Macrophage Accumulation Near Injured Neuronal Cell Bodies Is Necessary And

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MACROPHAGE ACCUMULATION NEAR INJURED

NEURONAL CELL BODIES IS NECESSARY AND

SUFFICIENT FOR PERIPHERAL AXON

REGENERATION

JON PAUL NIEMI

Submitted in partial fulfillment of the requirements

for the degree of Doctor of Philosophy

Dissertation Advisor: Dr. Richard E. Zigmond

Department of Neurosciences

CASE WESTERN RESERVE UNIVERSITY

January 2017 CASE WESTERN RESERVE UNIVERSITY

SCHOOL OF GRADUATE STUDIES

We hereby approve the thesis/dissertation of

Jon Paul Niemi

candidate for the degree of Doctor of Philosophy *.

Committee Chair

Dr. Heather Broihier

Committee Member

Dr. Richard E. Zigmond

Committee Member

Dr. Gary Landreth

Committee Member

Dr. Ruth Siegel

Date of Defense

July 11th, 2016

*We also certify that written approval has been obtained

for any proprietary material contained there in Dedication

To my wife, my son, and my parents, thank you for your love and support.

Table of Contents

DEDICATION ...... iI

TABLE OF CONTENTS ...... ii I

LIST OF FIGURES ...... ivV

ACKNOWLEDGMENTS ...... vIiI

CHAPTER 1: GENERAL INTRODUCTION ...... 1

1.1 PERIPHERAL AND CENTRAL NERVOUS SYSTEM INJURY ...... 2

1.2 EXTRINSIC MECHANISMS SUPPORTING PNS REGENERATION ...... 3

1.3 SLOW WALLERIAN DEGENERATION MOUSE (WLDS) ...... 5

1.4 INTRINSIC MECHANISMS SUPPORTING PNS REGENERATION ...... 10

1.5 MACROPHAGE RESPONSE TO INJURY ...... 16

1.6 MACROPHAGE POLARIZATION ...... 20

1.7 ROLE OF MACROPHAGES IN AXONAL REGENERATION ...... 23

1.8 FOCUS OF THESIS ...... 28

CHAPTER 2: A CRITICAL ROLE FOR MACROPHAGES NEAR

AXOTOMIZED CELL BODIES IN STIMULATING NERVE REGENERATION

...... 29

2.1 ACKNOWLEDGEMENTS ...... 30

2.2 ABSTRACT ...... 31

2.3 INTRODUCTION ...... 32

2.4 MATERIALS AND METHODS ...... 33

2.5 RESULTS ...... 39

2.6 DISCUSSION ...... 45

CHAPTER 3: OVEREXPRESSION OF THE MONOCYTE CHEMOKINE CCL2

IN DORSAL ROOT GANGLION NEURONS CAUSES A CONDITIONING-LIKE

INCREASE IN NEURITE OUTGROWTH AND DOES SO VIA A STAT3

DEPEDNDENT MECHANISM ...... 70

3.1 ACKNOWLEDGEMENTS ...... 71

3.2 ABSTRACT ...... 72

3.3 INTRODUCTION ...... 73

3.4 MATERIALS AND METHODS ...... 75

3.5 RESULTS ...... 82

3.6 DISCUSSION ...... 89

CHAPTER 4: DISCUSSION ...... 110

4.1 CCL2: MORE THAN JUST A CHEMOKINE ...... 111

4.2 MACROPHAGES AND REGENERATION: NEW SITE OF ACTION ...... 115

4.3 PRO-REGENERATIVE MACROPHAGE MECHANISMS ...... 119

4.4 TRANSLATIONAL ASPECTS ...... 123

4.5 FUTURE DIRECTIONS ...... 126

4.6 CONCLUDING REMARKS ...... 128

REFERENCES ...... 130

iii

List of Figures

CHAPTER 2

FIGURE 2.1 MACROPHAGE ACCUMULATION IN THE SCIATIC NERVE OF WLDS MICE .... 52

FIGURE 2.2 MACROPHAGE ACCUMULATION IN THE SCG AND DRG OF WLDS MICE .... 54

FIGURE 2.3 CCL2 MRNA EXPRESSION IN WLDS AND CCR2 -/- MICE ...... 56

FIGURE 2.4 MACROPHAGE ACCUMULATION IN THE SCIATIC NERVE, DRG, AND SCG OF

CCR2 -/- MICE ...... 58

FIGURE 2.5 MYELIN CLEARANCE IN THE SCIATIC NERVE OF WT, WLDS, AND CCR2 -/-

MICE ...... 60

FIGURE 2.6 SALIVARY GLAND INNERVATION IN WT, WLDS, AND CCR2 -/- MICE ...... 62

FIGURE 2.7 DRG AND SCG CONDITIONING LESION RESPONSE IN EXPLANT CULTURE

FROM WT AND WLDS MICE ...... 64

FIGURE 2.8 DRG AND SCG CONDITIONING LESION RESPONSE IN EXPLANT CULTURE

FROM WT AND CCR2 -/- MICE ...... 66

FIGURE 2.9 DRG AND SCG CONDITIONING LESION RESPONSE IN DISSOCIATED

NEURONAL CELL CULTURE FROM WT, WLDS, AND CCR2 -/- MICE ...... 68

CHAPTER 3

FIGURE 3.1 YFP LOCALIZATION AND CCL2 MRNA OVEREXPRESSION IN DRGS

FOLLOWING INTRATHECAL INJECTION OF AAV5 ...... 96

FIGURE 3.2 TIME-DEPENDENT INCREASE IN MACROPHAGE ACCUMULATION IN L5 DRGS

FOLLOWING CCL2 OVEREXPRESSION ...... 98

FIGURE 3.3 CCL2 OVEREXPRESSION INDUCES A CONDITIONING-LIKE INCREASE IN

NEURITE OUTGROWTH ...... 100

FIGURE 3.4 MACROPHAGE POLARIZATION MARKERS IN THE DRG FOLLOWING CCL2

OVEREXPRESSION ...... 102

FIGURE 3.5 CCL2 OVEREXPRESSION IN CCR2 -/- MICE ...... 104

FIGURE 3.6 CCL2 OVEREXPRESSION INDUCED RAG EXPRESSION IN THE DRG ...... 106

FIGURE 3.7 PHARMACOLOGICAL STAT3 INHIBITION IN CCL2 OVEREXPRESSION

DISSOCIATED CELL CULTURE ...... 108

Acknowledgments

The work presented here would not have been possible without the help and support of many people. First, I would like to thank my advisor Dr. Richard E. Zigmond.

He has truly taught me the importance of having passion for the research you are performing. Richard has shown me that perseverance through the tougher times, whether it is a failed experiment or being turned down for funding, makes the victories that much more rewarding. As a mentor, he put an emphasis on being able to effectively communicate your science and encouraged me to present my research any chance I got, which has been integral in preparing me for a future in teaching and research.

I would also like to thank my committee members Dr. Heather Broihier, Dr. Gary

Landreth, and Dr. Ruth Siegel for their insight, direction, and support. Our meetings helped me see the big picture while I was lost in the details. I am truly grateful to have been mentored by such excellent and respected scientists.

I could not have completed this project, and had so much fun while doing it, without the help and support of the numerous members of the Zigmond Lab. Alicia

DeFrancesco, our lab manager, truly keeps the lab running smoothly. She is an excellent scientist and an even better person. Jane Lindborg was always there to help with experiments, give advice on data, and keep our lab decorated for Halloween year-round.

Many other members of the lab, including Dan Mandell, Lilinete Roldan-Hernandez,

Madeline Howarth, and Angela Filous, contributed their time, effort, and knowledge to this project. I will forever be honored to be considered a Lablander.

The Department of Neurosciences at Case Western Reserve University provides an excellent training environment for young scientists. I would like to thank the members of the Neurosciences Administrative Office. Nothing we do would be possible without

Katie Wervey, Pam Capasso, and Narlene Brown. I would also like to thank Maryanne

Pendergast, of the Neurosciences Imaging Facility, for her assistance and helpful guidance throughout this process. This department truly fosters collaboration between labs. One such collaboration, with Jared Cregg a member of the Silver Lab, was instrumental in the completion of this work. I would like to thank Jared for his assistance and helpful discussion over the last 6 years. I have no doubt that you will be an excellent scientist.

To my parents, Bob and Jeanne Niemi, thank you for everything you have done for me. From raising me in a loving home to encouraging me to reach for the stars, I could not have done this without you.

Most importantly, I have to thank my wife and best friend, Rachel. No one has had to endure more over the past 6 years through all of the ups and downs that come with research. I am forever grateful for her unwavering support and encoragement. She gave me the best gift in the entire world, our son, Braxton. As soon as he entered the world it instantly changed my life. Everything I do, day in and day out is for Braxton. The love and laughs Rachel and Braxton provide me with daily is all I need in life. I can’t wait to see what our future holds.

vii

Macrophage Accumulation Near Injured Neuronal Cell Bodies is Necessary and

Sufficient for Peripheral Axon Regeneration

Abstract

JON PAUL NIEMI

Neuroinflammation plays a critical role in peripheral nerve regeneration. An injury to the sciatic nerve leads to macrophage accumulation in the L5 dorsal root ganglion (DRG), an effect notably not seen when axons in the L5 dorsal root are injured.

We have found that macrophage accumulation around axotomized DRG cell bodies is necessary for the conditioning lesion response. After axotomy, DRG neurons upregulate expression of CCL2, a macrophage chemokine, which acts on the receptor CCR2. We found that in a CCR2 knockout mouse, macrophage accumulation is inhibited after injury in the distal sciatic nerve and in the DRG. To determine the effect of this lack of macrophage accumulation on regeneration, DRGs were placed in explant culture 1 week after a conditioning lesion, and the increase in outgrowth seen in previously lesioned

DRGs from wild type mice was not seen in CCR2 -/- mice. These data demonstrate the role CCL2/CCR2 signaling plays in mediating macrophage accumulation in DRGs and suggest a relationship between macrophage accumulation near neuronal cell bodies and the regenerative capacity of neurons. Axotomy leads to many changes in DRGs. To probe

viii the importance of CCL2 upregulation alone, we overexpressed CCL2 in DRG neurons of uninjured mice using a viral vector (AAV5) and asked whether this is sufficient to cause macrophage accumulation and to enhance regeneration or whether other injury-induced signals are necessary. Two different viral constructs were used: an experimental virus coding for CCL2 and a control virus coding for YFP. The viral constructs were delivered intrathecally. AAV5-CCL2 led to a time-dependent increase in both CCL2 mRNA expression and macrophage accumulation in the L5 DRG compared to the YFP control virus. In addition, CCL2 overexpression led to a conditioning-like increase in neurite outgrowth in both DRG explants and in dissociated DRG neurons. AAV-CCL2 also increased LIF mRNA in the DRG and increased neuronal phospho-STAT3. Blockade of

STAT3 activation by the inhibitor STATTIC completely ablated the CCL2 overexpression-induced increase in axonal regeneration. Together, our data suggests that neuronal CCL2 expression and macrophage accumulation within the DRG are both necessary and sufficient for peripheral axonal regeneration to occur.

Chapter 1: General Introduction

Sections reprinted with permission from Neuroscience, Copyright, 2015.

1.1 Peripheral and Central Nervous System Injury and Regeneration

It has long been known that the peripheral nervous system and central nervous system

(CNS) respond very differently to injury. Following an axonal injury in the peripheral nervous system (PNS), axons distal to the site of injury degenerate in a process referred to as Wallerian degeneration, while axons proximal to the site of injury extend growth cones and allow for the injured axons to regenerate (Waller, 1852; Zochodne, 2008).

Injury in the CNS, which is largely studied through injury to the optic nerve, spinal cord, and traumatic brain injury, result in dieback of axons from the injury site, extremely limited regeneration, and no recovery of function (Vargas and Barres, 2007; Benowitz and Yin, 2008; Cregg et al., 2014).

This discrepancy is highlighted by neurons which are located in the dorsal root ganglia (DRG), which extend one axonal branch into the PNS and another branch into the

CNS. Injury to the peripheral but not the central branch results in axonal regeneration

(Schwab and Bartholdi, 1996). This differing response can be explained by the presence of significant gene expression changes in injured neurons and extrinsic clearance mechanisms which remove inhibitory molecules which occur after PNS but not CNS injury (Zigmond, 2012; DeFrancesco-Lisowitz et al., 2014). Thus, understanding the mechanisms which support peripheral nerve regeneration could lead to novel therapeutic targets to promote functional recovery after CNS injury and may lead to the ability to achieve more complete regeneration after PNS injury.

Peripheral nerve injuries are common, with damage to at least one peripheral nerve found in almost 3% of trauma patients (Noble et al., 1998), resulting in approximately

200,000 new patients a year (Taylor et al., 2008). The only treatment for patients with nerve transections is surgical where the two ends of the damaged nerve are sutured together to promote directional regrowth back toward the denervated target. Despite advances in surgical techniques only 10% of patients recover movement in the injured limb (Scholz et al., 2009). This poor functional outcome is most often attributed to the slow regeneration of damaged axons, which has led researchers to seek novel approaches to enhance axon regeneration after injury.

1.2 Extrinsic Mechanisms Supporting PNS Regeneration

Nerve injuries are powerful stimuli that lead to profound cellular responses.

Following an injury, axons and myelin sheaths distal to the lesion site are degraded by a process known as Wallerian degeneration (Glass, 2004; Makwana and Raivich, 2005).

Myelin breaks down to vesicles, resulting in the collapse of the myelin sheath (Ngeow,

2010). Schwann cell cytoplasm withdraws from the myelin vesicles and significantly decreases the synthesis of myelin lipids and proteins between 12 and 48 hours post injury

(Ngeow, 2010). Such damage increases the permeability of the blood-nerve barrier, which allows for the recruitment of macrophages to the site of the injured nerve

(Harrisingh et al., 2004; Napoli et al., 2012). Infiltrating macrophages and injury- activated Schwann cells phagocytose the degenerative end products (Stoll and Muller,

1999). Wallerian degeneration takes place during the first few days after injury. During this stage, elimination of myelin sheaths is important, because it clears the regeneration- inhibitory factors associated with myelin (Skaper, 2005; Raivich and Makwana, 2007).

At the same time, retrograde degeneration also takes place at a short segment of the proximal nerve stump. The remaining axons in the proximal nerve also exhibit a

3 reduction in diameter, followed by chromatolysis at the neuron soma and dendritic arbor retraction (Hanz and Fainzilber, 2006; Navarro et al., 2007). Chromatolysis, characterized by the loss and dispersion of the Nissl bodies, reflects a reactive change in neuronal biochemistry and function, when the neurons shift their functions from the synthesis of proteins required for neurotransmission to those required for regenerative axon growth (Deumens et al., 2010).

Loss of axonal contact also triggers dedifferentiation and proliferation of Schwann cells in the distal nerve (Karanth et al., 2006; Navarro et al., 2007; Napoli et al., 2012).

Proliferated Schwann cells line up in bands of Bungner, which provide support for regenerating axons (Geuna et al., 2003; Geuna et al., 2009). Schwann cells not only pave the way for regenerating axons to grow, they also attract the injured axonal growth cones by secreting neurotrophic factors, such as nerve growth factor (NGF; Ngeow, 2010).

Proximal to the lesion, fine sprouts emerge (Witzel et al., 2005) and using distal endoneurial tubes as a guiding structure, elongate in association with Schwann cells toward targets (Stoll and Muller, 1999; Navarro et al., 2007; Cattin et al., 2015). In the absence of a guiding structure, regenerating axons may form neuromas, a growth composed of immature axonal sprouts and connective tissue (Cattin et al., 2015).

Finally, regenerated axons reconnect with their peripheral target tissue. Because several sprouts emerge from each parent axon, many sprouts will be withdrawn gradually during the maturation of the fiber (Witzel et al., 2005; Navarro et al., 2007). The regenerated axons will have a smaller caliber and shorter internodes than normal uninjured nerve structures (Geuna et al., 2009). For complete regeneration and functional recovery, the axons are expected to replace the distal nerve segment lost during

4 degeneration. However, more often than not, the regenerated axons do not innervate target tissues adequately or relay information from sensory receptors accurately, reducing the recovery of motor and sensory functions, especially when the lesion is severe (Choi et al., 2005; Bannerman and James, 2009). Usually, after nerve injury and repair, the diameter of regenerated axons, as well as their conduction velocity and excitability remain below normal levels. Consequently, this results in incomplete and inadequate functional recovery of reinnervated tissues (van Meeteren et al., 1997; Xiao et al., 2007).

1.3 Slow Wallerian Degeneration Mouse (Wlds)

Following axonal injury, the axon distal to the injury site no longer receives proteins and mRNA from the cell body (Conforti et al., 2014). Originally Wallerian degeneration was considered to be a passive process induced by the lack of nutrients and cell-body derived factors following injury (Waller, 1850). However, discovery of the slow

Wallerian degeneration mouse (Wlds) led to the realization that axons degenerate through an active process (Lunn et al., 1989). In this mutant mouse, axons distal to the site of injury remain intact and are able to conduct action potentials, when stimulated, for up to

10 times longer than their wild-type (WT) counterparts (Lunn et al., 1989; Coleman and

Freeman, 2010). Early studies on degeneration, following neurotrophin deprivation or injury, reported that Wallerian Degeneration occurs without activation of caspase-3, suggesting that axons degenerate through an active process separate from apoptosis

(Buckmaster et al., 1995; Finn et al., 2000; Raff et al., 2002); however, more recent work on embryonic sensory neurons in vitro indicates that caspase-3 and -6 are involved in axon degeneration caused by trophic factor withdrawal but not by axotomy (Simon et al.,

2012).

The Wlds mouse is characterized by a genetic mutation which brings two genes, E4 ubiquitin ligase (Ube4b) and nicotinamide mononucleotide adenylyl transferase

(Nmnat1), together to form a fusion protein as a result of mRNA splicing (Lyon et al.,

1993; Coleman et al., 1998). The mutation was identified as an 85kb tandem triplication likely involving nonhomologous recombination to form the genetic rearrangement

(Coleman et al., 1998). Ube4b is an ubiquitin ligase involved in multi-ubiquitin chain assembly (Koegl et al., 1999). Nmnat1 is a nuclear enzyme necessary for the production of nicotinamide adenine dinucleotide (NAD+) (Schweiger et al., 2001). NAD+ and nicotinamide adenine dinucleotide phosphate (NADP) are best known for their roles in the electron transport chain and energy metabolism. Additionally, recent work has described them as signal transducers for their use as precursors to signaling molecules and as substrates for protein modifications, such as protein kinase B (Akt) and mitogen- activated protein kinase (MAPK; Clerk et al., 1998; Griendling et al., 2000; Pollak et al.,

2007). The resulting Wlds protein contains the full length Nmnat1 protein with 70 amino acids from Ube4b on the N-terminal end, where only Nmnat1 is enzymatically active

(Laser et al., 2003). An additional 18 amino acids from the 5’ UTR of Nmnat1 are also included in the protein (Samsam et al., 2003). The unique genomic rearrangement that led to the creation of the Wlds protein maintains normal expression levels of endogenous

Ube4b and Nmnat1 genes (Gillingwater et al., 2002). The key question surrounding the

Wlds mouse has been how this fusion protein is able to protect axons from axonal degeneration.

A major observation in the Wlds mouse was a complete lack of early infiltrating macrophages into the nerve distal to the injury site, which was initially thought to be the

6 cause of the slow degeneration (Brown et al., 1991a). However, transplantation experiments that provided the mutant mouse with WT myeloid cells did not rescue the phenotype (Perry et al., 1990; Glass et al., 1993). This study provided evidence that the slow degeneration phenotype of the Wlds mouse was not the result of altered macrophages but instead was inherent to the neurons/axons themselves.

Much of the research on the Wlds mouse has focused on which part(s) of the fusion protein is critical for conveying its protective effects. Overexpression of the 70 amino acids from Ube4b or Nmnat1 alone did not convey axonal protection when tested in vivo

(Conforti et al., 2007), yet in vitro studies suggested Nmnat1 overexpression could protect severed neurites in primary culture (Araki et al., 2004; Wang et al., 2005a). This discrepancy led to a model that suggested the two major domains of the protein are both necessary for the phenotypic protection. To further address this, a variant of the Wlds protein was created that lacked the enzymatic activity of Nmnat1 and resulted in a significant loss of axonal protection (Conforti et al., 2009; Yahata et al., 2009). Thus,

Nmnat1 is necessary for the neuroprotective effects of the Wlds protein yet is not sufficient to convey this protection on its own, in vivo (Conforti et al., 2007).

Through the development of an antibody against the Wlds protein, it was shown that the protein localized almost entirely to the nucleus, with barely detectable levels in the cytoplasm or axon (Samsam et al., 2003; Gillingwater et al., 2006; Beirowski et al.,

2009). With a strong localization in the nucleus, Araki and colleagues suggested that increased Nmnat1 activity in the nucleus led to increased activation of the NAD-sensitive sirtuin-1 (SIRT1). However, it was discovered that if the Wlds protein’s cytoplasmic localization was enhanced by mutating the nuclear localization signal (NLS) in Nmnat1

(Beirowski et al., 2009) or if a mutant Nmnat1 (cytNmnat1), engineered to localize specifically to the cytoplasm and axon (Sasaki et al., 2009b), was expressed, axonal protection was significantly enhanced beyond what is seen in the original Wlds mutant.

This led to the realization that the Wlds protein was acting in the axon to delay Wallerian degeneration.

Two other orthologs of Nmnat1 exist in mammals. While Nmnat1 is localized to the nucleus (Schweiger et al., 2001), Nmnat2 is cytoplasmically localized (Yalowitz et al.,

2004), and Nmnat3 is found in mitochondria (Berger et al., 2005). Gilley and Coleman

(2010) demonstrated that Nmnat2 acts as a cell-body derived axonal survival factor.

Nmnat2 has a very short half-life and must be transported from the cell body down to the axon. Following injury, Nmnat2 levels are rapidly reduced in the distal axon, and this reduction coincides with axonal disintegration. Overexpression of Nmnat2, in vitro, can delay axonal degeneration (Gilley and Coleman, 2010; Milde et al., 2013). Nmnat1 has a much longer half-life and is not easily degraded. Thus, the Nmnat1 containing Wlds protein can substitute for the loss of endogenous Nmnat2 and delay axonal degeneration.

This suggests that the synthesis of NAD+ in the axon is critical for axonal survival.

However, it is not clear that NAD+ is neuroprotective. Even though Nmnat activity is significantly increased in the Wlds mouse (Orsomando et al., 2012), no detectable increase in NAD+ levels is observed (Mack et al., 2001; Araki et al., 2004). In addition, inhibition of the rate-limiting enzyme in NAD+ synthesis, nicotinamide phosphoribosyltransferase, does not have significant effects on the Wlds phenotype

(Conforti et al., 2009; Sasaki et al., 2009a). Yet, the enzymatic activity of Nmnat is necessary for its protective effects (Conforti et al., 2009).

Regeneration of axons in the Wlds mouse has been shown to be delayed and incomplete (Bisby and Chen, 1990; Myers et al., 1996; Sommer and Schafers, 1998; Shin et al., 2014). The increase in NGF in the sciatic nerve after transection is severely blunted in the Wlds mouse, and this has been raised as a possible cause of the regeneration deficiency (Brown et al., 1991b). However, a number of authors have indicated that NGF does not promote regeneration in adult animals (e.g., Diamond et al., 1987; Gloster and

Diamond, 1992, 1995; Shoemaker et al., 2006). More commonly, this regenerative deficiency is attributed to the delayed clearance of myelin and axonal debris (Bisby and

Chen, 1990). The idea that the Wlds mouse would show normal regeneration given a suitable substrate for growth was supported by the finding that facial motor and DRG neurons in Wlds mice show normal increases in growth associated protein-43 (GAP-43) following injury even though regeneration is delayed in vivo (Bisby et al., 1995).

Measurements of a variety of other cytokines and growth factors after axotomy have been shown to differ between WT and the Wlds mice. The decrease in ciliary neurotrophic factor (CNTF) normally seen is slowed in these mutant mice (Subang et al.,

1997). In addition, levels of TNF-α, IL-1α, and IL-1β are lower in Wlds mice (Shamash et al., 2002), as are levels of IL-6 (Reichert et al., 1996) and GM-CSF and IL-10 (Be'eri et al., 1998).

With much still unknown about the mechanism driving Wallerian degeneration, a large body of evidence built on the research conducted on the Wlds mouse suggests the importance of the NAD synthesis pathway in degeneration. The detailed mechanism of how the Wlds mutation conveys its axonal protection remains unknown.

1.4 .Intrinsic Mechanisms Supporting PNS Regeneration

The intrinsic growth capacity of a neuron is dependent on the multitude of changes that occur to the neuronal cell body following injury, termed the “cell body response”

(Grafstein, 1975). This response is centered around the significant increase in the synthesis of mRNAs and proteins (Watson, 1974). These gene expression changes mark a switch in the neuron allowing for the support of axonal regeneration while downregulating its focus on neuronal communication (Harkonen, 1964; Zigmond, 1997;

Chandran et al., 2016).

A significant increase in Ca2+ is seen immediately at the site of injury and is propagated towards the cell body. The Ca2+ wave is tightly regulated through the activation of the Ca2+ -sensitive protease, calpain, which helps to reseal the membrane and cut off the flow of Ca2+ into the axon (Gitler and Spira, 2002).

The Ca2+ wave is responsible for the activation of immediate early genes Jun and Fos

(West and Greenberg, 2011). Transcriptional analysis in response to nerve injury shows that most changes occur at later time points that do not coincide with the timeline of early injury-induced signals (Michaelevski et al., 2010b). However, the Ca2+ wave causes the nuclear export of histone deacetylase 5 (HDAC5) which increases histone acetylation and favors an increase in gene expression (Cho and Cavalli, 2012; Cho et al., 2013; Cho and

Cavalli, 2014).

The immediate early genes activated within hours of injury play a significant role in the regenerative capability of the neuron following injury (Patodia and Raivich, 2012b).

These include Jun, Fos, and activating transcription factor 3 (ATF3). These transcription

10 factors work in concert to form dimers that contribute to the formation of activator protein-1 (AP-1) transcription factor complexes, which can regulate genes associated with regeneration (Jochum et al., 2001). Jun plays a major role in regeneration as the overexpression of this transcription factor in cortical neurons is able to induce neurite outgrowth (Lerch et al., 2014). ATF3 is closely associated with regeneration as well in sensory, motor, and sympathetic neurons (Tsujino et al., 2000; Hyatt Sachs et al., 2007).

Retrograde transport, the slower injury signal, is most closely correlated with the gene expression changes that occur after PNS injury (Michaelevski et al., 2010b).

Administration of colchicine, a drug which blocks axonal transport, significantly delays morphological changes and regeneration associated gene (RAG) expression in injured neurons (Singer et al., 1982; Murphy et al., 1999). Proteomic analysis of these signaling complexes revealed a variety of proteins being trafficked including, JUN amino-terminal kinases (JNKs), signal transducer and activator of transcription 3 (STAT3), dual leucine zipper kinase (DLK), and gp130 cytokines (Michaelevski et al., 2010a).

This process requires de novo protein synthesis, as inhibition of translation prevents growth cone formation (Verma et al., 2005). Interestingly, if protein synthesis inhibitors are administered in vivo directly to the nerve, proximal to the site of injury, significant reductions in axonal regeneration are seen (Gaete et al., 1998). These studies indicate that axonally-synthesized proteins facilitate regeneration (Willis and Twiss, 2006). To further accentuate the importance of this retrograde signaling, axon-specific knockout of importin-β1, a critical scaffolding protein that allows for the association of transcription factors with the dynein motor protein, affects over half of all genes changed as part of the cell body response to injury (Perry et al., 2012).

Following injury to the sciatic nerve activation of STAT3 by phosphorylation of a specific tyrosine residue is seen in the nerve (Lee et al., 2004; Qiu et al., 2005). A correlative relationship between the activation of STAT3 and the ability for DRG neurons to regenerate following injury was demonstrated by showing that only an injury to the peripheral branch and not the dorsal root, activated STAT3 and exhibited regeneration

(Schwaiger et al., 2000). Transcriptome analyses of injured DRG neurons have also identified the STAT family of transcription factors, and more specifically STAT3, as an injury-induced locally translated axonal transcription factor (Michaelevski et al., 2010b;

Smith et al., 2011; Ben-Yaakov et al., 2012; Chandran et al., 2016). STAT3 is retrogradely transported back to the cell body through interaction with importin α and dynein (Ma and Cao, 2006; O'Brien and Nathanson, 2007; Ben-Yaakov et al., 2012).

STAT3 is a common transcription factor used by multiple pathways such as the gp130 cytokines (Zigmond, 2012). In addition to its role as a retrograde injury-induced signal,

STAT3 could be pivotal in integrating cytokine and neurotrophin signals to help promote regeneration (Corness et al., 1998; Rajan et al., 1998; Shadiack et al., 1998; Ng et al.,

2003). By examining sympathetic reinnervation of the heart following myocardial infarction, it was shown that NGF stimulates serine phosphorylation of STAT3, in contrast to the tyrosine phosphorylation induced by cytokines (Pellegrino and Habecker,

2013). Additionally, transfection studies of STAT3-null neurons suggested that two pools of STAT3, phosphorylated on either serine or tyrosine, are necessary for a maximal regenerative response (Selvaraj et al., 2012; Pellegrino and Habecker, 2013). However, an in vivo investigation into the importance of STAT3 in DRG regeneration revealed a phase specific role for STAT3 in the initiation, but not elongation of axonal regeneration

(Bareyre et al., 2011). This suggests that while STAT3 can support regeneration, it may not be necessary for it.

The gene expression changes occurring in peripheral neurons represents a shift in phenotype to support regeneration and not neurotransmission (Harkonen, 1964; Zigmond,

1997). This is supported by work in the sympathetic nervous system which showed increases in tubulin expression accompanied by a decrease in tyrosine hydroxylase expression (Cheah and Geffen, 1973; Koo et al., 1988; Sun and Zigmond, 1996a). Later studies showed the downregulation of a large number of synaptic transmission related molecules (Zhou et al., 1998, 2001; Boeshore et al., 2004). Large scale microarray and

RNA sequencing studies have provided a more complete analysis of the overall gene expression changes following peripheral nerve injury in the DRG, superior cervical ganglion (SCG), and facial motor nucleus (Costigan et al., 2002; Boeshore et al., 2004;

Zujovic et al., 2005; Kim et al., 2009; Michaelevski et al., 2010b; Ben-Yaakov et al.,

2012; Chandran et al., 2016).

A large portion of the identified RAGs encode proteins in one of several categories: cytoskeletal proteins and adaptors, metabolic enzymes, neuropeptides, cytokines and chemokines, neurotrophins, and transcription factors. Classic neurotransmitter systems are downregulated after axotomy (Grafstein and McQuarrie, 1978; Zigmond et al., 1996;

Patodia and Raivich, 2012a). Adhesion molecules and enzymes upregulated following nerve injury include α7β1 integrin, a cell surface receptor for laminins (Yao et al., 1996;

Ekstrom et al., 2003), and arginase 1, a key enzyme in polyamine synthesis (Boeshore et al., 2004; Schreiber et al., 2004; Deng et al., 2009). Peripheral injury-induced neuropeptides include vasoactive intestinal peptide (VIP) (Villar et al., 1989; Hyatt-Sachs

13 et al., 1993), galanin (Hokfelt et al., 1987; Mohney et al., 1994; Schreiber et al., 1994), and pituitary adenylate cyclase activating polypeptide (PACAP) (Moller et al., 1997;

Armstrong et al., 2008; Habecker et al., 2009). Multiple transcription factors are involved in injury induced signaling, and many show significantly increased expression following injury, including ATF3 (Tsujino et al., 2000; Hyatt Sachs et al., 2007) and the immediate early gene, JUN (Jenkins et al., 1993). Recently a systems-level analysis was completed to identify the transcriptional program that is observed after PNS but not CNS injury

(Chandran et al., 2016). The authors identified a core group of transcription factors that link the various regeneration-associated signaling pathways. The core transcription factors included, cJun, STAT3, Sox11, cMyc, Fos, Smad1, and others (Chandran et al.,

2016)

Immune molecules play a major role in the cell body response to injury, where cytokines such as leukemia inhibitory factor (LIF; Rao et al., 1993; Banner and Patterson,

1994; Sun et al., 1994; Gardiner et al., 2002), IL-6 (Habecker et al., 2009) and CCL2

(Schreiber et al., 2001; Subang and Richardson, 2001; Tanaka et al., 2004) are upregulated in response to axotomy. Gene expression for certain neurotrophic factors and their receptors are increased after nerve axotomy, as well.

The most recognized RAG is growth-associated protein 43(GAP-43). GAP-43 was originally identified as a rapidly transported axonal protein that is highly induced after sciatic nerve injury, hence its name (Skene and Willard, 1981). Its upregulation following nerve injury is correlated with significant regeneration (Skene and Willard, 1981; Katz and Black, 1986; Skene, 1989). Yet, only when GAP-43 is overexpressed with

14 cytoskeleton-associated protein-23 (CAP-23) does it allow for CNS axon regeneration, while it has minimal effects on its own (Bomze et al., 2001).

Out of the many genes whose expression is changed following peripheral nerve injury and are thus considered RAGs, only a handful have actually been directly implicated as being necessary or sufficient for regeneration.

Arginase 1 has been tied to axonal regeneration in the PNS (Gilad et al., 1996; Lee and Wolfe, 2000). Overexpression of arginase 1 in cerebellar neurons was sufficient to overcome the inhibition of MAG and myelin and allow for axonal elongation in vitro, similar to administration of cAMP. Additionally, blocking polyamine synthesis prior to administration of cAMP to cultures inhibited their ability to extend axons on inhibitory substrates (Cai et al., 2002).

ATF3 has become a reliable neuronal marker of injury (Tsujino et al., 2000; Hyatt

Sachs et al., 2007). Overexpression of ATF3 in uninjured DRG sensory neurons produced a significant increase in neurite outgrowth similar to a conditioning lesion

(Seijffers et al., 2006; Seijffers et al., 2007). This did not have an effect on central regeneration or growth on non-permissive substrates, such as MAG, suggesting its importance specifically in peripheral regeneration (Seijffers et al., 2007).

LIF, a member of the IL-6 cytokine family, has been linked to regeneration in the

SCG through activation of its signaling receptor gp130 (Habecker et al., 2009; Hyatt

Sachs et al., 2010). In the DRG, a LIF knockout mouse shows a significant decrease in

GAP-43 positive fibers, marking regenerating fibers, distal to the injury site 3 d, after a sciatic nerve crush, as well as, a 50% reduction in growth in vitro following a CL

(Cafferty et al., 2001). Small proline-rich repeat protein 1A (SPRR1A) shows a 60-fold increase in expression following peripheral injury (Bonilla et al., 2002). Overexpression of SPRR1A in sensory neurons induces significant increases in neurite outgrowth in vitro, while knockdown of this gene inhibits regeneration (Bonilla et al., 2002).

1.5 Macrophage Response to Injury

While the inflammatory response to injury in the PNS is complex and involves a number of immune cells, it is largely dominated by macrophages (Perry et al., 1987).

Macrophages respond to a peripheral axonal injury in two distinct compartments: the nerve distal to the injury site and the ganglion. Tissue or resident macrophages are present in the sciatic nerve, SCG, and DRG (Schreiber et al., 1995; Mueller et al., 2001;

Hu and McLachlan, 2003). While many details about the complete contribution of resident macrophages to the overall immune response to injury are unknown, their response has been measured in various tissues. Following a sciatic nerve crush, resident macrophages were shown to alter their morphology, suggesting inflammatory activation, and proliferate, with a maximal response observed 4 days after injury (Mueller et al.,

2001). Using a chimera created by transplanting bone marrow from a GFP-transgenic mouse, it was determined that resident macrophages increased 9.5-fold in the first 7 days after injury in the sciatic nerve distal to the crush site (Mueller et al., 2003). In two peripheral ganglia, the SCG and DRG, less quantitative measures have revealed little to no change in the number of resident macrophages after nerve injury (Lu and Richardson,

1993; Schreiber et al., 1995).

The largest contribution to macrophage accumulation within injured PNS tissues come from monocyte-derived hematogeneous macrophages. However, it should be noted

16 that the overlap of identifying markers between resident and hematogeneous macrophages make it difficult to isolate the populations from one another. In the sciatic nerve, hematogeneous macrophages outnumber the resident population 3:1 7 days after injury (Mueller et al., 2003). Infiltration of the sciatic nerve begins 48 hours after injury with peak accumulation occurring between 7 and 14 days after injury (Perry et al., 1987;

Nadeau et al., 2011). Macrophage numbers remain elevated in the nerve for at least 30 days after injury (Perry and Brown, 1992).

In peripheral ganglia, monocyte infiltration and macrophage accumulation follow a similar timeline to that seen in the sciatic nerve. In the SCG, hematogeneous macrophages are present at 48 hours after injury and remain for at least 2 weeks following injury to the ECN and ICN (Schreiber et al., 1995; Schreiber et al., 2002).

Lumbar DRGs show accumulation beginning 48 to 72 hours after sciatic nerve injury with an increased number of monocyte-derived macrophages remaining for at least 28 days (Lu and Richardson, 1993; Kwon et al., 2013). In both ganglia the peak accumulation is at 7 days post injury (Lu and Richardson, 1993; Schreiber et al., 1995).

The signal mediating entry into PNS tissue following injury is thought to be the chemokine (C-C motif) ligand 2 (CCL2), as a global knockout of the chemokine receptor

CCR2 yields a significant decrease in macrophage accumulation in the sciatic nerve following injury (Siebert et al., 2000). CCL2, also known as monocyte chemoattractant protein 1 (MCP-1), is a chemoattractant protein which recruits monocytes to sites of inflammation, infection, and injury. CCL2 also has the ability to act as a chemokine for neutrophils and dendritic cells (Colonna et al., 2004; Reichel et al., 2009). Although

CCL2 can bind to several chemokine (C-C motif) receptors, including CCR1, CCR2, and

CCR4 (Savarin-Vuaillat and Ransohoff, 2007; White et al., 2007), CCR2 is the primary receptor (Jung et al., 2009). CCR2 specifically binds CCL2 with greater than 10 times higher affinity than CCL7 and doesn't bind CCL3 or chemokine (C-X-C motif) ligand

1(CX3CL1; Kurihara et al., 1997). Jung et al. (2009) used a transgenic mouse that labelled CCL2 and CCR2 with fluorescent tags and showed that CCL2 binds with CCR2 leading to endocytosis of the CCL2/CCR2 bound receptor-ligand complex. Using transfected cells, application of a specific CCR2 antagonist could completely block the appearance of CCL2/CCR2 localization to vesicles.

Injury induced chemokine expression is thought to be responsible for monocyte- derived macrophage accumulation in both the nerve and the ganglia. Following axonal injury, Schwann cells upregulate Jun and undergo a process known as dedifferentiation, putting the cells into a non-myelinating and immature state (Jessen and Mirsky, 2008).

Dedifferentiated Schwann cells begin to express various pro-inflammatory cytokines and chemokines, within 6 hours of injury, including tumor necrosis factor alpha (TNFα), IL-

1, and CCL2 (Subang and Richardson, 2001; Shamash et al., 2002). Napoli et al. (2012) showed that expression and activation of an inducible Raf kinase in myelinating Schwann cells was sufficient to cause dedifferentiation, CCL2 expression, and macrophage accumulation in the sciatic nerve in the absence of injury. However, studies in zebrafish have shown that effective recruitment of macrophages can be achieved in the absence of

Schwann cells (Rosenberg et al., 2012).

CCL2 expression in primary sensory neurons, located in DRGs, has been significantly studied following nerve injury, as the DRG is a popular model system. An early finding showed that CCL2 was up-regulated rapidly in DRG neurons after sciatic

18 nerve lesion (Tanaka et al., 2004). Zhang and De Koninck (2006) then showed that after nerve constriction, a common model of neuropathic pain, CCL2 was induced in both small- and large- diameter neurons that also expressed the transcription factor ATF-3, a marker for axonal injury. This suggested that CCL2 was increased mainly in injured neurons. However, others have showed that CCL2 is produced by both damaged

(Lumbar level 5; L5) and undamaged (L4) primary sensory neurons after ligation and transaction of the L5 spinal nerve (Thacker et al., 2009). CCL2 is also induced in DRG neurons in other injury models, including sciatic nerve demyelination (Jung et al., 2009), chronic compression of the DRG (White et al., 2005b), and chronic constriction of the sciatic nerve (Jeon et al., 2009), as well as in an inflammatory pain brought on by adjuvant injection into the hind paw (Jeon et al., 2008). Neuronal expression of CCL2 is not restricted to sensory neurons, as sympathetic neurons in the SCG also upregulate the chemokine after nerve injury (Schreiber et al., 2001). Further evidence suggests that

CCL2 may also be expressed by satellite glial cells after injury (Jeon et al., 2009). The localization of CCL2 solely to injured DRG neurons may be contested, but its upregulation following injury is universally accepted.

CCR2 is also upregulated after peripheral nerve injury in the DRG (Jung and Miller,

2008). Staining shows that CCR2 is partially colocalized with CCL2, suggesting a possible autocrine/paracrine role for CCL2/CCR2 signaling within the DRG (Jung et al.,

2009). In situ hybridization shows that compression injury to the sciatic nerve the DRG induces CCR2 mRNA expression in neurons and non-neuronal cells in both compressed

(L4 and L5) and non-compressed (L3) DRGs next to the injury (White et al., 2005a).

CCR2 is also up-regulated in the DRG after sciatic nerve demyelination (White et al.,

2009).

1.6 Macrophage Polarization

As an essential component of the innate immune response to infection or injury, macrophages are capable of taking on a variety of activation states with a range of function (Gordon and Taylor, 2005; Martinez et al., 2008; Mosser and Edwards, 2008). In response to various cues (e.g., microbial products, damaged cells, activated lymphocytes, cytokines) or under different pathophysiologic conditions, macrophages can acquire distinct functional states by undergoing different phenotypic polarizations (Cassetta et al.,

2011). These differential activation states have most commonly been described using the nomenclature M1 and M2 to describe a purely pro-inflammatory or anti-inflammatory macrophage activation state, respectively (Gordon and Taylor, 2005). Recently, this nomenclature has fallen out of favor and is now replaced with a more descriptive terminology, for the purposes of this paper, the terms M1 and M2 will be used to describe macrophages that are primarily pro- or anti-inflammatory, respectively (Martinez and

Gordon, 2014). The M1/pro-inflammatory phenotype is stimulated by microbial products or pro-inflammatory cytokines, including but not limited to IFN-γ, TNF, or Toll-like receptor (TLR) ligands (Benoit et al., 2008; Lawrence and Natoli, 2011). The typical characteristics of M1 macrophages include high antigen presentation, high production of

IL-12 and IL-23, and increased production of nitric oxide (NO) and reactive oxygen species (Mantovani et al., 2004). In contrast, M2-type macrophages are anti- inflammatory in nature and are observed in healing-type circumstances without infections and express high levels of IL-10, and IL-1R antagonist (Gordon and Martinez, 2010). M2

20 macrophages are characterized by the upregulation of Dectin-1, mannose receptor/CD206, various scavenger receptors, CD163, and CCR2 (Gordon, 2003; Gordon and Martinez, 2010). Instead of generating nitric oxides, M2 macrophages produce ornithine and polyamines through the arginase pathway (Edwards et al., 2006; Pesce et al., 2009). Nitric oxide production through inducible nitric oxide synthase (iNOS) by M1 macrophages and the polyamine synthesis through arginase-1 by M2 macrophages have been regarded by some investigators as the most characteristic molecules distinguishing the phenotypes (Martinez and Gordon, 2014). Another distinguishing feature is the polarizing factors that induce a pro-inflammatory versus an anti-inflammatory activation state in macrophages. Work with bone marrow-derived and peritoneal macrophages in vitro has shown that interferon-gamma (IFNγ) stimulates a M1 macrophage polarization, whereas IL-4 and IL-13 stimulate a M2 activation state, a model that is frequently utilized to assess differences between these two extreme macrophage polarizations (Sica and Mantovani, 2012).

Inflammatory M1 macrophages produce many other pro-inflammatory cytokines like

TNFα, IL-1, IL-6, and IL-12 (Mosser, 2003; Mosser and Edwards, 2008), while M2 macrophages generate anti-inflammatory cytokines such as IL-10 and IL-1 receptor antagonist (Verreck et al., 2004; Park-Min et al., 2005). Additional signatures of M2 phenotype, such as YM1 (a member of the chitinase family) and FIZZ1 (resistin-like molecule 1 alpha) are also identified (Raes et al., 2002). M1 macrophages promote a Th1 response and possess strong activity in response to bacteria and tumors, while M2 macrophages are involved in parasitic containment and promotion of the Th2 response,

21 tissue remodeling, and immune tolerance (Mantovani et al., 2002; Martinez et al., 2008;

Mantovani et al., 2013).

A synchronized action of various inflammatory modulators, signaling molecules, and transcription factors are involved in regulating macrophage polarization. At the cellular level, although M1 and M2 macrophage activities exist without T or B cell influence

(Sica and Mantovani, 2012), specialized or polarized T cells (Th1, Th2, TRegs) do play a role in macrophage polarized activation (Adams and Hamilton, 1984; Paulnock, 1992).

Canonical STAT signaling is a central pathway in modulating macrophage polarization.

Activation of STAT signaling pathways by IFNs and TLR signaling will skew macrophage function toward the M1 phenotype (via STAT1), while activation of

IRF/STAT (via STAT6) signaling pathways by IL-4 and IL-13 will skew macrophage function toward the M2 phenotype (Kovarik et al., 1998; Szanto et al., 2010). Signals initiated by glucocorticoids, apoptotic cell-released molecules, and immune complexes can also significantly affect macrophage functional status (Mantovani et al., 2004;

Mosser and Edwards, 2008). In addition, macrophage polarization is modulated by local microenvironmental conditions such as hypoxia (Blouin et al., 2004). More importantly,

M1–M2 polarization of macrophages is a highly dynamic process highlighted by the fact that polarization can be reversed under physiological and pathological conditions

(Porcheray et al., 2005; Lumeng et al., 2007). In the course of various pathophysiological settings, the same signaling pathway can be involved in either M1 or M2 polarization of macrophages. The molecular mechanisms that govern the phenotype switch of macrophages, however, remains incompletely understood. Moreover, imbalances of macrophage M1–M2 polarization are associated with various diseases. Disease

22 conditions are frequently associated with polarization of macrophage activation, with classically activated M1 macrophages implicated in initiating and sustaining inflammation and M2 macrophages associated with resolution of chronic inflammation

(Mosser and Edwards, 2008).

1.7 Role of Macrophages in Axonal Regeneration

The macrophage response to peripheral nerve injury has been shown to play a functional role in axonal regeneration. The macrophage is best classified as a primary phagocyte, whose purpose is to engulf and digest debris, dying or dead cells, and foreign pathogens (Aderem and Underhill, 1999). It is this role as a professional phagocyte that is cited as the primary role in which macrophages support the process of axonal regeneration in the PNS through their support of the degenerative process. Early work into the peripheral injury response, by Augustus Waller described changes observed in cranial nerves after injury (He Waller, 1850; Stoll et al., 2002). The changes described have been termed Wallerian degeneration and include the rapid disintegration of the distal nerve after injury and the subsequent influx of immune cells that aid in ridding the area of debris resulting from this breakdown. However, in the CNS, Wallerian degeneration occurs much more slowly and myelin, which can inhibit regeneration, can persist for years (for review see Vargas and Barres, 2007).

The macrophage plays a significant role in this degenerative process. Both resident and inflammatory monocyte-derived macrophages are involved in the clearance of axonal and myelin debris in peripheral nerves, distal to the site of injury (Griffin et al., 1992;

Bruck, 1997; Hirata et al., 1999; Stoll and Muller, 1999; Hirata and Kawabuchi, 2002).

The debris resulting from the degeneration of the axons, no longer connected to their cell

23 bodies, and myelin is inhibitory to axons regenerating from the nerve proximal to the injury site (Caroni and Schwab, 1988; McKerracher et al., 1994; Mukhopadhyay et al.,

1994). Schwann cells, which dedifferentiate into an immature state following injury, phagocytose myelin and axonal debris prior to hematogeneous macrophage accumulation in the nerve (Liu, 1974; Stoll et al., 1989; Reichert et al., 1994). However, the macrophage plays the primary role in this clearance process, as blockade of entry into the nerve significantly impairs Wallerian degeneration and debris clearance (Vougioukas et al., 1998; Liu et al., 2000; Siebert et al., 2000; Gray et al., 2007; Barrette et al., 2008).

Inhibition of macrophage accumulation in the sciatic nerve after injury, and the subsequent impairment in Wallerian degeneration has also been shown to significantly decrease axonal regeneration. Barrette et al. (2008) demonstrated that continuous delivery of ganciclovir to a mouse expressing CD11b driven thymidine kinase in myeloid cells significantly decreased the number of granulocytes, inflammatory monocytes, and macrophages present in the sciatic nerve after injury. As a result myelin clearance, measured by luxol fast blue staining, and functional regeneration were significantly delayed. Another study, using the Wlds mouse, showed that Wallerian degeneration and macrophage accumulation were significantly delayed in the sciatic nerve after transection, as was subsequent regeneration (Lunn et al., 1989). Both studies concluded that Wallerian degeneration and clearance of myelin and axonal debris by hematogeneous macrophages is a prerequisite for peripheral axon regeneration (Lunn et al., 1989;

Barrette et al., 2008).

Macrophages also perform other functions within the nerve that support the growth of axons after injury, outside of their role as a phagocyte. Nerve growth factor (NGF) is a

24 critical axon guidance molecule and target-derived survival factor for sensory and sympathetic neurons during development (Campenot, 1977; Otten et al., 1980; Crowley et al., 1994; Patel et al., 2000; Harrington and Ginty, 2013). After injury in the peripheral nervous system, the loss of target derived NGF serves as an injury signal to the neuronal cell body (Zigmond, 1997; Shadiack et al., 1998; Shadiack et al., 2001; Shoemaker et al.,

2006). NGF mRNA shows a biphasic increase in response to nerve injury, where distal to the injury site NGF message increases at 6 hours and again at 3 days after injury where it remains elevated for at least 21 days (Heumann et al., 1984; Heumann et al., 1987a;

Taniuchi et al., 1988). This second and long-lasting increase in NGF is thought to be dependent upon macrophage entry into the distal nerve compartment, as blockade of hematogeneous macrophage accumulation or ex vivo nerve culture (prior to monocyte extravasation into the nerve) both show significant attenuation of NGF expression

(Heumann et al., 1987a; Heumann et al., 1987b; Brown et al., 1991b). While NGF is not necessary for the survival of sensory neurons after injury, numerous studies have shown that both NGF and brain-derived neurotrophic factor can stimulate significant axonal outgrowth and regeneration (Lindsay, 1988; Oudega and Hagg, 1996; Cai et al., 1999).

Recent work has highlighted another role for macrophages in supporting axonal regeneration after a nerve transection. Following nerve transection, the bridge of new tissue that forms between the nerve stumps is composed of dense extracellular matrix and inflammatory cells, with monocyte-derived macrophages the major cell type within the bridge (Cattin et al., 2015). As the bridge is initially without blood vessels, it becomes hypoxic and this is sensed specifically by the macrophages, which secrete vascular endothelial growth factor (VEGF), which is necessary and sufficient to induce the

25 vascularization of the bridge. The new blood vessels along with other unidentified macrophage-derived factors facilitate Schwann cell migration into the tissue (Cattin et al.,

2015). The Schwann cells use the vasculature as tracks to direct Schwann cell migration into the bridge, and the Schwann cells are then used as tracks to guide the regenerating axons into the distal nerve.

While the function of macrophages in peripheral ganglia following nerve injury remains elusive, indirect evidence in the DRG and work in the optic nerve suggest a role in axonal regeneration. Since there is very little cell death and degeneration occurring in peripheral ganglia after injury the phagocytic function of macrophages is likely not be utilized in the cell body compartment after axonal injury suggesting an alternate function

(Tessler et al., 1985; Himes and Tessler, 1989). Lu and Richardson (1991) showed that inflammation around DRG neuronal cell bodies increased regeneration of dorsal root axons following a crush injury. Injection of C. parvum into the L5 DRG, to create a local inflammatory response, significantly increased dorsal root regeneration. This inflammatory response, which resulted in significant macrophage accumulation, also significantly increased the expression of the RAGs GAP-43, c-Jun, and calcitonin gene- related peptide (CGRP; Lu and Richardson, 1995). To highlight the involvement of macrophages in this effect, peritoneal macrophages were isolated and directly injected into the DRG. Injection of macrophages alone was able to significantly increase dorsal root axon regeneration, suggesting that the normal macrophage accumulation in response to injury may be involved in the ability of peripheral neurons to regenerate (Lu and

Richardson, 1991, 1993). Inflammation around DRG neuronal cell bodies has also been shown to help facilitate regeneration of sensory axons past the dorsal root entry zone and

26 into the spinal cord following dorsal root injury. Injection of zymosan, a fungal ligand for toll-like receptor 2, was injected into the L5 DRG and induced the extravasation of monocytes into the DRG. Zymosan injection into the DRG, in combination with

Chondroitinase ABC treatment of the dorsal root entry zone, allowed for significant axonal regeneration (Steinmetz et al., 2005).

Macrophages can be polarized into different functional phenotypes in response to different environmental cues (Gordon, 2003; Gordon and Martinez, 2010; Murray and

Wynn, 2011; Murray et al., 2014); Wang et al. (2014). It has been proposed that M2- polarized macrophages make essential contributions to neural repair processes, such as neurogenesis, angiogenesis, remyelination, and axon regeneration (Kigerl et al., 2009; Hu et al., 2015; Kwon et al., 2015). Kigerl et al. (2009) showed that conditioned medium from M2 polarized bone marrow-derived macrophages, stimulated with IL-4, induced significant neurite outgrowth from sensory neurons on an inhibitory substrate.

M1polarized macrophage conditioned medium, stimulated with interferon-gamma

(IFNγ), did not induce increased neurite outgrowth (Kigerl et al., 2009; Gensel et al.,

2012). Another recent study also showed that CCL2 and sialic acid-binding Ig-like lectin-9 treatment promotes the M2-like activation of macrophages which release pro- regenerative factors that can result in significant neurite outgrowth of cerebellar neurons on an inhibitor substrate (Matsubara et al., 2015). Thus, an anti-inflammatory, or M2, phenotype may be a critical factor in the regeneration-promoting effects of macrophages.

Following injury to the optic nerve, like all CNS lesions, retinal ganglion cells

(RGC) are unable to regenerate their axons (Liu et al., 2011; Cregg et al., 2014).

However, stimulation of an inflammatory response around the retinal ganglion cells,

27 through lens injury or zymosan injection into the eye results in increased regeneration of axons in the optic nerve following a crush injury (Leon et al., 2000; Benowitz and

Popovich, 2011). It was later shown that this inflammation-induced regeneration was dependent upon macrophages and their expression of oncomodulin (Yin et al., 2003; Yin et al., 2006; Yin et al., 2009). The topic of macrophage involvement in optic nerve injury is a contested topic. Recent evidence from the same laboratory found that neutrophils may be the primary source of oncomodulin in the retina (Kurimoto et al., 2013).

1.8 Focus of Thesis

While exogenous macrophage addition and stimulation of inflammation around neuronal cell bodies have been implicated in peripheral nerve regeneration, no one has looked at the role of endogenous macrophage accumulation in ganglia as it pertains to regeneration. In this thesis, we present data that suggests that blockade of the endogenous injury-induced accumulation of macrophages in peripheral ganglia results in the complete ablation of the conditioning lesion response. We show that CCL2/CCR2 signaling is necessary for the accumulation of macrophages in ganglia after injury. Furthermore, utilizing viral overexpression of CCL2 in uninjured DRG neurons, we show that macrophage accumulation in the DRG is sufficient to increase the regenerative capacity of sensory neurons. Finally, we describe an initial mechanism that mediates macrophage stimulation of axonal regeneration through the LIF/STAT3 pathway.

Chapter 2: A Critical Role for Macrophages Near Axotomized Neuronal Cell Bodies

in Stimulating Nerve Regeneration

Jon P. Niemi1, Alicia DeFrancesco-Lisowitz1, Lilinete Roldán-Hernández1,2, Jane

A. Lindborg1, Daniel Mandell1, and Richard E. Zigmond1*

1Department of Neurosciences and 2Department of Graduate Studies, Case Western

Reserve University, Cleveland, Ohio 44106-4975, U.S.A.

Reprinted with permission from Journal of Neuroscience, Copyright, 2013.

2.1 Acknowledgements

This research was supported by grants DK097223, NS017512, and P30EY11373 from the National Institutes of Health. J.P.N and J.A.N. were supported by training grant

NS067431, and L.R.-H was supported by GM075207. The authors declare no competing financial interests. We thank Heather Butler and Kathryn Franke for maintaining the Wlds and CCR2 -/- breeding colonies, Anna Yakubenko for genotyping the mutant animals,

Maryanne Pendergast for advice on imaging, and Jared Cregg for helpful comments on the manuscript. We also thank Dr. Jeff Milbrandt, Washington University, and Dr.

Timothy Kern, Case Western Reserve University, for providing our original breeding pairs of the Wlds and the CCR2 -/- mice, respectively.

2.2 Abstract

Macrophages have been implicated in peripheral nerve regeneration for some time, supposedly through their involvement in Wallerian degeneration, the process by which the distal nerve degenerates after axotomy and is cleared by phagocytosis. Thus, in several studies in which macrophage accumulation in the distal nerve was reduced and

Wallerian degeneration inhibited, regeneration was delayed. This interpretation, however, ignores the more recent findings that macrophages also accumulate around axotomized cell bodies. The function of macrophage action at this second site has not been clear. In two mutant strains of mice, the Wlds mouse and the chemokine receptor CCR2 knockout mouse, we report that macrophage accumulation after axotomy was abolished in both the

DRG and the distal sciatic nerve. To measure neurite outgrowth, DRG neurons were given a conditioning lesion, and outgrowth was measured in vitro 7 d later in the absence of the distal nerve segment. The increased growth normally seen after a conditioning lesion did not occur in either Wlds or CCR2 -/- mice. In the SCG, particularly in Wlds mice, macrophage accumulation was reduced but not abolished after axotomy. In SCG neurons from Wlds mice, the conditioning lesion response was unchanged; however, in

CCR2 -/- mice where the effect on macrophage accumulation was greater, SCG neurite outgrowth was significantly reduced. These results indicate that macrophages affect neurite outgrowth by acting at the level of peripheral ganglia in addition to any effects they might produce by facilitation of Wallerian degeneration.

2.3 Introduction

In the peripheral nervous system unlike in the central nervous system, the axon segment distal to a lesion degenerates rapidly (Wallerian degeneration), and the proximal segment regenerates. Both degeneration and regeneration involve the interaction of axons with non-neuronal cells, including macrophages and Schwann cells. Wallerian degeneration begins rapidly after axonal damage and was thought to result from general blockade of axonal transport (Joseph, 1973); however, this simple view changed with the discovery of the slow Wallerian degeneration (Wlds) mouse in which degeneration takes weeks to begin (Lunn et al., 1989).

Early studies on the role of macrophages in Wallerian degeneration, reviewed by

Perry (1994), include observations by Ramon Y Cajal that blood borne cells play a role in clearing myelin. While peripheral nerves, like other tissues, contain “resident” macrophages (Oldfors, 1980), primarily “infiltrating” monocytes have been implicated in

Wallerian degeneration (e.g., Beuche and Friede, 1984; Lunn et al., 1989), and the slow

Wallerian degeneration in the Wlds mouse is accompanied by delayed monocyte influx

(Lunn et al., 1989; Hall, 1993).

The idea that macrophages also play a role in nerve regeneration and do so by their effects on Wallerian degeneration has been based on two arguments. First, blockade of macrophage accumulation in the distal nerve and the resulting slow Wallerian degeneration have been shown to be associated in several studies with slow nerve regeneration (Brown et al., 1991b; Dailey et al., 1998; Barrette et al., 2010). Secondly, axons do not grow into intact nerves (Langley and Anderson, 1904; Brown et al., 1991a;

Bedi et al., 1992; Agius and Cochard, 1998; Luk et al., 2003). Proposed rationales for these effects include physical obstruction of axonal elongation into the non-degenerating nerve segment, lack of clearance of growth inhibitory factors, and decreased secretion of neurotrophic factors by the distal nerve (Bisby and Chen, 1990; Perry and Brown, 1992;

Chen and Bisby, 1993), all three of which could result from decreased monocyte influx

(Lunn et al., 1989; Brown et al., 1991b; Barrette et al., 2008).

These arguments, however, do not take into account the fact that a second site of macrophage accumulation exists after axonal injury. Our laboratory and Lu and

Richardson demonstrated in the 1990s that infiltration also occurs around axotomized neuronal cell bodies (Lu and Richardson, 1993; Schreiber et al., 1995), an effect preceded by expression of the macrophage chemokine CCL2 by these neurons (Schreiber et al.,

2001; Tanaka et al., 2004). We, therefore, examined this second site of macrophage accumulation in two mutant mouse strains in which accumulation in the distal nerve is inhibited. Finding a clear inhibition also in peripheral ganglia, we tested the hypothesis, which is generally assumed to be true, that neurite outgrowth from sensory and sympathetic neurons from these mutant animals would be normal if they were cultured on a permissive substrate. Our findings contradict this hypothesis and instead strongly support the novel proposal that macrophages act within peripheral ganglia to promote nerve regeneration independent of any effects on Wallerian degeneration.

2.4 Materials and Methods

Animal surgeries. Eight to twelve week old male wild type (WT) mice

[C57BL/6NHsd (Harlan Laboratories, Frederick MD) or C57BL/6J (Jackson

Laboratories, Bar Harbor ME)] and mutant mice [C57BL/6OlaHsd-Wlds (Wlds; Harlan

Laboratories)] or C57B6.129S4-Ccr2tm1Ifc/J (CCR2 -/-; Jackson Laboratories)] were utilized for this study. The animals were housed under a 12 h:12 h light:dark cycle with ad libitum access to food and water. The SCG and the fifth lumbar DRG (L5 DRG) were axotomized unilaterally under isoflurane anesthesia by transecting the postganglionic axons of the SCG (the internal and external carotid nerves) near their exits from the ganglion and the sciatic nerve at mid-thigh level. For the sciatic nerve, a 2 mm piece of the distal nerve segment was then removed. The contralateral sciatic nerve and ganglia served as internal controls. Six, twenty-four or forty-eight hours or 7 d later, the animals were sacrificed by CO2 inhalation, and the SCG, L5 DRG, sciatic nerves, and submandibular glands, were removed for immunohistochemical or molecular biological analysis. All surgical procedures were approved by the Case Western Reserve

University’s Institutional Animal Care and Use Committee.

Immunohistochemistry (IHC). Axotomized and contralateral SCG, L5 DRG, and sciatic nerves from Wlds, CCR2 -/-, and WT mice were removed 7 d after unilateral axotomy, and the ganglia were desheathed and fixed by immersion in 4% paraformaldehyde. The tissues were cryoprotected in 30% sucrose and embedded in

Tissue-Tek O.C.T. compound (Electron Microscopy Sciences; Hatfield, PA). IHC was performed on 10 µm cryostat sections. For quantification of macrophages, a rat monoclonal antibody to CD11b (also known as Mac1, CR3, and integrin αM; 1:100;

Millipore; Billerica, MA) was incubated with tissue sections overnight at 4oC. While

CD11b can stain both macrophages and neutrophils, few neutrophils are found in the peripheral nervous system 7 d after nerve injury (Perry, 1994; Lindborg, DeFrancesco-

Lisowitz, and Zigmond, unpublished observations; Nadeau et al., 2011). After washing, the sections were incubated in DyLight 549 secondary antibody (1:400; Jackson

ImmunoResearch Laboratories, Inc., West Grove, PA) for 1 h. To assess Wallerian degeneration in the sympathetic system, submandibular glands were dissected 7 d after

SCG axotomy and prepared for IHC as described above. Submandibular gland sections were incubated with a rabbit polyclonal antibody to tyrosine hydroxylase (TH, 1:500;

Pel-Freez; Rogers, AR) for 2 h at room temperature. After washing, the sections were incubated with a Cy3 secondary antibody (1:400; Jackson ImmunoResearch Laboratories,

Inc.) for 45 min. In all experiments, sections not exposed to the primary antiserum were included for each experimental group. Images were captured at 25x magnification using

SimplePCI software (Hamamatsu Corporation; Bridgewater, NJ) and then quantified using MetaMorph software (Version 7.6.3.0, Molecular Devices; Downingtown, PA).

The area of the section that was stained is expressed as a percent of the total area examined. The CD11b data were further analyzed, by expressing each ipsilateral ganglion as a percent increase over the contralateral ganglion in the same animal.

Real Time PCR (RT-PCR). The expression of CCL2 mRNA was analyzed by RT-

PCR. The procedures followed are described in Habecker et al. (2009). Six, twenty-four and forty-eight hours after unilateral axotomy, ipsilateral and contralateral ganglia from

Wlds and WT mice were removed and stored in RNAlater (Life Technologies; Grand

Island, NY) at 4oC. RNA was extracted from pairs of ipsilateral and contralateral ganglia using the Ambion RNAqueous micro kit. Three to six samples were included for each time point. Total RNA was quantified, and 400 ng were reverse transcribed using a High

Capacity cDNA Reverse Transcription Kit (Applied Biosystems; Carlsbad, CA). RT-

PCR was performed in an ABI Step-One Plus, using prevalidated TaqMan expression assays (CCL2, Mm00441242_m1; GAPDH, Mm99999915_g1; Applied Biosystems), and samples were assayed in triplicate. CCL2 mRNA values were normalized for each sample to the mRNA glyceraldehyde-3-phosphate dehydrogenase (GAPDH), which served as an internal control.

Ganglion explants. To assess the outgrowth of peripheral neurons in response to injury, we evaluated neurite outgrowth in explanted ganglia after a conditioning lesion

(Shoemaker et al., 2005). Seven days after unilateral axotomy, the ipsilateral and contralateral SCGs and L5 DRGs from Wlds, CCR2 -/- and WT mice were removed, desheathed, placed on coverslips, and overlaid with 7.5 µl Matrigel (Becton Dickinson,

Franklin Lakes, NJ). Culture plates were placed in a 37oC incubator for 5 min to allow gelling of the Matrigel before adding 1 ml F12 medium with the additives described in

Hyatt Sachs et al. (2010). Phase-contrast images of neurite outgrowth from each SCG and each DRG were captured at 24 and 48 h after explantation using an Axiovert 405 M microscope at 10x magnification. Neurite outgrowth was assessed using MetaMorph software by measuring the distance between the edge of the ganglion and the leading tip of the longest 20 processes in each explant.

Ganglion dissociated cell cultures. To assess neurite outgrowth by isolated sensory neurons, we dissociated and cultured neurons from DRGs (Sachs et al., 2007).

Seven days after unilateral transection of the sciatic nerve, ipsilateral and contralateral L5

DRGs from Wlds, CCR2 -/-, and WT mice were removed and desheathed. Except where noted all reagents used in the DRG and SCG dissociations and culture were from Sigma-

Aldrich (St. Louis, MO). Ganglia were incubated in 0.125% collagenase A at 37°C for

1.5 h. The cells were then dissociated by gentle trituration using a P200 pipet in

Neurobasal A medium containing 2% B-27 serum free supplement, 2 mM glutamine

(both from Invitrogen, Life Technologies), 10 U/ml penicillin, and 10 µg/ml streptomycin. The dissociated cells were purified following the procedure of Gavazzi et al. (1999), by centrifugation through 15% BSA at 600 rpm for 6 min. Neurons were resuspended in Neurobasal A containing 50 µg/ml DNase (Type I) and centrifuged at

1000 rpm for 2 min. Supernatant was removed, and cells were resuspended in

Neurobasal A. Cells were gently plated (1 DRG/coverslip) onto 22 mm coverslips coated with 0.01% poly-L-lysine and 10 µg/ml laminin in a 6-well culture plate. Cells were allowed to adhere undisturbed for 20 min. Each coverslip was then overlaid with 2 ml of

Neurobasal A and cultured for 24 h at 37°C in 95% air/5% CO2.

To assess neurite outgrowth by isolated sympathetic neurons, we cultured dissociated SCGs (Hyatt Sachs et al., 2010). Seven days after unilateral axotomy, ipsilateral and contralateral SCGs were removed, desheathed, cut into 6 pieces, and incubated in 0.1% collagenase A at 37°C for 40 min. After washing, SCG were incubated in 0.1% trypsin at 37°C for 10 min. Cells were then dissociated in F12/Coon’s medium containing 340 ng/ml tri-iodo thyronine, 60 ng/ml progesterone, 400 ng/ml L-thyroxine,

38 ng/ml sodium selenite, 16 µg/ml putrescine,10 U/ml penicillin, 10 µg/ml streptomycin, and 3.5% Path-4 BSA (MP Biomedicals, Aurora, OH) by gentle trituration using a P200 pipet. Cells were gently plated (half a SCG/coverslip) onto 22 mm coverslips as described above for DRG neurons and allowed to adhere. Each coverslip was then overlaid with 2 ml of F12/Coon’s and cultured for 24 h at 37°C in 95% air/5%

CO2.

Analysis of neurite outgrowth in cultured neurons. Cells were fixed in 4% paraformaldehyde for 20 min at room temperature, washed in PBS, and labeled with a mouse monoclonal antibody to βIII tubulin (1:900 for DRG and 1:800 for SCG;

Promega, Madison, WI), followed by a 45 min incubation in an AlexaFluor488 labeled secondary (1:400, Molecular Probes, Grand Island, NY). Coverslips were placed onto slides with FluoroGel (Electron Microscopy Sciences). Neurons were visualized with a

Leitz epifluorescence microscope (Leica Microsystems Inc., Bannockburn, IL) at 10 or

25x and images were captured with a Hamamatsu ORCA 100 cooled CCD camera interfaced with SimplePCI software (Hamamatsu Corp., Bridgewater, NJ). MetaMorph was used to measure the longest neurite from each βIII tubulin positive neuron with a process of at least 1.5 times the diameter of the cell body.

Myelin visualization. To assess Wallerian degeneration in the sciatic nerve, myelin with visualized 7 days after unilateral axotomy. Cryostat sections were re- hydrated in distilled water for 5 min and dehydrated in 35% and 70% ethanol for 5 min each. They were then incubated in 0.1% Luxol Fast Blue (LFB) overnight at 60o C followed by washes in 95% ethanol and distilled water. The sections were then de-stained in 0.05% lithium carbonate for 30 sec, rinsed in 70% ethanol, and incubated in 70%, 95% and 100% ethanol for 5 min each. Finally the sections were transferred to xylene for 5 min and examined under a light microscope. The extent of staining was quantified as described above for TH.

Statistics. Data are expressed as the means + S.E.M. and were analyzed by one- way ANOVA (PCR data and comparisons of the ratio of CD11b staining between axotomized and contralateral tissues) or two-way ANOVA followed by Tukey’s post-hoc

38 test. P values less than 0.05 were considered statistically significant.

2.5 Results

Decreased axotomy-induced macrophage influx into sensory and sympathetic ganglia in the Wlds mouse

As indicated previously, the Wlds mouse exhibits little macrophage accumulation in the distal sciatic nerve segment one week after nerve transection (Lunn et al., 1989; Brown et al., 1991b). Later it was shown that this effect represented a delay, rather than a block, in macrophage accumulation (Sommer and Schafers, 1998). As a positive control for our immunohistochemical experiments in peripheral ganglia, we also examined the accumulation of macrophages in the distal segment of the sciatic nerve 7 d after nerve transection in WT and Wlds mice. Whereas a large increase in macrophages was found in the nerves of WT animals, no significant increase was found in the Wlds mouse, confirming these earlier studies (Fig.2.1a-f).

If the phenotype of the Wlds mouse after axotomy was entirely restricted to the distal nerve segment, as has generally been assumed, there would be no reason to expect a change in monocyte entry into the L5 DRG, which is about 2 cm away from the injury site. Nevertheless, whereas a 4-fold increase in macrophage accumulation was seen in the

DRGs of WT mice 7 d after axotomy (Fig. 2.2a-d), no significant increase was seen in

Wlds mice (Fig. 2.2a,b,e,f).

We next explored this phenomenon in a sympathetic ganglion, the SCG. Here, as in the DRGs, the increase in macrophage influx into the ganglia in WT mice was substantial (3.6-fold; Fig. 2.2g). There was also a significant increase in SCGs from Wlds

39 mice (Fig. 2.2g), however, the density of macrophages was significantly less than in WT ganglia (Fig. 2.2g,i-l). Thus, similar to the reduced accumulation of macrophages in the distal nerve stump after axonal transection, there is a reduced accumulation in regions of axotomized neuronal cell bodies in Wlds mice. The difference between the results in

DRGs and SCGs can be seen most dramatically by considering the percentage increases in macrophage staining between the WT and Wlds mice (Fig. 2.2b, h).

Regulation of CCL2 mRNA in peripheral ganglia after axotomy

What might account for the decreased macrophage accumulation in these ganglia? As already noted, CCL2 mRNA is increased in neurons in both sensory and sympathetic ganglia within hours after axotomy, and can, therefore, be considered a RAG (Schreiber et al., 2001; Costigan et al., 2002; Boeshore et al., 2004; Tanaka et al., 2004). We sought to determine if expression of this gene differed in WT and Wlds mice. L5 DRGs were examined 6, 24, and 48 h after the sciatic nerve was transected unilaterally. CCL2 mRNA was not detected in contralateral ganglia from either genotype at any of the three time points examined. In the axotomized DRGs, CCL2 mRNA was detected in both genotypes at all-time points; however, the levels in the ganglia from WT mice were significantly higher than in Wlds mice at 24 and 48 h and the difference was very close to significance at 6 h (Fig. 2.3a).

CCL2 mRNA was also undetected in contralateral SCGs of both genotypes. The elevation of this chemokine after axotomy was significantly greater in SCGs from WT animals compared to those from Wlds animals at all-time points examined (Fig. 2.3b).

These data indicate that the difference in CCL2 expression is likely to be the cause of the

40 reduced accumulation of macrophages in axotomized ganglia in the Wlds mouse.

Macrophage accumulation in sensory and sympathetic ganglia in the CCR2 -/- mouse

CCL2 produces its chemotactic effect on circulating monocytes via the CCR2 receptor

(Deshmane et al., 2009). For example, this receptor has been shown to be important for monocyte infiltration into the lesioned sciatic nerve (Siebert et al., 2000). Nevertheless, there are no data available on whether monocyte entry into peripheral ganglia after axotomy is dependent on CCL2, CCR2, or both of them. We examined this question in

WT mice and in mice in which CCR2 had been knocked out.

As a starting point, we examined macrophage accumulation in the distal segment of the sciatic nerve 7 d after nerve transection in WT and CCR2 -/- mice. In our hands, the increase in macrophage accumulation after axotomy was totally abolished in the knockout animals (Fig. 2.4a-e). Furthermore, as shown already, in WT animals 7 d after unilateral nerve transection, macrophage accumulation in the ipsilateral L5 DRG is dramatically increased (Fig. 2.4f,g,h); however, no significant increase was seen under the same conditions in the CCR2 -/- mouse (Fig. 2.4f,i,j). In the SCG, while there was an increase in macrophage accumulation in ganglia from CCR2 -/- mice (Fig. 2.4k,n,o), the density of macrophages was significantly smaller than that seen in ganglia from WT animals (Fig. 2.4k,l,m). These data raise the possibility that in the SCG a second chemokine in addition to CCL2 might be involved in bringing monocytes into the ganglion after axotomy.

The level of CCL2 mRNA was also determined in DRGs and SCGs in CCR2 -/- mice 24 h after axotomy. No differences were found between WT and CCR2 -/- mice

(Fig. 2.3c).

Wallerian degeneration in Wlds and CCR2 -/- mice

Wallerian degeneration was compared in the mutant mice to that in WT mice. As originally shown by Lunn et al. (1989) and replicated by others, little Wallerian degeneration was seen in Wlds mice one week after transection of the sciatic nerve (Fig.

2.5a,c,f), measured in our experiments by the disappearance of staining for myelin proteins. Since it is widely believed that normal Wallerian degeneration is dependent on infiltrating monocytes into the distal nerve (e.g., Bruck, 1997; Dailey et al., 1998; Luk et al., 2003; Barrette et al., 2008; Vargas et al., 2010; Gaudet et al., 2011; Rotshenker, 2011) and since there was no significant influx of macrophages in the sciatic nerves of the

CCR2 -/- mice 7 d after transection (Fig. 2.4a), we assumed that Wallerian degeneration would be slow in these animals. However, the clearance of myelin proteins in CCR2 -/- mice at 7 d was similar to that in WT mice and dramatically different from that in the

Wlds mice (Fig. 2.5a,d,g).

As of yet, there have been no reports of the relative speed of Wallerian degeneration in the sympathetic nervous system among WT, Wlds and CCR2 -/- mice.

Therefore, we determined if sympathetic axons from Wlds and CCR2 -/- mice in fact exhibit slow Wallerian degeneration. For this purpose, we examined the sympathetic innervation of one of the main targets of the SCG, the submandibular gland (Flett and

Bell, 1991), using immunohistochemistry for TH, the cytoplasmic enzyme that catalyzes the rate-limiting step in catecholamine biosynthesis. This approach was chosen because

TH is localized specifically in sympathetic neurons. Seven days after the internal and

42 external carotid nerves were severed, TH-immunoreactivity was no longer detectable in the submandibular glands of WT mice (Fig. 2.6a,b,e); however, considerable staining persisted in the glands from Wlds mice (Fig. 2.6a,c,f). In contrast, the disappearance of

TH-immunoreactivity in CCR2 -/- mice was similar to that seen in WT mice (Fig.

2.6a,d,g).

These data on Wallerian degeneration in the sciatic nerve and in the submandibular gland are of considerable interest in interpretation of results on neurite outgrowth presented in the following sections since both Wlds and CCR2 -/- mice have no significant increase in macrophage accumulation in the distal nerve segment only the

Wlds mice have slow Wallerian degeneration.

The conditioning lesion effect in DRG and SCG explants from Wlds mice

It has been established that regeneration of the sciatic nerve after nerve crush is decreased, or perhaps delayed, in Wlds mice. As already noted, this decrease has been attributed to the slow Wallerian degeneration that occurs in these animals. Based on this universally accepted view, neurite outgrowth from peripheral ganglia placed in a permissive growth environment would be expected to be normal. Nevertheless, given our finding of a decreased expression of a RAG in Wlds neurons in sensory and sympathetic ganglia and a change in the non-neuronal cellular environment of the axotomized neurons, we wondered whether these neurons might also exhibit a smaller increase in outgrowth in vitro in response to axotomy. To examine this possibility, we first looked at neurite outgrowth in explant cultures (i.e., in the absence of the distal nerve stump) 7 d after a conditioning lesion. Both DRGs and SCGs from WT animals have been shown to

43 have increased outgrowth under these conditions (Ekstrom et al., 2003; Shoemaker et al.,

2005; Hyatt Sachs et al., 2010).

As expected, DRG neurons from WT animals 7 d following a conditioning lesion exhibited an increase in neurite outgrowth both 24 and 48 h after explantation (Fig. 2.7a- c); however, in ganglia from Wlds mice, there was no significant effect of the conditioning lesion at either time (Fig. 2.7a,b,d). In striking contrast, in the SCG, comparable conditioning lesion responses were observed in ganglia from both WT (2.3- fold increase) and Wlds (2.3-fold increase) mice at 48 h, and the magnitudes of these responses were not significantly different (Fig. 2.7e-h).

The conditioning lesion effect in the DRG and SCG from CCR2 -/- mice

Our data establish that macrophage accumulation in peripheral ganglia after axotomy is abolished or significantly reduced in both Wlds and CCR2 -/- mice and that the conditioning lesion response is abolished in explants of the DRGs from Wlds mice. We, therefore, examined the growth response to a conditioning lesion in the CCR2 -/- mice.

As expected, there was an increase in outgrowth in response to a conditioning lesion in both the DRG (Fig. 2.8a-c) and the SCG (Fig. 2.8e-g) explants in ganglia taken from WT animals. Strikingly, however, this conditioning lesion effect was abolished in both types of ganglia taken from CCR2 -/- mice (Fig. 2.8a,b,d and Fig. 2.8e,f,h). These results indicate a strong relationship between macrophage accumulation in ganglia and the conditioning lesion response, and they provide a case in which inhibition of the response can be completely dissociated from changes in the rate of Wallerian degeneration.

Examination of the conditioning lesion effect in dissociated neurons from both Wlds

44 and CCR2 -/- mice

To determine whether the decreased neurite outgrowth seen in explants represented, at least in part, a neuron autonomous effect, we examined the conditioning lesion response in dissociated neuronal cultures. Demonstration that a conditioning lesion response in

WT neurons can be seen in dissociated DRG and SCG cultures was first observed by Hu-

Tsai et al. (1994) and Shoemaker et al. (2005), respectively.

The longest neurite was measured from every βIII tubulin immunostained neuron that had a neurite at least 1.5 times the diameter of the cell body. As we have previously reported (Shoemaker et al., 2005; Sachs et al., 2007), 7 d after a conditioning lesion, there was a substantial increase in neurite outgrowth from both sensory and sympathetic neurons from WT animals (Fig. 2.9a,b,e, and Fig. 2.9h,i,l, respectively). In dissociated neurons from DRGs of both Wlds and CCR2 -/- mice, this conditioning lesion effect was significantly reduced; nevertheless, the effect was not abolished (Fig. 2.9a). In neurons from SCGs of Wlds mice, no change in the conditioning lesion effect was seen compared to that in WT mice (Fig. 2.9h,j,m). In sympathetic neurons from CCR2 -/- mice, the conditioning lesion effect was significantly reduced, but again not abolished (Fig.

2.9h,k,n). Thus, under conditions in which macrophage accumulation in peripheral ganglia is substantially reduced, the conditioning lesion response in neurons dissociated from those ganglia is reduced.

2.6 Discussion

Macrophages are known to foster nerve regeneration; however, virtually all previous studies have assumed they do so by accumulating in the distal nerve segment,

45 promoting Wallerian degeneration, and perhaps triggering growth factor expression. Such studies include investigations of the Wlds mouse (e.g., Brown et al., 1991b) and of blockade of monocyte infiltration into the distal nerve, for example, by means of complement depletion (Dailey et al., 1998) or insertion of a thymidine kinase transgene on the CD11b promoter (Barrette et al., 2008). Under these conditions both Wallerian degeneration and nerve regeneration are inhibited. Though it has been known for about

20 years that macrophages also accumulate in axotomized sympathetic and sensory ganglia after axotomy (Lu and Richardson, 1993; Schreiber et al., 1995), no consideration has been given to the possibility that such accumulation was also blocked in these studies.

Macrophage accumulation in the lesioned sciatic nerve depends on CCL2 expression by Schwann cells (Perrin et al., 2005) and by the expression of its receptor

CCR2 on monocytes (Siebert et al., 2000; Abbadie et al., 2003). In addition, in peripheral ganglia macrophage accumulation is preceded by CCL2 expression by the axotomized neurons (Schreiber et al., 2001; Tanaka et al., 2004). We report here that axotomy- induced macrophage accumulation is totally absent in the L5 DRG in the Wlds mouse and that this is accompanied by a substantially smaller increase in CCL2 mRNA compared to that seen in WT ganglia. In the Wlds SCG, macrophage accumulation is also reduced, but it is not abolished in spite of a reduced upregulation of CCL2 mRNA similar to that in the

DRG. These results suggest that a second chemokine, in addition to CCL2, operates in the SCG and a number of candidates exist for this molecule (e.g., see Surmi and Hasty,

2010; Ingersoll et al., 2011). When we tested the importance of CCL2/CCR2 signaling further by looking at CCR2 -/- mice, we again found a total blockade of macrophage

46 accumulation in the DRG and a less robust but significant increase in the SCG.

To test whether these changes in macrophage accumulation in ganglia affect neurite outgrowth, we utilized the conditioning lesion response. In this response, a prior lesion to a nerve stimulates the regenerative response to a subsequent lesion in vivo

(McQuarrie and Grafstein, 1973). The conditioning lesion also increases neurite outgrowth in sensory and sympathetic neurons in explant or dissociated cultures (Hu-Tsai et al., 1994; Edstrom et al., 1996; White et al., 1996; Shoemaker et al., 2005). We used these in vitro approaches with the Wlds mice because interpretation of in vivo experiments would be complicated by the influence of the non-degenerating distal nerve segment, which as noted is assumed to inhibit regeneration.

In the Wlds DRG explants, neurite outgrowth does not increase after a conditioning lesion. In the SCG, on the other hand, no difference was seen in the conditioning lesion response between WT and Wlds mice. A likely explanation is that, whereas macrophage accumulation in the DRG after axotomy was completely abolished in the Wlds mouse, it was only somewhat diminished in the SCG. This relationship between the extent of macrophage accumulation in these two peripheral ganglia and subsequent neurite outgrowth supports our hypothesis that monocyte infiltration into peripheral ganglia plays a crucial role in triggering the conditioning lesion response.

The results of neurite outgrowth obtained in dissociated cell cultures bear both similarities and differences with those from explants. For example, in both explants and cell cultures, Wlds DRG neurons, but not SCG neurons, exhibited less neurite outgrowth after a conditioning lesion compared to WT animals. Nevertheless, the conditioning

47 lesion effect was not abolished in cell cultures from DRGs as it was in explants.

In both DRG and SCG explant cultures from CCR2 -/- mice, no conditioning lesion effect was seen. In dissociated sensory and sympathetic CCR2 -/- neurons, the conditioning lesion effect was significantly reduced, but it was not abolished. Presumably the differences between explant and cell cultures reflect the disruption of the cellular relationships between neurons and non-neuronal cells and a reduction in the presence of non-neuronal cells (e.g., satellite/Schwann cells, macrophages and other leukocytes, endothelial cells, and fibroblasts). Based on these results, we hypothesize that the effect of macrophages on the conditioning lesion response results in part from a stimulatory neuron autonomous event and in part from an effect on a non-neuronal cell type, perhaps inhibitory in nature.

A peripheral conditioning lesion also promotes regeneration of the lesioned central processes of DRG neurons (Richardson and Verge, 1987; Chong et al., 1999;

Neumann and Woolf, 1999). In a recent article, Salegio et al. showed that this effect is blocked after intravenous injections of clodronate liposomes, a treatment which decreases the number of circulating monocytes (Salegio et al., 2011). However, given the systemic effect of these liposomes, macrophages would be decreased at both the CNS and the PNS injury sites along with a decrease in the DRGs. Thus, no conclusion can be drawn from the study as to the importance of monocyte entry specifically into the DRG. However, Lu and Richardson (1991) did report that producing inflammation within the DRG or injecting peritoneal macrophages directly into these ganglia stimulated regeneration of the central, though not the peripheral, process of DRG neurons.

With few exceptions, studies on the Wlds mouse have focused on understanding the mechanism of the ~10-fold slower degeneration of the distal axonal segment that occurs after axotomy compared to that in WT axons. The expectation from early studies on regeneration in the Wlds mouse has been that delayed regeneration is the direct result of delayed degeneration and that no effects should be seen on neurite outgrowth of Wlds neurons in culture. Another argument against the idea that the delayed regeneration represents a change in the growth response of Wlds neurons was the finding that expression of three RAGs (c-JUN, growth-associated protein (GAP)-43, and the medium neurofilament protein) did not differ between mutant and WT animals (Brown et al.,

1994; Gold et al., 1994; Bisby et al., 1995). Of course, these are only three out of the hundreds of genes whose expression is altered after axotomy (Costigan et al., 2002;

Boeshore et al., 2004). Our finding that the injury-induced upregulation of CCL2 is lessened in the Wlds mouse in sensory and sympathetic neurons is the first demonstration of a change in expression of a RAG in neurons in these mutants.

Our data also strongly suggest that CCL2 acting via CCR2 expressed on monocytes is involved in monocyte entry into peripheral ganglia. There are, however, two caveats in interpreting our CCR2 -/- data. First, it is known that CCR2 is involved in the exit of monocytes from the bone marrow (Ingersoll et al., 2011). Therefore, the extent that knocking out this receptor diminishes monocyte entry into the blood stream and the extent to which it blocks entry from the blood stream into particular tissues must be resolved with further experiments. Second, it has been demonstrated that CCR2 receptors are present on DRG neurons in addition to being present on macrophages (White et al.,

2009), and, in a single study, CCL2 was shown to promote neurite outgrowth from

49 neurons of the statoacoustic ganglion of the inner ear (Bianchi et al., 2005). Thus CCL2 might have a direct growth promoting effect on sensory neurons.

In the CCR2 -/- mice, no conditioning lesion effect was seen in either the explanted DRG or SCG. The importance of macrophage action in axotomized ganglia is perhaps most directly demonstrated in these animals in which Wallerian degeneration appears to be similar to that in WT animals. While all of our outgrowth experiments were designed to minimize any possible inhibition of regeneration produced by the distal nerve stump, we found in addition and to our surprise, normal Wallerian degeneration in the

CCR2 -/- mice in spite of the fact that we found no accumulation of macrophages in the distal nerve. This finding that infiltrating monocytes are not required for normal

Wallerian degeneration is contrary to the view presented in three recent reviews (Gaudet et al., 2011; Rotshenker, 2011; Wang et al., 2012); however, it is supported by a single older study of Perry et al. (1995) in which mice were subjected to whole body irradiation.

Our results also seem in conflict with those reported by Barrette et al. (2008; see Fig. 6 of their paper), in which the clearance of LFB staining in their thymidine kinase mutants was substantially reduced. Further studies are required to determine the basis of these differences.

Many questions remain concerning macrophage action within ganglia. For example, we do not know the phenotype of the hematogenous macrophages in these ganglia (e.g., M1 or M2), what these cells might be releasing that stimulates neurite outgrowth, and whether other immune cells participate in this stimulation (Gordon and

Martinez, 2010; Gaudet et al., 2011; Gensel et al., 2012; Kurimoto et al., 2012). The growth promoting effect of lens injury on retinal ganglion cells though initially attributed

50 to macrophages is now thought to be mediated by either neutrophils or astrocytes (Leon et al., 2000; Leibinger et al., 2009; Kurimoto et al., 2012). Another question of interest is whether microglia could play a similar role in the CNS as macrophages do in the PNS, a possibility raised by the recent findings of Shokouhi et al. (2010).

In summary, our study involves four key findings. First, it is highly likely that

CCL2/CCR2 signaling is important in the infiltration of monocytes into peripheral ganglia after axotomy. Second, the decreased sensory nerve regeneration seen by others in Wlds mice involves both a decrease in the neurons’ growth response, as well as probably the long hypothesized influence of extrinsic factors. Third, Wallerian degeneration can proceed in the absence of infiltrating monocytes. Fourth, macrophage accumulation in peripheral ganglia after injury plays an essential role in the response of neurons to a conditioning lesion. Together these findings establish an important new site of macrophage action in promoting nerve regeneration.

Figure 2.1

Figure 2.1. Macrophage accumulation is significantly increased in the distal segment of the transected sciatic nerve in WT mice but not in Wlds mice. The sciatic nerve in WT and Wlds mice was transected unilaterally (Axotomy), and, 7 d later, the nerve segment distal to the site of transection was immunostained with a CD11b antibody to visualize macrophages. A comparable segment of the contralateral nerve was also examined. The extent of staining was quantitated as a percentage of the section area that was stained (a).

The number of nerves examined in each group is given within each bar of the histogram.

The staining in the axotomized nerve is also expressed as a mean ratio of staining in the ipsilateral and contralateral nerve for each animal X 100 (b). Micrographs are shown for contralateral (c,e) and axotomized (d,f) nerves from WT (c,d) and Wlds (e,f) animals. The data presented here and in all subsequent figures are means + S.E.M. ** p < 0.001. Scale bar = 20 µm.

Figure 2.2

Figure 2.2. The macrophage population in L5 dorsal root ganglia (DRGs) and superior cervical ganglia (SCGs) following unilateral axotomy is significantly diminished in Wlds mice compared to WT mice. The tissues examined were from the same groups of animals. One week after unilateral transection of the relevant nerves, the ipsilateral and contralateral ganglia were removed and immunostained for CD11b positive cells. The data are presented both as the percent of the section area that was stained (a,g) and as the mean ratio of staining in the ipsilateral and contralateral ganglia for each animal X 100

(b,h). Micrographs are shown for contralateral DRGs (c,e) and SCGs (i,k) and axotomized DRGs (d,f) and SCGs (j,l) from WT (c,d,i,j) and Wlds (e,f,k,l) mice. * p <

0.05, ** p < 0.001. Scale bar = 20 µm.

Figure 2.3

Figure 2.3. The axotomy-induced induction of CCL2 is diminished in DRGs (a) and

SCGs (b) in Wlds mice compared to WT mice. Six, twenty-four, and forty-eight hours after transection of the relevant nerves, axotomized and contralateral L5 DRGs and SCGs were extracted, and CCL2 mRNA was measured by RT-PCR. No detectable CCL2 mRNA was found in the contralateral ganglia. CCL2 mRNA was also measured in axotomized DRGs and SCGs from WT and CCR2 -/- mice (c). The data are normalized to GAPDH mRNA for each sample. * p < 0.05.

Figure 2.4

Figure 2.4. The axotomy-induced accumulation of macrophages in the sciatic nerve,

DRG, and SCG is diminished in CCR2 -/- mice compared to WT. The data show the extent of CD11b staining in sciatic nerves from WT and CCR2 -/- mice 7 d after unilateral axotomy (a). No effect of axotomy was seen in the CCR2 -/- animals.

Representative micrographs are shown from contralateral (b,d) and axotomized (c,e) sciatic nerves from WT (b,c) and CCR2 -/- (d,e) animals. Axotomy induced macrophage accumulation was abolished also in L5 DRGs from the same animals (f). Micrographs are shown from contralateral (g,i) and axotomized (h,j) DRGs from WT (g,h) and CCR2 -/-

(i,j) animals. Axotomy induced macrophage accumulation is lessened, but not abolished, in the SCGs from CCR2 -/- mice (k). Micrographs are shown from contralateral (l,n) and axotomized (m,o) SCGs from WT (l,m) and CCR2 -/- (n,o) animals. * p < 0.05, ** p <

0.001. Scale bars = 20 μm.

Figure 2.5

Figure 2.5. Seven days after the sciatic nerve was unilaterally transected, changes in reactivity for myelin proteins were determined in nerves from WT, Wlds and CCR2 -/- mice by staining with luxol fast blue. The distal nerve segments from WT and CCR2 -/- mice showed significantly less myelin staining compared to contralateral nerves, whereas axotomized nerves from Wlds mice retained >80% of myelin reactivity compared to contralateral nerves (a). The micrographs represent sections from the ipsilateral (e,f,g) and contralateral (b,c,d) nerves from WT, Wlds, and CCR2 -/- mice, respectively.* p <

0.05, ** p < 0.001. Scale bar = 20 µm.

Figure 2.6

Figure 2.6. Sympathetic nerve fibers in an autonomic target in the Wlds mouse, but not the CCR2 -/- mouse, exhibit slow Wallerian degeneration (a). To determine whether sympathetic neurons in vivo in the Wlds and CCR2 -/- mouse exhibit slow Wallerian degeneration, the internal and external carotid nerves were transected and, 7 d later, the ipsilateral and contralateral submandibular glands were stained for TH in WT, Wlds, and

CCR2 -/- mice. The micrographs represent sections from the ipsilateral (e,f,g) and contralateral (b,c,d) nerves from WT, Wlds, and CCR2 -/- mice, respectively. * p < 0.05,

** p < 0.001. Scale bar = 20 µm.

Figure 2.7

Figure 2.7. The conditioning lesion effect is abolished in DRG, but not SCG, explants from Wlds mice. Seven days after unilateral sciatic nerve transection, DRGs were placed in explant culture and neurite outgrowth was measured after 24 h (a) and 48 h (b). The phase micrographs are of individual conditioned DRG explants from WT (c) and Wlds (d) mice at 48 h. The arrows point to the endings of individual neurites. SCGs were examined under the same conditions. Phase micrographs are shown for conditioned SCG explants from WT (g) and Wlds (h) mice after 48 h. In the SCG no significant difference was seen between the two genotypes (e,f). * p < 0.05, ** p < 0.001. Scale bars = 100 µm.

Figure 2.8

Figure 2.8. The conditioning lesion effect is abolished in DRG and SCG explants from

CCR2 -/- mice. Seven days after unilateral sciatic nerve transection, DRGs were placed in explant culture and neurite outgrowth was measured after 24 h (a) and 48 h (b). The representative phase micrographs are of individual conditioned DRG explants from WT

Examination of SCG explants from CCR2 -/- mice revealed no conditioning lesion effect at 24 h (e) or 48 h (f). Representative micrographs of individual conditioned SCGs are shown for WT (g) and CCR2 -/- (h) mice at 48 h. * p < 0.05, ** p < 0.001. Scale bars =

100 µm.

Figure 2.9

Figure 2.9. Neurite outgrowth from dissociated cells after 24 h in culture from DRG (a- g) and SCG (h-n) neurons. The sciatic nerve and the internal and external carotid nerves were transected unilaterally in WT, Wlds and CCR2 -/- mice. Seven days later the DRGs and SCGs were removed, dissociated and cultured for 24 h. Neurite outgrowth was measured and expressed as mean length of the longest neurite from each neuron. Greater than 50 neurons were measured for each group, with the exception of the CCR2 -/- sham

DRG group in which 21 neurons were measured. A conditioning lesion response was observed in all three genotypes, but in DRGs the response was significantly less than WT in both mutants (a). In SCGs the magnitude of conditioning lesion response was similar in Wlds and WT mice but was significantly less in CCR2 -/- mice (h). Representative micrographs of single DRG neurons from contralateral (b,c,d) and conditioned (e,f,g) ganglia from WT (b,e), Wlds (c,f), and CCR2 -/- (d,g) mice. Representative micrographs of single SCG neurons from contralateral (i,j,k) and conditioned (l,m,n) ganglia from WT

(i,l), Wlds (j,m), and CCR2 -/- (k,n) mice. ** p < 0.001. # indicates that the mean length of CCR2 -/- contralateral SCG neurons was significantly longer than WT (p = 0.04). All scale bars = 100 µm.

Chapter 3: Overexpression of the Monocyte Chemokine CCL2 in Dorsal Root

Ganglion Neurons Causes a Conditioning-Like Increase in Neurite Outgrowth and

Does So via a STAT3 Dependent Mechanism

Jon P. Niemi, Alicia DeFrancesco-Lisowitz, Jared Cregg, Madeline Howarth, and

Richard E. Zigmond

Department of Neurosciences, Case Western Reserve University, Cleveland, OH

44106-4975, U.S.A.

Reprinted with permission from Experimental Neurology, Copyright, 2016.

3.1 Acknowledgements

This work was supported by a National Institutes of Health grant DK097223 to R.E.Z.

J.P.N was supported by a training grant NS077888. Breeding and genotyping of animals were carried out by the Case Western Reserve Visual Sciences Specialized Animal

Research Core, and imaging of the DRG dissociated cell cultures was conducted with the assistance of Scott Howell of the Visual Sciences Microscopy and Digital Imaging Core

(EY11373). The authors thank Yi-Lan Weng (Johns Hopkins University) for invaluable assistance in the design and creation of the AAV5-CCL2. We also thank Dr. Stephen

Strittmatter and Dr. William Cafferty (Yale University) for the SPRR1a antibody. Finally we thank Dr. Molly Ingersoll, Jane Lindborg and Dr. Angela Filous for helpful discussions and comments on the manuscript.

3.2 Abstract

Neuroinflammation plays a critical role in the regeneration of peripheral nerves following axotomy. An injury to the sciatic nerve leads to significant macrophage accumulation in the L5 DRG, an effect not seen when the dorsal root is injured. We recently demonstrated that this accumulation around axotomized cell bodies is necessary for a peripheral conditioning lesion response to occur. Here we asked whether overexpression of the monocyte chemokine CCL2 specifically in DRG neurons of uninjured mice is sufficient to cause macrophage accumulation and to enhance regeneration or whether other injury- derived signals are required. AAV5-EF1α-CCL2 was injected intrathecally, and this injection led to a time-dependent increase in CCL2 mRNA expression and macrophage accumulation in L5 DRG, with a maximal response at 3 weeks post-injection. These changes led to a conditioning-like increase in neurite outgrowth in DRG explant and dissociated cell cultures. This increase in regeneration was dependent upon CCL2 acting through its primary receptor CCR2. When CCL2 was overexpressed in CCR2 -/- mice, macrophage accumulation and enhanced regeneration were not observed. To address the mechanism by which CCL2 overexpression enhances regeneration, we tested for elevated expression of regeneration-associated genes in these animals. Surprisingly, we found that

CCL2 overexpression led to a selective increase in LIF mRNA and neuronal phosphorylated STAT3 (pSTAT3) in L5 DRGs, with no change in expression seen in other RAGs such as GAP-43. Blockade of STAT3 phosphorylation by each of two different inhibitors prevented the increase in neurite outgrowth. Thus, CCL2 overexpression is sufficient to induce macrophage accumulation in uninjured L5 DRGs

72 and increase the regenerative capacity of DRG neurons via a STAT3-dependent mechanism.

3.3 Introduction

Injury to peripheral nerves activates an inflammatory response that plays a significant role in axonal regeneration and functional recovery (Popovich and Longbrake,

2008; Bastien and Lacroix, 2014; DeFrancesco-Lisowitz et al., 2014). In response to axotomy, both injured peripheral neuronal cell bodies and Schwann cells distal to the injury site upregulate and release the C-C class chemokine 2 (CCL2), also known as monocyte chemoattractant protein-1 (MCP-1; Toews et al., 1998; Schreiber et al., 2001;

Subang and Richardson, 2001; Tanaka et al., 2004). As a result of the expression of this chemokine, monocytes extravasate into both peripheral ganglia and the nerve distal to the injury site, and macrophage accumulation in these areas remains elevated for several weeks (Perry et al., 1987; Lu and Richardson, 1993; Schreiber et al., 1995). Traditionally, investigation of this inflammatory process has focused on actions in the nerve at, or distal to, the site of injury. After axotomy, the distal nerve undergoes Wallerian degeneration, a process that includes fragmentation of the axon and myelin sheath and phagocytosis of debris by macrophages and Schwann cells (Rotshenker, 2011). Clearance of this debris is considered to be a prerequisite for regeneration, as blockade of macrophage accumulation yields incomplete degeneration and suboptimal regeneration and functional recovery

(Bisby and Chen, 1990; Brown et al., 1991b; Brown et al., 1994; Reichert et al., 1994;

Dahlin, 1995; Dailey et al., 1998; Barrette et al., 2008).

It should be noted, however, that in these experiments macrophage accumulation in peripheral ganglia was also almost certainly blocked and this might account for some

73 of the decreased regeneration (Niemi et al., 2013). In fact, recent evidence has shown that the inflammatory response near axotomized neuronal cell bodies is necessary for axonal regeneration following a peripheral nerve injury. Utilizing a global knockout for the C-C class chemokine receptor 2 (CCR2), the primary receptor for CCL2, we showed that if macrophage accumulation in the DRG does not occur then axonal regeneration is significantly diminished (Niemi et al., 2013). A related study showed that intrathecal administration of minocycline, which blocked injury-induced ganglionic macrophage accumulation, also reduced regenerative capabilities of DRG neurons (Kwon et al.,

2013).

Exogenous activation of inflammation near neuronal cell bodies has also been shown to increase the regenerative capacity of neurons. Injection of C. Parvum into the

DRG, creating a local inflammatory response characterized by macrophage accumulation, significantly increased expression of GAP-43 and CGRP, two RAGs, and axonal regeneration after a dorsal root crush injury, though interestingly not after a sciatic nerve injury (Lu and Richardson, 1991, 1993, 1995).

The most widely studied example of inflammation-induced regeneration is in the optic nerve. Retinal ganglion cells (RGCs) are normally unable to regenerate their axons following optic nerve injury, but are able to do so after inducing an inflammatory reaction in the eye (Leon et al., 2000; Yin et al., 2003). Intraocular injection of zymosan leads to a dramatic increase in the expression of oncomodulin (OCM), a protein that has been proposed to play a key role in inflammation-induced regeneration in the optic nerve

(Yin et al., 2006; Kurimoto et al., 2010). Other investigators have shown that lens injury-

74 induced optic nerve is dependent upon the upregulation and actions of two gp130 cytokines, CNTF and LIF (Leibinger et al., 2009).

While exogenous activation of local inflammation near neuronal cell bodies is able to greatly increase the regenerative capacity of those neurons, the mechanism by which this occurs and the specific role macrophages play in this response are poorly understood. The inflammatory responses activated by zymosan, and other inflammatory agents, are complex and involve several immune cell types, including CD4-positive cells, neutrophils, and macrophages (Gantner et al., 2003; Kurimoto et al., 2013; Baldwin et al.,

2015). We sought to determine if macrophage accumulation alone, without an axonal injury, could increase the regenerative capacity of DRG neurons. We addressed this question by overexpressing the monocyte chemokine CCL2 in uninjured lumbar DRGs and examined whether this singular change was sufficient to cause macrophage accumulation and increased axonal regeneration.

3.4 Materials and Methods

Generation and production of an adeno-associated virus (AAV5) CCL2 overexpression vector. Mouse cDNA encoding the open reading frame of CCL2

(Origene; Rockville, MD) was cloned into a pAAV shuttle vector containing the AAV inverted terminal repeats with an expression cassette composed of the human elongation factor-1 alpha (EF1α) promoter, a multiple cloning site, the woodchuck hepatitis virus post-transcriptional regulatory element (WPRE), and a polyadenylation signal. The CCL2 insert was generated by PCR using the following primers: Forward 5’ATA GTC GAC

ATG CAG GTC CCT GTC ATG CTT CTG G 3’ and Reverse 5’ ATA AAG CTT CTA

GTT CAC TGT CAC ACT GGT CAC TCC TA 3’. PCR products were digested by SalI

75 and EcoRV and subcloned into the MCS of the pAAV shuttle vector to generate the pAAV-EF1α-CCL2 vector. Recombinant AAV5 vectors were generated by transient co- transfection of HEK-293T cells with 37.5 µg of total DNA containing 7.5 µg of pAAV-

EF1α-CCL2, 7.5 µg of AAV5 rep-cap plasmid, and 22.5 µg of pAD helper plasmid using the calcium phosphate method. Seventy-two hours after transfection, cells were collected and lysed with 10% chloroform and AAV particles were purified using PEG8000

(Promega; Madison, WI) and benzonase nuclease (Life Technologies; Carlsbad, CA) and concentrated using an Ultra-4 centrifugal filter (EMD/Millipore; Billerica, MA). Titers were determined by quantitative PCR for viral genomic copies extracted from proteinase

K-treated viral particles using the following primers: Forward: 5’ATA GTC GAC ATG

CAG GTC CCT GTC ATG CTT CTG G 3’ and Reverse 5’ TTG TAG CTC TCC AGC

CTA CTC ATT GG 3’. For the present experiments, the following vectors were produced: AAV5-EF1α-CCL2: 2.2x1013 GC/ml and AAV5-EF1α-eYFP: 1.52x1013

GC/ml.

Animals and Sciatic Nerve Transection Injury (SNI). Eight to twelve-week-old male wild type (WT) mice [C57BL/6J (Jackson Laboratories, Bar Harbor ME)] and mutant mice [C57B6.129S4-Ccr2tm1Ifc/J (CCR2 -/-; Jackson Laboratories)] were utilized for this study. The animals were housed under a 12 h:12 h light:dark cycle with ad libitum access to food and water. SNI was performed on WT mice under isoflurane anesthesia. Briefly, the right sciatic nerve was exposed and transected proximal to the trifurcation and the wound closed with a wound clip. The left sciatic nerve was then exposed but not transected to serve as an internal control. One, two, three, or four weeks after intrathecal injection or 7 d after SNI, the animals were sacrificed by CO2 inhalation,

76 and the lumbar-level DRGs, sciatic nerves, and spinal cords were removed for immunohistochemical or molecular biological analysis. All animal procedures were approved by the Case Western Reserve University’s Institutional Animal Care and Use

Committee.

L5 laminectomy and intrathecal injection. The intrathecal injection method was modified from Parikh et al. (2011). Briefly, mice were anesthetized using isoflurane. A small incision was made in the back skin of the mouse and the spinal column was exposed. The site of viral injection was between lumbar levels L5 and L6, where a small laminectomy was performed to expose the dura. The injection was performed with a borosilicate glass capillary (WPI; Sarasota, FL) pulled to a fine point, attached by polyethylene tubing (Thermo Fisher Scientific; Waltham, MA) to a Hamilton syringe.

The glass capillary remained at midline and was slowly inserted underneath the dura and further advanced in the subarachnoid space. Ten microliters of AAV5-EF1α-CCL2 or

AAV5-EF1α- eYFP were slowly injected using 1% trypan blue added to the AAV particle solution to visualize the injection, and the capillary remained in place for 2 min after the injection. The paraspinal muscles and fascia were repositioned, and the incision closed with wound clips. One, two, three, or four weeks after injection mice were sacrificed and lumbar DRG were harvested for qPCR, IHC, or explant culture.

Immunohistochemistry (IHC). Generally, lumbar DRGs from CCR2 -/- and WT mice were removed, and the ganglia were desheathed and fixed by immersion in 4% paraformaldehyde. The tissues were cryoprotected in 30% sucrose and embedded in

Tissue-Tek O.C.T. compound (Electron Microscopy Sciences; Hatfield, PA). IHC was performed on 10 µm cryostat sections. For quantification of macrophages, a rat

77 monoclonal antibody to CD11b (also known as Mac1, CR3, and integrin αM; 1:100;

EMD/Millipore) or CD68 (1:200; AbD Serotec, Oxford, UK) was incubated with tissue sections overnight at 4oC. Given that the same effect was observed with CD11b and

CD68, the cells labeled by either of these markers will be referred to as macrophages throughout the paper. For quantification of RAG proteins, a rabbit monoclonal antibody to pSTAT3 (Y705; 1:100; Cell Signaling, Danvers, MA), a rabbit polyclonal antibody to

ATF3 (1:200; Santa Cruz Biotechnology; Dallas, TX), or a rabbit polyclonal antibody to small proline-rich repeat protein 1a (SPRR1a), generously provided by Dr. Stephen

Strittmatter and Dr. William Cafferty (Yale University), was incubated with tissue sections overnight at 4°C. For YFP detection, a rabbit antibody against GFP (1:500; Life

Technologies) was incubated with tissue sections for 3 d at 4°C. After washing, the sections were incubated in Cy3 secondary antibody (1:400; Jackson ImmunoResearch

Laboratories, Inc.; West Grove, PA) or AlexaFluor 488 secondary antibody (1:500; Life

Technologies) for 1 h. In all experiments, sections not exposed to the primary antibody were included for each experimental group. Images were captured at 25x magnification using HCImage software (Hamamatsu Corporation; Bridgewater, NJ) and then quantified using MetaMorph software (Version 7.6.3.0, Molecular Devices; Downingtown, PA).

Data are represented as the percentage of the imaged area that was positively immuno- labeled.

Real time PCR (RT-PCR). The expression of CCL2 mRNA was analyzed by RT-

PCR. One, two, three, or four weeks after intrathecal injection both L5 DRGs were removed and stored in RNAlater (Life Technologies) at 4oC. RNA was extracted from pairs of ganglia using the Ambion RNAqueous micro kit. Five samples were included for

78 each time point. Total RNA was quantified and 400 ng were reverse transcribed using a

High Capacity cDNA Reverse Transcription Kit (Applied Biosystems; Carlsbad, CA).

RT-PCR was performed in an ABI Step-One Plus, using prevalidated TaqMan expression assays [CCL2, Mm00441242; glyceraldehyde 3-phosphate dehydrogenase (GAPDH),

Mm99999915; ATF3, Mm00476032; GAP-43, Mm00500404; Jun, Mm00495062;

Smad1, Mm00484723; Sox11, Mm01281943; galanin, Mm00439056; interleukin (IL)-6,

Mm00446190; LIF, Mm00434762; iNos, Mm00440502; CD86, Mm00444543; CD16,

Mm00438875; arginase 1, Mm00475988; CD206, Mm00485148; Fizz1, Mm00445109;

Ym1, Mm00657889; Applied Biosystems], and samples were assayed in triplicate.

Relative expression was determined using the Comparative Ct Model (ΔΔCt) with

GAPDH as the housekeeping gene.

DRG explants. To assess the outgrowth of peripheral neurons in response to injury, we evaluated neurite outgrowth in explanted ganglia (Shoemaker et al., 2005).

Three weeks after intrathecal injection of the virus, L5 DRGs from uninjured CCR2 -/- and WT mice were removed, desheathed, placed on coverslips, and overlaid with 7.5 µl

Matrigel (Becton Dickinson, Franklin Lakes, NJ). Culture plates were placed in an incubator at 37oC for 5 min to allow gelling of the Matrigel before adding 1 ml F12 medium with the additives described in Hyatt Sachs et al. (2010). Phase-contrast images of neurite outgrowth from each DRG were captured at 24 and 48 h after explantation using an Axiovert 405 M microscope at 10x magnification. Neurite outgrowth was assessed using MetaMorph software by measuring the distance between the edge of the ganglion and the leading tip of the longest 20 processes in each explant. The average length of the 20 longest neurites is taken for each ganglion. At 48 h, explants were fixed

79 and labeled with an antibody against βIII tubulin (1:500; Promega) to visualize the outgrowth. Representative images were taken at 10x. In some experiments, 200 ng/ml of recombinant mouse CCL2 (R&D Systems; Minneapolis, MN) was added to the culture medium at the time of plating.

DRG dissociated cell culture. To assess neurite outgrowth from isolated sensory neurons, we dissociated and cultured DRG neurons from uninjured WT mice or mice injected with AAV5-CCL2 or AAV5-YFP (Sachs et al., 2007). L5 DRGs were removed, cleaned and desheathed. Except where noted all reagents used in the DRG dissociations and culture were from Sigma-Aldrich (St. Louis, MO). Ganglia were incubated in

0.125% collagenase A at 37°C for 1.5 h. The cells were then dissociated by gentle trituration using a P200 pipet in Neurobasal A medium containing 2% B-27 serum free supplement, 2 mM glutamine (both from Life Technologies), 10 U/ml penicillin, and 10

µg/ml streptomycin. The dissociated cells were purified following the procedure of

Gavazzi et al. (1999), by centrifugation through 15% BSA at 600 rpm for 6 min.

Neurons were resuspended in Neurobasal A containing 50 µg/ml DNase (Type I) and centrifuged at 1000 rpm for 2 min. Supernatants were removed, and cells were resuspended in Neurobasal A. Cells were gently plated (1/2 DRG/coverslip) onto 12 mm coverslips coated with 0.01% poly-L-lysine and 10 µg/ml laminin in a 12-well culture plate. Cells were allowed to adhere undisturbed for 20 min. Each coverslip was then overlaid with 1 ml of Neurobasal A and cultured for 24 h at 37°C in 95% air/5% CO2. In some experiments, 200 ng/ml of recombinant mouse CCL2 (R&D Systems) was added to the culture medium at the time of plating.

Inhibitors of signal transducer and activator of transcription 3 (STAT3)

80 phosphorylation: AG490 and STATTIC. Two inhibitors of STAT3 phosphorylation were utilized in DRG dissociated cell culture to assess the role STAT3 plays in CCL2 overexpression-induced neurite outgrowth. AG490 (50 µM; EMD Millipore; Meydan et al., 1996), a Janus kinase (JAK) inhibitor, and STATTIC (10 µM; EMD Millipore;

Schust et al., 2006), a STAT3 SH2 domain binding inhibitor, were used. L5 DRGs from mice 3 weeks after intrathecal injection with AAV5-YFP or AAV5-CCL2 were removed, dissociated and cultured as described above. At the time of plating, cells were cultured in

Neurobasal A medium containing AG490 (50 µM), STATTIC (10 µM), or DMSO (0.17

µl/ml) for 24 h.

Technologies, Grand Island). Coverslips were placed onto slides with FluoroGel

(Electron Microscopy Sciences). Fourteen-bit images were collected on a Leica DMI

6000 B inverted microscope (Leica Microsystems; Wetzlar, Germany) using a Retiga

Aqua blue camera (Q-imaging; Vancouver, British Columbia). Briefly, the entire coverslip was imaged at 10x and all resulting images were stitched together using the scan slide function in MetaMorph Imaging Software (Molecular Devices) to generate a full resolution composite image. The composite image was then subjected to neurite outgrowth analysis using the neurite outgrowth module in MetaMorph software. The longest neurite from each βIII tubulin-positive neuron with a process of at least 1.5 times the diameter of the cell body was measured.

Analysis of pixel intensity of pSTAT3 in cultured neurons. Cells were fixed in 4% paraformaldehyde for 20 min at room temperature, washed in PBS, and labeled with a mouse monoclonal antibody to βIII tubulin (1:900; Promega) and a polyclonal antibody against pSTAT3 (Y705; 1:80; Cell Signaling), followed by a 1 h incubation in an

AlexaFluor 488 labeled secondary antibody (1:400, Life Technologies) to visualize βIII tubulin and an AlexaFluor 647 labeled secondary antibody (1:200; Jackson Immuno) to visualize pSTAT3. Coverslips were placed onto slides with FluoroGel (Electron

Microscopy Sciences) and imaged as described above. The longest neurite from each βIII tubulin-positive neuron with a process of at least 1.5 times the diameter of the cell body was measured using the MetaMorph software. Data for neurite outgrowth was then expressed as the average length of the longest neurite for each group. To assess the average pixel intensity of pSTAT3 in culture, the nucleus of each neuron was outlined and the average pixel intensity within the nucleus was measured using the MetaMorph software. The same neurons were measured for both neurite outgrowth and pSTAT3 nuclear pixel intensity.

Statistics. Data are expressed as the means + S.E.M. and were analyzed by one- way or two-way ANOVA followed by Tukey’s post-hoc test. P values less than 0.05 were considered statistically significant.

3.5 Results

Intrathecal injection of AAV5-CCL2 results in a time-dependent overexpression of

CCL2 mRNA in uninjured L5 DRGs

To overexpress the chemokine CCL2 in uninjured DRGs, we constructed an

AAV5 vector encoding for CCL2 driven by the EF1α promoter (AAV5-CCL2; Fig.

3.1A). Intrathecal delivery of various AAV serotypes has been shown to allow for infection of DRG neurons (Wang et al., 2005b; Iwamoto et al., 2009; Parikh et al., 2011), and a screen of AAV serotypes revealed AAV5 as the most efficient for infection of

DRG neurons (Mason et al., 2010). To test the efficiency and localization of intrathecal delivery of AAV5, we first injected an AAV5 vector expressing eYFP driven by the

EF1α promoter (AAV5-YFP; Fig. 3.1B-E’). Three weeks after injection of 10 µl of high- titer virus, lumbar DRGs, lumbar spinal cord, and sciatic nerves were removed and immuno-labeled with an antibody against YFP to assess infection efficiency and localization. Lumbar DRGs showed significant YFP labeling of neuronal cell bodies with minor localization to axons (Fig. 3.1B-C’), resulting in a DRG neuron labeling efficiency of 37.2% ± 5.7%. Only sparse axonal YFP labeling was found in the sciatic nerve (Fig.

3.1D). AAV5-YFP also resulted in significant labeling of neurons in the lumbar spinal cord (Fig. 3.1E), with some specificity to neurons and processes in the dorsal horn (Fig.

3.1E’).

Intrathecal administration of AAV5-CCL2 resulted in a time-dependent overexpression of CCL2 mRNA in L5 DRGs (Fig. 3.1F). By 1 week after delivery,

CCL2 mRNA showed a 2.9-fold increase and by 3 weeks there was a 9.1-fold overexpression compared to AAV5-YFP controls. This level of overexpression is comparable to the injury-induced increase in CCL2 mRNA in the L5 DRG seen at 1 d after sciatic nerve transection (Niemi et al., 2013). The intrathecal delivery method was chosen specifically because it allows for significant CCL2 overexpression in DRGs, while leaving the ganglia uninjured.

CCL2 overexpression leads to increased CD11b-positive macrophage accumulation

Immuno-labeling with CD11b, a common marker employed for detecting macrophages, was used to assess if CCL2 overexpression was sufficient to increase macrophage accumulation within L5 DRGs. The immune response to injury, and specifically macrophage accumulation near injured neuronal cell bodies, has been shown to be an important mediator of regenerative capacity (Lu and Richardson, 1991; Kwon et al., 2013; Niemi et al., 2013). A time-dependent increase in CD11b-positive macrophages was seen after injection of AAV5-CCL2 compared to AAV5-YFP controls

(Fig. 3.2A-E). AAV5-YFP administration did not alter CD11b-positive macrophage accumulation compared to uninjured WT L5 DRGs (data not shown). Two weeks after viral delivery, there was a 3-fold increase in macrophage accumulation which increased to 4-fold by 3 weeks in AAV5-CCL2 mice compared to AAV5-YFP controls (Fig. 3.2A).

This increase in CD11b-positive macrophages following CCL2 overexpression is similar to the injury-induced increase seen in WT mice at 7 d (Niemi et al., 2013). We also utilized a second macrophage marker, CD68, and saw a time-dependent increase in CD68 labeling compared to YFP controls (Fig. 3.2F-J). CD68 showed considerably more staining than CD11b, which could indicate that some of the CD68-positive macrophages are CD11b-negative.

Peripheral nerve injury has been shown to induce monocyte entry into ganglia through CCL2/CCR2 signaling (Niemi et al., 2013). The present data show that CCL2 overexpression is sufficient to cause macrophage accumulation within the L5 DRG even in the absence of nerve injury.

CCL2 overexpression and macrophage accumulation cause a conditioning-like increase in neurite outgrowth in vitro

To test if CCL2 overexpression and the subsequent increase in macrophage accumulation in the absence of injury enhance the regenerative capacity of DRG neurons,

L5 DRGs from AAV5-CCL2 and AAV5-YFP mice were placed in explant culture 3 weeks after intrathecal injection. After 48 h in culture, DRG explants overexpressing

CCL2 showed significantly longer neurite outgrowth than DRGs from mice receiving the control virus (Fig. 3.3A-C). We also measured regeneration in DRG dissociated cell cultures. We have found that dissociation of the DRG does not completely eliminate glial cells from the culture (data not shown); however, it does disrupt the proximity of non- neuronal cells and neurons. After 24 h in culture, DRG neurons overexpressing CCL2 grew significantly longer neurites than neurons from mice receiving the control virus

(Fig. 3.3D-F). To examine whether the increased regenerative responses observed from both explant and dissociated cell cultures might be due to a direct effect of CCL2 acting on the DRG neurons at the time of regeneration, recombinant mouse CCL2 protein (200 ng/ml; Bianchi et al., 2005) was added to DRG explant cultures from WT mice.

Exogenous CCL2 addition to culture medium did not increase the length of axon regeneration after 48 h in culture when compared to controls (Fig. 3.3G-I). The lack of an effect of exogenous CCL2 on neurite outgrowth could be due to inefficient penetration of the protein into the explants. To allow for better exposure of DRG neurons to CCL2, exogenous CCL2 was added to DRG dissociated cell cultures. However, exogenous

CCL2 addition still had no effect on neurite outgrowth after 24 h in culture (Fig. 3.3J-L).

This suggests that CCL2 does not act directly on neurons to induce increased

85 regeneration. Thus, CCL2 exerts a conditioning-like effect to increase the regenerative capacity of neurons in vivo, prior to placing them in culture.

Macrophages have been shown to increase the regenerative response of DRG neurons through releasable factors (Kigerl et al., 2009); however, this ability was dependent upon the activation state of the macrophages. Macrophages stimulated to an anti-inflammatory state were growth-promoting, and pro-inflammatory macrophages were growth-inhibiting (Gordon and Taylor, 2005; Kigerl et al., 2009). Given, the increased regeneration we see with CCL2 overexpression, we sought to determine if macrophages in our system expressed an activation state which would be expected to support regeneration. We assayed the mRNA expression of a subset of anti-inflammatory

(arginase 1, CD206, Fizz 1, Ym 1) and pro-inflammatory (iNos, CD86, CD16) macrophage markers in L5 DRG (Fig. 3.4A; Gordon and Taylor, 2005; Martinez and

Gordon, 2014). Expression of the anti-inflammatory markers CD206 and Fizz 1 significantly increased, 3.1 and 11.2-fold, respectively, 3 weeks after injection of AAV5-

CCL2 compared to AAV5-YFP. mRNA levels for the pro-inflammatory macrophage markers did not change with CCL2 overexpression. This profile of anti-inflammatory macrophage marker expression is similar, though not identical, to the expression observed in WT L5 DRGs 7 d after SNI (Fig. 3.4B).

CCL2 acts through CCR2 to elicit changes in macrophage accumulation and regeneration

To determine if the changes mediated by CCL2 overexpression require the chemokine receptor CCR2, AAV5-CCL2 was delivered to CCR2 -/- mice (Boring et al.,

1998). CCR2 is localized to a subpopulation of monocytes (Geissmann et al., 2003;

Taylor and Gordon, 2003) and is necessary for their chemotactic responses to CCL2.

CCR2 also plays an important role in monocyte exit from the bone marrow (Han et al.,

1998; Serbina and Pamer, 2006). Three weeks after intrathecal injection of AAV5-CCL2, a significant increase in CCL2 mRNA (>7 fold) was found in CCR2 -/- L5 DRGs compared to that after injection with AAV5-YFP (Fig. 3.5A). However, CCL2 overexpression did not yield a significant increase in CD11b-positive macrophages in the

L5 DRG 3 weeks after virus administration in CCR2 -/- mice (Fig. 3.5B). Furthermore, overexpression of CCL2 in CCR2 -/- mice did not result in increased axonal regeneration, measured at 24 and 48 h in DRG explant culture 3 weeks after intrathecal injection (Fig.

3.5C). These data show that CCL2 must act through its primary receptor CCR2 to elicit increases in macrophage accumulation and regeneration in the DRG.

CCL2 overexpression leads to a selective increase in LIF mRNA and activation of

STAT3

To begin to ascertain the mechanism by which CCL2 overexpression in DRG neurons could lead to a conditioning-like increase in neurite outgrowth, the expression of various RAGs were screened in L5 DRGs 3 weeks after intrathecal injection of AAV5-

CCL2 or AAV5-YFP. The mRNA expression of LIF, IL-6, GAP-43, JUN, ATF3, galanin, Smad1, and Sox11 were assayed using RT-PCR and quantified using the comparative Ct method with GAPDH as the housekeeping gene. CCL2 overexpression resulted in a significant increase in LIF expression (4-fold) compared to YFP controls

(Fig. 3.6A). None of the other mRNAs showed a significant change in ganglia overexpressing CCL2. This is in contrast to the RAG expression profile observed 7 d

87 after SNI where all of the RAGs measured, with the exception of LIF, show significant upregulation (Fig. 3.6B).

LIF signals through a heterodimeric receptor containing the signaling subunit gp130, and activates the transcription factor STAT3 by inducing its phosphorylation and nuclear translocation (Symes et al., 1994). Therefore, we next tested whether STAT3 phosphorylation was increased in DRG neurons following CCL2 overexpression. Three weeks after intrathecal injection of virus, a 3-fold increase in pSTAT3-positive labeling was seen in AAV5-CCL2 L5 DRGs compared to AAV5-YFP controls and CCR2 -/- mice overexpressing CCL2 (Fig. 3.6C-E,G). This increased activation of STAT3 in uninjured DRGs following CCL2 overexpression was nevertheless significantly lower than the activation of STAT3 7 d after sciatic nerve transection in WT mice (Fig.

3.6F,G). ATF3 and SPRR1a, two other well-known RAGs (Bonilla et al., 2002; Seijffers et al., 2007), were also assayed by IHC. Neither ATF3 (Fig. 3.6H-J,L) nor SPRR1a (Fig.

3.6M-O,Q) was upregulated following CCL2 overexpression, though both were upregulated 7 d after sciatic nerve transection (Fig. 3.6K,L,P,Q). Thus, the CCL2 overexpression-induced increase in axonal regeneration may result from activation of

STAT3. pSTAT3 is necessary for the conditioning-like increase in neurite outgrowth observed with CCL2 overexpression

To test whether the CCL2 overexpression-induced increase in neurite outgrowth is dependent upon STAT3 activation, we treated DRG cultured neurons with STAT3 phosphorylation inhibitors. We utilized the Janus kinase (JAK) inhibitor AG490 (Meydan

88 et al., 1996) and the SH2-binding inhibitor STATTIC (Schust et al., 2006). Previous literature has shown that conditioning lesion-induced neurite outgrowth can be abolished through inhibition of STAT3 activation with AG490 in culture (Liu and Snider, 2001) or in vivo (Qiu et al., 2005).

After 24 h in culture, DRG neurons overexpressing CCL2 grew significantly longer neurites and showed significantly higher intensity nuclear pSTAT3 labeling than neurons from mice receiving the control virus (Fig. 3.7A-H). The addition of AG490 or

STATTIC to the culture medium at the time of plating significantly reduced the CCL2 overexpression-induced neurite outgrowth compared to vehicle-treated CCL2 overexpressing neurons (Fig 3.7A). This reduction in neurite outgrowth, similiar to

AAV5-YFP levels, coincided with a significant reduction in the pSTAT3 labeling intensity following treatment with AG490 or STATTIC (Fig. 3.7B). These data suggest that the CCL2 overexpression induced increase in neurite outgrowth is dependent upon the activation of pSTAT3.

3.6 Discussion

In the present study, we have demonstrated that overexpression of CCL2 in uninjured

DRG neurons is sufficient to cause macrophage accumulation in the ganglion. We further showed that this overexpression alone results in a conditioning-like increase in neurite outgrowth, increased LIF mRNA, and activation of STAT3 in DRG neurons.

Sciatic nerve injury produces dramatic changes in DRG neurons and in non-neuronal cell populations both distal to the site of injury and in the ganglion (Lieberman, 1971;

Bastien and Lacroix, 2014). These include activation and proliferation of satellite glial

89 cells (Hanani et al., 2002), dedifferentiation and proliferation of Schwann cells distal to the injury site (Napoli et al., 2012), and macrophage accumulation distal to the site of injury and in the DRG (Perry and Brown, 1992; Lu and Richardson, 1993). In addition, the expression of hundreds of genes is altered (Costigan et al., 2002; Nagarajan et al.,

2002). Therefore, the fact that overexpression of one gene, CCL2, is sufficient to increase the regenerative capacity of DRG neurons is remarkable.

Macrophage stimulation of axonal regeneration

Studies on the immune consequences of nerve injury have focused on actions at or distal to the injury site, in particular the role of macrophages in Wallerian degeneration

(for reviews see Bruck, 1997; DeFrancesco-Lisowitz et al., 2014). The actions of multiple immune cells in the distal nerve also may have bearing on the process of regeneration

(e.g., Brown et al., 1991b; Vargas et al., 2010; Schmid et al., 2013). For example, inhibition of macrophage accumulation in the distal nerve following injury showed significant reductions in myelin clearance and significant delays in functional recovery of hind paw motor control (Barrette et al., 2008). Whether these two effects are causally related is not clear, however, because of the likelihood that in these experiments there were also decreases in macrophage accumulation in the DRG.

While macrophage accumulation in peripheral ganglia following axotomy was described in the 1990s (Lu and Richardson, 1993; Schreiber et al., 1995), their functions have not been clear. Injection of peritoneal macrophages or bacteria (C. parvum) into the

DRG to induce a local inflammatory response resulted in increases in RAG expression and increased regeneration after a dorsal root injury though not after sciatic nerve

90 transection (Lu and Richardson, 1993, 1995). Recent evidence has shown that inhibition of the injury-induced macrophage accumulation in lumbar DRGs results in the abolishment of the conditioning-lesion response both in vitro and in vivo (Kwon et al.,

2013; Niemi et al., 2013).

What remains unclear is how macrophages influence the regenerative capacity of peripheral neurons. Neuroinflammation is often referred to as a “double-edged sword” as illustrated by the divergent role macrophages play in two CNS injury models. Depletion of hematogeneous macrophages leads to improved functional recovery after spinal cord injury (Popovich et al., 1999). However, stimulation of an immune response in the eye in conjunction with optic nerve injury significantly increases regeneration of retinal ganglion cell axons, though the involvement of both neutrophils and macrophages has been implicated (Yin et al., 2003; Yin et al., 2006; Kurimoto et al., 2013). These differing effects presumably result from the heterogeneity of macrophage activation following nervous system injury. Upon entry into tissue, monocyte-derived macrophages can take on a spectrum of activation states (for review see Gordon and Taylor, 2005; Martinez and

Gordon, 2014). Representing the polar opposites on this spectrum, macrophage activation states have been described as pro-inflammatory/M1 or anti-inflammatory/M2 (Gordon and Taylor, 2005; Martinez and Gordon, 2014). Work by Kigerl et al. (2009) demonstrated that conditioned media from bone marrow-derived macrophages stimulated to an anti-inflammatory/M2 activation state with IL-4, induced significant increases in neurite outgrowth from DRG neurons on both growth-permissive and growth-inhibiting substrates. The identity of the factor(s) released from these M2 macrophages to stimulate axonal regeneration remains unknown. Given the increases in regeneration observed as a

91 result of CCL2 overexpression, we hypothesized that the macrophages in our system are anti-inflammatory in nature. The upregulation of CD206 and Fizz 1 mRNA, two anti- inflammatory macrophage markers, and the lack of change in three pro-inflammatory markers are consistent with this hypothesis. CD206, a mannose receptor, is a useful marker of anti-inflammatory/M2 polarization as pro-inflammatory/M1 macrophages do not express this receptor (Alan et al., 1984). Fizz-1, a cysteine-rich secreted protein, has been implicated in the regulation of extracellular matrix and in wound healing (Holcomb et al., 2000), and is specifically expressed by macrophages in response to IL-4 both in vivo and in vitro (Raes et al., 2002).

Activation of a regeneration-associated signaling pathway

In a search for the mechanism by which CCL2 overexpression leads to a conditioning-like increase in neurite outgrowth, we assayed the expression of various

RAGs, expecting to observe an increase in a number of mRNAs. Surprisingly of the eight

RAGs we examined, we found that only LIF was significantly increased 3 week after intrathecal delivery of AAV5-CCL2. In addition, the main signaling mechanism by which LIF acts, namely the phosphorylation of STAT3, increased as a result of CCL2 overexpression. This specificity in CCL2 overexpression-mediated RAG expression suggests that LIF and STAT3 may be responsible for the increase in axonal regeneration.

LIF has been implicated previously in conditioning lesion-induced sensory and sympathetic neuron regeneration (Cafferty et al., 2001; Hyatt Sachs et al., 2010).

Following sciatic nerve injury, LIF is expressed by Schwann cells in the nerve and is retrogradely transported to the DRG (Banner and Patterson, 1994; Curtis et al., 1994;

Thompson et al., 1997). In addition, LIF expression increases in non-neuronal cells in the axotomized ganglia (i.e., superior cervical ganglion), probably in satellite cells (Banner and Patterson, 1994; Sun et al., 1994). In our overexpression system, the cell expressing

LIF has yet to be identified. While the 4-fold increase in LIF mRNA seen in DRGs following CCL2 overexpression is impressive, LIF expression in ganglia following axotomy reaches increases of >100-fold (Banner and Patterson, 1994; Sun et al., 1996;

Habecker et al., 2009).

Axotomy induces the expression of both IL-6 and LIF in peripheral ganglia (Sun et al., 1994; Cafferty et al., 2001; Cafferty et al., 2004; Habecker et al., 2009). Thus, it was unexpected to see only LIF, and not also IL-6, mRNA increase with CCL2 overexpression. We also assayed the expression of two known gp130 cytokine/STAT3- dependent genes, galanin and GAP-43 (Rao et al., 1993; Qiu et al., 2005; Zigmond, 2012;

Ogai et al., 2014) and found that their expression remained unchanged following CCL2 overexpression. Although initially surprising, this may indicate that LIF-induced gene expression requires more than a 4-fold increase in the cytokine. Another common RAG,

ATF3, also remained unchanged following CCL2 overexpression. While ATF3 has been shown to play an important role in axon regeneration (Seijffers et al., 2006; Seijffers et al., 2007), its expression is not known to be gp130-dependent (Habecker et al., 2009).

Activation of the gp130 receptor subunit by LIF and other IL-6 family cytokines results in the activation of JAK2 and the phosphorylation of STAT3 (Heinrich et al.,

1998). STAT3 activation has been shown to play a major role in axon regeneration following injury. STAT3 is activated in DRG neurons after a sciatic nerve injury, but not after a dorsal column lesion (Qiu et al., 2005). Inhibition of STAT3 phosphorylation,

93 through administration of the JAK inhibitor AG490, results in significantly diminished

DRG axonal regeneration in vitro after a conditioning lesion (Liu and Snider, 2001), as well as significantly reduced conditioning lesion-dependent outgrowth after a dorsal column lesion (Qiu et al., 2005). Utilizing in vivo imaging and selective deletion of

STAT3 in DRG neurons, STAT3 was shown to play a role in the initiation of axonal regeneration following a sciatic nerve injury (Bareyre et al., 2011). Our finding that

CCL2 overexpression-induced increases in neurite outgrowth are abolished through the application of STAT3 phosphorylation inhibitors aligns with this previous work. Thus, the conditioning-like increase in neurite outgrowth caused by CCL2 overexpression occurs via a STAT3-dependent mechanism.

CCL2 action in the ganglia

CCL2 is upregulated and expressed by neurons in peripheral ganglia following nerve injury (Schreiber et al., 2001; Tanaka et al., 2004; Niemi et al., 2013). This upregulation and its action through the chemokine receptor, CCR2, are necessary for the injury-induced increase in macrophage accumulation in the DRG (Niemi et al., 2013).

CCL2 primarily acts as a monocyte chemokine by inducing CCR2+ monocyte extravasation into tissue (Geissmann et al., 2003). Yet, CCL2 has also been shown to contribute to the development and maintenance of pain by acting directly on sensory neurons (Abbadie et al., 2009), and it has been demonstrated that CCL2 can depolarize

DRG neurons, following pain-causing injuries, through direct action on the neurons

(White et al., 2005b). While CCL2 has also been shown to cause neurite outgrowth from embryonic avian statoacoustic ganglia, in vitro (Bianchi et al., 2005), in our preparation,

CCL2 does not act directly on neurons to stimulate an increase in outgrowth.

Previous studies have also suggested that CCL2 can stimulate IL-4 production, a cytokine implicated in anti-inflammatory macrophage activation (Karpus et al., 1997).

Evaluation of the general immune responses mounted in a CCL2 -/- mouse revealed decreased anti-inflammatory cytokine expression in response to bacterial immunization

(Gu et al., 2000). Furthermore, macrophages stimulated with M-CSF in vitro in the presence of a CCL2 blocking antibody show significant increases in M1 or pro- inflammatory marker expression, while the presence of CCL2 induced expression of the anti-inflammatory cytokine IL-10 (Sierra-Filardi et al., 2014). It is interesting to note that we have observed increased protein expression of M-CSF in peripheral ganglia following nerve injury (Niemi and Zigmond, unpublished observations). These data suggest that

CCL2 can lead to an anti-inflammatory stimulation of macrophages in addition to being chemotactic to CCR2-positive monocytes. This would favor a macrophage activation state which is capable of promoting axonal growth.

Conclusions

In conclusion, CCL2 overexpression induces macrophage accumulation in uninjured lumbar DRG and leads to a conditioning-like increase in axonal outgrowth.

CCL2 overexpression leads to activation of STAT3, a regeneration-associated signaling molecule. CCL2 must act through its primary receptor, CCR2, to elicit subsequent changes in macrophage accumulation and neurite outgrowth. Finally, the conditioning- like increase in neurite outgrowth induced by CCL2 overexpression is abolished through inhibition of STAT3 phosphorylation. Taken together, CCL2 overexpression in DRG neurons is sufficient to increase the regenerative capacity of DRG neurons through a

STAT3-dependent mechanism.

Figure 3.1

Figure 3.1. Intrathecal injection of AAV5 infects lumbar DRG neurons and leads to a time-dependent overexpression of CCL2. Schematic of AAV5-EF1α-CCL2 vector. We used AAV5-EF1α-eYFP as a control (A). Representative images of AAV5-YFP localization in L4 (B) and L5 DRG (C) at 10x, visualized by anti-YFP labeling. High magnification image of YFP labeling in L5 DRG at 25x (C’). AAV5-YFP injection led to a 37.2% ± 5.7% DRG neuron infection efficiency. Representative image of sparse YFP axonal labeling in the sciatic nerve (D). A representative image of YFP localization in the lumbar spinal cord at low magnification (E) and high magnification (E’) shows preferential expression in the dorsal horn. Injection of AAV5-CCL2 resulted in a time- dependent overexpression of CCL2 mRNA in L5 DRG quantified using the ΔΔCt method normalized to GAPDH (F). For images: Scale bar=100 µm. For RT-PCR: n=5 per group. *p < 0.05. **p < 0.001.

Figure 3.2

Figure 3.2. CCL2 overexpression causes a time-dependent increase in CD11b- and

CD68-positive macrophage accumulation in L5 DRGs. Immunohistochemical labeling of

DRG sections with an antibody against CD11b (A) or CD68 (F) shows significant increases at 2, 3, and 4 weeks after injection of AAV5-CCL2 compared to AAV5-YFP

(A,F). Data were quantified as a percentage of the tissue area positively immuno-labeled.

Representative images of CD11b staining in L5 DRG 1, 2, 3, and 4 weeks after intrathecal injection of AAV5-CCL2 (B-E). Representative images of CD68 staining in

L5 DRG 1, 2, 3, and 4 weeks after intrathecal injection of AAV5-CCL2 (G-J). Inset images taken at 63x. Scale bar=100 µm. n=5 per group. *p < 0.05. **p < 0.001.

Figure 3.3

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Figure 3.3. A conditioning-like increase in neurite outgrowth from DRG neurons is observed with in vivo CCL2 overexpression but not after exogenous CCL2 addition at the time of culturing. Quantification of neurite outgrowth in explant culture measured after

24 and 48 h, 3 weeks after intrathecal injection of AAV5-CCL2 or AAV5-YFP (A). Data is represented as the mean length, in microns, of the 20 longest neurites. Representative images of neurite outgrowth after 48 h in explant culture, visualized with βIII tubulin, for

AAV5-YFP (B) and AAV5-CCL2 (C). Quantification of neurite outgrowth in dissociated cell culture measured at 24 h, 3 weeks after intrathecal injection of AAV5-CCL2 or

AAV5-YFP (D). Representative images of neurite outgrowth in dissociated cell culture at

24 h for AAV5-YFP (E) and AAV5-CCL2 (F). Quantification of neurite outgrowth in

WT DRG explant culture measured after 24 and 48 h, with or without exogenous CCL2

(200 ng/mL) added to the culture medium at the time of plating (G). Representative images of neurite outgrowth after 48 h in culture for control (H) and exogenous CCL2 (I).

Quantification of neurite outgrowth in WT DRG dissociated cell culture at 24 h, with or without exogenous CCL2 (200 ng/mL) added to the culture medium at the time of plating

(J). Representative images of neurite outgrowth in dissociated cell culture for control (K) and exogenous CCL2 (L). Scale bar=100 µm. For CCL2 overexpression explant cultures: n=4 per group. *p < 0.05. For CCL2 overexpression dissociated cell cultures: n=180-251 neurons per group. **p < 0.001. For exogenous CCL2 explant cultures: n=6 per group.

For exogenous CCL2 dissociated cell cultures: n=253-397 neurons per group.

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Figure 3.4

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Figure 3.4. Two anti-inflammatory macrophage markers and no pro-inflammatory markers are significantly increased in uninjured L5 DRGs 3 weeks after intrathecal injection of AAV5-CCL2. Relative expression of mRNA for 4 anti-inflammatory

[arginase 1, CD206, Fizz1, and Ym1] and 3 pro-inflammatory [iNos, CD86, CD16] macrophage markers in L5 DRGs 3 weeks after injection of AAV5-CCL2 or AAV5-YFP

(A) or 7 d after SNI in WT mice compared to sham controls (B). Data was quantified using the ΔΔCt method normalized to GAPDH. n=5 per group. *p < 0.05. **p < 0.001.

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Figure 3.5

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Figure 3.5. Overexpression of CCL2 in CCR2 -/- mice does not cause macrophage accumulation or increased neurite outgrowth in L5 DRGs. Expression of CCL2 mRNA measured 3 weeks after intrathecal injection of AAV5-CCL2 or AAV5-YFP in CCR2 -/- mice, quantified using the ΔΔCt method normalizing to GAPDH (A).

Immunohistochemical labeling of DRG sections with an antibody against CD11b shows no difference between AAV5-CCL2 and AAV5-YFP 3 weeks after injection (B). Data was quantified as a percentage of the imaged area positively immuno-labeled.

Quantification of neurite outgrowth in explant culture measured at 24 and 48 h, 3 weeks after intrathecal injection of AAV5-CCL2 or AAV5-YFP in CCR2 -/- mice (C). Data are represented as the mean length, in microns of the 20 longest neurites. For RT-PCR: n=5 per group. **p < 0.001. For IHC: n=5 per group. For explant cultures: n=5 per group.

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Figure 3.6

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Figure 3.6. LIF mRNA and pSTAT3 are significantly increased in uninjured L5 DRGs 3 weeks after intrathecal injection of AAV5-CCL2. Relative expression of various RAGs in L5 DRGs 3 weeks after intrathecal virus infection of uninjured DRGs quantified using the ΔΔCt method normalizing to GAPDH (A). Relative expression of various RAGs in

L5 DRGs 7 d after SNI in WT mice compared to sham controls (B). Representative images of pSTAT3 staining in L5 DRGs 3 weeks after injection of AAV5-YFP (C),

AAV5-CCL2 with inset image taken at 63x (D), AAV5-CCL2 in CCR2 -/- mice (E), or 7 d after SNI in WT mice (F). Quantification of pSTAT3 staining represented as the percentage of the imaged area positively immuno-labeled (G). Representative images of

ATF3 staining in L5 DRGs 3 weeks after injection of AAV5-YFP (H), AAV5-CCL2 (I),

AAV5-CCL2 in CCR2 -/- mice (J), or 7 d after SNI in WT mice (K). Quantification of

ATF3 staining (L). Representative images of SPRR1a staining in L5 DRGs 3 weeks after injection of AAV5-YFP (M), AAV5-CCL2 (N), AAV5-CCL2 in CCR2 -/- mice (O), or 7 d after SNI in WT mice (P). Quantification of SPRR1a staining (Q).For RT-PCR: *p <

0.05. **p < 0.001. n=5 per group. For IHC: Scale bar=100 µm. n=8 per group. *p < 0.05.

**p < 0.001.

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Figure 3.7

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Figure 3.7. pSTAT3 is required for the CCL2 overexpression-induced increase in neurite outgrowth. Quantification of neurite outgrowth in DRG dissociated cell culture in the presence of inhibitors of STAT3 phosphorylation, AG490 (50 nM) or STATTIC (10 nM), or vehicle control, measured after 24 h, 3 weeks after intrathecal injection of AAV5-

CCL2 or AAV5-YFP (A). Quantification of pSTAT3 nuclear pixel intensity in dissociated cell culture in the presence of the STAT3 phosphorylation inhibitors, AG490 or STATTIC, or vehicle control, measured after 24 h, 3 weeks after intrathecal injection of AAV5-CCL2 or AAV5-YFP (B). Representative images of neurite outgrowth and pSTAT3 are shown for AAV5-YFP Vehicle (C-E), AAV5-CCL2 Vehicle (F-H), AAV5-

YFP AG490 (I-K), AAV5-CCL2 AG490 (L-N), AAV5-YFP STATTIC (O-Q), and

AAV5-CCL2 STATTIC (R-T). Insets are digitally magnifications of the neuronal cell body. Scale bar =100 µm. For neurite outgrowth: n=51-99 neurons per group. **p <

0.001. For pixel intensity: n=51-99 neurons per group. *p < 0.05.

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Chapter 4: Discussion

110

4.1 CCL2: More than just a Chemokine

CCL2 is defined as a chemokine for its ability to attract monocytes, macrophages, and other immune cells to sites of inflammation or injury (Cochran et al., 1983; Charo et al., 1994; Van Coillie et al., 1999; Bartoli et al., 2001). Our work clearly defines CCL2, and its cognate receptor CCR2 (Kurihara and Bravo, 1996), as the primary chemokine responsible for macrophage accumulation in peripheral ganglia and nerve after injury.

The absence of an injury-induced accumulation of macrophages in the DRG and sciatic nerve was seen following sciatic nerve transection in CCR2 -/- mice. Further evidence of

CCL2’s chemotactic capabilities was demonstrated when overexpression of CCL2 in uninjured DRG sensory neurons resulted in a time-dependent accumulation of macrophages equivalent to levels seen following injury.

Our overexpression data also suggests that CCL2 may play a role in the activation of macrophages and not just their recruitment. The monocytes recruited to the DRG in our CCL2 overexpression paradigm would likely experience fewer “activating signals” than those recruited in response to peripheral nerve injury. This creates a type of sterile inflammation that is lacking microbial or damage associated signals to drive the activation of macrophages (Chen and Nuñez, 2010). In response to an injury, numerous signals are released by the injured neuronal cell bodies which could play a role in the activation of macrophages accumulating in ganglia. These include the cytokines IL-6,

TNF-α, LIF, IL-1α, and IL-1β, among others (Curtis et al., 1994; Knoblach et al., 1999;

Cafferty et al., 2004; Ohtori et al., 2004; Allan et al., 2005; Zhang et al., 2007; Brazda et al., 2009).

111

Thus, CCL2 could be involved in the activation of macrophages into a pro- regenerative state, in addition to its involvement in recruiting the monocytes to the ganglia. Recent in vitro experiments examined this idea. Conditioned medium taken from macrophage-DRG neuron co-cultures stimulated with cyclic AMP (Kwon et al.) induces significant neurite outgrowth when placed onto naïve DRG dissociated cultures (Kwon et al., 2015). However, when a CCL2 function blocking antibody is added to the neuron- macrophage co-culture the conditioned medium no longer stimulates neurite outgrowth, an effect not seen if CCL2 is blocked only in the conditioned medium (Kwon et al.,

2015). This suggests that CCL2 is necessary for the production of pro-regenerative factors in the macrophage-DRG neuron co-culture, but does not act as a regenerative factor itself. These mirrors our findings which demonstrate that addition of murine CCL2 alone directly to WT DRG dissociated or explant culture does not stimulate an increase in neurite extension and thus acts indirectly to stimulate an increased regenerative response.

Additional confirmation of this fact is demonstrated by the injection of recombinant chemokines into the ganglia. Injection of recombinant CCL2, CCL3 (macrophage inflammatory protein 1-alpha), or CX3CL1 into the L5 DRG all elicited significant macrophage accumulation in the ganglia 1 week after injection, yet only DRGs injected with CCL2 showed an increased regenerative response, measured in DRG neuron dissociated culture (Kwon et al., 2015). An alternative explanation for this finding, is that these three chemokines recruit different populations of macrophages to the ganglia, which release different cytokines. It is known that patrolling monocytes, which express the receptor for CX3CL1 (CX3CR1), do not usually enter tissue in response to injury

(Auffray et al., 2007; Auffray et al., 2009; Thomas et al., 2015).

112

CCL2 is expressed by a wide array of cell types, but the main source of CCL2 after injury is Schwann cells and neurons (Schreiber et al., 2001; Chazaud et al., 2003;

Mahad and Ransohoff, 2003; Tanaka et al., 2004; Dewald et al., 2005; Napoli et al.,

2012). Besides its chemotactic effects on monocytes and other inflammatory cells it can promote the expression of regulators of macrophage activation and pro-inflammatory cytokine production through direct and indirect alteration of RNA regulatory elements

(Liang et al., 2008; Matsushita et al., 2009; Suzuki et al., 2011). CCL2 can act as a primary activator of Th2 responses and is thought to be a major contributing factor in asthma, rheumatoid arthritis, neurologic disorders such as multiple sclerosis, and type-II diabetes (Gu et al., 1999; Gu et al., 2000; Deshmane et al., 2009). CCL2 may be most studied in cancer, where it has been shown that tumor-derived CCL2 expression positively correlates with tumor-associated macrophage infiltration, tumor vascularization, angiogenesis, and anti-inflammatory macrophage activation/polarization

(O'Hayre et al., 2008; Melgarejo et al., 2009). Along this line, CCL2 expression shifts human peripheral blood CD11b-expressing mononuclear cells toward a CD206-positive

M2-polarized phenotype in culture (Roca et al., 2009). These data are supported by the fact that CCL2 or CCR2 deficient mice show higher dendritic cell and macrophage expression of TNF-a, but lower IL-10 expression than their WT counterparts (Chen et al.,

2011; van Zoelen et al., 2011). The mannose receptor, CD206, was initially identified as part of a protective mechanism against tissue damage during inflammation and was found to act by downregulating lysosomal hydrolases, both in vitro and in vivo (East and Isacke,

2002; Taylor et al., 2005). Mannose receptor activity is increased in cultured bone

113 marrow-derived macrophages in response to IL-4 and IL-13 and is inhibited by IFNγ

(East and Isacke, 2002; Gordon, 2003; Gordon and Martinez, 2010).

In our overexpression paradigm, mRNA for the macrophage activation markers,

CD206 and Fizz1, associated with an M2 phenotype are significantly upregulated in the

L5 DRG 3 weeks after CCL2 overexpression, whereas CD86, iNos, and CD16, markers of M1 activation, do not differ from YFP controls (Holcomb et al., 2000; Raes et al.,

2002; Gordon, 2003; Gordon and Martinez, 2010; Martinez and Gordon, 2014). We showed that the CCL2 overexpression-induced macrophage activation marker expression profile was similar to the mRNA expression of these markers in the DRG 7 days after sciatic nerve injury, where the anti-inflammatory macrophage markers arginase 1,

CD206, and Ym1 were all increased. This suggests that CCL2 may play a major role in facilitating this M2 macrophage activation profile, as CCL2 overexpression alone causes a profile very similar to that seen in the more complex environment created after injury.

In vitro studies using human blood monocytes illustrate the important role CCL2 plays in anti-inflammatory macrophage activation. Human monocytes stimulated with M-

CSF were found to take on an anti-inflammatory activation state and express high levels of CCL2 (Sierra-Filardi et al., 2014). Furthermore, it was shown that CCL2 enhanced the expression of IL-10 and repressed the expression of inflammatory cytokines, such as IL-6 in these macrophages, as use of a CCL2 function blocking antibody in the culture inhibited M2 macrophage activation (Sierra-Filardi et al., 2014). Additional work in human CD11b-expressing mononuclear cells revealed that culturing monocytes with

CCL2 significantly increased survival of the cells and resulted in an anti-inflammatory polarization of the cells (Roca et al., 2009). CCL2 was shown to significantly increase the

114 expression of CD206 in monocytes while simultaneously reducing the mRNA and protein expression of the pro-inflammatory cytokine, IL-6 (Roca et al., 2009).

Anti-inflammatory activation of macrophages has been shown to be pro- regenerative, while M1 or pro-inflammatory activation leads to a decreased axonal regeneration of axons in cultured CNS and PNS neurons. Kigerl et al. (2009) found that conditioned medium from IL-4 stimulated bone marrow-derived macrophages significantly increased the ability of DRG and cortical neurons to grow axons on inhibitory substrates. Conditioned medium from macrophages, stimulated with IFNγ, did not increase the regenerative response of neurons, but rather decreased the ability of axons to cross an inhibitory CSPG spot compared to control DRG neurons (Kigerl et al.,

2009). Our conclusion is that CCL2 overexpression leads to not only macrophage accumulation, but also a pro-regenerative activation of macrophages which results in a conditioning-like increase in neurite outgrowth. This is supported by our overexpression data in CCR2 -/- mice, where we are able to successfully overexpress CCL2 in L5 DRG neurons but do not see macrophage accumulation or increased axonal regeneration.

4.2 Macrophages and Regeneration: New Site of Action

Over the past three decades it has been demonstrated that exogenous inflammatory activation near injured neuronal cell bodies could increase the regenerative capacity of both peripheral and central nervous system neurons (Lu and Richardson, 1991; Hikawa et al., 1993; Hikawa and Takenaka, 1996; Steinmetz et al., 2005; Benowitz and Yin, 2008;

Kigerl et al., 2009; Benowitz and Yin, 2010; Benowitz and Popovich, 2011). Injection of peritoneal macrophages or stimulation of a local inflammatory response through the injection of zymosan or bacteria into the DRG was shown to increase axonal regeneration

115 following a crush injury of the dorsal root (Lu and Richardson, 1991; Steinmetz et al.,

2005). Stimulation of optic nerve regeneration was also seen when lens injury or zymosan injection into the eye was utilized to create a local inflammatory reaction near injured retinal ganglion cells (Leon et al., 2000; Fischer et al., 2001; Yin et al., 2003; Yin et al., 2006; Leibinger et al., 2009; Yin et al., 2009; Hauk et al., 2010). Yet, the role of the endogenous immune response and macrophage accumulation in peripheral ganglia following nerve injury remained unknown.

Previous work has focused on the actions of macrophages and other immune cells in the nerve after injury. In this location, at the site of injury and distal to it, macrophages, neutrophils, and B cells have all been shown to play a critical role in the ability of sensory axons to regenerate and allow for functional recovery (Anthony et al., 2007;

Morin et al., 2007; Barrette et al., 2008; Vargas et al., 2010; Wu et al., 2012; Cattin et al.,

2015). Macrophages are the prominent immune cell present in the sciatic nerve following crush or transection injury, where they remove axonal and myelin debris as well as release growth factors to support the regrowth of axons (Perry et al., 1987; Brown et al.,

1991b; Dahlin, 1995; Liu et al., 2000; Abbadie et al., 2003; Luk et al., 2003; Barrette et al., 2008). A similar influx and accumulation of hematogeneous macrophages occurs in peripheral ganglia after nerve injury, however, the endogenous immune response in the cell body compartment and its role in regeneration has not been studied as thoroughly (Lu and Richardson, 1993; Schreiber et al., 1995; Magnusson and Kanje, 1998).

The endogenous macrophage response to injury in peripheral ganglia has many characteristics that are suggestive of its possible role in promoting regeneration of peripheral axons. Macrophages only accumulate in peripheral ganglia following an injury

116 that result in regeneration. Injury to the central branch of the DRG, either by injuring the dorsal root or through a lesion of the dorsal column of the spinal cord, do not result in macrophage accumulation within the DRG (Perry et al., 1987; Kwon et al., 2013; Ton et al., 2013). This follows a similar profile to the neuronal expression of RAGs which are only upregulated after peripheral but not central axonal injury, a defining characteristic for being associated with regeneration (Smith and Skene, 1997; Schwaiger et al., 2000;

Bulsara et al., 2002; Costigan et al., 2002; Schmitt et al., 2003; Boeshore et al., 2004).

Furthermore, Lu and Richardson (1995) demonstrated that inducing a local inflammatory response dominated by macrophage accumulation in the DRG, through injection of c. parvum, increased neuronal mRNA expression of the RAGs GAP-43, cJun, and CGRP.

These data suggest that macrophages at the site of the ganglia could directly contribute to the gene expression changes necessary for peripheral regeneration.

Our data was the first definitive proof that the endogenous macrophage response to injury in peripheral ganglia significantly contributes to axonal regeneration. Through the use of two different mouse models, the Wlds and CCR2 -/- mice, we demonstrated that if hematogeneous macrophages accumulation is diminished in the DRG or SCG compared to WT mice, following peripheral nerve injury, than the conditioning lesion response is significantly impaired. The conditioning lesion response allows for a direct measurement of the growth capacity of neurons (Oblinger and Lasek, 1984; Shoemaker et al., 2005). The first conditioning injury, performed in vivo, allows injury induced signaling, changes in gene expression in injured neurons, and non-neuronal cell changes, including macrophage accumulation within the ganglia, to occur. The test lesion, occurring seven days later, results in a 5- to 10-fold increase in regeneration, measured by

117 axonal length, if and only if the proper signaling and gene expression changes have taken place in the injured neurons and their immediate environment. The impairment of the conditioning lesion response in the CCR2 -/- mouse was directly proportional to the impairment of macrophage accumulation in the ganglia. Macrophage accumulation in the

DRG, seven days after sciatic nerve injury, was completely inhibited in CCR2 -/- mice in comparison to WT mice. Macrophage accumulation in the SCG, however, was reduced by only 50% in the CCR2 -/- mouse, seven days after injury. The reduction seen in the conditioning lesion responses of these two ganglia, measured in explant and dissociated cell culture, directly related to the reduction of macrophage accumulation, where the conditioning lesion response in the DRG of CCR2 -/- mice was completely inhibited and the SCG showed a significant reduction but not complete loss of the conditioning lesion response. This idea was further supported by Kwon et al. (2013) who showed that application of the antibiotic minocycline to the DRG, to “deactivate” macrophages in the ganglia, resulted in significant impairment of macrophage accumulation in the DRG and the conditioning lesion response.

Previous work looking at the contribution of macrophages in the sciatic nerve to regeneration has used various strategies to inhibit their entry and accumulation within the nerve. These have ranged from the expression of thymidine kinase in CD11b expressing cells and ganciclovir administration to kill mononuclear cells, to the use of clodronate liposomes which are toxic to phagocytic cell populations (Liu et al., 2000; Barrette et al.,

2008). In these paradigms, it was shown that systemic elimination of monocytes and macrophages prevented macrophage accumulation within the sciatic nerve and resulted in delayed or decreased regeneration and functional recovery. From this, it was concluded

118 that the lack of macrophages in the nerve was directly responsible for this delayed regeneration, likely as a result of slowed or incomplete clearance of axonal and myelin debris (Brown et al., 1991b; Brown et al., 1992; Liu et al., 2000; Barrette et al., 2008).

However, these papers never addressed the likely effect that systemic ablation of macrophages would have on macrophage accumulation within peripheral ganglia. Given the work from our lab that defines a necessary role for macrophages in the ganglia, the delayed and incomplete regeneration exhibited in these previous studies may be a result of inhibiting macrophage action in both the ganglia and at the site of injury.

4.3 Pro-Regenerative Macrophage Mechanisms

In addition to illustrating a novel sight of action for macrophages following peripheral nerve injury, our work highlights a potential mechanism by which macrophage accumulation near injured neuronal cell bodies could facilitate increased axonal regeneration. While macrophage stimulation of axonal regeneration has been demonstrated in many ways, the mechanisms and molecules/cytokines involved remain elusive (Lu and Richardson, 1991, 1993, 1995; Kigerl et al., 2009; Alexander et al., 2012;

Gensel et al., 2012). CCL2 overexpression resulted in a conditioning-like increase in neurite outgrowth that is dependent upon the macrophage accumulation within the uninjured DRG. When we measured the mRNA expression of a small subset of well- defined RAGs including cytokines, transcription factors, growth-associated proteins, and neuropeptides, in DRG 3 week after CCL2 overexpression we expected to see multiple genes upregulated. What we found was a specific upregulation of the cytokine, LIF, and no change in the expression levels of the other genes. Although we isolated mRNA from the entire DRG and cannot identify the cell(s) expressing LIF, a recent paper used

119 magnetic cell separation to identify the expression of LIF and IL-6 in macrophages after sciatic nerve injury (Kwon et al., 2013). Interestingly, it was previously shown that LIF was primarily expressed in satellite glial cells in the SCG and DRG following nerve injury (Banner and Patterson, 1994; Sun et al., 1994) while IL-6 can be expressed by injured neurons in the DRG (Murphy et al., 1999). Thus, it is unlikely that macrophages are the sole source of these cytokines in the ganglia. A member of the IL-6 cytokine family, LIF acts through a heteromeric receptor made up of the signaling subunit, gp130, and the ligand binding subunit, LIF receptor which dimerize to form a signaling receptor unit (Slaets et al., 2008; Zigmond, 2012). Receptor activation leads to the activation of

STAT3, through phosphorylation, dimerization, and nuclear translocation (Kishimoto,

1994; Aaronson and Horvath, 2002). We also found that phosphorylated STAT3

(pSTAT3) was significantly expressed in neurons following CCL2 overexpression.

STAT3 activation plays a critical role in the initiation phases of regeneration and has been identified as a core transcription factor in peripheral nerve regeneration (Bareyre et al., 2011; Chandran et al., 2016). To ascertain if STAT3 activation played a critical role in the CCL2 overexpression-induced increase in neurite outgrowth we used pharmacological inhibition of STAT3 phosphorylation in dissociated cell culture. We found that STATTIC, a STAT3 phosphorylation inhibitor, completely abolished the increased neurite outgrowth seen after CCL2 overexpression. This suggests that the

LIF/STAT3 pathway may be critically responsible for macrophage induced increases in axon regeneration following peripheral nerve injury.

LIF has many other qualities that make it likely to be involved in the regeneration-promoting effects in our system. LIF is transported retrogradely by sensory

120 and sympathetic neurons; this transport is increased after nerve lesions (Bachoo et al.,

1992; Hendry, 1992b; Hendry, 1992a; Hendry et al., 1992; Curtis et al., 1994; Sun et al.,

1996; Sun and Zigmond, 1996b; Zigmond, 1997). LIF is absent from the adult mammalian nervous system but it is upregulated after injury to the sciatic nerve and axotomy of the SCG (Curtis et al., 1994; Sun et al., 1996; Sun and Zigmond, 1996b;

Habecker et al., 2009; Hyatt Sachs et al., 2010). LIF has been shown to regulate the expression of the neuropeptides in axotomized SCG and has been shown to be absolutely necessary for the conditioning lesion response in this ganglia (Habecker et al., 2009;

Hyatt Sachs et al., 2010).Another way LIF is involved in regeneration is through its role in the transected sciatic nerve by improving the conduction velocity of the regenerated nerve and the number of myelinated fibers (Tham et al., 1997). LIF is also involved in cell viability and supporting survival of sensory neurons (Murphy et al., 1991), possibly via direct mechanisms, since LIF binds specifically to DRG neurons in vitro (Hendry et al., 1992). Moreover it can also promote survival of sensory and motor neurons after axotomy in vivo (Tham et al., 1997). Exogenous addition of LIF after spinal cord injury has been shown to increase levels of NT-3 at the site of injury and induce sprouting

(Blesch et al., 1999). These findings support the hypothesis that ‘‘anti-inflammatory factors not only directly mediate regeneration, but can also indirectly regulate the nervous system response to injury by increasing the production of trophic factors via the cytokine/neurotrophin axis” (Golz et al., 2006). Using LIF knockout mice, it was shown that LIF plays an important role in the initial infiltration of inflammatory cells after cortical and sciatic nerve injury, acting as a chemotactic factor for macrophages and possible activator of microglia and astrocytes (Sugiura et al., 2000). LIF has also been

121 identified as one of the critical factors mediating lens injury-induced optic nerve regeneration (Leibinger et al., 2009) Thus, it is likely that LIF, alone or in concert with other cytokines, may be the critical macrophage-derived pro-regenerative factor in peripheral ganglia.

Given that anti-inflammatory macrophages have been shown to increase axonal regeneration and that our data indicates that macrophages in the DRG after injury or

CCL2 overexpression may be polarized towards an M2 state, other anti-inflammatory cytokines may also play a role in macrophage stimulated regeneration. IL-4 is a 30 kDa protein that is produced mainly by mature Th2 cells, mast cells, and macrophages

(Gordon, 2003; Gordon and Taylor, 2005; Gordon and Martinez, 2010). It has been shown that after sciatic nerve injury, there is a downregulation of IL-4 mRNA levels in injured DRGs within hours after injury, before slight increases in expression are seen at later time points (Uceyler et al., 2007; Üçeyler et al., 2008) IL-4 has been implicated in peripheral axon regeneration and has been found to promote facial motor neuron survival after axotomy through a mechanism involving STAT6 (Jones et al., 2005; Deboy et al.,

2006). While in vivo evidence for a direct role of IL-4 in axonal regeneration does not exist, work in DRG and retinal cell culture have revealed that IL-4 can directly induce neurite outgrowth in a dose-dependent manner (Volpert et al., 1998; Üçeyler et al., 2008)

IL-10 is the quintessential anti-inflammatory cytokine and is mainly produced by monocytes/macrophages and Th2 cells. It inhibits inflammatory monocytes and macrophages and modulates lymphocyte and neutrophil responses as well as cytokine production (Opal and DePalo, 2000; Moore et al., 2001). Treatment with IL-10 increases both cell survival and axonal regeneration after PNS injury. After facial nerve axotomy,

122 flow cytometry analyses showed that the levels of IL-10 were increased and provided a protective effect from cell death (Xin et al., 2008). IL-10 also exerts a neuroprotective effect on retinal ganglion cells by inhibiting apoptotic cell death through mechanism that involves activation of the STAT3 pathway (Boyd et al., 2003). Similarly, in vivo, IL-10 overexpression in the spinal cord using a herpes simplex virus to express IL-10, in both glia and neurons, or systemic IL-10 administration demonstrated increased neuronal survival and improved motor function after injury (Bethea et al., 1999; Plunkett et al.,

2001; Zhou et al., 2009). While there is little evidence for direct stimulation of axonal regeneration by IL-10, recent data showed that peripheral regeneration was significantly impaired in an IL-10 -/- mouse after sciatic nerve injury (Mietto et al., 2015)

TGF-b is mainly involved in inhibition of monocytes and macrophages and pro- inflammatory cytokine synthesis (Opal and DePalo, 2000). TGF-β has been directly implicated in axonal regeneration. Using DRG explants, administration of recombinant

TGF-β was found to promote neurite outgrowth (Schachtrup et al., 2010). TGF-β’s effect on neurite outgrowth may occur through activation of RhoA signaling, which has been shown to be a critical factor for peripheral nerve regeneration (Sebök et al., 1999; Cheng et al., 2008; Kohta et al., 2009). TGF-β is upregulated at early time points in both the sciatic nerve and the DRG following injury (Nath et al., 1998).

4.4 Translational Aspects

Our data show that CCL2-mediated macrophage activation and accumulation in the DRG can significantly increase axon regeneration. Kwon et al. (2015) demonstrated that CCL2 overexpression in the DRG was just as effective in stimulating regeneration of sensory axons after spinal cord injury as a peripheral preconditioning lesion (Richardson

123 and Issa, 1984). The CCL2 overexpression induced regeneration occurred when AAV5-

CCL2 was injected either 7 days prior to or 1 day after spinal cord injury (Kwon et al.,

2015). These data support studies showing that peripheral conditioning injury enhances regenerative responses whether the conditioning injury procedure is performed before or after the central lesion (Kadoya et al., 2009; Ylera et al., 2009). Our finding that overexpression of CCL2 results in a conditioning-like effect on axon growth suggests that it could be employed as a therapeutic strategy for various CNS injuries. Importantly, macrophage activation in the DRGs by overexpression of CCL2 did not result in general morphological changes or neuronal damage (Kwon et al., 2015), whereas zymosan- induced macrophage accumulation produced cytotoxicity in a previous study (Gensel et al., 2009). Thus, it is likely that the CCL2-induced macrophage accumulation may be more physiologic in nature than the chemically induced process in recapitulating the activation of macrophages and inflammatory reactions that occur after nerve injury. It was also shown that CCL2 overexpression induced no changes in the threshold for pain perception (Kwon et al., 2015). Thus, the therapeutic effects of CCL2 overexpression seem to be unaccompanied by undesirable side effects.

The route of administration used in our study, is therapeutically relevant, as it does not involve direct injection into nervous system tissue (Follett et al., 2004; Smith et al., 2008). Following injection of AAV5-YFP into the intrathecal space of the spinal cord we observed infection of neurons and axons in lumbar DRGs and the dorsal horn of the spinal cord. Considering a previous report showing involvement of CCL2 in regulating macrophage responses to remove myelin debris from damaged axons (Perrin et al., 2005), it is conceivable that CCL2 overexpressed in DRGs by AAV might enhance myelin

124 breakdown at the injury site and thereby further enhance regeneration of injured axons in the spinal cord (Vargas and Barres, 2007). Endogenous upregulation of CCL2 expression in the lesioned spinal cord is usually limited within several days after injury (Ma et al.,

2002; Perrin et al., 2005). Since Wallerian degeneration proceeds at a much slower pace in the CNS (Vargas and Barres, 2007), stable long-term expression of CCL2 in the spinal cord after injury could increase clearance of debris by recruiting and beneficially- activating more macrophages proximal to the site spinal cord injury.

Accurate identification of neuronal gene expression changes that are directly or indirectly triggered by macrophages could provide another option for therapeutic application of our findings. Chandran et al. (2016) recently demonstrated that a systems- level analysis of gene expression changes in axotomized peripheral neurons could be used to identify drug candidates to increase regeneration. After determining a core transcription factor network that is upregulated in response to sciatic nerve injury, the authors used bioinformatics and publicly available databases to identify small molecules that could induce similar gene expression profiles (Lamb et al., 2006). Using a small molecule gene expression database, the drug ambroxol was identified to activate a gene expression profile in a non-neuronal cell line comparable to what is observed in DRG neurons after injury. Ambroxol treatment increased axon regeneration of RGCs both in vitro and in vivo (Chandran et al., 2016). More work needs to be done to identify specific cytokines and genes involved in macrophage-mediated axon regeneration, however, this strategy holds great promise for the identification of novel therapeutic targets and treatments for spinal cord injury.

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4.5 Future Directions

Numerous questions remain to be answered regarding the mechanism by which macrophages can increase the regenerative capacity of peripheral neurons through action at the site of the neuronal cell body. We first want to define the RAGs whose neuronal expression is dependent upon macrophage accumulation within the ganglia. Peripheral axon regeneration is dependent upon a neuronal switch in gene expression which is defined by an upregulation of transcription factors and growth-associated genes and a downregulation of genes associated with neurotransmission (Smith and Skene, 1997).

Given the timeline of macrophage accumulation within the ganglia, where macrophage accumulation begins at 48 hours post-injury and is sustained for at least 2 to 4 weeks (Lu and Richardson, 1993; Schreiber et al., 1995; Kwon et al., 2013), macrophages may be involved in both initiation and maintenance of specific gene expression changes. To answer this we will perform an RNA sequencing time course analysis of WT and CCR2 -

/- DRGs at 0, 12, 24, and 48 hours, and 3, 7, and 14 days after sciatic nerve injury.

Differences in gene expression between WT and CCR2 -/- DRGs, specifically genes that are upregulated in WT and not in CCR2 -/- mice, could help define RAGs whose injury- induced expression is dependent upon macrophage accumulation within the ganglia. This bottom-up approach could ultimately help define the overall contribution the endogenous immune response plays in peripheral regeneration.

Another avenue of interrogation into this mechanism is to define the pro- regenerative cytokine(s) released by macrophages. Defining the cytokine profile present with in the ganglia, in vivo, may be hindered by the inability to accurately track the source of the cytokines. Neurons and satellite glial cells can also express and release

126 cytokines in response to nerve injury (Patterson, 1992; Banner and Patterson, 1994; Sun et al., 1994; Marz et al., 1998). We will instead use an in vitro system to define macrophage-derived pro-regenerative factors. Conditioned medium taken from bone marrow-derived macrophages, stimulated with IL-4 in culture, can induce significant neurite outgrowth from DRG and cortical neurons plated on inhibitory substrates (Kigerl et al., 2009). We will use this same paradigm to specifically identify the factors in the macrophage conditioned medium necessary to stimulate regeneration. Seventy-two hours after treating bone marrow-derived macrophages with IL-4, we will collect the medium and perform a proteomics analysis on it. This will identify all of the soluble factors released by the macrophages. We can then select the most likely candidates based on expression level and literature searches (to determine any molecules that have been previously implicated in regeneration). Systematic depletion of these factors can be done by adding specific function blocking antibodies to the conditioned medium and then testing the ability of the “depleted” medium to induce neurite outgrowth from DRG dissociated cell cultures. We can confirm expression of the identified macrophage- derived pro-regenerative factors in vivo by performing enzyme-linked immunosorbent assays (ELISA) of proteins isolated from the DRG following nerve injury. Identification of the genes regulated by macrophages after injury and the specific pro-regenerative factors necessary for macrophage mediated regeneration, could lead to the discovery of novel therapeutic targets and would greatly increase our overall knowledge of the regulators of peripheral regeneration.

Finally, we want to determine if CCL2 overexpression in CNS neurons could boost their regenerative capabilities in response to injury. Initial testing will be carried

127 out in the eye. Previous research indicated that creating a local inflammatory response around RGCs following optic nerve injury results in increased cell survival and significant optic nerve regeneration (Cui et al., 2009; Benowitz and Yin, 2010; Benowitz and Popovich, 2011). We will overexpress CCL2 in the retina prior to optic nerve injury, through direct eye injection of AAV2-CCL2, specifically designed to infect RGCs (Smith et al., 2009; Sun et al., 2011). Nerve regeneration will be measured in vivo at 14 days following optic nerve crush. Macrophage accumulation within the retina and aqueous humor will also be quantified. Based on our data in the DRG following AAV5-CCL2 administration, we would hypothesize that optic nerve regeneration and macrophage accumulation would significantly increase following CCL2 overexpression. Application of our macrophage studies to CNS injury is an important next step in determining just how powerful and translatable our findings are.

4.5 Concluding Remarks

In conclusion, our results demonstrate that macrophage accumulation in peripheral ganglia is both necessary and sufficient for axonal regeneration. Through the use of two mouse models, CCR2 -/- and Wlds mice, we showed that if hematogeneous macrophage accumulation in peripheral ganglia is inhibited following peripheral nerve injury, then the conditioning lesion response is significantly inhibited. These data define a new site of action where macrophages play a significant role in the ability of peripheral neurons to regrow their axons after injury.

Furthermore, we revealed that inducing macrophage accumulation in uninjured lumbar DRG induced a conditioning-like increase in neurite outgrowth. Viral

128 overexpression of the macrophage chemokine CCL2 resulted in a time-dependent increase macrophage accumulation in the DRG and significantly increased neurite outgrowth in both explant and dissociated cell culture. The increased neurite outgrowth was not seen if recombinant CCL2 was added directly to neurons in culture or when

CCL2 was overexpressed in CCR2 -/- mice where macrophage accumulation was not seen in the ganglia. In addition, we defined a possible mechanism by which macrophages may stimulate an increased regenerative response in sensory neurons. CCL2 overexpression-induced macrophage accumulation in uninjured DRGs resulted in a specific upregulation of the cytokine LIF as well as neuronal activation of STAT3.

Pharmacological blockade of STAT3 activation in culture completely abolished the increased conditioning-like increase in regeneration seen in our CCL2 overexpression paradigm.

This work has revealed a novel mechanism supporting peripheral axon regeneration. Further investigation into the specific proteins and signaling pathways that mediate neuron-macrophage interactions in peripheral nerve injury could result in new therapeutic strategies for CNS injuries.

129

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