE Healthcare) equilibrated with 20 mM Tris pH 7.5, 100 mM NaCl, and 1 mM DTT, using an Agilent 1100 HPLC system (Agilent Technologies). MALS was measured in line using a DAWN-HELEOS multi-angle light scattering detector and an

Optilab rEX refractive index detector. The scattering data were analyzed with the ASTRA software (Wyatt Technology, Santa Barbara, California).

Isothermal Titration Calorimetry

ITC measurements were carried out on a VP-ITC instrument (MicroCal,

Northampton, MA). Protein samples were dialyzed for 2 days against 20 mM HEPES pH

7.5, 100 mM NaCl, and 0.25 mM TCEP (ITC buffer). The LIC1433–458 peptide was resuspended in ITC buffer, followed by three cycles of lyophilization/resolubilization in

50% (v/v) methanol to remove any trifluoroacetic acid remaining after reverse-phase purification. The peptide was then resuspended in ITC buffer. The proteins (or LIC1433–

458 peptide) in the syringe were titrated at a concentration 10- to 20- fold higher than that of the proteins in the ITC cell of total volume 1.44 ml (as indicated in the figures).

Titrations consisted of 10 µl injections, lasting for 10 s, with an interval of 300–400 s between injections. The heat of binding was corrected for the heat of injection, determined by injecting proteins into buffer. Data were analyzed using the program

Origin (OriginLab, Northampton, MA). The temperature and parameters of the fit

(stoichiometry and affinity) of each experiment are given in the figures.

Rotary shadowing and electron microscopy

Full-length Hook1 was suspended in a solution containing 10 mM Tris pH 7.5, 50 mM NaCl, 1 mM DTT, and 50 % (v/v) glycerol. Samples were diluted to a concentration of 100 µg ml-1 in the same solution, and 1 µg of each sample were sprayed on a freshly split mica surface, dried for 1 h at room temperature. The samples were rotary shadowed with platinum at a 7° angle and replicated with carbon in a Balzers 410 freeze-fracture machine. Replicas were photographed at a magnification of 98,900 using a Philips 410 transmission electron microscope operating at 80 kV. The original images were obtained from areas at the edge of each droplet that showed distinct non- 96 aggregated molecules and a clear background. Images were analyzed using the ImageJ software (Schneider et al., 2012).

Crystallization, data collection and structure determination

-1 Hook31–160 at 10 mg ml in 10 mM Tris pH 7.4, 25 mM NaCl and 2 mM TCEP was mixed with 1.2 molar excess of the LIC1433–458 peptide at 4°C for 1 h. Crystal were obtained at 20°C using the hanging drop method. The crystallization drop consisted of a

1:1 (v/v) mixture of protein solution and well solution (1.44 M ammonium citrate tribasic, pH 6.25). Crystals were improved through consecutive rounds of micro-seeding. For data collection, crystals were flash frozen in liquid nitrogen from a cryo-solution consisting of crystallization buffer with addition of 30% (v/v) glycerol.

An x-ray diffraction dataset was collected at the Cornell High Energy Synchrotron

Source (CHESS) beamline F1. The diffraction data were indexed and scaled using the program HKL2000 (Otwinowski and Minor, 1997). A molecular replacement solution was obtained with the program Phenix (Adams et al., 2010) using PDB entry 5J8E (unbound

Hook domain of Hook3 (Schroeder and Vale, 2016)). Model building and refinement were carried out with the programs Coot and Phenix (Adams et al., 2010; Emsley et al.,

2010). Figures were generated with the program PyMOL (Schrödinger, New York City,

NY). Sequence alignments were carried out with the program MAFFT (Katoh et al.,

2002) and visualized using ESPript (Robert and Gouet, 2014). Data collection and refinement statistics are listed in Table 1.

Single molecule motility assays

The motility of dynein-dynactin-Hook31–552 or dynein-dynactin-BICD21–572 complexes from cell extracts were tracked using total internal reflection fluorescence

(TIRF) microscopy (Olenick et al., 2016). The motility assays were performed in flow chambers constructed with a glass slide and a silanized (PlusOne Repel Silane, GE 97

Healthcare) coverslip, held together with double sided adhesive tape and vacuum grease to form a ~15 µl chamber. A 1:40 dilution of monoclonal anti-β-tubulin antibody

(T5201, Sigma) was perfused into the chamber, which was subsequently blocked with

5% pluronic F-127 (Sigma-Aldrich, St. Louis, MO). Taxol-stabilized microtubules, labeled with HiLyte 488 or 647 (Cytoskeleton, Denver, CO) at a labeling ratio of 1:40, were flowed into the chamber and immobilized by interaction with anti-β-tubulin antibodies.

HeLa cells expressing Halo-tagged Hook31–552 or BICD21–572 were labeled with

TMR-HaloTag ligand (Promega, Madison, WI) 18–20 h post-transfection. Cells grown in

10 cm plates to 70–80% confluence were then lysed in 100 µl lysis buffer (40 mM

HEPES pH 7.4, 120 mM NaCl, 1 mM EDTA, 1 mM Mg-ATP, 0.1% Triton X-100, 1 mM

PMSF, 0.01 mg ml-1 TAME, 0.01 mg ml-1 leupeptin and 1 µg ml-1 pepistatin-A). Cell lysates were then clarified by centrifugation at 17,000g. Before flowing into the imaging chamber, the cell extracts were diluted in P12 buffer (12 mM PIPES pH 6.8, 1 mM

EGTA, 2 mM MgCl2, and 20 µM Taxol). Cells lysates were then further diluted in motility buffer (1x P12 buffer supplemented with 10 mM Mg-ATP, 0.3 mg ml-1 casein, 0.3 mg ml-1 bovine serum albumin and 10 mM DTT) and an oxygen scavenging system (0.5 mg ml-1 glucose oxidase, 470 U ml-1 catalase and 15 mg ml-1 glucose) and flowed into the chamber to be imaged.

The dynein-driven motility of single Halo-Hook31–552 or Halo-BICD21–572 positive molecules was then examined in the absence or the presence of varying concentrations of the Helix-1 (or Helix-1F447A,F448A) peptide, added to the motility buffer immediately prior to the addition of the cell lysates. All the movies (4 frames s-1) were acquired at room temperature using a Nikon TIRF system (Perkin Elmer, Waltham, MA) on an inverted Ti microscope equipped with a 100x objective and an ImageEM C9100-13 camera

(Hamamatsu Photonics, Hamamatsu City, Japan) controlled by Volocity software 98

(Improvision, Lexington, MA). Particle tracking was performed using the TrackMate plugin in the program Fiji (Schindelin et al., 2012). Particle runs were tracked only if both the start and end of a run were observable over the course of the movie. Runs on microtubule bundles were excluded from this analysis.

Lysosomal distribution analysis

HeLa cells transfected for 18–22 h with GFP, LIC1WT-GFP or LIC1F447A,F448A-GFP were fixed with PFA. Cells were then stained with anti-LAMP1 antibodies (rabbit polyclonal, ab24170, Abcam, Cambridge, UK) diluted 1:500, followed by anti-rabbit secondary antibodies, and mounted on glass coverslips with ProLong Gold anti-fade reagent (Invitrogen). Images were taken with an inverted epifluorescence microscope

(DMI6000B, Leica Camera AG, Wetzlar, Germany) using an Apochromat 63x, 1.4 NA oil immersion objective (Leica Camera AG).

Images were blinded before analysis, and the distribution of LAMP1-positive vesicles was scored using cells expressing GFP only. LAMP1 staining was scored

“abnormal” if the cells lacked the characteristic perinuclear lysosomal clustering and/or had several or a few enlarged LAMP1-positive vesicles dispersed throughout the cell.

Data availability

Atomic coordinates and structure factor amplitudes for the crystal structure of the

Hook3-LIC1 complex were deposited with the Protein Data Bank (PDB) under accession code 6B9H. Other data and materials are available from the corresponding author upon request.

Figure 3.S3 Table of primers used in this study

100

CHAPTER 4: Dynein activator Hook1 is required for trafficking of BDNF-signaling

endosomes in neurons

This chapter is adapted from:

Olenick, M. A., Dominguez, R., Holzbaur, E.L.F. (2018) Dynein activator Hook1 is required for trafficking of BDNF-signaling endosomes in neurons. Journal of Cell Biology, in revision.

101

I. Summary

Axonal transport is required for neuronal development and survival. Transport from the axon to the soma is driven by the molecular motor dynein, yet it remains unclear how dynein is spatially and temporally regulated for all retrograde cargo. We find that the dynein effector Hook1 mediates transport of TrkB-BDNF signaling endosomes in primary hippocampal neurons. Hook1 co-migrates with a subpopulation of Rab5 endosomes which exhibit retrograde motility with faster velocities than the overall Rab5 population. Knockdown of Hook1 did not affect the overall motility of Rab5, but significantly reduced the motility of BDNF-signaling endosomes. In microfluidic chambers, Hook1 depletion resulted in a significant decrease in the flux and processivity of BDNF-Qdots along the mid-axon, an effect specific for Hook1 but not Hook3. Further,

Hook1 knockdown inhibited downstream BDNF-TrkB signaling to the nucleus. Together, these studies support a model in which differential association with cargo-specific effectors efficiently regulates dynein in neurons.

102

II. Introduction

Axonal transport is vital for the maintenance and survival of neurons. Axons have a uniformly polarized microtubule array, in which the faster-growing plus-ends of microtubules are oriented toward the distal terminal. These microtubules serve as a highway for fast organelle trafficking, mediated by molecular motors. While multiple plus- end directed kinesins are responsible for delivery of cargo to the distal end of the axon, the minus-end directed motor cytoplasmic dynein is solely responsible for trafficking a wide variety of cargo back to the soma, including autophagosomes, endosomes, and mitochondria (Maday et al., 2014). These organelles not only differ in their lipid and protein compositions, but they also display distinct motility properties. It remains unclear how cytoplasmic dynein attaches to each of its cargos and how the motor is regulated to facilitate the precise trafficking of organelles to the soma.

Cytoplasmic dynein 1 (referred to here as dynein) is a 1.4 MDa AAA+ motor complex that drives the majority of minus-end directed motility in the cell. Alone, dynein is a flexible dimer with low processivity, taking many sideways or backwards steps along the microtubule lattice (Reck-Peterson et al. 2006; Ross et al. 2006). Processive motility is enhanced when dynein binds to dynactin, a 1 MDa multi-subunit complex that reorients the dynein dimer for proper recruitment and motility along microtubules (Ayloo et al., 2014; Zhang et al., 2017). While dynactin has been suggested to play a role in cargo interaction (Yeh et al., 2012; Zhang et al., 2011), adaptor and scaffolding proteins are required to link cargo to the dynein-dynactin motor complex (Fu and Holzbaur, 2014;

Kardon and Vale, 2009). Recently, a set of coiled coil effector proteins including BICD2,

Hook1/3, Spindly, FIP3 and NINL have been shown to enhance the dynein-dynactin interaction and induce superprocessive motility (McKenney et al., 2014; Olenick et al.,

2016; Redwine et al., 2017; Schlager et al., 2014a; Schroeder and Vale, 2016). BICD2, 103 the best characterized of these dynein effectors, has been shown to increase the affinity of dynein-dynactin interaction through coiled-coil contacts along the Arp1 filament that forms the core of dynactin (Chowdhury et al., 2015; Urnavicius et al., 2015). BICD2 also interacts with the N-terminal tail of the dynein heavy chain (Chowdhury et al., 2015;

Urnavicius et al., 2015) and the dynein light intermediate chain 1 (LIC1) (Lee et al.,

2018; Schroeder et al., 2014), leading to a stabilization of the dynein-dynactin-effector complex. Some dynein effectors can recruit two dynein dimers to a single dynactin, which further enhances the force and velocity of the motor complex (Grotjahn et al.,

2018; Urnavicius et al., 2018).

Another set of dynein effectors, Hook proteins (HookA or Hok1), were first characterized in filamentous fungi and shown to link dynein to early endosomes (Bielska et al., 2014; Zhang et al., 2014). In mammalian cells, three highly conserved Hook proteins are expressed: Hook1, Hook2, and Hook3. These proteins are characterized by an N-terminal Hook domain which binds LIC1 of dynein (Lee et al., 2018; Schroeder and

Vale, 2016). The Hook domain is followed by a central coiled-coil region and a less well- conserved C-terminal cargo binding domain (Bielska et al., 2014; Zhang et al., 2014). In vitro studies show that the binding of either Hook1 or Hook3 enhances the dynein- dynactin interaction leading to significant increases in velocity and run lengths

(McKenney et al., 2014; Olenick et al., 2016; Schroeder and Vale, 2016).

While Hook1 and Hook3 have been identified as dynein activators in vitro, the role of these proteins in dynein-mediated cargo transport in mammalian cells is less clear. Hook2 has been linked to centrosomal function and homeostasis (Guthrie et al.,

2009; Moynihan et al., 2009; Szebenyi et al., 2007), while Hook1 and Hook3 have been implicated in a variety of endosomal trafficking pathways, although there is still no clear consensus on the specific roles of each isoform (Luiro et al., 2004; Maldonado-Báez et 104 al., 2013; Xu et al., 2008). The highly polarized nature and spatial compartmentalization of neurons provide an excellent system to study the role of Hook proteins in endosomal transport. Initial work from Guo et al. (2016) suggested that Hook1 and Hook3 co- localize with retrograde Rab5a vesicles in hippocampal neurons and that knockdown of

Hook1 and Hook3 reduced the retrieval of transferrin receptor from the axon (Guo et al.

2016). These data support a potential role for Hook proteins in dynein-mediated trafficking in axons, prompting us to investigate this question in more detail.

In this study, we investigated the role of Hook1 in the dynein-driven transport of endosomes along the axons of hippocampal neurons. We found that Hook1 co-migrates with subpopulations of Rab5- and Rab7-positive endosomes. While loss of Hook1 did not significantly change the overall motility of Rab5- or Rab7-positive endosomes, Hook1 siRNA depletion significantly reduced the motility of a specific endosomal compartment that we identified as TrkB-BDNF signaling endosomes. The motility of TrkB-BDNF signaling endosomes is also lost if the interaction of Hook1 with dynein is disrupted by targeted mutations at the Hook1-LIC1 interface. In addition, Hook1 is enriched in the distal axon, distinct from the cellular distribution of other dynein effectors like RILP or

BICD2, suggesting a specific function in trafficking from the distal axon. Using microfluidic chambers to model the distal axonal transport of BDNF signaling endosomes, we found that knockdown of Hook1 significantly reduced the flux and the processivity of BDNF transport from the distal axon to the soma. In contrast, knockdown of Hook3 did not affect BDNF uptake or transport. Loss of Hook1 also produced a functional block in downstream BDNF-dependent signaling to the nucleus, which is vital for neuronal survival and maintenance. Overall, this work supports a model in which

Hook1 acts as a specific dynein effector for BDNF-signaling endosomes trafficking from

105 the distal axon to induce downstream signaling in the soma of primary hippocampal neurons.

106

III. Results

Hook1 co-migrates with the endosomal markers Rab5 and Rab7

To investigate the role of Hook1 in endosomal trafficking, we first expressed the early endosome marker Rab5-GFP or the late endosome marker Rab7-GFP, in primary rat hippocampal neurons to observe endosomal motility. Neurons were imaged 7-8 days in vitro (DIV) at 48-hours post-transfection using live cell confocal microscopy. Focusing on the axon, Rab5-GFP endosomes were found to be enriched in the distal axon, while

Rab7-GFP endosomes were found throughout the distal and mid-axon. In the axon,

Rab5-GFP endosomes were mainly stationary or bidirectional, with about 80% moving less than 10 microns in any net direction (Fig. 4.S1B). In contrast, 50% of Rab7-GFP endosomes displayed net retrograde motility (Fig. 4.S1E).

Next, Hook1-Halo was co-expressed with Rab5-GFP or Rab7-GFP to assess whether Hook1 co-migrates with a specific population of endosomes, since previous reports had found Hook proteins on Rab5-positive endosomes (Bielska et al., 2014; Guo et al., 2016; Zhang et al., 2014). Surprisingly, we observed co-migration of Hook1 with a sub-population not only of Rab5 endosomes but also a sub-population of Rab7 endosomes (Fig. 4.1A,B). Previous in vitro studies indicate that Hook1 enhances the processivity of dynein (Olenick et al. 2016), so we asked whether Hook1-positive endosomes displayed distinctive retrograde bias in their motility or were significantly more processive than the overall population of axonal endosomes. We found that

Hook1-positive Rab5 endosomes had an increased retrograde bias, as seen by an increased retrograde: anterograde ratio as compared to neurons expressing Rab5-GFP only (Fig. 4.1C,D). Hook1-positive Rab5 endosomes also showed an increase in retrograde-directed instantaneous velocity as compared to control Rab5-GFP neurons

(Fig. 4.1E). In comparison, Hook1-positive Rab7 endosomes had the same retrograde 107 bias and retrograde instantaneous velocity as seen in neurons expressing Rab7-GFP alone, where approximately 50% of organelles displayed fast, retrograde motility (Fig.

4.1F-H). Overall, this analysis indicates that motile endosomes positive for Hook1 were primarily fast, retrogradely moving organelles, indicative of processive dynein-mediated motility.

Figure 4.1 Hook1 co-migrates with Rab5- and Rab7-endosomes

A) Mid-axons of hippocampal neurons expressing Hook1-Halo and Rab5- or Rab7-GFP.

White arrows show co-localized Hook1 with indicated Rab-endosome. Scale= 4µm. B) 108

The percentage of endosomes with Hook1-colocalization. Scatter plot with mean ±

S.E.M. N: Rab5=17, Rab7=10 neurons. C) Kymograph of Rab5-GFP and Hook1-Halo motility in axon of hippocampal neuron. White arrows show co-migrating Hook1-Rab5 organelle. Scale= 4 µm, 20 sec. D) The ratio of Retrograde to Anterograde motility events in neurons expressing Rab5-GFP+Hook1-Halo or Rab5-GFP only. N: Rab5=19,

Rab5+Hk1=17 neurons. E) Cumulative histogram of retrograde instantaneous velocities of events in neurons expressing Rab5-GFP+Hook1-Halo or Rab5-GFP only. F)

Kymograph of Rab7-GFP and Hook1-Halo motility in axon of hippocampal neuron. White arrows show co-migrating Hook1-Rab7 organelle. Scale=4 µm, 20 sec. G) The ratio of retrograde to anterograde motility events in neurons expressing Rab7-GFP+Hook1-Halo or Rab7-GFP only. N: Rab7=18, Rab7+Hk1=9 neurons. H) Cumulative histogram of retrograde instantaneous velocities of events in neurons expressing Rab7-GFP+Hook1-

Halo or Rab7-GFP only.

Hook1 knockdown reduces the motility of BDNF-positive signaling endosomes

As Hook1 co-migrates with a subpopulation of Rab5- and Rab7-positive endosomes, we next asked if Hook1 depletion would alter the motility of these endosomes. We used a rat siRNA pool to knockdown Hook1 in cultured neurons; 60% depletion of Hook1 was observed in PC12 cells using this approach (Fig. 4.S1G,H).

Rab5-GFP or Rab7-GFP were transfected along with Hook1 siRNA and imaged 48-hour post transfection at DIV7-8 along the mid- or distal regions of the axon. Hook1 KD did not induce significant differences in the motile fraction or the directionality of either

Rab5- or Rab7-positive endosomes at a population level (Fig. 4.S1A-F). Hook1 KD did induce a trend toward decreased flux of Rab5 endosomes, but the effect was not significantly different than control (Fig. 4.S1C). 109

We hypothesized that Hook1 might play a more specific role in the transport of a subpopulation of neuronal endosomes, so we focused on signaling endosomes, which initiate distally and mature through the Rab5 and Rab7 endosomal pathway (Deinhardt et al., 2006; Ye et al., 2018). In neurons, neurotrophic factors such as BDNF bind to their transmembrane kinase receptors and are endocytosed to form signaling endosomes that undergo retrograde transport toward the nucleus, leading to changes in gene expression

(Cosker and Segal, 2014; Scott-Solomon and Kuruvilla, 2018). To focus on a signaling endosomal population in neurons, we expressed the neurotrophic receptor TrkB-RFP in control and Hook1 KD neurons. Upon Hook1 depletion, retrograde TrkB-endosomes displayed less processive motility as indicated with more pausing (66.6% retrograde events displayed pausing per neuron) than in control cells (32.2% retrograde events with pausing per neuron) (Fig. 4.2A).

To more directly measure the motility of signaling endosomes, we monitored the uptake and motility of the TrkB ligand, BDNF. First, we looked for co-localization of

BDNF with Hook1-Halo in neurons. Neurons were serum-starved for an hour and then

BDNF-biotin was added to neuronal cultures for at least an hour before fixation.

Approximately 48% of Hook1-Halo puncta co-localized with BDNF-Alexa633 in fixed neurons (Fig. 4.2B,C). Next, we investigated the motility of BDNF-conjugated quantum dots (BDNF-Qdots). BDNF-Qdot motility in axons was quantified as the percent of motile Qdots observed in a 2-minute video. Neurons with Hook1 KD displayed significantly reduced BDNF-Qdot motility compared to control neurons (Fig. 4.2D,E).

This motility defect could be rescued by expression of siRNA-resistant human Hook1-

Halo (Fig. 4.2D,E). As an additional control, we imaged mitochondrial motility in Hook1

KD neurons and found no differences in flux or morphology (Fig. 4.S1I). Together, these

110 results suggested Hook1 plays an important role in TrkB-BDNF signaling endosome motility in axons.

Figure 4.S1 Hook1 Knockdown does not significantly change Rab5 or Rab7 motility

A) Kymograph of Rab5-GFP in control or Hk1 KD neurons. Traced events below, color coded for ease of interpretation. Scale= 5 µm, 2 mins total. B) Motility fractioned into retrograde, anterograde, and non-motile events per neuron. Bar graph with mean ±

S.E.M. N: Rab5=23, Rab5+siHk1=25 neurons C) Flux of Rab5 organelles in control or

Hk1 KD neurons. Scatter plot with mean ± S.E.M. N: Rab5=23, Rab5+siHk1=25 neurons

D) Kymograph of Rab7-GFP in control or Hk1 KD neurons. Traced events below, color coded for ease of interpretation. Scale= 5 µm, 2 mins total. E) Motility fractioned into

111 retrograde, anterograde, and non-motile events per neuron. Bar graph with mean ±

S.E.M. N: Rab7=23, Rab7+siHk1=23 neurons. F) Flux of Rab7 organelles in control or

Hk1 KD neurons. Scatter plot with mean ± S.E.M. N: Rab7=23, Rab7+siHk1=23 neurons

G) Western blots of Hook1 and Hook3 siRNA KD in PC12 cells. H) Quantification of KD western blots, from three individual repeats. I) Kymographs of mitochondria in control and Hook1 KD cells. Scale= 5 µm.

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Figure 4.2 Hook1 KD reduces TrkB-BDNF signaling endosome motility

A) Kymographs of TrkB-RFP in control or Hk1 KD neurons. Traced events shown below, color coded for ease of interpretation. Black arrows show pausing in retrograde events.

Scale=10 µm, 1 min total. B) Colocalization images of BDNF-Alexa633 with Hook1-Halo in axon of hippocampal neuron. Arrows show Hook1 puncta co-localized with BDNF.

Scale bar=2 µm. C) Line scan through axon in image B. Arrows point to colocalized

Hook1 and BDNF. D) Quantification of BDNF motility. Bar graph with mean ± S.E.M.

Kruskal-Wallis one-way ANOVA. N: Mock=103, Hk1 KD=63, Hk1 KD+Hk1-Halo=55 neurons. E) Kymographs of BDNF-Qdots in control, Hk1 KD and Hk1 KD + Hk1-Halo

113 neurons. Traced events shown below, color coded for ease of interpretation. Scale=5

µm, 5 sec.

A direct interaction of Hook1 with dynein is important for signaling endosome motility

Dynein subunit LIC1 interacts with several dynein effectors including BICD2,

RILP, and FIP3 (Schroeder et al., 2014). Recent reports have found that the Hook domain in Hook1 and Hook3 also mediates a direct interaction with LIC1 and is important for Hook-mediated dynein processivity (Lee et al., 2018; Schroeder and Vale,

2016). To determine if the LIC1 interaction is important for signaling endosome motility in neurons, we analyzed two constructs: Hook1(Q149A, I156A) based on previous mutations in Hook3 shown to diminish the interaction with LIC1 (Schroeder and Vale,

2016), and Hook1(M146D, I156D), based on our recent structure of a Hook3-LIC1 complex (Lee et al., 2018) (Fig. 4.3A,B). We used TIRF microscopy to perform motility assays with single molecule resolution to test these mutant constructs and found that both Hook1(Q149A, I156A) and Hook1(M146D, I156D) significantly inhibited dynein- driven motility along microtubules (Fig. 4.3C,D). Next, we tested if mutating the binding interface of Hook1 and LIC1 would disrupt signaling endosome motility in hippocampal neurons. Similar to our results with Hook1 siRNA KD, BDNF-Qdot motility was significantly reduced in neurons expressing GFP-Hook1(Q149A, I156A) (Fig. 4.3E,F).

These results indicate that the interaction of Hook1 with LIC1 is not only important for in vitro motility but also during cargo transport of BDNF-signaling endosomes in neurons.

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Figure 4.3 Hook1 requires interaction with LIC1 for signaling endosome motility

A) Diagram of Hook1 domain structure with mutated constructs below. Red arrows indicate point mutations. B) Hook3 (yellow)-LIC1 helix (green) structure (PDB: 6B9H) with residues that were mutated highlighted in magenta (Hook1 residue numbering). C)

Kymographs of Halo-Hook1 constructs from single molecule TIRF motility assay. Arrows

115 indicate motile events. Scale bar= 5 µm and total length is 1 min. D) Quantification of

Hook1 motility in TIRF motility assay. Scatter plot with mean ± S.E.M. Kruskal-Wallis one-way ANOVA. N:14-15 videos, from three individual experiments E) Kymographs of

BDNF-Qdots in axons of hippocampal neurons. Traced events below, color coded for ease of interpretation. Scale= 5 µm, 5 sec. F) Quantification of BDNF motility. Bar graph with mean ± S.E.M., Mann-Whitney t-test. N: Mock= 31, Hk1(Q149A, I156A)= 46 neurons

Hook1 is enriched in the distal axon and is distinctly localized from other dynein effectors

There are now several dynein effectors implicated in the regulation of dynein- driven motility in cells (Fu & Holzbaur, 2014; Kardon & Vale, 2009). We hypothesized that the dynein effectors might display differential localization in neurons, reflecting distinct roles in intracellular transport. To address this question, we individually expressed Hook1-Halo, RILP-GFP, and BICD2-GFP in hippocampal neurons for 48- hours and then fixed the cells on DIV7-8. Confocal z-stacks were captured of the distal axon, mid-axon, and soma of neurons. Images were deconvolved, and the somal localization was scored as either punctate or cytoplasmic. Over 80% of BICD2- and

RILP-expressing neurons had an accumulation of these effectors in the soma, associated with either large puncta or clusters, while the distribution of Hook1 was predominantly cytoplasmic (Fig. 4.4A,B). In axons, dynein effectors were imaged either in the mid- or distal axon and puncta quantified per unit length, comparing levels along the same axon to assess relative enrichment of individual effectors. BICD2 had a generally sparse distribution in axons compared to RILP and Hook1. In contrast, RILP was enriched in the mid-axon and Hook1 was enriched in the distal axon (Fig. 4.4A,C). 116

The localization of these effectors is consistent with their proposed roles in organelle transport. BICD2 is linked to Rab6 vesicles (Matanis et al., 2002; Matsuto et al., 2015), which are mainly localized to the soma. RILP is a Rab7-adaptor (Cantalupo et al., 2001;

Johansson et al., 2007; Rocha et al., 2009; Wu et al., 2005), and Rab7 vesicles are enriched in the mid-axon and soma. In contrast, the enrichment of Hook1 in the distal axon suggests that this effector is involved early in the pathway of neurotrophic factor uptake and trafficking within signaling endosomes.

Figure 4.4 Hook1 localizes to distal axon, while other dynein effectors are enriched in other compartments

A) Representative images of neurons expressing indicated dynein effectors. Black arrows point to effector puncta. Scale bar=10 µm. B) Quantification of percentage of

117 neurons with puncta in soma. Bar graph with mean ± S.E.M., N= 3-4 individual experimental averages, 23-26 cells. C) Graphs of dynein effector enrichment in mid vs distal axons. Red lines indicate enrichment in distal region of axon. Black lines indicate enrichment in mid-axon. N: BICD2= 21, RILP= 27, Hook1= 27 neurons.

Hook1 depletion decreases the flux and processivity of BDNF-signaling endosomes from distal axons

Hook1 is enriched in the distal end of axons, similar to the enrichment of Rab5- early endosomes. Due to this localization, we asked whether Hook1 plays a role early in the transport of signaling endosomes from the distal axon. To better model the uptake and transport of BDNF from the distal axon, we utilized microfluidic devices in which axons grow through microchambers to the fluidly isolated axonal chamber, permitting

BDNF application only to the distal axons (Fig. 4.5A). Hippocampal neurons were electroporated with GFP fill and Hook1 siRNA prior to plating in the microfluidic devices.

Neurons were cultured for 7-8 days, allowing the axons to extend to the distal chamber.

Prior to imaging, BDNF-Qdots were added to the axonal chamber of the devices. We compared the motility properties of BDNF-Qdots in Hook1 KD and control neurons.

While BDNF-Qdots still exhibited a retrograde bias in motility upon Hook1 KD, the flux was greatly reduced compared to control conditions (Fig. 4.5B-D). In contrast, neurons transfected with Hook3 siRNA did not show this decrease in BDNF-Qdot flux (Fig. 4.5D).

In addition to decreased flux, we also noticed a difference in the size of BDNF-Qdot vesicles. Previous studies have shown that multiple neurotrophic factor-bound quantum dots may be internalized into a single signaling endosome (Cui et al., 2007). Thus, we measured the apparent size of BDNF-Qdots organelles as a measure of the number of internalized BDNF-Qdots. We found a significant reduction in organelle area in Hook1 118

KD neurons (0.39 µm2 with 0.30-0.48 95% CI) compared to control neurons (0.62 µm2 with 0.51-0.74 95% CI), suggesting fewer BDNF-Qdots are being endocytosed per vesicle. Overall, this work suggests that Hook1 depletion not only reduces the number of

BDNF-organelles trafficked down the axon but also reduces the load of individual organelles.

While significantly fewer BDNF-Qdots were observed to traffic along the axon in

Hook1-depleted neurons, we analyzed the motility properties of these organelles to see if loss of the dynein activator would reduce the processivity of signaling endosomes from the distal axon. In Hook1 KD cells, BDNF-Qdots showed less directed motility along the axon, with significantly more directional switching within individual runs (Fig. 4.5E). In addition, retrograde events also displayed increased pause duration and reduced net velocity (Fig. 4.5F,G). These results suggest that Hook1 does act as a dynein activator to increase the processivity of retrograde BDNF-signaling endosomes once they have been endocytosed.

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Figure 4.5 Hook1 KD reduces flux of BDNF from distal axon

A) Schematic of microfluidic (MF) device and experimental setup. B) Kymographs of

BDNF in MOCK, Hk1 KD, Hk3 KD neurons grown in MF device. Green arrows point to retrograde events. Scale= 10 µm, 10 sec. C) Motility fractioned into retrograde, anterograde and non-motile events per neuron. Bar graph with mean ± S.E.M., two-way

ANOVA. N: MOCK= 35, Hk1 KD= 34 neurons. D) Quantification of flux of BDNF-Qdots in mid-axons. Scatter plot with mean ± S.E.M, one-way ANOVA. N: MOCK= 54, Hk1

KD= 36, Hk3 KD= 29 neurons. E) Number of switches in BDNF-Qdot events. Scatter plot with mean ± S.E.M, Mann-Whitney t-test. N: MOCK= 110, Hk1 KD= 84 events. E) Pause duration of retrograde BDNF-Qdot events. Scatter plot with mean ± S.E.M, Mann-

Whitney t-test. N: MOCK= 57, Hk1 KD= 33 events. F) Net Velocity of retrograde BDNF-

Qdot events. Scatter plot with mean ± S.E.M, unpaired t-test. N: MOCK= 63, Hk1 KD=

34 events. 120

Since we observed reduced flux and impaired motility of signaling endosomes in

Hook1 KD neurons, we hypothesized there might be a resulting accumulation of BDNF-

Qdots at the distal ends of the axons due to loss of dynein transport (Figure 4.6A). We imaged the distal regions of axons after several washes to remove excess BDNF-Qdots.

Quantification of the area of BDNF-Qdot signal per unit area of axon showed no difference between Hook1 KD and control neurons (Fig. 4.6B,C). Similar results were seen when dynein motility was blocked with the dynein inhibitor Ciliobrevin D (Fig.

4.6B,C), suggesting there is down-regulation of TrkB endocytosis when motility is impaired, preventing distal accumulation. One way to reduce endocytosis of BDNF is to reduce the amount of its receptor TrkB at the plasma membrane. We measured the amount of TrkB on the surface of Hook1 KD or control distal axons. To look at the plasma membrane-associated TrkB, we used an antibody against a region of the TrkB extracellular domain, which is not conserved in other Trk proteins. After fixation, neurons were stained with anti-TrkB (aa 54-67) without permeabilization. There was no significant change in surface TrkB levels at the axon tips with Hook1 depletion, suggesting that a reduction in TrkB levels is not contributing to the reduced flux we observed with Hook1 KD (Fig. 6D,E). Together, these results suggest that endocytosis of BDNF is downregulated when dynein motility is impaired, which may constitute a potential mechanism to reduce the distal accumulation of cargos in axons.

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Figure 4.6 Impaired signaling endosome motility reduces BDNF endocytosis

A) Schematic model of two possible effects of Hook1 KD on distal axons. B)

Representative images of distal tips of axons with BDNF-Qdots. White arrows indicate

BDNF-Qdots, white dashed line is cell outline. Scale bar= 5 µm. C) Area of Qdots in the distal axons normalized by area of axon quantified. Scatter plot with mean ± S.E.M.,

Kruskal-Wallis one-way ANOVA. N: MOCK= 58, Hk1 KD= 46, CilioD=3 6 neurons. D)

Surface TrkB staining with anti-TrkB in distal axon of fixed neurons. Bottom panel (Anti-

Rabbit594) is secondary antibody only, as a control. Scale bar= 5 µm. E) Area of surface

TrkB per a micron of axonal length. Anti-Rabbit 594 condition is control for secondary antibody. Scatter plot with mean ± S.E.M., one-way ANOVA. N: Mock=40, Hk1 KD=32,

Anti-Rabbit594=10 neurons

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Hook1 KD reduces downstream BDNF-signaling to the nucleus

BDNF binds TrkB, which then recruits signaling kinases to produce transcriptional changes in the nucleus (Cosker and Segal, 2014; Mitre et al., 2017). The transport of signaling endosomes is important to produce the downstream signaling to the nucleus (Heerssen et al., 2004; Ye et al., 2003). We wondered if loss of Hook1 reduced downstream signaling to the nucleus due to reduced flux of BDNF. To measure downstream signaling, we monitored phosphorylated CREB levels in the nucleus, which has been previously shown to increase after treatment with BDNF (Watson et al., 2001).

Hook1 KD and control neurons were grown in culture for 7 days in microfluidic devices, then treated with 1nM BDNF for one hour before being fixed and stained with anti- pCREB (Ser133) antibody. Using epifluorescence microscopy, we imaged neurons with axons that grew through the microchannels to reach the axonal compartment. In control cells, BDNF-treated neurons had increased nuclear pCREB compared to non-treated

(NT) control neurons (Fig. 4.7A-C). In Hook1 KD neurons, BDNF treated cells did not show increased pCREB staining compared to non-treated cells (Fig. 4.7A-C). These results indicate that the reduced flux of BDNF impairs downstream signaling to the nucleus in Hook1-deficient neurons.

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Figure 4.7 Loss of Hook1 leads to loss of downstream signaling, measured by pCREB levels

A) Representative images of soma with pCREB staining. Scale bar= 10 µm. B)

Qualification of nuclear pCREB signaling. Scatter plot with mean ± S.E.M., one-way

ANOVA. N: MOCK, NT= 42, MOCK, +BDNF= 48, Hk1 KD, NT= 51, Hk1 KD, +BDNF=

124

55 somas. C) Normalized pCREB intensity to the non-treated MOCK condition per individual experiment. Bar graph with mean ± S.E.M, RM one-way ANOVA. N: three individual experiments. D) Model of neurotrophin uptake and transport from distal axon.

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IV. Discussion

Here, we found that Hook1 co-migrates with a subpopulation of Rab5 and Rab7 endosomes. Previously Hook proteins in fungal systems had been linked to early endosomal transport marked by Rab5, but these systems only express one Hook isoform (Bielska et al., 2014; Zhang et al., 2014). In mammalian systems, Hook1 has been linked to different aspects of the endosomal pathway. In HeLa cells, Hook proteins were found to interact with members of the HOPS complex and to be important for timely trafficking of EGF through endosomal compartments marked by EEA1, CD63 and

LAMP1, but this study simultaneously knocked down all three Hook isoforms, making it difficult to determine their individual roles (Xu et al., 2008). Another study using HeLa cells suggested that Hook1 interacts with CD147 to facilitate sorting into Rab22-positive recycling tubules (Maldonado-Báez et al., 2013). In COS-1 cells, Hook1 was found to interact with Rab7, Rab9, and Rab11 using immunoprecipitation (Luiro et al., 2004). The variety of results seen in these studies is likely due to the fluid nature of endosomal pathways and differential cellular demands on these pathways. In our work, Hook1 co- migrates primarily with fast, retrograde Rab5-positive vesicles in primary neurons, which suggests a role for Hook1 in activating the motility of these vesicles. Yet, Hook1 depletion produced only subtle effects on the dynamics of the axonal Rab5-positive endosomal population, suggesting a higher level of specificity than previously observed for dynein effectors that interact directly with Rab proteins, such as the interaction of

BICD proteins with Rab6 (Huynh and Vale, 2017; Matsuto et al., 2015; Schlager et al.,

2010; Terawaki et al., 2015) or RILP with Rab7 (Cantalupo et al., 2001; Johansson et al., 2007; Wu et al., 2005). Instead of cargo attachment through a Rab protein, Hook1 has been suggested to attach to cargo through C-terminal interactions with Fused Toes

(FTS) and FTS-Hook Interacting (FHIP) proteins (Guo et al., 2016; Xu et al., 2008; Yao 126 et al., 2014). It remains to be determined if Hook1 is linked to signaling endosomes by

FTS/FHIP or another protein complex in neuronal systems.

In this work, we found that Hook1 plays a role in signaling endosome processivity. Our previous work has shown that Hook1 increases dynein-dynactin processivity in vitro, with Hook1-bound dynein displaying higher velocities and longer run lengths than BICD2-associated motors (Olenick et al., 2016). Recent cryo-EM structures have shown Hook3 can recruit two dynein dimers per one dynactin complex (Urnavicius et al., 2018). Due to the high sequence similarity in the Hook domain and coiled-coil regions, it is likely that Hook1 functions in a similar manner. It is possible that the relatively high velocities observed for signaling endosome transport (averaging 1.4

µm/sec) are due to the incorporation of two dynein dimers into the dynein-dynactin-

Hook1 complex.

We also found that the interaction of Hook1 with dynein subunit LIC1 is essential for signaling endosome motility, a mechanism that is likely conserved in other dynein effectors regulating cargo transport in the cell. Recent structural work showed that a helix (aa 433-458) within the otherwise unstructured C-terminal region of LIC1 is a conserved interface for the binding of dynein effectors, including BICD2, Spindly and

Hook proteins (Lee et al., 2018). Since the LIC1 interaction region is conserved among dynein effectors, it is likely that competition for this binding site plays a role in regulating cargo transport. During our investigation of Hook1, we observed differential localization of the dynein effectors Hook1, BICD2, and RILP in hippocampal neurons. It is possible that the compartmentalization of neurons and the differential localization of dynein effectors locally regulates the competition of dynein effectors for dynein-dynactin binding sites.

127

In this study, we found that Hook1 plays a key role early in the transport of TrkB-

BDNF endosomes as reflected by Hook1 enrichment in the distal axon. This enrichment is not seen for RILP or BICD2. RILP was enriched in the mid-axon and soma, which is consistent with its role in mediating Rab7-endosomes motility (Cantalupo et al., 2001;

Johansson et al., 2007). It remains to be investigated if there is a transition or hand-off of dynein effectors as endosomal maturation occurs, with the removal of Rab5 and accumulation of Rab7. It is also possible that a given cargo could have a mixture of dynein effectors. While Hook and BICD proteins are dynein effectors that modulate dynein processivity, there is as yet no direct evidence of RILP acting as a dynein activator but only as an adaptor to recruit dynein to Rab7 cargo (Johansson et al., 2007;

Rocha et al., 2009). Thus, it is possible that cargo might recruit a dynein activator to induce greater force or velocity, along with dynein adaptors to increase motor binding sites on the organelle. It remains to be seen if there is coordination between dynein activators and adaptors to help maintain a constant processive dynein pool on a given organelle.

Using microfluidic devices, we showed that loss of Hook1 reduces the flux and processivity of BDNF signaling endosomes. However, this reduced flux to the cell body did not result in an accumulation of BDNF-Qdots in the distal axon. Inhibiting dynein motility with CiliobrevinD also showed similar BDNF-Qdots levels in the distal axon, suggesting that endocytosis is downregulated when endosomal motility is inhibited.

BDNF has been reported as a self-amplifying autocrine factor, which can signal to promote BDNF expression and increase TrkB membrane levels (Cheng et al., 2011).

Therefore, it seems likely that the loss of BDNF signaling could reduce TrkB surface levels and regulate endocytosis to prevent a buildup of BDNF endosomes at the distal tip. However, we detected no significant change in plasma membrane-associated TrkB 128 levels at the axon tip upon Hook1 depletion, suggesting that another mechanism might be at work. It is possible that the internalization of the TrkB-BDNF complex via endocytosis is tightly linked to the formation of a high-speed, highly processive Hook1- dependent transport compartment. Thus, if transport is blocked either by Hook1 depletion or dynein inhibition, internalization may also be down-regulated, preventing the distal accumulation of stalled signaling endosomes. Hook1 binding partners such as

Fused Toes (FTS) and FTS-Hook Interacting (FHIP) proteins (Guo et al., 2016; Xu et al.,

2008; Yao et al., 2014) may mediate this coordination of uptake and transport, an interesting question for future studies.

TrkB-BDNF signaling is important for neuronal survival, and disruption of signaling endosome trafficking has been found in models of neurodegenerative diseases including Huntington’s and Parkinson’s disease (Millecamps and Julien 2013). In

Huntington’s disease, the polyQ expanded huntingtin protein has been shown to impair

BDNF retrograde trafficking, leading to reduced neuronal survival (Gauthier et al., 2004).

Alpha-synuclein has also been shown to impair BDNF transport in a mouse model of

Parkinson’s disease (Fang et al., 2017). Currently, it is unclear how Hook proteins play a role in these neurodegenerative diseases, but Hook1 and Hook3 have been localized to tau aggregates, a pathological hallmark of Alzheimer’s disease, frontotemporal dementia, and other tauopathies (Herrmann et al., 2015). With our new understanding of

Hook1 as a dynein effector for BDNF transport in non-pathological states, the role of

Hook1 in disease states remains to be investigated in future studies.

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V. Materials and Methods

Plasmids and Reagents

Halo-tagged Hook1 constructs were generated from the human Hook1 sequence

(Uniprot code Q9UJC3) and using the HaloTag from the pHTN or pHTC Halo tag CMV- neo vector (Promega). GFP-Hook1 constructs were generated in the pEGFP vector.

Rab5-GFP was provided by M. Zerial (Max Planck Institute). Rab7-GFP was purchased from Addgene. TrkB-mRFP was provided by M. Chao (New York University). RILP-GFP was provided by J. Neefjes (Leiden University Medical Center). BICD2-GFP was provided by A. Akhmanova (Utrecht University). Empty pEGFP-N1 (Addgene) was used as a cell fill to identify neuronal morphology.

Antibodies used for biochemistry and Western blotting include: anti-Hook1(rabbit,

1:250, AbCam ab104514), anti-Hook3(rabbit, 1:1000, ProteinTech), anti-actin (mouse,

1:1000; Millipore). For IF experiments, anti-pCREB(Ser133) (rabbit, 1:1000; Cell

Signaling) and anti-TrkB (aa 55-67, rabbit, 1:1000; Millipore). For TIRF assays, monoclonal antibodies used were anti-β-tubulin (1:40, mouse; Sigma T5201). ON-

TARGETplus siRNA SMARTpool of 4 siRNAs for rat Hook1 or Hook3 was purchased from Dharmacon (GE LifeSciences).

Neuronal Culture

Embryonic day (E) 18 Sprague Dawley rat hippocampal neurons were obtained in suspension from the Neuron Culture Service Center at the University of Pennsylvania and plated on 35-mm glass-bottom dishes (MatTek) or 25mm coverslips (World

Precision). Dishes or coverslips were pre-coated with 0.5 mg/mL poly-L-lysine (Sigma)

24 hours prior to plating. Neurons were cultured at 37C with 5% CO2 in maintenance media that consisted of Neurobasal (Gibco) supplemented with 2mM GlutaMAX,

100units/mL penicillin, 100 mg/mL streptomycin and 2% B27 (ThermoFisher Scientific). 130

Every 3-4 days, 40% of the media was replaced with fresh maintenance media supplemented with 1 μM AraC.

Neuronal Imaging

Imaging was done at 7–8 DIV (days in vitro) with neurons transfected 24-48 hr before imaging. Neurons were transfected using Lipofectamine 2000 (Invitrogen) according to manufacturer’s instructions, with 0.3-1 µg of each DNA plasmid and for siRNA conditions, 45 pmol of siRNA. Control siRNA labeled with Cy5 was used to confirm transfection of siRNA with lipofectamine in cultured neurons. Neurons were imaged in low-fluorescence media (HibernateE, Brain Bits) supplemented with 2% B27 and 1% GlutaMax. For experiments with HaloTag constructs, neurons were labeled with

TMR HaloTag ligands according to manufacturer’s instructions (Promega). For mitochondria imaging, TMRE was added according to manufacturer’s protocol. Neurons were imaged in an environmental chamber at 37C on a spinning-disk confocal UltraView

VOX (Perkin Elmer) on an inverted Nikon Eclipase Ti microscope with the Prefect Focus system using apochromat 100 × 1.49 NA oil-immersion objective and a C9100-50

EMCCD camera (Hamamatsu) controlled by Volocity software (Perkin Elmer). Axons and dendrites were identified based on morphologic criteria as outlined in (Kaech and

Banker, 2006).

Microfluidic experiments

Round microfluidic devices of 450 um (Xona microfluidics) were used for axon isolation experiments. Devices were UV sterilized and attached to PLL coated imaging dishes (FluoroDish; World Precision Instruments) prior to plating. On day of plating, neurons were nucleofected with an Amaxa Neuclofector machine (Lonza) with

DNA/siRNA in similar quantities as described above. Cells were plated to one side of

131 microfluidic device at 4x105 cells per dish. Fresh maintenance media was added every 2 days (about 30%).

BDNF-Qdot experiments

Neurons were serum starved in unsupplemented Neurobasal (Gibco) for 2-

4hours before BDNF-Qdot addition. 50 nM hBDNF-biotin (Alomone Labs) was combined with 50 nM Quantum Dot ITK 655 Streptavidin conjugate (Invitrogen) for 1 hour on ice to generate BDNF-Qdots. After conjugation, BDNF-Qdots were added to neurons in unsupplemented Neurobasal to a final concentration of 0.25 nM for 1-2 hours. In microfluidic experiments, BDNF-Qdots was only added to axon side. For pCREB experiments, 1 nM unconjugated BDNF-biotin was added for 1 hour before fixing. For

Ciliobrevin D (EMD Millipore) conditions, 20µM Ciliobrevin D was added 10 mins prior to

BDNF-Qdot addition and was present throughout BDNF-Qdot treatment/imaging, with the same timescale as control conditions.

Immunofluorescence

Neurons cultured on 25 mm glass coverslips were fixed at 7-8 DIV in phosphate- buffered saline (PBS) containing 4% paraformaldehyde and 4% sucrose for 8 mins.

Coverslips were washed three times in PBS and blocked with cell block (PBS with 5% normal goat serum and 1% bovine serum albumin (BSA)). Primary antibodies were incubated for 2 hours at RT in cell block. After removing the primary antibodies, the coverslips were washed with PBS and incubated for 1hr at RT with fluorophore- conjugated secondary antibodies diluted in cell block. Following washes with PBS, the coverslips were mounted in ProLong Gold Antifade Mountant (ThermoFisher Scientific) on glass slides.

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TIRF Motility Assay

Single molecule TIRF Motility assays were performed as previously described in detail (Olenick et al., 2016). In brief, HeLa cells 18–20 h post-transfection of Halo-Hook1 constructs were labeled with the Halo ligand TMR (Promega) and lysed in buffer containing 40 mM HEPES, 1 mM EDTA, 120 mM NaCl, 0.1% Triton X-100, and 1 mM magnesium ATP (pH 7.4) supplemented with protease inhibitors. Cell lysates were diluted in assay buffer containing 10mMmagnesium ATP, 0.3 mg/ml bovine serum albumin, 0.3 mg/ml casein, 10 mM DTT, and an oxygen-scavenging system. Diluted cell lysates were then flowed into imaging chambers with taxol-stabilized microtubules immobilized to the coverslip with a tubulin antibody. TIRF movies were acquired at room temperature at 4 frames/s using the Nikon TIRF system (PerkinElmer Life Sciences) on an inverted Ti microscope with a 100x objective and an ImageEM C9100–13 camera

(Hamamatsu Photonics) controlled by Volocity software.

Western blotting

To test siRNA efficiency, PC12 cells were transfected at 70-80% confluency with

45 pmol of a pool of Hook1 or Hook3 siRNAs using Lipofectamine RNAiMAX (Invitrogen) and lysed 48 hours later. PC12 cells were lysed (50 mM Tris-HCl (pH 7.4), 150 mM

NaCl, 1% Triton-X100, and protease inhibitors) and clarified by centrifugation at 13.2k rpm for 10 min at 4°C. For all Western blot experiments, samples were boiled in denaturing buffer for 5 min and run on a SDS/PAGE gel to separate proteins.

Imaging Analysis

For motility analysis, kymographs were generated using the MultipleKymograph plugin for Fiji (NIH) and analyzed using custom MATLAB software (MathWorks) or by measurement tools in Fiji. For effector localization, images were deconvolved using

Huygens Professional software. Then effector puncta were counted by hand, and length 133 of axon was measured in Fiji. Area of BDNF-Qdot was measured using measurement and analysis functions in Volocity (Perkin Elmer). For pCREB signal, signal was measured by outlining nucleus with the Hoechst staining and measuring integrated intensity of pCREB in that area with Fiji.

Statistical methods

Statistics were performed in GraphPad Prism. Student’s t test or Mann-Whitney test was used when comparing two data sets, as indicated, while a one-way analysis of variance (ANOVA) was used with multiple data sets. For all experiments, data was analyzed from at least three independent replicates. Statistical significances noted as follows: NS=P>0.05, *=P≤0.05, **=P≤0.01, ***=P≤0.001

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CHAPTER 5: Discussion and Future Directions

Cytoplasmic dynein drives the majority of minus-end directed vesicular and organelle motility in the cell. Yet, it remains unclear how dynein is spatially and temporally regulated given the variety of cargo that must be properly localized to maintain cellular function. In this thesis, I investigated the role of mammalian Hook proteins, Hook1 and Hook3, as potential regulators of molecular motors. I found that

Hook proteins act as dynein effectors using biochemistry, optogenetic approaches. and single molecule TIRF motility assays. I found that interaction of dynein-dynactin requires the N-terminal domain of Hook proteins, which resembles the calponin-homology domain of EB proteins yet cannot bind directly to microtubules. I also found that both

Hook1 and Hook3 effectively activate cytoplasmic dynein, inducing longer run lengths and higher velocities than the previously characterized dynein activator, BICD2.

Together, these results suggest that dynein adaptors can differentially regulate dynein to allow for organelle-specific tuning of the motor for precise intracellular trafficking.

In collaboration with the Dominguez lab, I helped identify a conserved region of the dynein LIC1 that mediates interactions with unrelated dynein-dynactin effectors.

Quantitative binding studies of purified proteins mapped these interactions to a conserved helix within LIC1 and to N-terminal fragments of Hook1, Hook3, BICD2, and

Spindly. I found that the LIC1-dynein-effector interaction is vital for processive dynein- dynactin-effector motility with in vitro motility assays and LIC1 helix mutations impair lysosomal positioning in cells. The results revealed a conserved mechanism of effector interaction with dynein-dynactin required for processive motility.

In the final part of my thesis, I focused on the role of Hook proteins in neuronal transport. I found that the Hook1 mediates transport of TrkB-BDNF signaling endosomes 135 in primary hippocampal neurons. Hook1 co-migrates with a subpopulation of Rab5 and

Rab7 endosomes but knockdown of Hook1 did not affect the overall motility of Rab5 or

Rab7. In contrast, Hook knockdown significantly reduced the motility of BDNF-signaling endosomes. In microfluidic chambers, Hook1 depletion resulted in a significant decrease in the flux and processivity of BDNF-Qdots along the mid-axon, an effect specific for

Hook1 but not Hook3. Further, Hook1 knockdown inhibited downstream BDNF-TrkB signaling to the nucleus. Together, this work supports a model in which differential association with cargo-specific effectors efficiently regulates dynein in neurons.

Potential dynein activators

The list of known dynein activators is growing, but there are many known dynein adaptors that have not been studied as activators yet, including HAP1, TRAKs, JIPs and

RILP. The assays used and developed in my thesis, including the inducible dimerization assays, single molecule TIRF motility assays, and classic biochemistry techniques like co-IPs, can be used in future studies to identify dynein activators.

One of the many hurtles to previous studies on dynein activators has been autoinhibition of the protein. For BICD proteins, the C-terminal tail folds back on itself and blocks the coiled-coil region that interacts with dynein-dynactin, which is why most studies of BICD proteins as dynein activators use a N-terminal construct that lacks the last ~400 amino acids. For Hook proteins, truncating the C-terminal cargo binding domain also improved the frequency of motility events in single molecule TIRF motility assays. TRAK2 is also autoinhibited by a head-to-tail interaction (van Spronsen et al.,

2013). Due to the evidence for autoinhibition of effectors, the precise constructs used for identifying dynein activators will make or break a study.

136

Due to a combination of recent studies, the structural aspects important for dynein activator function have been identified. The most inclusive of these features is a long coiled-coil region. For BICD2, BICDR-1, and Hook3, the coiled-coil region of these effectors lays along the Arp1 filament of dynactin and acts as a scaffold for dynein interaction (Chowdhury et al., 2015; Grotjahn et al., 2018; Urnavicius et al., 2015, 2018).

All the identified activators have >200 amino acid coiled-coil stretch. In addition, many of these activators have a known interaction site for LIC1. In BICD2 and Spindly, the CC1 box motif is used for interaction with LIC1, while the Hook domain facilitates LIC1 interaction in Hook proteins (Lee et al., 2018). While they differ in sequence, these regions might provide very similar structural aspects, a symmetrical hydrophobic binding site, for interaction with LIC1. Future studies might identify other sequences that fold into a similar LIC1 binding site. The other feature of known activators is an interaction region for cargo/protein binding, usually on the opposite side from the dynein-dynactin interaction region. For Hook proteins, the C-terminal region is responsible for interacting with cargo proteins, like FTS and FHIP (Bielska et al., 2014; Zhang et al., 2014). For

BICD proteins, the C-terminal coiled-coil interacts with Rab6 (Liu et al., 2013; Schlager et al., 2010; Terawaki et al., 2015). Spindly and Rab11FIP3 also have C-terminal cargo interaction domains.

Using the common features of known dynein activators, I tried to identify some of these features in known dynein adaptors and designed constructs to test for activation of dynein motility. The most promising candidates, HAP1 and TRAK1/2 have a stretch of coiled-coil similar to other activators, as well as the CC1 box and Spindly motif. The other adaptors, RILP and JIP3 have shorter coiled-coil regions. In addition, RILP has previously been shown to interact with LIC1 (Schroeder et al., 2014). Initial experiments tested these candidates in α-p150 co-IP experiments in cell lines, similar to Figure 2.3. 137

HAP1, JIP3, TRAK1/2, and RILP displayed similar results to Hook proteins, where there was increased dynein (as seen by DIC signal) compared to control (Fig. 5.1A,B). I also preformed α-DIC co-IP experiments in cell lines and found that BICD2, HAP1, and JIP3 co-immunoprecipitated, while Hook1 did not (Fig. 5.1C). These results could reflect differences in effector interaction interface on the dynein-dynactin complex.

In addition to immunoprecipitation experiments, I tested some of these potential effector proteins in the inducible dimerization assay used in Figure 2.1. JIP3 showed no change after dimerization with peroxisomes, as seen in Figure 5.2. On the other hand,

HAP1 induced clustering of peroxisomes, but there was no significant redistribution to the perinuclear region, as was seen for BICD or Hook proteins in Chapter 2. Since HAP1 has been suggested to interact with Htt and GRIP1 to activate motors, I also tested

HAP1 in cells expressing Htt or GRIP1, but the results were similar to cells with HAP1 alone (Fig. 5.2). While the results for HAP1 are intriguing, further work is needed to understand the role of HAP1 as a potential dynein effector.

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Figure 5.1 Co-IP experiments with other dynein adaptors

A) Anti-p150 IPs from Cos7 cells with expressed dynein adaptors. B) Quantification of p150 IP experiments from A. C) Anti-DIC IPs from Cos7 cells with expressed dynein adaptors.

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Figure 5.2 Optogenetic recruitment assay with other dynein adaptors

Representative images of dynein adaptors in induced dimerization assay pre and post- dimerization. Scale bar=10µm.

Overall, the preliminary results described here provide a starting point for future work on these potential dynein effectors. The most intriguing results are the HAP1 dimerization results, which display induced clustering of the cargo after dimerization, but no motility was seen in the TIRF motility assay. It is possible that HAP1 is autoinhibited or needs additional interaction factors to activate motor motility. JIP3 and RILP are less likely to be effectors based off these initial results and the lack of conserved features of dynein activators (i.e. long coiled coil) but might coordinate with other activators in cells.

These ideas will be discussed more in the next section.

140

Figure 5.3 Summary of results for dynein effectors and adaptors

Outstanding questions on dynein effectors

Recent studies including my thesis work have shown that dynein effectors can modulate motility in different ways. In Chapter 2, I showed that Hook1 and Hook3 display faster velocities and longer run lengths than BICD2 (Olenick et al., 2016). Another study on BICD proteins has also shown that BICDR-1 displays faster velocities than BICD2

(Schlager et al., 2014b). In addition, BICDR-1 and Hook3 have increased velocity and force of dynein-dynactin-effector complex by recruiting two dynein dimers per one dynactin-effector complex (Urnavicius et al., 2018). Cryo-EM studies have also shown that interaction of effector coiled-coil regions with the dynactin filament are slightly different from each other and might lead to different motility properties (Urnavicius et al.,

2018).

Studies are now focused on understanding how the different activators affect step size and efficiency. Discussions in the field suggest that activators might have

141 similar step size but an increased rate of stepping and less backward steps to help produce the longer and faster runs of BICDR-1 and Hook proteins. Future work will also need to examine the force under load since these activators are involved in the motility of different sized cargo. Furthermore, it seems that the close proximity of two dynein dimers on one dynactin filament might also be vital to enhance velocity and force. There might be allosteric interactions between the two dynein dimers to help produce the superprocessive motility seen between Hook3 and BICDR-1. Continued work in these areas is needed to better understand differences and mechanisms of effector-mediated processivity.

Since there are multiple dynein effectors, the question to be tackled now is how the effectors are regulated to control dynein-mediated transport. There is indication that some effectors are autoinhibited until interaction with cargo, as suggested by the overlapping Rab6 interaction with the C-terminal coiled-coil that folds for autoinhibition

(Terawaki et al., 2015). In addition, expression changes might also play a role in regulating which effectors are being utilized at different stages in development, such as the down-regulation of BICDR-1 during development (Schlager et al., 2010). It is not clear what the expression pattern is for other dynein effectors, but many are enriched in the brain. There is some indication that the localization of effectors in neurons might help to regulate transport. In my thesis work, Hook1 was found enriched in distal tips and

RILP is enriched in the mid-axon, while BICD2 is generally distributed (Chapter 4).

TRAK proteins have also been shown to have distinct localization, with TRAK1 in axons and TRAK2 predominately in dendrites (van Spronsen et al., 2013). Other protein interactions with effectors and post-translational modifications like phosphorylation might inhibit interaction of effectors with the motor complex, but future work is needed to understand the regulation of dynein effectors. 142

Since there are many dynein effectors and adaptors, it remains to be seen how these proteins work together for proper intracellular transport. Most studies to date have looked at dynein effectors in isolation or within individual families. In my thesis work

(Chapter 4), I found that Hook1 is enriched in the distal axon and RILP is enriched in the mid-axon, a similar pattern to Rab5 and Rab7 vesicles, respectively. In preliminary work,

I observed Hook1 and RILP co-migrating in the axons of primary hippocampal neurons.

This was seen for a portion of migrating RILP events, suggesting that there is a population of vesicles with a mixture of dynein effectors present. During the maturation of endosomal vesicles from Rab5-positive to Rab7-positive, it is possible that vesicles might go from Rab5-Hook1-positive to Rab7-RILP-postive, in a dynein effector handoff model. It is also possible there is a mixture of dynein effectors on a given cargo, with some dynein activators and some dynein adaptors. Recent work has shown in vitro that the presence of Lis1 in the dynein-dynactin-BICD2 complex enhances the motility compared to complexes without Lis1 (Gutierrez et al., 2017). It is unclear if Lis1 can interact with other dynein-dynactin-effector complexes and if Lis1 works with dynein- dynactin-effector complexes in vivo. The coordination of dynein effectors is important to understand the regulation of dynein motility and remains to be studied.

Since the majority of dynein effectors are thought to dimers due to the coiled coil region, it is still unclear if effectors in the same family can form and function as heterodimers. A previous paper suggested that Hook1, Hook2 and Hook3 could all interact and function together in complex with FTS and FHIP (Xu et al., 2008). From my work in neurons, it is unlikely that these Hook proteins act together or in similar pathways since Hook3 knockdown does not produce the same effect as Hook1 knockdown. In addition, my preliminary work looking for heterodimerization of Hook1 and

143

Hook3 was inconclusive. Future work is needed to better understand if heterodimerization is physiology for dynein effectors like hook proteins.

Some cargos like mitochondria have bidirectional motility with links to both dynein and kinesin. HAP1, TRAKs, and JIPs have been suggested to act as motility switches, due to the overlapping interaction sites for dynein and kinesin on these dynein adaptors (Fu and Holzbaur, 2014). Interacting proteins and post-translational modifications like phosphorylation act as the cues to switch from one motor interaction to the other (Fu and Holzbaur, 2014). For Hook proteins, a study on filamentous fungus suggested that Hok1 might act as a switch between dynein and kinesin-3 motility

(Bielska et al., 2014). In my work, I did not find any evidence for mammalian Hook proteins acting as motor switches, but it is still possible that Hook proteins might be able to interaction with kinesins in other cell types.

Some dynein effectors and regulators have previously been implicated in disease and developmental conditions. Mutations in BICD2 have been linked to DCSMA

(dominant congenital spinal muscular atrophy) with LED and SMA (Lipka et al., 2013).

Lis1 mutants cause type 1 lissencephaly, a neurodevelopmental disease (Reiner et al.,

1993). Other dynein adaptors like HAP1 are linked to disease proteins, such as huntingtin in Huntington’s disease. Some of the other dynein effectors have been understudied at this point and have yet to be linked to disease states, but future work should be done to look at the role of dynein effectors in disease conditions.

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