THE ROLE OF SCHWANN CELLS IN

PERIPHERAL NERVE DEVELOPMENT IN ZEBRAFISH

A DISSERTATION

SUBMITTED TO THE DEPARTMENT OF DEVELOPMENTAL BIOLOGY

AND THE COMMITTEE ON GRADUATE STUDIES

OF STANFORD UNIVERSITY

IN PARTIAL FULFILLMENT OF THE REQUIREMENTS

FOR THE DEGREE OF

DOCTOR OF PHILOSOPHY

Alya Raphael

January 2011

© 2011 by Alya Rachel Raphael. All Rights Reserved. Re-distributed by Stanford University under license with the author.

This work is licensed under a Creative Commons Attribution- Noncommercial 3.0 United States License. http://creativecommons.org/licenses/by-nc/3.0/us/

This dissertation is online at: http://purl.stanford.edu/dm455gb2152

ii I certify that I have read this dissertation and that, in my opinion, it is fully adequate in scope and quality as a dissertation for the degree of Doctor of Philosophy.

William Talbot, Primary Adviser

I certify that I have read this dissertation and that, in my opinion, it is fully adequate in scope and quality as a dissertation for the degree of Doctor of Philosophy.

Ben Barres

I certify that I have read this dissertation and that, in my opinion, it is fully adequate in scope and quality as a dissertation for the degree of Doctor of Philosophy.

Thomas Clandinin

I certify that I have read this dissertation and that, in my opinion, it is fully adequate in scope and quality as a dissertation for the degree of Doctor of Philosophy.

David Kingsley

Approved for the Stanford University Committee on Graduate Studies. Patricia J. Gumport, Vice Provost Graduate Education

This signature page was generated electronically upon submission of this dissertation in electronic format. An original signed hard copy of the signature page is on file in University Archives.

iii ABSTRACT

Myelin is a specialized sheath that insulates axons and allows for the rapid conduction of action potentials. In the peripheral nervous system, myelin is made by glial cells called Schwann cells. Whereas Schwann cells have received attention mainly because of their role in generating myelin, they play many other important roles. Using the posterior lateral line nerve in zebrafish, I have investigated two aspects of Schwann cell development. First, I demonstrate a new role for Schwann cells in repositioning peripheral nerves. Second, I find that ErbB signaling and Schwann cell proliferation are required during radial sorting, a process during which Schwann cells change their interactions with axons and reorganize the structure of the nerve itself.

The posterior lateral line nerve is a prominent peripheral nerve in zebrafish that innervates sensory organs called neuromasts. I find that the posterior lateral line nerve initially grows out within the epidermis but is then rapidly transitioned across the basement membrane into the subepidermal space. Schwann cells are required for this process; in three different mutants that lack Schwann cells, the nerve is consistently mislocalized to the epidermis. This mislocalization results in significant disorganization of the nerve that worsens over time. When wildtype Schwann cells are transplanted into mutants lacking Schwann cells the position of the nerve is rescued. These results provide evidence that moving the posterior lateral line nerve out of the epidermis, across the basement membrane, protects the nerve from the migration of its targets.

iv

Before Schwann cells can make myelin around an axon, they must go through several developmental steps. Immediately after Schwann cells and axons have completed their migration, each Schwann cell associates with many axons. In order for myelination to proceed, however, a Schwann cell must associate with only a single axon. This transition, termed radial sorting, results in significant reorganization of the nerve. Neuregulin signaling from axons through ErbB receptors on Schwann cells controls many aspects of Schwann cell development and in mammals both

Neuregulin/ErbB signaling and Schwann cell proliferation have been implicated in radial sorting. This prompted me to investigate whether ErbB signaling was required to regulate Schwann cell proliferation during radial sorting. To test this I took advantage of the zebrafish model system and used small molecule inhibitors to inhibit either ErbB signaling or proliferation at the time when radial sorting is beginning. I find that ErbB signaling and Schwann cell proliferation are critical for radial sorting.

ErbB signaling was also required for Schwann cells to extend processes into the axon bundle, while proliferation was not. Therefore, I propose that ErbB signaling is directly required during radial sorting to regulate Schwann cell process extension, in addition to the previously established role of ErbB signaling in stimulating Schwann cell proliferation.

v ACKNOWLEDGEMENTS

This thesis could never have been completed without the help and support of many, many people. And the internet. Listing all of them would make for a document longer than the thesis itself, so, thank you everyone!

None of this would have been possible without the amazing support staff for

Developmental Biology. Thanks especially to Vanessa Bravo, Sue Elliott, and Yong

Chong. And very especially to Todd Galitz, who fixes everything, finds everything, and does it all with a smile on his face. I will miss stalking him through the halls of

Beckman by following the sounds of his laughter.

I spent at least half of my time at Stanford in the TEM facility, so I owe a major debt of gratitude to John Perrino and Lydia-Marie Jorbert for always making it such a welcoming, cheerful, and pleasant environment. And for answering all of my questions! And for being wonderful people.

I want to thank Will Talbot, for letting me join his lab and for giving me the freedom, time, and guidance to find my way to two projects that I really love. For all of his guidance and support and knowledge over the years, and especially in the final year as everything was coming together and finishing. I want to thank him especially for his amazing attention to detail and for going through every line of everything that I wrote, making the writing better and tighter and more scientific.

vi

Thanks to my committee: David Kingsley, Ben Barres, Tom Clandinin and my chair,

Kang Shen. Thanks to David Kingsley for always being in my head, making me a better scientist, asking me why I’m doing an experiment. Ben Barres for being the fastest email responder ever, and always remembering my science. Tom Clandinin for spending an entire afternoon teaching me how to use our microscope better – it has revolutionized the way we take images in our lab. Kang Shen for always being interested in what I’m doing, for his support and encouragement.

Thanks to Lucy Shapiro for really embracing me and welcoming me into the McShap lab family. And for all her advice and enthusiasm and warmth. Matt Scott for convincing me to come to Stanford and for all the stories and pictures. Sue

McConnell for making me a better teacher and speaker.

All the members of the Talbot lab throughout my time here have been fantastic – they are an incredibly fun and wise group of people. Thanks to everybody for all your help over the years and for dealing with my total insanity. And for generally putting things back in the right spot. Mrwah.

Dave Lyons for his infinite wisdom, good cheer, guidance, support, and for suggesting the experiment that led to chapter 3. Sara Mercurio for her endless cheerleading and for being the best landlady ever. Kelly Monk for all the songs, sage science and shopping advice, tasty meals, and telling me to go home. Tom Glenn for the evening

vii conversations about science and everything else. Marguax Bennett for being the most fun and enthusiastic person around. Harwin Sidik for bringing such joy and fun back into the lab – you’re going to do great! Tuky Reyes for always being unbelievably helpful and kind and taking amazing care of our fish. Chenelle Hill for making every day fun – my life is going to be so boring without you in it! Nitzan Sternheim for being an amazing role model. Matt Voas for being as an insane as I am, writing the best protocols imaginable, and for choreographing our show stopping rendition of the

Time Warp. Thanks to everyone else in the Talbot lab throughout the years: Ian,

Heather, Naomi, Isaac, Ali, Claudia, Stephen, Ryan, Jeanette, and Celia.

I think it is safe to say that Julie Perlin is just the best all around human being I have ever met. Her constant enthusiasm, support, kindness, and lunch company made coming to lab everyday a pleasure. She is the best friend anyone could ask for.

Thanks to the McShap lab poker gang, especially Franklin, Lisandra, Ayla, Leticia,

Grant, Anya, Katya, Antonio, Nikki, Paola, Monica, and Jen.

Thanks to my parents for always telling me that I could do anything I wanted to and for supporting me on my quest. Thanks for trying so hard (and mostly succeeding!) to understand what it is that I do, and why we care about Schwann cells. Thank you for always being there with guidance, support, and love.

viii I have the awesomest siblings on the planet – Naomi, Jake, and Goldie. It’s been so much fun to watch them grow into themselves and I’m so proud of who they are becoming. They always make me laugh…I’ll put it in a box, and put that box in another box, and then I’ll mail it to myself, and then I’ll smash it with a HAMMER!

If there’s one person that this really couldn’t have been done without, it would be

Esteban Toro. He’s been there for me since the beginning, and has been unbelievable generous with his encouragement, support, and advice. And love. Thanks for talking to me in the silly voice when I need cheering up and introducing me to all sorts of new and exciting foods and places and experiences. Thanks for making me a better, happier person.

Finally, I would like to dedicate this thesis work to my grandparents. To my maternal grandparents, Hal and Shirley Schneider, MomMom and PopPop, for being my biggest fans. PopPop for nurturing an early love for math and science and nature and

MomMom for teaching me to love words and crafts and order. And for Smallwood and blueberry pancakes. To my paternal grandmother, Nana Billie Raphael Sher, who had 14 grandchildren and made each one of us feel special. She died as I was finishing my thesis work and I miss her deeply.

ix TABLE OF CONTENTS

Chapter 1: General Introduction 1

The posterior lateral line nerve 3 Myelination in the peripheral nervous system 6 Schwann cell development 6 Radial sorting 7 Signals controlling radial sorting 9 Differentiated Schwann cells 11 Promyelinating Schwann cells 11 Myelinating Schwann cells 12 Non-myelinating Schwann cells 12 Neuregulin/ErbB signaling 13 Summary of research and future directions 15

Chapter 2: Schwann cells reposition a peripheral nerve to isolate it from postembryonic remodeling of its targets 21

Summary 22 Introduction 23 Materials and methods 26 Results 28 Discussion 33

Chapter 3: An analysis of ErbB signaling and Schwann cell proliferation during radial sorting in zebrafish 50

Summary 51 Introduction 52 Materials and methods 56 Results 58 Discussion 64

Appendices 84

References 98

x LIST OF ILLUSTRATIONS

Figure 1.1: Posterior lateral line development 17 Figure 1.2: Radial sorting 19

Figure 2.1: The PLLn invades the epidermal basement membrane 38 Figure 2.2: PLLn outgrowth is normal in Schwann cell deficient mutants 40 Figure 2.3: The PLLn is mislocalized in mutants lacking Schwann cells 42 Figure 2.4: Full length clones of transplanted wildtype Schwann cells can rescue nerve position in sox10/clst3 mutants 44 Figure 2.5: Shorter length clones of transplanted wildtype Schwann cells can partially rescue the position of sox10/clst3 mutant nerves 46 Figure 2.6: Nerves become disorganized during development in mutants lacking Schwann cells 48

Figure 3.1: ErbB signaling and Schwann cell proliferation are required for the expression of myelinating and promyelinating markers 68 Figure 3.2: Radial sorting is underway at 48 h and continues over time in the posterior lateral line nerve 70 Figure 3.3: ErbB signaling and Schwann cell proliferation are required for radial sorting 72 Figure 3.4: Another cell cycle inhibitor, camptothecin, also inhibits radial sorting 75 Figure 3.5: ErbB signaling and Schwann cell proliferation play distinct roles in radial sorting 77 Figure 3.6: Inhibiting ErbB signaling or proliferation arrests radial sorting 80 Figure 3.S1: Larvae treated with small molecule inhibitors 82

xi LIST OF APPENDICES

Appendix 1: Schwann cells are required for proper axon number in the PLLn 84 Appendix 2: αII-spectrin is required for myelination of motor axons 94

xii Chapter 1

General Introduction

1 INTRODUCTION

Myelin is a specialized membrane sheath that allows for the rapid conduction of action potentials along axons (Hodgkin, 1964). Myelin reduces capacitance of the axonal membrane, and allows for saltatory conduction (Stampfli, 1954). Defects in the myelin sheath have severe repercussions, and can result in diseases such as Multiple

Sclerosis and Charcot Marie Tooth disease (Berger et al., 2006; McQualter and

Bernard, 2007). Specialized glial cells, oligodendrocytes in the central nervous system and Schwann cells in the peripheral nervous system, wrap their membranes many times around a segment of an axon to create the myelin sheath (Bunge, 1968;

Geren and Raskind, 1953). Along its length, each axon is ensheathed by multiple myelin segments, which are separated by unmyelinated gaps called nodes of Ranvier

(Bunge, 1968; Tasaki, 1959). Oligodendrocytes are able to interact with and elaborate myelin sheaths around many different axons (Bunge, 1968). In contrast, Schwann cells myelinate only one segment of one axon (Bunge, 1968).

While making myelin is the most widely known function of Schwann cells, they play many other important roles during development (Jessen and Mirsky, 2005). Schwann cells are also required to provide trophic support for axons and remodel the developing nerve through the process of radial sorting (Jessen and Mirsky, 2005; Webster et al.,

1973). Recently, new roles for Schwann cells have emerged, including properly positioning the nodes of Ranvier, preventing the premature differentiation of sensory organs, fasciculation of the nerve, preventing oligodendrocytes from improperly

2 exiting the spinal cord, and repositioning peripheral nerves (Gilmour et al., 2002;

Grant et al., 2005; Kucenas et al., 2009; Raphael et al., 2010; Voas et al., 2009).

While much is known about the roles of Schwann cells, many questions remain unanswered, including how Schwann cells regulate nerve development and what signals control Schwann cell development and myelination. We chose to study

Schwann cell development in zebrafish for many reasons. Zebrafish are very tractable model organisms that rapidly develop, are fertilized externally, and are born in large clutches making them an excellent organism for genetic screens and studies of early development, including the development of the nervous system. In Chapter 2 of this thesis, I explore a new role for Schwann cells in correctly positioning a peripheral nerve. In Chapter 3, I explore the roles of ErbB signaling and Schwann cell proliferation during a specific stage of Schwann cell development, radial sorting.

THE POSTERIOR LATERAL LINE NERVE

We have chosen to study the role of Schwann cells in peripheral nerve development in the zebrafish posterior lateral line nerve (PLLn). This nerve has two main advantages: it is present and functional early in development and it is located superficially, allowing for easy visualization (Metcalfe, 1985; Metcalfe et al., 1985). Lateral line nerves are present along the head and trunk of fish and amphibians, and much of the early research on this nerve was conducted in frogs or newts (Ghysen and Dambly-

Chaudiere, 2007; Stone, 1922; Stone, 1935; Winklbauer, 1989). Lateral line nerves innervate sensory organs called neuromasts that allow organisms to sense changes in

3 water current. The lateral line system is important for a myriad of behaviors, including predator avoidance, prey localization, orientation of the fish in a water current, schooling, and sexual courtship (Ghysen and Dambly-Chaudiere, 2007). This sensory organ is often referred to as “sense at a distance.”

The posterior lateral line nerve in zebrafish begins as a very simple nerve that runs straight down either side of the trunk of the animal (Ghysen and Dambly-Chaudiere,

2007; Metcalfe, 1985; Metcalfe et al., 1985). It is derived from an ectodermal placode that is located posterior to the ear (Ghysen and Dambly-Chaudiere, 2007; Schlosser,

2002). The posterior portion of the placode will give rise to a migrating primordium that will migrate the length of the embryo, depositing clusters of pre-neuromasts cells and interneuromast cells along the way (Fig. 1.1; Metcalfe et al., 1985; Schlosser,

2002). The anterior portion will differentiate into the ganglion of the lateral line nerve, which contains all of the neurons that will innervate the posterior lateral line neuromasts (Schlosser, 2002). The axons from the posterior lateral line neurons will co-migrate with the migrating primordium (Gilmour et al., 2004). This is a unique form of axon outgrowth, such that the axon growth cones are buried within their migrating target. Schwann cells will coalesce with the ganglion and then migrate behind the outgrowing axons (Gilmour et al., 2002). By 48 hours post fertilization

(hpf), the primordium has reached the tip of the tail and divided into two to three more neuromasts (Ghysen and Dambly-Chaudiere, 2007). The axons and Schwann cells have also migrated to the tip of the tail, so the entire system is present and active by 48 hpf. Chapter 2 of this thesis explores how the lateral line nerve initially grows out

4 within the epidermis but then rapidly transitions across the epidermal basement membrane into the subepidermal space.

As time goes on, the primary neuromasts, those that were directly deposited by the migrating primordium, begin to migrate ventrally (Grant et al., 2005). However, the posterior lateral line nerve remains firmly anchored in its original position (Grant et al., 2005; Raphael et al., 2010). Axon branches follow the migrating neuromasts to maintain their innervation. Simultaneously, the interneuromast cells, which are chains of primordium-derived cells that link primary neuromasts, also begin to migrate ventrally (Grant et al., 2005). These cells begin to divide and produce new clusters of neuromast cells, giving rise to the secondary neuromasts. Axons will break away from the main body of the lateral line nerve and migrate towards these secondary neuromasts to innervate them (A. R. Raphael and W. S. Talbot, unpublished observation). Finally, neuromasts can bud to form “stitches” of neuromasts – rows of neuromasts running perpendicular to the lateral line nerve (Ghysen and Dambly-

Chaudiere, 2007). Thus, the complexity of the posterior lateral line system grows throughout the life of the fish. In Chapter 2, I propose a mechanism for how the initial patterning of the posterior lateral line nerve is protected from the extensive changes in size and complexity of the posterior lateral line system over time.

5 MYELINATION IN THE PERIPHERAL NERVOUS SYSTEM

Schwann Cell Development

Before a Schwann cell can make myelin, it must go through several developmental steps (Jessen and Mirsky, 2005). Schwann cells are derived from neural crest, which delaminate from the neural tube and migrate out to associate with neurons where they will differentiate into Schwann cell precursors (Jessen et al., 1994; Sauka-Spengler and Bronner, 2010). Schwann cell precursors comigrate with axons as they grow towards their targets (Dong et al., 1999). Once migration is complete, Schwann cell precursors will differentiate into immature Schwann cells (Jessen and Mirsky, 2005).

At this point, the immature Schwann cells are faced with a fate choice – whether to become a myelinating or non-myelinating Schwann cell. If a Schwann cell is ultimately associated with one large caliber axon, it will become a pro-myelinating

Schwann cell, initiate the myelination transcription pathway, and begin to wrap its cytoplasm around the axon to create a myelin sheath (Jessen and Mirsky, 2005;

Webster et al., 1973). In contrast, if a Schwann cell is interacting with several small caliber axons it will become a non-myelinating Schwann cell and ensheath each axon in a pocket of its cytoplasm, forming a Remak bundle (Aguayo and Bray, 1975;

Aguayo et al., 1976; Hahn et al., 1987; Jessen and Mirsky, 2005; Peters and Muir,

1959).

Throughout their development, Schwann cells undergo many signaling and physical changes (Jessen and Mirsky, 2005; Martin and Webster, 1973; Svaren and Meijer,

2008; Webster et al., 1973). Genes are up- and down-regulated to move the Schwann

6 cells through the developmental progression (Jessen and Mirsky, 2005; Svaren and

Meijer, 2008). In addition, a significant physical transformation must occur in order for a Schwann cell to make myelin (Webster et al., 1973). Schwann cell precursors and immature Schwann cells associate with many axons (Jessen and Mirsky, 2005;

Webster et al., 1973). In contrast, a myelinating Schwann cell only interacts with one segment of one axon (Webster, 1971). The process by which Schwann cells transition from interacting with many axons to just one is called radial sorting (Webster et al.,

1973).

Radial Sorting

The advent of transmission electron microscopy in the 1930s led to a revolution in the field of neurobiology, allowing scientists to explore the fine structure of the nervous system. Using this new technology, Geren discovered that in the peripheral nervous system the myelin sheath was made by Schwann cells, and not by the axons themselves (Geren and Raskind, 1953). More electron microscopy studies in the late

1950s found that Schwann cells are initially arrayed around axon bundles but then somehow transition to myelinating Schwann cells associated with one axon or non- myelinating Schwann cells that bundle axons into Remak bundles (Peters and Muir,

1959). But how do Schwann cells make this transition? The answer came from a series of elegant electron microscopy studies of developing nerves in rat performed by

Webster in 1973 (Webster et al., 1973). Using a time course of rat sciatic nerves,

Webster described a process wherein “axons to be myelinated appeared to progress radially from a bundle to a 1:1 relationship with a Schwann cell at the sheath’s outer

7 margin” (Webster et al., 1973). Thus the term “radial sorting” was born. Basically,

Schwann cells extend processes into the axon bundle, interacting with several axons, but ultimately choose one and sort it to the periphery of the axon bundle (Fig. 1.2).

These Schwann cell processes are actually long lamellar sheets that interact with axons along their lengths (Wanner et al., 2006; Webster et al., 1973).

Webster made several other interesting observations in his 1973 paper (Webster et al.,

1973). First, during the time when radial sorting is occurring, new axons are growing into the nerve. Additionally, the number of Schwann cells within the nerve also increases significantly during this time. While the nerve is initially one giant bundle of axons, this is subdivided over time into Schwann cell families, where Schwann cells surround small clusters of axons. Webster found that over time the number of these

Schwann cell families increased, but the number of axons within them decreased, suggesting that Schwann cells were subdividing the axons of the nerve into smaller and smaller bundles. Finally, Webster also observed that within a bundle of axons, those axons that were surrounded by other axons and not in contact with a Schwann cell were smaller than those that were contacting the surface of a Schwann cell and that axons segregated by Schwann cells were the largest. This suggested that the axons might be receiving trophic factors from Schwann cells, an observation later proven to be true (Jessen and Mirsky, 2005).

8 Signals controlling radial sorting

The first mutant with a radial sorting defect was published the same year as Webster’s paper. In dystrophic mice, which are a model of merosin-deficient congential muscular dystrophy, there are large bundles of unsorted axons that are only minimally invested by Schwann cell processes (Bradley and Jenkison, 1973). The dystrophic mice turned out to have a lesion in laminin-α2 (Sunada et al., 1994; Xu et al., 1994), giving the first suggestion that the basal lamina may be important for regulating radial sorting. In the past ten years, several more genes that control radial sorting have emerged, and many of them are tied to the basal lamina (Benninger et al., 2007; Chen and Strickland, 2003; Feltri et al., 2002; Grove et al., 2007; Nodari et al., 2007; Pereira et al., 2009; Yang et al., 2005; Yu et al., 2009; Yu et al., 2005). These genes seem to fall into two main classes: those that control Schwann cell proliferation and those that regulate Schwann cell process extension (Benninger et al., 2007; Feltri et al., 2002;

Nodari et al., 2007; Pereira et al., 2009; Yang et al., 2005). Interestingly, basal lamina proteins and receptors play roles in both (Chen and Strickland, 2003; Feltri et al.,

2002; Nodari et al., 2007; Yang et al., 2005; Yu et al., 2009; Yu et al., 2005).

It makes intuitive sense that Schwann cell proliferation would be important for radial sorting. Initially, there are many more axons than Schwann cells to myelinate them, and the number of Schwann cells must be carefully matched to the number of axon segments that will be myelinated (Jessen and Mirsky, 2005). This occurs not through the influx of new Schwann cells into the nerve, but by the proliferation of those

Schwann cells already present within the nerve (Jessen and Mirsky, 2005; Martin and

9 Webster, 1973). As such, there is a significant increase in Schwann cell proliferation during radial sorting (Jessen and Mirsky, 2005; Lyons et al., 2005; Martin and

Webster, 1973; Webster et al., 1973). Several genes that are essential for radial sorting have defects in Schwann cell proliferation (Benninger et al., 2007; Chen and

Strickland, 2003; Grove et al., 2007; Wallquist et al., 2005; Yang et al., 2005; Yu et al., 2009; Yu et al., 2005). In the absence of these genes, Schwann cell processes are still extended into the axon bundle, but radial sorting is arrested nonetheless. These genes include cdc42, focal adhesion kinase (FAK), laminin-2 (the laminin affected in the dystrophic mouse), laminin-8, and laminin-γ1 (Benninger et al., 2007; Chen and

Strickland, 2003; Grove et al., 2007; Wallquist et al., 2005; Yang et al., 2005; Yu et al., 2009; Yu et al., 2005).

It also makes sense that Schwann cells would need to coordinate process extension and stabilization for radial sorting. They must be able to extend processes into the axon bundle, interact with multiple axons, but ultimately wrap only one (Webster et al., 1973). Genes that are required for process extension during radial sorting include rac1, which is regulated downstream of β1-integrin, integrin linked kinase (ILK), and laminin-γ1 (Benninger et al., 2007; Feltri et al., 2002; Nodari et al., 2007; Pereira et al., 2009; Yu et al., 2005). In all of these cases, mutation of the gene in Schwann cells results in the inability of Schwann cells to insert processes into the axon bundle, and thus radial sorting is defective. Rac1 plays well-known roles in regulating the actin cytoskeleton and in vitro work has shown the actin cytoskeleton is involved in axon ensheathment (Etienne-Manneville and Hall, 2002; Fernandez-Valle et al., 1997).

10 Using Schwann cell/neuron co-cultures Fernandez-Valle et al., found that at high levels of an actin depolymerizing drug, cytochalasin D, Schwann cells are unable to interact with axons (Fernandez-Valle et al., 1997). At lower levels of cytochalasin D,

Schwann cells can elongate, interact with axons, and extend processes into the axon bundle but they are unable to segregate axons into 1:1 relationships.

Differentiated Schwann cells

Promyelinating Schwann cells

Once an axon has been radially sorted and is in a 1:1 relationship with a Schwann cell, the Schwann cell associated with the axon transitions from an immature Schwann cell to a promyelinating Schwann cell (Jessen and Mirsky, 2005). As a promyelinating

Schwann cell, the cell begins to express transcription factors, including Oct6 and

Krox20, that regulate the expression of myelin specific proteins (Arroyo et al., 1998;

Ghislain and Charnay, 2006; Monuki et al., 1989; Svaren and Meijer, 2008; Topilko et al., 1994). Oct6, Brn2, Krox20, and the recently discovered Gpr126 are required for

Schwann cells to transition from the promyelinating stage to the myelinating stage

(Bermingham et al., 1996; Jaegle et al., 2003; Jaegle et al., 1996; Jessen and Mirsky,

2005; Monk et al., 2009; Svaren and Meijer, 2008; Topilko et al., 1994). As such, mutations in these genes result in Schwann cells that are arrested with one and a half wraps of Schwann cell cytoplasm around an axon (Bermingham et al., 1996; Jaegle et al., 2003; Monk et al., 2009; Topilko et al., 1994).

11 Myelinating Schwann cells

The amount of myelin that is made around an axon is correlated to the diameter of the axon, such that large axons will have thicker myelin sheaths than smaller axons

(Donaldson and Hoke, 1905). A major signal that controls how much myelin a

Schwann cell will make is Neuregulin 1 type III (Nrg1 type III; Michailov et al., 2004;

Taveggia et al., 2005). Larger axons express higher levels of Nrg1 type III, which leads to a thicker myelin sheath. In contrast, smaller axons have lower levels of expression and therefore a thinner sheath. Driving higher levels of Nrg1 type III expression in smaller caliber axons is sufficient to cause them to be myelinated

(Taveggia et al., 2005). Even lower levels may lead to Remak bundle formation

(Nave and Salzer, 2006; Taveggia et al., 2005).

Non-myelinating Schwann cells

At the end of radial sorting, all of the large caliber axons are in a 1:1 relationship with a promyelinating or myelinating Schwann cell (Webster, 1971; Webster et al., 1973).

The remaining small caliber axons are bundled by non-myelinating Schwann cells into

Remak bundles (Hahn et al., 1987; Jessen and Mirsky, 2005; Peters and Muir, 1959).

Each small caliber axon will be ensheathed by a Schwann cell process, such that the many axons interacting with a non-myelinating Schwann cell in a Remak bundles are separated from other axons in the bundle (Aguayo and Bray, 1975; Aguayo et al.,

1976; Peters and Muir, 1959). Disruption of Remak bundles leads to defects in the axons that they ensheath (Chen et al., 2003). Nrg1 type III also plays a role in Remak

12 bundle formation; in Nrg1 type III +/- animals, Remak bundles are not properly invested by Schwann cell processes (Taveggia et al., 2005).

Transition from an immature Schwann cell to a promyelinating and then myelinating

Schwann cell or non-myelinating Schwann cell is accompanied by cell cycle exit

(Jessen and Mirsky, 2005). This is the only stage in the Schwann cell developmental program that is considered to be a terminal differentiation. However, in cases of injury or disease, Schwann cells are capable of dedifferentiating to the immature

Schwann cell state and reentering the cell cycle (Romine et al., 1976).

Neuregulin/ErbB Signaling

Neuregulin signaling from axons through their ErbB receptors on Schwann cells is turning out to be key in regulating several facets of Schwann cell development, including migration, proliferation, survival, myelination, and the formation of Remak bundles (Birchmeier and Nave, 2008; Dong et al., 1995; Garratt et al., 2000; Lyons et al., 2005; Michailov et al., 2004; Morrissey et al., 1995; Riethmacher et al., 1997;

Taveggia et al., 2005; Woldeyesus et al., 1999). Additionally, they play a role in radial sorting (Taveggia et al., 2005), which we further explore in Chapter 3. There are over fifteen neuregulin isoforms, although Nrg1 type III seems to be the main isoform active during Schwann cell development (Falls, 2003). All of the neuregulin

1 isoforms begin as transmembrane proteins that are then cleaved; Nrg1 type I and II are released as paracrine signals. In contrast, Nrg1 type III has a second

13 transmembrane domain, such that when the first transmembrane domain is cleaved, it remains anchored the plasma to the membrane as a juxtacrine signal.

There are four receptors in the ErbB tyrosine kinase receptor family, EGFR, ErbB2,

ErbB3, and ErbB4 (Citri et al., 2003). ErbB2 and ErbB3 are the receptors primarily expressed in Schwann cells (Birchmeier, 2009; Lyons et al., 2005). ErbB2 cannot bind to the Nrg ligand, but it does have kinase activity (Citri et al., 2003). In contrast,

ErbB3 can bind to the Nrg ligand, but lacks kinase activity. Therefore, only when the two receptor heterodimerize in the presence of the Nrg ligand can they autophosphorylate and induce downstream signaling.

How does one signal regulate so many disparate steps of Schwann cell development?

Perhaps the way in which the ligand is present results in distinct Schwann cell behaviors. For example, during radial sorting and Remak bundle formation, Schwann cells are presented with several sources of the Nrg ligand, originating from multiple axons (Taveggia et al., 2005). In contrast, during myelination, Schwann cells are receiving a single, high-level source of Nrg signaling from one axon (Michailov et al.,

2004; Taveggia et al., 2005). Recently, in vitro co-culture work revealed that there is a concentration dependent effect of adding ectopic Nrg to a myelinating co-culture, such that high levels inhibit myelination and low levels enhance it (Syed et al., 2010).

Additionally, there may be distinct intracellular responses to Nrg signaling downstream of the ErbB receptors. There are several molecules that are known to

14 signal downstream of Nrg, including PI3K/Akt, MEK/ERK, and Cdc42, and these may modulate the response of a Schwann cell to the same signal (Benninger et al.,

2007; Ogata et al., 2004; Syed et al., 2010). For example, during radial sorting Cdc42 is likely acting to regulate Schwann cell proliferation downstream of Nrg/ErbB signaling while Rac1 may be acting to regulating the cytoskeleton to coordinate process extension downstream of the same signal (Benninger et al., 2007). We explore the role of ErbB signaling during radial sorting in Chapter 3.

SUMMARY OF RESEARCH AND FUTURE DIRECTIONS

In Chapter 2 of this thesis, I describe a new role for Schwann cells in correctly positioning the posterior lateral line nerve. While the PLLn initially grows out within the epidermis, it rapidly transitions across the basement membrane into the subepidermal space. Schwann cells are required for this process, and in their absence significant disorganization of the nerve occurs. What genes control the movement of the PLLn across the basement membrane remain unknown, but are likely to include molecules that both degrade and rebuild a basement membrane. We hypothesize that moving the PLLn below the epidermal basement membrane protects it from the significant reorganization of its targets (the neuromasts) and continual growth of the fish. Many other sensory epithelia are organized such that the main bulk of innervating axons are separated from their targets by a basement membrane – an open question is whether this organizing principle serves a protective function in these other instances (Boulais and Misery, 2008; Fernandez et al., 1990; Nedelec et al., 2005;

Northcutt, 2004; Oakley and Witt, 2004; Purcell and Perachio, 1997; Si et al., 2003;

15 Winklbauer, 1989). Additionally, whether Schwann cells are responsible for repositioning peripheral nerves across a basement membrane in other systems remains unknown.

Chapter 3 of this thesis explores the role of ErbB signaling and Schwann cell proliferation during radial sorting. I find that while both ErbB signaling and Schwann cell proliferation are required for radial sorting, ErbB signaling plays a direct role in regulating Schwann cell process extension while Schwann cell proliferation does not.

What signals coordinate ErbB function during radial sorting remain unknown, but are likely to include Rac1 and Cdc42 (Benninger et al., 2007). I also wonder if Schwann cell proliferation is required during radial sorting merely to provide adequate numbers of Schwann cells or if a special terminal or polarized division is required before a

Schwann cell can commit to myelination and become terminally differentiated.

Finally, Nrg/ErbB signals control many aspects of Schwann cell development, and how this one signal is developmentally regulated and coordinated remains a mystery.

16 Figure 1.1: Posterior Lateral Line Development

17 Figure 1.1: Posterior Lateral Line Development

(A) An ectodermal placode that forms posterior to the ear gives rise to a migrating primordium (blue) and the ganglion of the posterior lateral line nerve (red, PLL nerve). Schwann cells (green) coalesce with the ganglion. (B) The primordium migrates along the horizontal myoseptum towards the tail. Axon growth cones (red) are buried within the primordium and axons trail behind it. Schwann cells migrate behind the outgrowing axons. (C) The primordium buds off groups of cells, (D) which are deposited as clusters of pre-neuromasts cells (blue with asterisk) that will give rise to the neuromasts. (E) At the completion of primordium migration to the tip of the tail, several neuromasts have been deposited and the PLL axons stretch all the way to the tail with Schwann cells associated with them. Cartoons are superimposed on a 28 hours postfertilization (hpf) embryo. Primordium migration begins at ~20 hpf and is complete by ~40 hpf (Kimmel et al., 1995).

18 Figure 1.2

19 Figure 1.2: Radial Sorting

(A) At the beginning of radial sorting, Schwann cells (yellow) are arrayed around the outside of the axon bundle (blue). Several genes have been found that are required for

Schwann cell process extension, and when they are mutated radial sorting arrests at this early stage. (B) Schwann cells insert their processes into the axon bundle, interacting with several axons. Several genes have been found that regulate Schwann cell proliferation during radial sorting; when these genes are mutated, radial sorting arrests with Schwann cell processes within the axon bundle. (C) The Schwann cell will pick one axon and withdraw all of its other processes. (D) Once a Schwann cell sorts axon to the periphery of the axon bundle it enters the promyelinating stage.

When genes required for the transition from promyelinating to myelinating Schwann cells are mutated, the Schwann cell arrests with one and a half wraps around the sorted axon.

20 Chapter 2

Schwann cells reposition a peripheral nerve to isolate it from

postembryonic remodeling of its targets

Alya R. Raphael, Julie R. Perlin, William S. Talbot

This chapter was originally published in Development (2010) 137(21): 3643-9.

Author contributions: A.R.R. performed all experiments with exception of the transplants, which were carried out by J.R.P. A.R.R. and W.S.T. designed the experiments and wrote the paper.

21 SUMMARY

While much is known about the initial construction of the peripheral nervous system

(PNS), less understood are the processes that maintain the position and connections of nerves during postembryonic growth. Here we show that the posterior lateral line nerve in zebrafish initially grows in the epidermis and then rapidly transitions across the epidermal basement membrane into the subepidermal space. Our experiments indicate that Schwann cells, which myelinate axons in the PNS, are required to reposition the nerve. In mutants lacking Schwann cells, the nerve is mislocalized and the axons remain in the epidermis. Transplanting wildtype Schwann cells into these mutants rescues the position of the nerve. Analysis of chimeric embryos suggests that the process of nerve relocalization involves two discrete steps – the degradation and recreation of the epidermal basement membrane. Although the outgrowth of axons is normal in mutants lacking Schwann cells, the nerve becomes severely disorganized at later stages. In wildtype embryos, exclusion of the nerve from the epidermis isolates axons from migration of their targets (sensory neuromasts) within the epidermis.

Without Schwann cells, axons remain within the epidermis and are dragged along with the migrating neuromasts. Our analysis of the posterior lateral line system defines a new process in which Schwann cells relocate a nerve beneath the epidermal basement membrane to insulate axons from the postembryonic remodeling of their targets.

22 INTRODUCTION

Many studies have investigated the mechanisms of axonal pathfinding that connect neurons to their targets in the developing embryo (Bashaw and Klein, 2010). Axonal growth cones migrate in response to attractive and repulsive cues, such as netrins, semaphorins, ephrins, and slits, to reach targets that are often significant distances from the neuronal cell body (Bashaw and Klein, 2010). Once the initial architecture of the nerve is established, it must be able to respond to changes that occur during postembryonic development, including extensive growth of the organism and changes in tissue morphology. Little is known about growth and remodeling of nerves and their targets after the initial outgrowth of axons. In addition, many cells types are present in mature peripheral nerves, including endothelial cells, fibroblasts, cells of the immune system, and Schwann cells (Gamble and Goldby, 1961; Jessen and Mirsky,

2005), and the roles of these non-neuronal cells that associate with axons in postembryonic remodeling are unclear.

Schwann cells are a significant component of peripheral nerves and play well established roles in ensheathing and myelinating axons (Jessen and Mirsky, 2005).

Additionally, Schwann cells provide trophic support for axons and present cues that cluster axonal sodium channels at the node of Ranvier (Jessen and Mirsky, 2005;

Poliak and Peles, 2003). Recently, new roles for Schwann cells have emerged, including elimination of ectopic sodium channels from the internode (Voas et al.,

2009), preventing the premature differentiation of sensory organs (Grant et al., 2005),

23 fasciculating the nerve (Gilmour et al., 2002), and preventing oligodendrocytes from exiting the spinal cord (Kucenas et al., 2009).

The posterior lateral line nerve (PLLn) is a prominent peripheral nerve in zebrafish that innervates sensory organs called neuromasts, which detect changes in water currents (Ghysen and Dambly-Chaudiere, 2007). The components of the posterior lateral line arise from an ectodermal placode, the posterior portion of which forms a primordium that migrates toward the posterior end of the embryo and deposits clusters of cells that will give rise to the neuromasts. The anterior half of the placode becomes neurons in the ganglion of the PLLn, which will innervate the neuromasts of the lateral line. An interesting variation of axonal pathfinding occurs in the PLLn, where the precursors of the targets (i.e. the primordium) and the associated axons grow out together (Gilmour et al., 2004). While Schwann cells are intimately associated with the axons from an early stage, they do not appear to play a role in axonal pathfinding

(Gilmour et al., 2002).

The posterior lateral line system (nerve and neuromasts) initially has a simple structure with only seven to eight neuromasts distributed in a line down the midbody of the larvae, but it rapidly becomes more complicated, containing hundreds of neuromasts widely distributed over the body of the adult (Ghysen and Dambly-

Chaudiere, 2007). Additionally, the fish grows considerably in size after the initial formation of the lateral line, so the PLLn must be able to compensate both for the increased size and complexity of the system. Finally, while the primordium migrates

24 within the epidermis (Metcalfe, 1985), which is the final position of the neuromasts, the mature nerve is embedded within the subepidermal space at the horizontal myoseptum, directly below the basement membrane of the epidermis (Voas et al.,

2009; Winklbauer, 1989). How these complex changes in tissue architecture occur is not well understood, including how the PLLn grows within the epidermis but is ultimately excluded from it.

To learn more about the initial formation and remodeling of the lateral line, we have investigated the development of the lateral line system in embryonic and early larval stages. Ultrastructural studies reveal that the entire nerve, including axons and

Schwann cells, is initially localized within the epidermis, superficial to the epidermal basement membrane. Shortly after axon outgrowth, the epidermal basement membrane is degraded and then reformed on the opposite side of the nerve, so that the nerve is repositioned to its mature location in the subepidermal space. The analysis of mutant and chimeric embryos shows that Schwann cells are required for this process of basement membrane degradation and regeneration. In mutants lacking Schwann cells, nerves become progressively disorganized as development proceeds, suggesting that the process of basement membrane invasion is essential to maintain the functional integrity of the nerve during postembryonic remodeling. These results define a new role for Schwann cells in remodeling tissues in the vicinity of nerves to ensure proper organization during postembryonic growth.

25 MATERIALS AND METHODS

Fish strains

Zebrafish embryos were raised at 28.5°C and were staged as described (Kimmel et al.,

1995). The erbb2st61, erbb3st48, and sox10/clst3 mutants and FoxD3:GFP(17) transgenic fish have been previously described (Gilmour et al., 2002; Kelsh and Eisen,

2000; Lyons et al., 2005; Pogoda et al., 2006).

Genotyping

The erbb2st61 and erbb3st48 mutations were genotyped as described (Lyons et al.,

2005). sox10/clst3 embryos were scored for lack of pigment and then genotyped for the presence of the t3 insertion using the following primers forward: 5’

TGAAGTCCGACGAGGAAGAT 3’ and reverse: 5’ CACAGCTTCCCCAGTGTTTT

3’, such that a fragment is not amplified in the mutants.

Transmission electron microscopy

Sample preparation for electron microscopy was performed as described in (Lyons et al., 2008) for 28 hours postfertilization (hpf), 3 days postfertilization (dpf), and 5 dpf embryos; a minimum of three nerves from three separate embryos were examined for each condition. Images were collected on a Jeol TEM1230 and pseudocolored using

Adobe Photoshop software.

Immunohistochemistry and live imaging

26 Antibody staining for acetylated tubulin was performed following standard methods as described in (Lyons et al., 2005). FoxD3:GFP(17) wildtype embryos were soaked in

1-phenyl-2-thiourea at a final concentration of 0.2 mM following gastrulation to prevent melanization of the pigment. For live imaging, embryos were anesthetized with 0.016% Tricaine (w/v) and were mounted in 1.5% low melting agarose.

Fluorescent images were collected on a Zeiss Pascal LSM5 confocal microscope.

Generation of genetic chimeras

Wildtype embryos carrying the FoxD3:GFP transgene were injected with 1% Texas red dextran. Labeled cells were transplanted at the blastula stage into sox10/clst3 mutant hosts. Embryos were scored live at 3 dpf and 5 dpf for GFP-positive Schwann cells along the PLLn and were imaged live as described above. Embryos were then removed from the agarose and processed for transmission electron microscopy (TEM).

27 RESULTS

The PLLn invades the subepidermal space

At 3 days postfertilization (dpf), the zebrafish PLLn is normally localized just beneath the basement membrane that separates the epidermis from the muscle (Voas et al.,

2009). To determine the localization of the PLLn during its initial outgrowth, we performed transmission electron microscopy. At 28 hours postfertilization (hpf), the axons and Schwann cells in the posterior segment of the nerve, which are close to the migrating primordium, are located within the epidermis (Fig. 2.1A,C). At more anterior locations farther from the primordium, the nerve can be found in a transitional state, with basement membrane both above and below the nerve (Fig. 2.1A,D). At later developmental stages, the nerve is located completely below the basement membrane of the epidermis (Fig. 2.1E). Thus, the PLLn transitions across a basement membrane from the epidermis into the subepidermal space during its development

(Fig. 2.1B). Additionally, this transition occurs in an anterior to posterior fashion, with more anterior segments of the nerve moving across the basement membrane prior to more posterior segments (Fig. 2.1A,B).

The PLLn does not cross the basement membrane in Schwann cell deficient mutants

Sox10 is essential for the development of Schwann cells and other neural crest derivatives (Kelsh et al., 2000; Kelsh and Eisen, 2000). In Schwann cells, ErbB2 and

ErbB3 form a heteromeric receptor that is required for glial proliferation, migration, and differentiation (Lyons et al., 2005; Monk and Talbot, 2009; Nave and Salzer,

28 2006). We have previously reported that in mutants lacking Schwann cells, including erbb2st61, erbb3st48 and sox10 (clst3) mutants, the PLLn is aberrantly localized in the epidermis at 3 dpf (Voas et al., 2009). To learn more about the development of the

PLLn in mutants lacking Schwann cells, we examined earlier and later stages. At 28 hpf, the PLLn axons are in the epidermis of erbb2st61 mutants (Fig. 2.2B), just as with wildtype siblings (Fig. 2.2A), and both can be found associated with pre-neuromast cells, indicating that initial outgrowth of the nerve is normal, even in the absence of

Schwann cells. At 5 dpf the wildtype nerve is properly located beneath the epidermal basement membrane (Fig. 2.3A), while the nerves of erbb2st61, sox10/clst3 and erbb3st48 mutants remain in the epidermis (Fig. 2.3B,C and data not shown). These results indicate that there is a complete failure of basement membrane invasion in the absence of Schwann cells. Finally, defasciculation of the nerve is evident in the mutants, as individual axons and small groups of axons are frequently observed separate from the main axon bundle (Fig 2.3C and data not shown). These results suggest that Schwann cells are essential for the nerve to cross the basement membrane, reach its mature position, and remain fasciculated in the larva.

Schwann cells are required for the PLLn to relocalize across the basement membrane

To further investigate whether Schwann cells are required to shift the PLLn from the epidermis into the subepidermal space, we performed transplantation experiments to generate genetic chimeras in which Schwann cell deficient mutants contained wildtype cells, including some Schwann cells. Cells from wildtype embryos carrying the

29 FoxD3:GFP(17) transgene, which expresses green fluorescent protein (GFP) in

Schwann cells and other neural crest derivatives (Gilmour et al., 2002), were transplanted into sox10/clst3 mutant hosts, which lack all Schwann cells in the PLLn.

Chimeric embryos were screened for the presence of GFP-expressing (wildtype)

Schwann cells in the PLLn and were then processed for electron microscopy to determine the localization of the nerve.

We analyzed 12 chimeras in which sox10/clst3 mutant embryos had wildtype Schwann cells covering the entire length of the PLLn (Fig. 2.4B). In these 12 chimeras, there were eight cases of complete rescue of the nerve position (Fig. 2.4C), such that the chimeric nerve occupied the mature wildtype position beneath the basement membrane. There were also four cases of partial rescue, three of which had nerves in the proper location beneath the basement membrane, but with some residual basement membrane remaining on the deep (i.e., muscle) side of the nerve (data not show, similar to Fig. 2.5C). In the fourth case of partial rescue, basement membrane surrounded the nerve on both sides, suggesting that the transplanted Schwann cells were unable to degrade the original basement membrane underneath the nerve, but were nonetheless able to initiate the formation of a new, superficial membrane (Fig.

2.4D). These results indicate that Schwann cells are required to correctly position the

PLLn within the subepidermal space.

We also analyzed four other chimeras in which sox10/clst3 mutant embryos had smaller clones of wildtype Schwann cells that covered only the anterior segment of the

PLLn (Fig. 2.5B). In the anterior segments of the PLLn, all of these had partial rescue of the nerve position, either with some residual deep basement membrane (Fig. 2.5C,

30 n=1), equal basement membrane on both sides (data not shown, similar to Fig. 2.4D, n=2), or location in the epidermis but surrounded by a circumferential basement membrane (Fig. 2.5E,F, n=1). The partial rescue of nerve position suggests that

Schwann cells were not present in sufficient numbers or at the correct stage to completely degrade the original basement membrane in these chimeras. We did not observe any evidence of rescue in the posterior segments of these nerves, in which axons were not associated with Schwann cells (Fig. 2.5D), indicating that Schwann cells can only rescue the portion of the nerve with which they are associated. Finally, in all 16 chimeras with full or partial Schwann cell coverage, myelination of the PLLn was rescued, indicating that sox10 acts cell autonomously in Schwann cells to regulate myelination (insets in Figs 2.4-2.5).

Movement across the epidermal basement membrane protects the integrity of the

PLL nerve

The foregoing results raise questions about the possible functions of nerve relocalization. It has previously been reported that by 3 dpf the neuromasts of the

PLLn begin to migrate ventrally within the epidermis (Grant et al., 2005). Because the

PLLn remains in the epidermis with the neuromasts in mutants lacking Schwann cells, we hypothesized that ventral migration of the neuromasts might drag the mislocalized lateral line nerve and cause the disorganization of the nerve that is observed in these mutants (Gilmour et al., 2002; Lyons et al., 2005; Pogoda et al., 2006) (Fig. 2.6). To test this hypothesis, we studied a time course of wildtype and Schwann cell mutant

(erbb2st61 and sox10/clst3; erbb3st48 data not shown) nerves (Fig. 2.6). Consistent with previous studies (Gilmour et al., 2002), wildtype and mutants nerves are

31 indistinguishable from each other at 48 hpf, a stage before the neuromasts begin to migrate (Fig. 2.6A,F,K). Beginning at 3 dpf, however, as neuromasts are migrating ventrally (Grant et al., 2005) (Fig. 2.6B), the mutant nerves began to undulate and defasciculate (Fig. 2.6G,L); this phenotype worsened over time (Fig. 2.6G-J, L-O). In the mutants the nerves were always the most severely disorganized in the anterior segments, where neuromasts first begin to migrate ventrally (Fig. 2.6). Additionally, while the wildtype neuromasts migrate ventrally away from the PLLn, which remains anchored at the horizontal myoseptum, mutant neuromasts are often found close to the disorganized nerve (arrowheads, Fig. 2.6). Finally, when mutant nerves at 5 dpf were studied by TEM, they often remained closely associated with neuromasts (10/19 mutant nerves examined, data not shown, Fig. 2.3B,C), whereas wildtype nerves were well separated from the neuromasts by this stage (Fig. 2.6 and data not shown). These results support the possibility that moving the PLLn from the epidermis into the subepidermal space is critical to protect the nerve from being pulled ventrally along with the migrating neuromasts in the larva (Fig. 2.6A-E).

32 DISCUSSION

As development progresses, organs that are initially built in a small embryo must grow to the adult size. The organism must be able to adapt to these changes in size while maintaining functionality of the organs. This is certainly true of the nervous system, where connections are initially established in the embryo and then maintained during extensive growth of the organism. Here we present a process by which the posterior lateral line nerve in zebrafish initially forms in the epidermis and then rapidly transitions out of the epidermis, below the epidermal basement membrane. Our evidence suggests that repositioning the nerve protects it from the remodeling of its targets in the epidermis and maintains its integrity during the continued growth of the fish.

Our analysis defines a new role for Schwann cells in breaching and reforming the epidermal basement membrane to properly position the lateral line nerve. We find that the wildtype PLLn, including Schwann cells and axons, initially grows within the epidermis but then crosses the epidermal basement membrane to invade the subepidermal space (Fig. 2.1). This transition occurs early in development, while the distal region of the nerve is still growing. Additionally, in mutants lacking Schwann cells, including erbb2st61, erbb3st48 and sox10/clst3, the nerve is aberrantly located within the epidermis, and this results in significant disorganization of the nerve (Fig.

2.3, 2.6). The phenotypic similarity of these mutants suggests that the abnormal nerve localization results from the loss of Schwann cells, rather than a specific function of any of these genes in the process of basement membrane remodeling itself.

33 Transplantation of wildtype Schwann cells into sox10/clst3 mutants restores the nerve to its normal position beneath the epidermal basement membrane (Fig. 2.4, 2.5).

Therefore, Schwann cells are required for the local degradation and immediate reformation of the basement membrane on the opposite side of the nerve.

Basement membrane invasion defines a mode of axon localization that is distinct from axonal growth cone pathfinding. In the PLLn, the axons grow out along with the primordia of their final targets (Gilmour et al., 2004), the neuromasts, in the epidermis and then are moved along their entire length by the Schwann cells associated with the nerve. In addition, the segments of the axon farthest from the growth cone are the first to be moved across the basement membrane. It appears that interactions between

Schwann cells and their local environment regulate the transition of the PLLn across the epidermal basement membrane. This most likely occurs through short-range or contact-dependent signaling, because Schwann cells are only competent to move segments of the nerve with which they are associated (Fig. 2.5C,D). In addition, it appears that this mode of nerve localization is distinct from the delamination of newborn neurons from ectodermal placodes to form sensory ganglia, including the

PLLn ganglia (Schlosser, 2002). In ganglion formation, neurons delaminate from the ectoderm and migrate into the mesenchyme to coalesce with neural crest cells (Baker and Bronner-Fraser, 2001; Shiau et al., 2008), whereas in the transition of the PLLn,

Schwann cells, which are neural crest derivatives, are present in the epidermis and move with the axons into the subepidermal space.

34 Our results demonstrate that localization of the lateral line nerve occurs in two key phases. First, axons grow with precursors of their target neuromasts, ensuring proper connectivity and function at early stages (Gilmour et al., 2004). Next, associated

Schwann cells relocalize the axons across the epidermal basement membrane. The postembryonic remodeling of the lateral line sensory organs begins with migration of neuromasts and interneuromast cells ventrally within the epidermis by 72 hpf (Grant et al., 2005) (Fig. 2.6). By this time in wild type, the nerve has crossed the basement membrane and is therefore compartmentalized from neuromast migration. In mutants lacking Schwann cells, however, the nerve remains in the epidermis and is towed along with ventrally migrating neuromasts, resulting in progressive defasciculation and mislocalization from the horizontal myoseptum. Our data suggest that the transition of the PLLn from the epidermis into the subepidermal space insulates the nerve from the extensive postembryonic remodeling of the lateral line neuromasts by anchoring the nerve at the horizontal myoseptum.

The transition of the nerve across the basement membrane involves two distinct processes: the basement membrane deep to the nerve is degraded while a new basement membrane superficial to the nerve forms. Schwann cells appear to play an active role in both processes, as Schwann cells are required for the degradation of the basement membrane and can induce the formation of a new basement membrane even when the old one has not been degraded or when the nerve is improperly localized

(Fig. 2.3, 2.4D, 2.5E,F). Because Schwann cells can express matrix metalloproteases

(MMPs) (Baker and Bronner-Fraser, 2001; Ferguson and Muir, 2000; La Fleur et al.,

35 1996; Lehmann et al., 2009; Mantuano et al., 2008), which cleave extracellular matrix components (Page-McCaw et al., 2007), and can secrete a basement membrane that surrounds mature nerves (Prockop and Kivirikko, 1995), the simplest possibility is that Schwann cells directly secrete a new basement membrane superficial to the PLLn while degrading the deep basement membrane.

MMPs mediate basement membrane invasion in many contexts, including leukocyte invasion and anchor cell invasion in C. elegans (Madsen and Sahai, 2010; Page-

McCaw et al., 2007; Sherwood et al., 2005; Yadav et al., 2003). Additionally,

Schwann cells activate MMP expression following nerve injury, and MMPs have been implicated in Schwann cell myelination (Ferguson and Muir, 2000; La Fleur et al.,

1996; Lehmann et al., 2009; Mantuano et al., 2008). Intriguingly, MMP-2 was recently shown to have a role in degrading the basement membrane to allow dendrite remodeling in Drosophila sensory neurons (Yasunaga et al., 2010). Despite the role of MMPs in these other systems and in other aspects of Schwann cell biology, our preliminary experiments suggest that other enzymes may be involved in remodeling the epidermal basement membrane, because application of a broad-spectrum MMP inhibitor did not alter repositioning of the nerve (unpublished observation).

The process of localizing a nerve below a basement membrane to protect it from the remodeling of its targets is likely to be a common feature of sensory epithelia. In many sensory structures, including the tongue, epidermis, nose, vestibular organ, and lateral line, innervated receptive organs or free nerve endings are located within the

36 epithelium, while the main nerve plexus is located below the epithelial basement membrane (Boulais and Misery, 2008; Fernandez et al., 1990; Nedelec et al., 2005;

Northcutt, 2004; Oakley and Witt, 2004; Purcell and Perachio, 1997; Si et al., 2003;

Winklbauer, 1989). These are tissues, with the exception of the inner ear hair cells in mammals, where there is frequent turnover of the epithelium or the sensory cells themselves. For example, taste receptors and olfactory neurons are constantly reborn and replaced throughout life and the epidermis is continually being sloughed off and replaced (Fuchs and Horsley, 2008; Nedelec et al., 2005; Oakley and Witt, 2004).

Perhaps separating innervating axons from their targets protects the nerve from the turnover of its targets (or, in the case of the nasal epithelium, the turnover of individual neurons). Thus, as we show here in the lateral line nerve, Schwann cells may be important for positioning nerves in other sensory organs.

37 Figure 2.1

38 Figure 2.1: The PLLn invades the epidermal basement membrane

(A) 28 hpf embryo with schematic of the PLLn superimposed. Schwann cells are green, axons and the ganglion are red, and the primordium and a pre-neuromast

(asterisk) are blue. Dashed lines refer to approximate location of cross sections shown in C and D. (B) Schematic representation of the transition of the PLLn across the epidermal basement membrane. Basement membrane is shown in black, Schwann cells in green, axons in red, epidermal cells in yellow; only one of two epidermal cell layers is shown. Dashed lines correspond to cross sections in C, D, and E. (C) At 28 hpf, the distal segments of axons and associated Schwann cells of the PLLn are located within the epidermis (n = 6). Arrowheads mark the epidermal basement membrane. (D) At locations more anterior to that shown in (C) at 28 hpf, the PLLn is found in a transition state, with basement membrane located on both sides of the nerve

(arrowheads), arrow indicates the end of the basement membrane (n = 6). (E) By 3 dpf, the PLLn is found at its mature location, medial to the epidermal basement membrane (arrowheads, n = 9). A neuromast is present within the epidermis. (C’, D’,

E’) Higher magnification of boxed regions in C, D, E, respectively. (C’’, D’’, E’’)

C’, D’, E’ without pseudocoloring. Scale bars in C, D are 2 µm; 5 µm in E; 1 µm in

C’, D’, E’. Schwann cells are pseudocolored green, axons red. Abbreviations: D, dorsal; V, ventral; A, anterior; P, posterior; L, lateral; M, medial; ep, epidermis; m, muscle; p, pigment.

39 Figure 2.2

40 Figure 2.2: PLLn outgrowth is normal in Schwann cell deficient mutants

(A, B) At 28 hpf, axons and Schwann cells of heterozygous siblings (A) and erbb2st61 mutants (B) are located within the epidermis (ep) (n = 3 and 6 respectively) and are associated with pre-neuromast cells (pn). (A’, B’) Higher magnification of boxed region in A and B, respectively. Arrowheads indicate the location of the epidermal basement membrane. Scale bars for A and B are 5 µm; 1 µm in A’ and B’. Schwann cells are pseudocolored green and axons red. Abbreviations: ep, epidermis; m, muscle; pn, pre-neuromast cell.

41 Figure 2.3

42 Figure 2.3: The PLLn is mislocalized in mutants lacking Schwann cells

(A) At 5 dpf, the PLLn of wildtype siblings has migrated from the epidermis (ep), across the basement membrane (arrowheads), into the muscle (m) (n = 28). (B, C) In contrast, at 5 dpf in the Schwann cell deficient mutants, erbb2st61 (B) and sox10/clst3

(C), the PLLn remains in the epidermis, outside of the basement membrane

(arrowheads, n = 7 and 6 respectively). Defasciculation of the mutant nerves is also observed (asterisk, C). (A’, B’, C’) Higher magnification of boxed region in A, B and

C, respectively. Scale bars are 2 µm in A, B and C; 1 µm in A’, B’ and C’. Schwann cells are pseudocolored green and axons red. Abbreviations: ep, epidermis; m, muscle; p, pigment; n, neuromast cell.

43 Figure 2.4

44 Figure 2.4: Full length clones of transplanted wildtype Schwann cells can rescue nerve position in sox10/clst3 mutants

(A) Wildtype 3 dpf embryo carrying the FoxD3:GFP(17) transgene, labeling the

PLLn, motor nerves, and some pigment cells. (B) FoxD3:GFP(17) cells have been transplanted into a sox10/clst3 mutant host creating a clone of wildtype Schwann cells that traverse the entire length of the PLLn (3 dpf). (C) Transplanted wildtype

Schwann cells can reposition the mutant nerve in the subepidermal space, below the basement membrane (arrowheads). (D) Transplanted wildtype Schwann cells partially rescue the mutant nerve, positioning it within the epidermal basement membrane

(arrowheads). Insets show the presence of myelinated axons in C and D. Scale bars are 200 µm for A and B; 2 µm for C and D. Abbreviations: ep, epidermis; m, muscle.

45 Figure 2.5

46 Figure 2.5: Shorter length clones of transplanted wildtype Schwann cells can partially rescue the position of sox10/clst3 mutant nerves

(A) Wildtype 5 dpf embryo carrying the FoxD3:GFP(17) transgene, labeling the PLLn

(dorsal, d, and midbody, m, tracts), motor nerves, and some pigment cells. (B) A clone of wildtype, FoxD3:GFP(17) cells that are present on the dorsal (d) and anterior portion of the midbody (mb) tract of the PLLn. (C, D) Micrographs of the anterior

(C) and posterior (D) portions of a partial length clone of rescued Schwann cells, as shown in (B). (C) Wildtype Schwann cells rescue the position of the mutant nerve, but a part of the old basement membrane remains (arrows). Inset in C shows the presence of myelinated axons. (D) The posterior portion of the nerve lacks Schwann cells and is not rescued, remaining within the epidermis. (E, F) In this chimera, wildtype Schwann cells failed to rescue the position of the PLLn, but induced the formation of an ectopic basement membrane surrounding the nerve in the epidermis

(arrows). (F) Higher magnification of boxed area in E. Arrowheads indicate the epidermal basement membrane; arrows indicate ectopic basement membrane. Scale bars are 200 µm for A and B; 2 µm for C, D and F; 5 µm for E. Abbreviations: d, dorsal tract; mb, midbody tract; ep, epidermis; m, muscle.

47 Figure 2.6

48 Figure 2.6: Nerves become disorganized during development in mutants lacking

Schwnan cells

(A-O) Antibody staining for acetylated tubulin shows the position of the PLLn. (A-E)

Wildtype nerves remain anchored at the horizontal myoseptum and run straight along the length of the embryo at all times examined (48 hpf, 3-6 dpf), but the neuromasts migrate ventrally over time (arrowheads). (F-O) Nerves in erbb2st61 (F-J) and sox10/clst3 (K-O) mutants appear normal at 48 hpf but become progressively disorganized (3-6 dpf) and often track with migrating neuromasts (arrowheads).

Excess neuromasts are present in nerves lacking Schwann cells (F-O). Asterisks indicate the presence of pigment cells. Scale bar is 50 µm.

49 Chapter 3

ErbB Signaling has a Role in Radial Sorting Independent of Schwann Cell

Proliferation

Alya R. Raphael, David A. Lyons, William S. Talbot

This paper was submitted for publication on 11/29/10.

Author contributions: A.R.R. carried out all experiments, A.R.R., D.A.L., and W.S.T. designed the experiments, A.R.R. and W.S.T. wrote the paper.

50 ABSTRACT

In the peripheral nervous system, Schwann cells make myelin, a specialized sheath that is essential for rapid axonal conduction of action potentials. Immature Schwann cells initially interact with many axons, but, through a process termed radial sorting, eventually interact with one segment of a single axon as promyelinating Schwann cells. Previous studies have identified two classes of genes that control radial sorting: those that mediate Schwann cell process extension and those that regulate Schwann cell proliferation. It has also been shown that ErbB signaling is required for Schwann cell proliferation, myelination, radial sorting, and the proper formation of unmyelinated Remak bundles. In light of the evidence that both ErbB signaling and

Schwann cell proliferation are required in radial sorting, we wondered if the primary function of ErbB signaling in this process is to regulate Schwann cell proliferation. To address this question, we applied small molecule inhibitors in vivo in zebrafish to independently block ErbB signaling and proliferation during stages at which radial sorting normally takes place. Ultrastructural analysis of treated animals revealed that both ErbB signaling and Schwann cell proliferation are required for radial sorting in vivo. ErbB signaling, however, is required for Schwann cell process extension, while

Schwann cell proliferation is not. These results provide in vivo evidence that ErbB signaling plays a direct role in process extension during radial sorting, in addition to its role in regulating Schwann cell proliferation.

51 INTRODUCTION

Myelin is an essential component of vertebrate nerves, insulating axons to allow for the rapid conduction of action potentials. In the peripheral nervous system, myelin is made by Schwann cells, which wrap their cytoplasm many times around an axon to create the myelin sheath. Schwann cell precursors migrate along axons and then differentiate into immature Schwann cells that are faced with a fate choice (Jessen and

Mirsky, 2005). Immature Schwann cells associated with large caliber axons will become promyelinating and then myelinating Schwann cells, while those that continue to associate with several small caliber axons will become non-myelinating (“Remak”)

Schwann cells (Jessen and Mirsky, 2005; Nave and Salzer, 2006).

Prior to myelination, Schwann cells must transition from interacting with many axons as immature Schwann cells to just one segment of one axon as promyelinating

Schwann cells. This occurs by a process termed radial sorting, during which immature

Schwann cells progressively “sort” large caliber axons to the periphery of an axon bundle and then myelinate them (Webster et al., 1973). Initially, immature Schwann cells are arrayed around the edges of bundles of axons within the larger nerve (Jessen and Mirsky, 2005; Webster et al., 1973). Next, Schwann cells insert long lamellar processes into the axon bundle, interacting with several axons. A premyelinating

Schwann cell will pick just one axon, generally a larger axon, and sort it to the periphery of the family. At this point, the Schwann cell will differentiate into a promyelinating Schwann cell, express stage-specific transcription factors including

Oct6/Pou3f1 and Krox20/Egr2, begin wrapping the axon, and then make myelin

52 (Svaren and Meijer, 2008). Small caliber axons will ultimately be ensheathed by non- myelinating Schwann cells in Remak bundles.

To ensure that all axons are either myelinated or ensheathed by the end of radial sorting, Schwann cells must coordinate the extension and stabilization of their processes, in addition to regulating the number of Schwann cells present (Jessen and

Mirsky, 2005; Webster et al., 1973). The latter involves extensive proliferation of

Schwann cells during the period of radial sorting (Webster et al., 1973). A signaling network, involving extracellular signals from the basal lamina as well as intracellular regulators of the cytoskeleton and transcription, that regulates radial sorting has emerged over the past several years. Molecules that effect Schwann cell process extension include Rac1, β1-integrin, integrin linked kinase (ILK), and laminin-γ1

(Benninger et al., 2007; Chen and Strickland, 2003; Feltri et al., 2002; Nodari et al.,

2007; Pereira et al., 2009; Yu et al., 2009; Yu et al., 2005). Molecules required for

Schwann cell proliferation during radial sorting include Cdc42, focal adhesion kinase

(FAK), laminin-2, laminin-8, and laminin-γ1 (Benninger et al., 2007; Chen and

Strickland, 2003; Grove et al., 2007; Yang et al., 2005; Yu et al., 2009; Yu et al.,

2005).

A key signal during Schwann cell development is Neuregulin (Nrg), which is expressed in axons and signals through ErbB receptors on Schwann cells (Nave and

Salzer, 2006). Nrg/ErbB signaling controls many steps of Schwann cell development, including migration, proliferation, survival, radial sorting, Remak bundle formation,

53 and myelination (Dong et al., 1995; Garratt et al., 2000; Jessen and Mirsky, 2005;

Lyons et al., 2005; Michailov et al., 2004; Morrissey et al., 1995; Nave and Salzer,

2006; Riethmacher et al., 1997; Taveggia et al., 2005; Woldeyesus et al., 1999).

Because mammalian studies have revealed the importance of Nrg signaling during radial sorting and myelination, we wondered if Nrg/ErbB signaling is simply required to regulate Schwann cell proliferation during radial sorting or whether it plays a more direct role (e.g. in process extension). A previous in vitro study found that the addition of excess Schwann cells to a co-culture containing Nrg1 Type III deficient neurons was not sufficient to rescue myelination, suggesting that Nrg/ErbB signaling may be regulating something other than proliferation during the initiation of myelination, possibly radial sorting (Taveggia et al., 2005). To test this directly, we independently blocked ErbB signaling and proliferation using small molecule inhibitors just prior to the onset of myelination in zebrafish. This allowed us to temporally control inhibition of ErbB signaling and proliferation, so that early events in Schwann cell development proceeded normally. Ultrastructural analysis of drug treated nerves revealed that both ErbB signaling and Schwann cell proliferation are required for radial sorting. In both cases, there is an arrest of radial sorting, such that over time no new axons are sorted. The analysis revealed that inhibiting ErbB signaling and Schwann cell proliferation affect radial sorting in different ways. In the absence of ErbB signaling, no Schwann cell processes are present within the axon bundle, axon bundles are not subdivided into smaller bundles, and fewer axons are contacted by each Schwann cell. In contrast, Schwann cells were able to extend

54 processes into axon bundles in control larvae and those treated with cell cycle inhibitors, despite the sorting defect in the latter. Our analysis reveals that ErbB signaling regulates Schwann cell process extension during radial sorting, in addition to its role in controlling Schwann cell numbers through proliferation.

55 MATERIALS AND METHODS

Fish care

Wildtype embryos (TL and WIK) were raised at 28.5°C and were staged according to

(Kimmel et al., 1995).

Small molecule inhibitors

Small molecule inhibitors were added directly to the embryo medium at 48 hours postfertilization (h) and enough extra dimethyl sulfoxide (DMSO) was added to bring the total concentration to 1%. Control embryos were treated with 1% DMSO from 48 h. Embryo medium and inhibitors were changed daily. The following small molecule inhibitors were diluted in DMSO and used at the given concentrations: broad spectrum ErbB inhibitor, AG1478 at 2 µM (EMD; Lyons et al., 2005); S-phase inhibitor aphidicolin at 150 µM (Sigma; Spadari et al., 1985); and G2/M transition inhibitor camptothecin at 500 nM (EMD; Jones et al., 1997).

Immunofluorescence and in situ hybridization

Immunofluorescence and in situ hybridization staining were carried out using standard techniques. Anti-acetylated tubulin was used at a concentration of 1:1000 (Sigma), anti-MBP was used at a concentration of 1:50 (Lyons et al., 2005), and anti- phosphohistone H3 was used at a concentration of 1:200 (Upstate). Alexa 488 or 568 secondary antibodies (Molecular Probes) were used at a concentration of 1:2000.

Images were collected on a Zeiss Pascal LSM5 confocal microscope. oct6 and krox20

56 in situ probes were previously described (Lyons et al., 2005; Pogoda et al., 2006;

Rauch et al., 2003).

Live imaging

Embryos were anesthetized with 0.016% Tricaine (w/v) and mounted in 1.5% low melting agarose for live imaging.

Transmission electron microscopy

Transmission electron microscopy (TEM) was performed as previously described

(Lyons et al., 2008). Images were collected with a JEOL1230 and pseudocolored using Adobe Photoshop software.

Statistical analysis

Standard deviations and unpaired t test analyses (two tailed) were performed using

Microsoft Excel software.

57 RESULTS

ErbB signaling and Schwann cell proliferation are required for the expression of promyelinating markers

We previously found that inhibiting either ErbB signaling using the small molecule inhibitor AG1478 or Schwann cell proliferation with aphidicolin and hydroxyurea was sufficient to reduce the expression of myelin basic protein (MBP) in the zebrafish posterior lateral line nerve (Lyons et al., 2005). Because ErbB signaling is required for Schwann cell migration, we began treating the zebrafish embryos with small molecule inhibitors at 48 hours postfertilization (h), by which time Schwann cells have completed their migration along the lateral line nerve (Lyons et al., 2005). A reduced concentration of AG1478 (2 µM versus 4 µM) and alternative methods for blocking the cell cycle yielded a similar reduction in MBP staining (Fig. 3.1A-D). Aphidicolin blocks the cell cycle at S phase, while camptothecin blocks it at the G2/M transition

(Jones et al., 1997; Spadari et al., 1985). Additionally, there was a marked decrease in phosphohistone H3 positive nuclei, which indicates a cell in M phase, in the treated larvae as compared to control (data not shown). We also examined the expression of oct6 and krox20, which are expressed during the promyelinating stage (Svaren and

Meijer, 2008). Similar to previous work, we found that krox20 mRNA expression is reduced upon inhibition of ErbB signaling (Lyons et al., 2005; Fig. 3.1E,F).

Expression of krox20 was also reduced upon cell cycle inhibition (Fig. 3.1E,G,H).

Expression of oct6, which acts upstream of krox20 in promyelinating Schwann cells

(Svaren and Meijer, 2008), was also reduced after all three treatments (Fig. 3.1I-K).

58 These results suggest that ErbB signaling and post-migratory cell division are required for Schwann cells to progress to the promyelinating stage.

Radial sorting has begun in the lateral line nerve by 48 h

Previous studies suggest that radial sorting is underway in the lateral line nerve by 48 h, as Schwann cells express oct6 by this time (Monk et al., 2009), but the ultrastructure of the nerve at this stage has not been extensively investigated. Our ultrastructural analysis reveals that radial sorting is already underway at 48 h, and proceeds in a process similar to that described in mammals. Schwann cell processes are evident within the axon bundle of the lateral line nerve, and axons can be found at several stages – unsorted, contacted by a Schwann cell process, encircled by a

Schwann cell process, or with several wraps of nascent (uncompacted) myelin (Fig.

3.2A). A “sorted” axon is generally defined as one that is in a one-to-one configuration with a Schwann cell – it has been radially sorted away from the main axon bundle and is completely surrounded by Schwann cell cytoplasm (Jessen and

Mirsky, 2005). For the purposes of our analysis, we defined “sorted” as any axon that was completely separated from other axons by Schwann cell cytoplasm that is not obviously contacting other axons. For example, in figure 3.2A, the axons labeled “S” and associated with blue pseudocolored Schwann cell processes are sorted. In contrast, those axons associated with the yellow or green Schwann cell processes are not sorted, even though they are completely segregated from other axons, because these Schwann cell processes are associated with two axons (Fig. 3.2A, axons marked by asterisks).

59

By 24 hours later, at 3 days postfertilization (d), there are several myelinated axons with compacted Schwann cell cytoplasm wrapped around the axon (Fig. 3.2B). The process of radial sorting is still ongoing, as evidenced by the presence of a large cluster of unsorted axons and Schwann cell processes within the axon bundle (Fig.

3.2B). At 3 d, unsorted axons are of a similar size, and the sorted axons are generally larger than unsorted axons (Fig. 3.2). At this stage, we did not observe mature Remak bundles in the region of the posterior lateral line nerve that we examined (at the level of the sixth somite; Fig. 3.2). There were large bundles of unmyelinated axons, but these were not individually ensheathed, as evidenced by the direct apposition of axon membranes and the lack of Schwann cell membrane between axons (Fig. 3.2).

ErbB signaling and Schwann cell proliferation are required for radial sorting

In order to analyze the roles of ErbB signaling and proliferation during radial sorting, we conducted ultrastructural analysis of lateral line nerves in larvae treated with

AG1478, aphidicolin, or camptothecin from 48 h (after Schwann cell migration is complete) to 4 d. In all cases, there was a reduction in the number of sorted and myelinated axons (Fig. 3.3, 3.4, 3.5A). It is important to note that large caliber axons were present but not myelinated in the experimental conditions, which shows that these treatments do not generally disrupt axonal growth (Fig. 3.3; unsorted large caliber axons are marked with a “+”). Also, in all the treatment conditions immature

Schwann cells were present, but could not continue radial sorting. These results

60 indicate that ErbB signaling and Schwann cell proliferation are required for radial sorting in zebrafish.

Closer examination revealed that nerves from ErbB inhibited and cell cycle inhibited zebrafish have distinct radial sorting phenotypes. In AG1478-treated nerves, no

Schwann cell processes were inserted into the axon bundle, whereas there were processes present within axon bundles from control, aphidicolin-, or camptothecin- treated animals (Fig. 3.5C). We also quantified the number of axon clusters within the nerve. When Schwann cell processes enter the axon bundle they often bisect the nerve, creating separate clusters of axons. At 4 d, nerves from AG1478-treated larvae consistently had only one axon cluster, while nerves from control and cell cycle inhibited larvae had from 1 to 5 axon clusters (Fig. 3.5D; sorted and myelinated axons were not included). Because Schwann cells did not extend processes into axon bundles in AG1478-treated fish at 4 d, there were significantly fewer axons in contact with Schwann cells in these animals than in control and cell cycle inhibited larvae

(Fig. 3.5E). These results indicate that ErbB signaling, but not cell division, is required for the insertion of Schwann cells processes within axon bundles during radial sorting.

To determine if the defect we observe after inhibition of ErbB signaling or cell proliferation is due to a delay in Schwann cell development, we looked at earlier and later time points. A radial sorting defect is also apparent in nerves that had been treated with either AG1478 or aphidicolin from 48 h to 3 d (Fig. 3.6A-C). A

61 significant decrease in sorted axons was already apparent in AG1478-treated nerves, while there was a trend towards fewer sorted axons in aphidicolin-treated nerves (Fig.

3.5A). In nerves from larvae treated with AG1478 from 48 h to 5 d, the radial sorting defect persists and no new axons are sorted (Figs. 3.5A and 3.6D,E). By 5 d, aphidicolin-treated larvae displayed severe developmental defects, and we were not able to examine the nerves at this stage (data not shown, treated embryos at 4 d shown in Fig. 3.S1). There is no significant difference between the number of sorted axons at

48 h in control embryos and the number of sorted axons in AG1478-, aphidicolin-, or camptothecin-treated nerves at the later time points examined (Fig. 3.5A). In addition, the number of axons in AG1478 or camptothecin treated nerves was similar to control

(Fig. 3.5B). There was a slight reduction in the number of axons in embryos treated with aphidicolin, but it was not large enough to explain the radial sorting defect we observe (Fig. 3.5B). This suggests that inhibiting ErbB signaling or the cell cycle from 48 h completely arrests of radial sorting, since no new axons are sorted at 3-5 d in treated animals, despite the presence of immature Schwann cells.

ErbB signaling is required for the elaboration of the myelin sheath

Nrg/ErbB signaling regulates the thickness of the myelin sheath in mammals (Garratt et al., 2000; Michailov et al., 2004; Taveggia et al., 2005). Analysis of the axons sorted (likely prior to treatment) in nerves from AG1478-treated larvae revealed that

ErbB signaling is also required for elaboration of the myelin sheath in zebrafish.

Myelinated axons in AG1478-treated larvae consistently had fewer wraps of Schwann cell cytoplasm than those in the control larvae (Fig. 3.3A’’, B’’, 6D’, E’’; 5-6 wraps

62 in controls at 5 d vs. 2-3 wraps in AG1478-treated larvae). Schwann cells lacking

ErbB signaling, however, were able to extrude the cytoplasm from the wraps of myelin that they did form, indicating that ErbB signaling is not required for the compaction of the myelin sheath, similar to the situation in mammals (Garratt et al.,

2000; Michailov et al., 2004; Taveggia et al., 2005).

63 DISCUSSION

To examine the requirements for ErbB signaling and cell division during radial sorting in vivo, we applied small molecule inhibitors after Schwann cells completed their initial migration along the lateral line nerve in zebrafish. By applying the inhibitors after Schwann cell migration is complete, we can focus on the specific function of

ErbB signaling and cell proliferation during radial sorting without confounding effects resulting from defects at earlier steps in Schwann cell migration. In these experiments, the entire animal is bathed in the inhibitors, but it is likely that the defects in radial sorting result from direct effects on Schwann cells. ErbB2 and ErbB3 receptors are expressed in Schwann cells, and these ErbB receptors act autonomously in Schwann cells in other contexts in zebrafish and mammals (Garratt et al., 2000;

Grant et al., 2005; Lyons et al., 2005; Nave and Salzer, 2006; Riethmacher et al.,

1997). Additionally, Schwann cells proliferate extensively before reaching the promyelinating stage in zebrafish and mammals, consistent with a requirement for

Schwann cell proliferation during radial sorting (Lyons et al., 2005; Webster et al.,

1973). Blockade of either ErbB signaling or cell division causes significant defects in radial sorting, but there is an important difference between these two treatments. ErbB signaling was required for the extension of Schwann cell processes into the axon bundle, whereas Schwann cell proliferation was not. Supporting and extending a previous in vitro study (Taveggia et al., 2005), our experiments provide in vivo evidence that ErbB signaling plays a role in regulating Schwann cell process extension during radial sorting, in addition to its more extensively characterized role in stimulating Schwann cell proliferation.

64

Previous work supports the idea that the levels of Nrg expressed on an axon determine the differentiated fates of the associated Schwann cells. High levels of Nrg signal coming from a large axon instruct a Schwann cell to differentiate as a myelinating cell, while low levels of Nrg from several small caliber axons lead to the formation of a

Remak bundle (Michailov et al., 2004; Nave and Salzer, 2006; Taveggia et al., 2005).

Additionally, the amount of Nrg produced in a large caliber axon determines the thickness of the myelin sheath (Michailov et al., 2004; Taveggia et al., 2005). Similar to these mammalian studies, inhibition of ErbB signaling in zebrafish blocks the elaboration of the myelin sheath (Fig. 3.3, 3.6; Lyons et al., 2005). Additionally, radial sorting arrests when ErbB signaling is blocked in vivo (Fig. 3.3, 3.5, 3.6).

Because the infiltration of an axon bundle during radial sorting is similar to ensheathment of axons in a Remak bundle, it is possible that the defects observed in radial sorting and Remak bundle formation in the absence of Nrg-ErbB signaling

(Taveggia et al., 2005) are caused by the failure of Schwann cells to properly extend processes into an axon bundle in the absence of ErbB signaling.

Radial sorting requires the coordination of Schwann cell process extension and

Schwann cell proliferation (Jessen and Mirsky, 2005; Webster et al., 1973). Schwann cell process extension and stabilization is required for Schwann cells to be able to subdivide the axon bundle and sort axons to the periphery (Benninger et al., 2007;

Chen and Strickland, 2003; Feltri et al., 2002; Nodari et al., 2007; Pereira et al., 2009;

Webster et al., 1973; Yu et al., 2009). Schwann cell proliferation is critical for

65 properly matching the number of Schwann cells to the number of axons that will be myelinated (Benninger et al., 2007; Grove et al., 2007; Jessen and Mirsky, 2005; Yang et al., 2005; Yu et al., 2005). Accordingly, many of the genes involved in radial sorting seem to primarily regulate one or the other of these Schwann cell functions

(Benninger et al., 2007; Feltri et al., 2002; Grove et al., 2007; Nodari et al., 2007;

Pereira et al., 2009; Yang et al., 2005). The group of genes that regulate Schwann cell process extension but not proliferation includes rac1, β1-integrin, and ILK (Benninger et al., 2007; Feltri et al., 2002; Nodari et al., 2007; Pereira et al., 2009). A second group of genes is required for proliferation but not process extension and includes cdc42, FAK, laminin-2, and laminin-8 (Benninger et al., 2007; Grove et al., 2007;

Yang et al., 2005). While it is useful to classify these radial sorting genes into two complementary groups, the two classes are not mutually exclusive, likely reflecting close coordination between Schwann cell proliferation and division. For example, laminin-γ1 is required for both process extension and Schwann cell proliferation

(Chen and Strickland, 2003; Yu et al., 2009; Yu et al., 2005), and our data suggest that

Nrg/ErbB signaling is also required for both process extension and proliferation during radial sorting.

In conclusion, we find that ErbB signaling is required for Schwann cell process extension in addition to regulating Schwann cell proliferation during radial sorting. It is important to note that other signals must also activate Schwann cell division, because these cells do divide—although in reduced numbers—when ErbB signaling is blocked (Lyons et al., 2005). Further studies will be needed to understand how

66 Schwann cell proliferation and process extension are coordinated during radial sorting.

Additionally, it is unclear whether Schwann cell proliferation is required during radial sorting solely to increase Schwann cell numbers, or whether there is role for a special terminal or polarized division before a Schwann cell can commit to myelination.

Finally, it is unclear how Neuregulin and other signals instruct a Schwann cell to withdraw some processes while maintaining one process around the axon it will later myelinate, and control the decision between Remak bundle formation and the continuation of radial sorting.

67 Figure 3.1

68 Figure 3.1: ErbB signaling and Schwann cell proliferation are required for the normal expression of myelinating and promyelinating markers.

Myelin basic protein (MBP, expressed in myelin) expression is reduced along the posterior lateral line nerve in animals treated with AG1478 (B), aphidicolin (C), and camptothecin (D) as compared to DMSO-treated controls (A). MBP is in green, acetylated tubulin (AcTub) marking axons is shown in red. MBP is shown alone in

A’-D’. krox20 and oct6 expression is also reduced in AG1478 (F, J), aphidicolin (G,

K), and camptothecin (H, L) treated nerves as compared to DMSO control nerves (E,

I).

69 Figure 3.2

70 Figure 3.2: Radial sorting is underway at 48 h and continues over time in the posterior lateral line nerve.

(A) Ultrastructural analysis of a 48 h lateral line nerve reveals that radial sorting is already underway. In this example, three axons are already sorted (s) and sequestered from other axons by an individual Schwann cell (pseudocolored blue or red). Two sorted axons already have two wraps of Schwann cell cytoplasm around them

(pseudocolored blue). Other Schwann cell processes (pseudocolored yellow and green) enter the axon bundle and interact with more than one axon (*). (A’) Same image as in (A) without pseudocoloring. (A’’-A’’’) Magnification of regions boxed in

A’. (B) By 3 d, several myelinated axons (m) are present within the posterior lateral line nerve; different pseudocolors indicate that each myelinated axon is associated with its own Schwann cell. A Schwann cell process pseudocolored in orange is present within the unsorted axon bundle. (B’) Same image as in (B) without pseudocoloring. (B’’-B’’’) Magnification of regions boxed in B’. Scale bars are 1

µm in A, A’, B, and B’ and 0.5 µm in A’’, 0.2 µm in A’’’ and B’’, and 0.3 µm in B’’’.

71 Figure 3.3

72 Figure 3.3: ErbB signaling and Schwann cell proliferation are required for radial sorting.

(A) Cross section through the posterior lateral line nerve in a control treated with

DMSO from 48 h to 4 d (DMSO 48h-4d) reveals several myelinated axons (m) and two sorted axons (s), Schwann cell cytoplasm pseudocolored blue. Two Schwann cell processes subdivide the large bundle of unsorted axons into three smaller clusters, one with only two axons at the top of the nerve. Shown without pseudocoloring in (A’) magnified image of boxed myelinated axon (A’’) or boxed sorted axon (A’’’). (B) In the nerve treated with AG1478 from 48 h to 4 d (AG1478 48h-4d) there is a significant reduction in the number of myelinated (m) and sorted axons. However, several large caliber axons (+) that are similar in size to myelinated axons in control nerves (A) remain in the large bundle of unsorted axons. Schwann cell cytoplasm pseudocolored in blue; no Schwann cell processes are present within the axon bundle; however, Schwann cell processes still surround the exterior of the bundle. (B’) Same image as in B shown without pseudocoloring, magnified boxed regions shown in B’’ and B’’’. Fewer wraps of myelin are present around the one myelinated axon as compared to the DMSO control (A’ versus B’). (C) Similar to nerves treated with

AG1478, nerves treated with aphidicolin from 48 h to 4 d (Aph 48h-4d) also have a reduction in the number of myelinated axons (m, boxed region magnified in C’).

However, sorted axons (s) are still present and Schwann cell processes (pseudocolored blue) have split the main axon bundle into three axon clusters. Like AG1478 treated nerves, there are several unsorted, large caliber axons present (+). (C’) Same image as in C shown without pseudocoloring, boxed regions magnified in C’’, C’’’. Scale bars

73 are 2 µm in A, A’ and C, C’, 1 µm in B, B’, 0.5 µm in A’’, A’’’, C’’, and C’’’, and

0.2 µm in B’’ and B’’’.

74 Figure 3.4

75 Figure 3.4: Another cell cycle inhibitor, camptothecin, also inhibits radial sorting.

(A) There are several myelinated axons (m, boxed region magnified in A’) and

Schwann cell processes have bisected the main axon bundle into one unsorted bundle at the bottom of the nerve and a small cluster with two axons at the top of the nerve in the DMSO control nerve at 4 d (DMSO 48h-4d). In contrast, in the camptothecin- treated nerve from 48 h to 4 d (B, Camp 48h-4d), there are fewer myelinated axons

(m, boxed region magnified in B’). One sorted axon is present (s) and Schwann cell cytoplasm has divided the main axon bundle into two sections, a large bundle to the left and a bundle of two axons to the right. Scale bars are 2 µm for A, 1 µm for B, and

0.5 µm for A’ and B’.

76 Figure 3.5

77 Figure 3.5: ErbB signaling and Schwann cell proliferation play distinct roles in radial sorting.

(A) The number of sorted axons was quantified for several treatment conditions at several time points. Control nerves are treated from 48 h with DMSO. There is a significant decrease in the number of sorted axons in AG1478 treated nerves at 3, 4, and 5 d (p = 5.8×10-5, p = 3.7×10-7, and p = 5.5×10-8, respectively). There is a significant decrease in the number of sorted axons in aphidicolin and camptothecin treated nerves at 4 d (p = 7.4×10-5 and p = 6.6×10-6, respectively). (B) The total number of axons per nerve was quantified for all of the treatment conditions. AG1478 and camptothecin treated nerves had similar axon numbers to control, while aphidicolin treated nerves had slightly reduced axon numbers (p = 0.001 and p = 0.007 for 3 and 4 d respectively). (C) The number of Schwann cell processes extended into the axon bundle was counted for each condition at 4 d and at 48 h in control nerves.

No Schwann cell processes were ever found in AG1478 treated nerves (p = 4.2×10-5), while all other treatments had processes extended. (D) The number of axon clusters within the nerve was counted for all treatments at 4 d and for the control at 48 h.

AG1478 consistently had only one large bundle of axons (p = 0.038), while the other treatments had 1-5 clusters. (E) The number of axons touched by a Schwann cell, including myelinated and sorted axons, was counted for each of the treatments at 4 d and for the control at 48 h. AG1478 treated nerves had significantly fewer axons being touched by a Schwann cell as compared to all other treatments (p = 0.0004).

Error bars show + 1 standard deviation. The number of nerves examined is as follows:

48 h control, 7; 3 d control, 6; 4 d control, 10; 5 d control, 7; 3 d AG1478, 6; 4 d

78 AG1478, 5; 5 d AG1478, 6; 3 d aphidicolin, 5; 4 d aphidicolin, 4; 4 d camptothecin, 7.

AG1478 and aphidicolin experiments were performed twice with similar results; results from one experiment shown. All p values are from two tailed unpaired t tests versus the age-matched control. * = p < 0.05, ** = p < 0.005, *** = p < 0.0005.

Abbreviations: Aph, aphidicolin; camp, camptothecin.

79 Figure 3.6

80 Figure 3.6: Inhibiting ErbB signaling or proliferation arrests radial sorting.

(A) In control nerves that have been treated with DMSO from 48 h to 3 d (DMSO 3d), a few myelinated axons (m) are present, as well as several sorted (s), and two axons that are being touched by the same Schwann cell process (*). In contrast, in nerves treated with AG1478 from 48 h to 3 d (AG1478 3d), there are myelinated axons present (m), but no additional sorted axons (B). In nerves treated with aphidicolin from 48 h to 3 d (Aph 3d), again myelinated axons are present (m) and a Schwann cell process is bisecting the main bundle of unsorted axons. (D) In nerves treated with

DMSO from 48 h to 5 d (DMSO 5d), there are many myelinated axons (m, boxed region in D’) and a few sorted axons (s). The bundle of unsorted axons has been split into two – a small bundle at the top of the nerve and a large bundle at the lower right.

(E) Nerves treated with AG1478 from 48 h have a similar number of myelinated axons (m, boxed region magnified in E’) as at 3 d (B), suggesting that radial sorting has been arrested. Myelinated axons in nerves treated with AG1478 have fewer wraps of myelin than those in the DMSO control (E’ versus D’). Scale bars are 1 µm in A-E and 0.5 µm in D’ and E’.

81 Figure 3.S1

82 Figure 3.S1: Larvae treated with small molecule inhibitors

(A-D) Zebrafish larvae treated with DMSO, AG1478, aphidicolin, or camptothecin from 48 h – 4 d were live imaged. (C-D) Aphidicolin and camptothecin treated larvae have heart edema.

83 APPENDIX 1

Schwann cells are required for proper axon number in the posterior lateral line nerve

Authors: Alya R Raphael, Julie R Perlin, William S Talbot.

Figure A1.1 is adapted from Voas MG, Glenn TD, Raphael AR, and Talbot WS.

(2009). Schwann cells inhibit ectopic clustering of axonal sodium channels. J.

Neurosci. 29(46), 14408-14. Additionally, the 3 days postfertilization (dpf) axon counts in figure A1.2 were also published in the above paper. The data shown is all my own work, with the exception of the transplants, which were performed by Julie R.

Perlin and analyzed by me.

While performing control experiments for the paper listed above, I found that there is a significant reduction in the number of axons in mutants lacking Schwann cells at 3 days postfertilization (3 dpf, sox10/clst3, erbb3st48, and erbb2st61, Fig. A1.1, A1.2A).

This is also true at 5 dpf (Fig. A1.2A). Strikingly, the axons that are lost are small caliber axons, while large caliber axons remain in the proper numbers (Fig. A1.2B).

However, at an earlier time point, while the nerve is growing out, there is no defect in axon number (28 hours postfertilization, Fig. A1.3). In order to see whether this was due to an autonomous loss of sox10 in neurons, or due to a loss of Schwann cells, we transplanted lineage labeled wildtype cells into sox10/clst3 mutant hosts. Wildtype

Schwann cells rescued the defects in axon number and also restored myelin (Fig.

84 A1.4), while wildtype neurons did not. From these data, we conclude that Schwann cells are required non-cell autonomously to regulate the number of axons within the posterior lateral line nerve (PLLn). How Schwann cells are exerting this effect remains unknown – future studies will explore the number, proliferation, and cell death of neurons within the posterior lateral line ganglion.

85 Figure A1.1

86 Figure A1.1: In mutants lacking Schwann cells, the posterior lateral line nerve is defasciculated and smaller.

(A) At 3 dpf, the posterior lateral line nerve (PLLn) in wildtype siblings has several large, myelinated axons (asterisks), and other smaller unsorted axons. (B-D) The

PLLn of mutants lacking Schwann cells, sox10/clst3, erbb3st48, and erbb2st61, has no myelin, is smaller, and defasciculated axons are present (arrowheads in B, C).

87 Figure A1.2

88 Figure A1.2: Mutants without Schwann cells are deficient for small caliber axons

(A) There is a significant reduction in the number of axons in nerves lacking Schwann cells (sox10/clst3 and erbb2st61) at 3 and 5 dpf (two tailed t tests against aged matched wildtype siblings, clst3 3 dpf p = 4.76×10-11, erbb2st61 3 dpf p = 9.89×10-8, clst3 5 dpf p

= 5.61×10-10, erbb2st61 5 dpf p = 4.39×10-8). (B) Small caliber axons are missing in mutants lacking Schwann cells. There is the normal number of large caliber axons

(≥0.1 µm2, typically myelinated in wildtype siblings) in mutants lacking Schwann cells, with a slight increase in large axons in erbb2st61 at 5 dpf (two tailed t test against age matched wildtype siblings, p = 0.045). In contrast at 3 and 5 dpf there is a significant decrease in the number of small caliber axons (<0.1 µm2) in sox10/clst3 and erbb2st61 mutants (two tailed t test against age matched wildtype siblings, clst3 3 dpf p

= 0.0002, erbb2st61 3 dpf p = 0.0008, clst3 5 dpf p = 9.35×10-5, erbb2st61 5 dpf p =

5.37×10-5). Error bars indicate +/- one standard deviation.

89 Figure A1.3

90 Figure A1.3: Axon number during initial outgrowth is normal in mutants lacking Schwann cells

At 28 hours postfertilization (hpf), a time at which the PLLn is growing out, wildtype

(A) and erbb2st61 (B) mutant nerves have similar numbers of axons. Quantified in (C).

The wildtype nerve (A) is surrounded by a Schwann cell process. Scale bars in A, B indicate 0.5 µm. Error bars indicate +/- one standard deviation.

91 Figure A1.4

92 Figure A1.4: Schwann cells are required non-cell autonomously to regulate axon number in the PLLn

(A) At 5 dpf, many myelinated axons are present in wild type nerves. (B) In clst3 mutant nerves, there are fewer axons and no myelinated axons. (C) Wildtype cells were transplanted into clst3 mutant hosts (wt → clst3). The presence of wildtype

Schwann cells rescues the number of axons and myelinated axons. Quantified in (D,

E). Scale bars are 2 µm.

93 APPENDIX 2

αII-spectrin is required for myelination of motor axons

This analysis was performed as part of a collaboration that resulted in the following manuscript that is in preparation: Keichiro Susuki, Alya R. Raphael, Yasuhiro

Ogawa, Michael C. Stankewich, Elior Peles, William S. Talbot, Matthew N. Rasband.

(2010) Spectrins couple neuron-glial interactions to the actin cytoskeleton for peripheral nerve myelination. The work shown below is all my own. Spectrins play well known roles in linking the plasma membrane to the cytoskeleton. Voas et al. previously found that αII-spectrin is required for proper node of Ranvier formation

(Voas et al., 2007). K. Susuki and M. Rasband found that in vitro spectrins are required for myelination. Since we had an αIIspnst60 mutant in hand and previous evidence had indicated that there might be a defect in myelination in motor nerves but not in the posterior lateral line nerve (Voas et al., 2007), we analyzed peripheral nerves in wildtype and αIIspnst60 mutants. We found that there is a significant decrease in the number of motor nerves with myelinated axons in αIIspnst60 mutants, but that there is no apparent defect in the posterior lateral line nerve (Fig. A2.1, Table

A2.1).

94 Figure A2.1

95 Figure A2.1: αII-spectrin is required for myelination of motor axons

(A) Wildtype motor nerves have 1-2 myelinated axons at 7 days postfertilization (dpf).

(B) Approximately 60% of all αIIspnst60 mutant motor nerves lack myelinated axons.

However, most large axons are surrounded by one wrap of Schwann cell cytoplasm

(D). (C, D) Larger view of a large axon from A and B. (E, F) Myelination of the posterior lateral line nerve (PLLn) is comparable between αIIspnst60 mutants (F) and heterozygous siblings (E). Scale bars are 1 µm for A,B and 0.5 µm for C-F.

Genotypes were confirmed by PCR assay as in (Voas et al., 2007) prior to TEM.

96 Table A2.1: αIIspnst60 mutant motor nerves have a reduction in myelinated axons at 7 and 9 dpf

Number of MN with 1 Average Average number myelinated axon/total number of of myelinated Genotype MN examined axons in the axons in the PLLn PLLn 7 αIIspnst60/+ 30/30 (100%) 66.2 (± 7.8) 15.8 (± 2.2) dpf αIIspnst60 15/36 (41.7%) 65.6 (± 5.1) 13.5 (± 1.7) 9 αIIspnst60/+ 36/36 (100%) 56.5 (± 5.54) 16.7 (± 1.75) dpf αIIspnst60 15/26 (57.7%) 48.5 (± 1.52)** 12.7 (± 1.86)**

By 9 dpf there is a defect in the number of axons and the number of myelinated axons in the PLLn (p = 0.007 and 0.003 respectively). ± one standard deviation is shown for the PLLn counts. For the motor nerves at 7 dpf 4 embryos were examined in mutants and heterozygous siblings, at 9 dpf 3 embryos each were examined. For the PLLn, at 7 dpf 5 nerves from 3 embryos were examined and at 9 dpf 6 nerves from 3 embryos were examined for heterozygous siblings and αIIspnst60 mutants.

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