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Home , Glia, Histology, Neuron, Satellite glial cell, Schwann cell, Sympathetic ganglia

NEURON-SATELLITE GLIAL INTERACTIONS IN SYMPATHETIC DEVELOPMENT

by Erica D. Boehm

A dissertation submitted to the Johns Hopkins University in conformity with the requirements for the degree of Doctor of Philosophy

Baltimore, Maryland July 2020

© 2020 Erica Boehm All rights reserved.

ABSTRACT

Glial cells play crucial roles in maintaining the stability and structure of the nervous system. Satellite glial cells are a loosely defined of glial cells that ensheathe neuronal cell bodies, , and of the peripheral nervous system

(Elfvin and Forsman 1978; Pannese 1981). Satellite glial cells are closely juxtaposed to peripheral with only 20nm of space between their membranes (Dixon 1969).

This close association suggests a tight coupling between the cells to allow for possible exchange of important , yet very little is known about and development. How neurons and glial cells co-develop to create this tightly knit unit remains undefined, as well as the functional consequences of disrupting these contacts.

Satellite glial cells are derived from the same population of cells that give rise to peripheral neurons, but do not begin differentiation and proliferation until has been completed (Hall and Landis 1992). A key signaling pathway involved in glial specification is the Delta/ (Tsarovina et al. 2008). However, recent studies also implicate Notch signaling in the maturation of through non- canonical Notch ligands such as Delta/Notch-like EGF-related (DNER) (Eiraku et al. 2005). Interestingly, it has been reported that levels of DNER in sympathetic neurons may be dependent on the target-derived growth factor, growth factor

(NGF), and this is prominent in sympathetic neurons at the in which satellite glial cells are developing (Deppmann et al. 2008; Hall and Landis 1992).

Here, we find that the close association of satellite glial cells with sympathetic neuronal cell bodies is mediated by neuronal expression of DNER. In mice, conditional deletion of DNER from sympathetic neurons disrupts -glia contacts. Loss of

ii DNER and/or neuron-glia contacts has profound effects on the neurons, resulting in hyper-innervation of targets, decreased neuron size, and an increase in activity.

Additionally, DNER expression is regulated by target-derived NGF. This suggests a tripartite system in which the development of neurons and glial cells may be coordinated by the same distant signal, and supports a role for neuron-satellite glia contacts in neuronal architecture and activity.

Advisor: Rejji Kuruvilla, Ph.D.

Second Reader: Haiqing Zhao, Ph.D.

Committee Members: Mark Van Doren, Ph.D.

Seth Blackshaw, Ph.D.

Hey-Kyoung Lee, Ph.D.

Haiqing Zhao, Ph.D.

iii PREFACE Nothing I have done in life prior to graduate school could have prepared me for the trials and errors of my time here. It has been an emotional and intellectual rollercoaster with twists, turns, and loops of failures and successes. I believe I have come off this ride a better, more confident person in my work life and in my personal life. I have discovered the aspects of science that I love as well as the aspects of it that I do not like so much, and the growth I have gone through as a person is something that I will carry with me for the rest of my life wherever it me.

During my time as a graduate student in Dr. Rejji Kuruvilla’s lab, I have learned how to think for myself, how to design experiments, a project, and mentor students with patience and kindness. I have gotten to dip my hands into many different techniques, some of which were very helpful while others did not bear fruit as we had hoped. I have learned how to deal with failure and disappointment when experiments didn’t work or when they gave confusing results, and I learned how to think outside the box to find new solutions to problems. My time here has transformed me into a person I never I could be; someone who can provide valuable insight during lab meetings, who can think critically about data being presented, and who figured out how to appreciate the small successes as much as the big ones.

I have a number of people to thank for my growth throughout graduate school.

First, I would like to thank my mentor, Rejji, who constantly kept pushing me to be better, think harder, and dig deeper for answers. I’d also like to thank the Mouse Tri-lab as a and the individuals within who have really helped me along the way.

Without the Mouse Tri-lab, I would not be the thoughtful person I am today. The unique

iv environment of the Tri-lab community nurtured my scientific curiosity. I would especially like to thank those who have helped me with my project experimentally and through their critical . Particularly I would like to give thanks to my lab mates, current and former, Dr. Alexis Ceasrine, Dr. Ami Patel, Aurelia Mapps, Blaine Connor,

Dr. Chantal Bodkin-Clarke, Dr. Chih-Ming Chen, Dr. Emily Scott-Solomon, Dr. Eugene

Lin, Dr. Jessica Houtz, and Nelmari Ruiz-Otero for helping with experiments, lending expert advice, and being shoulders to lean on. Without these people, I would not be the I am. I also want to thank a collaborator, Dr. Bryan Jones, for his contribution to my work. Without his help, this work would not be where it is today. Finally, I would like to thank the professors from the Mouse Tri-lab and my committee for their support and generosity throughout my time here. So, thank you to Dr. Haiqing Zhao, Dr. Hey-

Kyoung Lee, Dr. Mark van Doren, Dr. Samer Hatter, and Dr. Seth Blackshaw. I am grateful to everyone, friends, family, and colleagues, who supported me and kept me afloat while obtaining my Ph.D.

v TABLE OF CONTENTS

ABSTRACT…………………………………………………………………………...... ii

PREFACE………………………………………………………………………………..iv

TABLE OF CONTENTS…………………………………………………………..…...vi

LIST OF FIGURES………………………………………………...….…...... viii

LIST OF ABBREVIATIONS…………………………………………………...... x

CHAPTER ONE: INTRODUCTION…………………………………………..……....1

CHAPTER TWO: REGULATION AND EXPRESSION OF DNER IN THE

SYMPATHETIC NERVOUS SYSTEM……………………………………………....16

INTRODUCTION………………………………………………………………...17

RESULTS………………………………………………………………...... 22

DISCUSSION………………………………………………………………...... 31

METHODS………………………………………………………………...... 33

CHAPTER THREE: NEURONAL DNER IS REQUIRED FOR SYMPATHETIC-

SATELLITE GLIAL CELL CONTACTS……………………………………………40

INTRODUCTION………………………………………………………………...41

RESULTS………………………………………………………………...... 46

DISCUSSION………………………………………………………………...... 58

METHODS………………………………………………………………...... 61

CHAPTER FOUR: DISRUPTION OF NEURONAL AND

ACTIVITY IN DNER CKO MICE……………………………………………………67

INTRODUCTION………………………………………………………………...68

RESULTS………………………………………………………………...... 73

vi DISCUSSION………………………………………………………………...... 88

METHODS………………………………………………………………...... 93

CHAPTER FIVE: GENETIC ABLATION OF SATELLITE GLIAL CELLS

DURING SYMPATHETIC NERVOUS SYSTEM DEVELOPMENT……………100

INTRODUCTION…………………………………………………………….…101

RESULTS………………………………………………………………...... 105

DISCUSSION………………………………………………………………...... 115

METHODS………………………………………………………………...... 117

CHAPTER SIX: MOLECULAR CHANGES IN DNER CKO SYMPATHETIC

GANGLIA………………………………………………………..……………………123

INTRODUCTION…………………………………………………………...... 124

RESULTS………………………………………………………………...... 126

DISCUSSION………………………………………………………………...... 130

METHODS………………………………………………………………...... 131

CLOSING REMARKS………………………………………………………..…...….133

REFERENCES………………………………………………………..……...... 141

CURRICULUM VITAE………………………………………………………………168

vii LIST OF FIGURES

Chapter Two:

Figure 2.1. DNER is expressed in all three major compartments of sympathetic neurons…………………………………………………………………………………...26

Figure 2.2. DNER has a half-life around 4 hours and partially localizes to the cell surface membrane………………………………………………………………………………...28

Figure 2.3. DNER is regulated by NGF-TrkA signaling………………………………...29

Figure 2.4. Disruption of satellite glial cell sheaths in TrkA knockout …………………………………………………………………………………....30

Chapter Three:

Figure 3.1. DNER alters satellite glial cell morphology in vitro………………………...51

Figure 3.2. DNER-Fc is capable of activating Notch signaling in cultured satellite glial cells………………………………………………………………………………………52

Figure 3.3. DNER is knocked down in DNER cKO sympathetic ganglia……………….53

Figure 3.4. Cell proliferation is unaffected in neonatal DNER cKO sympathetic ganglia……………………………………………………………………………………54

Figure 3.5. Satellite glial cell ring structure is lost in sympathetic ganglia of DNER cKO mice………………………………………………………………………………………55

Figure 3.6. Sympathetic ganglia morphology is disrupted in DNER cKO mice………...56

Figure 3.7. Neuron-satellite glial cell contacts are lost in DNER cKO ganglia………….57

Chapter Four:

viii Figure 4.1. Loss of adrenergic biosynthetic pathway expression in DNER cKO ganglia……………………………………………………………………………………78

Figure 4.2. DNER cKO neurons are smaller in size……………………………………..79

Figure 4.3. Neuronal survival is unaffected in DNER cKO ………………….....80

Figure 4.4. DNER cKO neurons hyper-innervate the . …………………………….83

Figure 4.5. Salivary glands are hyper-innervated in DNER cKO mice………………….84

Figure 4.6. Cultured DNER cKO neurons do not exhibit changes in length or branching…………………………………………………………………………………85

Figure 4.7. DNER cKO neurons have a sustained response to ….86

Figure 4.8. Altered mitochondrial morphology in DNER cKO neurons………………...88

Chapter Five:

Figure 5.1. Increased cell death in DT-injected sympathetic ganglia…………………..111

Figure 5.2. No changes in neuronal cell number in DT-injected ganglia with 3 or 5 days of DT injection …………………………………………………………………………112

Figure 5.3. Loss of satellite glial cell ring structure in 5-day DT-injected sympathetic ganglia but no loss in neuronal identity.………………………………………………..113

Figure 5.4. Loss of satellite glial cells results in neuronal cell atrophy ..………...114

Figure 5.5. Loss of satellite glial cells during development results in hyper-innervation of sympathetic target tissues………………………………………………………………115

Figure 5.6. Sympathetic neurons have an altered response to depolarization with loss of satellite glial cells………………………………………………………………….……117

ix Chapter Six:

Figure 6.1. Molecular changes in DNER cKO sympathetic ganglia……………………128

Chapter Seven:

Figure 7.1. Tripartite interactions between peripheral target tissues, sympathetic neurons, and satellite glial cells during development…………………………………………….139

x LIST OF ABBREVIATIONS

Ascl1 (BHLH factor 1)

BDNF (-derived neurotrophic growth factor)

BLBP (brain binding )

BMP ( morphogenetic protein)

CHX (cycloheximide)

CNS ()

Cnx43 (-43)

CSL (CBF1–Suppressor of Hairless–LAG1)

DAPT (N-[N-(3,5-Difluorophenacetyl)-L-alanyl]-S-phenylglycine t-butyl ester)

DBH ( β-hydroxylase)

DNER (Delta/Notch-like EGF-related Receptor)

DRG (dorsal root )

DT (diphtheria toxin)

DTR (diphtheria toxin receptor)

EMT (epithelial-to-mesenchymal)

FGF ( growth factor)

GFAP (glial fibrillary acidic protein)

GLAST (glutamate-aspartate transporter)

GO-term (gene ontology term)

GS ()

Hand2 (heart and derivatives-expressed protein 2)

Hes (Hairy and Enhancer of split)

xi Hey (Hes Related Family BHLH With YRPW Motif)

HGF (hepatocyte growth factor)

IHC ()

Kir (inwardly rectifying channel)

NCC (neural crest cell)

NGF ()

NL ()

Nrg1 ( 1)

Nrp1 (neuropilin1)

NSC (neural )

NT3 ( 3)

NT4 (neurotrophin 4)

Phox2b (paired-like homeobox 2b)

PNS (peripheral nervous system)

PTP (protein phosphatase )

SCG (superior cervical ganglion)

Sema3a (semaphorin3a)

SER (smooth )

SIF (small intensely fluorescent cell)

TEM (transmission electron )

TH ()

xii Chapter One: Introduction

1

Sympathetic Nervous System Function and Formation

The mammalian nervous system comprises two separate but interconnected systems, the central and peripheral. The central nervous system (CNS) includes the brain and , while the peripheral nervous system (PNS) encompasses the neurons and associated cells throughout the rest of the body. Everything from the ability to regulate to feeling is regulated by the PNS. Defects in, or trauma to, this system can result in dozens of different diseases such as Guillain-Barre Strohl Syndrome,

Postural Orthostatic Tachycardia Syndrome, and as well as several peripheral neuropathies that affect nearly every throughout the body (Goldstein et al. 2002).

The PNS is divided into two groups: the sensory nervous system and the motor nervous system, which includes the somatic motor, sympathetic, and parasympathetic systems. The sympathetic nervous system is made up of pre- and postganglionic neurons and their associated glial cells: satellite glial cells, which ensheathe cell bodies, and

Schwann cells, which wrap around axonal projections. Preganglionic neuronal cell bodies are located in the thoracic and lumbar spinal cord, and make short projections to postganglionic neurons, which are clustered into discrete ganglia that lie parallel to the spinal cord along the length of the , forming the sympathetic chain. From the postganglion, neurons project millimeters to meters away to innervate distal target tissues such as the heart, salivary glands, and .

The sympathetic nervous system is globally responsible for organ tissue homeostasis throughout the body, as well as controlling the “fight or flight” response. In

2 dangerous or stressful situations, the brain sends a signal to preganglionic neurons in the spinal cord, which release the into synapses on postganglionic sympathetic neurons. Rapid activation of sympathetic neurons results in the release of at distal tissues, which acts through adrenergic receptors to regulate tissue function such as increasing heart rate, salivation, or sweat (McCorry

2007). The effects of norepinephrine on organ function help prepare the body to respond to the presented challenge, and precise innervation of target organs during development is essential.

Development of the sympathetic nervous system is a highly regulated process and many of the molecular players involved in early sympathetic nervous system formation have been described. Cells of the sympathetic nervous system arise from a common precursor during development, denominated neural crest cells (NCCs). Bone morphogenetic protein (BMP), (Fgf), and Wnt signaling from the induces NCCs to form from the ectoderm along the anterior-posterior axis at the junction between the neural plate and around embryonic day 8 (E8) in the mouse (Garcia-Castro, Marcelle, and Bronner-Fraser 2002; Monsoro-Burq, Fletcher, and

Harland 2003; Liem et al. 1995; Marchant et al. 1998). Early work demonstrated the plethora of cell types that are derived from NCCs. Cell fate tracing using injected dyes, or later on using genetic recombination, found that NCC’s represent a broad category of cells that give rise to the peripheral nervous system as well as bone, , cells, and (Huang and Saint-Jeannet 2004). While all neural crest cells initially express genes such as Sox10, they are categorized by their location along the anterior-posterior axis and the cell types they give rise to. There are cranial, cardiac,

3 vagral and sacral, and trunk NCCs. It is trunk neural crest cells which give rise to the sympathetic nervous system (Le Douarin and Teillet 1973). These cells, which lie along the anterior axis, delaminate from the neuroepithelium and migrate between the somites to their final destination (Rickmann, Fawcett, and Keynes 1985; Erickson, Loring, and

Lester 1989). In order for NCCs to delaminate and migrate they undergo an epithelial-to- mesenchymal (EMT) transition, which is, in part, achieved through the simultaneous activation of genes such as Sox9, Snai2, and FoxD3 (Teng et al. 2008; Cheung et al.

2005).

Trunk NCCs delaminate in three waves. The first and intermediate waves of NCC delamination give rise to cells of the dorsal root ganglia (DRG) and sympathetic chain, as well as chromaffin cells of the adrenal medulla (Loring and Erickson 1987). Cells of the first and intermediate waves migrate ventrally between the somites or into the sclerotome of the somite, respectively. In the intermediate wave, cells that stay in the sclerotome become part of the DRG while those that continue to migrate out give rise to sympathetic precursors (Serbedzija, Fraser, and Bronner-Fraser 1990). A series of attractive and repulsive cues signal to the cells destined to become the sympathetic chain to migrate towards the dorsal . Once at the dorsal aorta, then direct certain cells to begin to form the sympathetic chain.

Studies in mouse, rat, xenopus, and chick have elucidated many of the early factors involved in sympathetic chain formation. As the NCCs exit the neural tube,

Ephrin and Semaphorin signaling direct migration through the somites. Neuropilin1-

Semaphorin3a (Nrp1-Sema3a) signaling acts as a repulsive cue for migrating NCCs, which go on to form the sympathetic chain. Knockouts of either Sema3a, or the receptor

4 Nrp1, exhibit ectopic localization of the sympathetic chain (Kawasaki et al. 2002;

Schwarz, Maden, Davidson, et al. 2009; Schwarz, Maden, Vieira, et al. 2009). As the

NCCs migrate away from the sclerotome of the somite and get closer to the dorsal aorta,

Neuregulin 1 (Nrg1)-ErbB2/3 signaling starts to act as an attractive cue. Bone morphogenetic from the dorsal aorta induce expression of Nrg1 in the around the dorsal aorta, which acts as a chemoattractant for ErbB2/3- expressing NCCs (Saito et al. 2012). In mice where Nrg1, ErbB2, or ErbB3 are knocked out, ventral migration of NCCs is arrested, and in the ErbB3 knockout, the sympathetic chain is smaller in size (Britsch et al. 1998).

BMPs from the dorsal aorta additionally play a role in inducing formation from NCCs. BMPs -2, -4, and -7 have all been shown to induce sympathetic neuroblast formation through activation of transcription factors that provoke expression of characteristic neuronal markers (Shah, Groves, and Anderson 1996; Reissmann et al.

1996; Varley and Maxwell 1996; Schneider et al. 1999). Among the transcription factors turned on as a result of BMP signaling is paired-like homeobox 2B (Phox2b), BHLH transcription factor 1 (Ascl1), and heart and neural crest derivatives-expressed protein 2

(Hand2). Of these transcription factors, Phox2b is essential for sympathetic neuron differentiation. Global knockout of Phox2b results in loss of sympathetic progenitors and a failure to express other key transcriptions factors like Hand2 (Pattyn et al. 1999). In conditional knockouts where Phox2b is ablated directly following differentiation of , sympathetic neurons do not acquire expression of the adrenergic phenotype demonstrating that Phox2b regulates the expression of key in the noradrenaline biosynthetic pathway: tyrosine hydroxylase (TH) and dopamine ß-hydroxylase (DBH)

5 (Coppola et al. 2010). Phox2b regulates expression of TH and DBH both directly and indirectly through activation of other transcription factors. For example, Hand2, which lies downstream of Phox2b signaling, is also capable of controlling TH and DBH expression (Hendershot et al. 2008; Morikawa et al. 2007). As TH and DBH are key enzymes in the synthesis of norepinephrine, the main neurotransmitter of sympathetic neurons, expression of these two proteins is essential for sympathetic neuron function.

Following acquisition of the sympathetic neuron phenotype, the neurons must then extend axonal processes to innervate target tissues. Because the sympathetic neuron cell bodies are clustered into ganglia along the length of the spinal cord, axons must grow great distances in order to reach their peripheral targets. This process of growth and branching at distal organs has been heavily studied, and many of the crucial players involved have been elucidated over the years.

Neurotrophin Signaling in Sympathetic Neuron Development

Following differentiation of NCCs into sympathetic neuroblasts, a change in must be established in order for a single process to grow out of the ganglion as the axon. The players involved in the initiation of axon development have yet to be fully understood, though it is hypothesized that both autocrine and paracrine signaling may play a role. For example, autocrine hepatocyte growth factor (HGF) signaling has been implicated as one of the players in this process. Sympathetic neurons express both HGF and its receptor, the Met tyrosine , and application of HGF to cultures of sympathetic neurons induces axon outgrowth while inhibition of endogenous HGF via application in vitro prevents growth (Yang et al. 1998).

6 Following axon initiation, growth and survival of sympathetic neurons is largely dependent on a peripheral growth factor known as nerve growth factor (NGF). In 1956,

Drs. Rita Levi-Montalcini and Stanley Cohen first isolated NGF as a peptide able to support sympathetic and survival in vitro (Levi-Montalcini and Cohen

1956). It was later discovered that NGF controlled several aspects of neuronal development in the sympathetic nervous system including survival, axon growth, final target innervation, and establishment of synapses.

NGF became the prototypical member of a family of growth factors known as . Other family members include neurotrophin-3 (NT3), neurotrophin-4

(NT4), and brain-derived neurotrophic growth factor (BDNF). These growth factors are expressed and secreted by tissues targeted for innervation by neurons, and globally control growth and survival mechanisms. Neurotrophins bind with high affinity to cognate receptors of the kinase family: TrkA (NGF and NT3), TrkB

(BDNF), and TrkC (NT3/4); as well as a low affinity receptor, p75NTR (all) (Huang and

Reichardt 2003). In the sympathetic nervous system, both NT3 and NGF act to control neuron growth and survival.

Following initiation, axons grow along the vasculature towards peripheral target tissues. Extension of axons along the vasculature is mediated by several , including artemin and the neurotrophin NT3. Both artemin and NT3 are expressed by vessels and promote axon growth of sympathetic neurons in vitro. In knockout models of artemin or its receptors, Ret and GFR3, axon growth is stunted and misdirected while in NT3-/- mice axon extension along the vasculature is markedly reduced (Enomoto et al. 2001; Honma et al. 2002; Yan, Newgreen, and Young 2003;

7 Kuruvilla et al. 2004). However, some axons still reach peripheral target organs in each of these knockout models, suggesting that a combination of factors contribute to axon growth along the vasculature, and some of these factors may not be yet elucidated.

The contribution of NT3 to proximal axon growth creates a unique setup for sympathetic neuron development. Though the primary receptor for NT3 is TrkC, it also binds to TrkA. Indeed, TrkC expression is extremely low in sympathetic neurons and

NT3 activation of TrkA in sympathetic neurons is capable of promoting survival and neuritogenesis (Fagan et al. 1996; Belliveau et al. 1997). However, TrkA is the primary receptor and has a higher affinity for NGF (Chao 2003). As the axon grows along the vasculature, the first comes into contact with secreted NT3 molecules, but as the growth cone nears target tissues it comes into contact with secreted NGF to control final target innervation (Glebova and Ginty 2004). Thus, the activation of TrkA by both

NT3 and NGF contributes to overall axon growth during distinct points in development.

Additionally, although both NT3 and NGF bind to TrkA, the means by which they achieve axon growth and final target innervation, respectively, are likely distinct

(Belliveau et al. 1997). While the mechanisms governing NT3 activation of TrkA and proximal axon extension remain largely undefined, the mechanisms governing NGF have been deeply studied.

At the growth cone, NGF dimerizes, binds TrkA, and induces autophosphorylation of two TrkA monomers in order to trigger a series of downstream signaling pathways including MAPK, PI3K, and Ras. Immediate local changes in the axonal growth cone are in part due to activation of PI3K and inhibition of GSK-3β, which together regulate the plus-end binding protein APC (Zhou et al. 2004).

8 However, unlike NT3, when NGF binds TrkA, one of the signaling cascades triggered results in internalization of the ligand-receptor complex into a signaling endosome that is then transported to the soma where it promotes survival, extension, and formation (Kuruvilla et al. 2004). In addition, NGF-TrkA activates transcription factors such as CREB to control transcriptional events in the developing neuron.

Several hundred potential downstream transcriptional targets were recently identified through microarray analysis (Deppmann et al. 2008), however the roles of many of these genes in sympathetic nervous system development have yet to be described. Given the vast number of genes identified, one intriguing possibility is that

NGF-TrkA signaling may participate in other aspects of sympathetic ganglia development that have yet to be illuminated.

Glial cell specification in the sympathetic nervous system

While neurons are maturing and extending their axons, the two glial cell types in the sympathetic nervous system begin to develop. Of the two primary peripheral glial cell types, the general course of development has been well characterized.

Mature Schwann cells exist in two forms, myelinating and non-myelinating. In general myelinating Schwann cells are found around large diameter sensory axons while non- myelinating Schwann cells predominate around small diameter autonomic axons. The progression from neural crest cell to Schwann cell has been characterized, though many of the molecular players involved are still unknown. The generation of Schwann cells follows a three-step cellular transition model from neural crest cells to Schwann cell precursors to immature Schwann cells and finally to mature myelinating or non-

9 myelinating Schwann cells (Kettenmann, Kettenmann, and Ransom 2013). Schwann cell precursors first appear around E12 in the mouse sciatic nerve and are associated with axonal projections. As the axon extends, Schwann cell precursors differentiate into immature Schwann cells (E15/16). Around birth some Schwann cells undergo a process called radial sorting in which they form a 1:1 relationship with large diameter axons to later adopt the mature myelinating Schwann cell phenotypes (Jessen and Mirsky 2005).

Other immature Schwann cells form pockets around small diameter axons to create

Remak bundles, which can first be seen in the rat about three weeks after birth.

Signals originating from the neurons are vital in the development of Schwann cells. For example, axonally expressed Nrg1 is essential for the survival and migration of

Schwann cell precursors (Dong et al. 1995). These cells are lost in mouse knockout models of Nrg1 or its receptors, ErbB2 and ErbB3 (Riethmacher et al. 1997; Morris et al.

1999; Lyons et al. 2005). Additionally, Schwann cell precursors cannot survive in vitro without contact from axons and this contact is mediated through Nrg1 (Brennan et al.

2000). Signaling through Nrg1 to Schwann cell precursors may also be accomplished through activation of Notch, which assists in the survival effects of Nrg1 and interacts with Nrg1 during development, and helps push Schwann cell precursors into the immature Schwann cell fate (Schmid et al. 2003; Morrison et al. 2000). In vitro activation of Notch also pushes neural crest cells into a Schwann cell fate and Notch has long been implicated in neuron versus glial cell fate in the CNS (Wakamatsu, Maynard, and Weston

2000; Wang and Barres 2000). Whether these signals work together or concomitantly in

Schwann cell development remains to be elucidated, but it is clear that signals from axons are essential for Schwann cell development.

10 In contrast to Schwann cells, much less is known about the development of satellite glial cells. Following neuronal specification and initial axon extension, the other primary cell type within the sympathetic ganglia, the satellite glial cell, begins to develop.

Few studies have delved into the specifics of how this mysterious cell type arises. In the rat sympathetic nervous system, there is a period of rapid cell division amongst non- neuronal cells in the ganglia between E16.5 and E18.5 (about E14.5-E16.5 in the mouse).

During this period of cell division, the non-neuronal cells cluster into groups along the periphery of the ganglia, which then begin to disperse and migrate into the ganglia around birth (Hall and Landis 1992). Over the course of early postnatal development, these satellite glial cells contact and wrap around individual neurons. However, the exact timing of, and the mechanisms, governing ensheathment, as well as how the glial cell changes molecularly and morphologically during this development are all still open questions.

Neuron-glial cell interactions during central nervous system development

Similar to the peripheral nervous system, neurogenesis precedes glial cell development in the CNS, and in the case of (one of the primary glial cell types in the CNS), newly generated neurons have been shown to influence their development.

For example, activation of astrogenesis is in part achieved through extrinsic signaling by the IL-6 family of . Mice lacking the IL-6 family ligands, LIFRβ or gp130, in neural stem cells (NSCs, the primary progenitor population in the developing brain) have deficits in astrogenesis (Bugga et al. 1998; Nakashima et al. 1999; Koblar et al. 1998). Evidence suggests that activation of these ligands in vivo is achieved through

11 of the IL-6 cytokine, cardiotrophin-1, from newly born cortical neurons

(Barnabe-Heider et al. 2005). In addition to extrinsically secreted factors, cell contact molecules such as Notch also influence astrogenesis. In the telencephalon, committed

NSCs and newly born neurons express Notch ligands that activate Notch on neighboring

NSCs to confer an astrocytic fate potential through induction of the transcription factor nuclear factor I (Namihira et al. 2009).

Cell contact between astrocytes and neurons also influences astrocyte morphology. In the absence of neurons, cultured astrocytes from the early postnatal mouse exhibit a flattened shape, however, co-culturing of neurons and astrocytes induces the highly complex branched morphology typical of young astrocytes in vivo (Hatten 1985). More recent evidence additionally finds that expression of members from the neuroligin (NL) family of cell adhesion molecules in mouse cortical astrocytes (NL1, NL2, and NL3) mediates astrocyte morphology through contact with neuronal (Stogsdill et al. 2017).

Cell-cell contact is a two-way street, and contact signaling between astrocytes and neurons can also influence neuronal morphology. Most notably, astrocytes appear to play a key role in . Studies have found that cortical and retinal neurons make few synapses naturally in vitro, however, addition of astrocytes or astrocyte conditioned media to the culture system markedly increases the number of synapses (Ullian et al.

2001). This astrocytic control of synaptogenesis is achieved through both secreted and contact-dependent mechanisms (Baldwin and Eroglu 2017).

In addition to astrocyte development, neurons in the CNS also dictate development. are analogous to Schwann cells in the

12 PNS in their role of myelination of axons, however, they are morphologically and molecularly distinct. Despite these differences, oligodendrocytes also share similar mechanisms with Schwann cells for their development. For example, axonal contact with oligodendrocytes is vital for their development and proper ensheathment of processes around axons. Similar to Schwann cells, ErbB2 and Notch signaling play primary roles in oligodendrocyte specification and terminal differentiation around axons, respectively, and the ligands for these two receptors are expressed by neurons (Kim et al. 2003; Hu et al.

2003; Wang et al. 1998). Thus, during development of the CNS, cell contact is important for both neuron and glial cell maturation.

Is cell-cell contact important for the development of satellite glial cells?

Contact with axons is essential for Schwann cell development in the PNS and oligodendrocyte development in the CNS, and astrocyte development is in part achieved through contact with neurons. Given the importance of cell contact signaling between neurons and numerous types of glial cells in development, and the close association between satellite glial cells and sympathetic neurons, it stands to reason that contact with the neurons would be important for satellite glial cell development as well.

Though we don’t know much about the molecular profile of satellite glial cells, we can begin to parse out some of the players that may be involved in their development by using our vast knowledge of peripheral neurons. The timing of satellite glial cell specification coincides with two major events in sympathetic ganglia development: the time in which sympathetic neurons begin to receive preganglionic input from the spinal cord, and when their axonal processes have extended towards their peripheral targets

13 (Hall and Landis 1992). This observation brings up two intriguing hypotheses. The first hypothesis is that neuronal activity may modulate satellite glial cell development. Indeed, neuronal activity has been implicated in the development of glial cells in the central nervous system. For example, activity of neurons can influence oligodendrocyte differentiation and myelination via secreted factors such as glutamate and ATP (Wake,

Lee, and Fields 2011). Neuronal activity can also alter astrocyte and as well as astrocytic coverage of synapses where increased activity is correlated with increased astrocytic coverage (Farhy-Tselnicker and Allen 2018; Wake,

Lee, and Fields 2011; Hasel et al. 2017).

The second hypothesis is that a peripheral signal may regulate satellite glial cell development. At peripheral target tissues, sympathetic neurons come into contact with secreted NGF molecules during this time in development, which regulates gene transcription within the neurons, as discussed earlier. Previous work has shown that NGF can regulate the myelination of axons in both the PNS and CNS wherein NGF promotes

Schwann cell myelination around TrkA-expressing DRG neurons, and inhibits oligodendrocyte myelination through regulating the expression of LINGO-1 (Lee et al.

2007; Chan et al. 2004). We find that when TrkA is knocked out in sympathetic neurons there are defects in satellite glial cell ensheathment around neurons in sympathetic ganglia (Figure 2.4a,b). Therefore, we investigated the possibility that NGF-TrkA signaling in sympathetic neurons might regulate satellite glial cell development.

Here, we describe a pathway in which a peripheral signal, NGF, controls expression of a gene in neurons, Delta/Notch-like EGF-related Receptor (DNER), that instructs development of a distal cell type, satellite glial cells. We also reveal roles for

14 satellite glial cells in shaping sympathetic neuron morphology during development. We find that DNER is specifically expressed in neurons within sympathetic ganglia and is regulated by NGF-TrkA signaling. Conditional deletion of DNER from sympathetic neurons results in disrupted satellite glial cell contact with neurons with no overall loss of cell numbers within the ganglia. Furthermore, we find alterations in neuronal morphology. DNER knockout sympathetic neurons are smaller but have increased axonal innervation of target tissues. These morphological changes are accompanied by a downregulation of major enzymes in the norepinephrine biosynthetic pathway, TH and

DBH, in the cell bodies with no apparent changes in expression in axons. In addition to morphological changes, DNER knockout neurons exhibit a sustained calcium response to depolarization, which could explain the increase in target tissue innervation. We also find that ablation of satellite glial cells during development recapitulates some of the neuronal phenotypes observed in DNER knockout mice. Sympathetic neurons in glia-ablated mice also have decreased some size and downregulate expression of TH and DBH. These results suggest a tripartite system in which secretion of a growth factor from peripheral target tissues influences both neuron and glial cell development.

15 Chapter Two: Regulation and Expression of

DNER in the Sympathetic Nervous System

16

INTRODUCTION

NGF-TrkA signaling controls many aspects of sympathetic ganglia development, including regulating the expression of several hundred target genes in late embryonic and early postnatal development. Some of these genes assist in various aspects of neuronal development, such as axon branching, dendrite growth, and synapse formation, but the roles of many of these genes are still unknown. One gene called DNER, or Delta/Notch- like EGF-related Receptor, was pulled out of a microarray from the sympathetic ganglia of neonatal mice lacking NGF (Deppmann et al. 2008). DNER has been implicated as a player in several aspects of nervous system development from neuron and glial cell differentiation to neuronal neurite extension and glial cell maturation. However, its role in the sympathetic nervous system is unknown.

DNER is a transmembrane protein that with an extracellular domain composed of ten epidermal growth factor (EGF)-like repeat-containing domains which are highly homologous to Notch and its receptors, Delta1 and Jagged1. The 6th EGF-like repeat shares 60% homology with mouse Notch1, 60% homology with human Delta1, and 54% homology with mouse Jagged1. Similar to Delta1 and Jagged1, the majority of the DNER protein (737 amino ) exists in the extracellular domain with a 30 amino transmembrane domain and 70 amino acid intracellular domain (Eiraku et al. 2002).

Unlike Delta1 and Jagged1, the extracellular domain of DNER does not have a DSL

(Delta, Serrate, and Lag2) domain, which is necessary for Notch binding by DSL ligands

(D'Souza, Meloty-Kapella, and Weinmaster 2010). Yet despite this, the similarities between DNER, and Notch and its receptors are striking and as such, DNER has been

17 proposed to be a Notch ligand (Eiraku et al. 2005). Indeed, DNER is able to bind to

Notch1, and the first two EGF-like repeats of DNER’s extracellular domain are required for this interaction (Eiraku et al. 2005).

Activation of Notch by DSL-ligands triggers a canonical pathway in which the intracellular domain of Notch is cleaved by a gamma-secretase and translocated to the . In the nucleus the Notch intracellular domain binds to the CBF1–Suppressor of

Hairless–LAG1 (CSL) DNA binding complex and induces transcription of genes such as those of the Hairy and Enhancer of split (Hes) and Hes Related Family BHLH

Transcription Factor With YRPW Motif (Hey) families. Activation of Notch by other ligands can trigger non-canonical pathways, which are not fully understood but one of which involves the binding of the Notch intracellular domain to a protein called Deltex in order to sequester it in the , possibly by promoting binding to β-arrestin and thus activating a separate pathway (D'Souza, Meloty-Kapella, and Weinmaster 2010;

Mukherjee et al. 2005). DNER has been shown to activate both canonical and non- canonical Notch pathways (Du et al. 2018; Eiraku et al. 2005; Wang et al. 2019; Hsieh et al. 2013).

Since DNER was first described in 2002, studies have found several roles for the gene in nervous system development. For example, in global DNER knockout mice

Bergmann glial cell maturation was partially disrupted. DNER knockout Bergmann glial cell bodies were mis-localized in the molecular layer of the cerebellum and they extended fewer radial processes than their wildtype counterparts in the early postnatal brain. This process was dependent on non-canonical Notch signaling through the Deltex pathway

(Eiraku et al. 2005). Additionally, morpholino knockdown of DNER in

18 inhibited both neuronal and glial cell differentiation in a manner dependent on Deltex1.

However, zebrafish DNER was also found to inhibit neuronal progenitor proliferation in a manner independent of Notch (Hsieh et al. 2013), suggesting that DNER may act through multiple receptors. These studies indicate the role of DNER may differ in an and region-dependent manner and may be involved in both neuron and glial cell development.

In vitro studies have suggested that DNER may also play a role in neuritogenesis

(Fukazawa et al. 2008; Du et al. 2018). The localization of DNER to the plasma membrane promotes neurite extension in Purkinje and Neuro-2A cells (Fukazawa et al.

2008). Similarly, knockdown of DNER in cultures of neurons results in disrupted cell polarity and a decrease in dendritic length (Du et al. 2018). These studies demonstrate that the expression of DNER, specifically at the plasma membrane, positively influences neurite extension, and suggest that the regulation of DNER expression and localization may be important in its function.

Several studies have delved into DNER expression and localization primarily in the central nervous system where DNER is strongly expressed in cerebellar granule cells,

Purkinje cells, cortical and hippocampal pyramidal neurons, spiral and vestibular ganglion neurons, and in auditory and vestibular hair cells of the (Eiraku et al.

2002; Hartman et al. 2010). In hippocampal neuron and cultures DNER localizes in a somato-dendritic manner to the plasma membrane and within endosomes

(Eiraku et al. 2002). Targeting of DNER to dendrites is dependent on a tyrosine-based sorting motif on its intracellular C-terminal domain and seems to be directed by the coat- associated protein complex AP-1 (Eiraku et al. 2002). DNER lacking the last 16 amino

19 acids in its C-terminal domain exhibits increased surface expression as well as an increase in neurite extension (Fukazawa et al. 2008). Additionally, the C-terminal domain of DNER contains three putative tyrosine phosphorylation sites which are weakly phosphorylated and appear to be dephosphorylated by protein tyrosine phosphatase 

(PTP), which co-immunoprecipitates with DNER, to regulate its localization and function. Addition of pleiotrophin, an inactivator of PTP’s phosphatase activity, to cultured cells increases DNER phosphorylation and similar to cleavage of the C-terminal domain of DNER, increases surface expression and neurite extension (Kurisu et al.

2010). Conversely, disruption of DNER phosphorylation or it’s somato-dendritic membrane targeting results in diminished neurite extension in cultured Purkinje and

Neuro-2A cells (Fukazawa et al. 2008; Kurisu et al. 2010).

DNER is widely expressed and its localization is tightly regulated in neurons of the central nervous system. Disruption of DNER expression or localization affects both neuron and glial cell development in vitro and in vivo, however, DNER expression and localization in the sympathetic nervous system as well as how DNER is regulated has yet to be elucidated. DNER was identified via microarray as being potentially regulated by

NGF-TrkA signaling, which is an important developmental cue for sympathetic ganglia development. Given the known roles for DNER in central nervous system development, in both neuritogenesis and glial cell development, we wanted to investigate the role of

DNER in sympathetic ganglia development as a potential involved in neuron- satellite glial cell contact. Here, we demonstrate that DNER is expressed in developing and adult sympathetic neurons. In contrast to the central nervous system where DNER localizes in a somatodendritic manner, DNER localizes to the soma and axons in

20 sympathetic neurons. We find that DNER expression is largely cytoplasmic with only a small portion of DNER existing on the plasma membrane at any given time. We found that the half-life of DNER in sympathetic neurons is approximately 4 hours, and this short lifetime could contribute to reduced localization on the cell surface. Additionally, we demonstrate that DNER expression is regulated by the peripheral growth factor NGF in sympathetic neurons. Interestingly, we also find that loss of the NGF receptor, TrkA, in sympathetic neurons perturbs satellite glial cell ensheathment. The timing of DNER expression in sympathetic neurons, the regulation of DNER by NGF-TrkA, the expression of DNER on the plasma membrane, and the previously observed roles for

DNER in the CNS suggest that DNER may play a role in sympathetic ganglia development, specifically in regulation of neuron-glial cell contacts.

21 RESULTS

DNER is expressed in sympathetic neurons, but absent from satellite glial cells

DNER is thought to be a neuronal-specific non-canonical Notch ligand (Eiraku et al. 2002; Eiraku et al. 2005). It is highly expressed in neurons of the developing central nervous system and plays roles in neuron and glial cell development in mice and zebrafish. As the central and peripheral nervous systems share many of the same developmental cues, it stands to reason that DNER would also be present in peripheral neurons, but expression in this system has not yet been analyzed. Therefore, we first asked what DNER expression looks like in the sympathetic nervous system. Given

DNER’s role in development in the mouse cerebellum and the fact that DNER was pulled out of a screen from neonatal sympathetic ganglia, we asked if DNER was expressed during sympathetic nervous system development. To this end, we employed for DNER in the developing superior (SCG). DNER expression was first identified at E13.5, following neuron specification, and continued throughout development and into adulthood (Figure 2.1a), though expression of DNER appeared to peak during late embryogenesis.

Sympathetic ganglia are made up primarily by two cell types – sympathetic neurons and their surrounding satellite glial cells. Previous work in the central nervous system found DNER expression specifically in neurons where in the cerebellum DNER expression was strongly localized to Purkinje cells and absent from neighboring

Bergmann glial cells (Eiraku et al. 2002). Therefore, we asked if DNER was also only expressed in neurons of sympathetic ganglia. To determine cell-type specific expression we cultured neurons or satellite glial cells from P0.5 rat SCG and performed RT-PCR

22 (Figure 2.1b,c). DNER mRNA was detected only in sympathetic neuron cultures, and absent from satellite glial cells in agreement with what was found in the central nervous system. At the protein level, DNER was again found to localize within neuron cell bodies and axons in vivo and in vitro by immunohistochemistry (IHC) (Figure 2.1d-f).

Interestingly, DNER expression was localized in punctae to both the soma and in vitro and specifically was found in axons innervating the salivary glands. This is in contrast to hippocampal and Purkinje neurons where DNER was found strictly in the somato-dendritic compartments (Eiraku et al. 2002; Kurisu et al. 2010) and suggests

DNER localization is differentially regulated in the central and peripheral nervous systems.

As DNER is a transmembrane protein, we asked if it localized to the cell surface.

To address this question, we used a membrane-impermeable biotin to label the cell surface of cultured sympathetic neurons and then used Alexafluor-488-conjugated streptavidin and DNER to visualize DNER on the neuronal plasma membrane.

Co-localization analysis revealed a small percentage of DNER localized to the cell surface, while the rest appeared in punctae or perinuclear patches throughout the cytoplasm (Figure 2.2a-b). Indeed, studies in the central nervous system found the majority of DNER localized to endosomes with only a small percentage appearing on the surface at any given time (Kurisu et al. 2010). The localization of DNER to endosomes and perinuclear patches suggests that the protein is highly processed and may be rapidly turned over. In accordance with this observation, we used cycloheximide (CHX) to block translation in cultured sympathetic neurons for 0 to 4 hours and assessed DNER expression by western blot. We found that DNER has a relatively short half-life of

23 around 4 hours (Figure 2.2c-d). This rate was not affected by the presence of the growth factor NGF and correlates with a recent report that DNER protein is quickly turned over

(Yap et al. 2018). Together, these results highlight DNER as a rapidly turned over protein that localizes to all three major compartments of developing sympathetic neurons.

DNER is regulated by NGF-TrkA signaling

Given the onset of DNER expression during sympathetic nervous system development, and the marked downregulation of DNER in mice lacking NGF by microarray analysis (Deppmann et al. 2008), we directly assessed whether DNER expression was regulated by NGF-TrkA signaling. We first tested the necessity of NGF-

TrkA signaling for DNER expression using a conditional mouse knockout line in which the receptor, TrkA, was specifically deleted in sympathetic neurons. As NGF-TrkA is necessary for neuronal survival, we concomitantly deleted the pro-apoptotic factor, Bax, to generate TH-Cre;TrkAf/f;Bax-/- (TrkA;Bax dKO) mice. In this way, we depleted sympathetic neurons of the NGF-TrkA signal without affecting neuronal survival.

Strikingly, DNER expression was diminished in TrkA;Bax dKO sympathetic ganglia by western blot and IHC analysis (Figure 2.3a-d).

To then ask if NGF was sufficient to induce DNER expression, we cultured sympathetic neurons from P0.5 rats for 6-7 days in the presence of NGF to allow for axon growth, removed NGF for 48 hours, and either kept neurons depleted of NGF or added

NGF back for 16 hours, then assessed DNER transcript levels by qPCR. We found a two- fold increase in DNER expression in the presence of NGF (Figure 2.3e). Taken together,

24 these data indicate DNER, expressed in sympathetic neurons, is regulated by peripherally derived NGF.

Loss of TrkA in sympathetic neurons disrupts satellite glial cell ensheathment

Previous studies have implicated DNER in Bergmann glial cell development, so we hypothesized DNER may play a role in satellite glial cell development in the sympathetic nervous system. Since DNER is regulated by NGF-TrkA signaling, we first wanted to know if NGF-TrkA signaling in sympathetic ganglia plays a role in satellite glial cell development. To this end we used immunohistochemistry for tyrosine hydroxylase (TH) to mark sympathetic neurons and brain lipid binding protein (BLBP) to mark satellite glial cells. In the mouse sympathetic nervous system, BLBP-positive glial cells first appear at embryonic day 11 (E11). During , BLBP labels the progenitor of the two glial cell types in the peripheral nervous system, satellite glial cells and Schwann cells. However, BLBP expression is lost in mature Schwann cells and is maintained in satellite glial cells (Kurtz et al. 1994; Shi et al.

2008). Therefore, we used BLBP expression to assess satellite glial cell morphology in mice lacking TrkA at postnatal day 14 (P14). In control sympathetic ganglia, we found rings of satellite glial cells around individual neurons (Figure 2.4a). However, in TrkA knockout sympathetic ganglia, we found disruptions in the ring-like structures of the satellite glial cell sheath (Figure 2.4b) indicating that satellite glial cell ensheathment around sympathetic neurons is perturbed with loss of NGF-TrkA signaling.

25

26 Figure 2.1. DNER is expressed in all three major compartments of sympathetic neurons.

(a) Developmental time-course of DNER expression by in situ hybridization in wildtype mouse SCG shows DNER is expressed as early as E13.5 and continues into adulthood

(scale bar: 50um). (b) RT-PCR for DNER shows expression specifically in sympathetic neurons and absent from glial cells. (c) Cultured sympathetic neuron (green) and (d) satellite glial cell (blue) show DNER (magenta) expression in neuronal cell body and axon while absent from satellite glial cell. (e) DNER (green) expression in sympathetic neuron cell bodies (magenta) of P7 TH-Cre;TdTomatof/+ SCG (scale bar: 25um, inset scale bar: 10um). DNER is robustly expressed in a puntate pattern in the cytoplasm and perinuclear region within the neuron cell body. (f) Salivary glands from P7 TH-

Cre;TdTomatof/+ shows DNER (green) expression in sympathetic axons (magenta) (scale bar: 50um).

27

Figure 2.2. DNER has a half-life around 4 hours and partially localizes to the cell surface membrane.

(a,b) Cell surface biotinylation (magenta) shows about 20 percent of DNER (green) co- localizes to the neuronal (scale bar: 10um). (c) Western blot analysis of

DNER in neuron cultures treated with cycloheximide (CHX) for 0, 2, and 4 hours shows

DNER has a half-life around 4 hours that is unaffected by the presence of NGF. (d)

Quantification of western blots for DNER expression from cycloheximide treated neuron cultures (n=3 individual experiments).

28

Figure 2.3. DNER is regulated by NGF-TrkA signaling.

(a,b) DNER (white) expression is greatly reduced in TH-Cre;TrkAf/f;Bax-/- SCGs (TH: magenta, scale bar: 50um). (c,d) Western blot analysis shows DNER expression is diminished in TH-Cre;TrkAf/f;Bax-/- sympathetic ganglia (n=3 control, 3 TrkA;Bax dKO, p<0.01). (e) Addition of NGF to sympathetic neuron cultures increases DNER transcript levels (n=4, p<0.01).

29

Figure 2.4. Disruption of satellite glial cell sheaths in TrkA knockout sympathetic ganglia. (a) Immunohistochemistry for neurons (TH: green) and satellite glial cells

(BLBP: magenta in merge, black in inverted) from 1 month old TrkAf/f;Bax+/- and (b) TH-

Cre;TrkAf/f;Bax+/- sympathetic ganglia.

30 DISCUSSION

DNER was initially discovered as a transmembrane protein specifically expressed in neurons of the central nervous system. Our data demonstrates DNER is also expressed in sympathetic neurons of the peripheral nervous system. DNER largely exists in a punctate pattern with only a fraction of DNER protein appearing on the cell surface. Our experiments suggest that DNER is a rapidly turned over protein with a half-life of around

4 hours. DNER is a proposed Notch ligand. When canonical ligands like Delta and

Jagged bind Notch, they are known to internalize with the extracellular domain of Notch in order to be degraded (Ables et al. 2011). If DNER is indeed a Notch ligand, it is possible that DNER is also internalized and degraded upon binding to Notch, and thus for continued signaling, the protein must be continually translated and processed.

While DNER expression is restricted to the somato-dendritic compartment of neurons in the central nervous system, in sympathetic ganglia, DNER is also expressed in axons. This differential expression could be due to the presence of separate isoforms in the CNS versus PNS, or differential post-translational modifications of the protein. In support of these hypotheses, it was found that deletion of the intracellular domain of

DNER resulted in loss of compartmentalized sorting of the protein in cultured hippocampal neurons (Eiraku et al. 2002). Additionally, the C-terminal tail of DNER contains a tyrosine-based sorting motif (YXXO), which plays a role in directing DNER localization in hippocampal neuron cultures (Kurisu et al. 2010). Future experiments comparing DNER in sympathetic and hippocampal neurons would help to determine if

DNER exists in multiple isoforms or if the phosphorylation state of the DNER is

31 differentially regulated in central versus peripheral neurons and could help to address how DNER achieves distinct localization patterns.

Intriguingly, we found that DNER expression in sympathetic neurons is regulated by NGF-TrkA signaling. This is the first study to identify how DNER expression is regulated in neurons and raises the question of what the role of DNER is in sympathetic neurons. NGF-TrkA is a developmental signal turned on in post-mitotic neurons, and

DNER expression is observed simultaneous with the onset of this signal, which suggests that DNER may play a role in sympathetic ganglia development. While DNER is expressed specifically in the neurons, it is a transmembrane protein and likely signals to neighboring cells. Sympathetic glial cell development coincides with the onset of DNER expression, and DNER has been shown to instruct both neuron and glial cell maturation and differentiation in other systems (Du et al. 2018; Eiraku et al. 2005; Hsieh et al. 2013).

It therefore seems likely that DNER may play a similar role in sympathetic nervous system development. Indeed, we found that loss of TrkA in sympathetic neurons perturbs satellite glial cell ensheathment, and so we hypothesize this may be, in part, due to the loss of DNER and explore this possibility in the next chapter.

32

MATERIALS AND METHODS

Animal Husbandry

All animal procedures were performed in accordance with NIH and Johns Hopkins

University Animal Care and Use Committee (JHU ACUC) guidelines. Animals were kept in a 12-hour light:dark cycle with food and water ad libitum. Mice were kept in mixed

C57BL/6 and 129P backgrounds and animals of both sexes were used for analyses. TH-

Cre and TrkAF592A (TrkAf/f) were gifts from Dr. C. Gerfen (NIH), and Dr. D. Ginty

(Harvard ) respectively. Bax-/- mice were obtained from Jackson

Laboratory. Mice were crossed to generate TH-Cre;TrkAf/f;Bax-/- (TrkA;Bax dKO) mice.

TH-Cre;Bax-/- or TrkAf/f;Bax-/- mice were used as controls in all experiments. Sprague-

Dawley rats were purchased from Charles River (strain code: 400).

Isolation of sympathetic ganglia

Neurons from superior cervical ganglia (SCG) of Sprague-Dawley rat pups (P0.5) or mice of indicated genotypes and ages were dissected and enzymatically dissociated as previously described (Bodmer et al., 2011). Briefly, pups were anesthetized according to

JHU ACUC protocol. SCGs of pups were dissected and separated from surrounding tissue, incubated for 20 minutes at 37°C in DMEM AIR (DMEM F12, 12.5mM ,

1U/ml penicillin/streptomycin) and supplemented with 4mg/ml collagenase IV, 1mg/ml hyaluronidase, 0.6mg/ml DNase I, and 10mg/ml BSA. Ganglia were centrifuged, resuspended in DMEM air containing 3mg/ml trypsin, and incubated for an additional 30

33 minutes at 37°C. Cells were mechanically dissociated by tituration with a glass pipet, centrifuged, and resuspended in SCG media (DMEM supplemented with 10% FBS and

1U/ml penicillin/streptomycin) for glial cell cultures. Neuronal culture media was additionally supplemented with 100ng/ml NGF and 10mM Ara-C to prevent glial cell proliferation. For mass cultures, SCGs from 5-6 rats or 6-8 mice were plated on - coated 60mm or 35mm dishes, respectively. For sparse cultures, SCGs from 0.5-1 rats or

1-2 mice were plated on coverslips coated with Poly-D- mixture.

In situ hybridization

In situ hybridization was performed with 904-bp digoxigenin-labeled probes spanning 6-12 of mouse DNER. Sample tissues were fresh-frozen in 30% sucrose for a minimum of 3 days at 4°C to allow for cryoprotection, embedded in OCT, and serial sectioned at 12um. Sections were post-fixed in 4% PFA for 20 minutes followed by washes with PBS, and permeabilization with 0.1% TX-100 for 10 minutes. Sections were then washed again with PBS and acetylated with 0.25% acetic anhydride in 0.1M triethanolamine with 0.9% NaCl for 10 minutes. Following acetylation, sections were equilibrated in hybridization solution (50% formamide, 5x Denhardt’s, 250ug/ml MRE

600 tRNA, 5x chloride/sodium citrate (SSC) for 2 hours at room temperature, then hybridized overnight at 68°C in a hybridization oven with 2ug/ml of probe diluted in hybridization solution. Following hybridization, sections were washed with 0.2x SSC 3 for 20 min at 60°C, blocked for 1 hour at room temperature with 10% goat serum in buffer (0.1M Tris pH 7.5, 0.15M NaCl), and treated with anti-DIG ab (1:5000, Roche) overnight at 4°C. The reaction was carried out with NBT/BCIP

34 tablets for 4-6 hours, rinsed in PBS, fixed in 4% PFA, and mounted in AquaMount (EMD

Chemicals). Solutions were treated with diethylpyrocarbonate (DEPC) and autoclaved prior to use to remove Rnases.

Immunohistochemistry and Immunocytochemistry

For IHC, mice at indicated ages were fixed in 4% PFA for 4-16 hours at 4°C, cryoprotected in 30% sucrose in PBS, embedded in OCT, and serially sectioned (12-

30um) with a Thermo Microtom HM 550 cryostat. Sections were blocked with 5% DS serum and 0.1% TX-100 in PBS for 1 hour and incubated in primary antibody overnight at 4°C. Sections were then washed with PBS, incubated with secondary antibody for 1 hour at room temperature, washed again, and mounted with fluorimount plus DAPI.

For ICC, sparse cultures were fixed for 20 minutes with 4% PFA, washed in PBS, and blocked for 1 hour at room temperature with 1% BSA and 0.1%TX-100 in PBS. Primary antibodies were diluted in block and incubated on coverslips for 2 hours at room temperature or overnight at 4°C. Following primary, coverslips were washed with PBS, incubated with secondary antibody for 1 hour at room temperature, washed again, and mounted with fluorimount plus DAPI. Images for IHC and ICC were taken with a Zeiss

LSM 700 confocal .

Cycloheximide on DNER expression

Neurons from P0.5 rat pups were cultured in mass in SCG media plus 100ng NGF and

10mM AraC to rid cultures of glial cells for five days. NGF was then removed by

35 washing cultures with PBS and replacing media with low serum media (DMEM supplemented with 1% FBS, 1U/ml penicillin/streptomycin), Boc-Asp(Ome)-FMK (BAF

1:1000, BPS Bioscience, 27611-1), and sheep anti-NGF (1:1000, AB1542). After 36 hours, cultures were washed and treated with low serum media and 25uM cycloheximide for 2 or 4 hours in the presence of NGF (100ng) or sheep anti-NGF. Protein was collected by applying boiling laemmli buffer (50mM Tris pH 6.8, 0.5% SDS, 10% glycerol, 5% ß- mercaptoethanol, 0.0125% bromophenol blue) directed to neurons and western blot analysis was performed.

RT-PCR and quantitative real-time PCR

For NGF regulation of DNER, neurons from P0.5 rats were cultured in mass (4-6 rat ganglia per dish) and grown in DMEM supplemented with 10% FBS, 1U/ml penicillin/streptomycin, 100ng/ml NGF, and 10mM Ara-C for 6-7 days. Cells were then treated with low serum media (DMEM, 1% FBS, 1U/ml penicillin/streptomycin) supplemented with sheep anti-NGF (1:1000, AB1542), and Boc-Asp(Ome)-FMK (BAF

1:1000, BPS Bioscience, 27611-1). After 36 hours, media was replaced in one dish with low serum media plus NGF and BAF for 16 hours. Cultures were then washed with PBS and 250ul of Trizol was added for RNA extraction. RNA extraction was performed according to manufacturer’s protocol (Invitrogen 15596026) using phenol:.

For cDNA synthesis, 5ug of RNA were reverse transcribed with SuperScript IV Reverse

Transcription following manufacturer’s protocol (Thermo Scientific 18091050). qPCR was performed using Maxima SYBR green/ROX qPCR Master Mix (Thermo Scientific

36 K0222) and detected using Applied Biosystems StepOnePlus Real-Time PCR System

(cat 4376600).

Cell surface biotinylation

Sympathetic neurons from wildtype neonatal mouse pups were sparsely plated for 24 hours on coverslips. Cells were washed with 1x PBS, incubated on ice for 30 minutes with 1mg/mL of EZ-Link Sulfo-NHS-SS-Biotin (Thermo Scientific, 21331), then washed again with ice-cold 1x PBS plus 50mM prior to with 4% PFA for 20 minutes. Cells were immunostained and mounted with fluorimount. Analysis of co- localization of DNER with biotin was done using the FIJI plug-in Coloc-2 for the

Manders coefficient.

Western blot

SCGs from P0.5 mice of indicated genotypes were homogenized in laemmli buffer

(50mM Tris pH 6.8, 0.5% SDS, 10% glycerol, 5% ß-mercaptoethanol, 0.0125% bromophenol blue), run on an 8% SDS-PAGE gel, and protein transferred to a PVDF membrane. The membrane was blocked for 1 hour at room temperature with 5% milk diluted TBS-T buffer. Primary antibodies were diluted in Signal Enhancer HIKARI 250

(Nacalai, NU00102) and incubated overnight at 4C. Antibody solutions were saved and reused up to three times. Membranes were washed with 5% milk followed by TBS-T and then incubated in secondary antibody for 1 hour at room temperature. Membranes were washed again, signal was detected using Pierce ECL Plus Western Blotting

37 (Thermo Scientific, 32132), and imaged using a Typhoon 9410 Variable Mode Imager

(GE Healthcare).

Statistical analyses

Sample sizes were similar to those reported in previous publications (Armstrong et al.

2011; Patel et al. 2015). Data were collected randomly. For practical reasons, analyses of neuronal cell counts in mouse tissues were done in a semi-blinded manner such that the investigator was aware of the genotypes prior to the experiment, but conducted the and data analyses without knowing the genotypes of each sample. All Student’s t tests were performed assuming Gaussian distribution, two-tailed, unpaired, and a confidence interval of 95%. Statistical analyses were based on at least 3 independent experiments and described in the figure legends. All error bars represent the standard error of the mean (s.e.m).

Antibodies

Antibody Company Dilution Use Rabbit anti-BLBP Abcam, AB32423 1:200 IHC Goat anti-DNER Sigma, AF 2254 1:75 IHC 1:1000 Western blot Mouse anti-βIII- Sigma, T6880 1:500 IHC 1:2000 Western blot Mouse anti-tyrosine Sigma, T2928 1:500 IHC hydroxylase Rabbit anti-tyrosine Millipore, AB152 1:500 IHC hydroxylase 1:2000 Western blot Rabbit anti-PI3 Kinase (p85) Millipore, ABS234 1:2000 Western blot Streptavidin Alexafluor 488 Thermo Scientific, S11223 1:500 IHC Donkey anti-rabbit Thermo Scientific, 1:500 IHC Alexafluor488 A21206 Donkey anti-rabbit Thermo Scientific, 1:500 IHC Alexafluor546 A10040

38 Donkey anti-mouse IgG Thermo Scientific, 1:500 IHC Alexafluor647 A31571 Donkey anti-goat Thermo Scientific, 1:500 IHC Alexafluor488 A11055 Donkey anti-goat Thermo Scientific, 1:500 IHC Alexafluor555 A21432 ECL anti-mouse IgG HRP GE Healthcare, NA931V 1:2000 Western blot ECL anti-rabbit IgG HRP Sigma, NA934 1:2000 Western blot Donkey anti-goat HRP Santa Cruz, sc-2020 1:5000 Western blot

39 Chapter Three: Neuronal DNER is required for sympathetic neuron-satellite glial cell contacts

40

INTRODUCTION

The term glial cell encompasses a broad range of non-neuronal cells in the nervous system. Originally thought to simply be just supporting cells for neurons, the term glia was derived from the Greek word for glue. Over a century of work since their discovery in 1856 by Rudolph Virchow has uncovered an immense number of vital roles that glial cells play throughout the nervous system and has further divided the population into subgroups of distinct cell types, including but not limited to astrocytes, oligodendrocytes, , and Schwann cells. Studies have delved deeply into the functions and unique properties of these different glial cell populations, such as controlling the extracellular microenvironment, myelination, and providing support and protection to neurons. The vast majority of studies have focused on glial cells of the central nervous system – astrocytes, oligodendrocytes, and microglia – while glial cells of the peripheral nervous system have been less well studied. In the peripheral nervous system, the two glial cell types that predominate are Schwann cells and satellite glial cells. Schwann cells have been the primary focus of glial cell studies in the PNS for their roles in axon regeneration, myelination, and propagation (Kettenmann,

Kettenmann, and Ransom 2013). In contrast, very little is known about satellite glial cells.

Satellite glial cells are a loosely defined population of glial cells found to ensheathe neuronal cell bodies, dendrites, and synapses of peripheral neurons, including autonomic and sensory neurons (Elfvin and Forsman 1978; Pannese 1981).

Morphologically, the satellite glial cell appears as a ring-like structure around neurons in

41 the ganglia and is characterized by its thin cytoplasm and expression of the protein, brain lipid binding protein (BLBP). Electron microscopic studies, focused mostly in the sensory nervous system, have revealed the close proximity of satellite glial cells to individual neurons, with only 20nm of space between their membranes (Dixon 1969).

This close association suggests a tight coupling between the cells to allow for possible exchange of important nutrients, yet few studies have investigated the role of the satellite glial cell or even begun to explore how they develop.

During development, both neurons and glial cells of the peripheral nervous system arise from the same population of stem cells called neural crest cells, discussed in detail in Chapter One. Briefly, these cells migrate from the neural tube around embryonic day 9.5 (E9.5) in the mouse and, for the sympathetic nervous system, coalesce at the dorsal aorta where secretion of BMPs instructs a subset of neural crest cells to differentiate into sympathetic neurons. Following differentiation of neurons, around

E14.5, satellite glial cells begin to proliferate and mature (Hall and Landis 1992). This proliferation peaks embryonically but continues throughout the first two weeks of postnatal development as satellite glial cells begin to ensheathe their neuronal neighbors.

One study found that satellite glial cells express consistent levels of brain lipid binding protein (BLBP), but over time change expression of other proteins. The authors observed

Nestin expression high during early postnatal development that decreased over time while expression of went up, and thus proposed a model where satellite glial cells switch from a proliferative immature state to a mature satellite glial cell fate around two weeks after birth (P14) (Shi et al. 2008). Yet still the molecular players responsible for satellite glial cell maturation remain undefined.

42 Sympathetic neuron development has been well characterized and given the progression of development where sympathetic neurons arise first, followed by satellite glial cells, as well as the close association of the two cell types, we asked how neurons communicate to satellite glial cells to promote development. Gene expression in sympathetic neurons is strongly influenced by the peripheral signal NGF during development, and the onset of NGF signaling coincides with the start of satellite glial cell development. Additionally, we found that neuron-specific deletion of the NGF receptor,

TrkA results in defects in glial cell morphology in mice (Figure 1.1a,b). A study identified several hundred potential target genes of NGF via microarray analysis from

NGF-deficient mice (Deppmann et al. 2008). One gene identified was a proposed Notch ligand called Delta/Notch-like EGF-related receptor (DNER), discussed in more detail in

Chapter Two.

Notch signaling is one of the best characterized signaling pathways involved in nervous system development. Studies manipulating the Notch pathway in melanogaster, Xenopus, chick, zebrafish, and mice have all provided strong evidence for the role of this pathway in the neural and glial cell fate decisions in which cells expressing the Notch ligand undergo neuronal differentiation while the neighboring cells expressing the receptor retain cell stem-ness (Ables et al. 2011; Appel, Givan, and Eisen

2001). This classic model of has dominated the field, however, studies over the last few decades have demonstrated that precise control of how and when Notch signaling is activated is important beyond cell fate, specifically for glial cell differentiation. For example, Notch signaling induces differentiation of Schwann cells, retinal Müller glial cells, and radial glial cells (Wang and Barres 2000; Wang et al. 1998).

43 Additionally, it is hypothesized that Notch signaling may also act as an on/off switch to control when oligodendrocytes and Schwann cells myelinate axons (Wang and Barres

2000). This heterogeneity in roles of Notch signaling may be due to differences in timing of expression as well as ligand-receptor pairing. Notch ligands can trigger canonical and non-canonical signaling pathways. With canonical Notch signaling, the Notch intracellular domain is cleaved by a gamma-secretase, which results in the translocation of the Notch intracellular domain to the nucleus where it binds to the CSL DNA binding protein and turns on transcription. Though non-canonical Notch signaling is not fully understood, in one non-canonical pathway a protein called Deltex binds to the Notch intracellular domain in the cytoplasm and prevents translocation (D'Souza, Meloty-

Kapella, and Weinmaster 2010; Mukherjee et al. 2005). In mammals there are four Notch receptors (Notch1-4), six canonical ligands (Delta1-4, Jagged1, and Jagged2), and a number of non-canonical ligands, including DNER (D'Souza, Meloty-Kapella, and

Weinmaster 2010).

DNER was found to promote Bergmann glial cell development in the mouse cerebellum. DNER is expressed specifically in Purkinje cells, the specialized neurons of the cerebellum, while the proposed receptor, Notch1, is expressed in the surrounding

Bergmann glia population. Co-immunoprecipitation experiments reveal DNER and

Notch1 are capable of interacting with each other. In mice with the gene for DNER globally knocked out, Bergmann glial cells mis-localize in the molecular layer of the cerebellum and extend fewer radial processes. This phenotype could be rescued with injection of a dominant negative Deltex virus, but not with the Notch intracellular

44 domain, suggesting that DNER is acting through a non-canonical Deltex-dependent

Notch pathway in these glial cells (Eiraku et al. 2002; Eiraku et al. 2005).

We found that DNER is expressed in sympathetic neurons as early as E13.5 and continues throughout development and into adulthood (Figure 2.1a). Expression of

DNER therefore coincides with the onset of satellite glial cell development. Here, we show that neuronal DNER is important for neuron-satellite glial cell contacts in the developing sympathetic nervous system. In cultures of satellite glial cells from neonatal rat pups, addition of a soluble form of DNER induces structural changes in glial cell morphology in which they change from a spindle-like shape to a flattened and round shape. In mice lacking DNER in sympathetic neurons, satellite glial cells make fewer contacts with neurons with no changes in total glial cell number. Our results suggest that

DNER may act through Notch on satellite glial cells. Inhibition of Notch signaling by addition a gamma-secretase inhibitor to satellite glial cell cultures blocks the morphological changes induced by DNER. Conversely, treatment of cultured satellite glia with soluble DNER upregulates Notch target gene expression. Together, these data suggest that sympathetic neurons express DNER which signals to satellite glial cells, potentially through Notch receptors, to instruct satellite glial cell ensheathment.

45 RESULTS

DNER influences satellite glial cell morphology in vitro and can activate Notch signaling

We asked what the role of DNER is in sympathetic nervous system development.

DNER has previously been implicated in regulating the maturation of Bergmann glial cells. We found that DNER is regulated by NGF-TrkA signaling and that loss of TrkA in sympathetic neurons disrupts the satellite glial cell sheath. Therefore, we first asked if

DNER affected satellite glial cell morphology. We first turned to an in vitro approach.

We cultured satellite glial cells from P0-P1 rat pups and treated them with a soluble form of DNER (DNER-Fc) and used BLBP and phalloidin immunohistochemistry to assess glial morphology. We found that satellite glial cells in culture primarily existed in two forms – a spindle, bipolar-like shape (polarized) and a flat, rounded shape (non- polarized). In control cultures, satellite glial cells exhibited an equal ratio of bipolar spindle shaped cells to flat, rounded cells. With Dner-Fc treatment, significantly more flat, rounded satellite glial cells were observed within 16 hours relative to control cells

(Figure 3.1a,b,e). As DNER was shown to act through Notch signaling in Bergmann glial cell development, we then asked if this change in morphology was due to activation of Notch. To address this, we used the gamma-secretase inhibitor, DAPT, to block Notch signaling in cultures treated with DNER-Fc. Treatment with DAPT blocked the morphology change induced by DNER-Fc and kept the glial cells in a spindle, bipolar shape (Figure 3.1c-e). Interestingly, treatment with DAPT alone resulted in an increase in the number of spindle, bipolar shaped satellite glial cells compared to untreated

46 samples suggesting that some level of endogenous Notch signaling may be occurring in the cultures and affecting the ratio of spindle, bipolar to flat, rounded glial cells. These data suggest that DNER is sufficient to instruct satellite glial cell morphology.

DAPT is an indirect Notch inhibitor by inhibiting gamma-secretase. However, gamma-secretase also cleaves other transmembrane proteins besides Notch. Therefore, to determine if DNER-Fc is capable of activating the Notch pathway, we cultured satellite glial cells and treated with DNER-Fc or Delta-like-1-Fc (DLL1-Fc, a known canonical

Notch ligand) and performed qPCR for the Notch target genes, Hey1 and Hes1, as well as

Notch binding components Deltex-1 (Dtx1) and Rbpj. As expected, addition of DLL1-Fc upregulated the levels of Hey1 and Hes1 mRNA. We found that, though not significant,

DNER-Fc also appears to be capable of upregulating expression of these Notch effectors while transcription of the Notch binding partners, Dtx1 and Rbpj, are unaffected (Figure

3.2a). Taken together, these data suggest that DNER-Fc influences satellite glial cell morphology likely through Notch signaling.

DNER cKO sympathetic ganglia have impaired neuron-satellite glial cell contact

We next asked if DNER played a role in satellite glial cell development in vivo.

To this end, we generated DNER cKO mice in which DNER was specifically ablated from sympathetic neurons by crossing TH-Cre mice with DNER-floxed mice (TH-

Cre;DNERf/f). DNER cKO mice were born at expected Mendelian ratios, had no obvious morphological abnormalities, and survived through adulthood. DNER transcript levels were significantly reduced in sympathetic ganglia of DNER cKO mice by IHC and qPCR

(Figure 3.3a-c). As DNER expression is first observed during embryogenesis when

47 satellite glial cells begin proliferating and differentiating, we first asked if glial cell proliferation was affected. To address this, we injected the thymidine analog, EdU, into

P0 neonates and assessed proliferation 24 hours later. At one day after birth, we found no difference in the total number of EdU-positive cells, indicating that cell proliferation was unperturbed in the DNER cKO SCG at this time (Figure 3.4a-c). As the majority of sympathetic neurons are post-mitotic by birth, it can be assumed that any proliferating cells within the ganglia at this time are non-neuronal in . Additionally, we saw no discernible changes in overall ganglia size suggesting that any changes in DNER cKO ganglia might be morphological in nature.

To determine if DNER affected other aspects of sympathetic ganglia development, we examined the gross morphology of sympathetic ganglia from DNER cKO mice. At postnatal day 14 (P14), control neurons were surrounded by the distinct thin rings of satellite glial cells. However, in DNER cKO mice the satellite glial cell rings appeared incomplete with gaps in the BLBP staining, similar to the phenotypes observed with conditional loss of TrkA from sympathetic neurons, suggesting satellite glial cell development is disrupted in these mice (Figure 3.5a-b).

The lack of a complete satellite glial cell sheath around DNER cKO neurons could be due to changes in BLBP expression and localization, disruption in glial cell morphology, or a loss of glial cells. Therefore, to gain a better understanding of what changes may be occurring, we used western blot analysis for BLBP to assess expression levels as well as qPCR for glial cell markers. There were no changes in BLBP expression levels by western blot (Figure 3.5c,d), nor did we observe any changes in transcript levels of several glial cell markers including BLBP, Sox10, glutamine synthetase (GS),

48 the neurotrophin receptor p75NTR, and connexin-43 (Figure 3.5e). To then assess glial cell number, we used toluidine blue staining. We assessed cell number by counting cells with the distinct clumped heterochromatin nuclear morphology that is displayed by satellite glial cells. The majority of non-neuronal cells within sympathetic ganglia are satellite glial cells, however, it is important to note that these numbers may also reflect the small number of and Schwann cells that exist within the ganglia as their nuclei appear similar. Thus, our count reflects cells that are non-neuronal in nature. We found no difference in total numbers of non-neuronal cells between control and DNER cKO ganglia (Figure 3.6c).

To examine the of sympathetic ganglia, we used transmission electron microscopy. We found that gross ganglia morphology appeared disrupted in

DNER cKO mice (Figure 3.6a,b). Control ganglia exhibited tight compaction of neuronal somata and the majority of non-neuronal cells with clumped heterochromatic nuclei were closely juxtaposed to neurons, indicative of tightly associated satellite glial cells. In contrast, neuronal somata in DNER cKO ganglia were smaller in size and had more spacing between cells. Additionally, the majority of non-neuronal cell nuclei were found in clusters in the extracellular space between neurons and were not associated with neuronal somata, suggesting a loss in neuron-glial cell contact.

In the mature sympathetic nervous system, satellite glial cells form multiple layers around individual neurons, consisting of up to ten layers. To our knowledge, no study has conducted a careful analysis of exact neuron to satellite glial cell ratios in the mouse sympathetic nervous system, however, in the mouse spinal ganglia one study found an average of 5.51 satellite glial cells associated with nerve cell bodies (Ledda, De Palo, and

49 Pannese 2004). By transmission electron microscopy we noted a large number of neurons without associated glial cells and many glial cells did not appear to contact neurons in the

DNER cKO ganglia. Therefore, we asked if the number of glial cells associated with individual neurons was disrupted. In order to quantify the number of glial cells associated with individual neurons, we used toluidine blue staining of semi-thin plastic serial sections. We quantified glial cells associated with neurons in which the entire neuronal cell body could be followed through the sections. Glial cells were again identified by the clumped heterochromatin characteristic of their nuclei as well as the close juxtaposition of the nuclei to the neuronal somata. In control animals, most neurons counted had three or more glial cells associated. In contrast, most DNER cKO neurons had fewer than three associated glial cells (Figure 3.7a-c). These results indicate that loss of DNER in sympathetic neurons results in a disruption in neuron-satellite glial cell contacts without loss of glial cell number.

50

Figure 3.1. DNER alters satellite glial cell morphology in vitro.

(a) Cultured satellite glial cells (BLBP: green, Phalloidin: red) from neonate rats have a polarized shape (Scale bar: 10um). (b) Addition of DNER-Fc to satellite glial cell cultures reduces cell polarity and induces satellite glial cells to become flatter and rounder. (c-d) Inhibition of Notch signaling with DAPT blocks the effect of DNER-Fc on satellite glial cell morphology. (e) Quantification of cell polarity of satellite glial cells.

Polarity was determined as having a length:width ratio greater than 2. (n=8 for control and DNER-Fc, 4 for +DAPT conditions, 50-150 cells per experiment, one way ANOVA,

**p<0.01, ****p<0.0001).

51

Figure 3.2. DNER-Fc is capable of activating Notch signaling in cultured satellite glial cells.

(a) qPCR analysis of Notch effectors (Hey1, Hes1) and components of the Notch pathway (Dtx1, Rbpj) shows DNER-Fc and DLL1-Fc are able to activate transcription of

Notch effectors but do not affect Notch signaling components (n=3, student t-test, p- values shown or not significant).

52

Figure 3.3. DNER is knocked down in DNER cKO sympathetic ganglia.

(a) Control sympathetic ganglia have robust DNER expression (TH: magenta, DNER: grey, DAPI: blue; Scale bar: 50um). (b) DNER expression is lost in DNER cKO ganglia.

(c) DNER transcript is significantly reduced in DNER cKO sympathetic ganglia (n=3 control, 3 DNER cKO, student t-test, **p<0.01).

53

Figure 3.4. Cell proliferation is unaffected in neonatal DNER cKO sympathetic ganglia.

(a-b) EdU (green) was pulsed for 24 hours in neonatal control and DNER cKO mice

(BLBP: magenta, TH: teal). (c) No difference in the total number of EdU-positive cells in control versus DNER cKO sympathetic ganglia. (n=3 control, 4 DNER cKO, student t- test, not significant).

54

Figure 3.5. Satellite glial cell ring structure is lost in sympathetic ganglia of DNER cKO mice.

(a) P14 sympathetic ganglia from control animals have thin rings of satellite glial cells

(BLBP: white in merge, black alone) around neurons (TH: green) (Scale bar: 25um). (b)

DNER cKO satellite glial cells lack clear ring structures around neurons. (c,d) Western blot for BLBP. Expression is unchanged between control and DNER cKO (n=2). (e)

Major glial cell marker expression (BLBP, Sox10, GS, p75NTR, connexin-43) is not different between control and DNER cKO ganglia (n=3 control, 3 DNER cKO, student t- test, not significant).

55

Figure 3.6. Sympathetic ganglia morphology is disrupted in DNER cKO mice.

(a) Electron microscopic view of sympathetic ganglia from P14 control mice show tight packing of neurons (pseudocolored teal) and closely juxtaposed non-neuronal nuclei

(pseudocolored magenta) (scale bars: 10um). (b) Sympathetic ganglia from P14 DNER cKO mice show less dense packing of neurons with non-neuronal nuclei filling the space between and fewer juxtaposed to the neurons. (c) Total non-neuronal cells per mm2 shows no difference between control and DNER cKO ganglia (n=3 control, 3 DNER cKO, student t-test, not significant).

56

Figure 3.7. Neuron-satellite glial cell contacts are lost in DNER cKO ganglia.

(a) Toluidine blue staining of plastic serial sections of sympathetic ganglia from control mice with three non-neuronal nuclei (asterisk) juxtaposed to the neuronal membrane. (b)

Toluidine blue staining of plastic serial sections of sympathetic ganglia from DNER cKO mice with one non-neuronal nuclei (asterisk) juxtaposed to the neuronal membrane. (c)

Quantification of number of non-neuronal cells per neuron in control and DNER cKO mice (n=3 control, 3 DNER cKO, 11-16 neurons counted per animal, student t-test,

*p<0.05).

57 DISCUSSION

Here we show that in vitro DNER is sufficient to induce morphological changes in satellite glial cells in which they shift from a bipolar spindle shape to a flat, rounded shape. This change can be blocked by addition of the gamma-secretase and indirect

Notch inhibitor, DAPT. The non-polar shape of the satellite glial cells induced by DNER-

Fc is likely an important shift in cellular architecture during ensheathment of satellite glial cells around neurons. In the ganglia, satellite glial cells exhibit a thin ring of cytoplasm around neuronal cell bodies, dendrites, and synapses. The flattened and rounded shape of the non-polar glial cells could be indicative of a morphological change the glial cells go through in order to blanket around neurons. As DNER is a transmembrane protein, it stands to reason that this event would most likely occur at contact with the neurons. We hypothesize that prior to ensheathment around sympathetic neurons, satellite glial cells exist in a bipolar state and that upon contact with the neurons, and presumably DNER, they lose the polar shape and their cytoplasm thins in order to form a complete sheath around the neuron.

We generated a conditional knockout mouse of DNER in sympathetic neurons and found that DNER is required for neuron-satellite glial cell contact in the developing sympathetic nervous system. Immunohistochemical and electron microscopic analyses show that neurons lacking DNER have fewer associated glial cells and the thin glial cell sheathe around the neurons is disrupted. Interestingly, the effects of DNER loss on satellite glial cells appears to be only morphological as neither proliferation nor total non- neuronal cell number appear to be affected in DNER cKO ganglia. These data support the idea that the morphological change elicited by DNER-Fc in glial cell cultures is required

58 for the ensheathment process. However, it is difficult to ascertain satellite glial cell morphology in vivo with the techniques currently available. As satellite glial cells can form multiple sheaths around individual neurons and few specific markers are known, standard immunohistochemical analysis cannot differentiate one glial cell from another within the ganglia. Single-cell labeling of satellite glial cells or high resolution 3- dimensional electron microscopy may be useful to observe individual glial cell ensheathment around neurons in future studies.

In vitro treatment of glial cell cultures with DAPT increased the number of polarized glial cells and blocked the non-polarizing effect of DNER-Fc. A few studies have identified DNER as a ligand for Notch signaling while others have found that

DNER can act through Notch-independent pathways as well. An important step in the

Notch effector pathway is cleavage of the intracellular domain of Notch by gamma- secretase. DAPT is a gamma-secretase inhibitor and thus indirectly blocks Notch signaling, however, gamma-secretase cleaves several other transmembrane proteins as well, such as E-cadherin and ErbB-4. It is therefore possible that the effects of inhibiting gamma-secretase activity with DAPT on satellite glial cell cultures could due to inhibition of other targets, or a combination of the two. By qPCR, we found that DNER-

Fc is capable of activating canonical Notch targets of the Hes and Hey family, but whether or not this is the mechanism by which DNER-Fc influences glial cell morphology and if this is how DNER functions in vivo remains unknown. It would be of interest for future studies to delve into the mechanism of DNER action on satellite glial cells and whether or not DNER acts through Notch or an as of yet unknown receptor.

59 Finally, our conditional knockout mouse ablates DNER in sympathetic neurons and by electron microscopy we noted a decrease in the association between neurons and glia in the knockout ganglia. Further, knockout neurons appeared to be smaller in size and atrophied. These observations raise the question of the effects of DNER loss on the sympathetic neurons.

60 MATERIALS AND METHODS

Animal Husbandry

All animal procedures were performed in accordance with NIH and Johns Hopkins

University Animal Care and Use Committee (JHU ACUC) guidelines. Animals were kept in a 12-hour light:dark cycle with food and water ad libitum. Mice were kept in mixed

C57BL/6 and 129P backgrounds and animals of both sexes were used for analyses. TH-

Cre mice were gifts from Dr. C. Gerfen (NIH). Dnertm3a KO first allele (C57BL/6NTac-

Dnertm3a(EUCOMM)Hmgu/Ieg) mice were purchased from EMMA Repository (strain ID:

EM:08389). Dnertm3a mice were crossed to a ROSA26-FLPe mouse (Jackson strain number: 003946) to knock out the LacZ and neo cassette to generate the conditional

Dnertm3c allele (Dner-floxed) mice, which were then mated to TH-Cre in order to generate

TH-Cre;Dnerf/f (DNER cKO) mice. Littermate DNERf/f mice were used as controls in all experiments. Sprague-Dawley rats were purchased from Charles River (strain code: 400).

Isolation of sympathetic ganglia

Neurons from superior cervical ganglia (SCG) of Sprague-Dawley rat pups (P0.5) or mice of indicated genotypes and ages were dissected and enzymatically dissociated as previously described (Bodmer et al., 2011). Briefly, pups were anesthetized according to

JHU ACUC protocol. SCGs of pups were dissected and separated from surrounding tissue, incubated for 20 minutes at 37°C in DMEM AIR (DMEM F12, 12.5mM glucose,

1U/ml penicillin/streptomycin) and supplemented with 4mg/ml collagenase IV, 1mg/ml hyaluronidase, 0.6mg/ml DNase I, and 10mg/ml BSA. Ganglia were centrifuged, resuspended in DMEM air containing 3mg/ml trypsin, and incubated for an additional 30

61 minutes at 37°C. Cells were mechanically dissociated by tituration with a glass pipet, centrifuged, and resuspended in DMEM supplemented with 10% FBS and 1U/ml penicillin/streptomycin for glial cell cultures. Neuronal culture media was additionally supplemented with 100ng/ml NGF and 10mM Ara-C to prevent glial cell proliferation.

For mass cultures, SCGs from 5-6 rats or 6-8 mice were plated on collagen-coated 60mm or 35mm dishes, respectively. For sparse cultures, SCGs from 0.5-1 rats or 1-2 mice were plated on coverslips coated with Poly-D-laminin mixture.

Immunohistochemistry

For IHC, mice at indicated ages were fixed in 4% PFA for 4-16 hours at 4°C, cryoprotected in 30% sucrose in PBS, embedded in OCT, and serially sectioned (12-

30um) with a Thermo Microtom HM 550 cryostat. Sections were blocked with 5% DS serum and 0.1% TX-100 in PBS for 1 hour and incubated in primary antibody overnight at 4°C. Sections were then washed with PBS, incubated with secondary antibody for 1 hour at room temperature, washed again, and mounted with fluorimount plus DAPI. For proliferation analyses, P0.5 pups were injected with 50ug of EdU and sacked for IHC 24 hours later. IHC was conducted as described and prior to mounting, EdU was detected using Click-iT chemistry following the manufacturers protocol (Click-iT Plus EdU Cell

Proliferation Kit, Thermo Scientific C10637). Animals were serially sectioned and total number of EdU-positive cells were counted in every 5th section. Images were taken with a

Zeiss LSM 700 confocal microscope.

In vitro glial cell morphology

62 Glial cells were sparsely plated on glass coverslips and incubated for 16 hours with 2ug of DNER-Fc or PBS in the presence or absence of 10mM N-[N-(3,5-

Difluorophenacetyl)-L-alanyl]-S-phenylglycine t-butyl ester (DAPT). Cells were fixed for 20 minutes with 4% PFA, washed in PBS, and blocked for 1 hour at room temperature with 1% BSA and 0.1%TX-100 in PBS. Primary antibodies were diluted in block and incubated on coverslips for 2 hours at room temperature. Following primary, coverslips were washed with PBS, incubated with secondary antibody for 1 hour at room temperature, washed again, and mounted with fluorimount plus DAPI. Images were taken with a Zeiss LSM 700 confocal microscope. Glial cell morphology was determined using length:width ratio using ImageJ software analysis. A length:width ratio of greater than 2 was considered polarized.

Quantitative real-time PCR

For determination of Notch activation by DNER-Fc, glial cells were cultures in mass and incubated for 16 hours with 2ug DNER-Fc, DLL1-Fc, or PBS in DMEM supplemented with 10% FBS, 1U/ml penicillin/streptomycin. Cultures were then washed with PBS and

250ul of Trizol was added for RNA extraction. For sympathetic ganglia, 100ul of Trizol was used for one animal (two SCGs). RNA extraction was performed according to manufacturer's protocol (Invitrogen 15596026) using phenol:chloroform. For cDNA synthesis, 5ug of RNA were reverse transcribed with SuperScript IV Reverse

Transcription following manufacturer’s protocol (Thermo Scientific 18091050). qPCR was performed using Maxima SYBR green/ROX qPCR Master Mix (Thermo Scientific

63 K0222) and detected using Applied Biosystems StepOnePlus Real-Time PCR System

(cat 4376600).

Transmission Electron Microscopy

Tissues were fixed in 1% , 2.5% , 3% sucrose, 1 mM MgSO4 in 0.1 M phosphate or cacodylate buffer (pH 7.4). All tissues were then postfixed in 1% buffered with 1.5% potassium ferrocyanide. Tissues were embedded in

Eponate resin, sectioned at a thickness of 90 nm, and collected on single-slot grids with a carbon-coated Formvar film. Sections were poststained with uranyl acetate followed by lead citrate. Large-scale transmission electron microscopy (TEM) at 5,000x yielding a

2nm/pixel lateral resolution image. Images were then mosaiced, creating large-scale

TEM composite images as previously described (Anderson et al. 2009).

Toluidine Blue

Tissues were fixed in 3% formaldehyde, 1.5% glutaraldehyde, 2.5% sucrose, 5mM Ca2+ in 0.1M cacodylate buffer (pH 7.4), postfixed with 1% OsO4 then Kellenberger UA

(0.5% uranyl acetate). Tissues were embedded with Eponate resin, serial sectioned at

2um thickness with a glass knife, and collected on slides. Sections were stained with toluidine blue and imaged at 40x with Zeiss Microscope Axio Imager M1, stitched together with ImageJ and aligned using Reconstruct (Fiala JC (2005) Reconstruct: a free editor for serial section microscopy. J Microscopy 218:52-61.). Glial cell nuclei (defined as a nucleus with clumped heterochromatin) were counted if juxtaposed directly on

64 neuronal membrane for neurons in which the entire neuron could be followed through sections.

Statistical analyses

Sample sizes were similar to those reported in previous publications (Armstrong et al.

2011; Patel et al. 2015). Data were collected randomly. For practical reasons, analyses of neuronal cell counts in mouse tissues were done in a semi-blinded manner such that the investigator was aware of the genotypes prior to the experiment, but conducted the staining and data analyses without knowing the genotypes of each sample. All Student’s t tests were performed assuming Gaussian distribution, two-tailed, unpaired, and a confidence interval of 95%. Statistical analyses were based on at least 3 independent experiments and described in the figure legends. All error bars represent the standard error of the mean (s.e.m).

Antibodies

Antibody Company Dilution Use Mouse anti-α-tubulin Sigma, T9026 1:2000 Western blot Rabbit anti-BLBP Abcam, AB32423 1:200 IHC Mouse anti-BLBP Abcam, AB131137 1:500 IHC Goat anti-DNER Sigma, AF2254 1:75 IHC 1:1000 Western blot Mouse anti-tyrosine Sigma, T2928 1:500 IHC hydroxylase Rabbit anti-tyrosine Millipore, AB152 1:500 IHC hydroxylase Phalloidin-546 Life Tech, A22283 1:50 IHC Goat anti-rabbit Thermo Scientific, 1:500 IHC Alexafluor488 A11034 Donkey anti-rabbit Thermo Scientific, 1:500 IHC Alexafluor488 A21206

65 Donkey anti-goat Thermo Scientific, 1:500 IHC Alexafluor555 A21432 Goat anti-rabbit Thermo Scientific, 1:500 IHC Alexafluor546 A11035 Goat anti-rabbit Thermo Scientific, 1:500 IHC Alexafluor647 A21244 Goat anti-mouse IgG2b Thermo Scientific, 1:500 IHC Alexafluor488 A21141 Goat anti-mouse IgG2b Thermo Scientific, 1:500 IHC Alexafluor546 A21143 Goat anti-mouse IgGG1 Thermo Scientific, 1:500 IHC Alexafluor488 A21121 Goat anti-mouse IgGG1 Thermo Scientific, 1:500 IHC Alexafluor546 A21123 ECL anti-mouse IgG HRP GE Healthcare, 1:2000 Western blot NA931V ECL anti-rabbit IgG HRP Sigma, NA934 1:2000 Western blot

66 Chapter Four: Disruption of neuronal morphology and activity in DNER cKO mice

67 INTRODUCTION

Glial control of neuronal architecture and activity

Neurons are unique in structure and function from other cell types in the body.

They are highly polarized cells with three distinct compartments: the soma, dendrites, and axon. Surrounding and supporting nearly every compartment of neurons are glial cells, which encompasses a number of cell types including astrocytes, oligodendrocytes, microglia, Schwann cells, and satellite glial cells. Glial cells help shape and provide support to neurons and assist in their function. Some functions include control of the extracellular microenvironment, assistance in synaptic transmission, myelination of axons, and in response to (Zuchero and Barres 2015).

During development, neurons grow primary dendrites and axons. Astrocytes and other glial cells, such as oligodendrocytes and Schwann cells, can influence the extent of neurite growth in development and during injury. Many studies have identified glial- derived factors, such as secretion of neurotrophic factors and expression of adhesion molecules, that modulate neurite growth in numerous populations of neurons (Deumens et al. 2004; Bostrom et al. 2010; Altschuler et al. 1999; Yamamoto et al. 2006; Yadav et al. 2019). For example, astrocytes promote the extension of hippocampal neurites through expression of N-cadherin (Kanemaru et al. 2007). Interestingly, glial cells are also capable of inhibiting neurite growth. One study found that homogenous populations of both astrocytes or oligodendrocytes inhibit growth of neurites from spiral ganglion neurons (Jeon et al. 2011). Though the mechanism of this inhibition is unknown, it appears that glial cells possess the tools necessary to both promote and inhibit neurite growth in a manner dependent on neuronal type and the region in the body.

68 Neurons communicate with one another through synapses between axons and dendrites or axons and somas. The formation of synapses is a highly dynamic process, and in the central nervous system, glial cells play a key role in this development. For example, astrocytes form connections with neurons at synapses and play an active role in their formation. When cultured retinal ganglion cells form few synapses naturally, but the addition of astrocytes or astrocyte conditioned media to cultures significantly increases the number of synapses as well as synaptic activity (Pfrieger and Barres 1997; Ullian et al. 2001). Other studies have shown astrocytic control of synapses on motor neurons, hippocampal neurons, cerebellar neurons, and cortical neurons as well. The induction of synapse formation by astrocytes is achieved through secreted and contact-dependent mechanisms (Allen and Barres 2005). In addition to astrocytes, oligodendrocytes and

Schwann cells can also influence synaptic structure. For example, Schwann cells have been documented for their role in strengthening and eliminating synapses and axons in an activity-dependent manner (Allen and Barres 2005; Zuchero and Barres 2015).

Glial cells also play roles in modulating neuronal activity. For example, oligodendrocytes and Schwann cells myelinate axons in order to assist in action potential propagation (Zuchero and Barres 2015). Meanwhile, astrocytes cover between 60 and 90 percent of synapses in various regions of the brain and play a vital role in controlling the extracellular micro-environment and neuronal excitability (Farhy-Tselnicker and Allen

2018). During synaptic transmission, are released into the synapse and trigger an action potential in the receiving neuron which causes potassium levels to spike in the extracellular space. Astrocytes regulate the potassium concentration through expression of gap junctions and inwardly rectifying potassium channels (Allen and

69 Barres 2005). Removal of gap junctions from astrocytes leads to hyperactivation of hippocampal neurons and impairs (Pannasch et al. 2011).

Glial cells also regulate neuronal activity via neurotransmitter expression and removal. For example, the synthesis of some primary neurotransmitters - glutamate,

GABA, and glycine – is affected by the presence of astrocytes. This is best highlighted by the glutamate-glutamine cycle between neurons and astrocytes in which glutamate is released into the synapse, triggers a response in the post-synaptic cell, and is then taken up by astrocytes. Within the astrocyte glutamate is converted into glutamine and released back to the neuron. Once glutamine is taken up by the neuron it is finally converted back to glutamate and used for future events (Bak, Schousboe, and

Waagepetersen 2006). Without the of neurotransmitters like glutamate by astrocytes, neurons are subject to and subsequent neuronal damage. Studies using hippocampal slice cultures have demonstrated that inhibition of glutamate transporters leads to (Bonde et al. 2003; O'Shea et al. 2002).

Mechanisms of neurite growth and pruning

While glial cells play some roles in neuronal structure, the growth of neurites is also controlled by many different elements, such as growth factors that vary depending on the region of the nervous system and type of neuron. Axon initiation and growth of sympathetic neurons is achieved through several molecules such as HGF, artemin, NT3, and NGF (discussed in more detail in Chapter One). Once at the target tissues, however, axons undergo fine tuning in order to determine final branching and synapse formation.

Growth factors like neurotrophins play a major role in the refinement of axons at target

70 tissues through promotion of axon collateral branch formation and axonal pruning. For example, NGF was found to initiate axon collateral branching from DRG axons in vitro through the reorganization of filaments via the PI3K pathway (Gallo and

Letourneau 1998; Ketschek and Gallo 2010). In vivo expression of the beta subunit of

NGF in sympathetic neurons increased the number of sympathetic fibers innervating target tissues but reduced the number of terminal branches which could be reversed by expression of NGF in the target tissues. This demonstrated a requirement for strict NGF expression levels in order to define innervation density and patterning of target tissue

(Hoyle et al. 1993). Additionally, the survival of sympathetic axons and neurons is dictated through competition for limiting amounts target derived NGF. However, growth factors are not the only players in this process - neuronal activity is another major player in axon refinement in the nervous system.

During development and plasticity, neuronal activity can both positively and negatively influence the level of target innervation. Spontaneous spikes of neuronal activity via Ca2+ transients promote axon branching in developing neurons of the , , , and spinal cord prior to electrical input (Feller 1999).

Ca2+ transients may be caused by electrical activity through synapses or through extracellular signals. For example, -1 stimulation of cortical neurons increases Ca2+ transients that increase axon branching (Tang and Kalil 2005). Additional electrical activity through the activation of synaptic inputs can also influence axonal branching and synapse strength. For example, silencing activity of sympathetic neuron input to the in rats via exposure to constant light results in a reduction of axonal fibers

(Calinescu et al. 2011). Additionally, inhibiting neuronal activity in slice cultures of

71 thalamocortical neurons also reduces axon branching and synapse formation while inducing neuronal activity increases axon branching (Matsumoto et al. 2016; Uesaka et al. 2007). Though it has been well established that neuronal activity can shape axon branching and synaptic inputs, the mechanisms for this process remain elusive.

Here we show that loss of DNER in sympathetic neurons alters neuronal architecture and metabolism. Sympathetic neurons in DNER cKO mice exhibit atrophying of the neuronal cell body without a loss in total cell number. This decreased soma size is accompanied by a decrease in norepinephrine biosynthetic enzyme expression, specifically in the cell body compartment. Intriguingly, we also find abnormal mitochondrial morphology in which DNER cKO mitochondria were elongated and possessed a higher cristae density. These changes in the neuron cell body compartment suggest compromised sympathetic neuronal metabolism. We also note that upon depolarization, calcium transients in DNER cKO neurons persist longer than control neurons. It is possible that the changes in neuronal activity have consequences on neuronal metabolism, or that the metabolic changes causes alterations in activity. More studies are needed to determine if these phenotypes are connected. Finally, though we observe alterations in neuronal metabolism in the cell body compartment, we find that

DNER cKO axons hyper-innervate their target tissues. Target tissue hyper-innervation and maintenance of axonal TH levels may be a compensatory mechanism for losses observed in the cell body compartment in order to maintain metabolic homeostasis.

Overall, we observe alterations in sympathetic neuron structure and activity with loss of

DNER.

72 RESULTS

Sympathetic neurons lacking DNER are smaller and down-regulate noradrenergic enzymes

We generated DNER cKO mice in which DNER was specifically ablated from

TH-positive sympathetic neurons. Previously we examined sympathetic ganglia morphology via IHC and TEM and found that in addition to defects in neuron-satellite glial cell contacts, tyrosine hydroxylase (TH) levels appeared to be downregulated

(Figure 3.3) and overall ganglia morphology appeared altered in DNER cKO mice

(Figure 3.5). Given these observations and the fact that our conditional knockout line is in neurons themselves, we asked if neuronal development was also affected. We first examined tyrosine hydroxylase levels in sympathetic ganglia by qPCR and found a significant reduction in TH transcript levels (Figure 4.1a-c). Tyrosine hydroxylase is the rate-limiting enzyme of norepinephrine, the neurotransmitter used by sympathetic neurons. So, we then asked if the downregulation was specific to TH or if other enzymes in the norepinephrine biosynthetic pathway were also affected. We again performed qPCR for the enzyme responsible for synthesis of norepinephrine, dopamine β- hydroxylase (DBH), and found a similar reduction in DBH levels (Figure 4.1c). As neurotransmitter synthesis is a hallmark of neuronal identity, we examined levels of several other important neuronal identity markers: TrkA, Isl1, and Phox2b. By qPCR we found no significant difference in levels of those markers, suggesting that the downregulation of TH and DBH is a specific effect on the neurotransmitter machinery

(Figure 4.1c). To then ask if downregulation of TH and DBH affected overall norepinephrine levels, we measured norepinephrine by ELISA from sympathetic ganglia

73 and one of the major target tissues, the salivary glands. Despite the downregulation of enzymes in the biosynthetic pathway, we found no difference in total norepinephrine levels (Figure 4.1d).

In addition to the downregulation of TH, neuronal cell bodies appeared smaller by immunohistochemistry and transmission electron microscopy, so we next examined neuronal cell size. To this end, we used TrkA to visualize the neuronal cell bodies and determined the area of individual neurons at multiple depths within the

P14 sympathetic ganglia. In control animals, sympathetic neurons ranged from just under

100um in area to over 300um with an average size of 202.8um. In contrast, DNER cKO sympathetic neurons had significantly more smaller diameter neurons at the expense of larger diameter neurons with an average size of 158um (Figure 4.2a-d). The smaller size of the neurons suggests they may be atrophying, and so we asked if neuronal survival was affected. To this end we used Nissl stain to visualize and count sympathetic neurons at several time points throughout postnatal development (P1, P7, P14, and P28). There was no discernable difference in neuronal number at any time point tested between control and DNER cKO mice, suggesting that the smaller neuron size is not affecting cell survival (Figure 4.3a-c).

Another hallmark of neuronal atrophy is the retraction or loss of neurites.

Therefore, we next asked if sympathetic innervation was compromised. To examine target innervation, we used iDISCO whole-tissue processing of sympathetic target organs using tyrosine hydroxylase as a marker. Given the shrinkage of neuronal cell bodies, we hypothesized that sympathetic innervation would be decreased. Unexpectedly, we found that sympathetic targets were hyper-innervated in DNER cKO mice (Figure 4.4a,b,e).

74 During development, sympathetic axons are overproduced and pruned over time. The nervous system is still developing at two weeks after birth. Therefore, to determine if the hyper-innervation phenotype persists, we also examined sympathetic innervation in one- month old animals. Again, we found the heart to be hyper-innervated (Figure 4.4c-e).

Notably, we used TH as a marker for sympathetic innervation. In the cell bodies of DNER cKO neurons, TH was downregulated. However, in the heart TH levels appeared normal. It is possible that there is a compensatory upregulation of TH in the axons, which could emphasize the presence of smaller innervating fibers in DNER cKO mice that are not visible in control tissues and thus explain the increased innervation.

Indeed, in animals where TH was partially ablated from adult pars compacta neurons, reduction of TH levels occurred more slowly in axons than cell bodies, suggesting that axonal TH may be more stable (Tokuoka et al. 2011).

Therefore, to determine if target tissues in DNER cKO mice are truly hyper-innervated or if it is simply an upregulation of TH in the axons, we looked at innervation of the salivary glands using both TH and a pan-neuronal marker, Tuj1. Both TH and Tuj1 levels were increased in the salivary glands of DNER cKO mice (Figure 4.5a-f). These data suggest that loss of DNER in sympathetic neurons results in hyper-innervation of target tissues.

To determine if the hyper-innervation was an intrinsic effect due to the loss of

DNER in sympathetic neurons, we cultured control and knockout neurons for 16 hours and assessed neurite outgrowth and branching. We found no difference in neurite length or number of branches between control and DNER cKO neurons (Figure 4.6a-d), which suggests the hyper-innervation of target tissues is not due to an intrinsic change within the neurons or the direct loss of DNER itself, and may be due to some extrinsic factor.

75 Taken together, these data indicate a role for DNER in sympathetic neuron size and innervation, but not survival.

Depolarization of DNER cKO neurons leads to a sustained calcium response

Glial cell ensheathment of synapses and around the neuronal soma have been described to regulate neuronal activity (Yadav et al. 2019; Yamamoto et al. 2006).

Additionally, enhanced neuronal activity could contribute to excessive axonal outgrowth

(Calinescu et al. 2011). With the loss of glial cell contacts and hyper-innervation of target tissues in DNER cKO ganglia, we considered that neuronal activity may be affected.

Upon depolarization of neurons there is a sharp rise in intracellular calcium levels which can be exploited using calcium indicators to visualize neuronal activity in vitro.

Therefore, to determine if activity of sympathetic neurons is affected in DNER cKO mice, we used calcium imaging of neuronal explants isolated from control or knockout mice. The use of sympathetic ganglia explants allows for the assessment of neuronal activity while retaining neuron-glial cell contact. Several studies have noted, however, that glial cells undergo significant changes in vitro and migrate out of explants within a few days of culturing (Chamley, Mark, and Burnstock 1972; Hanani 2010; Belzer,

Shraer, and Hanani 2010). To avoid this possibility, we used a short culturing time of 16- hours and loaded sympathetic explants with the calcium indicator dye, Fluo4-AM, which is taken up by cells within 40 minutes of exposure. We then induced depolarization using

50mM KCl and assessed the change in calcium levels from individual neurons within the explants. Neurons were visually identified by cell body shape, size, and the presence of neurites. Upon depolarization, calcium levels in control neurons spiked rapidly and

76 quickly decreased back to baseline. Strikingly, we found that DNER cKO neurons spiked similarly to control neurons but retained higher calcium levels for a longer period of time, indicating a sustained calcium response (Figure 4.7a-c). These results suggest that neuronal loss of DNER alters neuronal activity.

DNER cKO neurons have altered mitochondrial morphology

Regulation of neuronal metabolism, neuronal activity, and axon branching are all complex processes. Countless proteins and play roles in ensuring that neurons are healthy and function properly. One of the primary players in cellular metabolism is the mitochondria, which not only provides the majority of ATP for cellular , but also acts as a calcium store and sink. In particular, mitochondria play a primary role in calcium buffering following depolarization by uptake of calcium from the cytosol.

Mitochondria are large organelles with varying structures and sizes depending on the cell type. As neurons are some of the most metabolically active cell types in the body, they require a lot of ATP and are often densely packed with mitochondria. Interestingly, during observation of the sympathetic ganglia by TEM, we noticed changes in mitochondria in DNER cKO neuronal cell bodies in which the mitochondria appeared elongated and electron-dense (Figure 4.8a,b). We therefore assessed the mitochondria more closely. We first determined mitochondrial content and found the total number of mitochondria in the cell bodies of sympathetic neurons were slightly increased in DNER cKO mice (Figure 4.8c). We then assessed mitochondrial morphology. In control neurons, mitochondria were mostly seen as large and round while in DNER cKO neurons the mitochondria were often small and elongated (Figure 4.8d-g). In addition to the

77 change in shape of the mitochondria in knockout neurons, they were also found to be more electron-dense with a 50 percent increase in cristae density (Figure 4.8h). Cristae are the inner folds of mitochondria where the majority of metabolic enzymes and channels are located. Increased cristae density and changes in mitochondrial morphology have been observed in cells under hypoxic stress. In DNER cKO neurons these changes could be a consequence of the smaller neuron size, persistent activation, and increased innervation density as a way for the neurons to keep up with a higher metabolic demand.

78

Figure 4.1. Loss of noradrenergic biosynthetic pathway enzyme expression in DNER cKO ganglia.

(a) Tyrosine hydroxylase (TH) immunostaining (white) from control and (b) DNER cKO

SCGs shows decreased levels of TH in mutant animals. (c) qPCR for TH, dopamine β- hydroxylase (DBH), TrkA, Isl1, and Phox2b transcript levels from control and DNER cKO SCGS. Noradrenergic biosynthesis enzymes, TH and DBH, levels are significantly decreased in DNER cKO SCG while neuronal identity markers TrkA, Isl1, and Phox2b are unchanged (n=6 control, 6 DNER cKO animals, student t-test, ***p<0.001,

****p<0.0001). (d) Norepinephrine content from SCGs and (e) salivary glands of control and DNER cKO animals shows no significant difference (n=16 control, 19 DNER cKO

SCGs; n=13 control, 12 DNER cKO salivary glands, student t-test, not significant).

79

Figure 4.2. DNER cKO neurons are smaller in size.

(a) TrkA immunostaining (magenta) from control and (b) DNER cKO SCGs shows larger neurons in control animals. (c) Quantification of neuronal size binned and shown as percent of neurons with an area of less than 100um, 100-200um, 200-300um, and greater than 300um. (d) Average neuronal area per animal. (n=3 control, 3 DNER cKO with 247-

328 neurons counted per animal, student t-test, *p<0.05).

80

81 Figure 4.3. Neuronal survival is unaffected in DNER cKO animals.

(a) Nissl stain from P14 control and (b) DNER cKO SCG (scale bar: 50um). (c) Total neuronal number from P1, P7, P14, and P28 SCGs shows no change in neuron number between control and DNER cKO animals. (n=4 control, 4 DNER cKO at P1; n=6 control,

5 DNER cKO at P7; n=3 control, 4 DNER cKO at P14; n=3 control, 3 DNER cKO at P28, student t-test, not significant).

82

Figure 4.4. DNER cKO neurons hyper-innervate the heart.

(a) Tyrosine hydroxylase staining (white) from control and (b) DNER cKO shows increased innervation in mutant animals at P14 (scale bar: 50um). (c) Innervation of P30 hearts from control and (d) DNER cKO animals shows persistent hyper-innervation of target tissues into adulthood. (e) Quantification of innervation from P14 and P30 hearts shown as percent of sex-matched littermate controls. (n=4 control, 4 DNER cKO hearts at

P14 and n=3 control, 4 DNER cKO hearts at P30, student t-test, *p<0.05).

83

Figure 4.5. Salivary glands are hyper-innervated in DNER cKO mice.

(a) Salivary glands from P14 control and (b) DNER cKO mice stained with TH (scale bar: 50um). (c) Quantification of TH innervation of salivary glands shows hyper- innervation in DNER cKO mice. (n=3 control, 3 DNER cKO mice, student t-test,

*p<0.05). (d) Tuj1 immunostaining of salivary glands from control and (e) DNER cKO mice (scale bar: 50um). (f) Quantification of Tuj1 innervation shows hyper-innervated

DNER cKO salivary glands. (n=3 control, 3 DNER cKO mice, student t-test, **p<0.01).

84

Figure 4.6. Cultured DNER cKO neurons do not exhibit changes in neurite length or branching.

(a) Sympathetic neurons cultured for 16 hours from control or (b) DNER cKO mice immunostained for TH (green) and BIII-tubulin (magenta) (scale bar: 25um). (c) Total length of the longest neurite and (d) number of branch points from control or DNER cKO neurons is unchanged (n=4 individual experiments, 15-46 neurons per experiment).

85

86 Figure 4.7. DNER cKO neurons exhibit a sustained calcium response to depolarization

(a) Graph of relative Fluo4-AM signal within control or DNER cKO neurons over time normalized to background levels. (b) Area under the curve (AUC) of Fluo4-AM response to depolarization. (c) Peak Fluo4-AM signal following depolarization of neurons with

50mM KCl. DNER cKO neuron calcium levels spike similarly but are maintained for an extended period of time compared to control neurons following depolarization.

(n=18 control, 18 DNER cKO neurons from 3 experiments, student t-test, n.s.).

87

88

Figure 4.8. Altered mitochondrial morphology in DNER cKO neurons.

(a) Mitochondrial content from control and (b) DNER cKO neurons (scale bar: 500um).

(c) Average mitochondrial content per um2 shows slightly higher mitochondrial content in DNER cKO neurons compared to controls (n=3 control, 4 DNER cKO, mitochondria measured from 15-21 neurons per animal). (d) High magnification of mitochondria from control and (e) DNER cKO neurons shows altered mitochondrial shape and cristae morphology in DNER cKO neurons (scale bar: 500um). (f) Mitochondria area in um2 of mitochondria from control and DNER cKO neurons, binned and represented as the percent of mitochondria with areas less than 0.1um2, between 0.1-0.2um2, and greater than 0.2um2. DNER cKO mitochondria are smaller than control mitochondria. (g) Percent of elongated mitochondria (defined as having a roundness index of less than 0.5). DNER cKO neurons have elongated mitochondria. (h) Cristae density of mitochondria shown as a percent of control shows 50% increase in cristae density of mitochondria from DNER cKO neurons. (For graphs f-h: n=3 control, 4 DNER cKO animals with 89-132 mitochondria measured from 15-21 neurons per animal, student t-test, *p<0.05,

**p<0.01).

89 DISCUSSION

We found that DNER cKO neurons were smaller than control wildtype neurons on average and had decreased levels of TH and DBH in the cell bodies while levels in axons appeared unchanged. Despite the downregulation of key enzymes in the norepinephrine biosynthetic pathway, there were no changes in norepinephrine levels in either the cell body or axonal tissues. Studies have previously demonstrated that partial loss of TH, the rate-limiting enzyme of dopamine, , and norepinephrine biosynthesis, does not drastically change overall neurotransmitter expression. In TH heterozygous animals where activity of the enzyme was significantly decreased there was only a modest loss of expression in neonates. Interestingly, norepinephrine levels in the TH heterozygote neonates were only decreased in the brain while the levels in the body matched those of wildtype animals, and levels of all three catecholamine’s were normal in adult heterozygous animals (Kobayashi et al. 1995). In another study, regulation of TH protein expression was found to differ between cell bodies and axons in the substantia nigra where knockdown of TH via adenovirus expression of Cre in TH-floxed mice resulted in a faster loss of TH in the cell bodies than the axons. This study also found that despite even a 50% reduction in TH protein levels, the amount of dopamine produced was relatively unchanged (Tokuoka et al. 2011). These studies suggest that multiple regulatory mechanisms exist to maintain catecholamine levels even when there is appreciable loss of enzymes involved in their biosynthesis. Additionally, the slower loss of TH protein in axons of neurons where TH was conditionally ablated suggests that the protein may be more stable in axons than in cell bodies or there could be TH synthesis in axons given the presence of axonal TH mRNA, which could explain the reduction of TH

90 levels in the cell bodies of DNER cKO neurons whereas levels were apparently normal in axons.

Despite smaller soma sizes, we found no loss of neuron number and target tissues were surprisingly hyper-innervated in DNER cKO mice. Regulation of cell size is a poorly understood subject, but several intrinsic and extrinsic factors including access to certain growth factors, nutrients, and the overall genome size all play roles (Lloyd 2013).

These factors often affect signaling pathways and metabolic activity in cells to determine how large or small a cell may be. One of the primary signaling pathways involved in cell size determination is PI3K/mTOR/AKT signaling (Saxton and Sabatini 2017). In neurons where the tumor suppressor gene, PTEN (a negative regulator of mTOR), is knocked out, there is overactivation of mTOR and increased phosphorylated AKT which results in enlarged soma size as well as aberrant dendritic and axonal growth (Kwon et al. 2003;

Kwon et al. 2001; Backman et al. 2001). Inhibition of mTOR in animals with conditional deletion of PTEN in cerebellar and neurons prevents neuronal hypertrophy

(Kwon et al. 2006). Interestingly, a recent paper found DNER could act through

PI3K/AKT signaling in tissue (Wang et al. 2019). Though we observed opposing effects of DNER knockout on neuronal cell bodies versus axons, it would be of interest to determine the state of PI3K/mTOR/AKT signaling in these animals.

While abnormal mTOR signaling provides one potential mechanism for the changes in soma size and/or hyper-innervation, other possibilities exist that could explain increased axonal growth. One possible explanation is that the hyperinnervation is a result of overexcitation of DNER cKO neurons. We found that in response to depolarization,

DNER cKO neurons exhibit a sustained calcium response. This hyperactivation of

91 sympathetic neurons could induce excessive axonal sprouting. Neuronal activity has long been associated with fine tuning of axon branches and calcium itself plays roles in gene activation and cytoskeletal remodeling (Feller 1999). Increased intracellular calcium in response to depolarization could result in hypertrophy of axons, through one or more of these processes. Another possible explanation for excessive axon branching is changes in growth factor expression. Target-derived NGF is the primary dictator of final target innervation by sympathetic neurons, and alterations in NGF expression levels can modify innervation patterns (Korsching and Thoenen 1983). Intriguingly, it is possible that the over-activation of the neurons caused changes in distal NGF levels as sympathetic activity has been linked to growth factor expression levels in the pineal gland (Calinescu et al. 2011). How the loss of DNER contributes to neuronal hyperactivation is unclear, however, it is possible that downstream signaling from DNER could play a role in dysregulation of calcium dynamics and excitability of sympathetic neurons. In vivo measures of neuronal activity, growth factor expression, and exploration of DNER signaling components might provide additional support for these hypotheses.

Finally, we observed abnormal mitochondrial morphology in nearly 30 percent of

DNER cKO neurons compared to controls. Mitochondrial morphology and function are vital for neuronal health and survival. Alterations in mitochondria are linked to many neurological disorders including Parkinson’s disease, Alzheimer’s, and amyotrophic lateral sclerosis (Johri and Beal 2012). As the powerhouses of the cell, mitochondria provide the largest source of energy for cells. With neurons being some of the most energetically demanding cells in the body, proper mitochondrial function is essential. We found abnormally long mitochondria with increased cristae density, which could indicate

92 that DNER cKO neurons are more metabolically active as the primary transporters and channels involved in mitochondrial energy production lie within the cristae (Galloway and Yoon 2013). Indeed, increased cristae density in is correlated with increased energy production (Nielsen et al. 2017). Measurement of mitochondrial output and dynamics in future studies might shed light on this interesting observation.

While the effects of DNER on neuronal architecture and function could be separate phenomena’s, one intriguing hypothesis is that these effects are interconnected.

Hyperactivation is stressful to neurons over time. In order to compensate for extended activation, the neurons likely have to expend extra energy which could result in changes to metabolic machinery as we’ve observed with the DNER cKO mitochondria. In fact, similar changes in mitochondria have been previously reported in cell cultures of mouse embryonic fibroblasts during starvation where mitochondria elongated and had increased cristae density. It was hypothesized that these mitochondrial changes were a compensatory mechanism for these cells to prevent by maintaining ATP levels under stress (Gomes, Di Benedetto, and Scorrano 2011; Gomes and Scorrano 2011).

Therefore, it is possible that the changes in DNER cKO neurons might be a stress response due to over-activation.

An additional layer of complexity is added to these hypotheses when we take into account the lack of satellite glial cell contact with DNER cKO neurons. Glial cells have been well documented to control the neuronal microenvironment, and astrocytes in particular have been demonstrated to play active roles in regulating neuronal activity at synapses. Given the parallels between satellite glial cells and astrocytes in their molecular makeup and the fact that both cell types ensheathe synapses, it is important to parse out

93 whether or not the changes observed in DNER cKO neurons is a result of knockdown of

DNER itself and some role DNER may be playing within the neurons, or if these changes are a consequence of the loss of glial cell contact.

94 MATERIALS AND METHODS

Animal Husbandry

All animal procedures were performed in accordance with NIH and Johns Hopkins

University Animal Care and Use Committee (JHU ACUC) guidelines. Animals were kept in a 12-hour light:dark cycle with food and water ad libitum. Mice were kept in mixed

C57BL/6 and 129P backgrounds and animals of both sexes were used for analyses. TH-

Cre mice were gifts from Dr. C. Gerfen (NIH). Dnertm3a KO first allele (C57BL/6NTac-

Dnertm3a(EUCOMM)Hmgu/Ieg) mice were purchased from EMMA Repository (strain ID:

EM:08389). Dnertm3a mice were crossed to a ROSA26-FLPe mouse (Jackson strain number: 003946) to knock out the LacZ and neo cassette to generate the conditional

Dnertm3c allele (Dner-floxed) mice, which were then mated to TH-Cre in order to generate

TH-Cre;Dnerf/f (DNER cKO) mice. Littermate Dnerf/f mice were used as controls in all experiments and where applicable same sex animals were compared to each other.

Sprague-Dawley rats were purchased from Charles River (strain code: 400).

In vitro neurite extension

Neurons from superior cervical ganglia (SCG) of mice of indicated genotypes were dissected between P2 and P4, and enzymatically dissociated as previously described

(Bodmer et al., 2011). Briefly, pups were anesthetized according to JHU ACUC protocol.

SCGs of pups were dissected and separated from surrounding tissue, incubated for 20 minutes at 37°C in DMEM AIR (DMEM F12, 12.5mM glucose, 1U/ml penicillin/streptomycin) and supplemented with 4mg/ml collagenase IV, 1mg/ml hyaluronidase, 0.6mg/ml DNase I, and 10mg/ml BSA. Ganglia were centrifuged,

95 resuspended in DMEM air containing 3mg/ml trypsin, and incubated for an additional 30 minutes at 37°C. Cells were mechanically dissociated by tituration with a glass pipet, centrifuged, and resuspended in DMEM supplemented with 10% FBS, 1U/ml penicillin/streptomycin, and 30ng/ml NGF on coverslips coated with Poly-D-laminin mixture. Neurons were cultured for 16 hours prior to fixation with 4% PFA for 20 minutes, washed with PBS, and blocked with 1% BSA in PBS. Primary antibodies were diluted in block and incubated at room temperature for 2-3 hours. Following washes with

PBS, cells were additionally incubated in secondary antibodies for hour at room temp, washed, and mounted with fluorimount plus DAPI. Confocal images were taken at 20x with a Zeiss LSM 700 confocal microscope. Image analysis was performed using ImageJ.

Number of branches and neurite length were quantified where the length of the longest neurite was determined. At least 15 neurons from four individual experiments were used.

Immunohistochemistry and neuronal cell size

For IHC, mice at indicated ages were fixed in 4% PFA for 4-16 hours at 4°C, cryoprotected in 30% sucrose in PBS, embedded in OCT, and serially sectioned (12-

30um) with a Thermo Microtom HM 550 cryostat. Sections were blocked with 5% DS serum and 0.1% TX-100 in PBS for 1 hour and incubated in primary antibody overnight at 4°C. Sections were then washed with PBS, incubated with secondary antibody for 1 hour at room temperature, washed again, and mounted with fluorimount plus DAPI.

Images were taken with a Zeiss LSM 700 confocal microscope. ImageJ was used for cell size quantifications. For quantifications, 247-328 neurons from three individual animals were counted and binned according to size.

96

Neuron cell counts

P0, P7, P14, and P28 control and DNER cKO mice were prepped for immunohistochemistry as described above and serially sectioned at 12um thickness.

Sections were stained with a solution containing 0.5% (Nissl). Cells with characteristic neuronal morphology and visible nucleoli were counted in every fifth

Nissl-stained section. Total cell number was calculated by multiplying raw counts by 2.5 as previously described.

Quantitative real-time PCR

Sympathetic ganglia were homogenized in 100ul of Trizol for one animal (two SCGs).

RNA extraction was performed according to manufacturer's protocol (Invitrogen

15596026) using phenol:chloroform. For cDNA synthesis, 5ug of RNA were reverse transcribed with SuperScript IV Reverse Transcription following manufacturer’s protocol

(Thermo Scientific 18091050). qPCR was performed using Maxima SYBR green/ROX qPCR Master Mix (Thermo Scientific K0222) and detected using Applied Biosystems

StepOnePlus Real-Time PCR System (cat 4376600).

Transmission Electron Microscopy

Tissues were fixed in 3% formaldehyde, 1.5% glutaraldehyde, 2.5% sucrose, 5mM Ca2+ in 0.1M cacodylate buffer (pH 7.4), postfixed with 1% OsO4 then Kellenberger UA

(0.5% uranyl acetate). Tissues were embedded in Eponate resin, sectioned at a thickness of 90 nm, and collected on copper grids. Mitochondria were imaged using an FEI Tecnai-

97 12 TWIN transmission operating at 100 kV and a SIS MegaView III wide-angle camera at 26,500x magnification. For analysis, mitochondria were imaged from at least three neurons in five randomly selected grids. Area, cristae density, and mitochondrial content were determined using ImageJ.

iDISCO whole mount tissue staining iDISCO-based tissue clearing for whole mount immunostaining of organs from P14 and

P28 mice was performed as previously described. Briefly, hearts were fixed in 4%PFA, then dehydrated by methanol series (20-100%) and incubated overnight in 66% dichloromethane (DCM)/33% methanol. Samples were then bleached with 5% H2O2 in methanol at 4°C overnight, then rehydrated, permeabilized with 0.2%TritonX-100 for two hours, and followed by overnight permeabilization with 0.16% TritonX-

100/20%DMSO/0.3M glycine in PBS. Samples were incubated in blocking solution

(0.17% TritonX-100/10% DMSO/6% Normal Goat Serum in PBS) for 24 hours and incubated with rabbit anti-TH (1:500) in 0.2% Tween-20/0.001% heparin/5% DMSO/3% normal goat serum in PBS at 37°C for 72 hours. Samples were then washed with 0.2%

Tween-20/0.001% heparin in PBS and incubated with anti-rabbit Alexa-647 secondary antibody (1:500) in 0.2% Tween-20/0.001% heparin/3% normal goat serum in PBS. After

48 hours, organs were again washed with 0.2% Tween-20/0.001% heparin in PBS and dehydrated in methanol. Samples were cleared by successive washes in 66% DCM/ 33% methanol, 100% DCM and 100% Dibenzyl Ether. Hearts were tile scanned (3x3) at 10x using a Zeiss LSM 700 confocal microscope, and innervation density was determined

98 using raw integrated density of the TH signal. Innervation was normalized as a percent of control in which DNER cKO hearts were compared to litter and sex-matched controls.

In vivo norepinephrine measurement

SCGs and salivary glands from P0-P2 control and knocked animals were homogenized in

0.1N HCl, 1mM EDTA solution. Norepinephrine levels were measured by ELISA according to the manufacturer’s protocol (LDN Noradrenaline Research ELISA, BA E-

5200).

Calcium Imaging

Sympathetic explants from indicated genotypes and ages were cultured on collagen coated MatTek 35mm, No. 1.5 coverslip, 20mm glass diameter dishes in SCG media plus

100ng NGF for 14-16 hours. Cells were then washed in calcium imaging buffer (145mM

NaCl, 5mM KCl, 1mM MgCl2, 1mM CaCl2, 10mM glucose, 10mM sodium-HEPES, pH

7.4) and incubated with Fluo4-AM for 40 minutes. Cultures were again washed several times and then incubated in calcium imaging buffer on a stage incubator (37°C and 5%

CO2) for equilibration prior to imaging. Single plane images were taken every 3 seconds at 40x magnification on a Zeiss Axio Observer Yokogawa CSU-X1 spinning disk confocal equipped with dual Evolve EMCCDs and 405, 488, 555, and 633 nm lasers. Zen

Blue (2012) image collection software was used to maintain experimental parameters between individual experiments. Concentrated KCl (final concentration of 50mM) was added by hand after approximately 2 minutes of imaging for background collection, and imaging continued for 8 additional minutes following depolarization.

99 Videos were analyzed using ImageJ. Raw integrated density of the calcium signal was determined and normalized to area for each neuron. Area under the curve and the peak calcium response were calculated for each neuron and plotted separately.

Statistical analyses

Sample sizes were similar to those reported in previous publications (Armstrong et al.

2011; Patel et al. 2015). Data were collected randomly. For practical reasons, analyses of neuronal cell counts and cell size in mouse tissues were done in a semi-blinded manner such that the investigator was aware of the genotypes prior to the experiment, but conducted the staining and data analyses without knowing the genotypes of each sample.

All Student’s t tests were performed assuming Gaussian distribution, two-tailed, unpaired, and a confidence interval of 95%. Statistical analyses were based on at least 3 independent experiments and described in the figure legends. All error bars represent the standard error of the mean (s.e.m).

Antibodies

Antibody Company Dilution Use Rabbit anti-TrkA Millipore, 06-574 1:200 IHC Mouse anti-tyrosine Sigma, T2928 1:500 IHC hydroxylase Rabbit anti-tyrosine Millipore, AB152 1:500 IHC, ICC, hydroxylase iDISCO Mouse anti-βIII-tubulin Sigma, T8660 1:500 ICC Goat anti-rabbit Thermo Scientific, 1:500 IHC Alexafluor488 A11034 Goat anti-rabbit Thermo Scientific, 1:500 IHC Alexafluor546 A11035 Goat anti-rabbit Thermo Scientific, 1:500 IHC Alexafluor647 A21244

100 Goat anti-mouse IgG2b Thermo Scientific, 1:500 IHC Alexafluor546 A21143 Goat anti-mouse IgGG1 Thermo Scientific, 1:500 IHC Alexafluor488 A21121 Goat anti-mouse IgGG1 Thermo Scientific, 1:500 IHC Alexafluor546 A21123

101 Chapter Five: Genetic ablation of satellite glial

cells during sympathetic nervous system

development.

102 INTRODUCTION

Neurons and glial cells interact with each other in almost every aspect of nervous system development and function. In both the neocortex and the peripheral nervous system, neurons and glial cells are derived from the same initial pool of multipotent stem cells, and in the neocortex glial cells have been shown to assist in neuronal migration. For example, during formation of the neocortex, radial glial cells initially provide a stem cell pool for the generation of cortical neurons. Following the generation of neuronal progenitors, radial glial cells undergo a gliogenic switch to begin generating astrocytes and oligodendrocyte precursors (Farhy-Tselnicker and Allen 2018). Analogously, in the peripheral nervous system, neural crest cells generate neurons prior to . In addition to acting as a stem cell pool, radial glial cells also provide a scaffold for newly generated neurons to migrate towards the pial surface of the developing brain. Similarly, newborn granule cells use Bergmann glial cell processes to migrate in the developing cerebellum (Tohgo et al. 2006). These observations demonstrate the importance of neuron-glia contacts during development but contact between neurons and glial cells is also vital for nervous system function.

In the adult nervous system, glial cells associate with nearly every compartment of the neuron. At the axon, oligodendrocytes in the CNS and Schwann cells in the PNS form sheathes that assist in action potential propagation and assist in axon regeneration. Schwann cells are also known to prune excess axon tracts in an activity- dependent manner. Meanwhile astrocytes, the most abundant glial cell in the central nervous system, contact neuronal cell bodies, dendrites, and synapses to influence synapse formation and pruning as well as activity through neurotransmitter recycling and

103 uptake, and potassium buffering (Sofroniew and Vinters 2010). Analogously, satellite glial cells form a complete sheath around peripheral neuron cell bodies, dendrites, and synapses (Hanani 2010).

With their proximity to the cell body and the fact that they ensheathe synapses, satellite glial cells have been compared to astrocytes in the CNS. In fact, satellite glial cells express many of the same proteins as astrocytes, many of which are vital for astrocytic function. For example, satellite glial cells and astrocytes both express and inwardly-rectifying potassium channels (Hanani 2010). Astrocytes use these proteins in order to take in excess potassium during an action potential and buffer the through gap junctions between adjacent astrocytes. Dye injection into satellite glial cells has demonstrated their ability to couple in a similar manner, and in a sensory nervous system injury model upregulation of gap junctions in satellite glial cells increased coupling between the glial cells as well as between glia and neurons leading to (Hanani 2010; Kim et al. 2016).

Astrocytes also play roles in neurotransmitter recycling. For example, at glutamatergic synapses, astrocytes express the glutamate-aspartate transporter (GLAST) which uptakes excess glutamate at the synapse during activity to prevent overstimulation of neurons. The astrocytes turn the glutamate into glutamine through the enzyme glutamine synthetase, which can then be recycled back to the pre-synaptic neurons to produce more glutamate for the next excitation event (Bak, Schousboe, and

Waagepetersen 2006). The expression of neurotransmitter transporters and receptors is poorly understood in satellite glial cells of the sympathetic nervous system, but expression of glutamine synthetase and GLAST has been observed in sensory satellite

104 glial cells (Berger and Hediger 2000; Hanani 2005). It is therefore likely that satellite glial cells of the sympathetic nervous express acetylcholine transporters, though this has not yet been investigated.

Satellite glial cells have also been proposed to form a nerve-blood barrier similar to the blood-brain-barrier formed in part by astrocytes in the CNS (Kiernan 1996). The evidence for such a barrier is confusing and contradicting. For example, satellite glia may play a role in providing a blood-nerve barrier, though the evidence for this is contradicting and controversial. Injection of dyes into sympathetic ganglia has produced differing results where some researchers have found that dyes are able to label near the neuron surface while others found they were not able to (Hanani 2010). The permeability of the ganglia to dyes may be dependent on the region within the ganglia as one study found the presence of injected lanthanum in the extravascular space near small intensely fluorescent cells (SIF), but these ions were excluded from the areas around principal neurons (Chau and Lu 1996).

Finally, satellite glial cells have also been implicated in injury response that is parallel in many ways to reactive by astrocytes in the CNS. Upon injury to CNS neurons, astrocytes undergo both morphological and molecular changes. Electron microscopic studies have observed morphological changes to satellite glial cells following of sympathetic . One group found significant changes in satellite glial cell structure within 1-3 weeks of injury to sympathetic nerves. The satellite glial cell sheath thickened while the number of glial cell layers decreased (Dixon 1969).

Several groups found satellite glial cells extend processes into the space between retracting nerve terminals and postsynaptic neurons following injury, which suggests a

105 possible role for these cells in the synaptic stripping process (Matthews and Nelson 1975;

De Stefano et al. 2007; Del Signore et al. 2004). Others noted molecular changes in satellite glial cells following neuronal injury similar to astrocytes. One hallmark of the astrocytic reactive gliosis is the upregulation of glial fibrillary acidic protein (GFAP).

Interestingly, satellite glial cells in sympathetic ganglia naturally express low levels of

GFAP, but upon insult to sympathetic neurons several groups noted a dramatic increase in GFAP levels as well as another glial marker, vimentin (Del Signore et al. 2006; Elfvin et al. 1987; Hu and McLachlan 2004). The biological basis for these changes is still unknown, but they highlight the need for further study into the roles satellite glial cells play in sympathetic ganglia.

We previously found changes to sympathetic neuron architecture and function as well as loss of neuron-satellite glial cell contacts following neuronal deletion of DNER.

We asked if the neuronal changes observed were due to direct deletion of DNER or if they are secondary to the loss of glial cell contact. We used a diphtheria toxin (DT) to specifically ablate satellite glial cells, which express the diphtheria toxin receptor (DTR) following injection under the BLBP promoter (BLBP-CreER). With a modest increase in cell death following a 3-day injection paradigm, we found that loss of satellite glial cells caused sympathetic neuron cell body atrophy with no significant loss of numbers, and a downregulation in the expression of norepinephrine biosynthetic machinery. These results suggest a potential role for satellite glial cells in modulating neuronal metabolism.

106 RESULTS

The effect of satellite glial cell loss on sympathetic neurons

Careful characterization of sympathetic ganglia in DNER cKO mice revealed disruptions in both neuron and glial cell development, but it remains to be seen if these phenotypes are connected or distinct. One attractive hypothesis is that lack of satellite glial cell contacts with sympathetic neurons results in hyper-activation of neurons which leads to the increased innervation and mitochondrial phenotypes. To determine if this was the case, we used a genetic model to abolish satellite glial cells during development. We generated BLBP-CreER;iDTR mice, in which the receptor for diphtheria toxin (DT) is expressed in BLBP-positive cells upon tamoxifen administration (Buch et al. 2005).

When subjected to diphtheria toxin, translation in the cells expressing the receptor will cease and the cells will die. As BLBP is a glial cell marker and mice do not naturally express the receptor for DT, changes in sympathetic neurons can be attributed to the lack of glial cells. BLBP-CreER;iDTR pups were injected with tamoxifen at postnatal days 4 and 5 to induce CreER translocation to the nucleus and subsequent expression of DTR. At postnatal day 11, 12, and 13 pups were given doses of either vehicle (PBS) or 300ng of

DT and sacrificed at P14 (Figure 5.1a), the same developmental time in which the DNER cKO phenotypes were observed.

To first ensure that satellite glial cells were dying, we assessed cell death by flow cytometry. Sympathetic ganglia were dissociated using papain and incubated with a fluorescent-tagged Annexin-V to assess and a necrosis dye. There was considerably more apoptotic cell death observed from DT-injected sympathetic ganglia versus control (27.09% versus 13.69% apoptotic cells) (Figure 5.1b,c). It is important to

107 note that we could not distinguish between neurons and glial cells by flow cytometry.

However, when we assessed neuronal cell number by Nissl staining, we found no difference in total number of neurons (Figure 5.2a-c). Given that neurons and satellite glial cells make up the majority of sympathetic ganglia, and the fact that BLBP is expressed only in glial cells, we can infer that the increased cell death is likely due to glial cells. Despite the increase in cell death, observed changes in satellite glial cells and neurons with 3 days of injection were minor. Therefore, we extended the paradigm to 5 days of injection and all characterizations from here on were done with this paradigm. Of note, 5 days of DT injections did not alter neuronal cell number (Figure 5.2d-f).

To further characterize the effects of DT-injection on satellite glial cells, we used

IHC to visualize satellite glial cell morphology using BLBP expression. In vehicle injected sympathetic ganglia, we found strong BLBP expression in ring-like structures around TH-positive sympathetic neurons as observed in previous control animals (Figure

5.3a). In contrast, DT-injected sympathetic ganglia exhibited lower BLBP expression in patches with only a few complete ring-like structures (Figure 5.3b). These data suggest effective glial cell ablation with DT injection.

We then asked if ablation of satellite glial cells resulted in similar neuronal phenotypes to those observed in DNER cKO mice. To this end, we assessed neurotransmitter machinery expression, neuronal atrophy, activity, and target tissue innervation. In DNER cKO sympathetic ganglia, expression of the norepinephrine biosynthetic machinery was downregulated. We therefore assessed TH and DBH expression in DT-injected animals. By IHC, TH expression appeared normal between control and DT-injected animals (Figure 5.2c). In agreement with this observation, we

108 found no observable changes in TH or DBH transcript levels in DT-injected ganglia

(Figure 5.3c). Similarly, there with no changes in other neuronal markers, TrkA and

Phox2b (Figure 5.3c), suggesting that loss of satellite glial cells does not affect sympathetic neuron identity, at least following 5 days of DT injection.

We next asked if loss of satellite glial cells affects neuronal size and target innervation. Using TrkA IHC, we assessed neuronal cell size and found an increase in smaller diameter neurons at the expense of large diameter neurons in DT-injected animals

(Figure 5.4a-c) similar to what was observed in DNER cKO mice. In addition to neuronal cell body atrophy in DNER cKO mice, target tissues were hyper-innervated.

Given the similar reduction in neuron soma size in DT-injected sympathetic ganglia, we assessed sympathetic innervation of the hearts and salivary glands of DT-injected animals. Again, we used iDISCO whole-tissue staining for TH in the hearts of control and DT-injected mice. We found a modest increase in innervation of the hearts of DT- injected animals compared to controls (Figure 5.5a-c). This is in contrast to the striking increase in sympathetic innervation of the heart in DNER cKO mice. As injection of DT did not ablate all satellite glial cells, it is possible that neurons innervating the hearts of

DT-injected animals retained glial cell contacts. Additionally, our injection paradigm might be too short to observe dramatic changes in sympathetic innervation of the heart and an extended glial cell ablation paradigm may elucidate further changes in innervation. Interestingly, while innervation of the heart was only modestly increased in glial cell ablated mice, sympathetic innervation of the salivary glands was significantly increased using TH as a marker (Figure 5.5d-f). Salivary gland hyper-innervation was also observed using Tuj1 (Figure 5.5g-i), confirming that we are observing tissue hyper-

109 innervation and not just an increase in TH levels, similar to what we observed in DNER cKO mice. Together, these results suggest that neuronal cell body atrophy and target tissue hyper-innervation phenotypes observed in DNER cKO mice are secondary to the loss of neuron-satellite glial cell contact.

Finally, we assessed if loss of satellite glial cells in sympathetic ganglia affected neuronal activity. To this end, we employed the same paradigm used for DNER cKO neuron activity by measuring calcium levels using Fluo4-AM from control and DT- injected sympathetic explants in response to depolarization. Similar to what was observed in DNER cKO mice, neurons from DT-injected animals exhibited a sustained calcium response to depolarization (Figure 5.6a,b). However, in contrast to DNER cKO mice, calcium levels in neurons from DT-injected animals spiked higher than controls (Figure

5.6a,c). In DNER cKO mice, neuron-satellite glial cell association was disrupted, but satellite glial cells were still present in similar numbers to control animals. With DT- injected animals, we induced satellite glial cell death in sympathetic ganglia. The observed differences in calcium responses to depolarization may be explained by the physical loss of satellite glial cells versus loss of neuron-satellite glial cell association.

110

Figure 5.1. Increased cell death in DT-injected sympathetic ganglia.

(a) Schematic of satellite glial cell ablation paradigm. (b) Flow cytometry of cells from control ganglia and (c) DT-injected ganglia treated with a far-red necrosis dye and

Annexin-V conjugated to Enzo-gold shows more apoptotic cell death in DT-injected ganglia (13.69% control, 27.09% DT-injected).

111

Figure 5.2. No changes in neuronal cell number in DT-injected ganglia with 3 or 5 days of DT injection.

(a) Nissl staining of control and (b) 3-day DT-injected sympathetic ganglia. (c)

Quantification of total neuronal cell number shows no difference between ganglia (n=2 control, 3 3-day DT-injected ganglia). (d) Nissl staining of control and (e) 5-day DT- injected sympathetic ganglia. (f) Quantification of total neuronal cell number shows no difference between ganglia (n=3 control, 3 5-day DT-injected ganglia).

112

Figure 5.3. Loss of satellite glial cell ring structure in 5-day DT-injected sympathetic ganglia but no loss in neuronal identity.

(a) IHC of control and (b) 5-day DT-injected sympathetic ganglia for TH (green) and

BLBP (white in merge, black inverted on right). Loss of satellite glial cell rings and apparent downregulation in BLBP protein expression in DT-injected ganglia (scale bar:

25um). (c) Transcript levels for neuronal markers, TH, DBH, TrkA, and Phox2b show no changes in 5-day DT-injected ganglia (n=5 control, 6 DT-injected, student t-test, n.s.).

113

Figure 5.4. Loss of satellite glial cells results in neuronal cell body atrophy.

(a) TrkA IHC for control and (b) 5-day DT-injected sympathetic ganglia shows smaller neurons in DT-injected animals. (c) Quantification of neuronal size binned and shown as percent of neurons with an area of less than 100um, 100-200um, 200-300um, and greater than 300um. (n=3 control, 3 5-day DT-injected with 93-215 neurons counted per animal, student t-test, *p<0.05).

114

115 Figure 5.5. Loss of satellite glial cells during development results in hyper- innervation of sympathetic target tissues.

(a) Tyrosine hydroxylase staining (black) from control and (b) 5-day DT-injected hearts shows slightly increased innervation in DT-injected animals. Scale bar: 50um. (c)

Quantification of innervation shown as percent of sex-matched littermate controls. n=3 control, 4 5-day DT-injected hearts. (d) TH immunohistochemistry of salivary glands from control and (e) 5-day DT-injected animals. (f) Quantification by raw integrated density shows increased salivary gland innervation by TH in DT-injected animals. (n=5 control, 6 DT-injected animals, student t-test, *p<0.05). (g) Tuj1 immunohistochemistry of salivary glands from control and (h) 5-day DT-injected animals. (i) Quantification by raw integrated density shows increased salivary gland innervation by Tuj1 in DT-injected animals. (n=5 control, 6 DT-injected animals, student t-test, *p<0.05).

116

Figure 5.6. Sympathetic neurons have an altered response to depolarization with loss of satellite glial cells.

(a) Graph of relative Fluo4-AM signal over time normalized to background levels. (b)

Area under the curve (AUC) of Fluo4-AM response to depolarization. (c) Peak Fluo4-

AM signal following depolarization of neurons with 50mM KCl. DT-injected neuron calcium levels spike higher and are maintained for an extended period of time compared to control neurons following depolarization.

117 DISCUSSION

Following deletion of DNER from sympathetic neurons, we observed losses in neuron-glial cell contact as well as changes in neuronal architecture and function. We sought to determine which changes in neuronal structure were due to loss of DNER versus loss of satellite glial cells by specifically ablating satellite glial cells during development. We found that similar to DNER cKO mice, satellite glial cell ablated mice exhibited neuronal cell body atrophying and hyper-innervation of target tissues.

However, unlike in DNER cKO mice, we found no observable changes in levels of TH or

DBH. It is important to note that our glial cell ablated paradigm did not kill all satellite glial cells. After three days of DT injection, we observed only a modest increase in cell death from 13.69% to 27.09%. While extending the injection paradigm to five days allowed us to observe some neuronal changes, we cannot exclude the possibility that stronger ablation of satellite glial cells or an extended glial cell ablation paradigm may eventually alter sympathetic neuron transmitter machinery expression. Future studies should address this possibility.

The neuronal cell body atrophy and target tissue hyper-innervation phenotypes highlight a possible role for satellite glial cells in regulating neuronal structure and health.

Cell size is highly regulated in the body and large shifts in neuronal cell size could be indicative of changes in neuronal health and metabolism. It is possible that loss of satellite glial cell contacts with sympathetic neurons places stress on the neurons, which in turn leads to atrophying. In support of this, neuronal cell body atrophy is observed with aging and in disease states (Gemmell et al. 2012; Smith et al. 1999; Swaab et al. 1994).

Additionally, it is interesting to note that the number of satellite glial cells associated with

118 individual neurons is correlated with the neuronal cell body size (Hanani 2010).

However, it is unclear if neuronal size influences the number of satellite glial cell sheathes or vice versa. Careful dissection of how neuron size influences glial cell number around neurons, or vice versa, could help to elucidate this question as well as the identification of molecules that could be involved in this process.

As we observed in DNER cKO mice, the atrophying of the neuronal cell body is contrasted with an increase in target tissue innervation. This suggests that excessive target tissue innervation may also be a result of loss of satellite glial cell contact with neurons. Satellite glial cell-neuron contact may influence target tissue innervation in a number of ways. The first possibility is that contact signaling from satellite glial cells may provide a regulatory cue that assists in control of axonal sprouting. Indeed, astrocytes, oligodendrocytes, and Schwann cells have all been implicated in axon growth, innervation, and synapse formation. Satellite glial cells differ from other glial cell types in that they exist only in the cell body compartment of the sympathetic nervous system and as such control of distal innervation patterns might seem counterintuitive, however, recent evidence in drosophila suggest that soma-associated glial cells can influence neurite branching. Soma glial cell loss results in excessive neurite branching (Yadav et al.

2019), as does disruption of Nrg-Ank signaling between drosophila sensory neurons and glial cells (Yamamoto et al. 2006).

The second possibility, as discussed briefly in chapter four, is that target tissue hyper-innervation may be secondary to the loss of satellite glial cells as a consequence of increased neuron activity. Interestingly the excess neurite extension in drosophila without soma glial cells is correlated with aberrant neuron activity (Yadav et al. 2019). It was

119 postulated that loss of contacts with soma glia increased neuron sensitivity to stimuli which resulted in hyperactivation and increased neurite growth. The connection between neuronal activity and axon branching is limited and evidence suggests that activity can influence axon growth negatively and positively depending on the system. We similarly found neuron hyperactivity in response to depolarization with satellite glial cell ablation in addition to hyper-innervation. Therefore, we cannot discount the possibility that these phenotypes are connected, and it would be of interest for future research to determine if satellite glial cell influence on neuronal activity and axon branching are associated.

120 MATERIALS AND METHODS

Animal Husbandry

All animal procedures were performed in accordance with NIH and Johns Hopkins

University Animal Care and Use Committee (JHU ACUC) guidelines. Animals were kept in a 12-hour light:dark cycle with food and water ad libitum. Mice were kept in mixed

C57BL/6 and 129P backgrounds and animals of both sexes were used for analyses. iDTR

(C57BL/6-Gt(ROSA)26Sortm1(HBEGF)Awai/J, stock #007900) mice were obtained from Jax

Laboratory and initially generated by Ari Waisman, Johannes Gutenberg University of

Mainz (Buch et al. 2005). BLBP-CreER mice were generously provided by Dr. Toshihiko

Hosoya (RIKEN Brain Science Institute; Saitama, Japan). BLBP-CreER;iDTR were generated by mating BLBP-CreER with iDTR homozygous mice. Experiments were carried out with BLBP-CreER;iDTR+/-. TH-Cre mice were gifts from Dr. C. Gerfen (NIH).

Dnertm3a KO first allele (C57BL/6NTac-Dnertm3a(EUCOMM)Hmgu/Ieg) mice were purchased from EMMA Repository (strain ID: EM:08389). Dnertm3a mice were crossed to a

ROSA26-FLPe mouse (Jackson strain number: 003946) to knock out the LacZ and neo cassette to generate the conditional Dnertm3c allele (Dner-floxed) mice, which were then mated to TH-Cre in order to generate TH-Cre;Dnerf/f (DNER cKO) mice. Littermate

Dnerf/f mice were used as controls in all experiments and where applicable same sex animals were compared to each other.

Induction of satellite glial cell ablation

BLBP-CreER;iDTR pups were injected subcutaneously with 20ul of 20mg/mL tamoxifen

(Sigma, T5648) at P4 and P5. Following a 6-day chase, pups were injected

121 intraperitoneally with 300ng of diphtheria toxin (Sigma, D0564) or PBS for 3 consecutive days (P11, P12, and P13) and were sacrificed on the 4th day (P14) for analyses.

Flow cytometry

Sympathetic ganglia from control and DT injected animals were incubated in 30units/ml

Papain in HBSS for 30 minutes at 37°C, washed with HBSS, then titurated and filtered.

Cells were analyzed for necrosis and apoptosis detection using a GFP-Certified

Apoptosis/Necrosis Detection Kit (Enzo Life Sciences, ENZ-51002) according to manufacturer’s protocol. Flow cytometry was done using a BD FACSCanto High- throughput Flow Cytometer (BD Biosciences). Data were analyzed using FCS Express 7

(De Novo).

Immunohistochemistry and neuronal cell size

For IHC, mice at indicated ages were fixed in 4% PFA for 4-16 hours at 4°C, cryoprotected in 30% sucrose in PBS, embedded in OCT, and serially sectioned (12-

30um) with a Thermo Microtom HM 550 cryostat. Sections were blocked with 5% DS serum and 0.1% TX-100 in PBS for 1 hour and incubated in primary antibody overnight at 4°C. Sections were then washed with PBS, incubated with secondary antibody for 1 hour at room temperature, washed again, and mounted with fluorimount plus DAPI.

Images were taken with a Zeiss LSM 700 confocal microscope. ImageJ was used for cell size quantifications. For quantifications, 247-328 neurons from three individual animals were counted and binned according to size.

122

Neuron cell counts

P14 control and DT-injected mice were prepped for immunohistochemistry as described above and serially sectioned at 12um thickness. Sections were stained with a solution containing 0.5% cresyl violet (Nissl). Cells with characteristic neuronal morphology and visible nucleoli were counted in every fifth Nissl-stained section. Total cell number was calculated by multiplying raw counts by 2.5 as previously described.

Quantitative real-time PCR

Sympathetic ganglia were homogenized in 100ul of Trizol for one animal (two SCGs).

RNA extraction was performed according to manufacturer's protocol (Invitrogen

15596026) using phenol:chloroform. For cDNA synthesis, 5ug of RNA were reverse transcribed with SuperScript IV Reverse Transcription following manufacturer’s protocol

(Thermo Scientific 18091050). qPCR was performed using Maxima SYBR green/ROX qPCR Master Mix (Thermo Scientific K0222) and detected using Applied Biosystems

StepOnePlus Real-Time PCR System (cat 4376600).

iDISCO whole mount tissue staining iDISCO-based tissue clearing for whole mount immunostaining of organs from P14 and

P28 mice was performed as previously described. Briefly, hearts were fixed in 4%PFA, then dehydrated by methanol series (20-100%) and incubated overnight in 66% dichloromethane (DCM)/33% methanol. Samples were then bleached with 5% H2O2 in methanol at 4°C overnight, then rehydrated, permeabilized with 0.2%TritonX-100 for

123 two hours and followed by overnight permeabilization with 0.16% TritonX-

100/20%DMSO/0.3M glycine in PBS. Samples were incubated in blocking solution

(0.17% TritonX-100/10% DMSO/6% Normal Goat Serum in PBS) for 24 hours and incubated with rabbit anti-TH (1:500) in 0.2% Tween-20/0.001% heparin/5% DMSO/3% normal goat serum in PBS at 37°C for 72 hours. Samples were then washed with 0.2%

Tween-20/0.001% heparin in PBS and incubated with anti-rabbit Alexa-647 secondary antibody (1:500) in 0.2% Tween-20/0.001% heparin/3% normal goat serum in PBS. After

48 hours, organs were again washed with 0.2% Tween-20/0.001% heparin in PBS and dehydrated in methanol. Samples were cleared by successive washes in 66% DCM/ 33% methanol, 100% DCM and 100% Dibenzyl Ether. Hearts were tile scanned (3x3) at 10x using a Zeiss LSM 700 confocal microscope, and innervation density was determined using raw integrated density of the TH signal. Innervation was normalized as a percent of control in which Dner cKO hearts were compared to litter and sex-matched controls.

Calcium Imaging

Sympathetic explants from indicated genotypes and ages were cultured on collagen coated MatTek 35mm, No. 1.5 coverslip, 20mm glass diameter dishes in SCG media plus

100ng NGF for 14-16 hours. Cells were then washed in calcium imaging buffer (145mM

NaCl, 5mM KCl, 1mM MgCl2, 1mM CaCl2, 10mM glucose, 10mM sodium-HEPES, pH

7.4) and incubated with Fluo4-AM for 40 minutes. Cultures were again washed several times and then incubated in calcium imaging buffer on a stage incubator (37°C and 5%

CO2) for equilibration prior to imaging. Single plane images were taken every 3 seconds at 40x magnification on a Zeiss Axio Observer Yokogawa CSU-X1 spinning disk

124 confocal equipped with dual Evolve EMCCDs and 405, 488, 555, and 633 nm lasers. Zen

Blue (2012) image collection software was used to maintain experimental parameters between individual experiments. Concentrated KCl (final concentration of 50mM) was added by hand after approximately 2 minutes of imaging for background fluorescence collection, and imaging continued for 8 additional minutes following depolarization.

Videos were analyzed using ImageJ. Raw integrated density of the calcium signal was determined and normalized to area for each neuron. Area under the curve and the peak calcium response were calculated for each neuron and plotted separately.

Statistical analyses

Sample sizes were similar to those reported in previous publications (Armstrong et al.

2011; Patel et al. 2015). Data were collected randomly. For practical reasons, analyses of neuronal cell counts in mouse tissues were done in a semi-blinded manner such that the investigator was aware of the genotypes prior to the experiment, but conducted the staining and data analyses without knowing the genotypes of each sample. All Student’s t tests were performed assuming Gaussian distribution, two-tailed, unpaired, and a confidence interval of 95%. Statistical analyses were based on at least 3 independent experiments and described in the figure legends. All error bars represent the standard error of the mean (s.e.m).

Antibodies

Antibody Company Dilution Use Rabbit anti-BLBP Abcam, AB32423 1:200 IHC Mouse anti-BLBP Abcam, AB131137 1:500 IHC

125 Mouse anti-tyrosine Sigma, T2928 1:500 IHC hydroxylase Rabbit anti-tyrosine Millipore, AB152 1:500 IHC hydroxylase Goat anti-rabbit Thermo Scientific, 1:500 IHC Alexafluor488 A11034 Goat anti-rabbit Thermo Scientific, 1:500 IHC Alexafluor546 A11035 Goat anti-rabbit Thermo Scientific, 1:500 IHC Alexafluor647 A21244 Goat anti-mouse IgG2b Thermo Scientific, 1:500 IHC Alexafluor488 A21141 Goat anti-mouse IgG2b Thermo Scientific, 1:500 IHC Alexafluor546 A21143 Goat anti-mouse IgGG1 Thermo Scientific, 1:500 IHC Alexafluor488 A21121 Goat anti-mouse IgGG1 Thermo Scientific, 1:500 IHC Alexafluor546 A21123

126 CHAPTER SIX: Molecular Changes in DNER

cKO sympathetic ganglia

127 INTRODUCTION

Loss of DNER in sympathetic neurons reduces neuron-glia contacts and alters neuronal architecture and activity. Previous studies have identified DNER as a potential

Notch ligand, however, others have noted that DNER may act through other receptors as well. In co-immunopreciptation experiments, DNER is pulled down with Notch and

DNER can activate both canonical and non-canonical Notch signaling (Eiraku et al.

2005). A recent study in breast cancer tissue found that DNER can also interact with the

PI3K/AKT pathway, and in the zebrafish nervous system, DNER controls neural progenitor proliferation via an unidentified Notch-independent pathway (Hsieh et al.

2013; Wang et al. 2019). Given the numerous phenotypes observed in DNER cKO sympathetic ganglia and the limited data on the downstream signaling pathways of

DNER, we sought to identify any molecular changes in DNER cKO ganglia that could help explain the observed phenotypes.

Sympathetic ganglia are comprised primarily of neurons and satellite glial cells.

We used single cell drop-sequencing in order to parse out the molecular changes in both neurons and glia from DNER cKO mice. The use of single cell drop-sequencing allows us to determine the transcriptome of thousands of individual cells for a relatively cheap cost and in a high-throughput manner. The technique involves capturing single cells and a unique molecular barcode inside tiny aqueous droplets and then compartmentalizing them into a nanoliter-sized chamber (Macosko et al. 2015). By using single cell drop- sequencing, we can determine the transcriptomes of individual neurons and glial cells, and the use of the molecular barcode allows for the identification of an individual cell’s origin so that we may compare cells from control versus DNER cKO animals. This

128 strategy will allow us to identify molecular changes in neurons and glial cells from DNER cKO sympathetic ganglia that could help to explain the phenotypes we observed.

Through the use of single cell drop-sequencing, we identified 46 differentially regulated genes within neurons and 108 differently regulated genes within glial cells. The overall molecular changes were modest between control and DNER cKO ganglia, but we identified a few interesting transcripts that might be of interest for future studies. The most highly downregulated gene in glial cells was Cyr61, which has been implicated in migration and adhesion of cells. The loss of Cyr61 may help to explain the loss neuron- glial cell contacts in DNER cKO mice. Additionally, a long non-coding RNA Meg3 was downregulated in neurons and has been implicated in neurite growth. It would be exciting to look into the role of some of these genes in sympathetic ganglia development in future studies.

129 RESULTS

Molecular changes in DNER cKO sympathetic ganglia

We found changes in both neuron and glial cell morphology in DNER cKO sympathetic ganglia. Several of the neuronal phenotypes were found to be due to the loss of glial cell contact with sympathetic neurons as they were recapitulated in a satellite glial cell ablation mouse model. However, the mechanism of how DNER is functioning between sympathetic neurons and satellite glial cells is still a mystery. We used single cell drop sequencing in order to gain a better idea of how DNER may be functioning and what changes are going on molecularly in both neurons and glia in response to loss of

DNER. Sympathetic ganglia from P14 control and DNER cKO mice were used for analysis. We obtained 16,464 cells in total between the two genotypes (7837 control and

8627 DNER cKO). UMAP plotting of all the cells revealed 13 distinct clusters with 3 neuron and 4 glial cell groups that clustered together into two larger, but spatially discrete clusters (Figure 6.1a). The remaining 6 clusters were small and represented populations of fibroblasts, , endothelial cells, , and one cluster of glial cell progenitors. By comparing expression between control and DNER cKO ganglia, we found 46 differentially regulated neuronal genes (16 up, 30 down) and 108 differentially regulated glial cell genes (60 upregulated, 48 down) (Figure 6.1b). The differentially regulated genes were sorted into gene ontology terms (GO-terms). Within neurons we found genes involved in organization and organismal growth to be upregulated and genes involved in translation and activity as downregulated

(Figure 6.1c). In contrast, we found an upregulation in genes involved in translation, regulation of neuron development, organismal growth, and interestingly, Notch signaling,

130 as well as a down regulation in and genes in the glial cell cluster

(Figure 6.1d). We pulled out DNER as downregulated in neurons of DNER knockout animals as additional confirmation of our knockdown and it was amongst the top differentially regulated genes (Figure 6.1e). By identifying differentially regulated genes in the DNER knockout sympathetic ganglia, future studies will hopefully begin to parse out the mechanism by which DNER functions as well as some additional genes that may be important in sympathetic ganglia development.

131

132 Figure 6.1 Molecular changes in DNER cKO sympathetic ganglia.

(a) UMAP plot of all sequenced 16,464 cells (control and DNER cKO combined) separated by clusters of neurons (clusters 2, 5, and 12), glia (clusters 0, 1, 3, and 4), and other cell types. (b) UMAP plot of control (7837 cells) and DNER cKO (8627 cells) overlaid on top of each other. (c) GO-term analysis of differentially regulated neurons and (d) glial cells. (e) Table of select differentially regulated genes within neuron and glial cell clusters.

133 DISCUSSION

We identified molecular changes in both neurons and satellite glial cells of sympathetic ganglia from DNER cKO mice. Overall, transcriptional changes within both populations of cells were modest at best, which could suggest that our phenotypes are not due to large changes in overall transcriptional gene expression. It is interesting to note that a few changes in translation machinery were picked out in both glial and neuronal populations, so it is possible that larger changes may be observed at the protein level.

Despite the modest changes, a few genes stick out as particularly interesting. Among them, Cyr61 and Meg3 were downregulated in glial cells and neurons respectively. Cyr61 is secreted by Muller glial cells in the retina and promotes survival of photoreceptor cells in explants (Kucharska et al. 2014). This molecule has also been implicated in migration and adhesion of fibroblasts (Chen, Chen, and Lau 2001; Grzeszkiewicz et al. 2001; Chen,

Chen, and Lau 2000). Cyr61 was the most highly downregulated gene in satellite glial cells of DNER cKO ganglia and considering the loss of satellite glial cell contact with sympathetic neurons in these animals, it could be that Cyr61 plays a migration and adhesive role in satellite glial cells as well. Also, of note is the downregulation of Meg3 in DNER knockout sympathetic neurons. Meg3 is a long non-coding RNA implicated in neuronal plasticity through regulation of PI3K signaling as well as nerve growth through

Wnt signaling (Tan et al. 2017). Loss of Meg3 inhibits neuronal apoptosis and enhances nerve growth in rats following cerebral ischemia-reperfusion injury in part through elevation of growth factors (You and You 2019). Downregulation of Meg3 in DNER cKO mice could account for the hyper-innervation of target tissues and it would be of interest to explore this possibility in the future.

134 MATERIALS AND METHODS

Dissection of sympathetic ganglia

Sympathetic ganglia from 3 control or 3 DNER cKO animals were pooled together for each round of drop sequencing. Ganglia were incubated in 50units Papain diluted in

HBSS plus HEPES for 20 min at 37°C. Ganglia were then washed with HBSS plus

HEPES and incubated for an additional 20 min at 37°C with 1.5mg/mL collagenase in

HBSS plus HEPES. Following the second incubation, ganglia were washed again and then suspended and titurated in DMEM AIR (DMEM F12 supplemented with 12.5mM glucose and 1U/ml penicillin/streptomycin). Cells were centrifuged and resuspended in fresh DMEM AIR to a concentration of 100 cells/ul.

Drop-Seq single cell partitioning, library preparation, and sequencing

Single cell suspensions were diluted to a concentration of 100 cells/ul and processed with the Drop-Seq protocol as previously described (Macosko et al. 2015) including the following modifications. Flow rates for cells, beads, and oil were optimized for aquapel- treated PDMS devices purchased from FlowJem (cells and beads: 2,300ul/hour, oil:

13,000ul/hour). Up to two samples were processed in series, with single cell suspensions and stable emulsions held on ice until all collections were completed (no more than 1H) before proceeding immediately with reverse transcription. cDNA amplification was performed using 4000 beads/reaction with a total of 15 cycles of PCR. Tagmentation was performed as previously described and up to 5 libraries were multiplexed for sequencing on an Illumina NextSeq500 platform.

135 Drop-Seq data preprocessing

Reads were demultiplexed and aligned to the mouse genome (mm10), and digital gene expression matrices were generated using the Drop-Seq Tools 2.0.0 pipeline

(https://github.com/broadinstitute/Drop-seq/releases).

PCA, clustering, and differential gene expression analysis

Digital gene expression matrices for all samples were imported and processed with the

Seurat package (Satija, Farrell, Gennert, Schier, & Regev, 2015; Stuart et al., 2019). Cells containing fewer than 250 genes or greater than 10% mitochondrial gene content were removed prior to data normalization and scaling. Principal component analysis was used to identify major sources of variation within the dataset, and gene loadings within each

PC were manually inspected to ensure the capture of biologically relevant signals. For the initial round of clustering, 25 PCs were included as input to clustering and dimensionality reduction, resulting in the identification of 13 major cell classes. Cell types were identified by the overlay of canonical marker genes. Following cell type identification, each major cluster was isolated and analyzed using iterative rounds of PCA to identify finer substructure among each class. Differential gene expression testing (Wilcox rank sum test) was used to identify transcripts up or down regulated in control vs. mutant samples.

136 Chapter Seven: Closing Remarks

137 CLOSING REMARKS

This study provides evidence for cross- between sympathetic neurons and satellite glial cells during development. Loss of DNER in sympathetic neurons resulted in loss of neuron-glial cell contact during development and had profound effects on neuronal architecture, resulting in prolonged neuronal activation following stimulation, abnormal mitochondrial morphology, atrophying of the neuronal cell body, and hyper-innervation of target tissues. These effects on sympathetic neurons were, in part, due to the loss of satellite glial cells as ablation of satellite glial cells during development using diphtheria toxin resulted in mostly overlapping phenotypes such as atrophying of the neuronal cell body, hyper-innervation of target tissues, and altered neuronal function (Figure 7.1). There were a few notable differences between loss of

DNER versus ablation of satellite glial cells on sympathetic neurons; wherein loss of

DNER but not loss of satellite glial cells resulted in concomitant downregulation of neurotransmitter machinery levels, and ablation of satellite glial cells but not deletion of

DNER resulted in a profound increase in calcium levels within stimulated neurons. While the observations made in this study provide a basis for neuron-satellite glial cell cross- communication during development, future studies should further examine the specific role of satellite glial cells on instructing neuronal development.

Nerve growth factor as a master developmental regulator

Over the last several decades, studies have found NGF to be a master regulator for many aspects of sympathetic nervous system development. At the peripheral target tissues NGF promotes axonal growth and final target innervation; while back at the soma

138 NGF has been found to regulate gene transcription, neuronal survival, dendrite extension, and synapse formation. NGF has even been shown to promote myelination of axons. We have added to the field’s knowledge of NGF’s role in development by showing that NGF-

TrkA signaling is also important for satellite glial cell development. We found that satellite glial cell ensheathment of sympathetic neuron cell bodies is disrupted in mice lacking TrkA, and that this is at least in part due to the regulation of DNER expression.

Our characterization of satellite glial cells in the TrkA knockout mice was superficial and limited. It would be of interest for future studies to delve further into the role of NGF-

TrkA signaling in satellite glial cell development, addressing questions such as: does loss of NGF-TrkA signaling in neurons affect satellite glial cell number, proliferation, or death, and what other factors does NGF-TrkA regulate that may play a role in satellite glial cell development?

Communication between sympathetic neurons and satellite glial cells during development

Our data provides the first clue for how neurons and satellite glial cells communicate during development. Previous work in sympathetic nervous system development has largely ignored the presence of the closely associated neuronal neighbors, satellite glial cells. Our current understanding of the development of these cells comes from observational studies which identified the general progression of satellite glial cell development. We expanded upon this early work by identifying a gene,

DNER, that partially directs the glial cell ensheathment process. Loss of DNER in sympathetic neurons results in fewer satellite glial cells being associated with individual

139 neurons early in postnatal development. In vitro, DNER induces a morphological change in satellite glial cells in which their membranes expand from a bipolar shape to a flattened, round shape. We hypothesize that this change resembles a thinning of the glial cell cytoplasm that needs to occur as the glial cells wrap around the neuronal cell bodies and dendrites.

This work provides a basis for understanding how satellite glial cells develop around sympathetic neurons, however, many questions remain. While DNER has been shown to act through noncanonical Notch signaling in some systems, others have suggested that it may also act through other pathways, such as PI3K (Eiraku et al. 2005;

Wang et al. 2019; Hsieh et al. 2013). We found that indirectly blocking Notch signaling through the use of the gamma-secretase inhibitor, DAPT, could prevent the morphological change elicited by DNER treatment of satellite glial cells in vitro.

However, single cell sequencing from DNER cKO sympathetic ganglia did not show any strong downregulation of known Notch effectors, and in fact some Notch-associated genes were upregulated in DNER cKO glial cells. It would be of interest to further characterize possible DNER binding partners and it’s downstream signaling in satellite glial cells in the future.

More broadly, we have yet to identify what factors initially trigger glial cell differentiation as well as what players are involved in their proliferation. We found no defects in glial cell number or early postnatal glia proliferation DNER cKO ganglia.

Some literature suggests that molecules such as Sox2 and FGF2 may play roles in this early development of satellite glial cells as Sox2 promotes survival of satellite glia in vitro and FGF2 has been implicated in glial cell proliferation during development and

140 regeneration (Koike et al. 2015; Ribeiro-Resende et al. 2012; Grothe, Haastert, and

Jungnickel 2006; Klimaschewski, Meisinger, and Grothe 1999). Future studies may look into those molecules as potential players. Finally, loss of DNER did not result in a full loss of satellite glial cell ensheathment which suggests it may only be one piece to a larger puzzle. It would be interesting to explore other potential molecules involved in this process with a focus on cell adhesion molecules given the close association of the two cell types.

Insights into satellite glial cell function in sympathetic ganglia

To date few studies have investigated the roles of satellite glial cells in sympathetic ganglia. Focus on satellite glial cell function has primarily been in the sensory nervous system where their contribution to chronic pain is starting to be appreciated. Within sympathetic ganglia, several roles for satellite glial cells have been proposed. Suggested roles for satellite glia include evidence for a possible blood-nerve barrier, satellite glia regulation of the extracellular space around neurons and synapses through expression of neurotransmitter transporters, inwardly rectifying potassium (Kir) channels, and purinergic receptors, and in the engulfment of dying cells (Hanani 2010).

The expression of Kir channels in satellite glial cells suggests a role for these cells in potassium buffering similar to astrocytes in the CNS. Indeed, expression of Kir channels in satellite glia appears to be dependent on activity of sympathetic neurons, supporting the hypothesis that these cells may buffer the extra-synaptic space (Konishi 1996).

Overall, these potential roles have implicated satellite glial cells in regulation of the neuronal microenvironment similar to astrocytes yet the evidence in still limited.

141 Through our study of satellite glial cell development, we found evidence for new roles of these cells in shaping sympathetic neuron architecture and function. By disrupting the satellite glial cell sheathe either through deletion of DNER in sympathetic neurons or through direct ablation of satellite glial cells themselves, we found defects in neuronal structure and activity. Deprived of satellite glial cell support, sympathetic neurons were smaller. The atrophying of sympathetic neuron cell bodies with loss of satellite glial cells supports the idea that satellite glia are involved in regulating the neuronal microenvironment. Without support from surrounding glia, sympathetic neurons are vulnerable to atrophy. There are several possible explanations for this phenomenon.

One hypothesis is that satellite glial cells may provide trophic support to neurons through secretion of growth factors and nutrients, such as being a source for energy. Studies have found that satellite glial cells can express NGF under injury conditions and secretion of cytokines has been noted as well.

Another attractive hypothesis is that satellite glial cells play an important role in uptake of excess neurotransmitters and ions from synapses in order to prevent cytotoxicity of neurons. Satellite glia are known to express potassium channels and gap junctions similar to astrocytes, which play this role in the CNS, though expression of neurotransmitter reuptake machinery hasn’t yet been determined. In support of this, we found that loss of satellite glial cells altered the neuronal response to depolarization.

Without satellite glial cell support, either through loss of DNER or more dramatically through direct ablation of glial cells, sympathetic neurons exhibited a prolonged calcium response upon induced depolarization. This observation could be due to a loss in glial cell potassium buffering.

142 Lastly, we observed that loss of DNER and loss of satellite glial cell contact with sympathetic neurons resulted in hyperinnervation of target tissues. This suggests a possible role for satellite glial cells in influencing the innervation patterns of the neurons they contact. Whether this influence is driven through direct contact signaling and regulatory cues or potentially secondary through loss of control over the extracellular microenvironment is unknown, however, this intriguing possibility should be subject to further research.

Our work was limited to primarily describing satellite glial cell influence on sympathetic ganglia structure. While we have added to the growing literature exploring satellite glial cells, the functions and development of these cells remains largely unknown. Our data indicates that this area of research should explored more in depth and that these cells likely play important roles in sympathetic nervous system development, architecture, and function.

143

144 Figure 7.1. Tripartite interactions between peripheral target tissues, sympathetic neurons, and satellite glial cells during development.

In development of the sympathetic nervous system, growth factor signaling from peripheral target tissues, through secretion of NGF, controls development of sympathetic neurons and satellite glial cells. NGF-TrkA within the neurons turns on expression of

DNER, which signals to satellite glial cells and instructs ensheathment around sympathetic neuron cell bodies. Loss of DNER, or direct ablation of satellite glial cells through diphtheria toxin receptor expression and subsequent diphtheria toxin administration, results in atrophy of the neuronal cell body, aberrant neuron activity, and hyper-innervation of target tissues. Additionally, loss of DNER, but not ablation of satellite glial cells, also causes decreased expression of the norepinephrine biosynthetic machinery.

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167 Erica Boehm Phone: (603) 965-5676 | [email protected]

SUMMARY OF QUALIFICATIONS Experienced research scientist with advanced technical skills in cellular, molecular, and neurobiology, and . Strong written and oral communication skills through interactions with diverse audiences, representing all backgrounds and ages. Effective at project management and working collaboratively within a team or independently.

EDUCATION Ph.D. in Biology | January 2020 | Johns Hopkins University Thesis: Neuron-satellite glia interactions in sympathetic nervous system development • Johns Hopkins Department of Biology Retreat Poster Award – October 2017 • Relevant courses taken: Biology Science Writing, Writing Articles and Technical Reports, Communication for

B.S. in Biochemistry | May 2013 | University of New England Minor: neuroscience, summa cum laude • University of New England Department of Chemistry and Physics Outstanding Student, Biochemistry – April 2013 • University of New England Summer Undergraduate Research Experience Scholarship – June 2012 • University of New England Department of Chemistry and Physics Outstanding Student, Service – April 2012

PROFESSIONAL EXPERIENCE Medical Writer-Editor | DRT Strategies | June 2020 - current • Provide writing and editing assistance to FDA Center for Drug Evaluation and Research in contract with DRT Strategies

Graduate Student Researcher | Johns Hopkins University | Lab of Dr. Rejji Kuruvilla | May 2014-June 2020 • Pioneered and led a new area of research in the lab looking at satellite glial cell development in the sympathetic nervous system. • Developed and conducted experiments for analysis of sympathetic satellite glial cells via transmission electron microscopy, immunohistochemistry, and primary culture systems. • Presented research at yearly departmental seminars, through departmental poster sessions, and at a national Gordon Conference. • Worked closely with peers and collaborators to contribute to the publication of three peer- reviewed articles with one additional review article in preparation for Nature Reviews Neuroscience, and one first-author publication also in preparation. • Relied upon to edit grant applications and papers for lab members and colleagues. • Acted as a mentor for 2 high school, 4 undergraduate, and 4 graduate students, resulting in successful poster presentations and short seminars within the department and at a national leadership conference. • Led biochemistry and lab courses and was a teaching assistant for five years in , biochemistry, and .

Undergraduate Researcher | University of New England | Lab of Dr. Steven Johnson | May 2012-May 2013

168 • Cloned, expressed, and purified the putative phytase, PXO_04860, and developed assays to assess function. • Awarded a grant to conduct summer research. • Presented research at a summer symposium to a broad audience.

Undergraduate Researcher | University of New England | Lab of Dr. Stephen Fox | January 2011-May 2012 • Used naphthyridine derivatives to bridge two sulfur groups between a di-copper bridge as a model for the Copper (A) electron transfer molecule.

OUTREACH AND LEADERSHIP EXPERIENCE

Maryland Zoo in Baltimore March 2016 - Present • Promote conservation efforts and science through education tables and animal handling.

Graduate Student Representative January 2017 - December • Acted as a Biology representative for graduate student rights 2018 and interest. FREELANCE AND SAMPLE WRITING

Male Sex Have a Role in Asthma: https://www.hopkinsmedicine.org/news/newsroom/news-releases/male-sex-hormones- have-a-role-in-asthma

Tips to Avoid the Dreaded Burn Out: https://tipbox.abcam.com/tips-to-avoid-the-dreaded- burn-out/

RESEARCH PUBLICATIONS

Ceasrine AM., Lin EE., Lumelsky DN., Ruiz Otero N, Boehm ED., Kuruvilla R. (2018). Tamoxifen improves glucose tolerances in a delivery, sex, and strain-dependent manner in mice. Endocrinology, https://doi.org/10.1210/en.2018-00985

Otis JP., Zeitun EM., Theirer JH., Anderson JL., Brown AC., Boehm ED., Cerchione DM., Ceasrine AM., Arraham-Davidi I., Templehof H., Yaniv K., Farber SA. (2015) Zebrafish as a model for apolipoprotein biology: comprehensive expression analysis and a role for ApoA-IV in regulating food intake. Dis Model Mech. 8(3), 295-309.

Patel A., Yamashita N., Ascano M., Bodmer D., Boehm E., Bodkin-Clarke C., Ryu YK., Kuruvilla R. (2015) RCAN1 links impaired neurotrophin trafficking to abberant development of the sympathetic nervous system in Down Syndrome. Nature , 2015, Dec 14;6:10119

PRESENTATIONS

Boehm E., Mapps A., Connor B., Kuruvilla R. (2017) Neuron-satellite glia interactions in the sympathetic nervous system. Glial Biology: Functional Interactions Among Glia and Neurons, Gordon Conference, Ventura, CA. Poster.

Boehm E., Mapps A., Connor B., Kuruvilla R. (2017) Neuron-glia interactions in sympathetic nervous system development. Johns Hopkins Department of Biology Retreat, Hershey, PA. Poster.

169

Boehm E., Maille R., Johnson S. (2012) Cloning and expression of a putative phytase from Xanthamonas Oryzae in E. coli for use as an animal feed additive. University of New England Summer Undergraduate Research Symposium, Biddeford, ME. Poster.

TEACHING EXPERIENCE

Tutor, Monument City Tutoring December 2017 – June • Tutor high school students in SAT/ACT/SAT II Subject test prep 2020 Private Tutor September 2017 – June • Tutored high school students in general chemistry and 2018 trigonometry

REFERENCES

Available upon request.

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