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A natural diketopiperazine stimulates axon sprouting and sensory recovery following dorsal rhizotomy Wong, Jennifer Wai Jing 2010

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A NATURAL DIKETOPIPERAZINE STIMULATES AXON SPROUTING AND SENSORY RECOVERY FOLLOWING DORSAL RHIZOTOMY  by JENNIFER WAI JING WONG B.Sc., The University of British Columbia, 2004  A THESIS SUBMITTED IN PARTIAL FULFILLMENT OF THE REQUIREMENTS FOR THE DEGREE OF DOCTOR OF PHILOSOPHY in  THE FACULTY OF MEDICINE (Neuroscience)  THE UNIVERSITY OF BRITISH COLUMBIA (Vancouver)  January 2010  © Jennifer Wai Jing Wong, 2010  ABSTRACT During prenatal and early postnatal development, the mammalian nervous system has the remarkable ability to build its intricate array of connections and circuitries with the help of a variety guidance cues. When the nervous system matures, it appears to lose its ability to rebuild damaged connections following traumatic insults. This can be attributed in part to the expression of inhibitory molecules that hinder axon regeneration and reconnection. The goal of my work was to identify novel compounds that can stimulate axon outgrowth in the unfavourable environment of the adult central nervous system (CNS) by manipulating the axon outgrowth machinery in the neuronal growth cone. To isolate compounds of therapeutic potential, we first developed a novel high-throughput screening technology to rapidly identify candidate neurite outgrowth promoting molecules from a bioactive marine sponge extract library. Using the highthroughput screening technology, we identified a natural diketopiperazine DKP101516 that demonstrated robust axon outgrowth promoting activity through the phosphotidyl-3-inositol kinase (PI3K) signalling pathway. Further in vivo studies revealed that while DKP101516 did not stimulate afferent regeneration, it markedly enhanced intraspinal axon sprouting following dorsal rhizotomy. Lastly, behavioural studies suggest that DKP10516 also promoted rapid and transient recovery in mechanosensation, concomitant to the sprouting of VGLUT1 positive mechanosensory afferents. Collectively, our data suggest that DKP101516 may be a promising therapeutic to stimulate axon repair and functional recovery following injuries in the CNS.  ii  TABLE OF CONTENTS Abstract………………………………………………………………………………….……….ii Table of Contents……………………………………..………………………………………....iii List of Tables……………………………………………...…………………………….…..…...vi List of Figures………………………………...……………..………………………..….……..vii List of Abbreviations………………………..…………………..………………………...…….ix Acknowledgements………………………….………………………………………..…………xi Dedication…………………………………..………………………….………………………..xii Co-Authorship Statement………………...…….………………...…………………………...xiii 1  Introduction……………………………………………….……………………………...1 1.1  Introduction……………………………………………...………………………...2  1.2  Injury to the Nervous System………………………………………………….…. 4  1.3  Axon Repair Strategies….……………………………………………...…………9  1.4  Manipulating the Neuronal Growth Cone………………………………………..17  1.5  Rat Dorsal Rhizotomy- Model for Regeneration………………………………...30  1.6  Experimental Objectives.……………………………………………………….. 40  1.7  Bibliography……………………………………………………………………..42  2  Development of a High-throughput Screen…..……………………………………… 60 2.1  Introduction……………………………………………………………………....61  2.2  Methodology……………………………………………………………………..63  2.3  Results…………………………………………………………………………... 67  2.4  Discussion………………………………………………………………………..77  2.5  Bibliography……………………………………………………………………..85  iii  3  DKP101516 Stimulates Axon Sprouting in the Spinal Cord…………………...…… 87 3.1  Introduction……………………………………………………………………... 88  3.2  Methodology……………………………………………………………………. 89  3.3  Results…………………………………………………………………………... 98  3.4  Discussion………………………………………………………………………122  3.5  Bibliography……………………………………………………………………128  4  Transient Effects of DKP101516 on Axon Sprouting and Sensory Function..……131 4.1  Introduction…………………………………………………………………….132  4.2  Methodology……………………………………………………………………135  4.3  Results…………………………………………………………………………..139  4.4  Discussion………………………………………………………………………162  4.5  Bibliography…………………………………………………………………… 178  5  Conclusion and Discussion …………….…………...………….…………….……… 182 5.1  Summary of Results…………………………………………………………….182  5.2  Discussion……………………………………………………………………....183 5.2.1  Effects of DKP101516 on the Sensory Circuitry…………………………...183  5.2.2  Non-neuronal Effects of DKP101516……………………………………....185  5.2.3  DKP101516 Mechanism of Action………………………………………....186  5.2.4  Therapeutic Applications- Promise and Problems………………………….188  5.3  Future Directions………………………………………………………………. 190 5.3.1  Elucidating the Mechanism of DKP101516 Mediated Microglia Reactivity………………………………………………………………….. .190  5.3.2  Minimizing the Detrimental Effects of DKP101516…………...…………..191 iv  5.3.3  Improvements on the High-Throughput Screen…………………………… 193  5.4  Conclusion……………………………………………………………………... 194  5.5  Bibliography…………………………………………………………………… 196  v  LIST OF TABLES Table 2.1.  Bioactive extracts detected from the high-throughput wound-healing assay…..71  vi  LIST OF FIGURES Figure 1.1  Sensory Circuitry………………………………………………………………...32  Figure 2.1.  High-throughput cell migration assay……………………………………………69  Figure 2.2  Candidate extracts promote neurite outgrowth in DRG explants………………..73  Figure 2.3  Candidate extracts stimulate neurite outgrowth on inhibitory substrates………..75  Figure 2.4  Purification of neurite outgrowth promoting compound DKP101516 (JH)……..78  Figure 2.5  DKP10516 (JH) demonstrates neurite outgrowth effects on various substrates... 80  Figure 3.1  DKP101516 stimulates neurite outgrowth and branching in a dose-dependent manner……………………………………………………………………………90  Figure 3.2  DKP101516 overcomes the inhibitory effects of myelin and CSPG …………..100  Figure 3.3  DKP101516 mediates its effects through the PI3K pathway…………………...103  Figure 3.4  DKP101516 stimulates afferent sprouting within the peripheral root…………. 106  Figure 3.5  DKP101516 enhances astrogliosis and neurocan expression in the DREZ…….109  Figure 3.6.  DKP101516 mediated neurite outgrowth is significantly reduced by high concentrations of neurocan………………………………………………….….112  Figure 3.7  DKP101516 stimulates the sprouting of intact CGRP positive afferents………115  Figure 3.8  DKP101516 stimulates the sprouting of SERT positive projections…………...118  Figure 3.9  DKP101516 stimulates the sprouting of TH positive afferents………………...120  Figure 4.1  Axonal projections involved in spontaneous sensory changes following C7/C8  DRI……………………………………………………………………………………………..141 Figure 4.2  DKP101516 accelerates mechanosensory recovery following C7/8 DRI…...…143  Figure 4.3  DKP101516 accelerates mechanosensory recovery following C7/8 DRI……...145  Figure 4.4  DKP101516 does not affect pain response to radiant heat and pressure following C7/8 DRI………………………………………………………………………..148  vii  Figure 4.5  DKP101516 stimulates transient sprouting of mechanosensory afferents (VGLUT-1)……………………………………………………………………..150  Figure 4.6  DKP101516 stimulates transient sprouting of CGRP positive nociceptive afferents……………………………………………………………154  Figure 4.7  DKP101516 stimulates transient sprouting of descending supraspinal projections………………………………………………………….157  Figure 4.8  DKP101516 stimulates transient sprouting of inhibitory interneurons………...160  Figure 4.9  DKP101516 mediates negative glial effects in the long-term………………….164  Figure 4.10  DKP101516 promotes the delayed expansion of CD11b positive cells in the cuneate fasciculus…………………………………………………..166  Figure 4.11  Summary of the effects of DKP101516 treatment following C7/C8 DRI ……..171  Figure 4.12  Timeline illustrating the effects of DKP101516 from 0-23 days post DRI…….173  viii  LIST OF ABBREVIATIONS AKAP APC Arp2/3 BDNF cAMP CGRP CLIP170 CNP CNS CRMP2 CSPG CST CTB DAG DKP DMEM DMSO DREZ DRG DRI EGFR Erk GFAP GSK3β HEK293 FBS IGF IL-6 IP3 IQGAP MAG MAIP MAP1b MAPK MLCK NCAM NF200 NGF NPY NT-3 N-WASP PAK PDL PI3K PI3Ki PIP2  A-kinase anchoring Protein adenomatous polyposis coli actin related protein 2/3 brain derived neurotrophic factor cyclic adenosine monophosphate calcitonin gene related protein cytoplasmic linker protein 170 2',3'-cyclic nucleotide 3'-phosphodiesterase central nervous system collapsin-response mediator protein 2 chondroitin sulfate proteoglycan corticospinal tract B fragment of cholera toxin diacylglycerol diketopiperazine Dulbecco’s minimal essential medium dimethylsulfate oxide dorsal root entry zone dorsal root ganglion dorsal root injury epithelial growth factor receptor extracellular related kinase glial fibrillary acid protein glycogen synthase kinase-3-beta human embryonic kidney cells fetal bovine serum insulin like growth factor interleukin-6 inositol-3-phosphate IQ motif containing GTPase activating protein myelin associated glycoprotein myelin associated inhibitory proteins microtubule associated protein-1 mitogen activated protein kinase myosin light chain kinase neural cell adhesion molecule neurofilament 200 nerve growth factor neuropeptide Y neurotrophin 3 neuronal Wiscott-Aldrich Syndrome protein p21 activated kinase poly-D-lysine phosphotidyl-3-inositol kinase phosphotidyl-3-inositol kinase inhibitor phosphatidyl-3,4-inositol-bisphosphate ix  PIP3 PKA PKB PKC PLCγ PLL PLP PNS PP1 ROCK SCAR SCI SNgR TGFβ1 TNF VEGF VGAT VGCC VGLUT-1 WAVE  phosphatidyl-3,4,5-inositol-triphosphate protein kinase A protein kinase B protein kinase C phospholipase C gamma poly-L-lysine proteolipid protein peripheral nervous system protein phosphatase 1 Rho kinase suppressor of cAMP receptor mutation spinal cord injury soluble nogo receptor transforming growth factor beta 1 tumor necrosis factor vascular endothelial growth factor vesicular GABA-ergic transporter-1 voltage gated calcium channels vesicular glutaminergic transporter-1 WASP family verprolin-homologous protein  x  ACKNOWLEDGEMENTS I would like to thank my supervisor Dr. Tim O’Connor for giving me an opportunity to embark on a drug research project. I would also like to extend my deepest gratitude to Dr. Matt Ramer for his ongoing support in all the in vivo studies, as well Jacquelyn Cragg and Dr. Lowell McPhail for their technical support. I would also like to thank Dr. Michel Roberge and Dr. Cal Roskelley for their invaluable advice during the development of the high-throughput screen. Lastly, I would like to thank Dr. Wolfram Tetzlaff, and other members of the committee, for their encouragement and helpful questions to keep my project on track.  xi  DEDICATION  To my parents  xii  CO-AUTHORSHIP STATEMENT Chapters 3 and 4 are written with the help of Dr. Matt Ramer and members of the Ramer lab. During the study, Dr. Matt Ramer helped with the surgeries and experimental design for both chapters 3 and 4, while Dr. Lowell McPhail helped with most of the surgeries for chapter 3. Lastly, Jacquelyn Cragg has helped with most of behavioural studies discussed in chapter 4.  xiii  CHAPTER 1  Introduction: The Sum of All Parts  1  1.1 INTRODUCTION The mammalian nervous system is an intricate circuitry consisting of billions of axonal connections. This complex network is established for the most part during embryonic and prenatal development, and to some extent during early postnatal development. Although understanding the process behind nervous system development has been a quest since the time of Santiago Ramón y Cajal (Hamburger, 1980; de Castro et al., 2007), we have only scratched the surface of understanding how this occurs. Our current understanding today suggests that the nervous system circuitry is established by the combined action of a variety of neurotrophic factors that promote axon outgrowth, and a myriad of guidance cues that guide axons to their appropriate targets (Yu & Bargmann, 2001; de Castro et al., 2007).  Although the nervous system has the remarkable ability to build and connect itself during development, its ability to repair itself following trauma is compromised. Its inability to repair itself becomes increasingly apparent as the nervous system matures and ages (Hafidi et al., 1999). In light of the current prevalence of spinal cord and brain injuries, and their debilitating consequences, a crucial question that arises is why does the nervous system not repair itself? What changes have occurred that prevent the adult nervous system from reconnecting itself by recapitulating axon development? In order to address these questions, I will review the macroscopic components of the mature nervous system and the changes that may occur following injury.  The mammalian nervous system is a complex circuitry that processes sensory information, conducts movements and regulates the body’s autonomic and homeostatic functions. Functionally, the nervous system can be divided roughly into the peripheral and central nervous systems. Macroscopically, the peripheral circuitry is built not only by neurons that make the 2  connections, but also by supporting glial cells known as Schwann cells. Specifically, Schwann cells form a myelin sheath around peripheral axons necessary for effective nerve conduction. In addition to their role in nerve conduction, Schwann cells produce a variety of growth factors and cell adhesion molecules that support neuronal survival and axon regeneration following damages to the peripheral nerve (Bunge, 1994; Martini, 1994).  The central nervous system (CNS) on the other hand is composed of the grey matter and white matter. In general, the grey matter is comprised of neuronal cell bodies, while the white matter is comprised of axonal projections. At the cellular level, the CNS circuitry is typically composed not only of various neuronal cell types, but also a number of supportive glial cells. Generally, neuronal cell types include projection neurons that make up the key axonal projections in the spinal cord and between higher order centers in the brain, as well as interneurons. Supportive glial cells include astrocytes, oligodendrocytes and microglia. Astrocytes contribute to the blood brain barrier, and are important for maintaining metabolic homeostasis in the central nervous system necessary to protect neurons from oxidative stress (Dienel & Cruz, 2003; Pellerin et al., 2007). Oligodendrocytes on the other hand play an important role in forming the myelin sheath necessary for rapid saltatory conduction of nerve impulses along projection axons, as well as maintaining axonal survival and integrity through the expression of glial specific proteins such as PLP [proteolipid protein], CNP [2',3'-cyclic nucleotide 3'-phosphodiesterase] and MAG [myelin associated glycoprotein] (Nave & Trapp, 2008). Lastly, microglia are endogenous macrophages in the CNS that play an important role in neuroprotection, clearing cell debris following injury, and recruiting the necessary innate immune response to eliminate foreign antigens or damaged cells (Garden & Moller, 2008).  3  1.2. INJURY TO THE NERVOUS SYSTEM Following injury, damaged axons demonstrate a remarkable ability to spontaneously initiate axon regeneration (Spira et al., 2001; Zheng et al., 2001; Chierzi et al., 2005). However, the success of axon regeneration depends in part on the availability of key signaling molecules such as cell adhesion molecules and growth factors necessary to support regeneration (Nieke & Schachner, 1985; Bixby et al., 1988; Martini et al., 1994; Werner et al., 2000; Oliveira et al., 2004; McGraw et al., 2005; Lindholm et al., 1987; Meyer et al., 1992; Curtis et al., 1994; Akazawa et al., 2004; Islamov et al., 2004). Other studies have also suggested that the expression of key regeneration associated genes may also influence the success of axon regeneration (Kobayashi et al., 1997; Cafferty et al., 2004).  1.2.1. Injury in the Peripheral Nervous System In the peripheral nervous system, damaged axons can successfully regenerate and reconnect with their original target. This can be attributed to the Schwann cells’ response to peripheral nerve injury, specifically by increasing the expression of numerous growth-promoting molecules. Permissive cell adhesion molecules such as neural cell adhesion molecule (NCAM), L1, and Ncadherin expressed on the surface of Schwann cells provide a substrate supporting axon outgrowth (Nieke & Schachner, 1985; Bixby et al., 1988). Similarly, the extracellular matrix molecule laminin and its receptors integrin, galectin and beta-2 microglobulin, are necessary for making the basal lamina more permissive for axon regeneration (Martini et al., 1994; Werner et al., 2000; Oliveira et al., 2004; McGraw et al., 2005). In addition, neurotrophic factors and growth factors including nerve growth factor (NGF), brain derived neurotrophic factors (BDNF), insulin like growth factors (IGF), vascular endothelial growth factor (VEGF), transforming growth factor beta-1 (TGFβ1), and neurokines, provide positive signals to stimulate axon  4  regeneration (Lindholm et al., 1987; Meyer et al., 1992; Curtis et al., 1994; Akazawa et al., 2004; Islamov et al., 2004). Collectively, Schwann cells play a pivotal role in mediating successful axon regeneration following peripheral nerve injury.  1.2.2. Injury in the Central Nervous System Following axon injury in the central nervous system, axons do not readily regenerate. The lack of axonal regeneration could be attributed to the non-permissive conditions in the adult CNS. To that end, pioneering studies by a number of groups in the 1980s have demonstrated that injured CNS axons could readily regenerate in the presence of permissive conditions such as that found in peripheral nerve grafts (David & Aguayo, 1981; Schwab & Theonen 1985). These findings have spurred a series of molecular studies to better understand the molecular basis underlying the inhibitory nature of the adult CNS. These studies have lead to the identification of the classical CNS derived inhibitory molecules including myelin associated inhibitory proteins (MAIPs) and chondroitin sulfate proteoglycans (CSPGs).  1.2.2.a. The Discovery of MAIPs The Schwab group was the first to discover that myelin in the CNS white matter is a potent inhibitor of axonal outgrowth (Caroni & Schwab, 1988; Schnell et al., 1990). Initial in vitro studies revealed that myelin demonstrated “non-permissive” effects on fibroblast spreading and neurite outgrowth in culture (Caroni & Schwab, 1988). From further extraction of the CNS white matter yielded a 250kD substrate bound protein that demonstrated “non-permissive” effects on fibroblast spreading and neurite outgrowth (Caroni & Schwab, 1988). Moreover, they found that rat anti-sera raised against the 250kD protein could effectively neutralize its non-permissive effects on neurite outgrowth (Caroni & Schwab, 1988). From the rat anti-sera, a monoclonal  5  antibody (IN-1) specific for the 250kD protein was identified (Caroni & Schwab, 1988). Further in vivo studies using a rat corticospinal tract (CST) transection model revealed that while CST axons do not readily regenerate in control animals, significant CST sprouting across the lesion was observed in IN-1 treated animals (Schnell & Schwab, 1990). Recent molecular advances have enabled the 250kD protein to be cloned (Chen et al., 2000). The cloning studies revealed that the ~250kD protein, now referred to as Nogo, is a member of the reticulon (RTN) family of proteins. Nogo consists of 3 splice variants NogoA, NogoB, and NogoC. NogoA in particular is expressed mostly by oligodendrocytes in white matter of the adult CNS, and demonstrates potent inhibitory effects on neurite outgrowth (Chen et al., 2000). Further molecular analysis revealed that NogoA has 3 inhibitory epitopes, with the conserved Nogo-66 loop region in the C-terminal being the most potent (Oertle et al., 2003; Grandpre et al., 2002). The Stritmatter group further identified a receptor for the Nogo-66 region, referred to as nogo receptor (NgR). NgR is a 443residue glycosyl-phosphatidylinositol-linked leucine-rich repeat glycoprotein (Fournier et al., 2001; Grandpre et al., 2002). Moreover, NgR is shown to bind a number of other later identified MAIPs including myelin-associated glycoprotein (MAG) and OMGp (oligodendrocyte myelin glycoprotein) (Barton et al., 2003; Wong et al., 2002; Niederost et al., 2002; Yiu & He, 2003). NgR forms a receptor complex with p75 neurotrophin receptor to activate downstream signaling pathways such as RhoA to inhibit neurite outgrowth (Wong et al., 2002; Yiu & He, 2003).  NogoA as well as other MAIPs function to inhibit spontaneous sprouting of axons in the adult CNS. Indeed, ectopic application of IN-1 antibodies into the CNS enhanced the spontaneous sprouting of a number of axonal populations including the corticospinal axons as well as Purkinje cell axons from the cerebellum (Buffo et al., 2000; Bareyre et al., 2002). IN-1 treatment also resulted in the upregulation of immediate early regeneration associated genes such as  6  GAP43 (Bareyre et al., 2002).  A key issue with this spontaneous sprouting is the possibility of generating aberrant connections that may result in undesired behavioural consequences. For example, previous studies have correlated aberrant sprouting of calcitonin-gene related peptide (CGRP) expressing nociceptive afferents in the spinal cord with the development of profound dysfunction in pain processing and autonomic function (Weaver et al., 1997; Christensen & Holsebosch, 1999; Lindsey et al., 2000; Mills et al., 2001b; Ackery et al., 2007). Therefore, by limiting spontaneous axonal sprouting, MAIPs may be key proteins in maintaining functional integrity of the adult CNS. However, this may be detrimental following axonal injury as spontaneous axon sprouting could be a potential mechanism through which functional reconnections could be made. For example, spontaneous sprouting of corticospinal axons following CST transection has resulted in recovery in motor function (Weidner et al., 2000). Moreover, IN-1 mediated spontaneous sprouting of corticospinal and serotonergic axons following injury has been correlated with recovery in motor and sensory functions (Bregman et al., 1995; Thallmair et al., 1998; Raineteau et al., 2001; Merkler et al., 2001; Li & Stritmatter, 2003). Lastly, systemic deletion of NogoA in knockout mice significantly enhanced axonal sprouting and motor recovery following spinal cord injury (Simeon et al., 2003; Kim et al., 2003). In conclusion, it appears that axon sprouting could be harnessed to stimulate functional repair following spinal cord injury.  The above findings have lead to the progressive development of molecular approaches to inhibit myelin signaling including the use of IN-1 antibodies, NgR peptide antagonists, as well as downstream signal manipulations (Schwab, 2004), which will be described further in Section 1.3. These molecular strategies have shown promising results in rat spinal cord injury models  7  (Thallmair et al., 1998; Lehman et al., 1999; Baeryre et al., 2002; Grandpre et al., 2002; Fournier et al., 2002, 2003; Dergham et al., 2002; Li & Stritmatter, 2003; Merkler et al., 2001; Fouad et al., 2001).  1.2.2.b. CSPGs Contribute to the Inhibitory Conditions in the Injured CNS CNS injury is often followed by the development of an astroglial scar consisting of a mass of hypertrophic astrocytes. The glial scar is inhibitory to axonal outgrowth, and its non-permissive nature is partly attributed to the upregulation chondroitin sulfate proteoglycans (CSPGs) at the site of the lesion (Pindzola et al., 1993; Lemons et al., 1999). Previous results have shown that removal of CSPGs from the injury site with the spinal infusion of a bacterial enzyme chondroitinase ABC enhanced axonal regeneration and sprouting (Lemons et al., 1999). Similarly, a number of studies have demonstrated that chondroitinase ABC treatment and its effects on axonal regeneration and sprouting corresponded with robust sensory and motor recovery following spinal cord transection (Bradbury et al., 2002; Barritt et al., 2006). However, the Bradbury group further demonstrated that spontaneous sprouting of particularly the CGRP+ nociceptive afferents resulted in the development of abnormal pain responses to non-noxious mechanical and thermal stimulation (Baritt et al., 2006). In conclusion, these findings suggest that similar to MAIPs, CSPGs are key for maintaining the functional integrity of the adult CNS and that they may hinder axonal sprouting and functional recovery following axonal injury in the CNS. It remains to be seen whether axon-sprouting effects of chondroitinase ABC treatment could be appropriately harnessed to promote functional recovery following spinal cord injury.  1.2.2.c. Glial Changes following CNS injury Although the damaged axon successfully initiates regeneration, the inhibitory environment in the  8  central nervous system and its exacerbation by a number of injury-mediated secondary events prevent axons from reconnecting to their targets (Fawcett & Asher, 1999; Zhang et al., 1997; Crowe et al., 1997; Niederost et al., 2002; Monnier et al., 2003; McPhail et al., 2007). These post-injury events could be described as a series of glial changes that take place following injury. As mentioned above, a classical phenomenon following spinal cord injury is the development of an astroglial scar, which consists of a mass of hypertrophic astroglial cells expressing CSPGs as well as a number of inhibitory molecules hindering axonal regeneration (Lemons et al., 1999; Fawcett & Asher 1999). Another well observed glial change following injury is the activation and recruitment of microglia/macrophages to the lesion site. Microglia/macrophages are crucial for the clearance of axonal and myelin debris following axonal injury, in a process known as Wallerian degeneration. This process often results in the release of myelin debris carrying MAIPs, further inhibiting axonal regeneration and sprouting in the injured CNS (McPhail et al., 2007). Recent studies by Silver’s group have demonstrated that reactive microglia/macrophages drive active axonal retraction or dieback through a contact dependent mechanism (Horn et al., 2009). Lastly, microglia/macrophages are classical mediators of the inflammatory response, which typically results in secondary tissue damage in the CNS (Garden & Moller, 2002). In conclusion, the variety of glial changes that occur following injury make a significant contribution to the multiple levels of inhibition in the CNS impeding axonal repair.  1.3. AXON REPAIR STRATEGIES The inhibitory environment in the adult CNS appears to be a major obstacle hindering the success of axon regeneration and compensatory axon sprouting (Li et al., 2004, Li & Strittmatter, 2003; Bradbury et al., 2002; MacDermid et al., 2005; Cafferty et al., 2008; Barritt et al., 2006). In order to overcome this obstacle, current strategies are focused not only to make the CNS  9  environment more favourable for axon outgrowth, but to also stimulate axon outgrowth by manipulating neurons intrinsically.  1.3.1. Axon Regeneration in the CNS In order to make the CNS environment more permissive for axon regeneration, one approach is to remove the inhibitory molecules in the CNS that may hinder axon regeneration. Previous studies by the Schwab and Strittmatter groups have shown that the use of the IN-1 antibodiy and molecular antagonists to neutralize inhibitory myelin associated molecules (Schnell & Schwab, 1990), or soluble NgR to antagonize the common Nogo receptor (NgR) complex, can stimulate robust axon regeneration and motor recovery following spinal cord hemisection (Li et al., 2004, Fournier et al., 2002; Li & Strittmatter, 2003). Similarly, a number of studies in the past have shown that following spinal cord lesion, the degradation of CSPGs using the enzyme chondroitinase ABC can stimulate robust regeneration and sprouting of the corticospinal and dorsal column axons, as well as restoration of both sensory and motor functions (Pindzola et al., 1993; Lemons et al., 1999; Bradbury et al., 2002; Barritt et al., 2006). Despite promising results, the beneficial effects of these interventions are limited due to a number of pitfalls. For example, ChABC activity is short-lived in vivo, and its growth promoting effects may be hindered by the rapid re-expression of CSPGs in the long-term (Laabs et al., 2007). On the other hand, due to the presence of multiple functionally redundant NgR isoforms in the CNS, nogo receptor antagonists may not completely neutralize the effects of myelin associated inhibitory proteins (Li et al., 2005). Likewise, knockout studies have revealed the presence of multiple isoforms of Nogo in the CNS and neutralization of one does not completely abolish the non-permissive nature of the CNS white matter (Zheng et al., 2003; Kim et al., 2003). Moreover, with the discovery of additional types of MAIPs including MAG and OmgP, and proteoglycans such as V2 and  10  brevican (Schwab, 2004; Niedorost et al., 1999; Filbin et al., 2003), it is apparent that multiple antagonists may be required to fully eliminate the inhibitory nature of the injured CNS. Therefore, approaches to antagonize MAIPs, and to remove CSPGs, should be complemented with other strategies to stimulate axon repair. Below are additional examples of other approaches.  One strategy to stimulate axon repair is to provide positive signals such as growth factors (Kobayashi et al., 1997; Tucker et al., 2008; McGraw et al., 2004). To stimulate axon regeneration, one approach is to introduce neurotrophins to the neurons either directly at the cell body or at the axon terminals. For example, application of brain derived neurotrophic factor (BDNF) and neurotrophin 4/5 (NT4/5) directly to the cell body resulted in the upregulation of regeneration-associated proteins and the increased propensity of axon regeneration and sprouting (Hiebert et al., 2002). However, because a number of studies revealed that neurotrophins can mediate neuronal cell death and axon retraction via the p75 neurotrophin receptor that is upregulated in the injured CNS (Kaplan & Miller, 2000; Hagg et al., 2007), using neurotrophins to stimulate axon regeneration in the CNS may not be recommended.  Another strategy to encourage axon regeneration involves filling the lesion center with a substrate that is more permissive to axon regeneration. To achieve this, one approach is to transplant stem cells into the cavity and then allow the stem cells to differentiate into the appropriate glial cells necessary to support axon regeneration and tissue repair (Ramer et al., 2005). Some of the stem cells used for transplants include neural stem cells, olfactory ensheathing cells, mesenchymal stem cells, skin derived progenitors and embryonic stem cells (Richter & Roskams, 2008; Parr et al., 2007; Biernaskie et al., 2007; McDonald & Howard,  11  2002; Coutts & Keirstead, 2008). Other approaches include the transplantation of neonatal astroglial cells embedded in collagen scaffolds (Deumanns et al., 2006) as well as neurotrophin expressing fibroblasts (Jin et al., 2002; Tobia et al., 2003) to create a permissive environment for axons to regenerate. Although these approaches significantly reduced the size of the cavity and stimulated robust axon regeneration into the lesion (Ramer et al., 2004; Richter et al., 2005; Richter & Roskams, 2008; Jin et al., 2002; Tobia et al., 2003; Deumens et al., 2006; Biernaskie et al., 2007), few axons were observed to reconnect to their final target. Further observations showed that regenerating axons tended to remain within the permissive transplant and failed to grow into the distal spinal cord (Jin et al., 2002; Tobias et al., 2003; Deumens et al., 2006; Biernaskie et al., 2007; Richter & Roskams, 2008). Indeed, these studies demonstrate that the absence of a continuous permissive substrate in the CNS is a key aspect preventing axon regeneration.  1.3.2.a. Spontaneous Axon Sprouting in the CNS Although the manipulation of the CNS inhibitory environment is often enough to encourage functional recovery (Bradbury et al., 2002; Barritt et al., 2006; Li & Strittmatter, 2003; Li et al., 2004), an interesting question that arises is whether the functional recovery is the direct consequence of axon regeneration. Intriguingly, subsequent structural analyses have revealed significant sprouting of not only regenerating axons, but of intact axonal projections as well (Bradbury et al., 2002; Barritt et al., 2006). Two alternative interpretations could arise from these observations. One is that the regenerating axons are responsible for the functional recovery observed possibly by reconnecting to their appropriate targets. However, in order to achieve successful repair, the regenerating axons would have had to grow through the spinal cord lesion, and then regenerate the long distance to their appropriate target. Given the significant distance  12  and time required for regenerating axons to reach their target, the observed functional recovery may not necessarily be the direct consequence of axon regeneration. Another interpretation is that functional recovery may be the result of the spontaneous sprouting of intact axonal projections outside the site of lesion. Indeed, studies by the Tuszynski group have shown that spontaneous sprouting of intact axons in the motor circuitry does play an important role in motor recovery (Weidner et al., 2001). In their studies, they showed that following corticospinal tract lesion, spontaneous recovery in motor function occurred. Furthermore, lesioning of these intact projections resulted in the complete abolishment of spontaneous motor recovery (Weidner et al., 2001). They therefore concluded that spontaneous recovery results from the sprouting of intact corticospinal projections (Weidner et al., 2001).  A similar phenomenon was observed in the sensory circuitry following dorsal root injury (DRI). Studies by Darian-Smith revealed that dorsal rhizotomy in primates resulted in spontaneous reorganization of the somatosensory receptive fields, which was attributed to the sprouting of intact sensory projections at the level of the spinal cord (Darian-Smith & Brown, 2000; DarianSmith, 2004). The structural reorganization in the sensory circuitry coincided with the apparent recovery in tactile mechanosensation in the affected hand (Darian-Smith & Ciferri, 2005). Similarly, recent studies by the Ramer group have shown dorsal rhizotomy in rats resulted in the spontaneous intraspinal sprouting that often resulted in recovery from neuropathic pain as well as restoration of mechanosensation (Ramer et al., 2004; Ramer et al., 2007). However, the spontaneous mechanosensory recovery was often observed appreciably after dorsal root injury, and in most cases the recovery was partial (Ramer et al., 2007). This finding suggests that while spontaneous intraspinal axon sprouting may not be sufficient for complete functional recovery, a potential strategy to enhance functional recovery may be to stimulate sprouting of intact axons.  13  1.3.2.b Axon Plasticity- A Potential Mode of Repair? Although axon regeneration is largely compromised in the CNS, there are examples of spontaneous reorganization in the CNS circuitry that can restore the function of the damaged connections (Weidner et al., 2001; Darian-Smith, 2004). This is apparently achieved through the compensatory sprouting of intact axonal projections, which serves to take over the function of their damaged counterparts. The axon sprouting is stimulated by injury related factors including neurotrophins such as NT-3 and BDNF (Chen et al., 2008; Ramer et al., 2007, Soril et al., 2008). However, the extent of axon sprouting is limited by inhibitory molecules in the CNS, which are previously shown to prevent spontaneous aberrant axon sprouting that may negatively alter the circuitry of the intact CNS (MacDermid et al., 2005; Barritt et al., 2006; Cafferty et al., 2008; Bareyre et al., 2002). Consequences of aberrant axonal sprouting as described in Section 1.2 include the development of neuropathic pain and autonomic dysfunction. In the injured CNS however, these inhibitors are impediments that prevent spontaneous axon reorganization and the associated restoration of sensory and motor function (Weidner et al., 2001; Darian-Smith, 2000; Ramer et al., 2004, 2007; Barritt et al., 2006; Cafferty et al., 2008).  In conclusion, it appears that spontaneous axonal sprouting in the adult CNS could result in beneficial and detrimental consequences. To harness the beneficial influences on axonal sprouting, future approaches may involve restricting axonal sprouting to only a specific population of axons needed to restore a specific motor or sensory function (Fouad & Tse, 2008). On the other hand, the Fouad group (and others previously) has shown that exercise or rehabilitation could be a means for triggering the appropriate spontaneous axonal sprouting and functional reconnections (Wiesel et al., 1965; Josephson et al., 2003; Gomez-Pinilla et al., 2002).  14  Lastly, interventions designed to enhance axonal sprouting should be complemented with the appropriate rehabilitative training to further enhance functional recovery.  1.3.2.c. Strategies to Stimulate Axon Sprouting A number of studies have revealed that removing and/or neutralizing inhibitory molecules results in significantly greater intraspinal axon sprouting and concomitant improvements in sensory and/or motor function (Bradbury et al., 2002; Massey et al., 2006; Barritt et al., 2006; Raineteau et al., 2001; Cafferty et al., 2008; MacDermid et al., 2004; Ramer et al. 2004, 2007). For example, spinal infusion of the chondroitinase ABC to remove inhibitory CSPGs in the spinal cord was effective in stimulating significant axon sprouting and recovery in sensory and motor functions following spinal cord lesions (Bradbury et al., 2002; Massey et al., 2006; Barritt et al., 2006). Likewise, neutralization of inhibitory myelin associated proteins by spinally infusing a myelin specific antibody, IN-1, resulted in long-lasting CST sprouting and reorganization (Raineteau et al., 2001; Bareyre et al., 2002). Similarly, after dorsal rhizotomy the removal of CSPGs by chondroitinase ABC spinal infusion resulted in robust axon sprouting in the dorsal horn (Cafferty et al., 2008). Furthermore, the neutralization of the myelin associated inhibitory molecules by infusing the soluble nogo receptor (sNgR) into the spinal cord also resulted in increased axon sprouting of sensory projections into the deafferented dorsal horn (MacDermid et al., 2004), which has been implicated in spontaneous recovery from cold pain as well as improvements in mechanosensory function (Ramer et al. 2004, 2007). The beneficial effects of the above interventions, however, may be attenuated by the previously mentioned limitations of ChABC and nogo antagonists in vivo (Laabs et al., 2007; Li et al., 2005).  As an alternative approach, the introduction of neurotrophic factors such as NT-3 and BDNF into  15  the spinal cord, either by direct infusion, viral gene therapy or with cellular/fibrin vehicle, has been previously shown to promote robust axon sprouting in the spinal cord (Tobias et al., 2003; Taylor et al., 2006, Chen et al., 2006) and a concomitant recovery of motor function (Shumsky et al., 2005; Mitsui et al., 2005). Similar studies revealed that intrathecal delivery of neurotrophins into the spinal cord promoted intraspinal sprouting of the sensory circuitry that resulted in partial sensory recovery (Tang et al., 2004; Ramer et al., 2007). However, studies by the Tetzlaff group have shown that after prolonged infusion of NT-3 into the spinal cord following partial corticospinal tract lesion, the compensatory sprouting of intact corticospinal tract axons appears to be selectively diminished (Hagg et al., 2007). Similar studies by the Ramer group have also demonstrated that endogenous neurotrophins such as BDNF and NT-4 appear to selectively inhibit the sprouting of VGLUT1-positive mechanosensory afferents, markedly reducing mechanosensory recovery (Ramer et al., 2007; Soril et al., 2008). The negative effects of prolonged neurotrophin treatment may be attributed to neurotrophin mediated activation of p75 neurotrophin receptor that is upregulated in most neuronal subtypes in the adult central nervous system, particularly following injury (Hannila & Kawaja, 2005; Scott et al., 2005). The p75 receptor activates a series of signaling cascades including the JNK-p53-Bax cell death pathway and the RhoA signaling pathway that reduces neuronal survival, and inhibits axon regeneration and sprouting (Scott et al., 2005; Kaplan & Miller, 2000). To avoid this problem, perhaps a more prudent strategy may be to target downstream effectors through which neurotrophins stimulate axon sprouting.  16  1.4. MANIPULATING THE NEURONAL GROWTH CONE The spinal cord repair strategies described above are focused on making the CNS environment permissive to axon outgrowth, and on enhancing regenerative potential of CNS axons by stimulating the expression of growth promoting genes (Kobayshi et al., 1997; Hiebert et al., 2002). Although degrading and/or neutralizing inhibitory molecules can encourage axon regeneration and sprouting, the effects are often limited in vivo. For example, the effects of ChABC treatment are attenuated due to the limited lifetime of ChABC in vivo well as the robust re-expression of the CSPGs (Laabs et al., 2007). On the other hand, the effects of NgR antagonists are limited due to the presence of multiple splice forms of NgR that are functionally redundant, and may not all be inhibited by one antagonist (Li et al., 2005). Furthermore, while neurotrophins encourage the expression of growth promoting genes to stimulate axon growth, recent studies demonstrating the detrimental effects of prolonged neurotrophin treatment suggest that there is much to learn about neurotrophins before they can be safely implemented for axon repair (Scott et al., 2005; Hagg et al., 2007). To improve upon the above strategies to promote axon repair in the CNS, a complementary approach may be to manipulate the machinery that is involved in axon outgrowth and branching. In the section below, I will review our current understanding of the mechanisms underyling axon outgrowth and branching.  1.4.1. Neuronal Growth Cone- Machinery of Axon Outgrowth and Branching Axon outgrowth is initiated by a specialized structure at the tip of growing axons and branches, referred to as the neuronal growth cone. A key function of the neuronal growth cone is to guide growing axons or axon branches as they pathfind to their appropriate targets during development (Tessier-Lavigne & Goodman, 1996). The growth cone consists of three basic regions: peripheral (P), intermediate (I) and central (C) regions (Dent & Gertler, 2003). The P region is the leading  17  edge of the growth cone consisting of dynamic meshworks and bundles of F-actin that support lamellipodial and filopodial protrusions. Towards the center (C region) of the growth cone, the dynamic F-actin structures give way to more stable structures supported mainly by microtubules (Dent & Gertler, 2003). The transition between the P and C regions is the intermediate or transitional zone where F-actin and microtubules interact (Dent & Gertler, 2003; Rodriguez et al., 2003). As the P region advances through F-actin polymerization, microtubules invade into the intermediate zone to promote axon consolidation (Dickson, 2001; Rodriguez et al., 2003; Dent & Gertler, 2003). The balance of F-actin assembly and disassembly governs the rate of axon/branch growth or retraction (Dickson, 2001; Luo, 2002).  Growth cones sense and respond to a variety of guidance cues that are expressed in the developing embryo (Tessier-Lavigne & Goodman, 1996). Guidance cues can include instructional guidance cues that can attract or repulse growing axons, and permissive guidance cues or substrates that support or inhibit axon outgrowth (Tessier-Lavigne & Goodman, 1996). In the mature nervous system, guidance cues expressed following axonal injury mediate very different effects in the PNS and CNS. For example, injury in the peripheral nervous system stimulates the expression of permissive guidance cues such as NCAM and laminin to enhance growth cone motility (Menager et al., 2004), supporting successful axon regeneration. In contrast, the central nervous system not only lacks the positive guidance cues to support axon regeneration and pathfinding, but also expresses a number of inhibitory guidance cues including ephrins and semaphorins (Niclou et al., 2006). Although the re-expression of ephrins and semaphorins has been shown to guide regenerating axons to their targets following optic nerve injury (Pasterkamp, 1999b; Rodger et al., 2004), a number of studies in spinal cord injury have demonstrated that ephrins and ephrin receptors (EphA4, EphrinB2, EphrinB3), and semaphorins  18  (Sema 3A, 4D and 5A) are among the inhibitory guidance cues expressed in the glial scar (Fabes et al., 2006; Pasterkamp et al., 1999a; Pasterkamp and Verhaagen, 2001; Goldberg et al., 2004; Moreau-Fauvarque et al., 2003). Moreover, there is evidence that ephrins and semaphorins in the glial scar may act in concert with CSPGs and/or myelin associated inhibitors (such as Nogo and MAG) to inhibit growth cone motility, and therefore hinder axon regeneration (Benson et al., 2005; De wit et al., 2005; Schweigreiter and Bandtlow, 2006).  1.4.2. Signaling Pathways Regulating Growth Cone Motility Intriguingly, the motility of the neuronal growth cone is regulated by a set of signaling pathways that regulate cell motility. Conserved signaling components between motile cells and growth cones include the Rho family of GTPases (Dickson, 2001; Gallo & Letourneau, 2004), and upstream calcium (Wen et al., 2004; Gomez & Zheng, 2006; To et al., 2007, 2008), cyclic adenosine monophosphate (cAMP) (Han et al., 2007), as well as phosphotidyl-inositol-3 kinase signaling (Menager et al., 2004; Orlava et al., 2007; Wolf et al., 2008).  1.4.2.a. Rho Family of GTPases Neuronal growth cones respond to permissive and inhibitory guidance cues through intracellular switches that regulate F-actin and microtubule dynamics. These switches are the Rho family of GTPases that are activated by binding guanosine triphosphate (GTP). Members of the Rho family of GTPases include RhoA, Rac and Cdc42. Generally, RhoA has been implicated in inhibiting growth cone motility, while Rac and Cdc42 are known to stimulate growth cone motility (Dickson, 2001).  When the growth cone encounters permissive or attractive guidance cues, the growth cone  19  responds by activating the specific Rho family GTPases (Rac and Cdc42) to enhance F-actin assembly (Dickson, 2001). Similar to the role of Rac/Cdc42 in migrating cells, Rac/Cdc42 stimulates growth cone motility and axon consolidation possibly through their influence on downstream targets that typically regulate cell motility: F-actin nucleating complexes, F-actin recycling machinery, as well as on F-actin retrograde flow (Dickson, 2001; Meyer & Feldman, 2002; Schmidt et al., 2002). Specifically, Rac and Cdc42 stimulate the nucleation of new F-actin branches by recruiting the Arp2/3 (actin-related protein) complex through adaptor proteins such as N-WASP (neuronal Wiscott-Aldrich Syndrome protein), WAVE (WASP family verprolinhomologous protein) and SCAR (suppressor of cAMP receptor mutation) (Miki et al., 1996; Rohatgi et al. 1999; Yarar et al. 1999; Machesky et al., 1999; Dickson, 2001; Meyer & Feldman, 2002). Conversely, Rac/Cdc42 may also stimulate nucleation of unbranched F-actin filaments by the activation of the formin mDia (Pruyne et al., 2006). Rac and Cdc42 also stimulate the growth of F-actin filaments by recruiting proteins such as Ena-VASP and profilin to promote F-actin assembly (Goldberg et al., 2000; Nakegawa et al., 2001; Meyer & Feldman, 2002). Furthermore, Rac/Cdc42 directly modulates F-actin recycling through Pak (p21 activated kinase) mediated inhibition of LIM kinase and the downstream actin depolymerizing factor (ADF)/Cofilin, favoring actin protrusion in the leading edge (Chan et al., 2000; Meyer & Feldman, 2002; Samiere et al., 2003). Lastly, Rac/Cdc42 inhibits F-actin retrograde flow by negatively modulating myosin light chain kinase (MLCK) through Pak (Sanders et al., 1999; Schmidt et al., 2002; Brzeska et al., 2004). Collectively, Rac/Cdc42 is likely to stimulate growth cone motility by promoting F-actin nucleation and assembly, and inhibiting actin recycling and retrograde flow to favor the protrusion of F-actin filled processes at the leading edge of the neuronal growth cone (Meyer & Feldman, 2002). Finally, the Rac/Cdc42 in the protrusive F-actin rich leading edge may also recruit microtubule binding adaptor proteins such as IQGAP [IQ motif containing  20  GTPase activating protein] and CLIP170 (cytoplasmic linker protein 170) to promote microtubule capture and axon consolidation (Rodriguez et al., 2003).  On the contrary, when the growth cone encounters inhibitory guidance cues, it responds by activating RhoA (Dickson et al., 2001; Zhang et al., 2003). RhoA can mediate growth cone collapse by triggering F-actin retraction and microtubule disassembly (Zhang et al., 2003; To et al., 2008). Similar to motile cells, RhoA may mediate growth cone collapse by engaging the downstream effector Rho Kinase (ROCK) that activates myosin mediated retrograde F-actin flow, resulting in F-actin retraction and growth cone collapse (Dickson, 2001; Bzreska et al., 2004). Conversely, the RhoA/ROCK pathway may mediate growth cone collapse through MLCK mediated activation of bipolar myosin II mini-filament structures that are localized to the C and I regions of neuronal growth cones (Zhang et al., 2003; Amano et al., 1996). The resulting myosin II activation stimulates the actinomyosin contractility that mediates F-actin kinking/buckling, and potentiates severing of F-actin bundles in the growth cone intermediate zone (Medeiros et al., 2006). Lastly, ROCK may phosphorylate and inhibit the microtubule assembling protein, collapsin-response mediator protein 2 (CRMP2), resulting in microtubule disassembly (Goshima et al., 1995; Dickson, 2001; Mimura et al., 2006). Conversely, RhoA may also stabilize microtubule filaments in the leading edge through the activation of mDia, hindering microtubule assembly necessary for axon consolidation (Bartolini et al., 2008; Rodriguez et al., 2003). Overall, RhoA signaling counteracts the effects of Rac and Cdc42 on neuronal growth cone motility (Dickson et al., 2001; Meyer & Feldman, 2002).  1.4.2.b. Upstream Signaling Components Rho family GTPases are modulated by a number of upstream signaling components including  21  calcium, cAMP/PKA and PI3K. These signaling molecules are a crucial bridge between a guidance cue receptor and the growth cone’s cytoskeletal machinery. For instance, calcium plays an important role in mediating an attractive growth cone response to netrins during axon development (Hong et al., 2000; Gomez & Zheng, 2006). In the adult CNS however, calcium signaling also plays an important role in the repulsive growth cone response to inhibitory molecules in the CNS, including CSPGs and MAIPs (Gomez & Zheng, 2006; Henley & Poo, 2004). In addition to calcium, cAMP/PKA signaling has been previously demonstrated to mediate attractive growth cone response to factors such as glutamate and brain derived neurotrophic factor [BDNF] (Han et al., 2007). Finally, PI3K signaling has been shown to stimulate growth cone motility and axon outgrowth in response to neurotrophic factors and guidance cues (Menager et al., 2004; Wolf et al., 2008). Thus, in order to manipulate the aforementioned signaling players to favour growth cone motility and axon outgrowth, the first step is to better understand how each of the signaling effectors regulates growth cone motility.  Calcium is a conserved signaling molecule that regulates not only growth cone motility, but also motility in most cells (Collins & Meyer, 2009). It is found mostly in the extracellular environment or in the growth cone’s intracellular store, with little found in the growth cone cytoplasm (Collins & Meyer, 2009). Extracellular calcium may enter the neuronal growth cone through a family of voltage gated calcium channels known as the transient receptor potential channels (TRPCs), which are activated by guidance cues including netrins and brain derived neurotrophic factor (BDNF) (Shim et al., 2005; Wang et al., 2005; Li et al., 2005). On the other hand, calcium from intracellular stores may enter the cytosol through inositol 3 phosphate (IP3) gated calcium channels (Akiyama et al., 2009). The calcium entering the neuronal growth cone then activates a range of signaling effectors that mediate either an attractive or repulsive growth  22  cone response, or in a broader sense either enhanced growth cone motility or growth cone collapse (Henley & Poo, 2004; Gomez & Zheng, 2006). The extent of calcium influx relative to the baseline calcium level in the growth cone is a key regulator of the growth cone response (Kater & Mill, 1991; Henley & Poo, 2004). For instance, small or large calcium influxes relative to growth cone baseline calcium levels activate the calcium effectors calcineurin and calpain respectively (Gomez & Zheng, 2006), resulting in growth cone collapse by the following mechanisms: 1) Calcineurin dephosphorylates and activates protein phosphatase 1 (PP1) to reduce filopodia and veil protrusion (Wen et al., 2004); 2) Calpain on the other hand stimulates veil retraction through inhibition of the integrin adhesion complex (Robles et al., 2003; Gomez & Zheng, 2006). On the contrary, medium levels of calcium influxes relative to basal calcium levels in the growth cone results in growth cone attraction or enhancement in growth cone motility (Henley & Poo, 2004). Medium calcium influxes typically activate calmodulin kinase II (CAMKII), which in turn activates Rac/Cdc42 and downstream effects on growth cone motility (Henley & Poo, 2004). Furthermore, medium calcium influxes are also crucial for maintaining PI3K-Akt signaling to mediate growth cone motility (Zheng et al., 2008). However, in response to inhibitory guidance cues in the adult CNS, the resulting medium calcium influxes activate PKC, which in turn activates RhoA to mediate growth cone repulsion or collapse (Hasegawa et al., 2004; Henley & Poo, 2004; Sivansakaran et al., 2008).  Calcium signaling can also be regulated by cAMP levels in the neuronal growth cone (Lohof et al., 1992). Previous studies by Mu-ming Poo’s laboratory have shown that BDNF mediated cAMP elevation in the neuronal growth cone is responsible for converting a repulsive growth cone response into an attractive one (Ming et al., 1997; Song & Poo, 1999). Further studies revealed that cAMP activates the downstream effector protein kinase A (PKA). In the growth  23  cone, the most abundant type II protein kinase A is localized to the leading edge filopodia through its interaction with A kinase anchoring proteins [AKAPs] (Han et al., 2007). In response to cAMP elevation, PKA phosphorylates one of the recently elucidated effectors protein phosphatase 1 [PP1] (Han et al., 2007). As previously mentioned, PP1 is activated by a calcium effector calcineurin, and has been implicated in growth cone repulsion (Wen et al., 2004). Intriguingly, PKA appears to inhibit growth cone repulsion and/or collapse through the phosphorylation and deactivation of PP1 (Han et al., 2007). Overall, it appears that cAMP/PKA signaling counteracts calcineurin mediated growth cone repulsion or collapse (Han et al., 2007), turning the repulsive growth cone response into an attractive one. It is therefore conceivable that cAMP/PKA mediated changes in growth behaviour may convert a collapsed growth cone into a motile one necessary for successful axon regeneration and/or sprouting.  In addition to cAMP signaling, PI3K signaling appears to play a beneficial role in promoting growth cone motility. PI3K is a lipid kinase that converts the membrane phospholipid phosphatidyl-3,4-inositol-bisphosphate (PIP2) into phosphatidyl-3,4,5-triphosphate (PIP3). PI3K activity in particular has been implicated in the polarization and motility of migrating cells (Kolsch et al., 2008). Similar studies have demonstrated that PI3K in the neuronal growth cone also plays a similar role in promoting growth cone motility (Menager et al., 2004; Orlava et al., 2007; Wolf et al., 2008). Specifically, PI3K activity and PIP3 accumulation in the neuronal growth cone has been implicated in laminin induced axon polarization and outgrowth (Menager et al., 2004; Da Silva et al., 2005), and was shown to mediate attractive growth cone response to the developmental guidance cues including Wnt, netrins and slits (Wolf et al., 2008; Quinn & Wadsworth, 2008). Further characterization revealed that PI3K stimulates growth cone motility and axon consolidation through the recruitment of conserved signaling pathways that regulate F-  24  actin and microtubules (Menager et al., 2004; Da Silva et al., 2005; Zhou et al., 2004; Kim et al., 2006; Orlava et al., 2007; Etienne-Mannville & Hall, 2003). Specifically, PI3K stimulates Factin assembly through the activation of Rac and Cdc42, resulting in filopodial and lamellipodial protrusions in the growth cone’s leading edge (Dickson, 2001; Menager et al., 2004; Da Silva et al., 2005; Quinn & Wadsworth, 2008). PI3K also enhances microtubule assembly by inhibiting the glycogen synthase kinase-3 beta (GSK3-beta), resulting in the disinhibition of the downstream microtubule binding proteins involved in microtubule assembly including collapsin response mediator protein (CRMP-2), microtubule associated protein 1b(MAP1b), adenomatous polyposis coli (APC), and tau (Zhou et al., 2004; Kim et al., 2006; Orlava et al., 2007). Other proteins recruited by PI3K signaling may include the polarity protein complex (aPKC, mPar3, mPar6) necessary to re-organize the cytoskeleton with a distinctive front-back polarity characterized by a dynamic actin-filled leading edge and a trailing edge supported by the microtubule network (Etienne-Mannville & Hall, 2003). Lastly, PI3K also inhibits signaling components responsible for growth cone collapse and repulsion. Specifically, PI3K activation in the growth cone has been demonstrated to inhibit RhoA signaling, impeding the downstream effects of RhoA on growth cone collapse (Da Silva et al., 2005; Eickholt et al., 2007).  1.4.3. Growth Cone response to CNS Derived Inhibitory Molecules A hallmark of regeneration failure is the dystrophic growth cone, which is often referred to as a “frustrated” or “collapsed” growth cone incapable of driving further axonal growth into the unfavourable CNS environment (Tom et al., 2004). The typical features of the dystrophic growth cone include its bulbous and vacuolated structure, as well as the absence of a motile leading edge (Tom et al., 2004). Surprisingly, the dystrophic growth cone is not inert, but rather a highly dynamic structure that may still drive axon growth if provided with a more permissive  25  environment (Tom et al., 2004). It is therefore thought that the dystrophic growth cone is likely a chronically collapsed growth cone resulting from the continuous presence of inhibitory molecules in the CNS environment.  At a molecular level, myelin associated molecules (Nogo, MAG and OMGp) and CSPGs mediate growth collapse through the convergent activation of the Nogo receptor complex, which consists of primarily the glycophosphoinositol (GPI)-linked Nogo receptor (NgR) and its coreceptor p75 neurotrophic receptor (p75NTR) (McKerracher & Winton, 2002; Yamashita et al., 2002). Additional components recently identified in the Nogo receptor complex include the coreceptors Lingo-1 and the Taj/TROY from the orphan tumor necrosis factor receptor family (Schwab et al., 2006; Mi et al., 2004; Park et al., 2005). The receptor complex activates the downstream RhoA signaling to mediate growth cone collapse via the downstream targets as described in Section 1.4.2a, rendering the growth cone “dystrophic” or incapable of driving axon outgrowth (Niedorst et al., 2002; Monnier et al., 2003; McKerracher & Winton, 2002; Schweigreiter et al., 2004; Simvansakaran et al., 2004). Interestingly, a number of studies have shown that the intrinsic levels of calcium and cyclic-adenyl-monophosphate (cAMP) in the neuronal growth cone can collectively modulate the growth cone’s response to inhibitory myelin associated inhibitors and CSPGs (Snider et al., 2002; McKerracher & Winton, 2002; Wen et al., 2004; Henley et al., 2006; Henley & Poo, 2004). Increasing cAMP levels have been classically shown to reverse the collapsing effects of repulsive or inhibitory molecules on the neuronal growth cone (Snider et al., 2002), possibly by inhibiting RhoA through the downstream effector PKA (McKerracher & Winton, 2002). On the other hand, local changes to calcium levels can regulate the growth cone’s response to myelin associated molecules and CSPGs, where the level of calcium selectively activates calcium effectors mentioned earlier that either promote growth  26  cone repulsion or attraction (Wen et al., 2002). Overall, myelin associated proteins and CSPGs mediate growth cone collapse not only through the activation of NgR complex, but also by the combined actions of intrinsic secondary messengers cAMP and calcium in the neuronal growth cone (Henley & Poo, 2004; McKerracher & Winton, 2002). Importantly, these studies demonstrate that manipulation of these secondary messengers in the neuronal growth cone can help overcome the inhibitory effects of myelin associated inhibitors and CSPGs in the CNS.  1.4.4. Stimulating Growth Cone Motility Since the neuronal growth cone plays a pivotal role in axon outgrowth and branching, a promising strategy to promote axon regeneration and/or sprouting in the nervous system is to manipulate signaling pathways that regulate growth cone motility. A well-known approach is to inhibit signaling pathways through which inhibitory CNS molecules collapse the neuronal growth cone. Indeed, successful strategies have involved pharmacologically inhibiting signaling pathways in the growth cone that act downstream of CSPGs and MAIPs (Chan et al., 2005; Ramer et al., 2004; Sivasankaran et al., 2004). These interventions were observed to stimulate robust axon regeneration and sprouting (Chan et al., 2005; Ramer et al., 2004; Sivansankaran et al., 2004), and in some cases enhanced functional recovery (Chan et al., 2005; Ramer et al., 2004). However, some of these interventions such as the inhibition of ROCK, resulted in an apparent dose dependent increase in astrogliosis and upregulation of CSPGs, which counteracts the beneficial effects on axon regeneration (Chan et al., 2007). This raises a key issue that many of the signaling pathways regulating growth cone motility may be conserved in other cells (Irino et al., 2008; John et al., 2004; Etienne-Manneville & Hall, 2001). Moreover, it is highly likely that these signaling components such as ROCK may activate other signaling effectors that have not yet been elucidated. Thus, manipulating these pathways may result in undesired extraneous  27  effects. It is evident that further characterization of the neuronal growth cone may be required to identify novel signaling components that specifically regulate growth cone motility and not other unknown signaling effectors. One approach may be to explore signaling pathways in the neuronal growth cone by screening for novel compounds that may promote growth cone motility.  1.4.5. Screening for Novel Compounds to Enhance Growth Cone Motility Because the availability of compounds that can effectively stimulate growth cone motility are limited, I approached this problem by creating a screen to rapidly identify natural bioactive compounds that can stimulate growth cone motility, and hence axon outgrowth. As mentioned earlier, the neuronal growth cone has many characteristics that are similar to migrating cells, including front-back polarity, as well as conserved cytoskeletal and signaling components that regulate structure and motility (Dickson, 2001). Because of these similarities, migrating cells can be a useful model to study the signaling mechanisms underlying growth cone motility. A key component of any high-throughput screen is a library of bioactive compounds or extracts preferably with previously demonstrated effects on cell motility. A good source of these bioactive compounds is the marine extract library from the Dr. Raymond Andersen’s group at the UBC Department of Chemistry, as a number of compounds from this library have previously been shown to have a wide range of biological activities including anti-bacterial effects, cytotoxic activities, and anti-migratory effects on motile cells (Belarbi et al., 2003; Keyzers et al., 2008; Hadashcik et al., 2008; Carr et al., 2008; McHardy et al., 2005; Roskelley et al., 2001). Many of these compounds have been previously used for environmental applications such as antifouling (Li et al., 2004), as well as medicinal uses such as anti-cancer or immuno-modulatory therapies (Roskelley et al., 2001; Hadaschk et al., 2008; McHardy et al., 2005; Carr et al., 2008). Of particular interest is the marine compound Motuporamine C, which not only effectively  28  inhibits the migration (or metastasis) of breast cancer cells in vitro via activation of RhoA (Roskelley et al., 2001), but also inhibits neurite outgrowth in dorsal root ganglion explants via RhoA mediated growth cone collapse (To et al., 2006, 2008). These studies demonstrate for the first time that marine derived Motuporamine C can inhibit cancer cell motility and neurite outgrowth via a conserved pathway RhoA. The neuronal effects of Motuporamine C suggest that natural marine compounds are likely to possess cell migration and/or neurite outgrowth modulatory activities.  It has become clear that neuronal growth cones and motile cells can be regulated by similar signaling pathways, as indicated by their similar response to Motuporamine C, a compound derived from the marine extract library. Therefore, a possible approach to identify marine compounds with neurite outgrowth promoting activity may be to use motile cells as a model for the neuronal growth cone. Since many cell lines such as human embryonic kidney (HEK) 293 cells are highly motile, a distinctive advantage with using motile cell lines is the relative ease of culturing compared to primary neurons. Furthermore, a number of existing cell-migration assays for motile cells have been established (Rodriguez et al., 2005; Valster et al., 2005; Liang et al., 2007), and can be easily automated for high-throughput screening, allowing for rapid identification of candidates among the thousands of extracts (over millions of compounds) in the marine extract library. Among cell migration assays used thus far, the classical wound-healing assay is an ideal choice by virtue of its simplistic design. Briefly, the wound-healing assay can be done with a confluent layer of cells, where a scratch using a blunt instrument is made in the confluent layer, and subsequent cell migration back into the scratch is assessed (Rodriguez et al., 2005; Liang et al., 2007). While this approach may be beneficial in identifying compounds that stimulate neurite outgrowth, any positive compound ultimately will need to be tested in vivo to  29  ascertain its effectiveness in promoting axon regeneration or sprouting.  1.5. RAT DORSAL RHIZOTOMY- MODEL FOR REGENERATION A relevant model to evaluate axon regeneration across the regenerative barrier is the rat dorsal rhizotomy or dorsal root injury (DRI) model, which involves selective injury of sensory spinal roots. An advantage of this model is that the regenerative barrier is clearly demarcated at the PNS/CNS interface at the dorsal root entry zone (DREZ), allowing straightforward evaluation of axon regeneration across the regenerative barrier (Ramer et al., 2001). Unlike complete brachial plexus injury, selective or partial DRI can result in specific sensory deficits that can be used to evaluate the behavioural effects of the axon outgrowth promoting compounds that are being tested. For example, single lumbar dorsal rhizotomy was reported to elicit touch and mechanosensory allodynia (Li et al., 2000; Eschenfelder et al., 2000). Likewise, the rhizotomy of the 7th and 8th cervical roots (C7/C8 DRI) has been shown to result in profound and highly reproducible sensory dysfunction particularly in the transduction of pain and low-threshold mechanosensory stimuli (Ramer et al., 2004; Ramer et al., 2007). Specifically, C7/C8 DRI typically results in hypersensitivity to innocuous cold stimulation, which becomes apparent by five days post DRI and spontaneously recovers 20 days later (Ramer et al., 2004). At the same time, C7/C8 DRI abolishes the transduction of low-threshold mechanosensory stimuli immediately following injury, which partially recovers 8-10 days later (Ramer et al., 2007). The abovementioned sensory deficits associated with dorsal root injury are well correlated to the observed changes to the sensory circuitry (Darian-Smith, 2000; Ramer et al., 2004). Below I review the key functional components of the sensory circuitry that are affected by dorsal root injury.  30  1.5.1. Sensory Input- Primary Afferents and Second Order Projection Neurons Briefly, primary afferents in a dorsal root include a heterogeneous population of sensory axons that originate from sensory neurons located in the dorsal root ganglia (DRG) situated beside the spinal cord (Devor, 1999). These sensory neurons are pseudo-unipolar neurons that receive sensory information from receptors through one axon, and relay the sensory information to the spinal cord through the other axon (Figure 1.1). Among the sensory axons in the dorsal roots are large diameter afferents and small diameter afferents (Millan, 1999). Large diameter afferents include myelinated Aα and Aβ afferents, as well as the lightly myelinated Aδ afferents. Small diameter afferents on the other hand include slow conducting unmyelinated C afferents. Functionally, Aδ and C afferents respond to high-threshold stimuli, and are responsible for relaying noxious stimuli to projection neurons in laminae I and II of the dorsal horn (Bessou & Perl, 1969, Millan, 1999). The lightly myelinated Aδ fibers also conduct at intermediate velocities, and may therefore respond to a wide range of innocuous to noxious thermal and mechanical stimuli (Burgess and Perl, 1967; Light and Perl, 1979). In contrast, Aα and Aβ afferents respond to low threshold stimuli, and are responsible for relaying low-threshold mechanosensation (touch sensation), proprioception, and non-noxious thermal sensations to laminae III-VI of the dorsal horn (Woolf, 1987; Koerber et al., 1995; Millan, 1999).  In the dorsal horn, second-order sensory projection neurons make up the ascending sensory projections in the spinal cord (Figure 1.1). Nociceptive projection neurons populate laminae I, and the deeper laminae III-IV (Todd, 2002). The cell bodies of superficial projection neurons and dendrites of deeper projection neurons colocalize with the nociceptive terminals from C and Aδ  31  Figure 1.1. Sensory Circuitry. Sensory input from the periphery is transmitted through pseudounipolar sensory neurons located in the dorsal root ganglion (DRG). Within the DRG is a heterogeneous population of sensory neurons that are responsive to various sensory modalities. Nociceptive sensory neurons project small diameter lightly myelinated Aδ and C afferents to the superficial dorsal horn, terminating in laminae I and II. Sensory neurons responsive to innocuous stimuli project large diameter Aα and Aβ afferents to deeper regions of the dorsal horn, terminating in laminae III-VI. Nociceptive Aδ and C afferents directly activate nociceptive projection neurons in laminae I, III and IV, while Aα and Aβ afferents directly activate dorsal column projection neurons located in laminae IV-V of the dorsal horn. Aβ, Aδ and C afferents also mono- or poly-synaptically activate WDR neurons in laminae V-VI via interneurons and/or dendritic connections. Nociceptive projection neurons and WDR afferents cross the midline and travel rostrally to the thalamus along the contralateral spinothalamic tract. Dorsal column projection neurons on the other hand travel rostrally to the brainstem via the ascending dorsal column.  32  33  afferents located in lamina I (Lawson et al., 1997), suggesting that these projection neurons likely facilitate transduction of afferent input (Naim et al., 1997; Todd et al., 2002). There is also another population of projection neurons known as ‘wide dynamic range’ (WDR) neurons that are located in laminae V-VI (Ness and Gebhart, 1990; Gebhart, 1995). Unlike the nociceptive projection neurons, WDR neurons are responsive to innocuous and noxious stimuli, and may be activated by C, Aδ and Aβ afferents directly (monosynaptically) or via interneurons (polysynaptically) (Light and Kavookjian, 1988; De Koninck et al., 1992). Lastly, there is also another population of projection neurons in laminae IV-V often referred to as dorsal column projection neurons (Dykes & Craig, 1998). Electrophysiological studies suggest that these neurons are responsive to inputs from mechansensory afferents including Aα and Aβ afferents that terminate in laminae III-VI of the dorsal horn (Dykes & Criag, 1998; Woolf, 1987; Koerber et al., 1995; Millan, 1999).  Overall, the nociceptive projection neurons and WDR neurons make up axonal projections that cross the midline and travel rostrally via the lateral spinothalamic tract to posteriolateral nucleus of the thalamus (Craig et al., 1994). On the other hand, dorsal column projection neurons project directly to the brainstem nuclei (cuneate and gracile nuclei) via the dorsal and/or dorsolateral funiculus (Dykes & Craig, 1998).  1.5.2. Resident Interneurons in the Dorsal Horn In the spinal cord, sensory processing is modulated by inhibitory and excitatory interneurons that make up the majority of dorsal horn neurons (Szentagothai, 1964). Inhibitory interneurons include γ-aminobutyric acid (GABA) or glycinergic interneurons in the dorsal horn (McLaughlin et al., 1975; Campistron et al., 1986; Todd & Sullivan, 1990). The glycinergic phenotypes are  34  typically in laminae III-IV, while GABAergic interneurons populate most of the dorsal horn (Todd & Sullivan, 1990). Both glycinergic and GABAergic neurons synapse onto the presynaptic nociceptive afferents, and post-synaptic projection neurons (Todd, 1990; Powell and Todd, 1992). On the other hand, excitatory glutaminergic interneurons are located in laminae IIV of the dorsal horn. Because markers that identify glutaminergic interneurons such as vesicular glutamate transporter 2 (VGLUT2) are shared with other axonal projections in the dorsal horn including primary afferent terminals (Todd & Spike, 1992; Todd et al., 2003), the anatomical locations of these interneurons are only speculative. Presumptive neuropeptide markers including somatostatin and enkephalin identify these neurons by virtue of their lack of colocalization with markers identifying primary afferent and inhibitory interneurons (Todd et al., 2003). These excitatory interneurons are thought to provide polysynaptic excitation of dorsal horn projection neurons (Light & Kavookjian, 1988; De Koninck et al., 1992). Overall, the combined actions of both the inhibitory and excitatory interneurons provide dynamic regulation over sensory processing in the dorsal horn.  1.5.3. Descending Supraspinal Projections Afferent input is also modulated by a variety of descending monoaminergic inputs from the brainstem. Some of these inputs include serotonergic, dopaminergic, and noradrenergic/ adrenergic systems (Millan, 2002). Serotonergic projections originate from neurons located in the raphe nucleus in the brainstem, specifically within the area of the rostral ventromedial medulla (RVM) (Millan, 2002). Serotonergic varicosities identified by their expression of the serotonin transporter (SERT) are distributed in laminae I-II (MacDemid et al., 2004; Ramer et al., 2004; Ramer et al., 2007). On the other hand, dopaminergic and noradrenergic/adrenergic projections originate from the substantia nigra pars compacta and locus coeruleus respectively  35  (Millan, 2002). While both populations can be identified by their expression of tyrosine hydroxylase (TH), only noradrenergic/adrenergic varicosities can be identified by their exclusive expression of dopamine-β-hydroxylase (Armstrong et al., 1981; Verney et al., 1982). Immunoreactivity of both enzymes is uniformly expressed throughout the spinal cord with the exception of lamina II (MacDermid et al., 2004; Ramer et al., 2004).  Functionally, monoaminergic projections are classical modulators of pain transmission (Millan, 2002). Monoaminergic projections may synapse onto pre-synaptic nociceptive terminals to negatively modulate the release of noxious neurotransmitters such as glutamate and substance P, and therefore inhibit pain transmission (Millan, 1999, 2002). Conversely, monoaminergic projections may also enhance pain transmission by activating second-order nociceptive projection neurons either directly (monosynaptically) or indirectly (polysynaptically) via excitatory interneurons (Millan, 1999). Overall, the modulatory influences of descending monoaminergic projections on pain transmission are likely to depend on the subtype of receptors that are activated in pre-synaptic versus post-synaptic terminals (Millan et al., 1996; Millan, 2002).  1.5.4. Consequences of Dorsal Rhizotomy As described earlier, injury to the cervical C7 and C8 dorsal roots can result in a range of sensory dysfunctions including the development of cold hypersensitivity and the loss of low threshold mechanosensation (Ramer et al., 2004; Ramer et al., 2007; Soril et al., 2008). The loss of lowthreshold mechanosensation immediately following injury can be attributed to the loss mechanosensory inputs in the C7/C8 segment (Ramer et al., 2007). The development of cold hypersensitivity on the other hand is much more complex, and may involve changes in synaptic  36  input to the nociceptive projection neurons (Eschenfelder et al., 2000). For example, normal afferent input in the 5th lumbar (L5) dorsal root has been suggested to directly suppress presynaptic input from adjacent roots (Wall & Devor, 1983). The loss of the afferent input may remove the suppression of the adjacent roots, allowing acute activation of nociceptive projection neurons and the development of mechanical hypersensitivity (Eschenfelder et al., 2000). It is speculative that cold hypersensitivity following C7/C8 DRI may occur in a similar fashion. On the other hand, the changes in synaptic input could be attributed to a number of injury related events including the spontaneous reorganization of intact sensory projections (Erschenfelder et al., 2000; Millan, 2002; Ramer et al., 2004).  Afferent regeneration is inhibited by a number of glial events that take place following DRI. First, astrogliosis occurs within 1-day post DRI, particularly in the dorsal root entry zone (DREZ) demarcating the boundary between the spinal cord and the dorsal root (Ramer et al., 2001). The resulting glial scar expresses inhibitory CSPGs that prevent the injured afferents from regenerating back into the spinal cord (Ramer et al., 2001; McPhail et al., 2005). This is followed by Wallerian degeneration of the injured roots projecting into the spinal cord characterized by macrophage recruitment and/or activation in the peripheral root. This results in delayed microglia/macrophage reactivity in the spinal cord specifically in the dorsal funiculus (dorsal column/white matter) and dorsal horn after 7 days post DRI (Ramer et al., 2001; Liu et al., 1998). The onset of Wallerian degeneration and microglia/macrophage reactivity in the dorsal funiculus, and the resulting protracted myelin clearance, have been implicated as a secondary regenerative barrier impeding afferent reconnection (Ramer et al., 2001; McPhail et al., 2007).  A key feature of dorsal rhizotomy is the robust sprouting and/or reorganization of the sensory  37  circuitry (Darian-Smith, 2004; Ramer et al., 2004; MacDermid et al., 2004; Scott et al., 2005). Among the axon populations where active reorganization takes place include intact adjacent afferents (McNeill et al., 1990, 1991; Darian-Smith, 2004), and uninjured descending monaminergic axons (Kinkead et al., 1998; Ramer et al., 2004, 2007; MacDermid et al., 2004; Scott et al., 2005). Functionally, the sprouting of intact mechanosensory afferents from adjacent roots was previously shown to restore peripheral cutaneous and mechanical sensation (DarianSmith and Brown, 2000). Similarly, studies by the Ramer group also suggested that the sprouting of adjacent intact mechanosensory afferents contribute to the recovery in low-threshold mechanosensation following C7/C8 DRI (Ramer et al., 2007). In contrast, the sprouting of intact nociceptive afferents may result in aberrant nociceptive transmission (Eschenfelder et al., 2000). The sprouting of descending monoaminergic projections on the other hand is thought to modulate nociceptive input to dorsal horn projection neurons, given the role of monoaminergic projections in regulating pain processing (Millan, 2002). Studies by the Ramer group further demonstrated that accelerated recovery from cold pain could be achieved by increasing monaminergic input to deafferented dorsal horn neuron, concomitant to serotonergic axon sprouting (Ramer et al., 2004).  1.5.5. Manipulations to Enhance Axon Plasticity and Sensory Recovery Following DRI Axon sprouting following DRI is attenuated by the expression of inhibitory CSPGs and myelin associated inhibitory proteins in the adult central nervous system (MacDermid et al., 2004; Cafferty et al., 2008). As described earlier, removal or neutralization of CSPG and myelin associated inhibitory proteins enhanced axon sprouting in the dorsal horn, which is associated with sensory recovery (MacDermid et al., 2004; Cafferty et al., 2008). Specifically, studies by the Ramer group demonstrated that neutralizing the effects of myelin on axon sprouting using the  38  soluble nogo receptor (NgR) resulted in robust sprouting of intact adjacent primary afferents and descending monoaminergic interneurons in the dorsal horn. Likewise, studies by the McMahon group have shown that removal of CSPGs with the spinal infusion of the enzyme chondroitinase ABC (ChABC) stimulated robust axon sprouting following DRI. Using the dorsal root injury model that injures specifically the 5th, 6th and 8th cervical roots (C5, C6, C8) and the 1st thoracic root (T1), and leaves the 7th cervical root (C7) intact, they elegantly demonstrated that ChABC greatly enhanced the sprouting of intact C7 afferents into the deafferented dorsal horn (Cafferty et al., 2008). Further behavioural analysis was able to correlate the sprouting of C7 afferents with the maintenance of low-threshold mechanosensation that was lost following dorsal rhizotomy (Cafferty et al., 2008).  Overcoming the effects of the abovementioned CNS inhibitory molecules using classical neurotrophins such as neurotrophin-3 (NT-3) has also resulted in the successful regeneration of the injured root across the DREZ (Ramer et al., 2001). Electrophysiological studies have revealed that these regenerated afferents are indeed functional, and are likely to restore sensory deficits that have arisen from DRI (Ramer et al., 2001). Intriguingly, later studies revealed that antagonizing the NT-3 receptor TrkB appeared to enhance the sprouting of intact afferents, which was correlated with the apparent accelerated recovery in mechanosensation (Ramer et al., 2007). Overall, these studies demonstrate that sensory recovery relies not only on afferent regeneration, but also on the sprouting of intact afferents.  Lastly, the use of Rho kinase (ROCK) inhibitor to enhance the axon sprouting and/or reorganization following DRI has also been reported to result in remarkable improvements in sensory function (Ramer et al., 2004). As mentioned earlier, the RhoA/ROCK pathway is  39  convergently activated by CSPGs and myelin associated inhibitors (such as NogoA), and inhibits axon outgrowth and/or sprouting by triggering growth cone collapse (Schweigreiter et al., 2004; McKerracher & Winton, 2002). Studies by the Ramer group have shown that spinal infusion of the ROCK inhibitor following C7/C8 DRI can accelerate the sprouting of descending monoaminergic projections (Ramer et al., 2004). Moreover, the intraspinal sprouting can be correlated with the accelerated recovery from cold hyperalgesia (Ramer et al., 2004). It can therefore be speculated that manipulations of the neuronal growth cone may also be used to stimulate axon sprouting and functional recovery following dorsal rhizotomy.  1.6. EXPERIMENTAL OBJECTIVES Although current strategies are focused on manipulating the inhibitory environment in the astroglial scar to stimulate axon regeneration in the CNS (MacDermid et al., 2004; Cafferty et al., 2008; Bradbury et al., 2002; Li et al., 2004), few strategies are focused on intrinsically manipulating the neuronal growth cone to stimulate axon repair (Chan et al., 2005; Ramer et al., 2004). With few compounds available to manipulate the neuronal growth cone, the purpose of my research was to identify and characterize novel compounds that can promote growth cone motility. The thesis is divided into 3 key aims: 1) To develop a high-throughput screen to rapidly identify bioactive compounds that can stimulate growth cone motility and hence neurite outgrowth. 2) To evaluate neurite/axon outgrowth-promoting effects of a purified candidate compound, DKP101516, using in vitro and in vivo models of regeneration. 3) To determine whether axon plasticity mediated by DKP101516 following C7/C8 dorsal rhizotomy would result in sensory improvements.  40  Overall, these studies would validate whether manipulations of the neuronal growth cone can stimulate functional axon regeneration and/or sprouting in the injured central nervous system. 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NGF-Induced Axon Growth Is Mediated by Localized Inactivation of GSK-3β and Functions of the Microtubule Plus End Binding Protein APC. Neuron. 42, 897–912. 231. Zheng, B., Ho, C., Li, S., Keirstead, H., Steward, O., Tessier-Lavigne, M. 2003. Lack of enhanced spinal regeneration in Nogo-deficient mice. Neuron 38, 213-224.  59  CHAPTER 2  Developing a High-Throughput Screen 2  2  A version of this chapter has been published. Wong J.W., Brastianos, H.C., Andersen, R.J., O’Connor, T.P. (2008) A High-throughput Screen to Identify Novel Compounds to Promote Neurite Outgrowth. J Neurosci. Methods. 169, 34-42.  60  2.1. INTRODUCTION Following spinal cord trauma, numerous inhibitory molecules are expressed at the injury site and prevent axon regeneration. These inhibitors include chondroitin sulfate proteoglycan (CSPG) and myelin associated inhibitory proteins (MAIPs) that inhibit axon regeneration by activating RhoA (Niederost et al., 2002; Monnier et al., 2003). Other inhibitors include a number of repulsive guidance molecules that are expressed in the glial scar (Silver & Miller, 2004; Yiu & He 2006). Interventions to treat spinal cord injury include both extrinsic and intrinsic manipulations. Extrinsic manipulations include the removal of inhibitory molecules present in the glial scar using various enzymes and blocking peptides (Bradbury et al., 2002; Yang et al., 2006; Li et al., 2004; MacDermid et al., 2006). Intrinsic interventions involve the manipulation of signaling pathways in the neuronal cell body and/or growth cone. For example, neurotrophic factors are often used to activate Trk signaling pathways that promote neuronal survival and axon regeneration (Ramer et al., 2001; Kobayashi et al., 1997; Hiebert et al., 2002). Other more specific methods include blocking signaling pathways in the neuronal growth cone that act downstream of inhibitory molecules such as the RhoA kinase (Chan et al., 2005) and PKC pathway (Sivasankaran et al., 2004).  While most intrinsic interventions described above are focused on blocking signaling pathways that act downstream of inhibitory molecules, there are few interventions that aim to enhance axon outgrowth by stimulating growth cone motility. Indeed, one of the major obstacles inhibiting axon regeneration is the reduced intrinsic capacity of mature axons to grow (Kobayashi et al., 1997). Treatments that enhance axon outgrowth may thus be useful to improve the success of axon regeneration. To identify novel compounds that can stimulate neurite  61  outgrowth, we have developed a high-throughput screen to rapidly identify natural compounds that can promote neurite outgrowth.  A conventional method for identifying novel compounds that can enhance axon outgrowth is to test the effects of these compounds on primary neuronal cultures. One of the most popular and well-characterized systems for neurite outgrowth assays is the chick dorsal root ganglion (DRG) explant. Although this model is often feasible for testing a small sample of drugs, the low efficiency of this type of screen due to laborious preparation and low throughput limits the likelihood of identifying a positive candidate. To increase the probability of finding neurite outgrowth-promoting compounds, we devised a high-throughput screen using the motile cell as a general model for the neuronal growth cone, which is a key motile structure at the tip of growing axons and/or neurite through which outgrowth takes place (Dickson, 2001). The rationale of this approach is that many of the basic mechanisms underlying growth cone motility and the consequent axon outgrowth are also conserved in the motile cell. As discussed previously in our introduction, both systems share a dynamic leading edge consisting of filopodial and lamellipodial protrusions, and a trailing edge where F-actin meshworks and bundles are disassembled (Dickson, 2001). Furthermore, similar signaling pathways such as the Rho family GTPases are used in both systems to regulate the balance of F-actin assembly and disassembly (Dickson, 2001). With these similarities in mind, we designed a primary high-throughput wound healing assay and a secondary neurite outgrowth assay to rapidly identify novel compounds that can promote neurite outgrowth. The rapid screening of the bioactive marine extract library using high-throughput wound healing assay has lead to the identification of a marine compound DKP101516 (JH) that not only promotes cell migration, but also robust neurite outgrowth in dorsal root ganglion (DRG) neurons cultured on permissive and inhibitory substrates.  62  2.2 METHODOLOGY 2.2.1. Cell Cultures HEK 293 cells were cultured to confluency in 100mm cell culture plates (Falcon) in DMEM supplemented with 5% FBS (Sigma). Once confluent, the cells were mechanically sheared off the plate and collected in fresh medium. The cells were then plated in 96 well plates (Falcon) at a density of 100,000 cells per well, and incubated at 37 degrees Celsius in 5% CO2 for 24 hours to allow the cells to reach confluency.  2.2.2. High-throughput Migration Assay The high-throughput migration assay was performed with a Biogrid robot, which consists of a 96-pin tool, and designated racks where 96 well plates containing the extracts (source) and the cells (destination) can be loaded. Confluent 96 well plates were first loaded into the ‘destination’ racks (Biogrid), and were subjected to automated scratching and extract transfer procedures. For the automated scratching procedure, the following settings were employed. Briefly, the 0.7mm 96-pin tool is set to insert into the destination plate and to stop at the height of 0mm where the pins touch the bottom of the plate. Upon contact with the plate, the tool wiggles for 5 sec to make a scratch in the confluent layer of cells. Immediately following the scratch, 96 well plates containing an array of marine extracts were loaded into the ‘source’ racks. To add extracts directly from the library (source) to the cells (destination), the 0.7mm 96 pin tool picks up 30microliter droplets per pin of extract from each well of the source plate, and transfers the droplet into the corresponding wells in the destination plate. At the destination plate, the pin stops at the height of 3mm and wiggles for 5 seconds to gently stir in the extracts into the cell culture medium. Immediately after adding the extracts, the cells were incubated for 18 hours at 37 ºC in  63  5% CO2 to allow cell reinvasion back into the scratch. The cells were then fixed in 3.7% formaldehyde and imaged using an inverted microscope.  The wells were screened manually to identify positive extracts that promote cell migration. In the future, this step could be automated and done with relative ease. Based on preliminary experiments, we found that HEK 293 cells fail to completely reinvade into the scratch area following a time period of 18 hours. This short period of time is not compatible for multiple cell division (~24 hour per division) required to cover the scratch, suggesting that our results are unlikely to be affected by cell proliferation. We identified positive extracts as those that can drive complete cell reinvasion into the scratch within the 18-hour period. To ensure consistency in our results, the screens for each plate of extracts were repeated at least 3 times.  2.2.3. DRG Explants Dorsal root ganglia were isolated from embryonic day 8 and 14 chick embryos, and cultured in DMEM (Sigma Aldrich; Oakville, ON) supplemented with 10% FBS and 50ng/mL NGF (Invitrogen; Burlington, ON). The explants were cultured on glass coverslips coated with either poly-L-lysine (Sigma Aldrich; Oakville, ON), CSPG (Chemicon; Temecula, CA) or myelin substrates. Specifically, E8 DRG explants were cultured on 100µg/mL poly-L-lysine (PLL), or CSPG substrate consisting of a mixture of 4µg/mL CSPG and 20µg/mL PLL. Because neurons do not respond to myelin until embryonic day 13, we cultured E14 chick DRG explants on myelin substrates consisting of a mixture of 6µg/mL myelin (isolated from mouse brains as previously described by Norton & Poduslo, 1973) and 20µg/mL PLL. Following 2 hours of incubation to allow the explants to stick onto the coverslips, the explants were treated with various positive extracts and incubated for an additional 24 hours.  64  2.2.4. Dissociated DRG Neuronal Cultures Dissociated dorsal root ganglion neurons were isolated from E14 chick embryos. Briefly, dorsal root ganglia (DRGs) from the lumbar-sacral region of the spinal cord were dissected out and the roots and meninges were removed. The DRGs were then dissociated in 0.25% trypsin and 0.05% collagenase in Dulbecco’s minimal essential medium (DMEM- Sigma Aldrich; Oakville, ON). The dissociated DRG neurons were collected in DMEM supplemented with B27, 5% FBS and 50ng/mL nerve growth factor (NGF) (Invitrogen; Burlington, ON), 1mM glutamine and 0.3% glucose (Sigma Aldrich; Oakville ON). The cell suspension was filtered through a 1-part percoll gradient (30% percoll), and pure DRG neurons were collected at the bottom of the gradient. DRG neurons were then seeded at a density of 12,000 cells per well in 24 well plates containing 12mm glass coverslips coated with 100µg/mL PDL, or myelin and CSPG substrates described above. Serum was withdrawn from the cultures after 4 hours of incubation. After an additional 30 minutes on incubation, DRG neurons were treated with either vehicle or positive extracts and incubated for an additional 48 hrs.  2.2.5. Immunocytochemistry Explants were initially fixed in 3.7% formaldehyde for 15 minutes, washed 3X with PBS, and permeabilized with PBS + 0.1% Triton X-100. Explants were then labeled with 1:500 rabbit antineurofilament antibody or 1:500 rabbit anti-tubulin III (TUJ-1) overnight at 4 ºC. Following incubation with primary antibody, the explants were washed 3X with PBS, and incubated with the secondary antibody cy3 anti-rabbit IgG (1:500; Jackson Immunoresearch, West Grove, PA) for 1 hour at room temperature. After incubation with the secondary antibody, the explants were washed again with PBS and mounted on glass slides.  65  2.2.6. Measuring Neurite Outgrowth DRG explants were imaged with an inverted fluorescent microscope (Nikon; Mississauga, ON). Images were taken with a CCD digital camera (Princeton Instruments; Trenton, NJ), and processed with Metaview imaging software (Molecular Devices; Sunnyvale, CA). The line scan program in the Metaview software was used to measure the length of at least 100 neurites per explant. Measurements were from 3 different explants for each treatment. For dissociated DRG neurons, the same software was used to measure the length of up to 3 longest neurite per neuron. A total of 20 neurons per treatment group were measured. Average neurite length ± SEM from vehicle and DKP101516 treated groups was then compared. One way ANOVA was performed using SPSS software (SPSS; Chicago, IL), where p<0.01 indicates statistical significance.  2.2.7. Purification of a Positive Compound DKP101516 DKP101516 was extracted from a laboratory culture of a bacterium isolated from a marine sediment sample collected in Dominica. It was originally grown on M1 agar and subsequent pans were also made of this agar. To make M1 agar, 10g of soluble starch, 2g of bacto peptone and 18g of agar were immersed in 1L of sterile seawater (30g/L NaCl in distilled water) and autoclaved. The autoclaved agar was dispensed into large stainless steel pans at 400 mL per pan and was subsequently incubated for 7 days before harvest. The cells and the agar were freeze dried for extraction. Twenty pans of the freeze dried bacteria were extracted five times with methanol (5 X 1.5L). The methanol extracts were combined and reduced in vacuo to give a golden brown solid (6g). The crude extract was then dissolved in 500 mL of a 90% methanol/10% water mixture and this was then followed by a partition with hexanes (3 X 200mL). Fractions obtained from hexane partitioning were tested for neurite outgrowth  66  promoting activity in DRG neurons to identify the active methanol/water fraction (800mg), which was then subjected to SephadexTM LH-20 size exclusion chromatography (eluent: 100% methanol). Again, fractions from the LH-20 column were systematically tested for neurite outgrowth promoting activity in DRG explants, and an active fraction of 122.3 mg was identified. This was followed by further purification on a gradient reverse phase Sep PakTM (H2O to Methanol) where the active fraction eluted after 60% water/40% methanol (7.6 mg). The crude brown solid was finally purified with reverse phase HPLC (eluent: 1;1 water) yielded a pure DKP101516.  2.3. RESULTS 2.3.1. Candidate Extracts Identified in the High-throughput Migration Assay The high-throughput migration assay exploits the similarity between growth cone migration and cell migration to screen for extracts that may promote neurite outgrowth in DRG explants. Briefly, HEK293 cells were cultured in 96 well plates and grown to confluency. A robot was used to make a uniform scratch in the middle of each well, and extracts from a marine sponge and microbial culture extract library was added to individual wells. After 18 hours of incubation, the cells were fixed and screened manually for candidate extracts that increased cell motility and wound healing. Within the period of 18 hours, control cells failed to completely reinvade into the scratch, while cells treated with a positive candidate completely filled in the scratch (Fig. 2.1). Of the first 2500 extracts screened using this approach, we have identified 41 candidate extracts that promote cell migration (Table 2.1). The screen also revealed 49 ‘negative’ extracts with biological activity detrimental to cell migration. Of these 49 extract, 7 extracts significantly decreased the rate of cell migration by reducing leading edge extension, resulting in a significantly larger scratch relative to control (Table 2.1; n=7, Two-tailed Student’s t-test,  67  p<0.01). The remaining 41 extracts promote the typical hallmarks of cell death (denoted in italics in Table 2.1) in our cells such as cell rounding and detachment (Fig. 2.1b).  2.3.2. Secondary Screen on DRG Neurite Outgrowth We picked 5 candidate extracts of the 41 candidate positives identified in the high-throughput screen and tested them on embryonic day 8 (E8) chick dorsal root ganglion (DRG) explant cultures. Of the 5 extracts tested, we found three extracts (2 examples: AAC-A6 and AAE-B2) that display robust neurite outgrowth promoting activity on a permissive poly-L-lysine substrate (Fig. 2.2.a). There were also extracts (AAF-C11, AAG-C11) that did not significantly enhance neurite outgrowth (data not shown). Neurite length measurements reveal that DRG explants treated with positive extracts extend longer neurites compared to vehicle treated explants (Fig. 2.2.b). When these extracts were further tested on DRG explants cultured on inhibitory CSPG substrate, they promoted substantial neurite outgrowth (Fig. 2.3.a). Quantification of neurite length revealed that while vehicle treated explants failed to extend neurites, explants treated with the above three positives extended neurites to up to 160 microns on CSPG (Fig. 2.3.b). To further demonstrate that the positive extracts have potential to promote neurite regeneration, we next asked whether our extracts could promote neurite outgrowth even in the presence of inhibitors present in a spinal cord lesion, such as the myelin associated inhibitors. We therefore cultured chick DRG explants on myelin  68  Figure 2.1. High-throughput cell migration assay. (A) Schematic of the high-throughput screen.  The screen is an automated wound-healing assay on 96-well plates using the specialized 96-pin tool Biogrid robot. (B) The screen has identified a number of extracts with various biological activities. Some extracts promote a robust increase in the rate of cell migration, as indicated by the complete reinvasion of cells back into the scratch (positive). Other extracts either inhibit cell migration (negative) or promote cell death.  69  70  Table 2.1. Bioactive extracts detected from the high-throughput wound-healing assay.  Library Positive extracts  Negative extracts  AAC  A6, A9, D4, E1, E3, E4, D10  AAD  C1, B2, E2, C6, A7, C8, D9, D11, C12, A10  AAE  B2, C4, D9, D10  AAF  B2, C11, B11  B1, C2  AAG  E1, F6, C11  B1, B3, A4, D4, G4, B5, A6, C6, D7, D8, F8, F9, D10, D11, F1  AAH  E1, B8, C9  A1, B1, D3, C4, G4, C5, E7, E8, E9, B10, C10, B11, C11, G9, G11, G12  AAI  E1, B8, C9  A2, B4  AAJ  H1, C2, H4, D4  C4, D4, A6, B10, C10  AAK  B3, C3  AAL  B3  AAM  B8  AAO  E1  E6, E8  A9, B6, A7, B9, G9  Total positives: 41, total negatives: 49; italics denote cell death.  71  substrates. As expected, vehicle treated DRGs failed to extend neurites on the inhibitory myelin substrates (Fig. 2.3.). Among the 3 positive extracts tested, only AAC-A6 promoted neurite outgrowth to up to 100 µm on the inhibitory myelin substrate (Fig. 2.3.). Therefore, the candidate extract AAC-A6 identified in the high-throughput screen not only enhanced neurite outgrowth under permissive conditions, but also encouraged neurite outgrowth on inhibitory CSPG and myelin substrates.  2.3.3. Identification of a Novel Neurite Outgrowth Promoting Compound DKP101516 Of the three candidate extracts, only AAC-A6 promoted neurite outgrowth on both inhibitory CSPG and myelin substrates. Therefore, we identified a bioactive component of the candidate extract AAC-A6 by subjecting it to a series of purification procedures. Since overcoming myelin inhibition is the key aspect of AAC-A6 bioactivity in which we were interested, fractions obtained from each purification step were tested for their ability to promote neurite outgrowth on myelin substrates. Specifically, we systematically applied each fraction on E14 chick DRG explants cultured on myelin substrates and assessed outgrowth after 24 hours of incubation. Briefly, the initial purification of the crude extract AAC-A6 was subjected to Sephadex TM LH20 size exclusion chromatography to yield a fraction (soluble in 100% methanol) that demonstrated modest neurite outgrowth promoting activity on myelin substrate (Fig. 2.4.). Further purification with HPLC chromatography yielded a pure compound DKP101516 that demonstrates robust neurite outgrowth promoting activity in the presence of myelin (Fig. 2.4.).  2.3.4. DKP101516 Promotes DRG Neurite Outgrowth on Inhibitory Substrates Purified DKP101516 was synthesized, and its effects on neurite outgrowth were  72  Figure 2.2. Candidate extracts promote neurite outgrowth in DRG explants. (A) The positive extracts from the high-throughput screen are tested on chick dorsal root ganglion (DRG) explants. Neurofilament staining reveals that the positive extracts promoted robust increase in neurite outgrowth (Scale bar 100µm). (B) Bar graph representation of neurite outgrowth. AACA6 and AAE-B2 promoted a significant increase in neurite length relative to control (n=3 explants, 100 neurites measured per explant, *p<0.0001, One-way ANOVA, SNK post hoc analysis).  73  74  Figure 2.3. Candidate extracts stimulate neurite outgrowth on inhibitory substrates. (A) The positive extracts previously shown to enhance neurite outgrowth in DRG explants can also encourage neurite outgrowth on inhibitory substrates. Treatment with AAC-A6 and AAE-B2 promotes robust neurite outgrowth on CSPG. However, only AAC-A6 promotes significant neurite outgrowth on myelin substrate (Scale bar 100µm). (B) Quantification revealed that although AAC-A6 and AAE-B2 promoted neurite outgrowth on CSPG, only AAC-A6 promoted significant neurite outgrowth on myelin (n=3 explants, 100 neurites measured per explant, *p<0.001, One-way ANOVA, SNK post-hoc analysis).  75  76  characterized. Briefly, DRG explants were cultured on poly-L-lysine, CSPG, and myelin as described above. Compared to vehicle treatment, DKP101516 treatment significantly enhanced neurite outgrowth on a permissive poly-L-lysine substrate (Fig. 2.5b.). On myelin and CSPG substrates where neurite outgrowth was severely inhibited, DKP101516 treatment promotes robust neurite outgrowth to up to 250 microns (Fig. 2.5b.). Furthermore, the effects of DKP101516 alone appeared to be significantly more robust than the original extract AAC-A6 (Fig. 2.4.). Due to the variability in neurite outgrowth observed in DRG explant cultures, the neurite outgrowth-promoting effect of DKP101516 was further verified using dissociated DRG neurons.  Similar to the results obtained in DRG explants, DKP101516 stimulated significant neurite outgrowth in dissociated DRG neurons cultured on permissive (poly-L-lysine) and inhibitory (myelin and CSPG) substrates (Fig. 2.5a, c, n=20, One way ANOVA, p<0.001).  2.4. DISCUSSION Following CNS injury, numerous inhibitory molecules hinder the success of axon regeneration (Bradbury et al., 2002; Li et al., 2004; Silver & Miller, 2004; Yiu & He, 2006; MacDermid et al., 2005). To encourage axon regeneration, we have focused on identifying novel compounds that can enhance neurite outgrowth. A conventional approach would be to use chick DRG explants to screen for compounds that stimulate neurite outgrowth. As a well-characterized system, DRG explants are often used in screens because of the relative ease in isolating and culturing. However, because DRG explants are cultured manually, this system is laborious and not  77  Figure 2.4. Purification of neurite outgrowth promoting compound DKP101516 (JH). (A) AAC-A6 promotes robust neurite outgrowth in DRG explants cultured on inhibitory myelin substrates. To identify the active compound in this extract, AAC-A6 was purified using size exclusion chromatography. The active lipophilic fraction was further purified using HPLC chromatography to yield the pure compound, designated as JH, which was demonstrated to promote neurite outgrowth on inhibitory myelin substrates (Scale bar 100µm). (B) Quantification revealed that both AAC-A6 and JH can promote significant neurite outgrowth on myelin substrates (n=3 explants, 100 neurites measured per explant, *p<0.001, One-way ANOVA, SNK post-hoc analysis).  78  79  Figure 2.5. DKP10516 (JH) demonstrates neurite outgrowth effects on various substrates. (A) TUJ-1 staining revealed that JH stimulates neurite outgrowth in dissociated DRG neurons cultured on PLL, myelin and CSPG (Scale bar 50µm). The arrows indicate the three longest neurites per neuron that would have been measured for this neuron. (B) Quantification of neurite length in DRG explant cultures. Treatment with JH promoted significant increase in neurite length even on inhibitory substrates (n=2 explants, 100 neurites measured per explant, *p<0.0001, One-way ANOVA, SNK post-hoc analysis). (C) Quantification of neurite length in dissociated DRG neurons. Relative to control, treatment with JH promoted significantly increased neurite outgrowth relative to control on both permissive and inhibitory substrates (n=20, *p<0.001, One-way ANOVA, SNK post-hoc analysis).  80  81  amenable to high-throughput screening. Therefore, the use of DRG explants to screen for neurite-outgrowth-promoting compounds would limit the probability of finding positive candidates. One way to overcome this problem is to devise a high-throughput screen using an even simpler culture system such as mammalian cell lines. As previous studies have shown a conservation of mechanisms between neurite outgrowth and cell motility, we hypothesized that compounds stimulating cell motility are likely to promote neurite outgrowth as well. With this rationale, we established a high-throughput wound-healing assay using the highly motile mammalian cell line HEK293 as our primary assay to identify positive candidates. Unlike primary DRG cultures, HEK 293 cells are easy to culture, propagate and maintain over long periods of time. The robustness of HEK 293 cells is such that the wound-healing assay can be automated using robotics, therefore making high-throughput screening possible. In this study, we used the efficient high-throughput wound-healing assay as a high speed screening tool to complement the neurite outgrowth assay in a simple and robust two-part screening procedure. Our results reveal that the high-throughput assay is considerably more efficient than conventional wound healing assays in a number of ways. Furthermore, we demonstrate for the first time that a wound-healing assay can be used to successfully identify a compound that can stimulate neurite outgrowth even on inhibitory myelin and CSPG substrates.  A conventional use of the wound-healing assay is to screen for compounds that inhibit cell migration, and therefore impede metastasis of cancer cells. One good example is Motuporamine C, a compound demonstrated using wound-healing assays to inhibit breast cancer cell migration (McHardy et al., 2004; Williams et al., 2002). In our study, we demonstrate an unconventional use of the wound-healing assay to identify compounds that not only stimulate cell migration, but also growth cone motility and neurite outgrowth. A major caveat of this approach is that due to  82  cell type differences between non-neuronal cells and primary neurons, extracts that promote cell migration may not necessarily stimulate growth cone motility and neurite outgrowth. We have overcome this deficit by screening a large number of extracts using a high-throughput woundhealing assay, and therefore increase the probability of identifying extracts that stimulate both cell migration and growth cone motility. Compared to conventional wound healing assays that can test one extract at a time, our high-throughput screen can screen up to 600 extracts at a time. Furthermore, the Biogrid robot makes consistent scratches throughout, thus minimizing errors resulting from manual scratches used in conventional wound-healing assays. Finally, while a conventional wound-healing assay relies on manual cell counts, the consistency of the highthroughput wound-healing assay allows cell migration to be assessed in an all-or-none basis, where positive extracts are those that can trigger complete reinvasion of cells into the scratch area. This mode of screening can be automated, and can greatly increase the ease and efficiency of the screen. Indeed, our study provides the first example that our high-throughput woundhealing assay can identify multiple extracts that stimulate both cell migration and growth cone motility.  Although the high-throughput wound-healing assay can be used to identify compounds that stimulate neurite outgrowth, a major caveat is that the cell type used in this screen is nonneuronal. Therefore, positive candidates identified in the high-throughput screen have to be tested again on DRG explants to determine whether they indeed enhance neurite outgrowth. A more direct approach may be to use neuronal cell lines such as PC12 cells to screen for compounds that can stimulate neurite outgrowth. However, using exclusively neuronal cells in this screen would limit its application to identifying compounds that stimulate neurite outgrowth. In contrast, the high-throughput wound-healing assay can have many other uses in addition to  83  identifying compounds that stimulate cell migration or neurite outgrowth. Indeed, a number of studies have suggested that non-neuronal processes involved in spinal cord repair such as myelination, bridging and angiogenesis share a conserved cytoskeletal machinery that regulates cell motility (Liang et al., 2004; Thurnherr et al, 2006; Holtje et al., 2005; Soga et al., 2001). Therefore, it is possible that compounds promoting cell migration can also encourage the repair of spinal cord tissue through bridging, remyelination and angiogenesis. Given our success in identifying a neurite outgrowth-promoting compound using the high-throughput wound-healing assay, we suggest that this assay can also be useful in identifying compounds to encourage nonneuronal aspects of spinal cord repair.  84  2.5. BIBLIOGRAPHY 1.  Bradbury, E.J., Moon, L.D., Popat, R.J., King, V.R., Bennett, G.S., Patel, P.N., Fawcett, J.W., McMahon, S.B. 2002. Chondroitinase ABC promotes functional recovery after spinal cord injury. Nature. 416, 636-40.  2.  Chan, C.C., Khodarahmi, K., Liu, J., Sutherland, D., Oschipok, L.W., Steeves, J.D., Tetzlaff, W. 2005. Dose-dependent beneficial and detrimental effects of ROCK inhibitory Y27632 on axonal sprouting and functional recovery after rat spinal cord injury. Exp Neurol. 196(2), 352-64.  3.  Da Silva, J.S., Hasegawa, T., Miyagi, T., Dotti, C.G., Abad-Rodriguez, J. 2005. Asymmetric membrane ganglioside sialidase activity specifies axonal fate. Nat Neurosci. 8, 606-15.  4.  Dickson, B.J. 2001. Rho GTPases in growth cone guidance. Curr Opin Neurobiol. 11, 103-10..  5.  Hiebert, G.W., Khodarahmi, K., McGraw, J., Steeves, J.D., Tetzlaff, W. 2002. Brainderived neurotrophic factor applied to the motor cortex promotes sprouting of corticospinal fibers but not regeneration into a peripheral nerve transplant. J Neurosci Res. 69, 160-8.  6.  Holtje, M., Hoffmann, A., Hofmann, F., Mucke, C., Grosse, G., Van Rooijen, N., Kettenmann, H., Just, I., Ahnert-Hilger, G. 2005. Role of Rho GTPase in astrocyte morphology and migratory response during in vitro wound healing. J Neurochem. 95, 1237-48.  7.  Kobayashi, N.R., Fan, D.P., Giehl, K.M., Bedard, A.M., Wiegand, S.J., Tetzlaff, W. 1997. BDNF and NT-4/5 prevent atrophy of rat rubrospinal neurons after cervical axotomy, stimulate GAP-43 and Talpha1-tubulin mRNA expression, and promote axonal regeneration. J Neurosci. 17, 9583-95.  8.  Liang, X., Draghi, N.A., Resh, M.D. 2005. Signaling from integrins to Fyn to Rho family GTPases regulates morphologic differentiation of oligodendrocytes. J Neurosci. 24, 71409  9.  MacDermid, V.E., McPhail, L.T., Tsang, B., Rosenthal, A., Davies, A., Ramer, M.S., 2004. A soluble Nogo receptor differentially affects plasticity of spinally projecting axons. Eur. J. Neurosci. 20, 2567–2579.  10.  Monnier, P.P., Sierra, A., Schwab, J.M., Henke-Fahle, S., Mueller, B.K. 2003. The Rho/ROCK pathway mediates neurite growth-inhibitory activity associated with the chondroitin sulfate proteoglycans of the CNS glial scar. Mol Cell Neurosci 22, 319-30.  11.  Niederost, B., Oertle, T., Fritsche, J., McKinney, R.A., Bandtlow, C.E. 2002. Nogo-A and myelin-associated glycoprotein mediate neurite growth inhibition by antagonistic regulation of RhoA and Rac1.J Neurosci. 22, 10368-76. 85  12.  Norton, W.T., Poduslo, S.E. 1973. Myelination in rat brain: method of myelin isolation. J Neurochem 21, 749–57.  13.  Ramer, M.S., Bishop, T., Dockery, P., Mobarak, M.S., O'Leary, D., Fraher, J.P., Priestley, J.V., McMahon, S.B. 2002. Neurotrophin-3-mediated regeneration and recovery of proprioception following dorsal rhizotomy. Mol Cell Neurosci.19, 239-49.  14.  Silver, J., Miller, J.H. 2004. Regeneration beyond the glial scar. Nat Rev Neurosci. 5, 146156  15.  Sivasankaran, R., Pei, J., Wang, K.C., Zhang, Y.P., Shields, C.B., Xu, X.M., He, Z. 2004. PKC mediates inhibitory effects of myelin and chondroitin sulfate proteoglycans on axonal regeneration. Nat Neurosci. 7, 261-8.  16.  Soga, N., Namba, N., McAllister, S., Cornelius, L., Teitelbaum, S.L., Dowdy, S.F., Kawamura, J., Hruska, K.A. 2001. Rho family GTPases regulate VEGF-stimulated endothelial cell motility. Exp Cell Res. 269, 73-87  17.  Thurnherr, T., Benninger, Y., Wu, X., Chrostek, A., Krause, S.M., Nave, K.A., Franklin, R.J., Brakebusch, C., Suter, U., Relvas, J.B. 2006. Cdc42 and Rac1 signaling are both required for and act synergistically in the correct formation of myelin sheaths in the CNS. J Neurosci. 26, 10110-9.  18.  Yiu G, He Z. 2006. Glial inhibition of CNS axon regeneration. Nat Rev Neurosci.7, 61727.  86  CHAPTER 3  DKP101516 Stimulates Axon Sprouting in the Spinal Cord2  2  A version of this chapter has been published. Wong J.W., McPhail, L.T., Brastianos, H.C., Andersen, R.J., Ramer, M.S., O’Connor, T.P. (2008). A Novel Diketopiperazine Stimulate Sprouting of Spinally Projecting Axons. Exp Neurol. 214, 331-40.  87  3.1. INTRODUCTION In Chapter 2, we have identified a novel neurite outgrowth-promoting diketopiperazine (DKP101516) from marine extracts. Marine extracts have been previously shown to contain various biological activities. These activities include inhibition of tumor growth and metastasis. A good example is the marine extract derived compound Motuporamine C that demonstrated anti-tumor activities through inhibition of blood vessel growth into tumors, as well as modulation of tumor invasiveness (Roskelley et al., 2001). Intriguingly, studies in our lab have also demonstrated that Motuporamine C inhibits axon growth in dorsal root ganglion neurons by promoting growth cone collapse (To et al., 2006). Given the potent biological effects of marine extracts on neurons, we therefore screened a library containing 1000s of marine extracts for compounds that promote neurite outgrowth (Wong et al., 2008). Out of the 41 candidate extracts from the 2500 extracts that were screened, we have identified a neurite outgrowth promoting extract AAC-A6 originating from a marine bacterium (Wong et al., 2008). Further purification of this extract yielded a pure neurite outgrowth-promoting compound DKP101516.  Previous studies have demonstrated that DKPs not only play a role in microbial interactions in the marine environment (Li et al., 2006), but also demonstrate biological effects in mammalian systems (Zeng et al., 2005). This particular neurite outgrowth-promoting DKP, which we have named DKP101516, is a cyclic dipeptide consisting of the two amino acids phenylalanine and valine (Fig. 1D). Initial in vitro characterization reveals that DKP101516 is the first example of a DKP that can stimulate axon outgrowth and branching in cortical neurons in vitro, even in the presence of CSPG and MAIPs. We find that DKP101516 mediates its effects through the PI3K pathway, which is a known downstream target of neurotrophins that stimulates axon outgrowth and branching (Menager et al., 2004). Furthermore, our in vivo studies revealed that DKP101516 88  enhances the plasticity of CGRP positive primary afferents as well as descending monoaminergic projections in the spinal grey matter. Collectively, our data suggest a potential therapeutic use of DKP101516 to stimulate spinal cord repair through compensatory sprouting of intact axon projections.  3.2. METHODOLOGY 3.2.1. Synthesis of DKP101516 To obtain large amounts of DKP101516 necessary for extensive in vitro and in vivo analysis, DKP101516 was synthesized with the following protocol developed by Bull et al. (2001). Briefly, commercially available diketopiperazine (S)-(+)-3-isopropyl-2,5-piperazinedione was used as starting material. The diketopiperazine was reacted with benzyl bromide to produce a mixture of benzylated diketopiperazine. Isopropyl protective groups were used in the reaction to increase the yield of the benzylated diketopiperazine cyclo(S-Val-S-Phe) in the reaction mixture. Further chromatographic purification with a reversed phase Sep PakTM and reversed phase HPLC eluted the pure compound cyclo(S-Val-S-Phe) designated as DKP101516. The chemical structure of the final product was confirmed by 1H NMR spectroscopy.  3.2.2. Cell Cultures Cortical neurons were isolated from E14 chick embryos. Briefly, cortices were isolated and the meninges removed. The cortices were dissociated with 0.1% trypsin in Hank’s buffer salt solution (HBSS; Invitrogen; Burlington, ON) for 20 minutes. The dissociated cortical cells were collected in ice-cold HBSS and filtered through a 2-part percoll gradient (60%/15%). Cortical  89  Figure 3.1. DKP101516 stimulates neurite outgrowth and branching in a dose-dependent manner. (A) E14 chick cortical neurons were treated with various concentrations of DKP101516. Treatment with DKP101516 not only enhanced neurite outgrowth, but also stimulated neurite branching in a dose dependent manner. Immunostaining against beta-IIItubulin revealed that at the concentration of 16µM, DKP101516 stimulated an increase in the number of interstitial branches along the neurite, as indicated by the white arrows (see inset for high magnification). At 32µM, DKP101516 stimulated even more elaborate interstitial branching along the length of the neurite (branches indicated by white arrows). (B) Quantification of neurite length in 300 neurons from 15 individual experiments revealed that DKP101516 significantly increased neurite length (n=15, One-way ANOVA, p<0.0001). Comparisons between treatment groups reveal that DKP101516 stimulates neurite outgrowth in a dose dependent manner. The neurite outgrowth promoting effect of DKP101516 is observed to peak at 32 µΜ (n=15, One-way ANOVA, *p<0.05) (C) Quantification of number of branches per neurite length (every 300 microns) revealed that DKP101516 significantly enhanced neurite branching (n=15, One-way ANOVA, p<0.0001). Comparisons between treatment groups reveal that DKP101516 stimulates neurite branching in a dose dependent manner. The neurite branching effect of DKP101516 is observed to peak at 32 µΜ (n=15, One-way ANOVA, *p<0.05). (D) The chemical structure of the diketopiperazine DKP101516. DKP101516 consists of a cyclic dipeptide made up of the two amino acids phenylalanine and valine. The two amino acids are held together by two peptide bonds.  90  91  neurons at the 60%/15% interface were collected and washed 2X with ice-cold HBSS. Neurons were then suspended in Neurobasal (Invitrogen; Burlington, ON) supplemented with B27, 10% fetal bovine serum (FBS; Invitrogen; Burlington, ON), 1mM glutamine, 0.3% glucose, and 0.45% bicarbonate (Sigma Aldrich; Oakville, ON). The neuronal suspension was then seeded at a density of 12,000 cells per well in 24 well plates containing 12mm glass coverslips coated with poly-D-lysine (PDL; 100 µg/mL, Sigma Aldrich; Oakville, ON), myelin (6µg/mL myelin [isolated from mouse brains as previously described by Norton & Podusulo, 1973] in 20µg/mL PDL) or CSPG (8µg/mL CSPG mixture [Chemicon; Temecula, CA) in 20µg/mL PDL]. After 24 hours of incubation, serum was withdrawn from the cultures and the neurons were serum-starved for an additional 24 hours.  3.2.3. Immunocytochemistry Cell cultures were fixed in 3.7% formaldehyde for 20 minutes, and were washed 3 times with phosphate buffer saline (PBS) for 5 minutes each. The cells were then incubated overnight with the TUJ-1 antibody (host: mouse, 1:500, BioCan, Mississauga, ON) in the presence of 0.1% Triton X-100. Cells were again washed 3 times with PBS for 5 minutes prior to incubation with secondary fluorescent antibodies (1:500, Jackson Immunoresearch; West Grove, PA). After three more washes with PBS (5 minutes each), coverslips were mounted (1% propyl gallate in glycerol) on glass slides.  3.2.4. Outgrowth and Branching Assay To determine the effective dose of DKP101516, various concentrations of DKP101516 (040µM) were added to cortical neurons cultured on PDL. To further characterize the effects of DKP101516, the effective dosage of DKP101516, or DKP101516 combined with PI3K inhibitor 92  (PI3Ki) (LY294002; EMD, San Diego, CA) was added to cortical neurons cultured on various substrates (PDL, myelin and CSPG). For all of the above experiments, neurons were visualized by beta-III-tubulin immunostaining and digital images were taken at 20X magnification. Neurons with one neurite that exceeded the length of any other of its neurites were included in the quantification. The longest neurite of each neuron was digitally retraced by hand and the length was quantified using Image J (NIH). Branching was quantified by counting the number of branches for every 300 µm of neurite length. For determining the effective dose of DKP101516, a total of 300 neurons per treatment group were measured. For assessing whether DKP101516 can overcome the inhibitory effects of myelin and CSPG substrates, a total of 200 neurons per treatment were quantified. Finally, for determining whether PI3K inhibition could reduce the effects of DKP101516, total of 100 neurons per treatment group were quantified. All the above measurements are presented as average neurite length or branch number (± SEM).  3.2.5. Cell Extracts and Western Blotting 1X106 cortical neurons were treated with 2% DMSO (negative control), DKP101516 (treatment) or neurotrophin-3 (positive control) for 15 minutes. The cells were then lysed in ice-cold lysis buffer (150mM NaCl, 50mM Tris HCl, 5mM EDTA, 1% Triton X-100) containing 5mM sodium fluoride, 1mM sodium orthovanadate, and 20µg/mL protease inhibitor cocktail (Roche Diagnostics, Laval, QC). After 5 minutes of lysis, the cells were scraped off the plate, and the collected samples were centrifuged at 6000 rpm for 5 minutes at 4ºC. Protein samples collected from the supernatant were reduced in a loading buffer (60mM Tris-HCl, 2% SDS, 10% glycerol, 50mM DTT, 0.01% bromphenol blue) and denatured by boiling at 100ºC for 5 minutes. The samples were separated with SDS PAGE on an 8% polyacrylamide gel (BioRad; Hercules, CA), and the proteins were transferred onto a nitrocellulose membrane (BioRad, Hercules; CA). The  93  membrane was then processed with the primary rabbit antibodies against Erk, Akt, phospho-Erk, phospho-Akt (Cell Signaling Technology; Boston, MA) and tubulin (Chemicon; Temecula CA), followed by a secondary horse-radish peroxidase conjugated anti-rabbit antibody (Jackson Immunoresearch; West Grove, PA). Specific bands were detected using a chemiluminescent detection kit (BioRad; Hercules, CA).  3.2.6. Animal Surgery and Tissue Preparation All experiments were performed in accordance with Canadian Council on Animal Care guidelines, and approved by the University of British Columbia Animal Care Committee. Briefly, adult male Sprague-Dawley rats (n=14; 200-300g) were anaesthetized with ketamine hydrochloride (72mg/kg; Bimeda-MTC, Cambridge, ON, Canada) and xylazine hydrochloride (9.1mg/kg; Bayer Inc. Etobicoke, ON, Canada). A septuple cervical rhizotomy was performed to examine the effects of DKP101516 on axon regeneration and sprouting. The dura mater was opened to expose the left cervical dorsal roots (C4-T2) and the exposed roots were subjected to a complete crush injury (crushed three times, 10 s each, with fine No. 5 forceps). A pre-filled cannula was inserted through the atlanto-occipital membrane along the dorsal surface of the cord until the tip rested at mid-C6. The opposite end was attached to an osmotic minipump (Alzet; Cupertino, CA) containing either DKP101516 (2.5 mg/mL) or vehicle (50% DMSO, 50% PEG). The drug was delivered at a rate of 2.5µg per day. Five days post-surgery, the primary afferents were traced transganglionically with the injection of 0.5 µl of 1% CTB (B fragment of cholera toxin; List Biologic, Campbell, CA). Seven days post-surgery, the animals were killed with an overdose of chloral hydrate (1g/kg i.p.; Sigma Aldrich; Oakville, ON ) and perfused with 4% paraformaldehyde in 0.1M phosphate buffer. Their spinal cords were removed and post-fixed overnight. The tissue was then cryoprotected in 20% sucrose in 0.1M phosphate buffer for 24 h  94  at 4ºC, frozen at -80ºC, and cross-sectioned (16µm) on a cryostat.  3.2.7. Immunohistochemistry Spinal cord sections were immunohistochemically processed to visualize the following: tracer (CTB, host goat, 1:2000; Cedarlane; Burlington, ON), primary afferents (CGRP [calcitonin gene related peptide] host mouse, 1:2000; Chemicon; Temecula, CA), astrocytes (GFAP [glial fibrillary acid protein], host rabbit, 1:1000, Sigma; Neurocan, IF6 antibody, host mouse, 1:100, Developmental Studies Hybridoma Bank; Iowa City, IA), and descending serotonergic and monaminergic projections (SERT [serotonin transporter], host rabbit, 1:2000, Immunostar; TH [tyrosine hydroxylase], host sheep, 1:200, Chemicon; Temecula, CA). Briefly, sections were first blocked with 10% normal donkey serum in PBS for 20 minutes, followed by overnight incubation with primary antibody solutions (in PBS plus 0.1% Triton X-100). After three 15-min washes in PBS, secondary antibodies raised in donkey and conjugated with Cy3 or Alexa 488 (1:200; Chemicon; Temecula, CA) were applied for 2 hours. After another series of three 15-min washes, the slides were coverslipped and examined using a Zeiss (Jena, Germany) Axioplan II microscope. Digital images were captured using Northen Eclipse software (Empix Imaging; Mississauga, ON) via a digital camera (Qimaging; Surrey, BC).  3.2.8. Analysis of Axon Regeneration and Glial Scarring To assess the regeneration of primary afferents, sections were labeled for CTB antigen as described above. Five images per animal (n=5 per treatment group) were captured, and the number of CTB positive axon terminals that regenerated up to the dorsal root entry zone (DREZ) was counted manually. Due to the variability in CTB tracer uptake, the axon counts were confirmed with immunostaining for NF200, which labels 200kDa neurofilaments typically found  95  in large-calibre axons. The density of NF200 positive axons in the peripheral root was measured with the Image J software (NIH). The average measurements obtained from the vehicle and DKP101516 treated animals were used for comparison.  To analyze astrocyte reactivity, sections were labeled for GFAP as described above. Images of 5 randomly selected sections per animal (n=5 per treatment group) were captured and filtered, and the density of GFAP positive processes was quantified using Sigma Scan Pro (version 5.0; San Jose, CA) as previously described (MacDermid et al., 2004). Briefly, images were filtered using a Laplacian omnidirectional edge-detection filter to optimize the signal-to-noise ratio, and to correct for small variations in background staining across images. Resulting profiles of filtered images were selected with a threshold overlay to give all immunopositive pixels equal weight and brightness regardless of their brightness in the original images. Using the Sigma Scan tracing function, the density of GFAP positive astrocytic processes in the dorsal root entry zone (DREZ) was measured. Densitometric results were presented as means ± SEM.  To determine whether DKP101516 affected scar formation at the DREZ, two randomly selected sections per animal (n=5) were stained for neurocan, an inhibitory chondroitin sulfate proteoglycan. As before, the images were captured, and the filtered profiles were used to measure the density of neurocan positive processes in the CNS portion of the dorsal root. Densitometric results are presented as means ±SEM.  3.2.9. Analysis of Axon Sprouting The sprouting of intact primary afferents within the dorsal horn was quantified by measuring the density of CGRP positive axon terminals as a function of depth in the dorsal horn. Briefly, 5  96  randomly selected sections per animal (n=5 per treatment group) were captured and filtered as described above. The resulting threshold images were analyzed with Sigma Scan as follows. First, three 50 pixel wide strips were used to select the central third of the dorsal horn starting from the top of lamina I down to lamina V (at an approx depth of 700 µm). The intensity of CGRP-positive axon terminals threshold-selected within the 50 pixel wide strips were measured, and converted into density measurements (Ramer et al., 2007; Ramer et al., 2007; MacDermid et al., 2004). The average axon density in lamina I/II, III, IV and V was calculated for each animal, and plotted as means ± SEM.  The sprouting of descending monoaminergic projections into the dorsal horn was quantified by measuring the density of SERT and TH positive axon terminals as a function of depth in the dorsal horn. Briefly, 5 randomly selected sections per animal (n=7 per treatment group) were captured, filtered, and the resulting profiles of the filtered images were analyzed with Sigma Scan Pro (version 5.0) as described above. The average densities of SERT-positive and THpositive axon terminals in lamina I/II, III, IV, V were calculated for each animal, and were plotted as means ± SEM.  3.2.10. Statistics Data are reported as the mean ± SEM. All statistical analyses were performed by Sigma Stat (version 5.0). Student’s t-test was used to compare between control and treatment groups. For multiple comparisons between groups, one-way ANOVA was used. For all pair-wise comparisons between control and treatment groups, further analysis using the SNK post-hoc test was used. For all statistical outcomes, significance was set as p<0.05.  97  3.3. RESULTS 3.3.1. DKP101516 Promotes Neurite Outgrowth and Branching In our initial screen, DKP101516 demonstrated robust neurite outgrowth effects on chick dorsal root ganglion explants (Wong et al., 2008). In order to utilize this property of DKP101516 in spinal cord repair, we further characterized the effects of DKP101516 on CNS neurons. Using E14 chick cortical neurons, we assessed the effects of varying concentrations of DKP101516 on cortical neuron morphology. Specifically, cortical neurons cultured for two days in vitro were treated with DKP101516 for 2 days prior to fixation, followed by TUJ-1 immunostaining to label neurons (Fig. 3.1.). The length and morphology of the longest neurite were assessed. 300 neurons from 3 different experiments were included in quantification. Results revealed that DKP101516 significantly enhanced neurite outgrowth and branching (Figs. 3.1.B,C, n=15, p<0.001, One-way ANOVA). Comparisons between concentrations showing that DKP101516 stimulated neurite outgrowth and branching in a dose dependent manner (Figs. 3.1.B,C, n=15, *p<0.05, One-way ANOVA), which leveled off at the concentration of 32 µM.  3.3.2. DKP101516 Stimulates Neurite Outgrowth and Branching on Myelin and CSPG Substrates Following spinal cord injury, a spontaneous repair mechanism is inhibited by the endogenous expression of CSPG and MAIPs in the adult central nervous system. To determine whether DKP101516 stimulates axon repair in the adult CNS, it was of interest to determine whether DKP101516 can encourage neurite branching despite the presence of MAIP and CSPG. To examine this, E14 chick cortical neurons cultured on myelin and CSPG were treated with either vehicle (DMSO) or DKP101516. Neurons cultured on CSPG and myelin substrates typically had significantly shorter neurites, and fewer branches per neurite length compared to neurons  98  cultured on permissive substrates (Figs. 3.2.A-C, n=10, #p<0.05, one-way ANOVA, SNK post hoc analysis). Furthermore, DKP101516 treated neurons cultured on CSPG/myelin had significantly longer neurites and more branches per neurite length compared to vehicle treated neurons cultured CSPG/myelin (Figs. 3.2.A-C, n=10, *p<0.001, one-way ANOVA, SNK post hoc test). Intriguingly, DKP101516 appears to differentially overcome the inhibitory effects of MAIPs and CSPGs. In particular, DKP101516 completely overcame the inhibitory effects of MAIPs (Figs. 3.2., 3.3.). In contrast, DKP101516 only partially overcame the inhibitory effects of CSPGs on neurite outgrowth (Figs. 3.2., 3.3.). Based on the above results, we conclude that DKP101516 not only stimulates robust neurite outgrowth and branching, but also differentially overcomes the inhibitory effects of MAIPs and CSPGs.  3.3.3. DKP101516 Stimulates Neurite Outgrowth and Branching through the PI3K Pathway Similar to the effects of trophic factors ([Klimaschewski et al., 2004], and [Kobayshi et al., 1997]), DKP101516 stimulated robust neurite outgrowth and branching even on inhibitory substrates. We therefore asked whether DKP101516 mediates its effect by activating signaling effectors downstream of receptor tyrosine kinases, such as mitogen activated protein kinase (MAPK) and phosphotidylinositol-3- kinase (PI3K). To determine whether DKP101516 activates these pathways, we looked at the phosphorylation levels of MAPK and PI3K substrates extracellular receptor kinase (erk) and protein kinase B (PKB/Akt) respectively. Protein samples collected from cortical neurons previously treated with vehicle, DKP101516 or neurotrophin-3  99  Figure 3.2. DKP101516 overcomes the inhibitory effects of myelin and CSPG. (A) Cortical neurons were cultured on poly-D-lysine, and inhibitory substrates myelin and CSPG as indicated. Vehicle treated neurons cultured on poly-D-lysine extended neurites with average lengths >200 µm, while vehicle treated neurons cultured on inhibitory myelin and CSPG substrates have significantly shorter neurites (with average neurite length ~100 µm). In contrast, DKP101516 (32µM) treated neurons demonstrated enhanced neurite outgrowth on poly-D-lysine, and significantly increased neurite outgrowth on myelin and CSPG (Scale bar= 100 µm). (B) Quantification of neurite length obtained from 200 neurons from 10 individual experiments. The presence of myelin and CSPG significantly reduced the average neurite length of vehicle treated neurons (denoted with #, p<0.05). DKP101516 treatment resulted in a significant increase in neurite outgrowth relative to vehicle treated neurons on both permissive and inhibitory substrates (n=10, One-way ANOVA, *p<0.001, SNK post-hoc analysis). (C) Quantification of neurite branching by counting the number of branches per 300 µm of neurite length. The presence of myelin and CSPG significantly reduced the number of branches (denoted with #, P<0.05). Treatment with DKP101516 significantly increased the number of neurite branches (denoted with *) even in the presence of myelin and CSPG (n=300, p<0.001, one-way ANOVA, SNK post-hoc analysis).  100  101  were separated with SDS-PAGE, transferred, and probed for Akt, Erk, phospho-Akt and phospho-Erk. Compared to vehicle treatment, NT-3 stimulated robust phosphorylation of both Akt and Erk (Fig. 3.3.A). However, DKP101516 stimulated only phosphorylation of Akt but not Erk (Fig. 3.3.A), suggesting that DKP101516 activates the PI3K pathway.  To further test whether DKP101516 stimulates neurite outgrowth and branching through the PI3K pathway, we next asked whether PI3K inhibition would abolish the effects of DKP101516. We therefore applied DKP101516 alone, or together with the PI3K inhibitor (PI3Ki) to cortical neuron cultures. As expected, treatment with DKP101516 alone enhanced both neurite outgrowth and branching even in the presence of MAIPs and CSPGs (Fig. 3.3.B). However, the addition of PI3Ki completely abolished the effect of DKP101516 on neurite outgrowth and branching (Fig. 3.3.B). Quantification of both neurite length and branching revealed that PI3Ki significantly reduced the effects of DKP101516 on neurite outgrowth and branching, even in the presence of MAIPs and CSPGs (Figs. 3.3.C,D, n=5, *p<0.001, One way ANOVA, SNK posthoc analysis). The neurite length and branching observed in neurons treated with PI3Ki +DKP101516 was comparable to vehicle treated neurons (Figs. 3.3.C,D). Based on these results, we conclude that DKP101516 mediates its effects through the PI3K pathway.  3.3.4. DKP101516 Stimulates the Sprouting of Primary Afferents within the Injured Dorsal Root  Since DKP101516 can promote axon outgrowth on myelin and CSPG in vitro, we next examined whether DKP101516 could stimulate axon regeneration across an inhibitory glial scar. The inhibitory glial scar is best modeled in septuple dorsal rhizotomy, where the boundary of the glial scar can be clearly defined as the PNS/CNS interface at the dorsal root entry zone (DREZ). Briefly, 5 male Sprague-Dawley rats per treatment group were subjected to septuple dorsal 102  Figure 3.3. DKP101516 mediates its effects through the PI3K pathway. (A) Application of DKP101516 stimulated robust increase only in Akt phosphorylation (substrate for PI3K), but not Erk phosphorylation (substrate for MAPK). (B) Compared to vehicle treated neurons, DKP101516 (32µM) treated neurons cultured on PDL have markedly longer neurites and more branches. However, application of DKP101516 together with a PI3K inhibitor (PI3Ki [1µM LY294002]) reduced the neurite outgrowth and branching effects of DKP101516. (C) Quantification of neurite length of 100 neurons from 5 individual experiments. Compared to vehicle treatment, DKP101516 significantly increased neurite length even on myelin and CSPG substrates. However, application of DKP101516 with PI3Ki resulted in neurite lengths similar to control levels (n=5, *p<0.001, One way ANOVA, SNK post-hoc analysis). (D) Quantification of neurite branching by counting the number of branches per 300 µm of neurite length. Application of PI3Ki significantly reduced the effects of DKP101516 on neurite branching in permissive conditions, as well as in the presence of myelin and CSPG (n=5, *p<0.001, One way ANOVA, SNK post-hoc analysis).  103  104  rhizotomy in which roots C4-T2 were crushed completely (Fig. 3.4.a). Vehicle (DMSO) or DKP101516 was applied intrathecally for 7 days, during which primary afferents regenerated up to the PNS/CNS interface. Two days prior to sacrifice, the injured roots were traced transganglionically with a CTB tracer. Following sacrifice, cross-sections of the spinal cord were taken from C6-T1. In both vehicle treated and DKP101516 treated animals, CTB traced axons regenerated up to the DREZ (Fig. 3.4.b). Both treatment groups showed enlarged axon terminals at the DREZ, indicating that the axons failed to regenerate into the spinal cord (Fig. 3.4.b). However, DKP101516 treated animals had significantly more CTB traced axons in the peripheral compartment of the dorsal root compared to vehicle treated animals (Fig. 3.4.b,c, n=5, p<0.001, Student’s t-test). Further analysis of NF200 immunoreactivity, which identifies the axonal neurofilaments of large diameter axons, revealed that DKP101516 significantly increased the density of primary afferents in the injured root (Fig. 3.4.D,E, n=5, p<0.001, Student’s t-test). Therefore DKP101516 can enhance the sprouting of primary afferents in the peripheral root, however it failed to stimulate regeneration across the DREZ.  3.3.5. DKP101516 Treatment Stimulates Astrocyte Reactivity and Neurocan Expression A major barrier hindering axon regeneration is the formation of the glial scar following injury. Since DKP101516 can enhance primary afferent regeneration up to but not beyond the DREZ, we asked whether DKP101516 may be hindering axon penetration across the DREZ by stimulating astrogliosis. To test this hypothesis, C6-C7 spinal cord cross-sections of animals in control and treatment groups were processed with GFAP immunostaining to assess the degree of astrocyte reactivity in the DREZ. On both the injured (ipsilateral) and uninjured (contralateral) side of the spinal cord, DKP101516 treatment promoted a marked increase in GFAP staining intensity in the DREZ compared to vehicle treatment (Fig. 3.5.A). Quantification of GFAP  105  Figure 3.4. DKP101516 stimulates afferent sprouting within the peripheral root. (A) Schematic of septuple dorsal rhizotomy. Following complete crush injuries of C4-T2 dorsal roots as indicated, DKP101516 or vehicle was applied intrathecally for a 7-day period. (B) Injured afferents are traced transganglionically with cholera toxin B (CTB). In both vehicle and DKP101516 treated animals, regenerating CTB afferents are observed to terminate at the dorsal root entry zone. Compared to vehicle treatment, DKP10516 treatment appears to markedly increase the number of CTB traced axons in the peripheral root. (C) Axon counts revealed that the average number of CTB traced axon terminals is significantly greater in DKP10516 treated animals compared to vehicle treated animals (n=5, *p<0.05, One-way ANOVA). (D) Labelling with an axonal marker NF200 revealed that compared to vehicle treatment, DKP101516 treatment markedly increased the sprouting of injured afferents within the peripheral root. (E) Densitometric measurements of NF200 axons in the peripheral revealed that DKP101516 treatment stimulated significantly greater axon sprouting in the peripheral root compared to vehicle treatment (n=5, *p<0.05, One-way ANOVA). Dottedl ines outline dorsal roots and spinal cord.  106  107  positive process density revealed that DKP101516 promoted a significant increase in GFAP expression in the DREZ on both the injured and uninjured sides of the spinal cord (Fig. 3.5.B, n=5, p<0.05, One way ANOVA). Therefore, DKP101516 enhanced astrocyte reactivity in a manner that was independent of injury.  Previous studies have reported that astrocyte reactivity detected by increased GFAP expression may not necessarily be inhibitory to axon regeneration. Indeed, reactive (proliferative) astrocytes may also promote spinal cord repair by bridging the lesion site (Davies et al., 2006; Pancalet et al., 2006). Therefore, to determine whether DKP101516 mediated astrogliosis would hinder axon regeneration, we examined whether the astrocytes express inhibitory molecules such as the CSPG neurocan. Previous studies have shown that neurocan is expressed in the glial scar following injury, and is inhibitory to neurite outgrowth in dorsal root ganglion explants (Sango et al., 2003). To determine whether the failure in primary afferent regeneration is the result of neurocan expression in the DREZ, C6-C7 spinal cross-sections of animals in both control and treatment groups were immunostained for neurocan. Compared to vehicle treated animals, DKP101516 treated animals had more intense neurocan staining in the DREZ only on the injured (ipsilateral) side of spinal cord (Fig. 3.5.C). Densitometric measurements of neurocan-positive processes in the DREZ revealed that relative to vehicle treatment, DKP101516 treatment significantly enhanced neurocan expression only on the injured side of the spinal cord (n=5, p<0.05, One way ANOVA). The data suggest that although DKP101516 can increase astrocyte reactivity independent of injury, DKP101516 promotes neurocan expression through an injurydependent mechanism.  To determine whether DKP101516 induced neurocan overexpression in the DREZ is responsible  108  Figure 3.5. DKP101516 enhances astrogliosis and neurocan expression in the DREZ. (A) Compared to vehicle treatment, DKP101516 markedly increased the density of GFAP immunoreactivity in the dorsal root entry zone both ipsilateral and contralateral to the DRI. (B) Densitometric measurements demonstrate DKP101516 mediated increase in GFAP immunoreactivity is significantly greater compared to vehicle treatment (n=5, *p<0.05, One-way ANOVA). (C) Neurocan labeling revealed that DKP101516 markedly increased neurocan expression in the DREZ compared to vehicle treated animals. The effects of DKP101516 on neurocan expression is only restricted to DREZ ipsilateral to the DRI. (D) Densitometric measurements revealed that DKP101516 treatment significantly increases neurocan immunoreactivity only in the DREZ ispilateral to DRI (n=5, *p<0.05, One-way ANOVA). Dotted lines outline the dorsal roots and spinal cord.  109  110  for inhibiting primary afferent regeneration, we tested whether DKP101516 can stimulate DRG neurons in culture to extend neurites on various concentrations of neurocan substrates. Our results revealed that DKP101516 demonstrates limited potential in overcoming the inhibitory effects of neurocan (Fig. 3.6). Specifically, DKP was ineffective in enhancing neurite outgrowth on concentrations of neurocan above 1µg/mL. Therefore, DKP101516 mediated overexpression of neurocan in the DREZ in vivo is likely to hinder the ability of DKP101516 to stimulate primary afferent regeneration across the glial scar.  3.3.6. DKP101516 Stimulates the Sprouting of Spinally Projecting Axons Our in vitro characterization has demonstrated that DKP101516 can stimulate robust outgrowth and branching even in the presence of myelin associated inhibitors and CSPG (Figs. 3.1., 3.2.). Therefore, we reasoned that DKP101516 is likely to stimulate sprouting of intact axonal projections in the spinal cord. To assess the sprouting of spinally projecting axons in vivo, we used a well-characterized septuple dorsal rhizotomy model (Fig. 3.4.A). In this model, C4-T2 dorsal roots were subjected to complete crush injuries. These injuries typically result in changes to the nociceptive circuitry, which is made up by the CGRP positive nociceptive afferents through which pain is transmitted to the CNS, as well as descending SERT positive and TH positive supraspinal axons through which pain signals are modulated at the level of the spinal cord. Following septuple dorsal rhizotomy, CGRP positive axons were largely depleted from the dorsal horn ipsilateral to the injury (MacDermid et al., 2004). The remaining CGRP positive terminals that innervate the lateral dorsal horn were spared nociceptive afferents from intact dorsal roots both rostral and caudal to the crushed roots (Hampton et al., 2007). Septuple dorsal rhizotomy also resulted in the spontaneous sprouting of descending SERT and TH positive supraspinal projections (Ramer et al., 2005). In the present study, DKP101516 intrathecal  111  Figure 3.6. DKP101516 mediated neurite outgrowth is significantly reduced by high concentrations of neurocan. (A) Dissociated DRG neurons were cultured on permissive PDL, or on various concentrations of inhibitory neurocan (0.1, 1, 10 mg/mL). Compared to vehicle treatment, DKP101516 stimulated a robust increase in neurite outgrowth on permissive PDL. However, the ability of DKP101516 to stimulate neurite outgrowth on neurocan progressively decreased with increasing concentrations of neurocan. (B) Quantification of neurite length. DKP101516 stimulated significant increase in neurite outgrowth only on permissive PDL or on neurocan substrates at concentrations below 0.1 mg/mL (n=20, *p<0.01, One-way ANOVA, SNK post-hoc analysis). (C) Quantification of the percentage of neurons extending neurites. DKP101516 stimulated significant increase in the proportion of neurons extending neurites only on permissive PDL (n=10, *p<0.01, One-way ANOVA, SNK post-hoc analysis).  112  113  treatment enhanced the sprouting of axon projections that make up the nociceptive circuitry.  1) CGRP Positive Afferents Previous studies have shown that sequestration of myelin-associated inhibitors in the CNS with soluble nogo receptor (sNgR) resulted in a small but significant increase in the density of CGRPpositive axons in the lateral dorsal horn (MacDermid et al., 2004). Since DKP101516 can overcome the inhibitory effects of MAIPs in vitro, we therefore asked whether DKP101516 could enhance the sprouting response of these small diameter afferents. Indeed, we found that relative to vehicle treatment, DKP101516 treatment markedly increased the density of CGRPpositive axon terminals specifically in the lateral dorsal horn (Fig. 3.7.A). Densitometric measurements revealed that relative to vehicle treated animals, DKP101516 treated animals had significantly greater CGRP-positive axon density in the lateral dorsal horn (n=5, p<0.01, Oneway ANOVA). Further SNK post-hoc analysis revealed that DKP101516 significantly increases CGRP-positive axon density in laminae I-II of the lateral dorsal horn (Fig. 3.7.B).  2) SERT and TH positive Projections- Ipsilateral to Injury Previous studies by McDermid et al (2004) have shown that sequestration of myelin associated inhibitors sNgR enhanced the sprouting of descending SERT and TH positive supraspinal projections. Since DKP101516 can overcome the inhibitory effects of myelin associated inhibitors in vitro, we next asked whether DKP101516 could stimulate the sprouting of descending SERT and TH positive projections. Briefly, C6-C7 spinal cord cross-sections from animals in both control and treatment groups were examined for markers of descending SERT and TH positive projections. The density of SERT-positive and TH-positive axon terminals in the dorsal horn was then used as a measurement for axon sprouting. Compared to vehicle  114  Figure 3.7. DKP101516 stimulates the sprouting of intact CGRP positive afferents. (A) C7 spinal cord cross-sections from vehicle treated and DKP101516 treated animals were labeled for CGRP (Scale bar: 500 µm). The compass indicates the orientation of the image. Relative to vehicle treatment, DKP101516 treatment resulted in a marked increase of CGRP-positive axon density specifically in the lateral dorsal horn. (B) Densitometric measurements of CGRP-positive axon density specifically in the lateral dorsal horn. Relative to vehicle treatment, DKP101516 treatment significantly increased the density of CGRP positive axons (n=5, *p<0.05, One-way ANOVA). Pairwise comparisons (SNK post-hoc analysis) between control and treatment across all 5 laminae revealed that DKP101516 significantly increases in CGRP-positive axon density only in lateral regions in laminae I-II. (C) Spinal cord schematic with the region of interest as indicated by box. The compass below the schematic depicts the orientation (D= dorsal, V= ventral, M= medial, L=lateral).  115  116  treatment, DKP101516 treatment significantly enhanced the density of SERT-positive and THpositive positive axons in the dorsal horn (Figs., 3.8., 3.9., n=7, p<0.05, One-way ANOVA). Pairwise comparisons between DKP101516 and vehicle treated animals across all 5 laminae using SNK post-hoc analysis confirmed that DKP101516 stimulated axon sprouting across all 5 laminae.  3) SERT and TH positive Projections- Contralateral to Injury Although our results showed that DKP101516 stimulates significant axon sprouting in the injured side of the spinal cord, it is unclear whether DKP101516 works by directly stimulating axon sprouting, or by potentiating dorsal rhizotomy induced axon sprouting. To differentiate between the effects of injury and the effects of DKP101516, we therefore asked whether DKP101516 can stimulate axon sprouting of both SERT-positive and TH-positive projections independent of the signals associated with injury. In the dorsal rhizotomy model, the contralateral uninjured side of the spinal cord was assessed to determine the effect of DKP101516 in the absence of injury. Briefly, SERT-positive and TH-positive axon densities in the dorsal horn on the uninjured side were quantified, and comparisons were made between DKP101516 and vehicle treated animals. Results revealed that DKP101516 treatment significantly increased SERT-positive and TH-positive axon densities even on the uninjured side of the spinal cord (Figs. 3.8., 3.9., n=7, p<0.05, One-way ANOVA). Further pair-wise comparisons between control and treatment groups across all 5 laminae using the SNK post-hoc test revealed that the effects of DKP101516 vary between different neuronal types. Specifically, DKP101516 stimulated significant sprouting of TH-positive projections across all 5 laminae. In contrast, DKP101516 stimulated significant sprouting of SERT-positive projections only in laminae I/II.  117  Figure 3.8. DKP101516 stimulates the sprouting of SERT positive projections. (A) C7 spinal cord cross-sections from vehicle treated and DKP101516 treated animals were labeled with antibody to SERT. Orientation of the image is as indicated (D=dorsal, V=ventral, M=medial, L=lateral). Sprouting of these projections can be detected as an increase in axon density observed in the dorsal horn. Relative to vehicle treatment, DKP101516 treatment stimulated a marked increase in axon density in the dorsal horn. (B, C) Quantification of axon density in the dorsal horn on both the injured (ipsilateral) and uninjured (contralateral) sides of the spinal cord. Measurements revealed that DKP101516 treated animals have significantly greater SERTpositive axon density on both sides of the spinal cord relative to vehicle treated animals (n=7,*p<0.05, One-way ANOVA). Pairwise comparisons between treatment and control using post-hoc analysis revealed that DKP101516 increases axon density across all 5 laminae only on the injured side. However, DKP101516 increased axon density only in laminae I/II on the uninjured side.  118  119  Figure 3.9. DKP101516 stimulates the sprouting of TH positive afferents. (A) C7 spinal cord cross-sections from vehicle treated and DKP101516 treated animals were labeled for TH. Orientation of the image is as indicated (D=dorsal, V=ventral, M=medial, L=lateral). Sprouting of these projections can be detected as increased in axon density observed in the dorsal horn. Relative to vehicle treatment, DKP101516 treatment stimulated a marked increase in axon density in the dorsal horn. (B,C) Quantification of axon density in the dorsal horn on both the injured (ipsilateral) and uninjured (contralateral) sides of the spinal cord. Measurements revealed that DKP101516 treated animals have significantly greater TH-positive axon density on both sides of the spinal cord relative to vehicle treated animals (n=7, *p<0.05, One-way ANOVA). Post hoc analysis confirmed that DKP101516 increases axon density across all 5 laminae.  120  121  3.4. DISCUSSION The adult mammalian central nervous system is an environment that does not favor axon outgrowth and plasticity. The expression of various inhibitory molecules including myelin associated inhibitory proteins (MAIPs) and chondroitin sulfate proteoglycans (CSPGs) work together to protect the integrity of existing connections by inhibiting aberrant sprouting and synaptogenesis (Romero et al., 2000; McGee et al., 2005). Following incomplete spinal cord injury, a certain degree of axon plasticity is required for reconnections to be made. However, the expression of MAIPs and CSPGs throughout the adult spinal cord inhibits the axon plasticity necessary for compensatory axon sprouting and reinnervation (MacDermid et al., 2004; Massey et al., 2006; Barritt et al., 2006). In this study, we have identified a diketopiperazine, DKP101516, which can stimulate axon outgrowth and branching even in the presence of MAIPs and CSPGs. In vivo characterization using the septuple dorsal rhizotomy model revealed that DKP101516 enhances the sprouting of injured primary afferents, as well as intact primary afferents and descending monoaminergic projections. Collectively, the characterization of DKP101516 thus far suggests that DKP101516 may be a potential therapeutic agent to encourage axon sprouting necessary for spinal cord repair.  3.4.1. DKP101516 Mediated Astrogliosis Hinders Axon Regeneration In vitro characterization has demonstrated that DKP101516 can stimulate axon outgrowth in cortical neurons even in the presence of CSPGs and MAIPs. Therefore, it was expected that DKP101516 might stimulate primary afferent regeneration across the glial scar in the DREZ following septuple dorsal rhizotomy. Interestingly, DKP101516 appeared to stimulate axon sprouting only up to the dorsal root entry zone. Further analysis of astrocyte reactivity and neurocan expression revealed that DKP101516 also enhances astrogliosis in the DREZ.  122  Although DKP101516 increased astrocyte reactivity in the DREZ of both the injured and uninjured roots, DKP101516 enhanced neurocan expression only in the injured root. Since DKP101516 demonstrates limited ability to overcome the inhibitory effects of neurocan (Fig. 3.6.), we suggest that DKP101516-mediated neurocan expression at the injury site may be a major factor behind the failure of DKP101516 to mediate the regeneration of primary afferents across the DREZ.  The observation that DKP101516 enhanced neurocan expression only on the injured side of the spinal cord suggests that DKP101516 requires an unknown signal resulting from the injury to trigger neurocan expression and the consequent inhibition of axon regeneration. This unknown signal may be from molecules such as cytokines and chemokines that are involved in secondary damage following injury (Ramer et al., 2005; Yiu & He, 2006). Previous studies have shown that the cytokine receptor epithelial growth factor receptor (EGFR) can trigger astrocyte reactivity and neurocan expression (Asher et al., 2000; Smith & Strunz, 2005), and that EGFR inhibition can attenuate the inhibitory effects of the glial scar (Erschbamer et al., 2007). This raises the possibility that DKP101516 works in combination with EGFR ligands in order to enhance neurocan expression at the site of injury. Therefore, EGFR inhibitors may abolish DKP101516 mediated neurocan expression at the site of injury, and therefore allow primary afferents to regenerate across the DREZ. Future studies will be needed to address this hypothesis.  3.4.2. DKP101516 Stimulates Axon Sprouting in the Adult Spinal Cord Intrathecal application of DKP101516 following septuple dorsal rhizotomy stimulated the sprouting of intact primary afferents in to the lateral dorsal horn, as well as the sprouting of descending monoaminergic projections into the dorsal horn on both the injured (ipsilateral) and  123  uninjured (contralateral) sides of the spinal cord. Interestingly, DKP101516 treatment appeared to result in similar outcomes as those resulting from the neutralization of myelin associated proteins with the soluble nogo receptor (sNgR).  1) CGRP Positive Afferents Similar to sNgR treatment (MacDermid et al., 2004), DKP101516 treatment enhances the sprouting of CGRP-positive primary afferents specifically in laminae I-II in the lateral dorsal horn. The lateral dorsal horn is normally innervated by CGRP-positive afferents from more rostral and caudal segments of the spinal cord (MacDermid et al., 2004; Hampton et al., 2007). Following septuple C4-T2 dorsal rhizotomy, the sprouting of CGRP afferents from intact C3 and T3 roots typically results in the invasion of CGRP positive terminals within the lateral dorsal horn of deafferented C4-T2 segments (Hampton et al., 2007). Since DKP101516 induced sprouting of CGRP positive afferents specifically in the lateral dorsal horn, we can conclude that these afferents originate from intact roots rostral and caudal to the crushed roots.  2) SERT and TH Positive Projections Dorsal rhizotomy typically results in the release of cytokines and growth factors that stimulate the sprouting of spinally projecting axons ipsilateral to the injury (McHugh & McHugh, 2000). To determine whether DKP101516 mediates its effects independent of injury, we measured the density of SERT and TH positive axons not only in the dorsal horn ispilateral to the injury, but also on the contralateral (uninjured) side of the spinal cord. Similar to sNgR treatment (MacDermid et al., 2004), our results show that DKP101516 can stimulate axon sprouting of SERT and TH positive axon projections both ipsilateal and contralateral to the dorsal root injury, indicating that DKP101516 stimulates axon sprouting independent of the effects of injury.  124  Further analysis with uninjured animals will verify whether DKP101516 can indeed stimulate axon sprouting in the absence of injury.  Intriguingly, DKP101516 stimulates more sprouting of SERT-positive projections compared to sNgR-mediated neutralization of myelin-associated proteins. SNgR treatment results in an enhanced sprouting of SERT-positive projections in layer I/II of the dorsal horn only on the ipsilateral side. In contrast, DKP101516 treatment stimulates the sprouting SERT-positive projections not only in deeper dorsal horn regions in the ipsilateral side, but also in laminae I/II in the contralateral side. Since DKP101516 intraspinal axon sprouting appears to exceed that observed in sNgR treatment, we suggest that DKP101516 may overcome not only the effects of MAIPs but also the effects of other molecules inhibiting axon sprouting.  3.4.5. Signals Regulating Axon Outgrowth and Branching Axon outgrowth and branching is initiated at the neuronal growth cone. Axon outgrowth begins with F-actin polymerization and the extension of F-actin rich processes in the leading edge of the neuronal growth cone located at the tip of growing axons. These F-actin rich protrusions trigger the recruitment of microtubules necessary to mediate axon elongation (Dent & Gertler, 2003). On the other hand, axon branching is initiated through various mechanisms that include splitting of the growth cone during embryonic development, or de novo branching along the axon in the mature CNS (Luo, 2002). Similar to the process of axon outgrowth, de novo branching begins with the accumulation of F-actin at specific point on the axon, followed by protrusion, microtubule recruitment and consolidation into an axon branch (Dent et al., 2003; Kornack & Giger, 2005). In the adult CNS, inhibitory molecules such as MAIPs and CSPGs inhibit axon branching by disrupting F-actin accumulation and microtubule assembly through the  125  PKC-RhoA-ROCK pathway (Sivansakaran et al., 2004; Hasegawa et al., 2004).  Our studies demonstrate that DKP101516 stimulates axon outgrowth and branching, and overrides the inhibitory effects of MAIPs and CSPGs. Further characterization revealed that DKP101516 mediates these effects in a PI3K dependent mechanism. PI3K is the key modulator of a wide range of signaling pathways that regulate axon morphology. Specifically, PI3K promotes axon outgrowth and branching by stimulating F-actin and microtubule assembly. First, F-actin assembly is initiated through PI3K mediated synthesis of phosphotidylinositol-3,4,5triphosphate (PIP3), which in turn activates Rac/Cdc42 and the downstream actin polymerizing machinery (Menager et al., 2004; Da Silva et al., 2005). Second, microtubule assembly is initiated through PI3K/PIP3 mediated recruitment of protein kinase B (PKB- also known as Akt), which in turn inhibits cdk5 and GSK3-beta. The inhibition of cdk5 and GSK3-beta disinhibits microtubule assembly necessary for axon growth and branch formation (Kornak and Giger, 2005). PI3K activation also modulates the key downstream effectors of MAIP and CSPG. Specifically, PI3K inhibits RhoA during the maturation of hippocampal neurons (Da Silva et al., 2005), suggesting that DKP101516 may overcome the effects of MAIP and CSPG on neurite outgrowth branching through PI3K mediated RhoA inhibition. In summary, we suggest that DKP101516 stimulates axon outgrowth and branching, and overcomes inhibitory effects of MAIPs and CSPGs, through multiple aspects of PI3K signaling.  3.4.6. DKP101516- A Candidate Drug to Stimulate CNS Repair Our studies reveal that DKP101516 stimulates robust sprouting of various axon projections even in the inhibitory conditions of the adult spinal cord. Since axon sprouting can lead to reinnervation and functional recovery following spinal cord injury (Z’Graggen et al., 1998; Li et  126  al., 2004; Chan et al., 2005; Ramer et al., 2006; Massey et al., 2006), we suggest that DKP101516 is likely to encourage spinal cord repair by stimulating compensatory sprouting of intact spinally projecting axons. Although DKP101516 appears to promote structural reconnections, it is unclear whether these connections are compensatory or detrimental to function. For instance, we have demonstrated that DKP101516 can stimulate robust sprouting of intact CGRP positive nociceptive afferents following dorsal rhizotomy. At a certain level, the increased sprouting of nociceptive afferents may restore pain sensation in response to mechanical and thermal sensory inputs. However, aberrant sprouting of these nociceptive projections may also result in the development of abnormal pain sensations to non-noxious stimuli (Nakamura & Myers, 2000). We have also demonstrated that DKP101516 can stimulate the sprouting of descending monoaminergic projections into the dorsal horn. Reinnervation by descending SERT and TH positive projections is likely to restore the appropriate modulation of nociceptive responses, thus preventing the onset of neuropathic pain. However, aberrant sprouting of these projections may also result in undesirable effects. For instance, sprouting of SERT positive projections in the dorsal horn may result in enhanced nociceptive responses possibly through ‘silent’ glutaminergic synapses (Li & Zhuo, 1998). In addition, sprouting of TH positive projections in the dorsal horn may enhance dopaminergic input into the autonomic nervous system, and therefore produce negative cardiovascular effects such as bradycardia and hypotension (Lalhou, 1998). In summary, our results suggest a potential use of DKP101516 to stimulate spinal cord repair by promoting compensatory sprouting of spinally projecting axons. However, behavioral studies are needed to determine whether DKP101516 mediated sprouting can make the appropriate connections required to restore normal sensory function.  127  3.5. BIBLIOGRAPHY 19.  Asher, R.A., Morgenstern, D.A., Fidler, P.S., Adcock, K.H., Oohira, A., Braistead, J.E., Levine, J.M., Margolis, R.U., Rogers, J.H., Fawcett, J.W. 2000. 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Chondroitinase ABC digestion of the perineuonoal net promotes functional collateral sprouting in the cuneate nucleus after cervical spinal cord injury. J. Neurosci. 26, 4406–4414.  37.  McGee, A.W., Yang, Y., Fischer, Q.S., Daw, N.W., Strittmatter, S.M., 2005. Experience driven plasticity of visual cortex limited by myelin and Nogo receptor. Science 309, 2222–2226.  38.  McHugh, J.M., McHugh, W.B., 2000. Pain: neuroanatomy, chemical mediators, and clinical implications. AACN Clin. Issues 11, 168–178.  39.  Menager, C., Arimura, N., Fukata, Y., Kaibuchi, K., 2004. PIP3 is involved in neuronal polarization and axon formation. J. Neurochem. 89, 109–118.  40.  Nakamura, S.I., Myers, R.R., 2000. Injury to dorsal root ganglia alters innervation of spinal cord dorsal horn lamina involved in nociception. Spine 25, 537–542.  41.  Ramer, L.M., Borisoff, J.F., Ramer, M.S., 2004. Rho-kinase inhibition enhances axon plasticity and attenuates cold hyperalgesia after dorsal rhizotomy. J. 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Scott, A.L., Borisoff, J.F., Ramer, M.S., 2005. Deafferentation and neurotrophin-mediated intraspinal sprouting: a central role for the p75 neurotrophin receptor. Eur. J. Neurosci. 21, 81–92.  47.  Sivasankaran, R., Pei, J., Wang, K.C., Zhang, Y.P., Shields, C.B., Xu, X.M., He, Z., 2004. PKC mediates inhibitory effects of myelin and chondroitin sulfate proteoglycans on axonal regeneration. Nat. Neurosci. 7, 261–268.  48.  Smith, G.M., Strunz, C. 2005. Growth factor and cytokine regulation of chondroitin sulfate proteoglycans by astrocytes. Glia. 52, 209-18.  49.  To, K.C., Loh, K.T., Roskelley, C.D., Andersen, R.J., O'Connor, T.P., 2006. The antiinvasive compound motuporamine C is a robust stimulator of neuronal growth cone collapse. Neuroscience 139, 1263–1274.  50.  Wong, J.W., Brastianos, H.C., Andersen, R.J., O'Connor, T.P., 2008. A high-throughput screen to identify novel compounds to promote neurite outgrowth. J. Neurosci. Methods 169, 34–42.  51.  Zeng, Y., Li, Q., Hanzlik, R.P., Aubé, J., 2005. Synthesis of a small library of diketopiperazines as potential inhibitors of calpain. Bioorg. Med. Chem. Lett. 15, 3034– 3038.  52.  Z'Graggen, W.J., Metz, G.A., Kartje, G.L., Thallmair, M., Schwab, M.E., 1998. Functional recovery and enhanced corticofugal plasticity after unilateral pyramidal tract lesion and blockade of myelin-associated neurite growth inhibitors in adult rats. J. Neurosci. 18, 4744–4757.  130  CHAPTER 4  Transient Effects of DKP101516 on Axon Sprouting and Sensory Function3  3  A version of this chapter is being prepared for submission. Wong. J.W., Craig, J., Andersen, R.J., Ramer, M.S., O’Connor, T.P. Transient Effects of DKP101516 on Axon Sprouting and Sensory Function. (Manuscript in Progress).  131  4.1 INTRODUCTION Our previous study demonstrated that although DKP101516 fails to stimulate axon regeneration due to its effects on the astroglial scar, DKP101516 appears to enhance the sprouting of intact spinally projecting axons that contribute to sensory processing (Wong et al., 2008). The question that arises is whether DKP101516 mediated intraspinal axon sprouting has any implications on sensory function. To address this question, let us revisit some of the previous work in motor and sensory systems illustrating how axon sprouting and reorganization following spinal cord lesions can influence functional recovery.  Previous studies by the Tuszynski group have shown that spontaneous axon sprouting in the motor circuitry does play an important role in motor recovery (Weidner et al., 2001). In their studies, they showed that following incomplete corticospinal tract lesion, spontaneous recovery in motor function occurred. Based on the axon-sprouting model, spontaneous recovery is thought to result from the sprouting of intact corticospinal axons. Accordingly, when the nearby corticospinal axons were lesioned, all improvements in motor function were abolished. The results clearly demonstrated that the spontaneous sprouting of intact axonal projections is necessary for spontaneous recovery in motor function. Studies by the Schwab and Bradbury groups further illustrate this point by performing a series of manipulations that make the CNS more permissive for axon sprouting, such as the removal of inhibitory CSPGs or neutralization of Nogo (an inhibitory myelin associated molecule) (Z’Graggen et al., 1998; Barritt et al., 2006; Raineteau et al., 2001, 2002). The resulting permissive environment enabled more robust sprouting and reorganization of the nearby intact corticospinal and rubrospinal tracts (CST and RST), which corresponded to a significant improvement in motor function following  132  corticospinal tract injuries in rats (Raineteau et al., 2001, 2002). Overall, these studies emphasized that the spontaneous axon sprouting and reorganization of the motor circuitry following corticospinal lesions is a valid mechanism through which motor function can be restored.  Spinal cord injury models are problematic since return of function cannot be unambiguously attributed to changes in motor function or sensation and perception. Dorsal root injury on the other hand, which selectively removes primary afferent input from the cord serves as a more appropriate model since changes in behaviour can be more definitively linked to rewiring of sensory circuitry (Ramer et al., 2004, 2007). The cervical dorsal root injury model was previously tested in primate models, which has revealed that similar to the motor circuitry, the sensory circuitry also undergoes plastic changes or reorganizations post-injury (Darian-Smith & Brown, 2000; Darian-Smith, 2004). These changes can be characterized as spontaneous reorganization of the cutaneous receptive fields in the somatosensory cortex that can be attributed to the spontaneous sprouting of intact primary afferents that are adjacent to the injured rootlets (Darian-Smith & Brown, 2000; Darian-Smith, 2004). Moreover, the observed structural plasticity corresponded to recovery in mechanosensory function characterized by improvement in manual dexterity of the affected hand (Darian-Smith & Ciferri, 2005). Indeed, these studies firmly established that axon sprouting and reorganization of the sensory circuitry is also a mechanism through which sensory function can be restored.  Similar studies by the Ramer lab have shown that the spontaneous intraspinal sprouting following dorsal rhizotomy in rats is associated with recovery from neuropathic cold-pain as well as partial restoration of mechanosensation (Ramer et al., 2004; Ramer et al., 2007). The  133  limitations in intraspinal axon sprouting and functional recovery can be attributed in part to molecules expressed in the central nervous system that inhibit axon outgrowth (Ramer et al., 2005). Indeed, neutralization of the myelin associated molecule nogo-66 with a soluble Nogo receptor chimeric molecule (sNgR) into the spinal cord resulted in significantly greater axon sprouting following dorsal rhizotomy (MacDermid et al., 2004). Similarly, the removal of CSPGs with spinal infusion of chondroitinase ABC markedly enhanced the sprouting of intact afferents into the deafferented regions of the dorsal horn, resulting in robust recovery in mechanosensory function (Cafferty et al., 2008). Lastly, manipulations that inhibit signaling pathways that work downstream of MAIPs and CSPGs, such as Rho kinase inhibition (McKerracher & Winton, 2002) resulted in robust intraspinal axon sprouting and behavioural recovery following dorsal root injury (Ramer et al., 2004).  Since the above studies have demonstrated a link between axon sprouting and functional recovery, a key question that arises is whether DKP101516 mediated axon sprouting translates to functional recovery. To address this question, we assessed the behavioral effects of DKP101516 using the C7/C8 dorsal root injury (DRI) model. C7/C8 DRI is a well-characterized model to study the effects of intraspinal axon sprouting on sensory function (Ramer et al., 2004; Ramer et al., 2007; Soril et al., 2008). Following C7/C8 DRI, animals experience sensory dysfunction in their affected forepaw, characterized by the loss of mechanosensation and development of cold hyperalgesia, which peak at 2 and 6 days post-injury (days post DRI) respectively (Ramer et al., 2004; Ramer et al., 2007). Mechanosensory function partially improves by 8 days post-DRI, and cold pain resolves approximately one week later. Previous studies attributed the recovery to the intraspinal sprouting of intact primary afferents, inhibitory interneurons, and descending monoaminergic projections (Ramer et al., 2004; Ramer et al., 2007). Since DKP101516  134  stimulates robust intraspinal axon sprouting, we hypothesized that the spinal infusion of DKP101516 following C7/C8 dorsal root injury (DRI) would help accelerate improvements in mechanosensation and/or normal pain perception.  Our results demonstrated that following C7/C8 DRI, DKP101516 treatment moderately accelerated recovery of low threshold mechanosensation. Histological analysis revealed that this coincided with the sprouting of intact axonal populations that make up the sensory circuitry. Interestingly, the failure of DKP101516 to maintain long-term behavioral effects coincided with apparent axonal loss and enhanced microglia/macrophage reactivity.  4.2. METHODOLOGY 4.2.1. Surgery All surgical procedures were conducted according to the guidelines of the Canadian Council for Animal Care and the University of British Columbia Animal Care Committee. Male Sprague– Dawley rats (200–250 g) were anaesthetized with ketamine hydrochloride (75 mg/kg, BimedaMTC, Cambridge, ON) and medetomidine hydrochloride (0.5 mg/kg, Novartis, Mississauga, ON), by intra-peritoneal (i.p.) injection. For C7/8 dorsal root injury (C7/8 DRI), the left C7 and C8 dorsal roots were exposed via hemilaminectomy and durotomy, and the exposed roots were subjected to a complete transection injury. A pre-filled cannula was inserted through the atlantooccipital membrane along the dorsal surface of the cord until the tip rested at mid-C6. The opposite end was attached to an osmotic minipump (Alzet; Cupertino, CA) containing either DKP101516 (2.5 mg/mL) or vehicle (50% DMSO, 50% PEG). The drug was delivered at a rate of 1.25µg per day for up to 14 days. A total of 28 animals were included in this study.  135  4.2.2. Tissue Processing 7 or 23 days post-injury (days post DRI), the animals were killed with an overdose of chloral hydrate (1g/kg i.p.; Sigma Aldrich; Oakville, ON ) and perfused with 4% paraformaldehyde in 0.1M phosphate buffer. Their spinal cords were removed and post-fixed for 2 hours. The tissue was then cryoprotected in 20% sucrose in 0.1M phosphate buffer for 24 h at 4ºC, frozen at -80ºC, and cross-sectioned (16µm) on a cryostat.  4.2.3. Behavioral Studies Behavioral testing was conducted blind. Naïve rats were first handled and acclimatized to the behavioral setup prior to pre-operative testing. Testing was performed on two pre-operative days (days −2 and −1), and on alternative even-numbered days up to 20 days post-C7/8 DRI, performed on both forepaws. Daily test scores for each test were averaged from three trials separated by 1 h. The animals were subjected to a battery of tests to assess for cold hypersensitivity, mechanical allodynia, thermal allodynia, as well as response to low threshold tactile stimulation.  An acetone test for the palmar forepaw was used to assess cold pain, as done previously (Soril et al., 2008). Briefly, the animals were individually placed in clear cages on a raised wire mesh bottom. A micropipette was used to squirt 10 µl of acetone from below the wire mesh bottom, and the duration of response to acetone applied to the palmar surface of both forepaws was measured. Cold pain responses were defined by the shaking, withdrawal, biting or licking of the tested forepaw. Behavioral scoring ranged from a minimum of 1 s (a brief withdrawal without excessive attention to the forepaw), to a maximum of 15 s. A prolonged withdrawal without accompanying biting, licking or shaking was assigned a score of 3 s, even if the paw remained  136  elevated beyond this time.  Mechanical allodynia was examined using the Dynamic Plantar Aesthesiometer (Ugo Basile, Comerio, Italy). Rats were placed in a raised cage with a wire mesh platform over the stimulator unit. The metal filament was applied to the center of the palmar surface of the forepaw, and upward force was increased from 1 to 50 gm over 7 sec. Force at withdrawal was recorded for both forepaws.  Thermal allodynia was also tested using the palmar radiant heat test (Ugo Basile, Camerio, Italy). Animals were again placed in clear cages, this time on a glass platform. An infrared light source below the glass was then positioned under the center of the forepaw, and withdrawal latency from heat stimulus onset was recorded for both forepaws.  Finally, low threshold tactile mechanosensation across the palmar surface of the forepaw was assessed using an adhesive removal test (Bradbury et al., 2002; Ramer et al., 2007). Circular stickers (6.4 mm in diameter) were applied to the palm of both forepaws, and the rat was placed in a cage with a clear front window for observation. The time required for the rat to sense the sticker, indicated by a quick paw shake or biting and removal of sticker, was measured for each paw to a maximum time of 150s. The maximum time of 150s was imposed to separate sticker sensation in the palm from accidental sticker removal during grooming. Trials in which the sticker was discovered during grooming before the 150s maximum were excluded.  4.2.4. Immunohistochemistry Spinal cord sections were immunohistochemically processed to visualize as previously described  137  (Wong et al., 2008). Briefly, sections were immunostained for nociceptive afferents (CGRP [calcitonin gene related peptide] host mouse, 1:2000; Chemicon; Temecula, CA), mechanosensitive afferents (VGLUT1 [vesicular glutamate transporter 1] host guinea pig, 1:4000; Chemicon; Temecula, CA), inhibitory interneurons (V-GAT [vesicular GABA transporter], host rabbit, 1:500, Synaptic Systems, Goettingen, Germany, and NPY [neuropeptide Y, host rabbit, 1:1000], host rabbit, 1:1000, Peninsula Laboratories Inc., San Carlos, CA), descending serotonergic and monaminergic projections (SERT [serotonin transporter], host rabbit, 1:2000, Immunostar; TH [tyrosine hydroxylase], host sheep, 1:200, Chemicon; Temecula, CA), activated microglia/macrophage (ED-1, host mouse, 1:500, Serotec Cedarlane Laboratories, Hornby, ON, Canada), microglia/macrophage marker (CD11b, host mouse, 1:500, Sigma Aldrich, Oakville, ON, Canada), and reactive astrocytes (GFAP [glial fibrillary acid protein], host Rabbit, 1:1000, Sigma Aldrich, Oakville, ON, Canada). The immunolabeled sections were coverslipped and examined using a Zeiss (Jena, Germany) Axioplan II microscope. Digital images were captured using Northen Eclipse software (Empix Imaging; Mississauga, ON) via a digital camera (Qimaging; Surrey, BC).  4.2.5. Measuring Axon Density Axon density was measured as described previously (Wong et al., 2008). Briefly, axon density in the dorsal horn of both ipsilateral (injured) and contralateral (uninjured) side of the spinal cord was measured using SigmaScan Pro (Version 5.0). Axon populations that were assessed in this study include the nociceptive CGRP positive afferents, mechanosensitive VGLUT1 positive afferents, and SERT and TH positive descending projections, and NPY and VGAT positive inhibitory interneurons. The average density of axon terminals in lamina I/II, III-V was calculated for each animal, and plotted as means ± SEM.  138  4.2.6. Measuring ED-1 and GFAP Immunoreactivity ED-1 and GFAP immunoreactivity were measured as follows. Five random optical layered images per animal were captured and filtered, and the resulting thresholded images were analyzed with SigmaScan. Briefly, selected images of the cuneate funiculus, dorsal root entry zone and laminae I/II of the ipsilateral dorsal horn were manually selected, and average density of these areas was measured. Densitometric measurements were presented as means ± SEMs.  4.2.7. Statistics Data are reported as the mean ± SEM. All statistical analyses were performed by Sigma Stat (version 5.0). Statistical differences in terminal densities were detected by one-way ANOVA, followed by Holm Sidak post-hoc tests where appropriate, between vehicle and DKP101516 treated groups. For all behavioral tests, significance in test score improvement compared to postsurgery scores for both treatment and control groups was measured using ANOVA on ranks followed by Holm Sidak post-hoc analysis.  4.3. RESULTS 4.3.1. DKP101516 Promotes Transient Functional Recovery Following C7/C8 DRI Previously, we demonstrated that DKP101516 treatment stimulates robust plasticity of intact nociceptive and mechanosensitive afferents, as well as descending monoaminergic projections (Wong et al., 2008). A number of studies have linked the plasticity of the aforementioned axonal projections to changes in sensory behavior (Figure 4.1). For instance, the sprouting of nociceptive afferents including CGRP-positive afferents has been attributed to the development of cold hyperalgesia (Ji et al., 2007). On the other hand, the sprouting of descending SERT and TH positive monoaminergic projections was shown to reduce cold hypersensitivity following 139  C7/C8 dorsal rhizotomy (Ramer et al., 2004). Similarly, the BDNF induced sprouting of GABAergic interneurons was associated with changes in cold hypersensitivity following C7/C8 DRI (Soril et al., 2008). Lastly, the sprouting of VGLUT1-positive afferents had been correlated with the robust recovery in low threshold tactile mechanosensation following C7/C8 dorsal rhizotomy (Ramer et al., 2007).  To determine whether DKP101516 mediated axon sprouting would accelerate the abovementioned changes in sensory function, we assessed the behavioral effects of DKP101516 using the well-characterized C7/C8 dorsal root injury (DRI) model. Specifically, we applied either vehicle or DKP101516 intrathecally for 14 days following C7/C8 DRI. We also performed a battery of standard behavioral tests to assess the effects of DKP101516 on cold pain, low threshold mechanosensation, as well as thermal and mechanical allodynia.  Vehicle treated C7/C8 DRI animals experienced cold pain in the affected forepaw starting at around 2-4 days post-injury (days post DRI), which peaked at around 8 days post DRI, followed by spontaneous recovery (Fig. 4.2 B). Vehicle treated C7/C8 animals also experienced loss of low threshold mechanosensation in their ipsilateral forepaws immediately following injury, after which partial spontaneous recovery occurred (Fig. 4.2.A). C7/C8 DRI did not affect thermal or mechanical withdrawal thresholds.  DKP101516 treated animals experienced significantly faster mechanosensory recovery: latencies to sticker detection were significantly different from the first post-operative day two days earlier than vehicle-treated controls (Figs. 4.2, 4.3). Differences between DKP101516 and vehicle treated animals were significant between days 6-10 post DRI, but was no longer apparent at 12  140  Figure 4.1. Axonal projections involved in spontaneous sensory changes following C7/C8 DRI. (A) Intact afferents originate from adjacent roots rostral or caudal to the injured roots. These afferents project up and down the spinal cord, and normally populate the lateral horn of the spinal cord at the level of the injured C7/C8 roots as indicated. The spontaneous sprouting of these afferents (particularly the blue mechanosensory afferents) into the deinnervated dorsal horn at C7/C8 has been correlated with spontaneous recovery in low-threshold mechanosensory function. (B) Descending monoaminergic projections including TH+ and SERT+ projections originate from the brainstem nuclei locus coerulus and raphe nucleus respectively. These projections descend along the dorsal column, and innervate the dorsal horn in the regions indicated. The spontaneous sprouting of these projections into the deinnervated dorsal horn at C7/C8 has been correlated with spontaneous recovery from cold hypersensitivity. (C) Inhibitory interneurons populate the dorsal horn throughout the spinal cord. The spontaneous sprouting of this population was correlated with spontaneous recovery from cold hypersensitivity.  141  142  Figure 4.2. Effects of DKP101516 on Mechanosensation and Cold Hypersensitivity following C7/8 DRI. C7/C8 DRI rats treated with vehicle or DKP101516 were subjected to the following sensory tests for mechanosensation and cold hypersensitivity. (A) Animals were assessed for detection of low-threshold mechanosensory stimulus (placement of sticker) in the affected forepaw. The animal’s response latency to the sticker was measured over the course of 20 days. Compared to vehicle treated animals, DKP101516 treated animals displayed significantly shorter sticker response latency at 8 days post injury (days post DRI) (n=7, *p<0.05, repeated measures ANOVA), but the effect was lost by 12 days post DRI. (B) Animals were assessed for their pain response to cold stimulus (acetone) in the affected forepaw, and pain response duration was measured over the course of 20 days. No difference between vehicle and DKP101516 treated animals was observed.  143  144  Figure 4.3. DKP101516 accelerates mechanosensory recovery following C7/8 DRI. (A) Vehicle treated animals lost sticker stimulus response immediately following C7/C8 DRI, but significantly recovered response to sticker stimulus at 8 days post DRI (n=7, *p<0.05, repeated measures ANOVA) (B) Similarly, DKP101516 treated animals lost sticker stimulus response immediately following C7/C8 DRI, but significantly recovered response to sticker stimulus at 6 days post DRI: 2 days faster compared to vehicle (n=7, *p<0.05, repeated measures ANOVA).  145  146  days post-DRI (Figs. 4.2, 4.3). Other behavior analyses of DKP101516 treated animals revealed no significant difference in cold pain, heat or mechanical allodynia compared to vehicle treated animals (Fig. 4.4).  Although intrathecal treatment with DKP101516 for 7 days lead to sprouting of intraspinal axons (Wong et al., 2008), which may be related to enhanced mechanosensory recovery (above results), DKP101516 lost its positive behavioral effects with longer treatment. In order to better understand the structural basis underlying the effects of DKP101516, we performed histological analysis of axonal and glial profiles in spinal cord tissues harvested at 7 and 23 days post DRI.  4.3.2. Effects of DKP101516 on the Sensory Circuitry 1) Primary Afferents: Previous characterization has revealed that following septuple dorsal rhizotomy, DKP101516 stimulated robust sprouting of a various primary afferents including nociceptive CGRP-positive afferents in the lateral dorsal horn (Wong et al., 2008) as well as mechanosensitive VGLUT1positive afferents in laminae I/II (data not shown). Therefore, we expected that DKP101516 treatment following C7/C8 dorsal root injury would enhance the sprouting of intact primary afferents situated in the lateral dorsal horn. Immunolabeling of C7/C8 spinal cord sections for CGRP and VGLUT1 demonstrated that at 7 days post DRI, DKP101516 treated animals had significantly greater density of CGRP and VGLUT1 positive axonal terminals than vehicletreated controls in the ipsilateral dorsal horn (Figs. 4.4A-D, 4.5A-D), in both the deafferentation gap and in the lateral dorsal horn. The axon sprouting in the lateral dorsal horn was consistent with our previous findings, indicating that these afferents originate from intact roots rostral and caudal to the crushed roots (Wong et al., 2008).  147  Figure 4.4. DKP101516 does not affect pain response to radiant heat and pressure following C7/8 DRI. Animals were assessed for their pain response radiant heat (A) and pressure (B) stimuli in the affected forepaw. Pain response duration was measured over the course of 20 days. Vehicle and DKP101516 treated animals were observed to respond similarly to radiant heat and pressure.  148  149  Figure 4.5. DKP101516 stimulates transient sprouting of VGLUT1 positive mechanosensory afferents. (A,B) Compared to the dorsal horn in vehicle treated C7/C8 DRI animals at 7 days post DRI, we observed markedly greater density of VGLUT1 positive mechanosensory afferents in the ipsilateral dorsal horn of DKP101516 treated C7/C8 DRI animals at 7 days post DRI, particularly in the indicated lateral horn and deafferentation gap. (C,D) At 23 days post DRI, the density of VGLUT1 positive mechanosensory afferents in the dorsal horn of DKP101516 treated animals was markedly reduced to levels comparable to vehicle treated animals at same time point. (E) VGLUT1 positive terminals are from mechanosensory afferents originating from intact dorsal roots rostral or caudal to the injured roots. (F,G ) Further quantification confirmed that DKP101516 significantly enhanced sprouting of VGLUT1 positive afferents at 7 days post DRI in laminae III-V of the above-indicated regions of the dorsal horn (n=7, *p<0.05, one way ANOVA). (H, I ) However, there was no significant difference between DKP101516 and vehicle treated animals at 23 days post DRI. (Scale bar: 100 µm)  150  151  Intriguingly, in spinal cord tissue harvested at 23 days post DRI, we found that the density of VGLUT1 positive axons in the ipsilateral dorsal horn of both vehicle and DKP101516 was markedly reduced (Figs. 4.4 E-H). Furthermore, there was no significant difference in VGLUT1 immunoreactivity between vehicle and DKP101516 in both the deafferentation gap and lateral region of the ipsilateral dorsal horn, indicating that the sprouting of VGLUT1 positive afferents was no longer stimulated (Figs. 4.4 E-H). Similar analysis of CGRP positive population revealed DKP101516 treated animals had significantly lower density of CGRP in laminae I/II in the ipsilateral lateral dorsal horn compared to vehicle treated animals (Figs. 4.5 E-H). It is therefore evident that prolonged DKP101516 treatment from 7-23 days post DRI not only did not promote further axon sprouting, but also might not have maintained the afferent sprouts that were generated in the first 7 days of treatment.  2) Supraspinal Projections Following septuple dorsal rhizotomy, DKP101516 stimulated the robust sprouting of descending SERT- and TH-positive monoaminergic projections in both the ipsilateral and contralateral dorsal horn (Wong et al., 2008). We therefore expected that spinal infusion of DKP101516 following C7/C8 DRI is likely to enhance the sprouting of SERT and TH projections in the ipsilateral dorsal horn. In spinal cord tissue harvested at 7 days post DRI, DKP101516-treated animals had significantly greater density of TH-positive projections across all laminae in the ipsilateral dorsal horn compared to vehicle treated animals (Figs. 4.7 A, B, I). Likewise, DKP101516-treated animals had significantly greater density of SERT positive projections in laminae I/II of the ipsilateral dorsal horn compared to vehicle treated animals (Figs. 4.7 E, F, K).  152  Figure 4.6. DKP101516 stimulates transient sprouting of CGRP positive nociceptive afferents. (A,B) At 7 days post DRI, we observed a markedly greater density of CGRP positive afferents in the ispilateral dorsal horn of DKP101516 treated C7/C8 DRI animals compared to that of vehicle treated animals, particularly in the indicated deafferentation gap and lateral dorsal horn regions. (C,D) At 23 days post DRI, the density of CGRP positive nociceptive afferents in the dorsal horn of DKP101516 treated animals was markedly reduced. (E) CGRP positive terminals are from nociceptive afferents originating from intact dorsal roots rostral or caudal to the injured roots. (F,G) Further quantification revealed that DKP101516 significantly enhanced the sprouting of CGRP positive nociceptive afferents in laminae I/II of the above-indicated regions (n=7, *p<0.05, one way ANOVA). (H, I) Similar quantification at day 23 post DRI revealed that DKP101516 treated C7/C8 DRI animals had significantly fewer CGRP positive terminals in the lateral dorsal horn compared to vehicle treated C7/C8 DRI animals at the same time point (n=7, *p<0.05, one way ANOVA). Axon sprouting in the deafferentation gap did not differ significantly between vehicle and DKP101516 treated groups. (Scale bar: 100 µm)  153  154  In contrast, in spinal cord tissue collected at 23 days post DRI, we found that DKP101516 treated animals had significantly lower density of TH positive projections in laminae I/II in the ipsilateral dorsal horn compared to vehicle treated animals (Figs. 4.7 C, D, J). Interestingly, the density of SERT positive axons in the ipsilateral dorsal horn of both vehicle and DKP101516 was noticeably reduced (Figs. 4.7 G-L). Furthermore, comparison between DKP101516 and vehicle treated animals revealed that there was no significant difference in the density of SERTpositive projections in the ipsilateral dorsal horn (Figs. 4.7 G, H, L). Similar to the effects of DKP101516 on primary afferents, it is apparent that prolonged DKP101516 treatment was not sufficient to maintain the monoaminergic axon sprouts generated during initial exposure to DKP101516 (0-7 days post DRI).  3) Inhibitory Interneurons Another component of the sensory circuitry is the inhibitory GABAergic spinal interneurons that populate the dorsal horn of the spinal cord. These inhibitory neurons function to modulate the transmission of sensory signals to the brain (Zhou et al., 2007; Soril et al., 2008). As an important component of the sensory circuitry, a question that arises is whether DKP101516 would influence the plasticity of spinal interneurons. To address this question, we used immunohistochemical approaches to examine C7/C8 spinal cord sections for spinal interneuron markers such as vesicular GABA-ergic transporter-1 (VGAT) and neuropeptide Y (NPY). Analysis of spinal cord tissue harvested at 7 days post DRI revealed that compared to vehicle treatment, DKP101516-treated animals had significantly greater density of VGAT and NPY positive axon terminals only in laminae I/II of the ipsilateral dorsal horn (Figs. 4.8 A-D).  155  Figure 4.7. DKP101516 stimulates transient sprouting of descending supraspinal projections. (A-D) At 7 days post DRI, DKP101516 treated C7/C8 DRI animals had markedly greater density of SERT and TH positive positive supraspinal axon terminals in the ipsilateral dorsal horn. (E-H) In contrast, at 23 days post DRI, DKP101516 treated C7C8 DRI animals had markedly reduced density of TH positive axon terminals in the dorsal horn. (I-J) Further quantification demonstrated that at 7 days post DRI, DKP101516 significantly enhanced the sprouting of TH positive axons across all laminae of the ipsilateral dorsal horn (n=7, *p<0.05, one way ANOVA). However, at 23 days post, DRI DKP101516 significantly reduced TH positive axon sprouting in laminae I/II of the dorsal horn (n=7, *p<0.05, one way ANOVA). (K,L) Quantification revealed that at 7 days post DRI, DKP101516 significantly enhanced the sprouting of SERT positive projections in laminae I/II of the ipsilateral dorsal horn (n=7, *p<0.05, one way ANOVA). However at 23 days post DRI, there was no longer any significant difference in SERT positive axon density between vehicle and DKP101516 treated animals. (Scale bar: 100 µm)  156  157  However, in spinal cord tissue harvested at 23 days post DRI, both vehicle and DKP101516 treated animals appeared to have markedly fewer VGAT- and NPY- positive sprouts in the ipsilateral dorsal horn (Figs 4.8 C, D, J). Moreover, DKP101516-treated animals had significantly lower density of VGAT- and NPY-positive terminals in laminae I/II of the ipsilateral dorsal horn (Figs. 4.8 E-H). We therefore conclude that while short-term DKP101516 treatment stimulated the sprouting of inhibitory interneurons, these were not maintained with prolonged exposure to DKP101516.  4.3.3. Glial Effects of DKP101516 Behavioral and structural characterization of DKP101516 so far suggests that while short-term DKP101516 promoted significant sensory improvement by stimulating sprouting of sensory circuitry, prolonged exposure to DKP101516 did not maintain the sensory recovery associated with the initial effects of DKP101516 on axon sprouting. The long-term decline in axon sprouts could be the result of a number of possibilities, including potential neurotoxicity of intrathecal DKP101516 in vivo, and the possible effects of DKP101516 on the glial environment in the deafferented dorsal horn. In order to investigate the latter possibility, we examined the glial effects of intrathecal DKP101516.  We examined spinal cord tissue for GFAP and ED-1 immunoreactivity to assess astrocyte hypertrophy and microglia/macrophage reactivity respectively, with reference to the glial events that occur following dorsal root injury (DRI). Briefly, DRI promotes astroglial scarring (GFAP immunoreactivity) in the dorsal root entry zone (DREZ) within 3 days post injury (McPhail et al., 2005). This is followed by Wallerian degeneration of the injured afferents (Chew et al., 2008; Zhang et al., 2009), which typically results in the delayed microglia/macrophage reaction  158  Figure 4.8. DKP101516 stimulates transient sprouting of inhibitory interneurons. (A-D) At 7 days post DRI, DKP101516 treated C7/C8 DRI animals had markedly greater density of NPY and VGAT positive supraspinal axon terminals in the ipsilateral dorsal horn. (E-H) In contrast, at 23 days post DRI, DKP101516 treated C7/C8 DRI animals had markedly reduced density of NPY positive axon terminal in the lateral dorsal horn. (I-L) Further quantification demonstrated that at 7 days post DRI, DKP101516 enhanced the sprouting of NPY and VGAT positive axons in laminae I/II of the lateral dorsal horn (n=7, *p<0.05, one way ANOVA). However, at 23 days post DRI, there was no longer any significant difference in NPY and VGAT positive axon density between vehicle and DKP101516 treated animals. (Scale bar: 100 µm)  159  160  (characterized by increased ED-1 immunoreactivity) in the dorsal funiculus and ipsilateral dorsal horn starting 1-2 weeks post DRI (Liu et al., 1998; Ramer et al., 2001).  Comparisons between vehicle and DKP101516-treated animals revealed that at 7 days post DRI, DKP101516-treated animals had significantly greater GFAP immunoreactivity in the DREZ and the ipsilateral dorsal horn (Figs. 4.9 A, B, I), as well as a noticeable increase in GFAP positive processes in the cuneate fasciculus (Figs, 4.9 C, D). However, there appeared to be no difference in ED-1 immunoreactivity between vehicle and DKP101516 treated animals, which was consistently restricted to the peripheral dorsal root in both treatment groups (Figs. 4.9 A-D). Similar comparisons in tissue harvested at 23 days post DRI revealed no significant difference in GFAP immunoreactivity between DKP101516 and vehicle treated animals (Figs. 4.9 E-H, J). As expected, both groups of animals at this time point (23 days post DRI) have pronounce ED-1 immunoreactivity particularly in the cuneate fasciculus (Figs. 4.9 E-H). However compared to vehicle treated animals, DKP101516 treated animals appeared to have more ED-1 positive cells in the cuneate fasciculus (Figs. 4.9 E, F), and significantly greater ED-1 invasion into laminae I/II of the dorsal horn (Figs. 4.9 G, H, L), which coincided with the apparent decline in axon density in laminae I/II of the dorsal horn following prolonged intrathecal DKP101516 treatment.  To determine whether the increased ED-1 immunoreactivity was the result of microglia/macrophage reaction, we attempted to correspond ED-1 staining with the distribution of microglia/macrophage (general marker CD11b) in the spinal cord. In tissues harvested at 7days post DRI, both vehicle and DKP101516 treated animals showed no difference in the density and distribution of CD11b positive cells in the ipsilateral dorsal horn, cuneate fasciculus and central dorsal root regions (Figs. 4.10 A, B). However, at 23 days post DRI, we observed the  161  apparent expansion of CD11b positive cells particularly in the cuneate fasciculus in both vehicle and DKP101516 treated animals (Figs. 4.10 C, D). Moreover, DKP101516 treatment resulted in more robust expansion and infiltration of CD11b positive cells in the cuneate fasciculus, such that more CD11b positive cells invaded into the laminae I-II of the ipsilateral dorsal horn (Fig. 4.10 D). Overall, the area of CD11b positive cell expansion appeared to coincide with the observed ED-1 immunoreactivity associated with prolonged DKP101516 treatment. These findings suggest a possible involvement of microglia/macrophage reactivity in the observed axonal loss associated with prolonged DKP101516 treatment.  Collectively, it appears that while short-term DKP101516 treatment enhanced astrogliosis, longterm DKP101516 resulted in enhanced ED-1 immunoreactivity in the dorsal horn. This indicates that short-term DKP101516 treatment stimulates afferent and intraspinal axon sprouting, but not afferent regeneration due to its effects on astrogliosis. On the other hand, prolonged DKP101516 mediated increase in ED-1 immunoreactivity (which is indicative of microglia/macrophage reactivity) in laminae I/II of the dorsal horn may be involved in the detrimental effects of DKP101516 on axon sprouting and sensory function.  4.4 DISCUSSION 4.4.1. Beneficial and Detrimental Effects of DKP101516 Previous in vitro and in vivo characterization of DKP101516 suggests that this marine-derived compound not only enhances axon outgrowth in vitro, but also promotes robust axonal sprouting in the sensory circuitry following septuple dorsal rhizotomy. Similar to our previous findings, short-term DKP101516 treatment following C7/C8 DRI resulted in robust sprouting of primary  162  Figure 4.9. DKP101516 mediates negative glial effects in the long-term. (A-D) At 7 days post DRI, DKP101516 treated animals demonstrated much greater GFAP positive immunoreactivity in both the dorsal root entry zone (DR) and the dorsal horn (DH) compared to vehicle treated animals. As expected, ED-1 immunoreactivity was restricted in the peripheral dorsal root in both vehicle and DKP101516 treated animals. Closer examination (inset- C,D) of the cuneate funiculus (CF) and dorsal horn (DH) revealed that ED-1 positive cells had not yet entered the spinal cord. (E-H) However at 23 days post DRI, there were no longer any difference in GFAP immunoreactivity in the ipsilateral DR or DH between vehicle and DKP101516 treated animals. Closer examination (inset- G,H) of the CF and DH revealed marked differences in the distribution and density of ED-1 positive cells between vehicle and DKP101516 treated animals. Specifically, vehicle treated animals had the majority of ED-1 positive cells restricted in the CF, with only a few spots in the DH. In contrast, DKP101516 treated animals not only appeared to have markedly more ED-1 positive cells in the CF, but also significantly more ED-1 positive cells in laminae I-II of the DH. (I) Quantification at 7 days post DRI revealed that DKP101516 treatment significantly enhanced glial scarring characterized by increased density of GFAP positive processes (n =7, *p<0.05, one way ANOVA). (K) No difference in the number ED-1 positive cells was observed between DKP101516 and vehicle treated animals at 7days post DRI. (J) Quantification at 23 days post DRI revealed that there was no longer any significant difference in the density of GFAP positive processes between vehicle and DKP101516 treated animals. (L) However, DKP101516 treated animals at 23 days post DRI had significantly more ED-1 positive cells in the dorsal horn compared to vehicle treated animals (n=7, *p<0.05, one way ANOVA). (Scale bar: 100 µm).  163  164  Figure 4.10. DKP101516 promotes the delayed infiltration and/or expansion of CD11b positive cells in the cuneate fasciculus. CD11b immunoreactivity in the spinal cord at 7 and 23 days post DRI. (A,B) At 7 days post DRI, there was no observable difference between vehicle and DKP101516 treated animals. In both groups, uniform CD11b distribution was observed in the dorsal horn (DH) and cuneate funiculus (CF). (C,D) At 23 day post DRI, there was a marked increase in CD11b positive cells in the CF in both vehicle and DKP101516 treated groups. Furthermore, DKP101516 treatment appeared to result in the marked expansion of CD11b positive cells in the CF, as well as increased encroachment of these cells into laminae I-II of the DH.  165  166  afferents, inhibitory interneurons and descending supraspinal projections that make up the sensory circuitry. Functionally, DKP101516 mediated sprouting of VGLUT1 positive afferents coincided with the transient improvement in low threshold mechanosensory function at 7 days post DRI. On the other hand, the sprouting of CGRP positive nociceptive afferents, SERT- and TH-positive descending supraspinal projections and VGAT-positive inhibitory interneurons observed at 7 days post DRI were not correlated with any effects on cold hyperalgesia.  One of the reasons underlying the transient effects of DKP101516 on sensory behavior may be the observed axonal loss associated with prolonged DKP101516 treatment. Specifically, we find that prolonged DKP101516 treatment (7-14 days post DRI) appeared to terminate the sprouting effects of DKP101516 on intraspinal axons. Specifically, the sprouting of CGRP positive afferents, as well as SERT and VGAT positive projections in the deafferented dorsal horn was no longer stimulated. On the other hand, prolonged DKP101516 spinal infusion also resulted in the apparent loss of axon sprouts originating from intact mechanosensory afferents (VGLUT-1 positive) as well as from the lateral tegmental nucleus in the brain stem (TH positive populations).  Intriguingly, the apparent loss of axon sprouts following prolonged DKP101516 exposure appeared to coincide with the enhanced ED-1 immunoreactivity in laminae I/II of the ipsilateral dorsal horn. As an indicator of enhanced microglia/macrophage reactivity that has been previously implicated in axonal retraction (McPhail et al., 2007, Horn et al., 2009), we suggest that long-term DKP101516 treatment may have caused an increase in microglia/macrophage reactivity and consequently axon loss. However, the effects of DKP101516 on microglia/macrophage reactivity may not necessarily be detrimental, since previous studies have  167  demonstrated the positive role of injury-induced microglia/macrophage activation on axon growth (Krenz & Weaver, 2000; Batchelor et al., 2002; Cafferty et al., 2004). Furthermore, it is also likely that the detrimental effects of DKP101516 may be indicative of neurotoxicity. Since cell death in the spinal cord could also enhance ED-1 immunoreactivity (Liu et al., 1998), there is a possibility that prolonged intrathecal DKP101516 treatment may be toxic, and may have resulted in increased ED-1 immunoreactivity in the aftermath. These findings may warrant future investigation on the toxicity of DKP101516 using in vitro assays.  In addition to the potential toxicity of DKP101516, our studies also demonstrated that prolonged intrathecal application of vehicle (1:1 DMSO and polyethylene glycol [PEG] mixture) resulted in profound axon loss in the dorsal horn. At day 23 days post DRI, we observed a marked reduction in the density of various axon populations in the ipsilateral dorsal horn of vehicle treated animals, including VGLUT1 positive mechanosensory afferents, SERT positive supraspinal projections, and NPY positive inhibitory interneurons (Figs. 4.6, 4.7, 4.8). Since prolonged DMSO and/or PEG intrathecal treatment had been previously shown to be neurotoxic (Miracci et al., 2009; Cole & Shi, 2005), we suggest that the prolonged effects our vehicle may have also contributed to the detrimental changes in the spinal cord.  Based on the above results, we propose the following model illustrating the behavioral and structural effects of DKP101516 (Fig. 4.11.). At 0-7 days post DRI, DKP101516 stimulated sprouting of various sensory projections in the ipsilateral dorsal horn. At 7-23 days post DRI, DKP101516 treatment apparently resulted in enhanced microglia/macrophage reactivity in the ipsilateral dorsal horn and cuneate fasciculus that coincided with the loss of axon sprouts. Structurally, these events suggest that the detrimental effects of DKP101516 including increased  168  microglia/macrophage activation and profound axon loss . Behaviorally, the transient axon sprouting in DKP101516 treated animals was only sufficient to produce subtle improvements in mechanosensory function (Figure 4.12), as they were rapidly reduced, concomitant to the axon loss associated with prolonged DKP101516 treatment.  4.4.2. Effects of DKP101516 on Microglia/Macrophage Reactivity following C7/C8 DRI C7/C8 DRI is often followed by Wallerian degeneration, which is characterized by active retraction of intact afferents, and the delayed clearance of axonal remnants and myelin debris by microglia/macrophages that are recruited to the spinal cord (Horn et al., 2009). A number of studies have suggested that microglia/macrophage reactivity could directly mediate axonal retraction (McPhail et al., 2007; Horn et al., 2009). For example, time-lapse imaging studies revealed that reactive microglia/macrophage in culture can promote active retraction of injured afferents via a contact dependent mechanism (Horn et al., 2009). Intriguingly, the depletion of macrophages from circulation was shown to marked reduce macrophage infiltration into the spinal cord, suggesting that the ED-1 immunoreactivity in the spinal cord lesion represents not only microglia but also macrophages from the circulation (Horn et al., 2009).  In the present study, DKP101516 appears to enhance the number of ED-1+ reactive microglia/macrophages in the dorsal column and dorsal horn at 23 days post-DRI. As DKP101516 was identified from active extracts that can stimulate cell migration, it is likely that DKP101516 may stimulate not only microglia activation but also the infiltration of reactive  169  Figure 4.11. Summary of the effects of DKP101516 treatment following C7/C8 DRI. (A) Vehicle or DKP101516 was intrathecally applied immediately following C7/C8 DRI. (B) At 7 days post DRI, DKP101516 treatment enhanced the sprouting of regenerating afferents in the peripheral root, as well as spinally projecting axons. (C) At day 23 post DRI, DKP101516 treatment no longer stimulated the axon sprouting, but rather resulted in axonal loss. DKP101516 treatment also resulted in an increase in microglia/macrophage reactivity, which may contribute to axonal loss in the long-term.  170  171  Figure 4.12. Timeline illustrating the effects of DKP101516 from 0-23 days post DRI. 0-7 days: DKP101516 treatment enhanced the sprouting of spinally projecting axons, and stimulated astroglial scarring in the dorsal root, dorsal funiculus and dorsal horn. 7-23 days: Prolonged DKP101516 treatment resulted in the loss of axon sprouts, which correlated with the apparent increase in microglia/macrophage reactivity and infiltration into the dorsal horn.  172  173  macrophages from circulation the into the spinal cord. It is also likely that DKP101516 mediated microglia/macrophage activation and/or recruitment may contribute to the observed axonal loss at 23 days post-DRI.  4.4.3. Therapeutic Potential of DKP101516 1) Brachial Plexus Avulsion and Chronic Pain C7/C8 DRI is a typical animal model for deafferentation injuries and neuropathic pain in humans. Similar to that observed in the C7/C8 model in rats, typical symptoms that may arise following deafferentation in humans include the development of neuropathic pain as well as abnormal mechanosensation in the affected dermatome. In our study, we find that DKP101516 transiently promotes the recovery in mechanosensation in the affected forepaw of C7/C8 DRI rats. However, due to the axonal loss associated with DKP101516 treatment, we did not see lasting mechanosensory improvements, or any recovery from cold pain. We therefore suggest that further studies on how to best reduce the detrimental effects of DKP101516 would shed further light into the potential therapeutic use of DKP101516 in treating mechanosensory and pain disorders in human deafferentation.  2) Spinal Cord Injury In most clinical cases, spinal cord injury is often incomplete, and is usually coupled with compensatory axon sprouting and partial recovery in function (Weidner et al., 2001; DarianSmith, 2004). However, due to the unfavorable environment in the CNS, axon sprouting and  174  functional recovery is limited (Z’Graggen et al., 1998; Li et al., 2004; MacDermid et al., 2004; Massey et al., 2006; Barritt et al., 2006). In our C7/C8 DRI studies, we find that DKP101516 appears to stimulate transient sprouting of a wide range of axonal projections in the spinal cord, resulting in transient improvement in mechanosensory function.  Based on these studies, we strongly suggest that DKP101516 is likely to enhance the plasticity of most spinally projecting axons following incomplete spinal cord injury. We further suggest that DKP101516 mediated sprouting is likely to promote recovery in sensory, motor and autonomic functions following incomplete spinal cord injury. In order to better assess the therapeutic potential of DKP101516 for incomplete spinal cord injury, future investigation on how to best reduce the detrimental effects of DKP101516 may be necessary.  4.4.3. Conclusion and Future Directions In our present study, we find that DKP101516 demonstrated short-term beneficial effects and long-term detrimental effects following C7/C8 DRI in rats. Based on our histological analysis, we were also able to associate the beneficial effects with DKP101516 mediated axon sprouting between 0-7 days post DRI, and the detrimental effects with DKP101516 mediated axon loss and increase in ED-1 immunoreactivity at 7 –14 days post DRI. We therefore conclude DKP101516 is beneficial in the short-term, but detrimental in the long-term characterized by axon loss and enhanced microglia/macrophage reactivity.  In order to use DKP101516 as a potential therapeutic, a future direction is to look for ways to minimize the detrimental effects of DKP101516. One approach is to modify the timing and duration of DKP101516 treatment. For instance, we find that DKP101516 enhanced  175  microglia/macrophage reactivity, resulting in axon loss, and decline in sensory improvement. Perhaps one approach may be to not administer DKP101516 during the period when which microglia/macrophages are most robustly activated, typically at 7-14 days post DRI. A feasible method to achieve this may be to administer DKP101516 for only the first 7 days postsurgery prior to microglia/macrophage reaction. Since an ideal feature of novel therapeutics is the effectiveness of short-term treatment, the success of this study may expedite the development of DKP101516 as a novel therapeutic. However, this approach may not be feasible if continuous DKP101516 infusion is required to produce sprouting effects.  Another possible approach is to combine DKP101516 treatment with minocycline- a drug that inhibits microglia/macrophage activation (Tikka et al., 2001; Stirling et al., 2004). The advantage of this approach is that it extends the beneficial effects of DKP101516, while eliminating the detrimental effects associated with prolonged DKP101516 exposure. While this approach could further maximize the beneficial effects of DKP101516, a potential problem could be the complications in combining two different treatments in vivo. Furthermore, minocycline treatment may be minimally effective if the issue here is not microglia/macrophage reactivity, but rather the neurotoxicity of prolonged DKP101516 treatment.  Lastly, perhaps a fundamental question that needs to be addressed is whether intrathecal DKP101516 and/or the vehicle (1:1 DMSO and PEG mixture) could be toxic. To address this issue, future studies should deal with the previously reported toxicity of our vehicle (Cole & Shi, 2005). One strategy previously demonstrated to minimize/avoid the toxicity of prolonged PEG exposure is to perform pulsatile intrathecal administration (Cole & Shi, 2005), where a bolus of vehicle and/or DKP101516 are injected intrathecally at various time points over the course of the  176  experiment. Alternatively, we may explore a less toxic vehicle for DKP101516 delivery, perhaps by simply implanting a biopolymer (scaffold/foam) previously infused with DKP101516 into the intrathecal space at the level of dorsal root injury. On the other hand, since DKP101516 appears to be highly insoluble, it is possible that the low solubility of DKP101516 in vehicle/intrathecal space may contribute to the long-term detrimental effects of DKP101516 in vivo. Therefore, another future study may be to assess the solubility and distribution of DKP101516 during intrathecal treatment, and to reformulate DKP101516 such that it can be more soluble vivo. Clearly, future studies into administration protocols and formulation to maximize the beneficial effects of DKP101516 may be needed to further develop DKP101516 as a potential therapeutic.  177  4.5. BIBLIOGRAPHY 1.  Barritt, A.W., Davies, M., Marchand, F., Hartley, R., Grist, J., Yip, P., McMahon, S.B., Bradbury, E.J. 2006. 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Plasticity in intact A delta- and C-fibers contributes to cold hypersensitivity in neuropathic rats. Neuroscience. 150, 182-93.  16.  Krenz, N.R., Weaver, L.C. 2000. Nerve growth factor in glia and inflammatory cells of the injured rat spinal cord. J Neurochem. 74, 730-9.  17.  Li, S., Liu, B.P., Budel, S., Li, M., Ji, B., Walus, L., Li, W., Jirik, A., Rabacchi, S., Choi, E., Worley, D., Sah, D.W., Pepinsky, B., Lee, D., Relton, J., Strittmatter, S.M. 2004. Blockade of Nogo-66, myelin-associated glycoprotein, and oligodendrocyte myelin glycoprotein by soluble Nogo-66 receptor promotes axonal sprouting and recovery after spinal injury. J Neurosci. 24, 10511-20.  18.  Liu, L., Persson, J.K., Svensson, M., Aldskogius, H. 1998. Glial cell responses, complement, and clusterin in the central nervous system following dorsal root transection. Glia. 23, 221-38.  19.  MacDermid, V.E., McPhail, L.T., Tsang, B., Rosenthal, A., Davies, A., Ramer, M.S. 2004. A soluble Nogo receptor differentially affects plasticity of spinally projecting axons. Eur J Neurosci. 2004 20, 2567-79.  20.  Marcacci, G., Corazzelli, G., Becchimanzi, C., Arcamone, M., Capobianco, G., Russo, F., Frigeri, F., Pinto, A. 2009. DMSO-associated encephalopathy during autologous peripheral stem cell infusion: a predisposing role of preconditioning exposure to CNSpenetrating agents? Bone Marrow Transplant. (in press).  21.  Massey, J.M., Hubscher. C.H., Wagoner, M.R., Decker, J.A., Amps, J., Silver, J., Onifer, S.M. 2006. Chondroitinase ABC digestion of the perineuronal net promotes functional collateral sprouting in the cuneate nucleus after cervical spinal cord injury. J Neurosci. 26, 4406-14.  22.  McPhail, L.T., Stirling, D.P., Tetzlaff, W., Kwiecien, J.M., Ramer, M.S. 2004. The contribution of activated phagocytes and myelin degeneration to axonal retraction/dieback following spinal cord injury. Eur J Neurosci. 20, 1984-94.  23.  McPhail, L.T., Plunet, W.T., Das, P., Ramer, M.S. 2005. The astrocytic barrier to axonal regeneration at the dorsal root entry zone is induced by rhizotomy. Eur J Neurosci. 21, 267-70.  24.  McKerracher, L., Winton, M.J. 2002. Nogo on the go. Neuron. 36, 345-8.  179  25.  Milligan, E.D., Watkins, L.R. 2009. Pathological and protective roles of glia in chronic pain. Nature Reviews Neurosci. 10, 23-36.  26.  Raineteau, O., Fouad, K., Noth, P., Thallmair, M., Schwab, M.E. 2001. Functional switch between motor tracts in the presence of the mAb IN-1 in the adult rat. Proc Natl Acad Sci U S A. 98, 6929-34.  27.  Raineteau, O., Fouad, K., Bareyre, F.M., Schwab, M.E. 2002. Reorganization of descending motor tracts in the rat spinal cord. Eur J Neurosci. 16, 1761-71.  28.  Ramer, L.M., Borisoff, J.F., Ramer, M.S. 2004. Rho-kinase inhibition enhances axonal plasticity and attenuates cold hyperalgesia after dorsal rhizotomy. J Neurosci. 24, 10796805.  29.  Scott, A.L., Borisoff, J.F., Ramer, M.S. 2005. Deafferentation and neurotrophin-mediated intraspinal sprouting: a central role for the p75 neurotrophin receptor. Eur J Neurosci. 21, 81-92.  30.  Ramer, L.M., Ramer, M.S., Steeves, J.D. 2005. Setting the stage for functional repair of spinal cord injuries: a cast of thousands. Spinal Cord. 43, 134-61.  31.  Ramer, L.M., McPhail, L.T., Borisoff, J.F., Soril, L.J., Kaan, T.K., Lee, D., Saunders, J.W., Hwi, L.P., Ramer, M.S., 2007. Endogenous TrkB ligands suppress functional mechanosensory plasticity in the deafferented spinal cord. J. Neurosci. 27, 5812-5822.  32.  Ramer, M.S., Duraisingam, I., Priestley, J.V., McMahon, S.B. 2001. Two-tiered inhibition of axon regeneration at the dorsal root entry zone. J Neurosci. 21, 2651-60.  33.  Scholz, J., Abele, A., Marian, C., Häussler, A., Herbert, T.A., Woolf, C.J., Tegeder, I. 2008. Low-dose methotrexate reduces peripheral nerve injury-evoked spinal microglial activation and neuropathic pain behavior in rats. Pain. 138, 130-42.  34.  Sivasankaran, R., Pei, J., Wang, K.C., Zhang, Y.P., Shields, C.B., Xu, X.M., He, Z. 2004. PKC mediates inhibitory effects of myelin and chondroitin sulfate proteoglycans on axonal regeneration. Nat Neurosci.7, 261-8.  35.  Soril, L.J., Ramer, L.M., McPhail, L.T., Kaan, T.K., Ramer, M.S. 2008. Spinal brainderived neurotrophic factor governs neuroplasticity and recovery from coldhypersensitivity following dorsal rhizotomy. Pain. 138, 98-110.  36.  Stirling, D.P., Khodarahmi, K., Liu, J., McPhail, L.T., McBride, C.B., Steeves, J.D., Ramer, M.S., Tetzlaff, W. 2004. Minocycline treatment reduces delayed oligodendrocyte death, attenuates axonal dieback, and improves functional outcome after spinal cord injury. J Neurosci. 2, 2182-90  37.  Tikka, T., Fiebich, B.L., Goldsteins, G., Keinanen, R., Koistinaho, J. 2001. Minocycline, a tetracycline derivative, is neuroprotective against excitotoxicity by inhibiting activation and proliferation of microglia. J Neurosci. 21, 2580-8. 180  38.  Weidner, N., Ner, A., Salimi, N., Tuszynski, M.H. 2001. Spontaneous corticospinal axonal plasticity and functional recovery after adult central nervous system injury. Proc Natl Acad Sci U S A. 98, 3513-8.  39.  Wong, J.W., McPhail, L.T., Brastianos, H.C., Andersen, R.J., Ramer, M.S., O'Connor, T.P. 2008. A novel diketopiperazine stimulates sprouting of spinally projecting axons. Exp Neurol. 214, 331-40.  40.  Z'Graggen, W.J., Metz, G.A., Kartje, G.L., Thallmair, M., Schwab, M.E. 1998. Functional recovery and enhanced corticofugal plasticity after unilateral pyramidal tract lesion and blockade of myelin-associated neurite growth inhibitors in adult rats. J Neurosci. 18, 4744-57.  181  CHAPTER 5  Conclusion and Discussion  182  5.1. SUMMARY OF RESULTS In brief, the studies described in this thesis so far have demonstrated that the natural diketopiperazine, DKP101516, has robust axon outgrowth promoting activity in various neuronal cultures in vitro, and in the sensory circuitry in vivo. Specifically, Chapters 2 and 3 revealed that DKP101516 stimulated robust axon outgrowth and branching in vitro in both peripheral dorsal root ganglion neuron and cortical neuron cultures. In addition, spinal infusion of DKP101516 following septuple dorsal rhizotomy resulted in robust intraspinal axon sprouting. Further evaluation (Chapter 4) using a well characterized C7/C8 DRI model provided an affirmative link between DKP101516 mediated axon sprouting and sensory recovery. The positive results so far strongly suggest that DKP101516 may be a possible therapeutic for dorsal root injury (brachial plexus avulsion) and potentially incomplete spinal cord injury.  Although DKP101516 may appear to be a promising drug to stimulate axon sprouting and functional recovery following dorsal root injury, DKP101516 also demonstrates long-term detrimental effects. In the discussion below, I will discuss the positive and negative effects of DKP101516 treatment, and whether DKP101516 can be used to as a potential therapeutic for dorsal root injury and/or spinal cord contusion.  5.2. DISCUSSION 5.2.1. Effects of DKP101516 on the Sensory Circuitry The natural diketopiperazine DKP101516 identified in our study demonstrates robust neurite outgrowth promoting activity in vitro, as well as the ability to stimulate neurite outgrowth in the presence of inhibitory molecules in the CNS including MAIPs and CSPGs. Surprisingly, DKP101516 failed to promote axon regeneration following dorsal root injury possibly because of 183  its stimulatory effects on astrogliosis, which unfortunately strengthened the regenerative barrier. Fortunately for us, afferent regeneration following DRI may not necessarily be the only mechanism for functional recovery in the dorsal rhizotomy model. Indeed, numerous studies with the dorsal root injury model have demonstrated that the sprouting and reorganization of the sensory circuitry can result in sensory recovery (Darian-Smith, 2004; Ramer et al., 2004). We therefore analysed the effects of DKP101516 spinal infusion on axon sprouting in the deafferented dorsal horn.  Preliminary characterization of DKP101516 revealed that it can stimulate robust sprouting of a number of axonal populations in the sensory circuitry, including descending supraspinal projections as well as nociceptive and mechanosensitive afferents. True to our expectations, further behavioral analysis revealed that within the first 7 days following C7/8 DRI, DKP101516 promoted significant improvement in mechanosensory function that was lost following C7/C8 dorsal root injury. Intriguingly, the beneficial effects of DKP101516 on axon sprouting and sensory recovery appear to be short-lived. Specifically, the effects of DKP101516 on axon sprouting and sensory recovery were lost at later time points (day 14-23) following C7/C8 DRI. Since DKP101516 infusion with osmotic mini-pump only lasts for 14 days, a possible conclusion is that continuous infusion of DKP101516 may be necessary for sustained axon sprouting and functional recovery, and therefore the effects would be reversed upon depletion of DKP101516. However, because there is an enhanced and prolonged reactivity of microglia/macrophage (up to 23 days) an alternative explanation is that long-term DKP101516 infusion may ultimately promote detrimental effects characterized by axon loss, microglia/macrophage reactivity and decline in functional recovery.  184  5.2.2. Non-neuronal Effects of DKP101516 Studies in both Chapters 3 and 4 revealed that DKP101516 might indirectly exert inhibit axon regeneration in the short term, and axon sprouting in the long-term. These were attributed to the extraneous effects of DKP10516 on astrogliosis and microglia/macrophage reactivity respectively. Although astrogliosis and microglia/macrophage reactivity are classically regarded as negative events that hinder axon regeneration and sprouting (Ramer et al., 2001; McPhail et al., 2005; McPhail et al., 2007), a number of studies have revealed that these events can also promote axon regeneration/sprouting (Krenz & Weaver, 2000; Batchelor et al., 2002; Faulkner et al., 2004). Below, we will discuss the role of DKP101516 in the context of the multi-faceted role of astrogliosis and microglia/macrophage reactivity in response to CNS injury.  1) Astroglial Scarring Studies in Chapter 3 demonstrated that although DKP101516 can stimulate robust sprouting of the injured afferents within the peripheral root, DKP101516 failed to stimulate the regeneration of injured afferents back into the spinal cord. The most likely reason for this failure is that DKP101516 also enhances astrogliosis, which strengthens the physical and molecular barrier that hinders axon regeneration. Specifically, DKP101516 not only enhanced the density of GFAP positive astrocytic processes in the DREZ, but also increased the expression of the inhibitory CSPG neurocan in an injury-dependent manner. Since astrogliosis has been classically defined as one of the major barriers hindering axon regeneration, a reasonable interpretation is that DKP101516 mediated astrogliosis inhibited the afferent regeneration into the spinal cord. However, another study has also demonstrated a positive role of astrogliosis in neuroprotection and wound healing (Faulkner et al., 2004). Perhaps the culprit may not be the DKP101516 mediated increase in astrogliosis per se, but rather DKP101516 mediated upregulation of CSPGs  185  such as neurocan.  2) ED-1 immunoreactivity Studies in Chapter 4 revealed that the sprouting effects of DKP101516 in the dorsal horn occurred only transiently, while prolonged DKP101516 treatment resulted in axon loss as well as enhanced ED-1 immunoreactivity indicating increased microglia/macrophage activation. Microglia/macrophage activation in the dorsal root injury model functions to clear axonal debris from injured afferents, in the process known as Wallerian degeneration. The classical view of microglia/macrophage activation is detrimental, and is supported by previous studies demonstrating the role of microglia/macrophage activation on axon dieback (McPhail et al., 2007, Horn et al., 2009). Based on the classical model, we suggest that DKP101516 mediated increase in microglia/macrophage activation in the long-term likely results in the observed loss of axon sprouts in the dorsal horn. However, a number of studies have revealed that activated microglia/macrophage produces neurotrophins and cytokines that apparently promote axon growth (Krenz & Weaver, 2002; Cafferty et al., 2004), suggesting that DKP101516 mediated microglia/macrophage reactivity may not necessarily be responsible for axon loss in the dorsal horn. Alternatively, since cell death can also promote microglia/macrophage reactivity, DKP101516 mediated increase in ED-1 immunoreactivity can also be interpreted as an indication of toxicity.  5.2.3. DKP101516 Mechanism of Action 1) Activation of PI3K Signaling The high-throughput screen in Chapter 2 has resulted in the identification of a marine extract derived diketopiperazine, DKP101516, which is shown in Chapter 3 to mediate neurite  186  outgrowth promoting effect through the PI3K pathway. However, it is unclear whether DKP101516 activates PI3K signaling directly or via some other upstream pathway. Intriguingly, marine compounds include a wide range of bioactivity that targets intracellular signaling components such as calpain and protein tyrosine phosphatases (Zeng et al., 2008; Sun et al., 2007), or by modulating cellular signaling by regulating intracellular levels of calcium (McLeland et al., 2004; Louzao et al., 2007, 2008; Ares et al., 2009). Therefore, it is likely that DKP101516 may activate PI3K signaling by targeting PI3K directly, or by inhibiting PI3K antagonists SHIP and PLC-γ. On the other hand, marine compounds including the DKP family regulate the cell cytoskeleton by modulating membrane potential (McCleland et al., 2004; Louzao et al., 2007, 2008). This can be achieved either by inhibiting Na+/K+ ATPase pump or K+ channels (Louzao et al., 2007, 2008). The resulting depolarization enhances calcium influx via the voltage-gated calcium channels, which activates calcium effectors (such as PKC, calpain or calcineurin) that can modulate F-actin dynamics (Shim et al., 2005; Wang et al., 2005; Li et al., 2005; Henley & Poo, 2004). Quite possibly, our diketopiperazine DKP101516 may mediate its neurite outgrowth promoting effects possibly by modulating neuronal membrane potential, conceivably by blocking K+ channels or inhibiting Na+/K+ ATPase pump. The resulting depolarization and calcium influx generates baseline calcium levels that may favor PI3K-Akt signaling (Zheng et al., 2008), which enhances growth cone motility and subsequent neurite outgrowth (Menager et al., 2003).  2) Mechanism behind Axon Loss In Chapter 4, prolonged intrathecal DKP101516 treatment in vivo was observed to result in the apparent loss of axon sprouts. It is possible that DKP101516 mediated PI3K activation in the long-term may stimulate microglia/macrophage activation, which may contribute to axon loss.  187  Alternatively, our present study revealed that DKP101516 is poorly soluble in aqueous solution, and is optimally effective at a high concentration of 32 micromolar. Therefore, the low solubility of DKP101516 in intrathecal space and the potential toxicity of its high effective concentration may also contribute to the observed detrimental effects in the long-term .  5.2.4. Therapeutic Applications- Promise and Problems Dorsal root injury in humans, often referred to brachial plexus avulsion, results in loss in mechanosensory function as well as the development of neuropathic pain. Recognized mechanisms leading to sensory recovery include afferent regeneration and sprouting and/or reorganization of intact sensory projections (Darian-Smith, 2004; Ramer et al., 2001). Both afferent regeneration and axon sprouting are hindered by the expression of inhibitory molecules in the CNS, and by the delayed microglia/macrophage reactivity resulting from Wallerian degeneration of the injured roots (Ramer et al., 2001; McPhail et al., 2005; McPhail et al., 2007). In Chapter 4, the spinal infusion of DKP101516 following dorsal root injury promoted transient axon sprouting and accelerated mechanosensory recovery, which is eventually masked by the negative effects of DKP101516 in the long-term, characterized by axon loss and microglia/macrophage reactivity. Overall, DKP101516 demonstrates robust short-term effects on axon sprouting and sensory recovery following DRI in vivo, rendering it a promising therapeutic in the short-term. However, it is unclear whether short-term DKP101516 is sufficient to produce lasting effects on sensory recovery following dorsal root injury.  Similar to DRI, contusive spinal cord injury (SCI) also results in a similar series of aforementioned glial events that hinder axon regeneration and sprouting. The mechanism to elicit sensory and motor recovery following SCI is unclear, but a number of studies demonstrate that  188  sprouting and reorganization of intact spinal circuitry corresponded with improvements in motor function (Barritt et al., 2006; Massey et al., 2006). Given the robust sprouting effects of DKP101516 in DRI model, intrathecal treatment of DKP101516 following SCI is likely to enhance axon sprouting as well as robust recovery in sensory and motor function. These effects may however be hindered in the long-term by the detrimental effects of DKP101516 characterized by axon loss and microglia/macrophage reactivity.  The ultimate question is whether DKP101516 can be used as a therapeutic for CNS injuries. The answer depends on the nature of DKP101516 effects, particularly whether its detrimental effects can be effectively modulated to favor its beneficial effects on axon sprouting and functional recovery following DRI and/or SCI. To determine whether the therapeutic use of DKP101516 can be salvaged, further investigation is needed to elucidate the mechanism underlying the effects of DKP101516 mediated axon loss and microglia/macrophage reactivity. If prolonged DKP101516 directly mediates axon loss or stimulates microglia/macrophage reactivity, then DKP101516 may not be suitable for use as a therapeutic. If on the other hand DKP101516 mediates detrimental effects in the long-term merely because of its low solubility and/or interactions with the mildly toxic vehicle DMSO/PEG, we could easily salvage the therapeutic use of DKP101516. This can be achieved by chemical reformulations to make DKP101516 more soluble in non-toxic vehicles (i.e. saline), and more potent such that only low effective concentrations are required for axon sprouting effect. Therefore, until we elucidate the mechanism behind the detrimental effects of DKP101516 in vivo, DKP101516 remains to be a promising therapeutic to promote functional recovery following DRI/SCI.  189  5.3. FUTURE DIRECTIONS Our studies have demonstrated that despite the positive effects of DKP101516 on axon sprouting and mechanosensory following DRI, the detrimental effects of DKP101516 in the long-term may render it unsuitable for therapeutic use. In order to overcome this problem, future studies will be focused on 1) developing new strategies to minimize the detrimental effects of DKP101516, or 2) modifying the high-throughput screen to improve the success of finding a neurite outgrowth promoting compound with minimal extraneous or detrimental effects.  5.3.1. Elucidating the Mechanism of DKP101516 Mediated Microglia/Macrophage Reactivity In vivo In order to minimize the detrimental effects of DKP101516 in vivo, it is crucial to elucidate the mechanism through which DKP101516 enhances microglia/macrophage reactivity in vivo. As mentioned above, microglia/macrophage reactivity can be induced directly by DKP101516, or indirectly via DKP101516 mediated cell death. To determine whether DKP101516 directly stimulates microglia/macrophage reactivity, ex vivo binding assays or signaling assays can be used to determine whether DKP101516 stimulates signaling pathways that promote microglia/macrophage reactivity. Further in vitro analysis using microglia/macrophage cultures can also be used to assess the direct effects of DKP101516 on microglia/macrophage reactivity. It may also be worth exploring whether DKP101516 mediated PI3K activation might stimulate microglia/macrophage reactivity.  If DKP101516 does not have any direct influence on microglia/macropahge reactivity, the next stage may be to evaluate the toxicity and solubility of DKP101516 in vitro in various neuronal cultures. It may also be worth looking at whether prolonged DKP101516 treatment induces  190  excitotoxicity as described above, by assessing the effects of DKP101516 on neuronal membrane potential and ion flux with electrophysiological analysis such as whole cell or patch clamp recordings. Further DKP101516 mediated changes in baseline intracellular calcium concentration can be assessed with fluorescent calcium indicators such as Fluo-3. Calcium chelators such as BAPTA could then be employed to determine whether DKP10516 mediates its short-term neurite outgrowth and long-term detrimental effects through calcium signaling.  5.3.2. Minimizing the Detrimental Effects of DKP101516 1) Avoidance A simple and feasible approach is to apply DKP101516 not at the site of injury where astrogliosis and microglia/macrophage reactivity could be exacerbated, but closer to the deinnervated targets where DKP101516 can stimulate the compensatory sprouting of nearby intact axons and functional recovery. Indeed, studies by Massey et al (2006) have shown that following dorsal column lesion, application of chondroitinase ABC at the deinnervated brain stem targets resulted in compensatory axon sprouting of intact contralateral projections and functional recovery (Massey et al., 2006). We therefore suggest that DKP101516 can be used in a similar fashion to stimulate compensatory sprouting and functional recovery following spinal cord injury.  Another strategy may be to limit the duration of intrathecal DKP101516 treatment, thus minimizing the effects of prolonged DKP101516 treatment on microglia/macrophage reactivity. To achieve this, one approach is to spinally infuse DKP101516 continuously for up to 7 days post DRI prior to the onset of microglia/macrophage activation. However, a potential problem is that continuous intrathecal DKP101516 infusion may be necessary to prolong the sprouting  191  effects of DKP101516. As a consequence, early termination of DKP101516 treatment at 7 days post DRI could severely curtail its beneficial effects on axon sprouting and mechanosensory recovery. Therefore, a more effective strategy may be to apply DKP101516 with a pulsatile delivery method previously used to minimize the toxicity of prolonged intrathecal polyethylene glycol (PEG) infusion (Cole & Shi, 2005), specifically by intrathecally delivering a bolus of DKP101516 at various time points during the course of the experiment. The pulsatile delivery method would minimize the detrimental effects of prolonged DKP101516 exposure, while prolonging the beneficial effects of DKP101516 in the long run.  2) Combinatorial Approach One way to minimize the detrimental effects of DKP101516 is to combine DKP101516 treatment with other interventions. For instance, DKP101516 could be combined with interventions that eliminate inhibitory CSPGs from the glial scar such as chondroitinase ABC treatment; therefore reducing the negative influence of DKP101516 mediated astrogliosis and CSPG expression on axon regeneration. Conversely, DKP101516 treatment can also be combined with astrogliosis inhibitors such as epithelial growth factor receptor (EGFR) inhibitors to minimize astrogliosis and CSPG expression. Lastly, DKP101516 can be complemented with minocycline in order to minimize microglia/macrophage activation and its detrimental effects on spinal cord repair.  3) DKP101516 Modifications DKP101516 can also be modified chemically to render it more suitable for intrathecal applications. Since DKP101516 used in our studies was poorly soluble in aqueous solution and quite possibly toxic, we suggest chemical reformulations and modifications are required to  192  rectify these problems. Specifically, DKP101516 can be chemically reformulated to become soluble in non-toxic vehicles such as saline, and to remain soluble as it diffuses into intrathecal space. Furthermore, DKP101516 can be modified chemically to enhance its potency such that a lower effective concentration can produce the required axon sprouting effects. The enhanced potency would minimize solubility issues and potential toxicity associated with high concentrations of DKP101516. Lastly, since DKP101516 appears to demonstrate extraneous effects on astrogliosis and possibly microglia/macrophage reactivity, these effects could be eliminated by future chemical modification to render DKP101516 more target-specific.  5.3.3. Improvements on the High-Throughput Screen In theory, it is possible to salvage the therapeutic use of DKP101516 to stimulate CNS repair. However, given the immense efforts to implement the abovementioned strategies to hopefully render DKP101516 a useful therapeutic, it may be more feasible to consider other candidate compounds in the marine extract library. In order to improve our chances of finding a neurite outgrowth-promoting compound with minimal toxicity or extraneous glial effects, we suggest implementing the following modifications to our high-throughput screen.  Since DKP101516 identified from the high-throughput screen demonstrated detrimental effects in vivo, it may be prudent to determine whether the screening technology can be improved in the future to eliminate candidate compounds that may have the abovementioned undesired effects. Perhaps a potential downfall of our screening technology is that the screen is based on the conserved mechanism underlying cell migration and growth cone motility. Unfortunately, this conserved mechanism may be also responsible for other undesired glial effects in the spinal cord including astrogliosis and microglia/macrophage reactivity (Irino et al., 2008; John et al., 2004;  193  Etienne-Manneville & Hall, 2001). In order to minimize the chance of identifying and evaluating compounds with negative glial consequences similar to those observed in DKP101516, a feasible solution is to include additional screens to further evaluate non-neuronal effects of candidate compounds.  5.4. CONCLUSION Repairing the central nervous system is a complex problem for which there is no clear solution. Current interventions involve recreating an environment similar to that during CNS development or the adult PNS in which axon repair can occur successfully. Although these interventions have yielded promising results, the beneficial effects are often limited. Reducing the non-permissive nature of the adult CNS is a daunting task given the myriad inhibitory molecules in the CNS that are functionally redundant (Schwab, 2004; Deumanns et al., 2008). Often growth cone manipulations such as prolonged neurotrophin treatment or spinal infusion of ROCK inhibitors can result in deleterious consequences such as p75 induced cell death and enhanced astroglial scarring respectively (Scott et al., 2005; Chan et al., 2005). Likewise, our novel neurite outgrowth-promoting compound DKP101516 yielded promising results in vivo, but similar detrimental effects in the long-term. Specifically, we found that DKP101516 stimulates transient axon sprouting and functional recovery following dorsal rhizotomy via PI3K signaling, suggesting that growth cone manipulation is a feasible approach to identify novel drugs to stimulate functional reconnections in the CNS. However, the deleterious effects of DKP101516 in the long-term strongly indicates that signaling components such as PI3K may activate unknown signaling pathways that may contribute to undesired extraneous effects in vivo. Lastly, although DKP101516-mediated axonal sprouting in the short-term resulted in subtle improvements in low-threshold mechanosensory function, it is unclear whether the sprouting 194  effects of DKP101516 in other axonal populations would result in undesired behavioral effects such as neuropathic pain and autonomic dysfunction. 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