Regulatory Mechanisms of Semaphorin/Plexin/Mical-Mediated

REGULATORY MECHANISMS OF SEMAPHORIN/PLEXIN/MICAL-MEDIATED

F-ACTIN DISASSEMBLY AND CELLULAR REMODELING

APPROVED BY SUPERVISORY COMMITTEE

Jonathan Terman, Ph.D. (Advisor)

Jane Johnson, Ph.D. (Chair)

Neal Alto, Ph.D.

Helmut Kramer, Ph.D.

DEDICATION

I dedicate this dissertation to all the people who have guided me through this process and who have been instrumental in my success. Specifically, I am appreciative of my mentor,

Dr. Jonathan Terman, who encouraged me at every step and helped me become a critical scientific thinker. His guidance on my dissertation research has been invaluable to me. I would also like to express my deep gratitude to my committee members: Dr. Jane Johnson,

Dr. Neal Alto, and Dr. Helmut Kramer for their useful suggestions over the years which have helped direct and shape my research.

Furthermore, I am incredibly thankful for my wonderful colleagues in the Terman lab over the years – Chris Spaeth, Laura Alto, Gias Ahmed, Jimok Yoon, Gizem Yesilyurt, Heng

Wu, Ruei-Jiun Hung, and Taehong Yang – who have always been willing to listen to my ideas and discuss them with me, explain concepts or teach techniques to me, or help me in any way. I truly could not ask for better people to work with every day.

I am also extremely grateful for my family - for my parents, Steve and Vicky, and my sister, Sarah, who have always supported me and believed in me. Finally, I am especially grateful for my husband, Ryan, who has been a source of endless support, encouragement, love, and inspiration, and for my daughter, Corinne, who has brought me so much joy and happiness. I cannot thank you enough for all you have done to ensure my success.

REGULATORY MECHANISMS OF SEMAPHORIN/PLEXIN/MICAL-MEDIATED

F-ACTIN DISASSEMBLY AND CELLULAR REMODELING

SHANNON KAY GOOD RICH

DISSERTATION

Presented to the Faculty of the Graduate School of Biomedical Sciences

The University of Texas Southwestern Medical Center at Dallas

In Partial Fulfillment of the Requirements

For the Degree of

DOCTOR OF PHILOSOPHY

The University of Texas Southwestern Medical Center at Dallas

Dallas, Texas

May, 2017

Shannon Kay Good Rich, 2017

REGULATORY MECHANISMS OF SEMAPHORIN/PLEXIN/MICAL-MEDIATED

F-ACTIN DISASSEMBLY AND CELLULAR REMODELING

Publication No.

Shannon Kay Good Rich

The University of Texas Southwestern Medical Center at Dallas, 2017

Supervising Professor: Jonathan R. Terman, Ph.D.

Dynamic changes to the actin cytoskeleton modify the shape of cells and their membranous extensions, and underlie diverse developmental and functional events in multiple tissues including migration, navigation, and connectivity. Semaphorins, together with their Plexin receptors, are a large family of extracellular cues that trigger complex cytoskeletal rearrangements to direct these cellular phenomena, but the mechanisms regulating their effects are poorly understood. Emerging evidence identifies Mical, a conserved oxidoreductase (Redox) enzyme, as a critical component in Semaphorin/Plexin signaling through its post-translational oxidation of F-actin, which promotes actin instability and disassembly. How this Mical-mediated redox regulation of actin dynamics is locally

v positioned and coordinated with the activity of other actin regulatory proteins to achieve specific, targeted effects on the cytoskeleton remains unknown. Therefore, as a part of my dissertation research, I used a genetic assay to begin to address these questions and search for proteins that could alter Semaphorin/Plexin/Mical signaling effects on the cytoskeleton. In this dissertation, I present my discovery of a functional interplay between Mical and two critical new interactors – cofilin, a well-known ubiquitous F-actin regulatory protein, and

Sisyphus, an unconventional class XV myosin. With regards to cofilin, my in vivo genetic/functional assays reveal that cofilin activity is required for and enhances

Semaphorin/Plexin/Mical-dependent cytoskeletal rearrangements and morphological changes. Additionally, in vitro biochemical assays demonstrate that cofilin preferentially binds Mical-oxidized actin and accelerates its disassembly. Together, these findings indicate that cofilin and Mical act as a functional pair in both neuronal and non-neuronal cells to rapidly and efficiently disassemble actin filaments. Similarly, my results reveal that Sisyphus is necessary and sufficient for triggering Semaphorin/Plexin/Mical-dependent F-actin disassembly/cellular remodeling. Moreover, using in vivo functional assays, I find that

Sisyphus uses its myosin motor activity and the first MyTH4 domain of its C-terminal tail region to modify the subcellular localization of Mical. In this way, Sisyphus spatially controls Mical-dependent F-actin disassembly/cellular remodeling. Therefore, both cofilin and Sisyphus function to promote Mical-mediated F-actin disassembly; thereby, they act as critical regulators of Semaphorin/Plexin/Mical-mediated effects on cytoskeletal and morphological dynamics. Thus, my findings unveil novel molecular and biochemical mechanisms that orchestrate cellular, developmental, and neural biology.

TABLE OF CONTENTS

PRIOR PUBLICATIONS ...... ix

LIST OF FIGURES ...... x

LIST OF TABLES ...... xiii

LIST OF DEFINITIONS ...... xiv

CHAPTER ONE General Introduction ...... 1

CHAPTER TWO F-actin dismantling through a Redox-driven synergy between Mical and

cofilin ...... 20

Abstract ...... 20

Introduction ...... 21

Results ...... 23

Discussion ...... 31

Figures ...... 34

Materials and Methods ...... 61

CHAPTER THREE The Class XV Myosin Sisyphus Spatially Targets Mical to Direct

Semaphorin/Plexin-mediated Actin Disassembly and Cellular Remodeling ...... 74

Abstract ...... 74

Introduction ...... 75

Results ...... 77

Discussion ...... 88

Figures ...... 95

Materials and Methods ...... 122

vii

CHAPTER FOUR Summary and Conclusions...... 133

BIBLIOGRAPHY ...... 140

viii

PRIOR PUBLICATIONS

Sephton, C.F., Good, S.K., Atkin, S., Dewey, C.M., Mayer, P., Herz, J., and Yu, G. (2010). TDP-43 Is a Developmentally Regulated Protein Essential for Early Embryonic Development. J. Biol. Chem. 285, 6826–6834.

Dewey, C.M., Cenik, B., Sephton, C.F., Dries, D.R., Mayer, P., Good, S.K., Johnson, B.A., Herz, J., and Yu, G. (2011). TDP-43 Is Directed to Stress Granules by Sorbitol, a Novel Physiological Osmotic and Oxidative Stressor. Mol. Cell. Biol. 31, 1098–1108.

Grintsevich, E.E.*, Yesilyurt, H.G.*, Rich, S.K., Hung, R.-J., Terman, J.R., and Reisler, E. (2016). F-actin dismantling through a redox-driven synergy between Mical and cofilin. Nat Cell Biol 18, 876–885. (*Co-first authors)

LIST OF FIGURES

FIGURE 1.1. Mical-mediated Redox regulation of actin controls Semaphorin/Plexin F-actin

Disassembly ...... 19

FIGURE 2.1. Mical/F-actin dynamics are modulated by cofilin ...... 34

FIGURE 2.2 Cofilin slows F-actin oxidation by Mical but accelerates filament disassembly

...... 36

FIGURE 2.3 Further characterization of the interaction of Mical and cofilin in modulating

F-actin disassembly and the quantification of Mical-oxidized actin ...... 38

FIGURE 2.4 Further characterization of Mical-oxidized actin using a limited proteolysis

assay with subtilisin and an antibody directed against the Met-44 residue of

actin ...... 40

FIGURE 2.5 Mical-mediated oxidation of actin alters polymerization and weakens the

mechanical properties of filaments ...... 43

FIGURE 2.6 Mical oxidation of F-actin improves cofilin binding and results in accelerated

filament severing ...... 45

FIGURE 2.7 Mical oxidation of actin filaments accelerates their severing by yeast and

human cofilins ...... 47

FIGURE 2.8 Indication of the enhanced cofilin severing and binding to actin filaments

containing oxidized actin ...... 49

FIGURE 2.9 Further characterization of cofilin binding to Mical-oxidized actin ...... 51

FIGURE 2.10 Cofilin enhances Mical-mediated F-actin alterations in vivo ...... 53

FIGURE 2.11 Cofilin enhances Sema-Plexin-Mical repulsive axon guidance ...... 55

FIGURE 2.12 Further analysis of Cofilin’s effects on Mical and Semaphorin/Plexin-

mediated F-actin/cellular remodeling in vivo ...... 57

FIGURE 2.13 Uncropped gels ...... 59

FIGURE 3.1 The unconventional class XV myosin Sisyphus increases Mical-driven F-actin

disassembly/cellular remodeling ...... 95

FIGURE 3.2 Syph is required for Mical-dependent F-actin disassembly/cellular remodeling

...... 97

FIGURE 3.3 Further analysis of the effects of Syph and other myosins on

Semaphorin/Plexin/Mical-dependent bristle remodeling...... 99

FIGURE 3.4 Syph distributes Mical to direct F-actin disassembly and cellular remodeling ..

...... 102

FIGURE 3.5 Further analysis of the effects of Syph on changes in the cellular distribution

of Mical and Mical-mediated F-actin disassembly/cellular remodeling...... 104

FIGURE 3.6 The myosin motor activity of Syph locally targets Mical to disassemble F-

actin ...... 106

FIGURE 3.7 Further characterization of Syph motor mutants and their effects on Mical’s

localization and cellular morphology...... 108

FIGURE 3.8 Syph employs its first MyTH4 domain to redistribute Mical ...... 110

FIGURE 3.9 Further characterization that the PH-like and FERM domains are not required

for Syph-triggered redistribution of Mical ...... 112

FIGURE 3.10 Further characterization of Mical localization and cellular morphology in

bristles expressing Mical and Syph tail region domain deletions ...... 113

FIGURE 3.11 Sisyphus orchestrates Semaphorin/Plexin/Mical-induced morphological

changes by locally controlling Mical’s subcellular localization, thereby

influencing where and to what extent actin-based cellular remodeling occurs ..

...... 115

FIGURE 3.12 Mical and Syph regulate synaptic structure and F-actin muscle organization ...

...... 116

FIGURE 3.13 Larval expression of Syph enhancer trap lines ...... 118

FIGURE 3.14 Summary/working model: The unconventional myosin Syph spatially

regulates the localization of Mical to direct Semaphorin/Plexin/Mical-

mediated F-actin disassembly and cellular remodeling ...... 120

xii

LIST OF TABLES

TABLE 3.1 Primers used to generate the Syph tail region domain deletion transgenes ... 132

xiii

LIST OF DEFINITIONS

CAM - cell adhesion molecule

CH – calponin homology

CRMP – collapsin response mediator protein

DRG - dorsal root ganglion

EB1 - End binding 1

ECM – extracellular matrix

EGCG - (-)-epigallcatechin gallate

Eps8 - Epidermal growth factor receptor pathway substrate 8

ERM - Ezrin, Radixin, and Moesin

F-actin - filamentous actin

FAD – flavin adenine dinucleotide

FERM - protein Four-point-one, Ezrin, Radixin, Moesin

FM – flavoprotein monooxygenase

GAP - GTPase activating protein

GPI - glycosylphosphatidylinositol

LIM - Lin11, Isl-1, and Mec-3

Mical – Molecule interacting with CasL

MRTF-A – myocardin-related transcription factor-A

Msr – Methionine sulfoxide reductase

MST1 - Mammalian Ste20-related 1

MyTH4 – Myosin Tail Homology 4

xiv NADH - nicotinamide adenine dinucleotide

NADPH - nicotinamide adenine dinucleotide phosphate

NDR - nuclear Dbf2-related

NGF - nerve growth factor

NMJ – neuromuscular junction

Npn - neuropilin

PAK - protein p21 activated kinase

PH – pleckstrin homology

PHBH - p-hydroxybenzoate hydroxylase

PIR – Plexin-interacting region

PKA - protein kinase A

Plex - Plexin

PSI - plexin-semaphorin-integrin

RBD - Rho-binding domain

Redox - oxidoreductase

SelR – Selenoprotein R

Sema – Semaphorin sh2 - shaker-2

SH3 - SRC Homology 3

Syph - Sisyphus

VNC - ventral nerve cord

CHAPTER ONE

General Introduction

The ability of cells to move and change shape to adapt to their local environment underlies diverse cellular processes such as axon guidance, cell migration, and neural connectivity – with a final outcome that specifies the genesis and functional workings of each of the organ systems. These cellular behaviors and their ability to form different tissues are driven by the assembly, disassembly, and rearrangement of the actin filaments that underlie a cell’s structure (Blanchoin et al., 2014; Pollard and Cooper, 2009; Rottner and Stradal, 2011).

Controlling this actin reorganization and how it mediates cell motility and morphological changes are extracellular cues that bind diverse receptors on the cell’s surface (Bashaw and

Klein, 2010; Berzat and Hall, 2010; Kay et al., 2008; Swaney et al., 2010). In turn, these cues acting via their receptors trigger intracellular signaling pathways/networks that locally modify actin organization, as well as induce alterations to other structural and adhesive elements controlling morphology and movement such as microtubules and the remodeling of the plasma membrane (Berzat and Hall, 2010; Hung and Terman, 2011). These extracellular cues are known to act over both long and short distances, and likewise, depending on the cue and its receptor, the signal may either promote (such as through actin assembly and increased cellular adhesion) or inhibit (such as through actin disassembly and reduced cellular adhesion) local movement/growth and thereby direct cellular morphology and motility (Hung and Terman, 2011). Understanding the mechanisms by which cells move and change their shape has long occupied a critical place in biomedical research; however, the precise

1 2 mechanisms by which cues induce specific and targeted structural changes within a cell are still far from clear.

Over the past 25 years, attempts to understand the complex wiring of the nervous system has led to the discovery of many types and families of extracellular/guidance cues

(Hung and Terman, 2011). This work has identified that guidance cues are diverse and can be both small molecules and large proteins, with single family members or multiple family members, such as those present within such well-known families of cues as Semaphorins,

Netrins, Slits, and Ephrins (Bashaw and Klein, 2010; Berzat and Hall, 2010; Hung and

Terman, 2011; Kolodkin and Tessier-Lavigne, 2011). Likewise, these cues have been found to exert multiple different effects on cells – playing critical roles in regulating morphology, motility, and connectivity in both neuronal and non-neuronal cells. In neurons, for example, a specialized actin-rich structure at the tip of growing axons, the growth cone, responds to attractive (growth/motility promoting) and repulsive (growth/motility inhibiting) guidance cues in the environment. Then, through the poorly-understood action of these cues and their cell surface receptors, actin dynamics are controlled in a directed manner within the growth cone to precisely allow the extension and navigation of axons from the neuronal cell body to its proper target. In much the same way, it is now appreciated that many different types of cells and developmental processes depend on these extracellular guidance cues to undergo cytoskeletal reorganization (Kolodkin and Tessier-Lavigne, 2011). Thus, the characterization of these extracellular guidance cues, and the complex network of effector proteins and the cellular effects they specify is critical to understanding diverse biological phenomena including how neurons form and maintain their connections.

3 Semaphorins and Plexins

Semaphorins (Semas) are one of the largest families of extracellular guidance cues. To date,

30 different Semas have been identified, and these are divided into eight classes based on similarities in structure and amino acid sequence (Alto and Terman, 2017). Classes 1, 2, and

5 (Sema-5c only) are found in invertebrates, classes 3-7 are found in vertebrates, and class V are found in viruses (Semaphorin Nomenclature Committee, 1999; Alto and Terman, 2017).

Some Semas are secreted (classes 2, 3, and V), while others are transmembrane (classes 1, 4,

5, and 6) or glycosylphosphatidylinositol (GPI)-linked (class 7) proteins (Alto and Terman,

2017). While the domain structure of the different classes of Semas vary, all Sema family members have an N-terminal cysteine-rich Sema domain, and all but a few viral Semas have a cysteine-rich plexin-semaphorin-integrin (PSI) domain (Alto and Terman, 2017). Structural and functional studies indicate that the Sema domain mediates homophilic dimerization, which is essential for Sema’s function (Antipenko et al., 2003; Janssen et al., 2010, 2012;

Klostermann et al., 1998; Koppel and Raper, 1998; Liu et al., 2010; Love et al., 2003; Nogi et al., 2010). The Sema domain also mediates binding to various receptors (Alto and Terman,

2017; Sharma et al., 2012; Yazdani and Terman, 2006).

Early characterizations of the first Semaphorin family members to be identified

(Sema-1a, originally called Fasciclin IV, and Sema 3A, originally called Collapsin) revealed that Semas affect axon guidance (Kolodkin et al., 1992) and cause axon growth cone collapse

(Luo et al., 1993). Careful examination of Sema3A-induced growth cone collapse found that exposure of dorsal root ganglion (DRG) growth cones to Sema3A resulted in rapid F-actin disassembly and growth cone collapse (Fan et al., 1993) – indicating a role for Semas in

4 negatively regulating growth/motility through effects on the actin cytoskeleton. Subsequent identification and characterization of additional Sema proteins in both invertebrates and vertebrates further supported the notion that Semas act as repulsive axon guidance cues to mediate proper nervous system wiring/patterning (reviewed in Pasterkamp, 2012; Tran et al.,

2007; Yoshida, 2012). For example, characterization of Sema3A mutant (Sema3A-/-) mice determined that loss of Sema3A produces cranial and spinal nerves that are inappropriately defasciculated and have excessive branching into areas of the embryonic nervous system that would normally express Sema3A (Taniguchi et al., 1997). Similarly, motor neuron axons in

Sema3A-/- mutants reach the embryonic limb bud, which has distinct regions of Sema3A expression, too early in development – and aberrantly project in an abnormally defasciculated manner within it (Huber et al., 2005). Likewise, examination of Sema3F mutants (Sema3F-/-) revealed aberrant axon trajectories in the embryonic limb bud, such that axons projected into Sema3F-expressing areas that they normally avoid (Huber et al., 2005).

These studies indicated that Semas play inhibitory/repulsive roles to restrict axonal growth, branching, and defasciculation and keep specific axons from entering inappropriate areas.

Numerous additional studies have also shown that Semas can work to exert other repulsive/inhibitory effects (e.g., such as promoting defasciculation through repulsive effects between axons within a nerve bundle (Kolodkin and Tessier-Lavigne, 2011; Yazdani and

Terman, 2006)) and regulate multiple other aspects of neural connectivity – including neuronal migration, synapse formation, and dendrite morphology (Chen and Cheng, 2009;

Fiore and Püschel, 2003; Pasterkamp, 2012; Pasterkamp and Giger, 2009; Shen and Cowan,

2010; Tran et al., 2007; Yoshida, 2012). Growing evidence also links Semaphorins to

5 numerous diseases of the nervous system (Pasterkamp and Giger, 2009; Van Battum et al.,

2015) and to inhibition of axon regeneration after nerve injury (Fawcett et al., 2012; Giger et al., 2010; Pasterkamp and Giger, 2009).

It is also important to note that Semas are widely expressed and have physiological roles in other systems besides the nervous system. For example, Semaphorins play prominent roles in the immune, endocrine, circulatory, and musculoskeletal systems (Alto and Terman,

2017; Yazdani and Terman, 2006), where they are critical in immunity (Roney et al., 2013;

Suzuki et al., 2008; Takamatsu and Kumanogoh, 2012), vasculogenesis and angiogenesis

(Carmeliet and Tessier-Lavigne, 2005; Gelfand et al., 2009), and bone homeostasis (Harre and Schett, 2013; Kang and Kumanogoh, 2013; Kumanogoh and Kikutani, 2013), respectively. Semaphorins have also been prominently linked to cancer, where they act in both promotion and suppression of tumor progression, in a class-specific manner (Neufeld et al., 2016).

Semaphorins mediate their repulsive effects primarily through transmembrane Plexin

(Plex) receptors. There are 4 classes of Plexins in vertebrates (A, B, C, and D) and 2 in invertebrates (A and B). Additionally, Semas can directly bind other receptors including neuropilin (Npn) receptors; indeed, class 3 Semaphorins (except Sema3E) require Npns as co-receptors for signaling – working with Plexins as well as cell adhesion molecules (CAMs)

– such as Nr-CAM and L1-CAM (Alto and Terman, 2017; Sharma et al., 2012). Plexins induce changes in cell adhesion and cytoskeletal organization in response to Sema binding through a complex network of downstream intracellular effector proteins (Hota and Buck,

2012; Hung and Terman, 2011; Yang and Terman, 2013). In particular, a wide variety of

6 proteins have now been implicated in Sema/Plex-induced repulsion – including kinases, cyclic nucleotides, GTPases, cell adhesion molecules, and proteins known to modify the cytoskeleton (Bashaw and Klein, 2010; Yazdani and Terman, 2006). However, the precise signaling networks regulating and mediating Sema/Plex-induced repulsion are still being elucidated, especially concerning Sema/Plex-induced collapse and reorganization of F-actin.

Small GTPases and their regulators have become well appreciated for their involvement in regulating Semaphorin repulsion. In particular, Rho, Ras, and Rap GTPases, small GTPases known to affect integrin function and the cytoskeleton (Etienne-Manneville and Hall, 2002; Hall and Lalli, 2010; Kinbara et al., 2003), are particularly important for the activity and function of Plexins as well as for mediating aspects of Sema/Plex signaling

(Hota and Buck, 2012; Püschel, 2007; Yang and Terman, 2013). Plexins have a GTPase activating protein (GAP) homology domain by which they turn-off Ras and Rap family

GTPases, and this action is especially important for Sema/Plex-mediated repulsion through effects on integrin-mediated cell adhesion (Oinuma et al., 2004, 2006; Rohm et al., 2000;

Saito et al., 2009; Uesugi et al., 2009; Wang et al., 2012; Yang and Terman, 2012). GTP- bound (active) Ras and Rap proteins, including R-Ras, enhance integrin binding to their extracellular matrix (ECM) ligands which promotes and stabilizes cell adhesion (Kinbara et al., 2003). In Sema/Plex signaling, Plexin GAP activity reduces the levels of GTP-bound Ras

(Ito et al., 2006; Oinuma et al., 2004, 2006; Toyofuku et al., 2005; Uesugi et al., 2009).

Therefore, by turning off Ras activity, Plexins reduce adhesion to the extracellular matrix.

The GAP activity of Plexin A is regulated by protein kinase A (PKA), which phosphorylates

Plexin leading to the binding of 14-3-3ɛ (a phospho-serine/threonine binding protein) to

7 Plexin. The association of Plexin with 14-3-3ɛ prevents Ras from binding Plexin; thus, Plexin cannot regulate Ras activity (Yang and Terman, 2012). Plexins also have a Rho-binding domain (RBD) by which they associate with various Rho GTPase family members (Hota and

Buck, 2012). For example, Plexin B binds Rac through its RBD and is thought to sequester

Rac from its effector protein p21 activated kinase (PAK); thereby, inhibiting signaling cascades mediated by PAK (Driessens et al., 2001; Hu et al., 2001; Vikis et al., 2002), a serine/threonine kinase which is known to regulate cytoskeletal dynamics (Bokoch, 2003).

Along with Sema binding, binding of Rho GTPases to the RBD of Plexin is also thought to be critical for Plexin GAP activity (Hota and Buck, 2012; Negishi et al., 2005; Yang and

Terman, 2013).

The MICAL family of proteins – domain organization, interactions, and involvement in

Sema/Plex signaling

In the past 15 years, a unique family of actin regulatory proteins, the MICALs, have emerged as critical players in Sema/Plex-mediated repulsion. A yeast two-hybrid screen conducted to find proteins that bind and function with Plexin A identified a novel family of proteins, the

MICALs, as binding partners of the cytoplasmic region of Plexin A (Terman et al., 2002).

There are three mammalian MICAL family members (MICAL-1, MICAL-2, and MICAL-3), one Drosophila MICAL (Terman et al., 2002), and 5 Zebrafish MICALs – MICAL-1,

MICAL-2a, MICAL-2b, MICAL-3a, MICAL-3b (Xue et al., 2010). These family members are collectively referred to as MICAL(s) (Terman et al., 2002) and were named after the first

MICAL (Molecule interacting with CasL), human MICAL-1, which was found in a far-

8 western screen initiated to find proteins that bind CasL, an SRC Homology 3 (SH3)-domain containing adaptor protein (Suzuki et al., 2002).

MICALs are large proteins with several conserved domains present in all MICALs

(Figure 1.1a). At their N-termini, MICALs have a flavoprotein monooxygenase (FM or redox) domain characterized by specific flavin adenine dinucleotide (FAD) binding motifs

(GxGxxG, GD, DG) (Terman et al., 2002) that are common to flavoprotein monooxygenases

(also known as hydroxylases), a subclass of oxidoreductases (Figure 1.1a; (Eggink et al.,

1990; Eppink et al., 1997; Terman et al., 2002; Wierenga et al., 1986)). Through their catalytic activity, flavoprotein monooxygenases bind and use FAD in reduction-oxidation

(Redox) reactions to insert one atom of molecular oxygen into a substrate (Massey, 1995), but particular substrates vary depending on the specific flavoprotein monooxygenase and cannot be predicted based on sequence. Supporting the idea that MICAL binds FAD, the purified FM domain of Mical has a yellow color like other FAD-binding proteins and a similar absorption spectra as FAD (Terman et al., 2002). Additionally, two independent studies confirmed through structural analysis of this region of mouse MICAL-1 that MICAL is an FAD-binding protein (Nadella et al., 2005; Siebold et al., 2005). In addition to FAD, flavoprotein monooxygenases use nucleotides such as nicotinamide adenine dinucleotide

(NADH) or nicotinamide adenine dinucleotide phosphate (NADPH) as electron donors in redox reactions (Massey, 1995). Likewise, the FM domain of MICAL consumes NADPH indicating that NADPH is a coenzyme for MICAL (Hung et al., 2010; Nadella et al., 2005;

Schmidt et al., 2008; Zucchini et al., 2011). A related protein family to MICALs, the

MICAL-Likes, have similar domain structure to MICALs but lack the FM domain (Terman

9 et al., 2002; Xue et al., 2010). The known substrates and functionality of MICAL’s FM domain will be discussed in more detail in the section below (see “Mical is a redox effector of actin”).

Interestingly, unlike other known flavoprotein monooxygenases, MICALs have additional domains besides their catalytic region (Terman et al, 2002). Adjacent to the FM domain, MICALs have a Calponin Homology (CH) domain (Figure 1.1a). There are several types of CH domains (type-1,-2,-3), and they are found in proteins that interact with the cytoskeleton and function in cell-cell signaling (Gimona et al., 2002). Type-1 CH domains directly bind to actin whereas type-2 CH domains do not (Gimona et al., 2002). Structural determination of the MICAL-1 CH domain identified it as a type-2 CH domain (Sun et al.,

2006), and biochemical assays found no evidence that MICAL’s CH domain directly binds to actin (Hung et al., 2010; Sun et al., 2006). However, the CH domain is required for the proper in vivo positioning of Mical (Hung et al, 2010), and recent work has indicated that

MICAL’s CH domain modulates the catalytic activity of the FM domain, such that the enzymatic activity of purified MICAL-FMCH (only the FM and CH domains of MICAL) is enhanced compared to MICAL-FM (Alqassim et al., 2016).

C-terminal to the CH domain, MICALs have a LIM (Lin11, Isl-1, and Mec-3) domain

(Figure 1.1a). LIM domains contain two closely linked Zinc-finger domains and are found in proteins with diverse cellular functions (Zheng and Zhao, 2007). They have been shown to mediate protein-protein interactions (Bach, 2000), and this is true of the MICAL LIM domain as well (Schmidt et al., 2008; Zhou et al., 2011a). To date, amino acid stretches containing the LIM domain of MICAL-1 have been found to bind to NDR (nuclear Dbf2-

10 related) kinase and negatively regulate this pro-apoptotic protein by competing for NDR kinase binding with MST1 (Mammalian Ste20-related 1), an activator of NDR kinase (Zhou et al., 2011a). Additionally, collapsin response mediator protein 2 (CRMP2) interacts with a region containing the LIM domain of MICAL-1 and alters MICAL’s activity (Schmidt et al.,

2008).

The C-termini of MICALs contain proline-rich motifs (PxxP) and coiled-coil motifs

(Figure 1.1a). Proline-rich motifs are known to facilitate binding to SH3 domains

(Alexandropoulos et al., 1995; Kay et al., 2000; Williamson, 1994), and the proline-rich motif of MICAL-1 is known to associate with the SH3 domain of CasL (Suzuki et al., 2002).

The C-terminus of MICAL-1 mediates a direct interaction with vimentin, an intermediate filament protein, although the functional consequence of this interaction is unknown (Suzuki et al., 2002). Additionally, the C-termini of MICALs directly bind multiple Rab GTPases

(Fischer et al., 2005; Fukuda et al., 2008; Weide et al., 2003) such as Rab8A (Grigoriev et al., 2011). Specifically, MICAL-3 binds Rab8A, and the Rab6-interacting protein, ELKS, and promotes exocytic vesicle docking and fusion (Grigoriev et al., 2011). Plexin A also interacts with the C-termini of MICALs in a region referred to as the Plexin-interacting region (PIR) which contains a coiled coil motif with sequence similarity to the alpha-helical region of Ezrin, Radixin, and Moesin (ERM) proteins (Schmidt et al., 2008; Terman et al.,

2002).

The initial characterization of the interaction between Mical and Plexin (Terman et al., 2002) revealed that Mical is critical for Sema/Plex-mediated repulsion. Specifically,

Mical mutant embryos display axon guidance defects that resemble defects observed in

11 Plexin A and Sema-1a mutants, and Mical genetically interacts with Sema-1a and PlexA to specify axon guidance. Interestingly, point mutations disrupting the FAD binding regions of

Mical as well as neuronal expression of Mical FAD-binding mutants (glycine residues in the

FAD binding motif GxGxxG were mutated to tryptophan residues) also cause significant axon guidance and connectivity defects resembling those found in Mical mutants, indicating that the redox activity of Mical is necessary for Mical’s function in Sema/Plex-dependent axon guidance (Beuchle et al., 2007; Terman et al., 2002). Both studies in Drosophila and mammals indicate that MICALs co-immunoprecipitate with Plexin A (Schmidt et al., 2008;

Terman et al., 2002) and that more MICAL co-immunoprecipitates with PlexA in Sema3A treated cells versus controls (Schmidt et al., 2008). Furthermore, the C-terminal region of

MICAL containing the Plexin-interacting region binds MICAL near the LIM domain and inhibits MICAL’s enzymatic activity (Schmidt et al., 2008; Vitali et al., 2016). This work coupled with genetic studies (Hung et al., 2010) suggests a model where Semaphorin signaling promotes MICAL binding to Plexin, and the binding of Plexin to MICAL’s C- terminal region relieves autoinhibition of MICAL so that it is enzymatically active. In contrast, Mical does not appear to directly bind to other Plexins such as Plexin B, the Sema-

2a receptor. However, Mical and Plexin B genetically interact, such that Drosophila embryos heterozygous for both Plexin B and Mical loss-of-function mutations display defects in axon guidance, further indicating that Mical is a critical component of Sema-Plex mediated axon repulsion (Ayoob et al., 2006). The functional requirement for MICALs in Semaphorin signaling-mediated cytoskeletal reorganization and repulsion is not isolated to a specific

MICAL family member or to a particular class or class member of Semaphorins; in fact, all

12 of the mammalian MICALs have been implicated in Sema3A signaling (Aggarwal et al.,

2015; Hou et al., 2015; Morinaka et al., 2011; Pasterkamp et al., 2006; Schmidt et al., 2008;

Terman et al., 2002). Additionally, MICALs have been linked to Sema3F signaling

(Pasterkamp et al., 2006) and MICAL-3 has been implicated in Sema6A signaling (Bron et al., 2007).

MICAL is a redox effector of actin

Although early characterizations of Mical demonstrated its importance in Semaphorin signaling, the manner in which Mical mediated repulsion was not understood. In Drosophila melanogaster, adult Mical mutants were found to have morphologically defective mechanosensory bristles (Hung et al., 2010). As bristle development and morphology depend on the extension of an actin-rich process, and has long been used as a model for studying actin dynamics in vivo (Tilney and DeRosier, 2005), this finding offered an important clue that Mical could be regulating actin dynamics (Hung et al., 2010). Subsequent analysis using biochemical assays and genetic and cellular studies indicated that Mical directly binds actin, and through the activity of its FM (Redox) domain Mical disassembles F-actin and bundled

F-actin, and alters actin in such a way as to inhibit repolymerization (Hung et al., 2010).

Furthermore, genetic experiments demonstrated that Mical-dependent changes to F-actin organization and cellular morphology are dependent on Semaphorin/Plexin (Sema-1a/Plex

A) signaling, and likewise, that Mical mediates the F-actin reorganizational events induced by Semaphorin/Plexin signaling (Hung et al., 2010). Mammalian MICALs also bind actin and induce F-actin disassembly (e.g., (Lee et al., 2013; Lundquist et al., 2014; McDonald et

13 al., 2013; Zucchini et al., 2011)), indicating a conserved function of this protein family. It is also known that MICAL-mediated redox activity and F-actin disassembly is dependent on the coenzyme NADPH, and the binding of MICAL to F-actin initiates NADPH consumption which indicates that MICAL is activated by F-actin (Hung et al., 2010, 2011; Lee et al.,

2013; McDonald et al., 2013; Zucchini et al., 2011). Mass spectrometry analysis on Mical- treated actin also revealed that actin filaments serve as a substrate for Mical, which oxidizes specific conserved methionine (Met)-44 and Met-47 residues of actin (Figure 1.1b; (Hung et al., 2011)). These residues are situated within the D-loop of actin, which mediates binding between two actin monomers, and both Met-44 and Met-47 sit at the interface between the pointed end of one actin monomer and the barbed end of the actin monomer it binds

(Dominguez and Holmes, 2011; Fujii et al., 2010; Galkin et al., 2010; Hung et al., 2011;

Murakami et al., 2010; Oda et al., 2009; Sheterline et al., 1998). Additional analysis revealed that Mical redox activity directly severs F-actin, and actin filaments with a Met-44 to Leu-44 mutation are resistant to Mical-mediated actin disassembly (Hung et al., 2011). Thus, Mical- mediated oxidation of actin causes actin disassembly by disrupting the binding of actin subunits within a filament so that the filament falls apart (Figure 1.1b; (Hung et al., 2011)).

The MICALs therefore are proteins of unique function that use a Redox post- translational regulatory system to specifically target F-actin for disassembly. Interestingly, while the MICALs are the first known class of enzymes that specifically target selective methionine residues on proteins, methionine oxidation is known to yield two different stereoisomers, Met-S-sulfoxide (Met-S-O) and Met-R-sulfoxide (Met-R-O). Furthermore, it is known that Met-S-O and Met-R-O are reduced to Met by distinct methionine sulfoxide

14 reductases, Methionine sulfoxide reductase A (MsrA) and Selenoprotein R/Methionine sulfoxide reductase B (SelR/MsrB), which act stereospecifically to reduce Met-S-O and Met-

R-O, respectively (Kryukov et al., 2002; Stadtman et al., 2002, 2003; Ugarte et al., 2009).

Recently, biochemical and mass spectrometric studies identified that MICAL-oxidation of actin is reversible by SelR/MsrB (Figure 1.1b; (Hung et al., 2013; Lee et al., 2013)).

Functional studies also revealed that SelR/MsrB restores the ability of MICAL-treated actin to repolymerize (Hung et al., 2013; Lee et al., 2013) – and that SelR reverses Mical-mediated effects on actin in vivo and functions in axon guidance, neural connectivity, skeletal muscle patterning, and Semaphorin/Plexin/Mical-dependent cell-cell signaling (Hung et al., 2013).

These results therefore further indicate the specificity of MICAL-mediated oxidation of F- actin – and that Mical and SelR/MsrB comprise a previously unknown reversible Redox post-translational regulatory system specifically targeting actin (Figure 1.1b). Moreover, these studies reveal that the enzyme-driven interconversion of specific Met/oxidized Met residues (Met  Met(R)O), similar to the reversible phosphorylation of specific serine/threonine/tyrosine residues, provides a selective means to precisely modulate protein function. Therefore, while protein oxidation is usually thought to be harmful to cell health and protein function, MICAL and SelR/MrsB act as a biologically critical redox switch that regulates actin dynamics, Semaphorin/Plexin signaling-induced actin rearrangements, and multiple other cellular events (Hung et al., 2013; Lee et al., 2013).

15 MICAL in development and disease

In addition to the role of the MICALs in Sema/Plex-mediated axon guidance, growth cone collapse, and Drosophila mechanosenory/bristle development mentioned above (Ayoob et al.,

2006; Bron et al., 2007; Hung et al., 2010; Pasterkamp et al., 2006; Schmidt et al., 2008;

Terman et al., 2002), functional studies have also indicated that MICAL is required for F- actin organization and reorganization events that underlie numerous other developmental processes. For example, genetic studies have revealed that MICAL-1 regulates the targeting of immunoglobulin superfamily cell adhesion molecule (IgCAM)-containing vesicles to the membrane of growth cones through redox-dependent F-actin disassembly, and thereby mediates proper assembly of hippocampal mossy fiber circuitry (Van Battum et al., 2014).

Mical also regulates synaptic structure, muscle F-actin organization, and myofilament organization (Beuchle et al., 2007). In particular, at neuromuscular junctions (NMJs), Mical mutant synapses do not spread out normally along the post-synaptic muscles and boutons often cluster at the nerve entry point or synaptic branches (Beuchle et al., 2007). Mical- mediated effects on F-actin organization are also observed in skeletal muscles, and these are also thought to contribute to the structural defects of NMJ synapses (Beuchle et al., 2007). In sensory neurons, F-actin severing by Mical is also critical for the pruning of dendritic branches (Kirilly et al., 2009). Furthermore, MICAL-2 mediated disassembly of nuclear actin in cultured cells and DRG neurons regulates the levels of G-actin in the nucleus. This Mical- mediated regulation of nuclear actin controls the nuclear levels of myocardin-related transcription factor-A (MRTF-A), and in this way, MICAL-2 regulates MRTF-A-dependent gene transcription in response to nerve growth factor (NGF) and mediates neurite outgrowth

16 (Lundquist et al., 2014). Although many of the functional studies of MICAL have examined its role in the nervous system, MICALs are also expressed in many tissues including brain

(Ashida et al., 2006; Bron et al., 2007; Fischer et al., 2005; Kirilly et al., 2009; Morinaka et al., 2011; Pasterkamp et al., 2006; Terman et al., 2002; Weide et al., 2003; Xue et al., 2010), skeletal muscles (Ashida et al., 2006; Beuchle et al., 2007; Fischer et al., 2005; Terman et al.,

2002; Xue et al., 2010), heart (Ashida et al., 2006; Fischer et al., 2005; Xue et al., 2010), kidney (Ashida et al., 2006; Fischer et al., 2005; Suzuki et al., 2002), lung (Ashida et al.,

2006; Fischer et al., 2005; Suzuki et al., 2002), spleen (Suzuki et al., 2002), bone marrow

(Ashida et al., 2006), thymus (Ashida et al., 2006; Suzuki et al., 2002), liver (Ashida et al.,

2006; Fischer et al., 2005), testis (Ashida et al., 2006; Suzuki et al., 2002), and fibroblasts

(Hochman et al., 2006). Functional studies have also indicated a role for MICAL-mediated

F-actin alterations and cellular remodeling in heart development (Lundquist et al., 2014), vascular permeability (Hou et al., 2015), podocyte morphology and function (Aggarwal et al., 2015), and innate immunity (Lee et al., 2013). Thus, MICALs likely control cytoskeletal dynamics in a variety of cell types and cellular processes.

Increasingly, expression and functional studies have linked MICALs to numerous diseases and disorders throughout the body. In the nervous system, MICAL 1 and MICAL-2 have been implicated in Alzheimer’s disease (Müller et al., 2007), temporal lobe epilepsy

(Luo et al., 2011), and cerebral ischemia-induced brain damage (Hou et al., 2015).

Furthermore, MICALs may inhibit neuronal regeneration after spinal cord injury

(Pasterkamp et al., 2006). Additionally, MICAL-1 and MICAL-2 have been implicated in multiple types of cancer and in cancer growth (Ashida et al., 2006; Ho et al., 2012; Loria et

17 al., 2014; Mariotti et al., 2015), and genome-wide association or expression profiling studies have linked these MICALs to obesity (Li et al., 2013), liver disease (Chambers et al., 2011), and muscular dystrophy (Marotta et al., 2009). MICAL-1 is also required for Sema3A upregulation-mediated pathology in diabetic nephropathy (Aggarwal et al., 2015). Although not extensively studied or characterized, a few pharmacological inhibitors of

MICAL/MICAL-mediated effects have been identified. For example, (-)-epigallcatechin gallate (EGCG), a component of green tea known to inhibit two flavoprotein monooxygenases including p-hydroxybenzoate hydroxylase (PHBH; (Abe et al., 2000a,

2000b)) which is the flavoprotein monooxygenase most structurally similar to MICAL

(Nadella et al., 2005; Siebold et al., 2005), attenuates Sema3A and Sema3F-induced axonal repulsion of rat DRG explants, suggesting that this compound prevents MICAL-mediated F- actin disassembly (Pasterkamp et al., 2006; Terman et al., 2002). Additionally, a small- molecule compound, CCG-1423, which inhibits growth in several cancer cell lines (Evelyn et al., 2007), binds to MICAL-2 and inhibits the effects of MICAL in MRTF-A signaling

(Lundquist et al., 2014). Thus, MICALs could be a potential therapeutic target for numerous pathologies.

Summary of dissertation contributions

Mical is a critical and unique F-actin regulatory protein that serves to remodel the cytoskeleton downstream of Sema/Plex signaling as well as in other contexts. Mical’s activity and effects on F-actin must be tightly controlled to avoid abnormal changes in cell morphology and motility, but the mechanisms by which MICAL-mediated F-actin

18 disassembly and cellular remodeling are precisely regulated and facilitated are not well understood. In my thesis work I sought to investigate the mechanisms regulating Mical activity and function by identifying novel proteins that genetically and functionally interact with Mical and by characterizing how these interactions contribute to

Semaphorin/Plexin/Mical-mediated F-actin disassembly and cellular remodeling. Chapter 2 demonstrates through biochemical assays, genetic interaction studies, and imaging of cellular

F-actin that Mical works with cofilin/ADF, a well-known, ubiquitously-expressed actin regulatory protein, to dismantle F-actin in vitro and in vivo. This synergism between Mical and cofilin is required for Semaphorin/Plexin-mediated F-actin disassembly, cellular remodeling, and axon guidance. Chapter 3 identifies and characterizes a novel interaction between Mical and the class XV myosin, Sisyphus. Genetic interaction and localization studies suggest that Sisyphus contributes to Semaphorin/Plexin/Mical signaling-induced cytoskeletal changes by modifying Mical’s cellular localization, thereby influencing the location and extent of actin disruption.

Figure 1.1. Mical-mediated Redox regulation of actin controls Semaphorin/Plexin F- actin Disassembly. (a) The redox enzyme, Mical, contains multiple conserved domains/motifs. FM (Flavoprotein monooxygenase), CH (Calponin Homology), LIM (Lin11, Isl-1, and Mec-3), PIR (Plexin-interacting region), ERM (Ezrin, Radixin, Moesin). (b) Semaphorin/Plexin signaling leads to the activation of Mical which uses the co-enzyme NADPH in its Redox reactions. Mical uses actin filaments as a substrate, oxidizing specific methionine (Met) residues on actin (MetO) to disrupt the interactions between individual actin filament subunits and disassemble F-actin. SelR is a reductase enzyme that reverses Mical-mediated oxidation of actin.

CHAPTER TWO

F-actin dismantling through a Redox-driven synergy between Mical and cofilin

This work was previously published: Grintsevich, E.E.*, Yesilyurt, H.G.*, Rich, S.K., Hung, R.-J., Terman, J.R., and Reisler, E. (2016). F-actin dismantling through a redox-driven synergy between Mical and cofilin. Nat Cell Biol 18, 876–885. (*Co-first authors). I performed and defined the genetic interaction between cofilin, Mical, and Plexin, which provided the basis for further in vivo analysis. In addition, E.E.G and H.G.Y performed biochemical experiments, with R-J.H also characterizing Mical-oxidized actin. H.G.Y collected images and performed in vivo F-actin alterations analysis. R-J.H and J.R.T developed the strategy and characterization of the MetO-44 and wild-type Met-44 antibodies. J.R.T performed axon guidance assays and genetic interaction analysis with SelR and ActinM44L.

Abstract

Numerous cellular functions depend on actin filament (F-actin) disassembly. The best-characterized disassembly proteins, the ADF/cofilins/twinstar, sever filaments and recycle monomers to promote actin assembly. Cofilin is also a relatively weak actin disassembler, posing questions about mechanisms of cellular F-actin destabilization. Here we uncover a key link to targeted F-actin disassembly by finding that F-actin is efficiently dismantled through a post-translational-mediated synergism between cofilin and the actin- oxidizing enzyme Mical. We find that Mical-mediated oxidation of actin improves cofilin binding to filaments, where their combined effect dramatically accelerates F-actin disassembly compared to either effector alone. This synergism is also necessary and sufficient for F-actin disassembly in vivo, magnifying the effects of both Mical and cofilin on cellular remodeling, axon guidance, and Semaphorin/Plexin repulsion. Mical and cofilin, therefore, form a Redox-dependent synergistic pair that promotes F-actin instability by rapidly dismantling F-actin and generating post-translationally modified actin that has altered assembly properties.

20 21 Introduction

Multiple cellular behaviors depend on the rapid assembly and disassembly of the actin filament (F-actin) cytoskeleton (Blanchoin et al., 2014). Under cellular conditions, F- actin assembly is favored (Brieher, 2013; Rottner and Stradal, 2011), making it critical to clarify how targeted and rapid F-actin disassembly occurs. In addition, specific extracellular cues including repellents such as ephrins, slits, semaphorins, myelin-associated inhibitors, and Wnts selectively collapse F-actin networks (Bashaw and Klein, 2010; Hung and Terman,

2011; Kolodkin and Tessier-Lavigne, 2011), but their direct effectors are still enigmatic. The best-known F-actin disassembly proteins, the ubiquitous ADF/cofilins, sever actin filaments and recycle monomers with a net effect of promoting new actin assembly (Bernstein and

Bamburg, 2010; Bravo-Cordero et al., 2013; Brieher, 2013; Rottner and Stradal, 2011).

Moreover, cofilin’s relatively weak disassembly of actin (Andrianantoandro and Pollard,

2006; Chin et al., 2016; McCullough et al., 2011) further complicates the current understanding of cellular F-actin destabilization.

Recently, we identified an unusual class of F-actin regulatory proteins, the MICALs, which are multidomain Redox enzymes that induce F-actin disassembly via the direct post- translational oxidation of actin (Hung et al., 2010, 2011). Notably, this Mical-modified actin no longer assembles normally (Hung et al., 2011, 2013), differentiating Mical’s effects from that of other F-actin disassembly proteins (Brieher, 2013; Rottner and Stradal, 2011).

Cellular and in vivo work has also revealed that MICALs are widely-expressed in different tissues (Giridharan and Caplan, 2014; Hung and Terman, 2011; Vanoni et al., 2013; Wilson et al., 2016; Zhou et al., 2011b) and control multiple cellular behaviors including motility,

22 axon guidance, synaptogenesis, immune responses, cardiovascular integrity, muscle function, and tumorigenesis (Van Battum et al., 2014; Beuchle et al., 2007; Hou et al., 2015; Hung et al., 2010, 2011, 2013; Kirilly et al., 2009; Lee et al., 2013; Lundquist et al., 2014; Terman et al., 2002; Wilson et al., 2016). The MICALs have also been identified as working with different growth factors, adhesion molecules, and repulsive guidance cues to exert their effects (Aggarwal et al., 2015; Van Battum et al., 2014; Hou et al., 2015; Hung et al., 2010;

Lundquist et al., 2014; Schmidt et al., 2008; Terman et al., 2002). Yet, nothing is known of how MICALs integrate with other better-known actin regulatory proteins to direct actin cytoskeletal reorganization and cellular functions.

We now find that Mical synergizes with the ubiquitous actin regulatory protein cofilin to dramatically enhance the dismantling of actin filaments. This coupling between Mical and cofilin depends on the Redox-mediated post-translational alteration of actin. Mical oxidation of actin improves cofilin binding to filaments accelerating F-actin severing and disassembly by over an order of magnitude compared to either effector alone. This synergism also regulates F-actin disassembly in vivo and serves to remodel cells, wire the nervous system, and orchestrate Semaphorin/Plexin repulsive signaling. The Redox-dependent synergy between Mical and cofilin, therefore, rapidly disassembles F-actin and also generates oxidized actin that re-assembles abnormally. This collective action has a net effect of promoting F-actin instability, revealing a previously unknown pathway of cellular F-actin disassembly.

23 Results

Cofilin modulates Mical Redox-mediated F-actin disassembly

Mical Redox enzymes are a new type of actin regulator – one that controls filament dynamics via the direct post-translational oxidation of actin (Hung et al., 2010, 2011).

Specifically, the enzyme activity of MICALs is activated in the presence of their substrate F- actin, which triggers consumption of Mical’s coenzyme NADPH and stereospecific oxidation of actin’s methionine (M) 44 and M47 residues to induce F-actin disassembly (Figure 2.1a;

(Alqassim et al., 2016; Hung et al., 2010, 2011, 2013; Lee et al., 2013; Lundquist et al.,

2014; McDonald et al., 2013; Vitali et al., 2016; Zucchini et al., 2011)). Mical’s characteristic consumption of NADPH in an F-actin dependent manner has thus provided a simple biochemical test for proteins that may affect Mical’s activity. We found that the well- known actin regulatory protein – cofilin (Bernstein and Bamburg, 2010; Bravo-Cordero et al., 2013) – strongly suppressed the ability of F-actin to trigger Mical-mediated NADPH consumption (Figures 2.1b-c and 2.3a).

The ubiquitous actin depolymerizing/severing factor cofilin is known to change the conformation of the D-loop of actin (Galkin et al., 2011), which harbors Mical’s substrate residues M44 and M47 (Hung et al., 2011). These results, coupled with the observation that non-muscle human cofilin-1 is a relatively weak severer of F-actin (Andrianantoandro and

Pollard, 2006; Chin et al., 2016; McCullough et al., 2011), prompted our investigation of a possible interrelation between Mical and cofilin effects on actin. In light of our NADPH consumption results (Figure 2.1b-c), we first wondered if cofilin affected Mical’s ability to bind to its substrate F-actin. However, using co-sedimentation assays we did not observe any

24 difference in the ability of Mical to associate with F-actin in the presence or absence of cofilin (Figure 2.1d). Therefore, we tested if cofilin affected Mical’s ability to disassemble

F-actin. Strikingly, we found that preincubation of F-actin with cofilin, which alone only minimally affects F-actin disassembly under these conditions (Figure 2.1e), dramatically enhanced Mical-mediated F-actin disassembly (Figure 2.1f-g). The rate of disassembly was greater than the combined rates with cofilin and Mical added individually (Figure 2.1h), which was also confirmed by co-sedimentation (Figure 2.1i). This cooperation was not observed in the absence of NADPH (sees Figure 2.3b-e), which rules out the possibility that cofilin and Mical without its NADPH coenzyme form a complex that is more efficient in F- actin dismantling than its individual components. Thus, cofilin enhances Mical-mediated actin filament disassembly and their synergistic effect requires the NADPH-dependent Redox activity of Mical.

Cofilin synergizes with Mical to accelerate F-actin disassembly

We therefore reasoned that cofilin might enhance Mical-mediated F-actin disassembly by allowing Mical to more efficiently oxidize its M44 and M47 substrate residues on actin (and thereby consume less NADPH in the process). To test for this possibility it was important to develop an independent assay for M44/M47 actin oxidation, since NADPH consumption is not an accurate measure of Mical-mediated F-actin oxidation and occurs to some extent even in the absence of F-actin (Figure 2.1c and (Hung et al.,

2011)). We found that the enzyme subtilisin, which under limited proteolysis conditions cleaves unoxidized actin between M47 and G48 (Schwyter et al., 1989), does not cleave

25 Mical-oxidized actin under such conditions (Figures 2.2a, 2.3f, and 2.4a-c). Using this observation as an assay, we found that cofilin strongly decreased Mical’s rate of F-actin oxidation (Figure 2.2b). Furthermore, generating antibodies that specifically recognized the wild-type (unoxidized) M44 residue of actin (Figure 2.4d) and the Mical stereospecifically oxidized M44 residue of actin (MetO-44) (Figure 2.2c), allowed us to confirm that cofilin does not increase the efficiency of Mical-mediated F-actin oxidation, but actually suppresses it (Figures 2.2d and 2.4e). Comparison of the time courses of Mical-mediated F-actin oxidation (Figure 2.2b and d) and F-actin disassembly (Figure 2.2e, left; and (Hung et al.,

2011)), indicated that Mical rapidly (~ 1 min) oxidizes F-actin but it takes hundreds of seconds for Mical-oxidized actin to disassemble. Strikingly, the addition of cofilin dramatically accelerated the disassembly of Mical-oxidized actin filaments (Figure 2.2e, right). Thus, Mical rapidly oxidizes but only relatively slowly disassembles filaments, and cofilin markedly accelerates this disassembly. These results are also consistent with cofilin’s suppressive effects on Mical-mediated NADPH consumption and actin oxidation (Figures

2.1c and 2.2b,d), because they reveal that Mical and cofilin combine to rapidly disassemble

(i.e., deplete) F-actin – which is Mical’s substrate and triggers Mical’s NADPH consumption and actin oxidation activities (Figure 2.1a).

Mical-mediated oxidation of actin weakens the mechanical properties of filaments

To more directly monitor and quantify the effect of Mical oxidation of actin and its disassembly by cofilin, we purified Mical-oxidized actin (Materials and Methods and (Hung et al., 2011, 2013)). We found that Mical-oxidized actin forms filaments, but such filaments

26 have altered polymerization kinetics and a critical concentration of at least an order of magnitude higher than that of unmodified actin (1μM) (Figures 2.5a-e and 2.4f; see also

(Hung et al., 2011, 2013)). Specifically, purified Mical-oxidized actin did not exhibit noticeable polymerization at 1.1 μM (Figure 2.5a; (Hung et al., 2011, 2013)), but did polymerize to increasing levels when incubated at 2.2 μM, 3.3 μM and 4.4 μM (Figure 2.5b- d). However, we found that polymerization of Mical-oxidized actin proceeded after a longer nucleation phase than normal and (consistent with the higher critical concentration) reached lower plateau levels than observed for unmodified actin (Figure 2.5b-d). Notably, re-treating the purified Mical-oxidized actin with Mical/NADPH did not alter its polymerization properties (Figure 2.5d), indicating that Mical-oxidized actin is not significantly reduced during purification and storage. Thus, above its critical concentration values, Mical-oxidized actin polymerizes but with abnormal kinetics indicative of the inhibited nucleation phase.

Further analysis of purified Mical-oxidized actin revealed that it also copolymerized with unoxidized actin monomers (Figure 2.5f; see also Fig. S11C of (Hung et al., 2011)).

We employed subtilisin digestion to quantify the extent of Mical-oxidized actin incorporation into such copolymers (Figure 2.5f). This allowed us to form and examine copolymers containing different and well-determined fractions of Mical-oxidized actin. Our results revealed that unlike unoxidized filaments, Mical-oxidized actin filaments easily fragment upon minimal handling (gentle pipetting and mixing). Even copolymers composed of low amounts of Mical-oxidized actin (11%) had a significantly lower mechanical stability than non-oxidized actin filaments (Figure 2.5g-h). Therefore, Mical-oxidized actin copolymers have different mechanical properties than non-oxidized actin.

Cofilin accelerates the dismantling of Mical-oxidized actin filaments

To directly assess the effect of cofilin on the disassembly dynamics of filaments composed of Mical-oxidized actin, we polymerized purified Mical-oxidized actin and employed time-lapse TIRF microscopy. We first grew filaments composed of 100% Mical- oxidized actin from unoxidized F-actin seeds. Dramatically, such Mical-oxidized actin filaments were rapidly dismantled by the addition of cofilin within the solution exchange time (~30 s) (Figure 2.6a, lower right) but not upon addition of buffer (Figure 2.6a, lower middle panel). Under the same conditions, F-actin severing in the presence of

Mical/NADPH or cofilin only was much weaker (see Figures 2.7a and 2.6a [compare upper right to lower right]). Thus, these results confirmed our observations using both pyrene-actin and actin sedimentation assays (Figures 2.1e-i and 2.2e) and demonstrated that cofilin markedly accelerates Mical-mediated F-actin disassembly.

Mical-mediated oxidation of actin increases cofilin’s binding and severing of filaments

We also examined the effects of partial Mical-oxidation on cofilin-mediated F-actin disassembly by employing copolymers with known amounts of Mical-oxidized actin incorporated. We found that even “lightly oxidized” F-actin copolymers (11% Mical- oxidized actin) accelerated cofilin severing by more than an order of magnitude (22-fold) compared to that of unmodified control F-actin (Figures 2.6b-c, 2.7e, and Supplementary

Movies 1-2). Increasing the content of the Mical-oxidized actin in the copolymers further accelerated cofilin severing and disassembly (Supplementary Movies S3-S4, Figure 2.6a,

28 compare upper right to lower right), and this effect was not cofilin isoform specific since we also observed it with yeast cofilin (Figure 2.7b-d). Thus, the presence of Mical-oxidized actin makes cofilin much more efficient at F-actin disassembly. Furthermore, when assisted by cofilin, partial oxidation of actin filaments by Mical is sufficient for their fast disassembly.

Additional analysis using two-color TIRF microscopy and co-sedimentation also indicated improved cofilin binding to filaments containing Mical-oxidized actin when compared to unoxidized control filaments (Figure 2.6d-e, Supplementary Movies 5-6,

Figures 2.8 and 2.9). In light of the extremely rapid nature of cofilin severing of Mical- oxidized F-actin, we quantified this improved cofilin binding by employing F-actin composed of either Q41C actin (yeast) or ANP-modified (skeletal) F-actin, since they both become disassembly-resistant when cross-linked between residues 41 and 374 (Figure 2.9).

Using this disassembly-resistant F-actin, we found that more cofilin co-sediments with filaments containing Mical-oxidized actin in comparison to unoxidized cross-linked control filaments (Figures 2.5d-e and 2.9) Therefore, Mical-oxidized actin increases both cofilin binding to filaments and the rate and extent of cofilin-mediated F-actin disassembly.

Cofilin modulates Mical-mediated Redox-dependent F-actin disassembly and cellular remodeling in vivo

In view of these results, we wondered if Mical and cofilin might also work together in vivo. Both cofilin and Mical have widespread effects on the organization of actin in vivo

(reviewed in (Bernstein and Bamburg, 2010; Bravo-Cordero et al., 2013; Giridharan and

29 Caplan, 2014; Hung and Terman, 2011; Vanoni et al., 2013; Wilson et al., 2016; Zhou et al.,

2011b)). For instance, Mical is required to shape Drosophila bristles, which are well- characterized cells (Figure 2.10a) that provide a high-resolution model to study actin organization and dynamics in vivo (Hung and Terman, 2011; Sutherland and Witke, 1999;

Tilney and DeRosier, 2005). Cofilin (which is encoded by the twinstar gene in Drosophila) is also required for shaping Drosophila bristles (Chen et al., 2001). Thus, we employed the bristle model to assay the interaction between Mical and cofilin in vivo.

Elevating the levels of Mical specifically in bristle cells results in F-actin disassembly and cellular remodeling (Figure 2.10b) that is dependent on Mical's Redox activity and its

M44 substrate residue within actin (Hung et al., 2010, 2011, 2013). Notably, cofilin and

Mical exhibited overlapping localization patterns within developing bristles (Figure 2.10c) and removing even a single copy of cofilin (cofilin heterozygous mutants) significantly suppressed the F-actin reorganization and bristle remodeling effects that are dependent on

Mical (compare Figure 2.10d with Figure 2.10b; Figures 2.10f-g and 2.12a). Moreover, raising the levels of cofilin significantly enhanced Mical-mediated effects on F-actin and cellular morphology (compare Figure 2.10e with Figure 2.10b; Figure 2.10f-g). Further analysis revealed that cofilin’s effects on Mical-mediated F-actin reorganization in vivo were dependent on Mical’s M44 substrate residue within actin (Figure 2.10h). Similarly, SelR

(MsrB), which is an enzyme that reverses Mical-mediated oxidation of actin (Hung et al.,

2013; Lee et al., 2013), reversed cofilin’s ability to enhance Mical’s effects on F-actin reorganization (Figure 2.10h). Thus, Mical-mediated F-actin alterations in vivo, as in vitro, are modulated by cofilin.

30 Mical and cofilin synergize to drive Semaphorin-Plexin repulsive signaling and axon guidance

In light of our in vitro and in vivo results demonstrating a synergistic action between

Mical and cofilin, it is notable that Mical and cofilin exhibit widespread overlapping expression patterns (Van Troys et al., 2008; Wilson et al., 2016) and both mediate the effects of growth factors, adhesion molecules, and guidance cues on diverse cellular behaviors

(reviewed in (Bernstein and Bamburg, 2010; Bravo-Cordero et al., 2013; Giridharan and

Caplan, 2014; Hung and Terman, 2011; Vanoni et al., 2013; Wilson et al., 2016; Zhou et al.,

2011b)). For instance, Mical associates with Plexins, which are receptors for one of the largest families of guidance cues – the Semaphorins (Semas), and plays critical roles in

Semaphorin/Plexin repulsive signaling (reviewed in (Hung and Terman, 2011)). Cofilin has also been linked to Semaphorin repulsion (Aizawa et al., 2001; Bribián et al., 2014; Hu et al.,

2001; Hung and Terman, 2011; Myster et al., 2015; Witherden et al., 2012), but its role and mechanisms of action in this regard have remained poorly understood. Since Mical-mediated bristle actin remodeling occurs in response to Semaphorin/Plexin repulsive guidance signaling (Hung et al., 2010, 2013), we wondered if cofilin could also be linked with Mical in mediating Semaphorin/Plexin repulsion.

To test this hypothesis, we first employed the bristle system and our genetic experiments demonstrated that cofilin was necessary for Semaphorin/Plexin/Mical-mediated effects on cellular remodeling (Figure 2.12b-c). Next, we turned to in vivo axon guidance assays using the Drosophila model nervous system, where Semaphorins-Plexins (Sema-1a and Plexin A) serve as repulsive axon guidance cues-receptors and were first functionally

31 linked to Mical (Terman et al., 2002). Notably, we found that cofilin (tsr) mutants exhibit axon guidance defects that are similar to loss of Sema-1a, Plexin A, and Mical (Figures

2.11a-c and 2.12d; (Terman et al., 2002; Winberg et al., 1998; Yu et al., 1998)).

Furthermore, we observed transheterozygous genetic interactions between cofilin and Mical mutants (Figure 2.11c), indicating they function in the same signaling pathway to mediate axon guidance. Moreover, we found that increasing the levels of cofilin enhanced Sema-

Plexin-Mical repulsive axon guidance, while decreasing the levels of cofilin suppressed these guidance effects (Figure 2.11d-f). These results further support that Mical and cofilin work together in vivo, as in vitro, and indicate that their synergistic effects are also instrumental for

Semaphorin-Plexin repulsive signaling and axon guidance.

Discussion

Here we have found that Mical and cofilin function as a pair – synergizing in a

Redox-dependent post-translational manner to disassemble F-actin and to control different cellular behaviors. Specifically, cofilin is a well-established actin regulatory protein and a relatively weak severer of F-actin (Andrianantoandro and Pollard, 2006; Chin et al., 2016;

McCullough et al., 2011). In contrast, Mical family Redox enzymes have only recently emerged downstream of Semaphorin-Plexin repellents as actin disassembly factors acting via the direct post-translational oxidation of actin (Alqassim et al., 2016; Hung et al., 2010, 2011,

2013; Lee et al., 2013; Lundquist et al., 2014; McDonald et al., 2013; Terman et al., 2002;

Vitali et al., 2016; Wilson et al., 2016; Zucchini et al., 2011). Previous work has also revealed that Mical, whose C-terminus associates with the intracellular portion of the

32 Semaphorin transmembrane receptor plexin (Hung et al., 2010; Terman et al., 2002), binds with its N-terminal NADPH-dependent Redox domain to F-actin and selectively oxidizes actin’s methionine-44 and 47 residues (Figure 2.11g, left panel; (Hung et al., 2010, 2011,

2013)). We propose that Mical oxidation-induced changes in filament structure and/or dynamics improve cofilin’s binding to actin filaments (Figure 2.11g, middle panel). Herein, we also find that Mical-oxidized actin co-polymers have different properties than unoxidized actin filaments. It is also known that the severing of actin filaments by cofilin is related to the mechanical properties of F-actin (McCullough et al., 2011; Ngo et al., 2015; Suarez et al.,

2011). Our results support the idea that Mical uses oxidation to weaken the inter-actin (inter- protomer) contacts within filaments ((Hung et al., 2011), present study) and these alterations dramatically speed up cofilin’s ability to break/dismantle filaments (Figure 2.11g, right panel). These results, therefore, uncover a previously unknown pathway of cellular F-actin disassembly and also present an unusual type of biological synergistic interaction – one involving two different types of proteins (Mical and cofilin) and the Redox-dependent post- translational modification of a third protein (polymerized actin).

Our results also shed new light on the mechanisms of action of both Mical and cofilin. They support a model that Mical and cofilin have been evolutionarily selected to work in tandem to ensure that even a low level of Mical activity in the presence of cofilin would facilitate F-actin disassembly, and vice versa. Moreover, unlike F-actin disassembly by cofilin, which promotes actin turnover by recirculation of monomers for polymerization

(Brieher, 2013; Kiuchi et al., 2007), Mical post-translationally modifies actin, decreasing its capacity for re-polymerization until the oxidation is reversed (Figure 2.11g, right panel).

33 Thus, the Redox-driven synergy between Mical and cofilin not only rapidly disassembles F- actin but also generates post-translationally modified actin that re-assembles abnormally with a net effect of promoting F-actin instability. These results, therefore, provide important insights into how actin-based structures are rapidly and specifically dismantled in cells.

Given their widespread overlapping expression patterns (reviewed in (Van Troys et al., 2008;

Wilson et al., 2016)) and diverse effects on cellular behaviors (reviewed in (Bernstein and

Bamburg, 2010; Bravo-Cordero et al., 2013; Giridharan and Caplan, 2014; Hung and

Terman, 2011; Vanoni et al., 2013; Wilson et al., 2016; Zhou et al., 2011b)), this synergistic interaction between Mical and cofilin provides the molecular framework to rapidly dismantle multiple actin-based cellular structures.

34 Figures

Figure 2.1. Mical/F-actin dynamics are modulated by cofilin. (a) Mical (1) physically associates with its substrate F-actin (2), which triggers Mical’s conversion/consumption of its co-enzyme NADPH to NADP+ (3). Mical then oxidizes F-actin subunits on their M44 and M47 residues (4) triggering F-actin disassembly. For simplicity, the presence of molecular oxygen (O2) and flavin adenine dinucleotide (FAD) have been excluded from this diagram. (b-c) Mical’s enzymatic activity (as determined by conversion of NADPH to NADP+, which is measured by a change in absorbance at 340 nm [NADPH Consumption]) is markedly accelerated by F-actin, but not when cofilin is present. [Mical]=600nM, [NADPH]=200μM, actin and cofilin were used at equal molar concentrations. n=3 independent experiments per condition. Mean +/- standard error of the mean (SEM). (d) Sedimentation/Association of Mical with F-actin is not altered by the addition of cofilin. S, soluble (G-actin); P, pellet (F- actin). [Actin]=4.6 µM; [Cofilin]=4.6 µM; [Mical]=2.4 μM. No NADPH present; n=3 independent experiments per condition. Mean +/- SEM. (e-h) Pyrene–actin assays, where the fluorescence (407 nm) is higher in the polymerized state. (e) Cofilin alone (at 1:10 mole ratio

35 to actin) has minimal effects on F-actin disassembly (pH 6.8). (f-h) Mical/NADPH-mediated F-actin disassembly (f) is rapidly accelerated by cofilin (at 1:10 molar ratio to actin) (g), resulting in a substantial increase in the change in pyrene-actin fluorescence/min (h). (i) Sedimentation of F-actin following short incubation times (3 minutes) with Mical/NADPH and/or cofilin. Sedimentation of actin shows an increase in the soluble (disassembled) actin amount following Mical/NADPH+cofilin treatment in comparison to Mical/NADPH treatment alone. For (e-i), [Actin]=2.5 µM; [Cofilin]=0.25 µM; [Mical]=10 nM; [NADPH]=100 µM. n=3 independent experiments per condition. Mean +/- SEM. See also Figure 2.13 for uncropped gels of d and i.

Figure 2.2. Cofilin slows F-actin oxidation by Mical but accelerates filament disassembly. (a) Subtilisin digestion of actin to assess its oxidation by Mical. (Top): Schematic representation of limited proteolysis of unmodified and Mical-oxidized actin with subtilisin. (Bottom): Subtilisin cleavage occurs between residues 47 and 48 in the D-loop of actin in unmodified actin monomers (red arrowhead), but not in Mical-oxidized actin (ox-G- actin). Cleavage time (0-15 min) is indicated; n=8 preps of Mical-oxidized actin. (b) Cofilin decreases Mical-mediated oxidation of F-actin, as assayed by limited proteolysis with subtilisin. Top panel (Mical/NADPH): Mical oxidation of bare F-actin. Subtilisin cleavage of bare actin (actincleaved), which is diagnostic for unoxidized actin, was abolished within 1 min of the addition of Mical/NADPH (oxidation time) due to the accumulation of oxidized actin. Bottom panel (+Cofilin): Mical oxidation of F-actin-cofilin complex (1:1 molar ratio). Significant amounts of subtilisin-cleaved actin (unoxidized actin) were detected even 30 min after the addition of Mical/NADPH indicating that cofilin strongly suppresses Mical- mediated actin oxidation. Conditions: [Actin]=3.5μM; [Mical]=25nM; [NADPH]=100µM; [Cofilin]=3.5μM; zero time points correspond to the limited proteolysis of unoxidized (non

37 Mical/NADPH-treated) actin using this approach. (c) Characterization of an antibody that specifically recognizes Mical-oxidized actin (actinMetO-44). This antibody recognizes Mical- treated actin but not untreated actin or Mical-treated actin following incubation with SelR, a reductase enzyme that reverses Mical-mediated actin oxidation (Hung et al., 2013). SelRC124S is an enzymatically-dead version of SelR that does not reduce Mical-oxidized actin (Hung et al., 2013). Specifically, 2.3μM of actin (Drosophila actin 5C) was polymerized with either 600nM Mical alone (untreated actin) or 600nM Mical/100μM NADPH (Mical-treated actin) for 1 hour at room temperature. Mical-treated actin was then incubated with 2.4μM of SelR or 2.4μM of SelRC124S and samples were subjected to SDS-PAGE and Western blotting with the actinMetO-44 antibody (see also Figure 2.4d). Similar amounts of actin (lower panel) are present in all experiments. (d) Cofilin suppresses Mical-mediated oxidation of actin, as observed using the actinMetO-44 antibody. [Actin]=1.15µM; [Cofilin]=1.15µM; [Mical]=50nM; [NADPH]=100µM. (e) Mical induces F-actin disassembly (left), while the addition of cofilin (right) rapidly accelerates Mical/NADPH-mediated F-actin disassembly. [Actin]=2.5µM; [Cofilin]=0.25µM; [Mical]=10nM; [NADPH]=100µM. See also Figure 2.13 for uncropped gels/blots of a-d.

Figure 2.3. Further characterization of the interaction of Mical and cofilin in modulating F-actin disassembly and the quantification of Mical-oxidized actin. (a) Cofilin alone does not modify Mical’s enzymatic activity, unlike Mical’s substrate F-actin (Figure 2.1b-c; (Hung et al., 2011)), or cofilin in the presence of F-actin (Figure 2.1b-c). Mean +/- SEM. n=3 independent experiments per condition. [Mical]=600 nM, [NADPH]=200 µM. (b-e) Pyrene actin assays with different combinations of cofilin, Mical, NADPH, and cofilin buffer. Compare with Figure 2.1e-i. Note, that the enhanced effect of cofilin on Mical-mediated actin filaments disassembly is not observed in the absence of Mical’s coenzyme NADPH. [Actin]=2.5 µM; [Cofilin]=0.25 µM; [Mical]=10 nM;

39 [NADPH]=100 µM. n=3 independent experiments per condition. (f) Limited proteolysis assay with subtilisin allows quantification of Mical-oxidized actin. This assay is based on the unique feature of Mical-oxidized actin (ox-actin) - its resistance to limited proteolysis by subtilisin that normally occurs between residues 47 and 48 on actin (Schwyter et al., 1989). F-actin and F-actin complexes with its binding partner (such as cofilin) are oxidized by Mical in the presence of NADPH. At the selected time points, oxidation is stopped with a large excess of NADP+ (product of the reaction) and the actin depolymerizing reagent Kabiramide C (KabC) (Klenchin et al., 2003). The resulting samples are subjected to limited proteolysis with subtilisin. Subtilisin A (type VIII, Bacillus licheniformis, 11.8-12 units/mg solid) was purchased from Sigma (P5380). The amount of ox-actin in the resulting samples was determined by SDS-PAGE and densitometry analysis. See also Figure 2.4a-c.

Figure 2.4. Further characterization of Mical-oxidized actin using a limited proteolysis assay with subtilisin and an antibody directed against the Met-44 residue of actin. (a-c) Development of limited proteolysis-based assay for quantification of Mical-oxidized actin (see also Figure 2.3f). Assay was developed based on the following observations: 1) Mical oxidizes actin at residue 47 (D-loop) (Hung et al., 2011); 2) subtilisin cleaves actin between residues 47 and 48 (Schwyter et al., 1989); 3) under our experimental conditions Mical

41 oxidized actin (ox-actin) cannot be cleaved by subtilisin at position 47/48, as opposed to unmodified actin, G-actin-cofilin, and G-actin-KabC complex; 4) Mical oxidation of actin can be inhibited by a large excess of the product of the reaction (NADP+); and 5) a combination of NADP+ and the F-actin depolymerizing agent KabC (Klenchin et al., 2003) stops Mical oxidation of actin. (a). The small actin sequestering molecule KabC does not affect limited digestion (proteolysis) of actin by subtilisin compared to G-actin alone; ox- actin – Mical-oxidized G-actin; cofilin – human cofilin-1; Digestion time (in minutes) is indicated in the panel. Conditions: 2 mM Tris, pH 8, 0.2 mM CaCl2, 0.2 mM ATP, 1 mM DTT; 1:1000 subtilisin:actin w/w ratio. (b) Mical oxidation of actin/NADPH consumption can be stopped by an excess of NADP+ (product of the reaction) in combination with KabC. NADPH consumption was followed by a decrease of its fluorescence at 460 nm (excitation wavelength was 340 nm). [F-actin] = 3.5 μM, [NADP+] = 1.5 mM, [KabC] = 3.5 μM, buffer: 20 mM imidazole, pH 6.8, 50 mM KCl, 2 mM MgCl2, 0.2 mM EGTA, 0.2 mM ATP, 1 mM DTT. (c) G-actin-cofilin complexes have the same subtilisin digestion pattern as uncomplexed G-actin; cofilin – human cofilin-1. [F-actin] = 3.5 μM, [cofilin] = 3.5 μM, [NADP+] = 1.5 mM, [KabC] = 3.5 μM, 1:200 subtilisin:actin w/w ratio. Conditions are the same as in (b). (d-e) Cofilin suppresses the Mical-mediated oxidation of actin, as observed using an antibody directed against the Met-44 residue of actin. (d) Characterization of an antibody that specifically recognizes the Met-44 residue of actin. Mical oxidizes actin on its Met-44 and Met-47 residues, but the oxidation of the Met-44 is instrumental for inducing F- actin disassembly (Hung et al., 2011). This Met-44 residue of actin is conserved in all actins from yeast to humans (as are almost all of the surrounding residues) (Hung et al., 2011). While attempting to generate antibodies to Mical-oxidized actin (using a peptide of actin residues 38-51), we identified anti-sera that preferentially recognized untreated actin from Mical-treated actin. In particular, note that the antibody (actinMet-44 antibody) preferentially recognizes untreated (Mical only) versus treated (Mical/NADPH) actin (d1, upper panel). This antibody specifically recognizes the Met-44 residue of actin since it does not recognize actin when the Met-44 residue is substituted with Leucine (M44L actin; d2), but does recognize actin when the Met-47 residue is substituted with Leucine (M47L actin; d2). Moreover, Mical-treated actin is recognized by the actinMet-44 antibody following treatment with SelR (d3), the reductase that reverses Mical-mediated oxidation of actin (Hung et al., 2013). Note that similar amounts of Mical (d1, lower panel) and actin (d1-d3, lower panels) are present in all experiments. For western blotting, 2.3 μM of Drosophila actin (actin 5C) was polymerized with either 600 nM Mical alone (untreated [Mical only] actin) or 600 nM of Mical and 100 μM of NADPH (treated [Mical/NADPH] actin) for 1 hour at room temperature. Then, Mical-treated actin was also treated with 2.4 μM of SelR (that we had previously generated (Hung et al., 2013)) in a buffer containing 20 mM DTT and 10 mM MgCl2 for 1 hour at 37°C. All samples were mixed with SDS-PAGE loading buffer, boiled for 5 min, and loaded onto a 12% SDS-PAGE. After transferring the proteins to a PVDF membrane, this membrane was blocked with 5% non-fat milk in PBST buffer for 1 hour. This membrane was then incubated with either a pan actin antibody (C4; Millipore) or actinMet-44 antiserum for 1 hour at room temperature, and followed with standard Western blotting procedures. Other mutant actin proteins that we had previously generated (Hung et al., 2011) were also employed in our experiments, including both the Mical-resistant M44L

42 and M44LM47L actins and the M47L actin. (e) Cofilin suppresses the Mical-mediated oxidation of actin, as observed using this actinMet-44 antibody (note the increased presence of actin that is recognized by this antibody in the presence of cofilin). [Actin]=1.15 µM; [Cofilin]=1.15 µM; [Mical]=50 nM; [NADPH]=100 µM. (f) Examples of the gels used to determine the Cc of purified Mical-oxidized actin (ox-actin) are shown in Figure 2.5e (pH 8.0, quantified in Figure 2.5e, blue circles). Total concentrations of ox-actin are indicated for each sample. S – Supernatant, P – Pellet. The determined Cc was ≥1 μM, which is consistent with the data shown in Figure 2.5a-e. Thus, below its Cc values Mical-oxidized actin will not form filaments under polymerizing conditions. See also Figure 2.13 for uncropped gels of d-f.

Figure 2.5. Mical-mediated oxidation of actin alters polymerization and weakens the mechanical properties of filaments. (a-d) Purified Mical-oxidized actin can be induced to polymerize when incubated at high-enough concentrations, although with altered kinetics and extent. Pyrene-actin and cosedimentation (insets, a-b) assays show that purified Mical- oxidized actin (ox-actin) does not polymerize at 1.1μM (a; see also (Hung et al., 2011,

44 2013)), but does polymerize to increasing levels when at concentrations of 2.2μM and 4.4μM (b-c). (d) Re-treating purified Mical-oxidized actin with Mical/NADPH (lower arrowhead) does not alter its polymerization state (compare with untreated actin [green curve], upper arrowhead). [Mical]=600nM; [NADPH]=100µM. Representative SDS-PAGE gels: S, soluble (G-actin); P, pellet (F-actin). (e) Critical concentration (Cc) of Mical-oxidized actin (ox- actin) is at least one order of magnitude higher than that of unoxidized actin. For ox-actin, intersects of linear plots of concentrations of pelleted F-actin versus total actin with the abscissa yielded a Cc value at pH 7 of 1.1μM±0.25 standard deviation (SD) (n=3 independent ox-actin preps) (red circles) and had similar values at pH 6.8-8. Unoxidized actin Cc was close to 0.1µM. Linear fits are shown for Mical-oxidized and unoxidized actin in zoomed inset. (f) Quantification of copolymers content using the subtilisin limited proteolysis assay reveals that polymerization of actin mixtures containing unoxidized actins and 25% and 50% of Mical-oxidized actin (ox-actin) yielded copolymers with 10.8±3.2% and 27.7±1.6% ox-actin, respectively (mean+/-SD). (g-h) Copolymers of Mical-oxidized (11%) and unoxidized actin show decreased mechanical stability compared to unmodified actin. (g) No statistically significant differences in average length of non-oxidized versus 11% Mical-oxidized F-actin were observed when filaments were assembled in flow chambers (no mixing). n=3 independent measurements of 26-129 filaments per condition per repeat; Mean+/-SEM; NS (not significant) using Student’s t-test (two-tailed). (h) In contrast, even minimal handling (gentle pipetting and mixing) decreases the average length of Mical- oxidized actin (11%) copolymers much more than unoxidized F-actin. Filament length distributions. Student’s t-test (two-tailed). P=0.0059. Mean+/-SD. n=3 independent measurements of 69-188 unoxidized actin filaments (top panel) and 107-182 11% Mical- oxidized copolymer (bottom). See Figure 2.13 for uncropped gels of a-b, f.

Figure 2.6. Mical oxidation of F-actin improves cofilin binding and results in accelerated filament severing. (a) Rapid disassembly of fully oxidized actin by human cofilin. Unoxidized Cy3-F-actin seeds were introduced on the slide surface (red filaments, unoxidized) and extended with 100% Mical oxidized actin (ox-actin) labeled with Alexa488 (green stretches, oxidized). Addition of 10 nM human cofilin-1 (but not buffer) to such filaments resulted in full dismantling of Mical ox-actin stretches (green) within the mixing time (~30 sec), but unoxidized actin (red stretches) was not disassembled and stayed on the surface (bottom panel). No cofilin severing/fragmentation of control (unoxidized 2-colored filaments) was observed under identical conditions (top panel). Scale bar=10 μm. (b-c) Enhanced cofilin severing of Mical-oxidized actin containing filaments. (b) Severing events are indicated with magenta arrowheads. Top panel: Severing of F-actin with human cofilin-1 (100 nM) over time. Bottom panel: Severing of F-actin copolymers containing Mical- oxidized actin (11%) by human cofilin-1 (100 nM) over time. Scale bar=5 μm. (c) Quantification of cofilin severing of unoxidized F-actin and 11% Mical-oxidized (ox-actin) copolymers. Mean +/- standard deviation (SD). Number of filaments analyzed is 43-45 copolymers and 31-42 unoxidized polymers from each of 3 independent experiments (n=132 copolymers analyzed and 112 unoxidized polymers analyzed). The result of Student’s t-test (two-tailed) is shown (p=0.004). (d-e) Improved binding of cofilin to Mical-oxidized Cu2+- cross-linked Q41C F-actin, which is disassembly-resistant. (d) Representative SDS-PAGE gel of Mical-oxidized actin pelleted with cofilin. Before gel analysis disulfide cross-linking in Q41C actin was reversed with beta-mercaptoethanol. Actin (A). The bottom bands in the gel are cofilin (C). S, soluble (G-actin); P, pellet (F-actin). (e) Quantification of cofilin that co-sedimented with unoxidized and Mical-oxidized (ox-actin) Q41C cross-linked F-actin. Mean +/- SD. n=3 independent experiments per condition. The result of Student’s t-test (two-

46 tailed) is shown (p=0.0005). Also see Figure 2.9. See also Figure 2.13 for uncropped gel of d.

Figure 2.7. Mical oxidation of actin filaments accelerates their severing by yeast and human cofilins. (a) Severing of actin filaments in the presence of Mical/NADPH. Average of 3 independent trials is shown (analyzing 10-13 filaments each, n=3 movies). Filaments were oxidized on-slide by addition of Mical (55 nM) and NADPH (100 μM) into the flow chamber. Filaments were allowed to oxidize for 1 min then movies were recorded. Note, that the oxidation-induced severing of F-actin by Mical can be followed for minutes (see also Ref ((Hung et al., 2011)) as opposed to its almost instant disassembly when combined with cofilin (compare to Figure 2.6a). (b-d) Severing of unmodified F-actin (b) or its copolymer (11%) with Mical-oxidized actin (c) with yeast cofilin. Scale bar = 5 μm (d) Time dependant change in the average length of skeletal F-actin and 11% Mical-oxidized actin copolymers in the presence of yeast cofilin. A dramatic acceleration of severing by cofilin was observed with 11% Mical-oxidized actin copolymer (c-d). This confirms that accelerated cofilin severing of 11% Mical-oxidized copolymer is not isoform specific (see Figure 2.6b-c for comparison). Conditions: [Actin] = 1 μM (10% Cy3-maleimide labeled), [cofilin] = 3.3 nM, pH 6.8. (e) Determining the rates of human cofilin-1 severing of unmodified F-actin and its copolymer with Mical ox-actin from the TIRF data (related to the Figure 2.6c). The plots are showing averages of 3 independent repeats for each condition. Red solid lines correspond to the linear fits used to determine maximum cofilin severing rates in Figure 2.6c. Note, that in

48 the case of the copolymers with Mical-oxidized actin, cofilin severing efficiency decreases over time (open circles). This effect is due to substantial shortening of actin filaments upon their extensive severing by cofilin within the observation time. Cofilin severs long filaments more efficiently than short filaments (McCullough et al., 2011). Since under our conditions such shortening is insignificant in the control sample (unoxidized actin), the dependence of the number of severing events per μm versus time is linear. To determine the maximum severing rates, linear parts of the traces were fitted individually for each repeat and the slopes were averaged (results are shown in Figure 2.6c). n=3 independent experiments per condition. Conditions: [Actin] = 1 μM (10% Cy3 labeled), [Cofilin] = 100 nM, pH 6.8.

Figure 2.8. Indication of the enhanced cofilin severing and binding to actin filaments containing oxidized actin. This figure presents analysis of the data from Supplementary Movies 5 and 6. [Actin] = 0.6 μM, [yeast cofilin] = 38.5 nM, pH 6.8. One time point image (at 100 sec) from the corresponding movie is shown next to each graph (Scale bar = 10 μm). After background subtraction (Rolling Ball Radius algorithm, 10 pxl), total fluorescence intensity per frame was determined in both channels (actin-Alexa488 (green) and cofilinCy5 (red)) and plotted versus time. The increase in fluorescence intensity is due to an increase in actin polymer mass (Alexa488 channel) and the mass of cofilin bound to these polymers (Cy5 channel). Note, that in both (a) and (b) the fluorescence intensity corresponding to actin and cofilin increases non-linearly within the observation time. This effect is due to the formation of new barbed ends as a result of cofilin severing activity. This tendency is more pronounced in the case of Mical-oxidized actin (ox-actin) copolymers (note the differences in the y axis scale between (a) and (b)). We determined the rates of fluorescence intensity change in both channels for unmodified actin and its copolymer with Mical ox-actin (11%). Changes in fluorescence intensity per second (which can be approximated by linear fits

50 within chosen windows of time) are shown for two time intervals (0-90 sec and 130-225 sec) for each channel (solid black lines). Numbers shown next to each fit correspond to their slopes. Due to undersaturating concentrations of cofilin (38.5 nM), total actin fluorescence per frame increases faster than that of cofilin. Note, that in the control population (unmodified F-actin) actin-Alexa488 fluorescence increases ~3-4 fold faster ([1.5x103/0.5x103]=3 or [2.5x103/0.7x103]=3.6) than that of cofilinCy5 ((a), Supplementary Movie 5). However, in the copolymer sample, fluorescence of actin-Alexa488 increases only ~2 fold faster ([4.0x103/2.0x103]=2 or [9.6x103/4.6x103]=2.1) than that of cofilinCy5 ((b), Supplementary Movie 6). Thus, this decreased difference between cofilin and actin fluorescence in the Mical ox-actin copolymer sample is consistent with the presence of more cofilin associated with Mical-oxidized F-actin than with unmodified F-actin. This prompted our investigation of cofilin binding to the Mical-oxidized cross-linked F-actin (see Figure 2.6d-e and Figure 2.9).

Figure 2.9. Further characterization of cofilin binding to Mical-oxidized actin. (a-d) Enhanced cofilin binding to cross-linked Q41C F-actin after its oxidation by Mical (see also Figure 2.6d-e). (a) Positions of the cross-linked residues (41 (red) and C374 (blue)) in F- actin (PDB: 3J8A)(von der Ecken et al., 2015). Adjacent longitudinal protomers within the same actin strand are shown in grey. Actin third protomer from the opposite strand (lateral self-interacting interface) is shown in dark green. In the Q41C yeast actin mutant, residue 41 was mutated to cysteine which allows for disulfide cross-linking of this residue to native cysteine 374 on another protomer. (b) Representative SDS-PAGE gel under non-reducing (Coomassie stained) conditions demonstrates that Q41C actin is fully cross-linked by Cu2+ under our experimental conditions. Monomeric actin (left lane) is depleted upon CuSO4 addition (right lane) due to the formation of higher order actin species. (c) Cross-linked (XL) Q41C-F-actin can be fully oxidized by Mical. Efficiency of Q41C-XL oxidation by Mical was assessed by limited proteolysis with subtilisin. To this end, Q41C F-actin cross-linking was reversed with DTT after the indicated oxidation time (to allow it to appear as monomeric

52 actin on the gel). Based on limited proteolysis with subtilisin, the increase in intensity of the uncleaved actin band over time reports on the progress of actin oxidation by Mical (ox-actin). Q41C-XL F-actin can be efficiently oxidized by Mical in the presence of NADPH (close to completion in 15 min). For cosedimentation assays with cofilin (Figure 2.6d-e), Mical oxidation of Q41C-XL F-actin was carried out for 1 hour at room temperature to ensure full oxidation. (d) Yeast cofilin binds tighter to the Mical-oxidized cross-linked Q41C F-actin (3.5 μM) than the unoxidized control. Note the absence of unbound cofilin (at 2.5 and 3.5 μM concentration) in the supernatants of its complexes with ox-actin in contrast to the complexes with unoxidized actin. Example of the SDS-PAGE gel under reducing conditions is shown (Coomassie staining). After Mical oxidation and pelleting (P) with cofilin, Q41C actin cross-linking was reversed in the presence of 2-mercaptoethanol and it now appears as a single band on the gel (compare to (b, right)). Note, that judging by the absence of actin in supernatants (S), complete C41-C374 cross-linking produces actin polymers that are disassembly-resistant despite their full oxidation and cofilin presence. Enhanced cofilin binding to the oxidized Q41C-XL (compared to unoxidized) was observed at three different ratios of cofilin:actin (0.7:1; 1:1; 2:1). The ratio used in Figure 2.6d-e (1:1) is indicated with the red arrows. (e) Co-sedimentation indicates improved binding of human cofilin-1 to Mical-oxidized ANP cross-linked skeletal F-actin (30% ox-actin) compared to the unoxidized ANP cross-linked F-actin. Unoxidized and Mical-oxidized ANP-cross-linked F- actin was pelleted at high speed and analysed as described in the Methods. Conditions: [ANP-cross-linked F-actin] = 3.5 μM, [Cofilin] = 3.5 μM; Buffer: 20 mM imidazole, pH 6.8, 2 mM MgCl2, 0.2 mM EGTA, 150 mM KCl, 0.2 mM ATP, 0.5 mM DTT. Mean +/- SD (n=4 independent experiments per condition); the result of Student’s t-test (two-tailed) is shown (p=0.001). See also Figure 2.13 for uncropped gels of b-d.

Figure 2.10. Cofilin enhances Mical-mediated F-actin alterations in vivo. (a-b) Drosophila bristles are unbranched (a) but become branched as the result of F-actin disassembly and remodeling (b; arrowheads and drawings) when Mical is overexpressed specifically within them (Hung et al., 2010, 2011, 2013). (c) Mical (red; see also (Hung et al., 2010)) and cofilin/twinstar (green) are both expressed in bristle processes in overlapping patterns. Note also that cofilin is more widely distributed than Mical, which shows its highest distribution at the tip of the process. Scale bars=10 μm (d) Decreasing the levels of cofilin (cofilin/twinstar heterozygote genetic background [cofilin+/–]) suppresses Mical-induced F- actin reorganization/bristle branching (arrowheads and drawings). (e) Increasing the levels of cofilin/twinstar (bristle specific expression of a hyperactive cofilinS3A transgene [cofilin+++], which has no bristle effects on its own) enhances Mical-induced F-actin

54 reorganization/bristle branching (arrowheads). (f-g) Quantification of the data from b, d and e. n=20 bristle cells accessed across 20 animals per genotype, Mean +/- SEM, Student’s t- test (two-tailed); ***p=0.0008, ****p<0.0001. (h) Employing the genetic background described in (e), we find that mutating Mical's substrate residue on actin, the Met-44 residue, and expressing this mutant actin in bristles (actinM44L) suppresses cofilin’s effects on Mical. Likewise, expressing SelR (SelR+++), but not an enzyme dead version of SelR (SelRC124S), suppresses cofilin’s effects on Mical. n=40 bristle cells accessed across 10 animals per genotype. Mean +/- SEM, Student’s t-test (two-tailed); ****p<0.0001. All genotypes are heterozygous (B11-GAL4/+, UAS:Mical/+, mutations/+, and/or transgenes/+). One copy of UAS:GFPactin was used when visualizing F-actin. B11-GAL4, UAS:Mical, UAS:ActinM44L, UAS:SelR, UAS:GFPactin, and UAS:SelRC124S lines were as previously described (Hung et al., 2010, 2011, 2013). For Mical/cofilin expression analysis, we crossed UAS:mCherryMical, B11- GAL4 flies to cofilin/twinstar (tsr) GFP-trap lines and Mical/cofilin expression and localization was imaged in pupal progeny. We used the following cofilin (twinstar [tsr]) publicly available fly lines: tsrN121 (a loss-of-function/”knockout” adult lethal mutant due to P-element mediated imprecise excision in the tsr gene; (Johnson et al., 2008; Ng and Luo, 2004; Ren et al., 2007; Schottenfeld-Roames et al., 2014)) and UAS:tsrS3A (a constitutively active tsr transgene; (Ng and Luo, 2004; Stephan et al., 2012)). We also used the following GFP-trap tsr lines, all of which showed similar expression patterns: ZCL2393, tsrCPTI002237, and CC01393.

Figure 2.11. Cofilin enhances Sema-Plexin-Mical repulsive axon guidance. (a) In wild- type embryos, Drosophila intersegmental nerve b (ISNb) motor axons innervate muscles 6 and 7 (filled arrowhead) and muscles 12 and 13 (open arrowhead). This normal pattern of innervation is also depicted in the drawing. (b) In a cofilin (twinstar) homozygous mutant embryo (cofilin–/–, adult-lethal genotype), note the absence (filled arrowheads) or abnormal (open arrowhead) innervation of these muscles. (c) Quantification of the data from a-b, reveals that cofilin–/– mutant embryos exhibit significant ISNb axon guidance defects (left graph). Embryos with heterozygous mutations for both cofilin and Mical (cofilin+/– and

56 Mical+/–) also exhibit significant ISNb guidance defects in comparison to either heterozygote alone (right graph). Examination of another motor axon pathway (segmental nerve a [SNa]) revealed similar significant differences. n=100 hemisegments assessed across 10 animals per genotype, Mean +/- SEM, Chi-Square Test; ****p<0.0001. (d) Wild-type Drosophila central nervous system (CNS) axons exhibit a characteristic organizational pattern including three longitudinal connectives (1, 2, 3) composed of bundled Fasciclin II (1D4)-positive axons. Increasing the levels of PlexA in combination with Mical in neurons (Neuronal PlexA [PlexA+++] and Neuronal Mical [Mical+++]) alters the pathfinding of these longitudinal axons (e.g., arrow, arrowhead) and creates a sensitive genetic background to quantify CNS axonal pathfinding defects including discontinuous or missing 1st, 2nd, or 3rd CNS longitudinals and/or axons crossing the midline (see also (Ayoob et al., 2004; He et al., 2009; Hung et al., 2013; Yang and Terman, 2012)). Increasing the levels of cofilin (+Neuronal cofilin [cofilin+++]) enhances these PlexA-Mical dependent effects, while decreasing the levels of cofilin (+cofilin+/–) suppresses these PlexA-Mical dependent effects. Scale bar applies to all images. (e-f) Quantification of the data from d. n = 480 longitudinals accessed in 160 hemisegments within 10 animals per genotype, Mean +/- SEM, Student’s t-test (two- tailed); ****p<0.0001. All genotypes are heterozygous (ELAV-GAL4/+, UAS:HAPlexA/+, UAS:Mical/+, mutations/+, and/or transgenes/+). (g) A model based on our in vitro and in vivo results that Mical and cofilin form a Redox-driven synergistic pair to negatively affect the stability of the actin cytoskeleton and direct F-actin dismantling, cellular remodeling, axon guidance, and Semaphorin-Plexin repulsion. See also main text.

Figure 2.12. Further analysis of Cofilin’s effects on Mical and Semaphorin/Plexin- mediated F-actin/cellular remodeling in vivo. (a) Knockdown of cofilin specifically in bristles using an RNAi transgenic line specific for cofilin/twinstar suppresses Mical-induced F-actin reorganization/bristle branching. Compare with Figure 2.10g. Mean +/- SEM. Student’s t-test (two-tailed); ****P<0.0001. n=20 animals per genotype. Replicated in at least 2 independent experiments (separate crosses) per genotype. (b) Increasing the levels of Plexin (Plexin A [PlexA]) in bristles generates bristle branching (Hung et al., 2010). Decreasing the levels of cofilin (cofilin/twinstar heterozygote genetic background) suppresses these Plexin-induced F-actin reorganization/bristle branching effects. Percent of flies with branched bristles. Chi-Square Test; ****P<0.0001. n=40 animals per genotype. Replicated in at least 2 independent experiments (separate crosses) per genotype. (c) Increasing the levels of both Plexin and Mical within bristles generates increased F-actin remodeling/bristle branching in a Semaphorin-dependent manner (Hung et al., 2010), and these Semaphorin/Plexin/Mical-dependent effects are suppressed by decreasing the levels of cofilin (cofilin/twinstar heterozygote genetic background). Mean +/- SEM, Student’s t-test (two-tailed); ****P<0.0001. n=20 animals per genotype. Mean + standard error of the mean (SEM). Replicated in at least 2 independent experiments (separate crosses) per genotype. (d)

58 Knockdown of cofilin specifically in neurons using the ELAV-GAL4 driver and an RNAi transgenic line specific for cofilin/twinstar generates motor axon guidance defects similar to those described in Figure 2.11b-c. n=100 hemisegments assessed in 10 animals per genotype, Chi-Square Test; **** P< 0.0001. All genotypes in a-d are heterozygous (B11- GAL4/+, ELAV-GAL4/+, UAS:Mical/+, UAS:PlexA/+, tsr/+, and/or cofilin RNAi/+). The twinstar (tsr) RNAi knockdown lines employed were tsrHMS00534 (obtained from the Bloomington Drosophila Stock Center) and tsrKK108706 (obtained from the Vienna Drosophila Resource Center). The UAS:HAPlexA and UAS:Mical lines were as previously described (Hung et al., 2010, 2011, 2013). Other lines were as described in Figures 2.10 and 2.11.

Figure 2.13. Uncropped Gels. Uncropped gels for Figures 2.1d (a, red boxes), 2.1i (b), 2.2a (c), 2.2b[upper] (d, red boxes), 2.2b[lower] (e, red boxes), 2.2c[upper] (f), 2.2c[lower] (g, red

60 boxes), 2.2d (h), 2.5a-b[inserts] (i, red boxes), 2.5f (j, red boxes), 2.6d (k, red boxes), 2.4a (l), 2.4c (m, red boxes), 2.4d[upper left] (n), 2.4d[lower left] (o), 2.4d[upper middle and right] (p), 2.4d[lower middle and right] (q), 2.4e (r, red boxes), 2.4f[upper] (s), 2.4f[lower] (t), 2.9b (u, red boxes), 2.9c (v, red boxes), and 2.9d (w).

61 Materials and Methods

Protein purification.

Drosophila MicalredoxCH construct (referred to as Mical in this study) (Hung et al., 2010; Wu et al., 2016) rabbit skeletal actin (Spudich and Watt, 1971), Drosophila actin mutant

M44L/M47L (Hung et al., 2011), yeast actin (Grintsevich et al., 2008), and human cofilin-1

(Grintsevich and Reisler, 2014) were expressed and purified as previously described. Yeast cofilin was expressed and purified essentially as described (Bobkov et al., 2002). In brief, yeast cofilin expression was induced at OD600=0.8 with 1mM IPTG and carried-out for 4 hours at 37°C. Cell lysate was loaded on QAE-52 column equilibrated with 20mM Tris–HCl

(pH7.5 at 4°C) containing 1mM DTT, 0.2mM PMSF and cofilin was eluted with linear gradient of NaCl (0-500mM) in 5 column volumes. Cofilin containing fractions were then gel-filtered on HiLoad 16/60 Superdex 75 (Amersham Biosciences) column equilibrated with

20mM Tris–HCl (pH7.5 at 4°C), 200mM NaCl, 1mM DTT, 0.2mM PMSF. Purified cofilin was stored at -80°C.

Mical-oxidized actin purification.

Rabbit skeletal G-actin was polymerized at 20μM for 1 hour at room temperature (RT)

(buffer composition: 5mM Tris, 0.2mM CaCl2, 0.5mM DTT, 0.2mM ATP, 2mM MgCl2,

50mM KCl, pH8). Polymerized F-actin was then diluted to 2μM and supplemented with

NADPH (0.4mM) and Mical 0.2μM (10:1 molar ratio, actin to Mical, unless stated otherwise). Mical-oxidation of actin was carried out for 2 hours at RT. Oxidation efficiency under chosen conditions was confirmed by mass spectrometry. After 2 hours any residual F-

62 actin was pelleted at 100,000g for 20 min at 4°C. Resulting supernatant containing Mical- oxidized actin (ox-actin) was dialyzed overnight into buffer G (GB2): 2mM Tris, 0.2mM

CaCl2, 0.5mM DTT, 0.2mM ATP, pH 8. Actin was gel filtered using Superdex S200 16/60 column. Efficiency of oxidation was confirmed in subtilisin digestion assay (Figure 2.4a).

Mical-treated/oxidized pyrene actin was purified as described (Hung et al., 2011,

2013). To examine repolymerization of Mical-treated actin, the purified actin was resuspended to 2.3μM in GB5, and polymerization was initiated with 2X polymerization buffer (10mM Tris-HCl pH 7.5, 0.1M KCl, 4mM MgCl2, 2mM EGTA, 0.4mM ATP, 1mM

DTT) to get a final concentration of 1.15μM actin. To further test the ability of Mical-treated actin to repolymerize, the purified actin was resuspended to 4.4μM or 8.8μM in GB5, and polymerization was initiated as described above with 2X polymerization buffer (to yield a final concentration of actin at 2.2μM and 4.4μM, respectively). To determine whether Mical- oxidized actin might be reduced during its purification and storage, purified Mical-treated actin was polymerized (as described above), and then re-treated with 600nM of Mical and

200μM NADPH. Polymerization was monitored using either fluorescence or sedimentation assays (described below).

Critical concentration (Cc) determination.

To determine the Cc, Mg-ATP-ox-actin was polymerized for 1 hour at RT by adding 10X polymerizing buffer, pH 6.8, 7.0, 7.5 or 8.0. Samples were diluted then into their corresponding 1X polymerizing buffer (pH 6.8 - 8.0), followed by 4°C overnight incubation.

Supernatants and pellets were separated by ultracentrifugation (TLA100, 62K, 30 min, 4°C)

63 and analyzed by SDS-PAGE. Gels were stained with Coomassie Blue and densitometry was performed using Scion Image software. The intersects of these linear plots of pelleted actin

([F-actin]) versus total actin ([Actin(total)]) with the abscissa yielded Cc in μM. The following buffers were used for Cc experiments: pH 6.8: 20mM imidazole, pH 6.8, 2mM

MgCl2, 0.2mM EGTA, 50mM KCl, 0.2mM ATP, 0.5mM DTT; pH 7.0: 10mM Hepes, pH

7.0, 2mM MgCl2, 0.4mM EGTA, 50mM KCl, 0.2mM ATP, 1mM DTT; pH 7.5: 5mM Tris, pH 7.5, 2mM MgCl2, 0.2mM EGTA, 50mM KCl, 0.2mM ATP, 0.5mM DTT; pH 8.0: 10mM

Tris, pH 8.0, 2mM MgCl2, 50mM KCl, 1mM EGTA, 0.2mM ATP, 0.5mM DTT.

Protein labeling.

Pyrene-labeled rabbit skeletal actin (RSA) (obtained from Cytoskeleton, Inc). RSA was labeled with Cy3-maleimide in thiol-free GB5 (5mM Tris, pH 8, 0.2mM CaCl2, 0.2mM ATP) using standard approach that included 1) actin polymerization with 2mM MgCl2 and 50mM

KCl for 30-60 min at RT; 2) labeling with Cy3 dye (1:1.5 (actin:dye) molar ratio for 2 hours on ice or overnight followed by addition of 1mM DTT; 3) pelleting (TLA110 rotor at 85,000 rpm for 20 min at 4°C); 4) depolymerization on dialysis followed by gel-filtration (Superdex

S200 10/300 GL). Extent of labeling was calculated using extinction coefficient ε550=130,000

M-1cm-1. Actin labeling with Alexa488-succinimidyl ester (SE) (Molecular Probes) was done essentially as described (Mahaffy and Pollard, 2006) but Alexa488SE dye was added to F- actin in 3-7 fold excess (overnight, 4°C) and then carried out as described above for Cy3 actin. Actin concentration was measured by Bradford assay or by quantitative gels

(Coomassie staining) employing known concentrations of unlabeled RSA as standard.

Alexa488SE-actin was used to obtain 100% oxidized labeled ox-actin (GB2, 70:1

(actin:Mical) molar ratio, 100μM NADPH, 1 hour at RT). The resulting actin was dialyzed overnight against GB2 then centrifuged (TLA100 rotor, 90,000 rpm, 30 min, 4°C). Oxidation was confirmed by limited proteolysis with subtilisin.

Yeast cofilin-KCK construct (for C-terminal labeling) was modified with Cy5- maleimide in buffer C: 5mM Tris, pH 7, 0.2mM CaCl2, 50μM TCEP (1:1.5 (cofilin:Cy5- maleimide) molar ratio, 15 min at RT). Excess dye was removed using Zeba Desalt Spin

Column (Pierce) equilibrated buffer C supplemented with 1mM DTT. Extinction coefficient of Cy5 was corrected for the solvent conditions (DMF vs [buffer C+1mM DTT]) as

-1 -1 described (Grintsevich et al., 2010) and was estimated ε643=121,420 M cm . Total concentration of labeled cofilin was measured by Bradford assay, using unlabeled WT yeast cofilin as a standard.

NADPH consumption.

Different RSA concentrations (unlabeled; Cytoskeleton, Inc.) were polymerized as described

(Hung et al., 2011). Each polymer sample (or actin buffer only) was then preincubated with cofilin (human cofilin-1; Cytoskeleton, Inc.) or cofilin buffer (10mM Tris pH 8.0, 10mM

NaCl, 5% sucrose, 1% dextran, 1mM DTT) at pH of 6.8. NADPH consumption was measured essentially as described (Hung et al., 2011) with the decrease in the reduced form of NADPH determined from the decreased light absorption at 340 nm or alternatively

(Figure 2.4b), by the decreased fluorescence signal at 460 nm (when excited at 340 nm).

65 Actin disassembly assays.

Standard pyrene-actin and co-sedimentation assays using RSA (pyrene-labeled or unlabeled;

Cytoskeleton, Inc.) were performed as described (Hung et al., 2010, 2011, 2013) with minor modifications to adjust sample pH. Actin in GB5 buffer was mixed with 10X polymerization buffer (pH 6.8, 200mM Imidazole, 500mM KCl, 20mM MgCl2, 2mM EGTA) to yield 10µM actin. This mixture was then incubated on ice overnight for actin polymerization and diluted the next day to 2.5µM actin. Then, each polymer sample was incubated with cofilin (or cofilin buffer), Mical (or Mical buffer), and/or NADPH, at pH of 6.8. In some cases, as described in the figures, the polymers were preincubated with cofilin (or cofilin buffer) or

Mical (or Mical buffer). For pyrene-labeled actin, the fluorescence intensity was monitored immediately and over time at 407 nm (excitation at 365 nm) by a fluorescence spectrophotometer (Spectra max M2, Molecular Devices) as described (Hung et al., 2011).

For co-sedimentation assays, the intensity of each of the stained bands in the pellet and soluble fraction was quantified by densitometry using Image J (NIH) (Hung et al., 2011) or

Scion Image software.

Subtilisin limited proteolysis assay.

Actin was polymerized at pH 6.8 for 1 hour at RT. Next, F-actin (3.5μM) was mixed with hCofilin-1 (3.5μM) or buffer (control) to form complexes. Samples were supplemented with

NADPH (0.1mM). After removing unoxidized controls, reactions were started by addition of

25-50nM Mical. Aliquots of the samples were removed at selected time points and oxidation was stopped by addition of 1.5mM NADP+ and 3.5μM of Kabiramide C (KabC, marine

66 macrolide toxin, a kind gift from Dr. Gerard Marriott) (Klenchin et al., 2003). NADPH and

KabC were also added to the control unoxidized actin/complexes. Samples were incubated overnight at 4°C for complete actin depolymerization. Then, reaction mixtures were digested with subtilisin (limited proteolysis conditions) at 1:200 subtilisin:actin w/w ratio for 20-30 min at RT. Subtilisin stock was prepared in 2mM Tris, 0.2mM CaCl2 and used within 9 min.

Limited digestion was started by adding 1μl of subtilisin solution to the samples (25μl) arranged in random order and stopped with PMSF (1mM). Samples were analyzed by SDS-

PAGE (Coomassie stain). Densitometry analysis was performed using Scion Image software.

Increased amounts of uncleaved actin reported on the accumulation of Mical-oxidized actin.

After making corrections for undercleaved actin in unoxidized controls (~5-14%), the amount of Mical-oxidized actin was plotted vs oxidation time. We elected to use subtilisin:actin ratio that yields slightly undercleaved preparation in order to restrict proteolysis to a single site (47/48) on actin. We have found that a higher ratio of subtilisin to actin is needed for limited digestion of actin samples depolymerized under F-buffer conditions. For limited digestion of G-actin/G-actin-KabC in GB (pH 8) by subtilisin, we routinely used 1:1000 (w/w) ratio of subtilisin:actin.

Using this assay we quantified the amount of Mical-oxidized actin incorporated into copolymers under conditions closely mimicking those of our TIRF experiments (pH 6.8,

1μM of total actin, 30 min polymerization at RT). After polymerization, F-actin was pelleted

(TLA110 rotor, 150,000g, 30 min, 4°C). Pellets were resuspended in 100μl of GB2 and depolymerized overnight by dialysis (against GB2), followed by a second high speed spin

(TLA100 rotor, 150,000g, 30 min, 4°C). Alternatively, samples were depolymerized by

67 adding KabC (30μM) followed by a 3 hour dialysis against GB2. Limited proteolysis with subtilisin was carried out at 1:1000 subtilisin:actin w/w ratio, for 15-20 min, at RT and stopped with PMSF (1mM). The resulting samples were analyzed by SDS-PAGE

(Coomassie staining). The fraction of ox-actin in the sample was determined (Figure 2.5f) as described above.

Actin Met-44 and MetO-44 Specific Antibodies.

We generated an antibody that preferentially recognized the unoxidized Met-44 residue of actin (Figure 2.4d1-d3). We also generated an antibody that specifically recognized the oxidized Met-44 residue on actin (MetO-44) (Figure 2.2c). These antibodies were used to observe the effect of cofilin on Mical-mediated oxidation of actin, by incubating 1.15µM F- actin for 1 hour at RT with 1.15µM cofilin or cofilin buffer only. Then, 50nM Mical and

100µM NADPH were added and the reaction was stopped at 1, 3, 5, and 10 minutes (or 1 hour) by adding loading buffer containing β-mercaptoethanol and boiling samples for 5 minutes. For Western blotting, all samples were loaded into a 12% SDS-PAGE, transferred to PVDF membrane, blocked with 5% non-fat milk/TBST buffer for 1 hour and then incubated for 1 hour with antiserum (pan actin antibody [C4; Millipore, 1:1000]; Actin Met-

44 and Actin MetO-44 antibodies [1:500]).

TIRF microscopy assays.

Copolymers of Mical-oxidized and unmodified actin were formed in flow chambers assembled with 25x75x1 glass slides (Fisherfinest, Premium Slides, Superfrost, 12-544-7)

68 and 2230-1.5 glass coverslips (Fisherbrand, 12-544-A). Coverslips were treated with polylysine PEG solution (1.25mg/ml in H2O) for 4 min, rinsed 3 times with water and air- dried. Single flow chambers (V~30μl) were assembled using two layers of permanent double-sided Scotch tape. Before each experiment the flow chamber was treated with 2 chamber volumes (CV) of 1% Pluronic F127 solution (Sigma, P2443) (Gurel et al., 2014) for

3 min then equilibrated with 5 CV of 1xTIRF imaging buffer (20mM imidazole, 2mM

MgCl2, 50mM KCl, 0.2mM EGTA (pH 6.8) supplemented with 50mM DTT, 0.2mM ATP,

0.05mg/ml caseine, 20mM glucose, 0.25mg/ml glucose oxidase, 50μM catalase, 0.5% methyl cellulose). G-actin mixtures (10% Cy3b-maleimide labeled) were incubated for 3 min at RT with Mg/EGTA exchange buffer (0.1mM EGTA, 50μM MgCl2) and the resulting mixture (3

CV) was introduced into the flow chamber. After 15 min of on-slide polymerization, the excess of actin monomers was removed with 1 CV of 1xTIRF imaging buffer. Since cofilin binds weaker to ADP-Pi-F-actin (compared to ADP-F-actin) (Muhlrad et al., 2006; Suarez et al., 2011), filaments were aged on the surface to allow for phosphate release. For severing experiments with yeast actin, at least 3 fields were imaged between minutes 29 and 30 to determine the average filaments length before cofilin severing. At 30 min time point from the start of actin polymerization, cofilin/buffer (2 CV) was introduced into the flow chamber and movies of severing were recorded. Copolymers containing 11% ox-actin weren’t severed upon buffer additions within the monitoring time. Images were acquired every 5 sec.

Filaments were imaged using DMI6000 TIRF microscope (Leica).

Average filament length of on-slide grown ox-actin copolymers (0 and 11% oxidized) was compared to those pre-polymerized in tubes and applied to the polyK surface (related to

69 Figure 2.5h). On-slide polymerization was carried out as described above. Images were collected after 16-17 min from the beginning of polymerization (immediately after the buffer wash, at 15 min of on-slide polymerization) (Figure 2.5g). Experiments with pre- polymerized ox-actin copolymers (0% and 11% oxidized) were performed as follows.

Coverslips were treated with 1mg/ml polyK for 3 min, rinsed with mQ water and air-dried.

Mg-ATP-G-actin (15% Cy3b labeled) was polymerized at 10μM by 1xKMEI6.8 buffer

(20mM imidazole, 2mM MgCl2, 50mM KCl, 0.2mM EGTA (pH 6.8)). F-actin samples were diluted to 8μM and mixed by pipetting up and down 2 times. The resulting mixtures were incubated 5 min at RT, followed by one step dilution into 1xKMEI6.8 buffer supplemented with 100mM DTT and 1μM phalloidin and mixing by inversion. Filaments’ length was measured manually using JFilament plugin to Fiji (JFilament 2D).

Severing of fully oxidized actin by human cofilin-1 was examined as follows (related to Figure 2.5a). Flow chambers were assembled as described above. Untethered filaments were imaged on Pluronic F127-coated surface (Gurel et al., 2014) as described. F-actin-Cy3- maleimide (15% labeled) was polymerized in 1xKMEI6.8 buffer overnight at 4°C and used as F-actin seeds. Mical-oxidized (15% Alexa488SE) or unoxidized actin was mixed with

10nM of Cy3-F-actin seeds in 1xTIRF imaging buffer and polymerized for 20 min on slides.

After 20 min, unpolymerized monomers were washed with 1xTIRF buffer. To test for severing, 10nM of human cofilin-1 in 1xTIRF buffer was introduced into the flow chambers.

Filaments fragmentation induced by Mical in the presence of NADPH was monitored under the same conditions (Figure 2.7a). Unoxidized filaments (15% Alexa488SE) were grown in

70 the flow chambers then Mical (55nM)/NADPH (100μM) were introduced into the flow chamber simultaneously washing out the remaining actin monomers.

Cofilin clustering on intact filaments and copolymers with ox-actin (11%) was imaged using two-color TIRF microscopy. Actin (Alexa488-SE, 23% labeled) and yeast cofilin (Cy5-maleimide labeled) were copolymerized in flow chambers prepared as described above and imaged, unattached, on Pluronic F127-coated surface (Gurel et al., 2014). Co- polymerization was started by simultaneous addition of cofilin-Cy5 and polymerizing salts to

Mg-ATP-G-actin.

Analysis of cofilin severing.

Fiji (Image J) software was employed for movie processing. Background subtraction was done using rolling ball radius algorithm (routinely 10 pxls). Total filaments’ length (L0) was estimated using the first frame recorded after cofilin addition (L0 (μm)=sum of the lengths of all filaments in frame #1). Filaments’ length was measured manually using JFilament plugin to Fiji (JFilament 2D). Bundled filaments were excluded from the analysis. To quantify cofilin severing of F-actin, the number of severing events (cuts) was counted manually for each frame, starting from frame #2. Cumulative number of cuts divided by L0 (cuts/μm) was plotted versus time. Linear segments of the obtained dependencies were used to determine the rates of F-actin severing by cofilin (Figure 2.7e). Between 31 and 45 filaments were analyzed in each movie.

71 F-actin cross-linking.

Disulfide cross-linking of Q41C yeast actin mutant was carried out as follows. DTT-free Ca-

ATP-Q41C actin was polymerized by adding 0.1 volume of polymerizing buffer (20mM imidazole, 50mM KCl, 2mM MgCl2) for 1 hour at RT. Disulfide cross-linking in QC-F-actin was triggered by addition of CuSO4 solution (in water) to F-actin in 1:1.5 (actin:Cu) molar ratio and carried out for 1 hour at RT. Cross-linked samples were supplemented with 1mM

EGTA and dialyzed against 20mM imidazole, 50mM KCl, 2mM MgCl2, 1mM EGTA,

0.2mM ATP for 3 hours. Efficiency of cross-linking was confirmed by SDS-PAGE analysis under non-reducing conditions in the presence of NEM. Cross-linked and uncross-linked QC-

F-actin and its cofilin complexes were subjected to Mical-mediated oxidation (140:1, molar ratio Actin:Mical, thiol-free Mical preparation) in the presence of 0.1mM NADPH under non-reducing (DTT-free) conditions for 1 hour at RT. Co-sedimentation with cofilin was performed as described above (TLA100 rotor, 150,000g, 30 min, 4°C).

N-(4-azido-2-nitrophenyl) putrescine (ANP) cross-linking was carried out as described (Kim et al., 2002). In brief, thiol-free skeletal G-actin was incubated with ANP

(1:8, actin:ANP molar ratio) and transglutaminase (2 units per 1mg of actin) in DTT-free

GB2 (pH 8) for 2 hours at RT. Actin was centrifuged to remove any aggregates (21,000g, 10 min, 4°C) and then polymerized (20mM imidazole, pH 6.8, 50mM KCl, 2mM MgCl2,

0.2mM ATP, 100μM NADPH). Mical-mediated oxidation was started with the addition of thiol-free Mical to ANP-F-actin (1:140, Mical:actin molar ratio) for 1 hour at RT. Prior to photoactivation and cross-linking, the extent of Mical-mediated oxidation was assessed by subtilisin digestion. Cross-linking (between Gln 41-Cys 374) in Mical-oxidized and

72 unoxidized ANP-F-actin was triggered by UV exposure (20 min at RT) and stopped with

1mM DTT. Mical-oxidized and unoxidized ANP-F-actin yielded the same cross-linking patterns. Co-sedimentation with human cofilin-1 and gel analysis was performed as described above.

In vivo data.

Expression analysis, F-actin organization, and bristle cell remodeling was examined and quantified as described (Hung et al., 2010, 2013). Embryos were collected, processed, staged, dissected, and analyzed for axon guidance defects using an antibody to Fasciclin II (1:4, 1D4 supernatant (Van Vactor et al., 1993), Developmental Studies Hybridoma Bank) as described

(Huang et al., 2007; Terman et al., 2002; Yang and Terman, 2012; Yu et al., 1998). Males and females of Drosophila embryos, pupae, and adults were used.

Statistics and reproducibility.

For each representative image, gel, immunoblot, graph, movie, or in vivo experiment, the experiments were repeated at least two separate independent times and there were no limitations in repeatability. At least two independent protein purifications and multiple independent actin biochemical experiments were performed with similar results including reproducing the effects in Figure 2.1b and Figure 2.1g-h independently in both of our labs using non-overlapping/independent sets of reagents. No statistical method was used to predetermine the sample size, which was based on what is published in the field. Differences between experimental and control animal conditions were large, with little variability – and

73 so the sample size was larger than needed to ensure adequate power to detect an effect.

Animal studies were based on pre-established criteria to compare against age-matched animals. Animal experiments were not randomized. Animals of the correct genotype were determined and those collected of that genotype were included as data. For genetic experiments, in which the genotype needed to be determined based on different Drosophila genetic/chromosome markers, blinding was not employed. The figure legends list the sample size for each experiment. To the best of our knowledge the statistical tests are justified as appropriate. No cell lines were used in this study.

CHAPTER THREE

The Class XV Myosin Sisyphus Spatially Targets Mical to Direct Semaphorin/Plexin- mediated Actin Disassembly and Cellular Remodeling

Abstract

The mechanisms that specify the morphological features of neurons and their axonal and dendritic processes are still poorly understood – but their characterization is of crucial importance to deciphering the physiological and pathological underpinnings of neuronal connectivity. Semaphorins, for example, together with their Plexin receptors, serve as one of the largest families of extracellular effectors of neuronal wiring, but how they affect changes in cellular form and function is still enigmatic. Recently, our lab identified a conserved oxidoreductase (Redox) enzyme Mical that controls Semaphorin/Plexin effects on neuronal morphology by directly oxidizing and dismantling actin filaments. These results identify a new oxidation based signaling system controlling neural connectivity, but we know little of how this activity is regulated. To provide answers to this question, I have initiated a genetic screen and the results have identified the class XV myosin Sisyphus as a strong modulator of

Semaphorin/Plexin/Mical signaling. Sisyphus is required for Semaphorin/Plexin/Mical- mediated F-actin reorganization/cellular remodeling and increasing the levels of Sisyphus dramatically enhances these Semaphorin/Plexin/Mical-mediated effects. Further analysis reveals that Sisyphus functions to direct the specific subcellular localization of Mical – and this occurs through a mechanism dependent on Sisyphus’ motor activity. Additionally, I find that one of the two MyTH4 (Myosin Tail Homology 4) domains of Sisyphus is required for

74 75 Sisyphus to modify both Mical’s localization and its cellular effects. Collectively, these genetic and cellular studies indicate that Sisyphus orchestrates Semaphorin/Plexin/Mical- induced morphological changes by controlling Mical’s subcellular localization, thereby influencing where and to what extent actin-based cellular remodeling occurs.

Introduction

Diverse biological phenomena depend on the ability of cells to change their shape, position, and connectivity – such that mutations or other perturbations impacting this morphological restructuring generate numerous human pathologies and functional deficits. It has long been appreciated that these changes to cellular form and function are specifically and tightly regulated by the actions of extracellular cues impinging on cell surface receptors

(Berzat and Hall, 2010; Kay et al., 2008; Stossel, 1989, 1993; Swaney et al., 2010; Tessier-

Lavigne and Goodman, 1996). In turn, these cell surface receptors signal to control the organization of the actin and microtubule cytoskeletons (Berzat and Hall, 2010; Hung and

Terman, 2011). A vast array of players that regulate cytoskeletal changes downstream of extracellular cues and their receptors have now been identified, but how these proteins come together to specifically drive cellular remodeling is poorly understood (Lappalainen, 2016;

Pollard and Cooper, 2009). Moreover, even less is known about how these morphological changes are spatially controlled in a directed manner to orchestrate specific types of remodeling such as cellular process extension and guidance.

To aid in the understanding of how extracellular signals affect the cytoskeletal elements underlying cellular behaviors, our lab has been employing simple high-resolution

76 single cell in vivo genetic and cellular models and one of the largest families of extracellular cues, the Semaphorins (Semas). Semaphorins are best known to negatively regulate movement and growth (Alto and Terman, 2017) – and emerging data has shown links to disorders of the nervous system (Pasterkamp and Giger, 2009), as well as non-neuronal pathologies including cancer, cardiovascular abnormalities, and immunocompromising diseases (Nasarre et al., 2014; Neufeld et al., 2012). Yet, how Semaphorins affect cellular form and function remains far from clear. Recently, our lab uncovered that Semaphorin signaling induces F-actin disassembly and cellular remodeling through a redox-driven mechanism mediated by Mical (Molecule Interacting with CasL), a conserved flavoprotein monooxygenase enzyme which binds to both the Semaphorin receptor Plexin and actin filaments (Hung et al., 2010, 2011; Terman et al., 2002). Through an enzymatic oxidoreductase (Redox) activity, Mical post-translationally oxidizes specific methionine residues at the pointed end of actin (Hung et al., 2011). This oxidation occurs along the interface of actin filament subunits, disrupting the interactions between individual actin filament subunits and resulting in F-actin disassembly-induced changes in cellular morphology (Grintsevich et al., 2016; Hung et al., 2011, 2013). The MICALs are now known to be essential for numerous cellular events that depend on F-actin reorganization in neuronal and non-neuronal cells – and are increasingly becoming linked to various diseases and human abnormalities (reviewed in (Giridharan and Caplan, 2014; Vanoni et al., 2013; Wilson et al., 2016; Zhou et al., 2011b)). Yet, little is known of how Mical’s activity is spatially regulated, including how it precisely and directionally orients the cytoskeletal and

77 morphological changes underlying such phenomena as cell migration, axon extension, and synaptic growth.

Using a genetic screening approach, I now identify that Mical interacts with the class

XV myosin Sisyphus (Myo10A), and requires this unconventional myosin to direct

Semaphorin/Plexin-mediated cytoskeletal disassembly and cellular remodeling. Genetic interaction and localization studies go on to indicate that Sisyphus controls Mical’s subcellular localization, thereby influencing where and to what extent actin-based cellular remodeling occurs. Moreover, these effects rely on Sisyphus’ use of its motor domain to position Mical, and by so doing precisely orient the remodeling of cellular processes. This work, therefore, provides important insights into how Semaphorin/Plexin/Mical effects are spatially regulated within actin-rich cellular protrusions.

Results

The unconventional Class XV myosin Sisyphus promotes Mical-driven actin reorganization and cellular remodeling

Mical regulates F-actin organization and morphology in numerous cells, including the mechanosensory bristles of Drosophila melanogaster (Hung et al., 2010; Wilson et al., 2016)

–which have long provided a high-resolution single cell model for studying the cellular, molecular, and biochemical mechanisms of actin-driven remodeling in vivo (Hung and

Terman, 2011; Sutherland and Witke, 1999; Tilney and DeRosier, 2005). In contrast to the slightly curved, unbranched bristles of wild-type flies (Figure 3.1a), elevating the levels of

Mical in bristles using the bristle specific B11-GAL4 line (subsequently referred to as Bristle

78 Mical) results in F-actin disassembly and remodeling of the bristle cell to produce an easily observable branch (Figure 3.1b). This Mical-induced branching effect is similar in location and length from cell-to-cell/animal-to-animal (Figure 3.1b) and is dependent on

Semaphorin/Plexin signaling (Hung et al., 2010). This easily observable, reproducible, and stereotypical branching pattern prompted others in the lab and me to initiate an enhancer- suppressor genetic screen to identify proteins that modify this branching phenotype and are involved in Semaphorin/Plexin/Mical-mediated F-actin disassembly and cellular remodeling

(Figure 3.1c; (Hung et al., 2013)). The screen itself is a simple one in which publicly available mutant and transgenic Drosophila fly lines are crossed to flies expressing Mical specifically in bristles (Bristle Mical) (Figure 3.1c). Progeny are then examined for modification (i.e., enhancement or suppression) of this Mical-dependent branching effect

(Figure 3.1c). During the screening process, I observed a striking enhancement of Mical- dependent bristle branching by the P element transposon P{XP}d05943 insertion fly line

(compare Figure 3.1b and 3.1d; Figure 3.1e). Such bristles had a significant increase in the number of branches (Figure 3.1d, f) – as well as noticeably longer branches (Figure 3.1d).

Thus, the P{XP}d05943 fly line was enhancing Mical’s effects on cellular remodeling.

I therefore examined the genomic position of the P{XP}d05943 transposon insertion and found that it is situated in the Drosophila Syph (myo10A) gene (Figure 3.2a), which codes for Sisyphus (Syph), an unconventional member of the myosin family of actin-based motor proteins. Moreover, the P{XP}d05943 transposon is positioned upstream of the Syph coding region and contains upstream activator sites (UAS) for the GAL4 transcription factor driver (Figure 3.2a) – indicating that P{XP}d05943, in combination with the B11-GAL4

79 driver, is likely to be increasing the expression of Syph to enhance Mical-dependent cellular remodeling. To directly test this hypothesis, I generated transgenic flies expressing Syph and indeed found that Syph significantly enhanced Mical-mediated cellular remodeling, resembling the effects I observed with the P{XP}d05943 transposon insertion (compare

Figure 3.1g to 3.1b and 3.1d; Figure 3.3a). Further analysis revealed that Syph also greatly enhanced the F-actin disassembly-induced alterations that underlie Mical-mediated cellular remodeling, dramatically reorganizing the filamentous actin structure of the bristle (Figure

3.1h-k). In contrast, raising the levels of other myosins such as class II (Mhc), class III

(ninaC), and class XVIII (mhcl) did not notably modify Mical-mediated actin disassembly/cellular remodeling (Figure 3.3b). Moreover, of all the different myosin mutant/transgenic lines I tested (from each of the different classes of myosins), Syph showed the strongest enhancement of Mical-dependent bristle branching. These results therefore indicate that the unconventional myosin Syph specifically increases Mical activity-dependent

F-actin reorganization and cellular remodeling.

Semaphorin/Plexin/Mical-mediated actin reorganization/cellular remodeling requires

Sisyphus

Myosin family members are actin-based molecular motors that are grouped into over

30 classes based on the similarity of their motor domain (Berg et al., 2001; Cheney et al.,

1993; Foth et al., 2006; Odronitz and Kollmar, 2007; Richards and Cavalier-Smith, 2005;

Sebé-Pedrós et al., 2014) – which binds actin and hydrolyzes ATP to move actin filaments, as well as myosins along these filaments (Hartman and Spudich, 2012). This actin-binding

80 motor domain is present in the N-terminal “head” region of all myosins, but myosins also have a variety of other domains located in their C-terminal “tail” region (Figure 3.2b). This

“tail” region serves to specify additional binding partners and spatiotemporal localization patterns for the myosins (Hartman and Spudich, 2012; Hartman et al., 2011; Woolner and

Bement, 2009). The 14 known Drosophila myosins fall into 10 different classes (I, II, III, V,

VI, VII, XV, XVIII, XX, XXII) (Figure 3.2b; (Berg et al., 2001; Cao et al., 2014; Foth et al.,

2006; Sebé-Pedrós et al., 2014; Tzolovsky et al., 2002)), with Syph being the only known class XV myosin in Drosophila. However, Syph has only been poorly characterized, since loss-of-function/”knock-out” genetic mutant lines have not been generated.

Therefore, to further examine the role of Syph in Mical-mediated cytoskeletal rearrangements, I used a FLP/FRT (site-specific) recombination technique (Parks et al.,

2004) to delete the entire coding region of Syph and generate a Syph mutant, Syph21J (Figure

3.2a). Characterizing these Syph21J mutants revealed that they reach third instar larval stages, but typically die before or shortly after pupation and rarely survive to adulthood (Figure

3.3c). I did note, however, a few adult escapers that had variable visible defects including effects on wings and bristles – including alterations to bristle length and morphology (Figure

3.3c). I thus sought to analyze the effects of decreasing the levels of Syph on Mical- dependent cellular remodeling. Notably, loss of one copy of Syph (Syph21J/+) strongly suppressed Mical activity-dependent cellular remodeling/actin alterations – generating branches that were typically very thin, short, or absent – and thereby decreasing both the number and length of bristle branches (Figure 3.2c-d). I also observed similar suppressive effects on Mical activity-dependent F-actin/cell morphological changes following bristle-

81 specific expression of a Syph RNAi (Figure 3.3d). Moreover, since Mical functions with

Semaphorin/Plexin repulsive guidance cues/receptors in bristle F-actin/cellular remodeling

(Hung et al., 2010, 2013), I wondered if Syph also played a role in Sema/Plexin repulsive signaling. My results revealed that increasing the levels of Syph (P{XP}d05943 line) in bristles, increased Semaphorin/Plexin-mediated actin/cellular remodeling, while reducing the levels of Syph (Bristle Syph RNAi) decreased Semaphorin/Plexin-mediated actin/cellular remodeling (Figure 3.2e). Furthermore, the Syph-induced increase in Semaphorin/Plexin- mediated actin/cellular remodeling was dependent on Mical and on the Plexin cytoplasmic

(intracellular) domain that interacts with Mical during Semaphorin signaling (Figure 3.3e).

Thus, Syph is required for Semaphorin/Plexin/Mical-dependent F-actin/cellular remodeling.

Sisyphus positions Mical to spatially direct F-actin disassembly/cellular remodeling

As a class XV myosin, Syph, along with class VII and class XXII myosins, are the

Drosophila members of the MyTH-FERM superclass of myosins, which are defined by the presence of MyTH4 (Myosin Tail Homology 4) and FERM (protein Four-point-one, Ezrin,

Radixin, Moesin) domains in their tail regions (Figure 3.2b). Members of the MyTH-FERM superclass have been defined based on their ability to regulate intracellular transport within actin-rich filopodia/cellular protrusions, with each MyTH-FERM protein differing in their subcellular localization and the particular cargo they carry (Weck et al., 2016). Specifically,

Syph and the mammalian class XV myosin, Myosin XV, have been shown to traffic specific protein cargo within and to the tips of actin-rich structures such as filopodia and stereocilia

(Belyantseva et al., 2005; Liu et al., 2008; Manor et al., 2011; Mauriac et al., 2017). I

82 therefore wondered if Syph might be regulating Semaphorin/Plexin/Mical-mediated actin disassembly/cellular remodeling by controlling Mical’s subcellular localization.

Mical-driven actin disassembly/bristle cell remodeling is not detectible along the entirety of the bristle process, but rather at distinct loci, which generate easily observable filopodia/branches (Figure 3.4a, arrows and arrowheads; (Hung et al., 2010)). Therefore, to better understand how this filopodial/branch formation is occurring, I imaged developing bristles over time to temporally follow both the remodeling bristle and the spatial localization of Mical within it. During the early stages of bristle extension, I noted that Mical generated an easily observable small branch at the tip of the growing bristle (Figure 3.4a [~34 hrs, arrows]). Then, as development continued, the main bristle shaft elongated past this 1st branch point (Figure 3.4a [~36 hrs- ~42 hrs]), advancing until Mical-mediated actin disassembly occurred again at the bristle tip, generating a 2nd larger branch that was spatiotemporally distinct from the first (Figure 3.4a [~44 hrs, arrowhead]). Further analysis revealed that Mical localized to bristle tips during the course of this bristle extension (Figure

3.4a, red) – making forays into the 1st branch (Figure 3.4a, arrows, red), but maintaining the bulk of its localization to the main bristle shaft throughout its elongation and after formation of the 2nd branch (Figure 3.4a, red).

I next sought to determine if this stereotypical pattern of Mical localization and bristle remodeling was significantly altered by Syph. Co-expressing Syph with Mical did not result in any observable changes during the early stages of bristle development – in that Mical localized in a normal pattern within extending bristle tips (Figure 3.4b [~32 hrs, red]).

However, beginning at a similar time and position where the first Mical-dependent branch

83 formed (~34hrs), I noticed that Mical’s distribution began to change dramatically in the presence of Syph – spreading out from around the position of the first branch point – and generating a curved and altered orientation to the bristle tip, as well as additional branches with increased length (Figure 3.4b [~34 hrs – ~39 hrs] and compare to Figure 3.4a [~34 hrs – ~39 hrs]; Figure 3.5a). Furthermore, with increasing development, these Mical/Syph- dependent tip and branch alterations became even more pronounced (Figure 3.4b [~41 hrs], arrowhead) – forming a large, complex, and misoriented process containing Mical (Figure

3.4b [~41 - ~44 hrs], red) – while the typical vertical extension of the bristle was delayed in forming and was devoid of appreciable Mical localization (Figure 3.4b [~44 hrs, arrow and arrowhead] and compare to Figure 3.4a [~44hrs, arrowhead]). I also found that Syph was localized within these regions with Mical (Figure 3.5b) and strikingly, that this Syph- triggered broadened distribution of Mical dramatically overlapped with the increasing areas of F-actin disassembly that characterized Syph’s enhancement of Mical-mediated cellular remodeling (Figure 3.4c, Figure 3.5c-d; see also Figure 3.1h-k; and see Figure 3.5e for

Syph only effects). Thus, Syph modifies the distribution of Mical, altering the positioning and extent of F-actin disassembly to spatiotemporally direct cellular growth and remodeling.

Sisyphus employs its myosin motor activity to target Mical and locally disassemble F-actin

My results reveal that Syph modifies Mical-mediated F-actin disassembly by altering

Mical’s localization, so I wondered if this action of Syph was due to effects on long-range transport of Mical from the cell body to the growing bristle tip. To address this question, I examined the localization of Mical in Syph “knock-out/known-down” mutants. Notably,

84 although reducing the levels of Syph (Syph21J/+; Bristle Syph RNAi) resulted in strong suppression of Mical activity-dependent F-actin disassembly/branching (Figure 3.6a; see also Figure 3.2d), it did not alter Mical’s localization to bristle tips (Figure 3.6b-c). Thus, my results indicate that Syph does not alter Mical activity-dependent cellular remodeling by regulating its long-range transport to the bristle tip.

I hypothesized therefore that Syph might be locally regulating the distribution of

Mical to direct its effects on actin disassembly. To test for this possibility, I disrupted the myosin motor activity of Syph by 1) deleting its entire motor domain (SyphΔMotor; Figure

3.6d) and 2) by making two point mutations (R213A and G434A) in the highly conserved switch I and switch II regions of the motor domain (SyphR213A,G434A; Figures 3.6d and 3.7a).

These point mutations are known to disrupt myosin motor activity (e.g., in mouse Myosin

XV and Dictyostelium discoideum Myosin II) by preventing ATP-hydrolysis, thereby keeping the myosin motor in a weak actin binding state and unable to move along actin filaments (Belyantseva et al., 2005; Sasaki et al., 1998; Shimada et al., 1997). Generating

Syph transgenic flies with this R213A and G434A motor mutation (SyphR213A,G434A) and Syph transgenic flies without the motor domain (SyphΔMotor) revealed that even without a functioning motor, Syph localized throughout the extending bristle process, as did wild-type

Syph (Figure 3.7b). However, these Syph motor mutants did not induce the same effects on

Mical’s localization and cellular remodeling as wild-type Syph. Specifically, while some branches formed in response to co-expression of Mical and the motor mutant forms of Syph

(Figures 3.6e and 3.7c-d), the redistribution of Mical that was a hallmark of wild-type Syph

(Figure 3.4b) was abolished by the motor mutant form of Syph (Figures 3.6e-f and 3.7e).

85 Likewise, the extensive Mical-mediated F-actin dependent cell morphological changes induced by Syph were suppressed by disrupting the motor domain (Figure 3.6e) – further indicating that the consequence of the Syph-induced relocalization of Mical is a greater area of F-actin/cellular remodeling. Thus, Syph uses its motor domain to locally direct Mical’s distribution to spatially target F-actin organization and cellular remodeling.

Sisyphus locally targets Mical-mediated actin disassembly through the use of its first MyTH4 domain

The Syph-containing class of myosins (Class XV myosins) have been shown to locally transport different cargo – which they do through the use of their motor domain and various tail domains that directly associate with specific cargo (Figure 3.8a; (Belyantseva et al., 2005; Liu et al., 2008; Manor et al., 2011)). Since Syph regulates the subcellular localization of Mical in a motor activity-dependent manner, I wondered if Syph might also be using one of its tail domains to locally transport Mical. To begin to address this question, I deleted the MyTH4, FERM, and PH-like tail region domains of Syph (SyphΔMPMF; Figure

3.8a) and made transgenic flies with this tail region deletion to examine its effect on Mical’s localization and cellular remodeling (Figure 3.8b-c). Dramatically, although SyphΔMPMF localized prominently to bristle processes in a manner similar to wild-type Syph (Figure 3.8b compared to Figure 3.7b), it did not relocalize Mical, or trigger the enhanced degree of

Mical-mediated cellular remodeling observed with wild-type Syph (Figure 3.8b-c). Thus, my results indicated that the MyTH4, FERM, and/or PH-like domains of Syph, similar to the

86 motor domain, were important for locally targeting Mical for F-actin disassembly/cellular remodeling.

To determine which of the tail region domain/s was required for Syph to locally target

Mical, I deleted the MyTH4, FERM, and PH-like domains individually, and generated additional transgenic fly lines to test their effects on Mical (Figure 3.8a). My results revealed that neither Syph’s PH-like nor FERM domains were required for Syph to locally target

Mical – such that Mical’s distribution in these deletion mutants (SyphΔPH-like and SyphΔFERM) was similar to that seen with wild-type Syph (Figures 3.8c and 3.9a-b). In contrast, deleting the MyTH4 domains (SyphΔMyTH4(1&2)), disrupted Syph’s ability to relocalize Mical, mimicking the effects on Mical-mediated local targeting and cellular remodeling that I saw with both the Syph motor mutant and SyphΔMPMF (Figures 3.8a,c and 3.10a). Similarly, since Syph has two MyTH4 domains, I deleted each of them individually (SyphΔMyTH4(1) and

SyphΔMyTH4(2)) – and transgenic flies for each of the two deletions revealed that it was the first

MyTH4 domain, the one most proximal to the motor domain (MyTH4(1); Figure 3.8a) that was required for Syph to redistribute Mical (Figures 3.8c-e and 3.10b). Moreover, further deletion analyses revealed that the presence of the first MyTH4 domain alone, among the different tail region domains, allowed Syph to relocalize Mical (SyphΔMF and SyphΔPMF;

Figures 3.8a,c and 3.10c-d). Thus, the first MyTH4 domain is necessary and sufficient among the tail region domains of Syph to direct Mical’s specific targeting – suggesting that

Mical is locally positioned by the specific myosin motor Syph to spatiotemporally induce F- actin disassembly/cellular remodeling (Figure 3.11a-b).

87 Syph regulates neural connectivity and muscle morphology, resembling Mical-mediated effects in these tissues

In addition to its role in the model bristle system, Mical regulates F-actin organization/cellular remodeling in numerous other tissues to direct axon guidance, synaptic connectivity, dendritic pruning, and muscle organization (reviewed in (Wilson et al., 2016)).

I therefore wondered if Syph might also function with Mical in some of these other tissues – and employed the developing Drosophila larval neuromuscular junction (NMJ; Figure

3.12a) to examine Syph and its role in the developing nervous and musculoskeletal systems.

Mical is broadly expressed in the nervous and musculoskeletal systems (Beuchle et al., 2007;

Terman et al., 2002) and Mical mutants have abnormal synapse structure at these NMJs – such that synaptic boutons are typically clustered together and synapses do not spread out normally along muscles (Figure 3.12b; (Beuchle et al., 2007; Hung et al., 2013)). Syph too, like Mical, is broadly expressed including within the nervous system and skeletal muscle

(Figure 3.13a-c; (Aradska et al., 2015; Graveley et al., 2011; Liu et al., 2008; Lye et al.,

2014; Ryder et al., 2009; Tzolovsky et al., 2002; Wasbrough et al., 2010)). Notably, Syph mutants also had abnormal synaptic structure with defects that resembled Mical mutants – such that NMJ synapse length was reduced in Syph mutants along muscles of the same size

(Figure 3.12d). Additionally, Mical mutants have disrupted actin and myofilament organization within skeletal muscles on the postsynaptic side of the NMJ (Beuchle et al.,

2007; Hung et al., 2013), and increasing Mical expression in muscles using a muscle-specific

GAL4 driver causes F-actin organizational defects (Figure 3.12c; (Hung et al., 2013)).

Likewise, I found that muscle-specific expression of Syph caused significant alterations in F-

88 actin organization, and also resembled Mical overexpression-induced muscle defects (Figure

3.12e). Together these data reveal roles for Syph in regulating neural connectivity and muscle morphology, resembling Mical’s effects and thereby suggesting that Syph may be working with Mical to achieve proper synapse development and muscle organization.

Discussion

Specific cellular behaviors such as establishing polarity, directing outgrowth, and properly navigating depend on the poorly-understood coordinated reorganization of the actin cytoskeleton in a spatial and temporal manner. I show here that such spatiotemporal coordination relies on a specific myosin – a member of the Class XV myosins – and its ability to locally target particular actin regulatory proteins within specific areas of the cell.

Specifically, my results reveal that the actin disassembly factor Mical is locally targeted to particular areas via its interactions with the class XV myosin, Sisyphus – and this targeting positions Mical to control the degree of F-actin disassembly and cellular remodeling that occurs in response to Semaphorin/Plexin repulsive signaling. My results also go on to demonstrate that Sisyphus uses its motor domain to locally target Mical, as well as its first tail-region MyTH4 domain. Moreover, I find that this myosin XV-Mical targeted F-actin disassembly is required for directing cellular remodeling and Semaphorin/Plexin repulsive signaling – and that Syph is important in specifying synaptic remodeling and muscle morphology.

Syph and mammalian class XV myosins have been shown to localize to sites where the actin cytoskeleton is dynamic, such as filopodia and hair cell stereocilia tips. They

89 interact with proteins that regulate the cytoskeleton, function in the formation/elongation of and F-actin organization within filopodia and stereocilia, and mediate cell-cell adhesion events (Anderson et al., 2000; Belyantseva et al., 2003, 2005; Liu et al., 2008; Manor et al.,

2011; Mauriac et al., 2017; Probst et al., 1998; Zampini et al., 2011). My results reveal a new mechanism for Syph regulating the cytoskeleton – through modifying the localization of the actin regulatory protein, Mical.

Class XV myosins are thought to primarily function as cargo transporters in actin-rich cellular protrusions based on the presence of a motor with a high duty ratio (Bird et al., 2014)

(i.e. the proportion of time spent strongly bound to actin during each ATPase cycle – a property required for processive movement on actin; (Krendel and Mooseker, 2005)) and on their observed effects in cellular systems. For example, Sisyphus has been shown to directly bind and traffic proteins within epithelial cell filopodia, such as the cell adhesion protein DE- cadherin and the microtubule plus-end-tracking protein EB1 (End binding 1) (Liu et al.,

2008). In mammals, Myosin XV directly binds the scaffolding protein whirlin and the actin binding and capping protein Eps8 (Epidermal growth factor receptor pathway substrate 8), and in a motor-dependent manner localizes to the tips of hair cell stereocilia along with these cargo (Belyantseva et al., 2005; Delprat et al., 2005; Manor et al., 2011). Additionally,

Myosin XV can traffic a complex of proteins through its interaction with whirlin (Mauriac et al., 2017). In shaker-2 (sh2) mice which have a point mutation in the motor domain of

Myosin XV, disruption of motor activity prevents the localization of Myosin XV cargo to its proper location at stereocilia tips and results in short stereocilia and profound hearing loss

(Belyantseva et al., 2005; Manor et al., 2011; Mauriac et al., 2017). In a similar way, my

90 results reveal that Syph’s motor domain is required for Syph to modify Mical’s localization.

Based on these findings and the known function of class XV myosins, I predict that the

Syph-induced effects on Mical are due to Syph’s function as a cargo transporter and that

Syph transports Mical within cellular protrusions - most likely through direct transport of

Mical or through transport of Mical as part of a complex of proteins, although vesicle transport is also a mode of intracellular trafficking for myosins (Krendel and Mooseker,

2005). Alternatively, Syph could indirectly alter Mical’s localization through an interaction with a currently unknown protein that modifies Mical’s subcellular localization. On the other hand, Mical’s normal distribution within muscles and neurons is disrupted in Mical mutants with a point mutation in the FAD binding motif required for Mical’s redox activity (Beuchle et al., 2007), suggesting that Mical’s activity may affect its localization in some way.

Therefore, it is possible that Syph could modify Mical’s localization by altering (either directly or indirectly) Mical’s activity. Future work should be aimed at exploring these possibilities to further eludicate the mechanism by with Syph directs Mical’s localization, such as by testing whether Mical and Syph directly bind through co-immunoprecipitation.

Localization studies also indicate that the first MyTH4 domain of Syph is required for

Syph to change Mical’s cellular distribution. While MyTH4 domains have been shown to interact with microtubules (Hirano et al., 2011; Narasimhulu and Reddy, 1998; Planelles-

Herrero et al., 2016; Weber et al., 2004), class XV myosin cargo that mediate cell adhesion or cytoskeleton regulation have not been found to bind this region. However, this domain has clear functional importance as indicated by numerous missense mutations in the first MyTH4 domain of the MYO15A gene that have been linked to non-syndromic autosomal recessive

91 deafness (DFNB3) (Shearer et al., 2009; Wang et al., 1998). Due to its role in binding microtubules, the requirement of the first MyTH4 domain for the Syph-induced effects on

Mical’s localization may signify the importance of an interaction between Syph and microtubules for the Syph-Mical interaction, through a currently unknown mechanism.

Alternatively, it’s intriguing to consider the possibility of the MyTH4 site as a new cargo binding site, and future work should be aimed at determining whether this domain indeed mediates direct binding of Mical to Syph.

Importantly, these results suggest a model where Syph contributes to Semaphorin signaling and cellular remodeling through an interaction with Mical, but they do not rule out the possibility that other unknown Syph cargo are contributing to the cellular remodeling observed, as Syph has been to shown to bind proteins that regulate both actin and microtubules (Liu et al., 2008). One cargo, EB1, has been shown to bind Plexin receptors and function in neurite outgrowth (Laht et al., 2012); and therefore, it might also contribute to

Semaphorin/Plexin/Mical-mediated cellular remodeling. In fact, the activity of other Syph cargo in this process could explain why some cellular remodeling was observed in all Syph domain deletion and Mical expressing bristles. Thus, future studies should be directed towards exploring what role, if any, these known Syph cargo might have in

Semaphorin/Plexin/Mical signaling.

Mammalian class XV myosins have been shown to be highly expressed in hair cells of the ear and in the pituitary gland and pituitary tumors (Liang et al., 1999; Lloyd et al.,

2001). Additionally, low expression has been observed in other tissues such as brain, liver, kidney, lung, pancreas, and skeletal muscle, but the function of Myosin XV in most of these

92 tissues is unknown (Liang et al., 1999). Here, results show that Syph plays a role in regulating F-actin organization in larval skeletal muscles. Although the mechanism is unclear, the similarity in muscle F-actin organization defects caused by Mical and Syph overexpression suggests that Syph may be regulating Mical’s localization/activity in muscles as well. This study also identified Syph as a critical component in regulating the length of synaptic innervation at NMJs, and future work should be aimed at understanding if Syph is regulating synapse length by acting presynaptically, postsynaptically, or both and if this effect is dependent on the interaction between Mical and Syph, as the similarity in Syph mutant and Mical mutant phenotypes suggests.

The findings in this study indicate an important functional interaction between Syph and Mical, which raises questions about the role of this interaction in cellular behaviors known to require class XV myosins. Mutations in human MYO15A are one of the leading cause of deafness (Rehman et al., 2016), and different isoforms of Myosin XV regulate the actin cytoskeleton and function in stereocilia elongation and maintenance (Anderson et al.,

2000; Belyantseva et al., 2003, 2005; Fang et al. 2015; Manor et al., 2011; Probst et al.,

1998). While there is no known role for Mical in stereocilia elongation or maintenance, mutations in a Methionine sulfoxide reductase, MsrB3, are linked to autosomal nonsyndromic deafness (DFNB74) and cause degeneration of stereocilia (Ahmed et al., 2011;

Kwon et al., 2014). In light of these findings, Kwon et al. (2014) suggest a possible link to redox regulation of actin in stereocilia maintenance, in which a MICAL family member could be a player given the known function of MICALs and Methionine sulfoxide reductase enzymes as a redox switch (Hung et al., 2013; Lee et al., 2013). MsrB3 was found to localize

93 at the base of the stereocilia (Kwon et al., 2014), in which case Mical would be expected to function at the base as well, while Myosin XV is localized at the tips (Belyantseva et al.,

2003, 2005; Rzadzinska et al., 2004). However, if redox regulation of actin occurs in stereocilia, Mical could also reasonably function at the tip as well, in concert with a different

MsrB protein, and participate in the localized actin turnover occurring at stereocilia tips which is required for stereocilia elongation and maintenance (Drummond et al., 2015;

Narayanan et al., 2015; Zhang et al., 2012). Although still highly speculative, a functional interaction between class XV myosins and MICALs in stereocilia elongation and maintenance is an intriguing avenue for future exploration as the precise control of actin turnover or actin dynamics in stereocilia is not well understood.

In conclusion, I find that Sisyphus is a novel regulator of Semaphorin/Plexin/Mical- mediated F-actin disassembly and cellular remodeling. Semaphorin repulsive signaling leads to the activation of Mical through its binding to the Semaphorin receptor Plexin, triggering

Mical-mediated F-actin disassembly and leading to an increase in the complexity of the cytoskeleton due to F-actin reorganization and branching (Figure 3.14a; (Hung et al., 2010,

2013; Terman et al., 2002)). My results now indicate that precisely controlling the subcellular localization of Mical is also critical for mediating the extent of Semaphorin’s F-actin disassembly/repulsive effects. Furthermore, my results suggest a model where Syph functions to promote Mical’s activity (F-actin disassembly) by directing Mical into new cellular regions (Figure 3.14b). I also find that this ability of Syph to locally modify Mical’s subcellular distribution occurs through a mechanism dependent on Syph’s motor domain and also its first MyTH4 domain (Figure 3.14b). One possible implication of these results and

94 this model is that Syph may be directing the movement of an active form of Mical, such as

Mical after it has been activated by Plexin which relieves autoinhibition of Mical (Schmidt et al., 2008). Additionally, this model suggests that Syph serves as an unconventional means of propagating the Semaphorin/Plexin/Mical signal within a local region by controlling how widespread Mical-mediated F-actin disassembly and cellular remodeling are within a cell.

Therefore, these findings offer new insights into how Semaphorin/Plexin/Mical-signaling induces localized actin disassembly and cellular remodeling and suggest that subtle manipulation of Mical’s localization can dramatically affect cell shape.

Thus, my findings provide insights into the spatial regulation of Mical and reveal that changes in Mical’s subcellular localization, driven by Syph, serve as a means of cellular remodeling. Additionally, these findings offer new clues about the function of the unconventional class XV myosin family members, the tissues in which they function, and their roles in regulating the actin cytoskeleton in the Semaphorin signaling pathway.

95 Figures

Figure 3.1. The unconventional class XV myosin Sisyphus increases Mical-driven F- actin disassembly/cellular remodeling. (a) Drosophila bristle cells project a long,

96 unbranched, F-actin rich (in green) cellular extension during pupal stages (left panel) – and a record of this extension and its F-actin driven morphological features is visible in the adult (right panel). (b) Expression of Mical specifically in bristles (Bristle Mical = UAS:Mical/+, B11-GAL4/+) induces alterations to F-actin organization (arrowhead, left panel) and a branched (arrowhead, middle and right panels) cellular morphology (c) Enhancer-suppressor genetic screen to identify modifiers of Mical-dependent bristle branching. The screening strategy involved (1) crossing female Bristle Mical flies to males carrying either a gain-of- function (GOF) or loss-of-function (LOF) mutation and (2) examining the number and length of the branches found on the left posterior scutellar (PSc) bristle of adult progeny. Bristles with a similar number and length of branches as Bristle Mical received a score of 0 (i.e., similar to the control), while bristles with decreased or increased numbers/lengths of branches compared to Bristle Mical were scored as suppressed (score = -1 or -2) or enhanced (score = +1 or +2), respectively. (d-f) Mical-dependent bristle branching is strongly enhanced by the P element transposon P{XP}d05943 insertion line – such that flies have increased number of branches per bristle (e.g., d, arrowhead [quantified in e and f]). Bristle Mical + P{XP}d05943 = UAS:Mical/+, B11-GAL4/+, P{XP}d05943/+. For e, n ≥ 60 bristles per genotype (1 bristle per animal). For f, n ≥ 40 bristles per genotype (1 bristle per animal). Mean ± standard error of the mean (s.e.m.). ****P <0.0001; unpaired t-test (two-tailed). Bristle P{XP}d05943 only flies (B11-GAL4/+, P{XP}d05943/+) have branchless bristles. (g- k) Bristle-specific expression of Sisyphus significantly enhances Mical activity-dependent F- actin disassembly and cellular remodeling. Actin is well organized along the bristle shaft in wild-type animals (h), but bristle-specific expression of Mical induces small regions of F- actin disassembly (i, arrows) and branching (i, arrowhead). Bristle-specific expression of Syph enhances Mical-mediated F-actin disassembly (j, arrow) and branching (g and j, arrowhead), such that a greater percent of the bristle process has disrupted actin when Syph and Mical levels are both elevated (k). ****P <0.0001; unpaired t-test (two-tailed); n ≥ 12 bristles per genotype (1-2 bristles assessed per animal). Mean ± s.e.m. Bristle Syph = UAS:Syph/+, B11-GAL4/+.

Figure 3.2. Syph is required for Mical-dependent F-actin disassembly/cellular remodeling. (a) Genomic map of Sisyphus with black boxes representing exons and gray boxes representing 5’ and 3’ UTR. Transposable elements inserted in Syph and utilized here are represented by triangles. A Syph mutant (Syph21J) was generated by FLP-mediated recombination between the FRT sites contained within two transposable elements, one (PBac{WH}f06507) upstream of the coding region and one (PBac{WH}Myo10Af03968) downstream of the coding region. As a result of the recombination, only a short non-coding

98 region of Syph remains in Syph21J mutants. (b) The Drosophila myosin family of proteins and their domain organization. Each myosin has a motor domain (head) and at least one IQ-motif (neck). In the C-terminal (tail) region, myosin family members contain various conserved domains. Cargo binding domains for class V and class VI are class-specific. N-terminal SH3- like is a myosin N-terminal domain with a SH3-like fold. TH1 (Tail Homology 1 (also referred to as a Basic domain)), SH3 (Src Homology 3), MyTH4 (Myosin Tail Homology 4) FERM (4.1 protein, Ezrin, Radixin, and Moesin), PH-like (Pleckstrin Homology-like), PDZ (PSD95, Dlg1, zo-1). Mhc (Myosin heavy chain), zip (zipper), ninaC (neither inactivation nor afterpotential C), didum (dilute Drosophila unconventional myosin), jar (jaguar), ck (crinkled), Syph (Sisyphus), mhcl (myosin heavy chain like), d (dachs). Domain organization shown here is based primarily on the Conserved Domain Database (CD-Search tool; (Marchler-Bauer et al., 2011). Additionally, IQ-motifs were based on ref: (Tzolovsky et al., 2002) (except for class XXII which was identified only recently (Sebé-Pedrós et al., 2014)). Although not identified through CDD, class VII myosins are also predicted to have a coiled- coil within their tail region, C-terminal to the IQ-motifs, and class VIIa is also predicted to have an N-terminal SH3-like domain (Kiehart et al., 2004; Yang et al., 2005, 2006). (c-d) Different heterozygous loss-of-function and RNAi mutants of Syph suppress Mical- dependent F-actin disassembly (c, left panel) and bristle branching (c, right panel [arrowhead] and drawings and d). Syph RNAi is a UAS:Syph RNAi line (P{TRiP.HMS02255}attP2). Syph21J/+ (Syph “knockout” mutant). For (d), n ≥ 30 bristles per genotype (1 bristle per animal); mean ± s.e.m. All genotypes are heterozygous (B11-GAL4, UAS:Mical/+ and mutations/+ or RNAi lines). (e) Syph modulates Plexin A-mediated cellular remodeling. Raising the levels of Syph (Bristle PlexA + P{XP}d05943 = UAS:HA- PlexA/+, B11-GAL4/+, P{XP}d05943/+) increases the percentage of flies with bristle branching induced by Plexin A (Bristle PlexA = UAS:HA-PlexA/+, B11-GAL4/+), while reducing Syph levels (Bristle PlexA + Bristle Syph RNAi = UAS:HA-PlexA/+, B11-GAL4/+, P{TRiP.HMS02255}attP2/+) decreases this Plexin A-dependent effect. n ≥ 30 animals per genotype.

Figure 3.3. Further analysis of the effects of Syph and other myosins on Semaphorin/Plexin/Mical-dependent bristle remodeling. (a) Bristle-specific expression of Mical (Bristle Mical) in combination with Syph (Bristle Syph) strongly enhances the branching effects observed in bristles expressing only Mical – with branches that are longer, thicker, and often more complex. Note that in the second tracing from the left on the bottom

100 row, the 2 posterior scutellar bristles are tangled together, so that 2 bristles are shown, instead of 1. Note also that the bristles (main shaft) are shorter and thicker (compare to Figure 3.1b). See also the image in Figure 3.1g. (b) Analysis of different putative myosin gain-of-function (GOF; green) and loss-of-function (LOF; red) mutants. Publicly available fly lines were obtained for analysis (i.e. lines that are predicted or known to increase (GOF) or decrease (LOF) the function of different myosins). Note that GOF lines were not publicly available for class III and VIIb myosins, and LOF and GOF lines were not available for class XXII. The analysis was conducted using the screening strategy outlined in Figure 3.1c and reveals that Mical activity-dependent bristle branching is most strongly enhanced by a GOF line (P{XP}d05943) for the class XV myosin, Sisyphus (note that data for the effects of Sisyphus are also shown in Figure 3.1e with the score not normalized to Bristle Mical). These results indicate that the strong enhancement of Mical activity-dependent bristle branching is not a property common to all myosins. Interestingly, note however, that GOF or LOF lines from a few additional myosins such as class VIIa (crinkled) LOF, class II (zipper (zip)) GOF and LOF, class VI (jaguar) GOF, and class V (dilute Drosophila unconventional myosin (didum)) LOF also moderately/strongly affect Mical-dependent branching. Note also that here the scores are normalized to the score given to the progeny of Bristle Mical and w1118 (wild-type) flies. (c) Analysis of the adult viability and phenotypes of the Syph21J mutant (see Figure 3.2a). The Syph gene is on the X chromosome, thus Syph21J mutant males (Syph21J/Y) have no functional Syph allele. At 25°C, Syph21J heterozygous females (Syph21J/FM7c) crossed to males with the FM7c balancer chromosome (FM7c/Y) rarely produce mutant males that survive to adulthood (0.34%). Stocks kept at 21°C also rarely contain adult mutant males. Therefore, only a handful of mutant escapers were examined. Based on the limited number of escapers, I determined that Syph21J/Y mutant males are fertile, but are small in overall size and have variable defects that were not apparent in every mutant – such as the presence of shorter and thicker interommatidial bristles (small bristles in the fly eye), a few missing bristles or morphologically defective bristles, shorter bristles on the abdomen of the fly, as well as a noticeable defect in the wings such that they appeared wrinkled. I also occasionally observed bristle morphology defects in flies with bristle-specific expression of Syph RNAi in Syph heterozygous flies (Bristle Syph RNAi; Syph21J/+; data not shown). It should also be noted that occasionally I noticed abnormalities indicating that X chromosome non- disjunction might be occurring at an increased frequency in crosses made with Syph21J heterozygous females (Syph21J/balancer chromosome). That is, whereas Syph mutant males (Syph21J/Y) are rarely viable as adults and have light orange eyes (due to a w+ containing transposable element), some crosses produced many male progeny without the balancer, but these males possessed the eye color (e.g. white, due to w-) of the father – indicating that the X-chromosome came from the father and was not Syph21J. However, I did not test for sterility as would be expected of an XO male in Drosophila (Griffiths et al., 2000); therefore, it is possible these effects are not due to non-disjunction. I noticed that this effect occurred less frequently when flies were balanced with the FM7c balancer rather than the FM7a balancer, so for this reason, FM7c balancers were used to maintain mutant stocks (this data is not shown). Syph localizes to mitotic spindles and is required for centromere clustering (Kwon et al., 2008). I predict that it may also play a role in meiosis – such that there might be increased risk of homologous chromosomes or chromatids failing to separate in Syph mutants. (d)

101 Bristle-specific expression of Syph RNAi (Bristle Syph RNAi = P{TRiP.HMS02255}attP2/+, B11-GAL4/+) also suppresses Mical-dependent bristle branching (d, arrowhead). n ≥ 30 bristles per genotype (1 bristle per animal); mean ± s.e.m. (e) Although elevating levels of Syph (by P{XP}d05943 line) increases the percentage of flies with PlexA-induced branching (see Figure 3.2e), this effect is suppressed by the loss of 1 copy of Mical (Bristle PlexA+ P{XP}d05943 + MicalDf = UAS:HA-PlexA/+, B11-GAL4/+, P{XP}d05943/+, MicalDf(3)- swp2/+ (a Mical deficiency line (Terman et al., 2002))), and Syph does not promote strong branching effects in flies expressing PlexAΔCyto, (Bristle PlexAΔCyto + P{XP}d05943 = PlexAΔCyto/+, B11-GAL4/+, + P{XP}d05943/+) which lacks the intracellular domain of Plexin needed to interact with Mical. n ≥ 24 animals per genotype.

102

Figure 3.4. Syph distributes Mical to direct F-actin disassembly and cellular remodeling. (a) Elevating Mical levels generates spatially and temporally distinct alterations in bristle morphology. Developing bristles were imaged over time in Bristle Mical flies

103 (UAS:mcherryMical/+, B11-GAL4/+) to temporally follow both the remodeling bristle and the spatial localization of Mical (red) within it. In this stereotypical time course, each image was taken from a different bristle but is typical of that stage (~ hour) of development. Bristles extend over the course of many hours, beginning at ~31 hours after puparium formation. Bristles with elevated levels of Mical often develop a branch (arrow) during the early stages of bristle extension (~34 hrs). The main vertical bristle shaft then continues to grow beyond this branch point (~36 hrs - ~42 hrs). Towards the end of bristle process extension, a second, larger, Mical-dependent branch (arrowhead) forms. During this time course of bristle development and cellular remodeling, Mical localizes to bristle tips from the beginning of their extension (~32 hrs), is at the tip when the first branch begins to form (~34 hrs), and stays primarily localized to the tip as the bristle continues its vertical extension over time (~36 hrs - 42 hrs). After formation of the second branch, Mical localizes near the tip and in the branch (~44 hrs). Note that this bristle (one of two posterior scutellar (PSc) bristles) crosses over the other PSc bristle as it extends, and in the images at ~39 hrs and ~42 hrs the crossing bristle is present and should not be confused for a branch. (b) Syph directs Mical’s distribution to enhance Mical-mediated bristle remodeling. Developing bristles in Bristle Mical + Bristle Syph (UAS:mcherryMical/+, UAS:SyphGFP/+, B11-GAL4/+) flies were imaged over time as described in (a). During the early stages of bristle development (~32hrs), Syph does not induce notable alterations to Mical’s distribution (red) or its effects on bristle morphology. However, with increasing development (~34 hrs – ~36 hrs), Mical in combination with Syph begins to affect vertical bristle extension, inducing deviations (open arrowheads) from the main bristle shaft (closed arrowheads) and a reoriented direction of bristle growth. Note, that these Mical/Syph-dependent bristle morphological alterations also correspond with an increase in the distribution of Mical (red), such that Mical spreads out within the bristle tip and newly-forming branches (~34 hrs – ~36 hrs). Note, also, that these changes in Mical’s localization – as well as Mical/Syph-triggered bristle remodeling – become more prominent with age (~39 hrs – ~41 hrs), with both increases to the length and number of bristle branches (e.g., arrows), and an enlargement to the size and circuitous nature of the non-vertical/reoriented bristle process (open arrowhead). Moreover, with further development (~44 hrs), note that vertical growth from the main shaft resumes (closed arrowhead), but with little Mical localized within it, while the non-vertical/reoriented process becomes a large branch (arrow) containing prominent Mical (red) within it. (c) Syph triggers a broadening of Mical-mediated F-actin disruption. (c, images) Note that high levels of Mical localization (red), correspond with low levels of F-actin (green). Representative images of a developing bristle expressing mcherryMical, GFPActin, and Syph (UAS:mcherryMical/+, UAS:Syph/+, UAS:GFPActin/+, B11-GAL4/+). (c, graphs) Areas within the bristle with high Mical levels (Mical localization) and with low levels of F-actin (Actin disruption) were both measured at the bristle tip/branches. (c, top graph) Distribution of Mical and F-actin disruptions. Compared to bristles with elevated levels of Mical only (Bristle mcherryMical), bristles with elevated levels of both Mical and Syph (Bristle mcherryMical + Bristle Syph) have increased areas of both Mical localization and F-actin disruptions. (c, bottom graph) Overlap of Mical’s localization with F-actin disassembly. Examining the percentage of overlap between Mical localization and F-actin disruptions reveals that when both Mical and Syph are expressed, F-actin is more disrupted in regions where Mical is localized.

104

Figure 3.5. Further analysis of the effects of Syph on changes in the cellular distribution of Mical and Mical-mediated F-actin disassembly/cellular remodeling. (a) Elevating the bristle levels of Syph alters the distribution of Mical, such that Mical is distributed in a greater percentage of the bristle process (as determined by measuring the area of high mCherryMical levels in bristles at ~ 38-40 hours after puparium formation as a percentage of the total bristle area). ***P = 0.0001; unpaired t-test (two-tailed); n ≥ 11 bristles per genotype (1-2 bristles assessed in an animal). Mean ± s.e.m. (b) Developing bristles with increased expression of Mical and Syph (UAS:mcherryMical/+, UAS:SyphGFP/+, B11-GAL4/+).

105 Syph (green) localizes throughout the bristle process, including in regions with high levels of Mical (red). (c-d) Syph induces dramatic changes in F-actin organization in Bristle Mical flies. In both c and d, representative images are shown of different bristles at different stages of development for each genotype, where, actin is visualized by expression of a GFPActin transgene. (c) In bristles with increased expression of Mical only (Bristle Mical), a small branch (arrow) with disrupted actin (i.e., low levels of F-actin, arrowhead) typically forms during the early extension of the bristle process (c, left panel). With increasing development, the bristle grows past this first branch point, and F-actin within the shaft is mostly well- organized into bundles (c, right panel). At this time point, Mical-mediated F-actin disassembly results in a small area of actin disruption (arrowhead) at the bristle tip, which precedes the formation of a new branch. (d) Compared to Bristle Mical only flies, in bristles with increased expression of Mical and Syph (Bristle Mical + Bristle Syph), there is an increase in F-actin disruptions (d, left panel, arrowhead). Mical/Syph-triggered increases in morphological alterations are also readily noticeable at this time point including that the newly-formed branch-like deviation (d, left panel, arrow) is larger in size than in Bristle Mical only flies, and no vertical extension/tip is present beyond the point of deviation (d, left panel). Note also that with increasing age, this increase in F-actin disruption (F-actin disassembly) becomes even more pronounced (d, right panel, the region of F-actin disruption is demarcated by arrowheads). (e) Increased expression of Syph only (Bristle Syph) is sufficient to generate morphological and F-actin alterations (drawings and images, and see also Figure 3.7b for Syph localization within the bristle process). I also noted that these defects are visibly different than those that occur in Bristle Mical only flies or when both Mical and Syph are expressed together in bristles. Note also that the widespread actin disruption (i.e. paucity of F-actin) that is characteristic of Syph and Mical expressing bristles (d, Figure 3.4c) is not observed. It should also be noted that some lines expressing Syph in bristles (P{XP}d05943 and Syph in pUASP) exhibited no bristle defects on their own (but enhanced Mical effects; Figure 3.1d-f and data not shown for Syph in pUASP). Transgenic lines I made of Syph in pUAST displayed higher levels of expression and showed the documented morphological defects. Also, as described in the Methods, it should be noted that I observed a single polymorphism in the large Syph transgene I obtained from the Parkhurst lab (Liu et al., 2008). I therefore reverted the particular base pair so as to match the sequence of the Syph transgene to the Syph published genome sequence. I made different transgenic flies containing either one or the other construct and the effects were indistinguishable.

106

Figure 3.6. The myosin motor activity of Syph locally targets Mical to disassemble F- actin. (a-c) Syph-induced changes in Mical localization and activity-dependent cellular remodeling are not due to long range transport of Mical to the bristle tip. (a-c) Reducing the bristle levels of Syph (Syph21J/+; Bristle Syph RNAi) suppresses Mical-dependent bristle branching (a; see also Figure 3.2d) but does not significantly alter Mical’s localization/distribution (red) to bristle tips (as compared to Bristle mcherryMical bristles) (b-c). Mical’s distribution was visualized using mcherryMical (Bristle mcherryMical) and the distribution of Mical in the bristle process was analyzed as a percentage of the total bristle cell process area. ns (not significant), P = 0.8083; unpaired t-test (two-tailed); n ≥ 12 bristles per genotype (1-2 bristles assessed per animal). Mean ± s.e.m. (d-f) Syph-induced changes in Mical’s localization and activity-dependent cellular remodeling are dependent on Syph’s motor activity. (d) Generation of two mutant transgenes that abolished the myosin motor function of Syph. SyphR213A,G434A contains 2 point mutations that block myosin motor

107 motility. SyphΔMotor lacks the motor domain. (e-f) Syph employs its myosin motor activity to target Mical and locally disassemble F-actin. (e, left panel) Loss of Syph’s motor activity suppresses the effects of Syph on Mical activity-dependent bristle remodeling, such that Mical-dependent branches are not typically as thick or as long as they are with wild-type Syph (compare to Figure 3.1g; see also Figure 3.7c compared to Figure 3.3a). (e, middle panels) Developing bristles in Bristle Mical + Bristle SyphΔMotor (UAS:mCherryMical/+, B11- GAL4/+, UAS:SyphΔMotor/+) flies were imaged over time as described in Figure 3.4a. SyphΔMotor does not redistribute Mical nor enhance Mical-mediated bristle remodeling in a similar manner as wild-type Syph. In particular, although branch formation is often observed (~34 hrs – ~36 hrs, arrows), Mical’s expanded distribution and the prominent non-vertical extension/bristle process reorientation that is a hallmark of wild-type Syph (see Figure 3.4b) does not occur with SyphΔMotor. Note also that unlike with wild-type Syph, Mical is not predominantly relocalized to the branches (arrows) that form with SyphΔMotor (~38 hrs – ~43 hrs, compare to Figure 3.4b), but remains prominently localized to the vertically extending bristle tip. (f) Quantification of the distribution of mCherryMical (the percentage of the total bristle cell process area with mcherryMical expression) in bristles ~38-40 hrs after puparium formation reveals that without a functioning myosin motor (SyphΔMotor), Syph does not induce prominent redistribution of Mical throughout the bristle ****P <0.0001; unpaired t- test (two-tailed); n ≥ 11 bristles per genotype (1-2 bristles assessed in an animal). Mean ± s.e.m.

108

Figure 3.7. Further characterization of Syph motor mutants and their effects on Mical’s localization and cellular morphology. (a) Sequence alignment of the Switch I and II regions (red boxes) of the myosin motor domains. Consensus sequences are in yellow. The switch I arginine and switch II glycine residues, which are mutated in SyphR213A,G434A, are demarcated in bold. Dd = Dictyostelium discoideum; Dm = Drosophila melanogaster; Mm = Mus musculus; Hs = Homo sapiens. The numbering of the amino acid residues corresponds to isoform C of Drosophila Syph, isoform 2 of mouse Myosin XV, and isoform 1 of human Myosin XV. (b) Wild-type Syph (Bristle Syph = UAS:SyphGFP/+, B11-GAL4/+) and the motor mutant forms of Syph (SyphΔMotor and SyphR213A,G434A) localize throughout bristle processes. (c-d) Loss of Syph’s motor activity suppresses the effects of Syph on Mical activity-dependent bristle remodeling, such that Mical-dependent branches are not typically as thick or as long as they are with wild-type Syph (compare to Figure 3.3a). (c) Tracings of adult bristles that developed from bristles co-expressing Mical and SyphΔMotor transgenes (Bristle Mical + Bristle SyphΔMotor = UAS:Mical/+, B11-GAL4/+, UAS:SyphΔMotor/+). (d) Tracings of adult bristles that developed from bristles co-expressing Mical and SyphR213A,G434A transgenes (Bristle Mical + Bristle SyphR213A,G434A = UAS:Mical/+, B11- GAL4/+, UAS:SyphR213A,G434A/+). (f) Quantification of the distribution of mCherryMical (the percentage of the total bristle cell process area with mcherryMical expression) in bristles ~38- 40 hrs after puparium formation. As in bristles co-expressing Mical and SyphΔMotor transgenes

109 (Figure 3.6f), quantification of the distribution of Mical in bristles co-expressing Mical and SyphR213A,G434A transgenes indicates that without a functioning motor, Syph is unable to alter the distribution of Mical. ****P <0.0001; unpaired t-test (two-tailed); n ≥ 11 bristles per genotype (1-2 bristles assessed in an animal). Mean ± s.e.m.

110

Figure 3.8. Syph employs its first MyTH4 domain to redistribute Mical. (a) Generation of GFP-tagged Syph transgenic lines containing a deletion of one or more conserved protein domains within the tail region of Syph. (b) Syph uses its conserved tail region to target Mical and induce Mical-dependent F-actin/cellular remodeling. (b, drawings) Deleting the tail region domains of Syph – the MyTH4(1), PH-like, MyTH4(2), and FERM domains (SyphΔMPMF) – suppresses the effects of Syph on Mical activity-dependent bristle remodeling,

111 such that no long, thick branches form (compare with Figure 3.3a). (b, right) Developing bristles in Bristle Mical + Bristle SyphΔMPMF (UAS:mCherryMical/+, B11-GAL4/+, UAS:SyphΔMPMF/+) flies were imaged as described in Figure 3.4a. Note that SyphΔMPMF (green) localizes throughout the bristle process. Note, also, that Mical (red) remains prominently localized to the vertically extending bristle tip, indicating that unlike wild-type Syph (see Figure 3.4b), SyphΔMPMF does not redistribute Mical and induce dramatic Mical- mediated bristle morphological changes. (c) Quantification of the distribution of Mical in bristles co-expressing Mical and one of the Syph domain deletion transgenes reveals that SyphΔMPMF, SyphΔMyTH4(1&2), and SyphΔMyTH4(1) alter the Syph-induced modification to Mical’s localization. Analysis of mCherryMical distribution was performed in the time frame when the Syph-triggered broadened distribution of Mical begins to occur (typically between 34 and 36 hours after puparium formation) and is presented as a percentage of bristles with an observable change in the distribution of Mical compared to Bristle mCherryMical only expressing flies (see Figure 3.4a). n ≥ 4 bristles per genotype. See also Figures 3.9 and 3.10 for corresponding images. (d) Syph requires its first MyTH4(1) domain to localize Mical and direct Mical-dependent F-actin/cellular remodeling. (d, drawings) Deleting the first MyTH4(1) domain of Syph (SyphΔMyTH4(1)) generates bristles with less severe defects than those observed in bristles co-expressing Mical and wild-type Syph (compare with Figure 3.3a). (d, middle/right panels) Developing bristles in Bristle Mical + Bristle SyphΔMyTH4(1) (UAS:mCherryMical/+, B11-GAL4/+, UAS:SyphΔMyTH4(1)/+) flies were imaged over time as described in Figure 3.4a. Note that SyphΔMyTH4(1) (green) has more of a punctate appearance than wild-type Syph (see Figure 3.7b) but localizes throughout the bristle process. Note, however, that Mical (red) is at the tip early in bristle process extension (~34 hrs) and remains at the tip with increasing development and bristle process extension (~39 hrs). Additionally, the dramatic Syph-triggered change in morphology that accompanies Syph-induced redistribution of Mical in bristles expressing Mical and wild-type Syph does not occur, so that bristles do not significantly deviate from the main bristle shaft, but maintain their vertical growth (similar to bristles without elevated levels of Syph, see Figure 3.4a). (e) Quantification of the distribution of mCherryMical (the percentage of the total bristle cell process area with mcherryMical expression) in bristles ~38-40 hrs after puparium formation reveals that without the first MyTH4 domain (SyphΔMyTH4(1)), Syph does not induce prominent redistribution of Mical throughout the bristle. **P = 0.0014; unpaired t-test (two- tailed); n ≥ 9 bristles per genotype (1-2 bristles assessed in an animal). Mean ± s.e.m.

112

Figure 3.9. Further characterization that the PH-like and FERM domains are not required for Syph-triggered redistribution of Mical. (a) The PH-like domain of Syph is not required for Syph to target Mical and induce Mical-dependent F-actin/cellular remodeling. (a, drawings) Co-expression of Mical and SyphΔPH-like transgenes in bristles produces highly branched bristles. Note that in the left-most drawing, the 2 posterior scutellar bristles are tangled together, which also occurs with wild-type Syph, so that 2 bristles are shown, instead of 1. (a, images) Developing bristles in Bristle Mical + Bristle SyphΔPH-like (UAS:mCherryMical/+, B11-GAL4/+, UAS:SyphΔPH-like/+) flies were imaged as described in Figure 3.4a. Note that SyphΔPH-like (green) localizes throughout the bristle process. Note that compared to Bristle Mical only expressing flies (Figure 3.4a), Mical’s distribution (red) is more spread out in SyphΔPH-like expressing bristles. These results indicate that the PH-like domain of Syph is not required for Syph to redistribute Mical. (b) The FERM domain of Syph is not required for Syph to target Mical and induce Mical-dependent F-actin/cellular remodeling. The SyphΔFERM transgene lacks the FERM domain (and the 7 amino acids C- terminal to the FERM domain, which make up the end of the protein). (b, drawings) Co- expression of Mical and SyphΔFERM transgenes in bristles produces highly morphologically- altered bristles with long, thick branches. (b, images) Developing bristles in Bristle Mical + Bristle SyphΔFERM (UAS:mCherryMical/+, B11-GAL4/+, UAS:SyphΔFERM/+) flies were imaged as described in Figure 3.4a. mCherryMical (red) and SyphΔFERM (green). Note that SyphΔFERM localizes throughout the bristle process. Note also that compared to Bristle Mical only expressing flies (Figure 3.4a), bristles expressing Mical and SyphΔFERM transgenes display alterations in Mical’s localization (red), so that Mical has a wider distribution.

113

Figure 3.10. Further characterization of Mical localization and cellular morphology in bristles expressing Mical and Syph tail region domain deletions. For a-d, developing bristles were imaged as described in Figure 3.4a. (a) A requirement for the MyTH4 domains of Syph to target Mical and induce Mical-dependent F-actin/cellular remodeling. The SyphΔMyTH4(1&2) transgene lacks both MyTH4 domains in the tail region of Syph. (a, drawings) The characteristic long, thick bristle branches that are a hallmark of co-expressing wild-type Syph and Mical (see Figure 3.3a), do not form in SyphΔMyTH4(1&2) flies. (a, images) Developing bristles in Bristle Mical + Bristle SyphΔMyTH4(1&2) (UAS:mCherryMical/+, B11- GAL4/+, UAS:SyphΔMyTH4(1&2)/+). mCherryMical (red) and SyphΔMyTH4(1&2) (green). Note that SyphΔMyTH4(1&2) localizes throughout the bristle process. Note also that without both MyTH4 domains (SyphΔMyTH4(1&2)), Syph (green) localization more closely resembles wild-type Syph localization rather than the punctate localization of SyphΔMyTH4(1) (compare with Figure 3.8d). However, similar to bristles co-expressing Mical and SyphΔMPMF (see Figure 3.8b), Mical (red) is localized concisely at the bristle tip in bristles co-expressing Mical and SyphΔMyTH4(1&2). This localization pattern contrasts with the spreading out of Mical observed in bristles that co-express Mical and wild-type Syph and are at a similar stage of development

114 (see Figure 3.4b, ~36 hrs). (b) The second MyTH4 domain is not required for Syph to target Mical and induce Mical-dependent F-actin/cellular remodeling. The second MyTH4 domain (MyTH4(2)) has been removed in the SyphΔMyTH4(2) transgene. (b, images) Developing bristles in Bristle Mical and Bristle SyphΔMyTH4(2) (UAS:mCherryMical/+, B11-GAL4/+, UAS:SyphΔMyTH4(2)/+) flies. Note that SyphΔMyTH4(2) (green) localizes throughout the bristle process. Note also that the Syph-triggered broadened distribution of Mical (red) and enhanced bristle morphological defects (drawings) occur with SyphΔMyTH4(2). (c-d) Syph deletion mutants containing the first MyTH4 domain are sufficient to redistribute Mical and enhance Mical-mediated bristle remodeling. The SyphΔMF transgene consists of a truncated form of Syph containing only the portion of Syph N-terminal to the second MyTH4 domain (i.e., it lacks both the MyTH4(2) and FERM domains). The SyphΔPMF transgene also consists of a truncated form of Syph containing only the portion of Syph N-terminal to the PH-Like domain (i.e., it lacks the PH-Like, MyTH4(2), and FERM domains). In both Bristle Mical and SyphΔMF (UAS:mCherryMical/+, B11-GAL4/+, UAS:SyphΔMF/+) flies (c) and Bristle Mical and SyphΔPMF (UAS:mCherryMical/+, B11-GAL4/+, UAS:SyphΔPMF/+) flies (d) a change in Mical’s distribution (red) and alterations in cell morphology that resemble bristles expressing Mical and wild-type Syph (drawings) are induced, although the alterations in cell morphology do appear less severe in SyphΔPMF expressing bristles.

115

Figure 3.11. Sisyphus orchestrates Semaphorin/Plexin/Mical-induced morphological changes by locally controlling Mical’s subcellular localization, thereby influencing where and to what extent actin-based cellular remodeling occurs. (a) Using the bristle model for cellular process shape and extension, I find that Mical (red) localizes to the bristle tip, and elevating the levels of Mical leads to Mical-dependent F-actin disassembly which leads to a new cellular branch/protrusion (left). With increasing development, the elongating bristle grows past this region of branch formation, and while I observe a small area of actin disruption at the tip as the bristle elongates, the F-actin is well organized within the bristle shaft distal to the branch point (right). (b) I also find using the model for cellular process shape and extension that elevating the levels of Syph in combination with Mical increases the amount of Mical-mediated actin disruptions around the vicinity of the first branch point, leading to the formation of a larger cellular branch/protrusion (left). With increasing development in the presence of elevated levels of Syph, Mical’s localization begins to spread out into more of the bristle process, dramatically increasing the degree of F-actin disruptions and bristle morphological change (right). Mutations to the motor domain of Syph and also to the first MyTH4 domain in Syph suppress these effects of Syph on Mical (as shown in a).

116

Figure 3.12. Mical and Syph regulate synaptic structure and F-actin muscle organization. (a) Schematic diagram of a subset of Drosophila larval neuromuscular junctions (NMJ) illustrating the stereotypical synaptic innervation of motor axons from the ISNb nerve branch (green) on muscles 6/7 and 12/13 (red boxes). (b) Unlike wild-type synapses which spread out along muscles 6/7 (left panel), Mical mutant synapses are abnormally short (right panel; (Beuchle et al., 2007; Hung et al., 2013)). Synapses were visualized with the aid of CD8-GFP-Shaker (Beuchle et al., 2007; Hung et al., 2013; Zito et al., 1999). (c) Increasing the expression of Mical in muscles (Muscle Mical = UAS:Mical/+, 24B-GAL4/+) causes defects in skeletal muscle F-actin organization such that the characteristic striated muscle pattern of F-actin is disrupted (Hung et al., 2013). (b-c) Reprinted by permission from Macmillan Publishers Ltd: [NATURE CELL BIOLOGY] (Hung, R.-J., Spaeth, C.S., Yesilyurt, H.G., and Terman, J.R. SelR Reverses Mical-mediated Oxidation of Actin to Regulate F-actin Dynamics. 15, 1445–1454.), copyright (2013). (d) Sisyphus mutants (Syph21J/Y) exhibit a decreased length of synaptic innervation – such that at larval neuromuscular junctions, synapses are not as spread out along the muscles (images, left graph). The width of muscle 6 (right graph) is not significantly different in the wild-type and Syph mutant larvae that were analyzed for the length of synaptic innervation – indicating that synapses of similar sized muscles were analyzed for length of synaptic innervation (left

117 graph). Muscles were labelled with phalloidin (F-actin), and presynaptic active zones were labelled with the nc82 (Bruchpilot (Brp)) antibody. ns (not significant), P = 0.1395; ***P = 0.0002; unpaired t-test (two-tailed); n ≥ 17 synapses/muscles per genotype (1-3 hemisegments assessed in an animal). Mean ± s.e.m. (e) Increasing the levels of Syph in muscles (Muscle Syph = UAS:SyphGFP/+, 24B-GAL4/+) causes significant alterations in F- actin organization (disruptions to the F-actin striation patterns). Syph (green) localizes to the site of these F-actin defects.

118

Figure 3.13. Larval expression of Syph enhancer trap lines. (a) Two publicly available fly lines have GAL4-containing transposable elements inserted either upstream of the Syph gene (NP0180 = P{GawB}NP0180 from Bloomington Stock Center stock: w* P{GawB}NP0180/FM7c) or within the Syph gene and upstream of the coding region (NP4100 = P{GawB}Myo10ANP4100 from Bloomington Stock Center stock:

119 y* w* P{GawB}Myo10ANP4100/ FM7c) – so as to indicate that GAL4 may be expressed where and when Syph is expressed. (b) The P{GawB}Myo10ANP4100 (NP4100) GAL4 line crossed to UAS:ActinGFP reveals GFP expression in the larval nervous system – in areas of the brain (left and middle panels) and ventral nerve cord (VNC; left and right panels). Strong expression is also observed in skeletal muscles (left panel) and trachea (not shown). (c) The P{GawB}NP0180 line (NP0180) GAL4 line (after crossing to UAS:ActinGFP) also exhibited GFP expression in the nervous system, although I noted differences in the intensity and patterning between the two lines.

120

Figure 3.14. Summary/working model: The unconventional myosin Syph spatially regulates the localization of Mical to direct Semaphorin/Plexin/Mical-mediated F-actin disassembly and cellular remodeling. (a-b) A summary/working model based on previous characterization of Semaphorin/Plexin/Mical-mediated F-actin disassembly/cellular remodeling (a) and my results identifying Syph as a modifier of these cytoskeletal changes through its effects on the localization of Mical – that is, the motor/MyTH4(1) domain- dependent redistribution of Mical (b). (a) Semaphorin/Plexin/Mical signaling leads to local F-actin disassembly through Mical-mediated oxidation of F-actin (left panel) and results in F- actin reorganization to generate a F-actin-dependent cellular protrusion/branch (right panel). (b) My results suggest that following Mical-mediated F-actin disassembly (as in a, left panel) Syph modifies the localization of Mical, and this change in Mical’s localization is accompanied by further Mical-mediated F-actin disassembly (b, left panel) and more extensive cellular remodeling occurs (b, right panel). It remains to be determined if this

121 Syph-triggered redistribution of Mical is due to direct binding and transport of Mical (as indicated by “?”). See also main text.

122 Materials and Methods

Mical activity-dependent bristle branching genetic screen.

In all cases, Bristle Mical = UAS:Mical/+, B11-GAL4/+. The enhancer-suppressor single-cell genetic screen based on Mical activity-dependent bristle branching was performed as described in (Hung et al., 2013), with a minor change in the scoring system. Here, bristles were scored on a 5-point scale with “0” being no enhancement or suppression of Mical activity-dependent bristle branching (i.e., bristles had a similar number and length of branches as Bristle Mical). Bristles with slight suppression of the Mical activity-dependent phenotype (i.e., a branch was present, but was shorter than in Bristle Mical flies) were scored as “-1,” and bristles with no branch were scored as “-2.” Bristles with increased length and/or number of branches were considered as enhanced Mical activity-dependent branching and were scored as “+1” or “+2”, depending on the degree of increase in branch length and/or number. For simplicity and consistency, only the left posterior scutellar bristle was scored in each fly, as this is a large, easily identifiable macrochaete (single bristle cell) on the thorax of the fly. Flies with EMS-induced mutations, transposable element insertions, or UAS-tagged transgenes were screened in this assay.

Screening of additional Myosins for effects on Mical activity-dependent bristle branching.

The analysis of other myosins for effects on Mical activity-dependent bristle branching was conducted as described above and as outlined in Figure 3.3b. The following fly lines were used: Class 1a: w1118; P{RSS}Myo31DF5-na-1552 (LOF) and y1 w67c23;

P{EPgy2}Myo31DFEY08859 (GOF), Class 1b: w1118; P{EP}Myo61FEP3325a

123 P{EP}Myo61FEP3325b (LOF) and P{GSV6}GS10898/TM3, Sb1 Ser1 (GOF), Class 1c: w1118; Mi{ET1}Myo95EMB06620 (LOF) and y1 w67c23; P{EPgy2}Myo95EEY22671/TM3, Sb1 Ser1

(GOF), Class II – Mhc: y1 w67c23; P{lacW}Mhck10423/CyO (LOF) and P{XP}d06530 (GOF),

Class II-zip: cn1 bw1 sp1 zip1/CyO (LOF) and y1 w67c23; P{GSV2}GS50077/SM1 (GOF), Class

III: w*; ninaC5 (LOF), Class V: y1; P{SUPor-P}didumKG04384/CyO (LOF) and w; Bl/CyO;

UAS-MyoV-GFP/TM2 (GOF; a kind gift from Anne Ephrussi (Krauss et al., 2009)), Class

VI: w*; P{wAR}jar1646/TM3, Sb1 Ser1 (LOF) and y1 w67c23; P{EPgy2}jarEY06619 (GOF), Class

VIIa: cn1 P{PZ}ck07130/CyO; ry506 (LOF) and PBac{WH}ckf04351 (GOF), Class VIIb: w1118;

P{RS3}Myo28B1CB-6333-3 (LOF), Class XV: y1 sc* v1; P{TRiP.HMS02255}attP2 (LOF) and

P{XP}d05943 (GOF), Class XVIII: PBac{RB}Mhcle03696 (LOF) and y1 w67c23;

P{EPgy2}EY00454 (GOF), Class XX: w*; dGC13 pr1 cn1/CyO (LOF) and w*; P{UAS- d.V5}50-5/CyO (GOF). Except where noted, flies were obtained from the Bloomington

Drosophila Stock Center, Kyoto Stock Center, and Exelixis Collection at Harvard Medical

School.

Transgenic and transposable element fly lines and molecular biology.

All Mical and Plexin A transgenic fly lines were as previously described (Hung et al., 2010,

2013). The Syph fly lines w1118 PBac{WH}Myo10Af03968 (Thibault et al., 2004) and y1 sc* v1; P{TRiP.HMS02255}attP2 (Syph RNAi) were obtained from the Bloomington

Drosophila Stock Center, P{XP}d05943 and PBac{WH}f06507 (Thibault et al., 2004) were obtained from the Exelixis Collection at Harvard Medical School.

124 To generate Syph transgenic flies, standard molecular biology techniques and reagents were employed. Drosophila embryo injections to generate transgenic flies from all

Syph constructs were performed by BestGene, Inc. Wild-type Syph transgenic flies were generated using a DNA construct, Sisyphus-GFP (a kind gift from Susan Parkhurst (Liu et al., 2008)). Sisyphus-GFP is comprised of GFP fused to the C-terminus of the Syph ORF

(yielding a 2602a.a. Syph protein) in pUASp. Sequencing of this DNA construct I obtained

(Liu et al., 2008) revealed a single nucleotide difference from the NCBI reference sequence

(NM_132441.3), resulting in a histamine (H) substitution for tyrosine (Y) at amino acid 280.

Site-directed mutagenesis using the QuikChange Lightning Site-Directed Mutagenesis kit

(Strategene) was employed to change H280 to Y280 using forward (5’ –

CGGACAAGTACTTCTATCTGAACC -3’) and reverse (5’ –

GGTTCAGATAGAAGTACTTGTCCG -3’) primers.

For optimal expression in Drosophila somatic cells using the UAS-GAL4 system

(Brand and Perrimon, 1993), Sisyphus-GFP (both the construct before (H280) and after site- directed mutagenesis (Y280), as described above) was subcloned from the pUASp vector

(optimized for expression in germline cells (Rørth, 1998)) into the pUAST vector using KpnI and XbaI restriction enzyme sites. I examined the effects of both transgenes and found that they were indistinguishable. With one exception (the adult bristle tracings shown in Figure

3.5e) all data shown utilizes the Y280 construct (SyphGFP or the untagged version of this construct (see below)). Eight independent SyphGFP (i.e., Y280 line in pUAST) transgenic lines and five independent SyphH280 (in pUAST) transgenic lines were generated and all showed similar phenotypes when expressed in bristles. To make an untagged version of Syph

125 in pUAST, a fragment was PCR-amplified from SyphGFP DNA in pUAST using a forward primer (5’ – CGTGTCATCCAGGCTAGCAT – 3’) that recognized a unique NheI site within Syph (amino acids 1606-1607) and a reverse primer (5’ -

CCTCTAGATTAGTTGAGCTCCCG – 3’) to engineer a Stop-XbaI site at the 3’ end of

Syph and then ligated into SyphGFP pUAST digested with NheI and XbaI – thereby removing

GFP. Three independent transgenic lines were made, and they all gave similar bristle phenotypes. Importantly, bristle expression of the untagged and GFP fusion forms of Syph caused similar phenotypes, indicating that the C-terminal GFP did not alter the function of

Syph. SyphR213A,G434A was made using the QuikChange Lightning Site-Directed Mutagenesis kit (Strategene) to sequentially introduce point mutations (R213 to A213, forward: 5’ –

GACAATAGTTCGCGGTTTGGAAAGTATCTGG – 3’, reverse: 5’ –

CCAGATACTTTCCAAACCGCGAACTATTGTC - 3’ and G434 to A434, forward: 5’ –

CTGGACATCTTTGCGTTCGAGGATTTGGC – 3’, reverse: 5’ –

GCCAAATCCTCGAACGCAAAGATGTCCAG – 3’) into SyphGFP pUAST in the conserved switch I (consensus sequence: NXNSSRFG) and switch II (consensus sequence:

DIXGFE) regions of the Syph motor domain. Two independent SyphR213A,G434A transgenic lines were generated and both showed similar bristle phenotypes.

The generation of all domain deletions was based on identification of conserved domains using the NCBI Conserved Domain Database (CD-Search tool; (Marchler-Bauer et al., 2011)). All Syph domain deletion DNA is in pUAST. Between 2-10 independent lines were generated for each construct, and the bristle phenotypes resulting from expression of each line were recorded. With a few exceptions, all independent lines of a given construct

126 showed similar phenotypes when expressed in bristles. Outliers were not used for subsequent experiments. The primers used to generate all domain deletion transgenes are listed in Table

3.1. SyphΔPMF (a.a.1-1788 of Syph fused to GFP; lacks the PH-like, MyTH4(2), and FERM domains) was generated in a two-step cloning process. First, a shortened SyphGFP pUAST construct (a.a.1-1607 of Syph fused to GFP) was created using PCR to amplify a NheI-GFP-

XbaI fragment which was then ligated into SyphGFP pUAST digested with NheI/XbaI, thereby producing a Syph construct lacking the C-terminal tail region containing the PH-like,

MyTH4(2), and FERM domains. To complete the construct, an additional (5’)SpeI/(3’)NheI-

DNA fragment coding for a.a.1606-1788 of Syph was PCR-amplified and ligated into the shortened SyphGFP pUAST digested with NheI. SyphΔPH-like (a.a.1-1788;1895-2602 of Syph fused to GFP) was generated by PCR-amplifying a (5’&3’)NheI-DNA fragment coding for a.a.1895-2602 of Syph and ligating the fragment into SyphΔPMF digested with NheI. SyphΔMF

(a.a.1-2019 of Syph fused to GFP; lacks the MyTH4(2) and FERM domains) was generated by PCR-amplifying a (5’)SpeI/(3’)NheI-DNA fragment coding for a.a.1606-2019 of Syph and ligating the fragment into the shortened SyphGFP pUAST construct at the NheI site.

SyphΔMyTH4(2) (a.a.1-2119; 2269-2602 of Syph fused to GFP) was generated by PCR- amplifying a (5’&3’)NheI-DNA fragment coding for a.a.2269-2602 of Syph and ligating the fragment into SyphΔMF digested with NheI. SyphΔFERM (a.a.1-2279 of Syph fused to GFP) was generated by PCR-amplifying a (5’)SpeI/(3’)NheI-DNA fragment coding for a.a.2120-2279 and ligating the fragment into SyphΔMF digested with NheI. SyphΔMyTH4(1) (a.a 1-922;1081-

2602 of Syph fused to GFP) was generated in a two-step process. First, a (5’)AvrII/(3’)NheI-

DNA fragment coding for a.a.260-922 was PCR-amplified and ligated into the SyphGFP

127 pUAST construct at unique AvrII and NheI sites within Syph. Second, a (5’)SpeI/(3’)NheI-

DNA fragment coding for a.a.1081-1607 was amplified and ligated into the vector generated in the first step digested with NheI. SyphΔMyTH4(1&2) (a.a.1-922;1081-2119; 2269-2602 of

Syph fused to GFP; lacking both MyTH4 domains) was generated by PCR-amplifying a

(5’)SpeI/(3’)XbaI-DNA fragment coding for a.a.1606-end of GFP from the SyphΔMyTH4(2) construct and ligating the fragment into SyphΔMyTH4(1) digested with NheI/XbaI. SyphΔMPMF

(a.a.1-922;1081-1788 of Syph fused to GFP; lacking all conserved tail domains) was generated by PCR-amplifying a (5’)SpeI/(3’)XbaI-DNA fragment coding for a.a.1606-end of

GFP from the SyphΔPMF construct and ligating the fragment into SyphΔMyTH4(1) digested with

NheI/XbaI. SyphΔMotor (a.a.1-63; 749-2602 of Syph fused to GFP) was generated by a two- step process. First a SyphΔNterm-motor construct (a.a.749-2602 of Syph fused to GFP) was generated by PCR-amplifying a (5’)KpnI/(3’)SfiI-DNA fragment coding for a.a.749-1847 of

Syph and ligating the fragment into SyphGFP digested with KpnI/SfiI. Then, a (5’&3’)KpnI-

DNA fragment coding for a.a.1-63 of Syph was PCR-amplified and ligated into SyphΔNterm- motor digested with KpnI.

Generation of Syph mutant flies.

FRT-containing transposable elements (PBac{WH}Myo10Af03968 and PBac{WH}f06507) inserted within Syph were utilized to delete the entire coding region of Syph through FLP- mediated recombination using the procedure described in (Parks et al., 2004). Successful deletion of the targeted-region was screened by PCR using WH transposon primers (WH5’ plus (reverse) and WH3’ plus (forward) - listed in (Parks et al., 2004)) together with genomic

128 primers (forward: 5’ - AGCCACGCATACACAAGCAC - 3’; reverse: 5’ –

ATGAGGACGAGGACGACGAG – 3’). Two deletion mutants were generated (Syph14B,

Syph21J). For further confirmation of these mutants, additional PCR was done using primers

(forward: 5’ – ATCAATCAACAATACACGCACC - 3’; reverse: 5’ -

AGGATGACAGCGATCAACTCA – 3’) to amplify a fragment including the transposable element and the genomic DNA flanking it. The DNA fragment was excised from an agarose gel using a QiaQuick gel extraction kit (Qiagen), cloned into pCR2.1-TOPO (ThermoFisher

Scientific), and sequenced to verify the deletion. Both deletion lines gave similar results in the Mical activity-dependent bristle branching assay, but for simplicity only Syph21J is shown here and was used for all subsequent analysis.

Adult bristle characterization, quantification, imaging, and drawings.

The bristles of recently emerged adults were examined using a dissecting stereomicroscope

(Leica S8 APO, 1.6X objective). During this examination, bristle morphology/defects were noted as described in (Hung et al., 2010). Additionally, the number of branches on the left posterior scutellar bristle of each fly was counted and recorded for quantification of branches per bristle. For quantification of bristle branching in Plexin overexpression flies (Bristle

PlexA = UAS:HA-PlexA), all 4 scutellar bristles were examined for branches. If 1 or more of the 4 scutellar bristles had branching, the fly was considered to have scutellar bristle branching. After examination, flies were stored in 70% ethanol for preservation. For imaging, the wings, legs, and abdomen were removed using Dumont #5 forceps (Ted Pella, Inc) leaving the head and thorax intact. The head of each fly was stuck to double-sided tape (3M)

129 on a slide. Bristle images were taken and compiled with a Zeiss Discovery M2 Bio stereoscope with a motorized zoom and focus and Zeiss Axiovision software and Extended

Focus Software (a gift from Bernard Lee). From these images, bristles were carefully traced in Microsoft PowerPoint using the “curve” tool to generate bristle drawings that accurately reflect bristle morphology. All images shown are of posterior scutellar bristles.

Pupal characterization and imaging.

Pupae were collected and placed on double-sided tape (3M) in Petri-dishes. Staging was done as described in (Bainbridge and Bownes, 1981). Genotyping was done with the aid of balancer chromosomes with the Tb1 marker, or using a Zeiss Discovery M2 Bio stereomicroscope and visualization of GFP balancer chromosomes or GFP or mCherry fusion proteins. Pupae were kept at 25°C until they reached the desired stage for imaging. Then, pupae were removed from their outer pupal case and whole pupae were mounted, dorsal side up, in depression well slides with VectaShield Mounting Medium (Vector Laboratories) and imaged within 4 hours with a Zeiss LSM510 confocal microscope using the x63 oil objective. Images were compiled using ZEN lite software (Zeiss).

F-actin organization and cellular distribution of Mical in bristles.

F-actin organization (observed using a GFPActin transgene) and Mical (observed using a mCherryMical transgene) distribution in bristles were calculated as a percentage of the total bristle process using images acquired from a Zeiss LSM510 confocal microscope. ImageJ software was used to trace (using the “polygon selection” tool) and measure the area of the

130 bristle process (including branches, but excluding the cell body). Similarly, the area of the bristle with low levels of F-actin (actin disruption) or high Mical expression (Mical localization) was traced and measured. As a means to delineate the regions of the bristle with high Mical expression, the “Fire” Lookup-table in ImageJ was utilized to highlight where

Mical was strongly expressed. All quantifications were done on posterior scutellar bristles and bristles of similar age (~38-40 hours after puparium formation), unless otherwise specified.

Larval neuromuscular junction innervation and muscle analysis.

Third instar larvae were collected from vial walls and dissected as previously described

(Budnik et al., 2006) to allow for visualization of the nervous system and muscles. Syph21J mutants from a Syph21J/FM7c, P{2xTb1 RFP}FM7c, sn+ stock were identified based on the lack of the Tb1 marker (balancer from Bloomington Drosophila Stock Center). Syph21J mutants were all males due to the lack of viable mutant males in the stock to produce homozygous females. After dissection, larvae were fixed in methanol for 20 minutes, then washed 3x in PBS. For histochemistry of synaptic studies, larvae were blocked with 5% Goat

Serum (Sigma-Aldrich) in PBS with 0.1% Triton X-100 (Fisher Scientific) for 1 hour at room temperature. Larvae were then incubated in a Brp (nc82) antibody (Developmental

Studies Hybridoma Bank; 1:500) in PBS for 1-2 hours at room temperature or overnight at

4°C, and then washed 5x in PBS with 0.1% Triton X-100. Next, larvae were incubated in

Alexa Fluor 635-conjugated phalloidin (ThermoFisher Scientific; 1:100) and goat anti-mouse

Alexa Fluor 488-conjugated IgG (ThermoFisher Scientific; 1:500) in PBS for 1-2 hours at

131 room temperature or overnight at 4°C, and then washed in PBS with 0.1% Triton X-100 prior to mounting on glass slides using VectaShield mounting medium (Vector Laboratories). For analysis of F-actin organization in muscles, larvae were incubated in Alexa Fluor 635- conjugated phalloidin (ThermoFisher Scientific;1:100). Larvae were imaged with a Zeiss

LSM510 confocal microscope using the x40 oil (synapses) and x20 (muscles) objectives.

Images were compiled using ZEN lite software (Zeiss). The length of synaptic innervation at muscles 6/7 was measured in ImageJ as previously described (Hung et al., 2013). NMJ synapse length is correlated with the size of the muscle being innervated (Balice-Gordon et al., 1990; Herrera et al., 1991; Li et al., 2002; Kuno et al., 1971; Nudell and Grinnell, 1983;

Ogata and Yamasaki, 1985; Wilkinson and Lunin, 1994; Wilkinson et al., 1992); therefore, muscle width was measured to make sure muscle size of all samples and genotypes was similar and synaptic innervation was comparable. Muscle width was measured in ImageJ using the “straight” tool to make a horizontal line from one side of muscle 6 to the other side, and the length of the line was measured. Quantification of F-actin organization defects was done by identifying the presence of abnormalities in the internal layers of muscles 6 and 7 (as previously described in (Beuchle et al., 2007; Hung et al., 2013)). Muscles were considered to have F-actin organization defects if any deviation in actin such as F-actin accumulation or disruptions in the normal striation pattern were present.

132

Table 3.1. Primers used to generate the Syph tail region domain deletion transgenes. Because of the strategy employed to make these transgenes, some primers were used more than once.

CHAPTER FOUR

Summary and Conclusions

Reorganization and remodeling of the cytoskeleton through actin assembly and disassembly drive cell motility and morphological changes. Extracellular guidance cues play an integral role in this process by binding cell surface receptors and initiating signaling cascades that activate or inactivate various actin regulatory proteins/effector proteins to promote actin assembly or disassembly (Berzat and Hall, 2010; Kay et al., 2008; Stossel,

1989, 1993; Swaney et al., 2010; Tessier-Lavigne and Goodman, 1996). However, the precise mechanisms by which actin reorganization is mediated and spatially regulated under these circumstances is still unclear. Recently the redox enzyme, Mical, was identified as a critical effector protein in signaling mediated by Semaphorin family guidance cues and their plexin receptors (Terman et al., 2002). Further characterizations indicated that

Semaphorin/Plexin-signaling activates Mical’s redox activity (Hung et al., 2010; Schmidt et al., 2008), and once activated, Mical oxidizes specific methionine residues of actin – leading to localized F-actin disassembly and subsequent cellular remodeling (Hung et al., 2011). My thesis work expands upon this model of Semaphorin/Plexin/Mical signaling by providing evidence of how additional proteins contribute to this pathway through functional interactions with Mical. Specifically, this work unveils novel functional interactions between

Mical and cofilin and Mical and Sisyphus and provides insights into mechanisms by which targeted Mical-mediated actin disassembly/cellular remodeling is precisely carried out and controlled.

133 134 Mical works with cofilin to rapidly disassemble F-actin and promote Semaphorin- mediated cellular remodeling

Although Mical directly disassembles F-actin downstream of Semaphorin by specific oxidation of actin (Hung et al., 2010, 2011), it was unclear if other actin regulatory proteins, especially those known to exert effects at similar sites within actin, affected Mical-mediated actin disassembly. Using in vitro and in vivo assays, my colleagues and I found that Mical works in synergy with cofilin, a well-known, ubiquitously-expressed actin regulatory protein

(Bernstein and Bamburg, 2010; Bravo-Cordero et al., 2013), to facilitate rapid F-actin disassembly and instability. In vitro assays with purified proteins showed that Mical oxidation of F-actin occurs rapidly and weakens the mechanical properties of F-actin. Cofilin preferentially binds Mical-oxidized F-actin and helps to rapidly disassemble the filaments at an exponentially faster rate than either effector protein alone. These findings improve our understanding of Mical oxidation-mediated effects on actin filaments. Likewise, by weakening the mechanical properties of actin and improving cofilin-actin binding, our results reveal that Mical-mediated oxidation of actin enhances cofilin’s ability to dismantle actin. In support of these in vitro findings, I found that cofilin and Mical also work together in a similar manner in vivo. Specifically, I found that altering levels of cofilin significantly modifies Mical activity-dependent F-actin disassembly and cellular remodeling – such that

Mical activity-dependent cellular remodeling is significantly suppressed in cofilin heterozygous mutants and significantly enhanced by constitutively active cofilin.

Additionally, we found that cofilin works with Mical in the Semaphorin signaling pathway to mediate proper cellular remodeling and axon guidance. These findings reveal new

135 mechanisms underlying Semaphorin/Plexin/Mical-mediated repulsive effects on the cytoskeleton, and they provide insights into how actin regulatory proteins work together in vivo to promote rapid and precise changes in the cytoskeleton downstream of extracellular guidance cues.

Mical’s subcellular localization is regulated by Sisyphus to spatially control

Semaphorin/Plexin/Mical-mediated actin disassembly

My findings also provide novel insights into the spatial control of

Semaphorin/Plexin/Mical-mediated F-actin disassembly and cellular remodeling. Using genetic assays, I found that an unconventional myosin, Sisyphus (Syph; a class XV family member) modulates and is required for Semaphorin/Plexin/Mical-mediated F-actin reorganization/cellular remodeling. Furthermore, my results revealed that Syph locally targets the subcellular distribution of Mical through a mechanism dependent on its myosin motor activity and one of the domains (the first MyTH4 domain) present in the tail region of

Syph. Moreover, this Syph-induced modification of Mical’s localization corresponded to dramatic changes in F-actin disassembly. Thus, my findings support that Syph locally regulates Mical-dependent F-actin disassembly and cellular remodeling by controlling the local distribution of Mical. Because Syph spatially regulates Mical, and cofilin is required for Mical-mediated actin disassembly/cellular remodeling, it will be interesting to examine if

Syph may also regulate the subcellular localization of cofilin to precisely direct rapid actin disassembly downstream of Semaphorin/Plexin-signaling.

136 My work also provides insights into the function of Syph and other class XV myosins. Class XV myosins in insects and mammals are known to function as cargo transporters in actin-rich cellular protrusions, and they have been implicated in the formation/elongation of and F-actin organization within these protrusions, although their exact roles in these processes are not clear (Anderson et al., 2000; Belyantseva et al., 2003,

2005; Liu et al., 2008; Manor et al., 2011; Mauriac et al., 2017; Probst et al., 1998). Here, I find that Syph can regulate actin disassembly through Mical, thus revealing a novel function of Syph. Furthermore, to characterize the functional interaction between Mical and Syph, I generated Syph “knockout” mutants (i.e. these mutants lack the coding region of the Syph gene) and numerous transgenic flies to enable tissue-specific expression of wild-type Syph, as well as other forms of Syph with various domain deletions, including a motor mutant form of Syph. Thus, my thesis work also provides genetic tools for further studies on the functions of Syph in Mical-mediated cytoskeletal rearrangements and in other contexts.

Broad implications and Future Directions

Mical is a critical actin regulatory protein that functions in numerous tissues including within the nervous system where it plays prominent roles in axon guidance, synaptogenesis, dendritic organization, and other cellular processes that underlie neural connectivity

(reviewed in Wilson et al., 2016; Zhou et al., 2011b). Mical and its reversible Redox mechanism to regulate actin dynamics has also been linked to a growing number of neuronal and non-neuronal diseases and disorders – including defects in neural connectivity and the inhibition of axon regeneration after injury (reviewed in Wilson et al., 2016). My work

137 herein, therefore, provides a new understanding of Mical and the mechanisms that promote and regulate Mical-mediated actin disassembly and cellular remodeling. Furthermore, it provides an important new understanding of how F-actin disassembly is regulated to drive the formation and function of neurons and other cells. Future work should determine if these new molecules and mechanisms serve as potential therapeutic targets to attenuate Mical activity-mediated pathologies.

Moreover, the identification and characterization of a functional interaction between

Mical and Sisyphus also posits that Mical might be critical in Sisyphus/Myosin XV- dependent physiological and pathological processes. Myosin XV transports cargo in actin- rich stereocilia, regulates the elongation and maintenance of stereocilia, and is required for hearing in mammals (Anderson et al., 2000; Belyantseva et al., 2003, 2005; Fang et al., 2015;

Manor et al., 2011; Mauriac et al., 2017; Probst et al., 1998; Wang et al., 1998). However, the mechanisms by which Myosin XV promote normal auditory function are still being elucidated. Furthermore, through its function as a cargo transporter, Sisyphus is required for the proper cell alignment and adhesion events underlying the morphogenetic process of dorsal closure in Drosophila (Liu et al., 2008) – a process resembling and often used as a model to understand vertebrate wound healing mechanisms (Belacortu and Paricio, 2011). As with my bristle model, these Syph-dependent processes also rely on the tight regulation of F- actin to direct and specify cellular process extension and morphology; thus, future work should explore the role of Mical in Myosin XV-related deafness and wound-healing.

Additionally, previous work has identified a role for Mical in regulating synaptic structure development (Beuchle et al., 2007; Hung et al., 2013), and I found that Sisyphus is

138 also required for this process. However, the mechanism by which either of these proteins regulate synapse structure remains unknown. Given my results using the bristle model system, it will be particularly interesting to examine if the movement of Mical to specific regions of the developing synapse, mediated by Sisyphus, is required to regulate the length of synaptic innervations. This finding is also exciting in that it identifies a novel role for class

XV myosins in neural development; whereas, a role for Sisyphus/class XV myosin in the nervous system was not previously known.

My findings demonstrate that Sisyphus and Mical are functionally interacting to remodel the cytoskeleton, and this occurs in the context of Semaphorin/Plexin signaling.

While these results identify a novel mechanism in spatially regulating actin dynamics and cellular remodeling downstream of Semaphorin/Plexin signaling to control cell shape and extension/motility (i.e. through the modification of the localization of Mical), they also spur many interesting questions and avenues for future exploration. For example, considering the known functional role of Sisyphus as an actin-based cargo transporter, it is of particular interest to determine if Sisyphus is directly transporting Mical and also what role other known Sisyphus/Myosin XV cargo play in Semaphorin/Plexin/Mical-mediated F-actin disassembly and cellular remodeling. Although Syph dramatically enhances Mical-mediated

F-actin disassembly to drive significant morphological changes, the complex cellular remodeling observed in response to Semaphorin/Plexin/Mical-mediated signaling is likely to also require actin polymerization - given that actin polymerization helps drive extension/motility and changes in cell shape (Pollard and Cooper, 2009), and that F-actin serves as the intracellular track for Sisyphus/Myosin XV-based transport. In stereocilia of the

139 inner ear, mammalian Myosin XV has been shown to transport or direct the specific localization of a complex of proteins that promote actin polymerization (Belyantseva et al.,

2005; Manor et al., 2011; Mauriac et al., 2017); therefore, it is interesting to consider that

Sisyphus might also be regulating actin polymerization and perhaps may be responsible for the localization of a large complex of actin regulatory proteins to precisely control actin dynamics. Additionally, a yeast two-hybrid screen found that Sisyphus binds several proteins that regulate microtubule dynamics – including Katanin 60 (a microtubule binding/severing protein) and EB1 (Liu et al., 2008). Furthermore, knockdown of Sisyphus slowed the movement of EB1 in cells (Liu et al., 2008), suggesting that Sisyphus does not just bind EB1, but controls its movement. EB1 binds to the distal plus tips of microtubules and acts as a molecular scaffold to recruit other microtubule regulatory proteins that regulate the stability of microtubules and promote microtubule polymerization. EB1 and other microtubule end- binding proteins also act as adaptor proteins to allow microtubules to bind other cellular components, including actin and the plasma membrane (reviewed in (Akhmanova and

Steinmetz, 2010, 2015)). Therefore, in addition to regulating the localization of Mical, as I have demonstrated, Sisyphus may also be responsible for the localization of other proteins to regulate cytoskeletal dynamics in the Semaphorin/Plexin signaling pathway. Thus, future exploration should also consider how the Sisyphus-Mical interaction is coordinated with other actin and microtubule regulatory proteins to induce specific alterations in the cytoskeleton.

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