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Intrinsic neuronal determinants of neurite regrowth Borisoff, Jaimie F. 2002

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INTRINSIC N E U R O N A L DETERMINANTS OF NEURITE R E G R O W T H by J A M I E F. BORISOFF B A . S c , The University of British Columbia, 1994 A THESIS SUBMITTED LN PARTIAL FULFILLMENT OF THE REQUIREMENTS FOR THE DEGREE OF DOCTOR OF PHILOSOPHY i n THE F A C U L T Y OF G R A D U A T E STUDIES (Graduate Program in Neuroscience) We accept this thesis as conforming to the required standard THE UNIVERSITY OF BRITISH C O L U M B I A September 2002 © Jaimie F. Borisoff, 2002 In p resen t i ng this thesis in partial fu l f i lment of t he requ i remen ts fo r an a d v a n c e d d e g r e e at the Univers i ty o f Bri t ish C o l u m b i a , I agree that t he Library shal l m a k e it f reely avai lable fo r re fe rence and s tudy. I fur ther agree that p e r m i s s i o n fo r ex tens i ve c o p y i n g o f this thesis fo r scho la r l y p u r p o s e s m a y b e g ran ted b y the h e a d o f m y depa r tmen t o r by his o r her representa t ives . It is u n d e r s t o o d that c o p y i n g o r pub l i ca t i on of this thesis for f inancia l ga in shal l no t b e a l l o w e d w i t h o u t m y wr i t t en p e r m i s s i o n . D e p a r t m e n t The Univers i ty o f Brit ish C o l u m b i a V a n c o u v e r , C a n a d a Da te D E - 6 (2/88) 11 ABSTRACT Traumatic injury to the central nervous system (CNS) results in neurological deficits, in part, due to axonal regeneration failure. This is functionally exemplified in spinal cord injury by motor and sensory paralysis. Regeneration failure has been attributed to several factors, including intrinsic neuronal limitations to regeneration, as well as numerous inhibitory molecules present in the injured CNS environment. The impact of intrinsic neuronal factors is investigated here. A transition from a permissive to a restrictive repair period exists in the developing chick on approximately embryonic day (E) 13, possibly due to the formation of an extrinsic inhibitory environment preventing axonal growth, and/or an intrinsic inability of mature neurons to regenerate. By fluorescent labeling of brainstem-spinal neurons in ovo, I was able to subsequently track the capacity of specific populations of young (E8) versus mature (El 7) brainstem-spinal neurons to regrow neurites (i.e. presumptive axons) in vitro on permissive growth substrates. When cultured on E8, robust neurite growth was observed from all brainstem populations examined. In contrast, when cultured on E l 7 , significant neurite growth was seen only from raphe-spinal neurons. Thus, brainstem-spinal neurite regrowth may dependent on both neuronal age and phenotype, suggesting that intrinsic neuronal properties may contribute to axonal regeneration failure. Because regeneration may depend on intrinsic neuronal properties, it may be beneficial to pharmacologically enhance the axonal growth capacity of neurons. Injured neurons respond characteristically (i.e. growth cone collapse or neurite retraction) to various molecules that inhibit axonal growth, including myelin proteins and chondroitin sulfate proteoglycans (CSPG). Accordingly, it is possible that intracellular signaling from several inhibitory molecules converge onto a common regulatory pathway of axonal growth inhibition, i.e. the Rho-GTPase. I tested in vitro whether pharmacological inhibition of a major downstream effector of Rho, Rho-kinase (ROCK), promoted neurite outgrowth of dorsal root ganglia (DRG) neurons grown on inhibitory substrates of aggrecan (a CSPG), myelin, and spinal cord cryosections. Indeed, R O C K inhibition promoted neurite outgrowth several-fold, as well as significantly altering the actin-based morphology of growth cones. The data support the notion that suppression of Rho-pathway activity may be a viable therapeutic avenue for enhancing axonal regeneration within the injured adult CNS. Ill T A B L E OF C O N T E N T S A B S T R A C T i i T A B L E OF CONTENTS i i i LIST OF FIGURES vi LIST OF ABBREVIATIONS vii STATEMENT OF ORIGINAL CONTRIBUTIONS viii A C K N O W L E D G E M E N T S ix C H A P T E R 1: G E N E R A L INTRODUCTION 1 EXTRINSIC CNS E N V I R O N M E N T A L LIMITATIONS TO REGENERATION 4 Myelin in White Matter Tracks Inhibits Axonal Growth 4 The Glial Scar Inhibits Axonal Growth 5 INTRINSIC N E U R O N A L LIMITATIONS TO REGENERATION 7 Age-Dependent Limitations to Regeneration 7 Cell-Specific Limitations to Regeneration 8 The Cell Body Response 10 A X O N A L GROWTH A N D RHO-GTPases 11 Axonal Growth Mechanisms 11 Actin Cytoskeletal Changes are Mediated by Rho-GTPases 15 Inhibitory Molecules Act Through Rho-GTPases 20 R A T I O N A L E A N D HYPOTHESES 24 IN VITRO MODELS OF SPINAL CORD REGENERATION 26 Explants of Identified Brainstem-Spinal Neurons 28 Explants of Dorsal Root Ganglia 31 C H A P T E R 2: R A P H E - S P I N A L NEURONS D I S P L A Y A N A G E - D E P E N D E N T D I F F E R E N T I A L C A P A C I T Y F O R N E U R I T E O U T G R O W T H C O M P A R E D TO O T H E R B R A I N S T E M - S P I N A L P O P U L A T I O N S 38 INTRODUCTION 39 iv M A T E R I A L S A N D METHODS ; 42 In Ovo Dye Application 42 Tissue Culture 42 Neurite Outgrowth Assay 43 Immunohistochemistry 43 Cell Viability Counts 44 RESULTS 44 Brainstem-Spinal Neuron Labeling 44 Brainstem-Spinal Neurite Outgrowth 45 Brainstem-Spinal Neuron Survival 53 5-HT Immunohistochemistry 56 DISCUSSION 59 Development of an In Vitro Brainstem-Spinal Neuron Model of Regeneration 59 Brainstem-Spinal Neurite Outgrowth is Age-Dependent 60 Differential Outgrowth From Mature Brainstem-Spinal Neurons 62 C H A P T E R 3: SUPPRESSION OF R H O - K I N A S E A C T I V I T Y P R O M O T E S N E U R I T E G R O W T H O N INHIBITORY CNS SUBSTRATES 64 INTRODUCTION 65 M A T E R I A L S A N D METHODS '. 67 Tissue Culture and Neurite Outgrowth Assay 67 Myelin Substrates 68 Spinal Cord Cryo-Sections 68 Growth Cone Measurements 69 Immunohistochemistry 70 Western Blots 70 R O C K Kinase Assay 70 RESULTS 71 R O C K is Expressed in Chick D R G 71 R O C K is Inhibited by Y-27632 71 Aggrecan Substrates Inhibit the Growth of D R G Neurites 74 V R O C K Inhibition Stimulates Neurite Outgrowth on Aggrecan 74 R O C K Inhibition Stimulates Neurite Outgrowth on Myelin and Spinal Cord 79 R O C K Inhibition Alters Growth Cone Morphology 84 DISCUSSION 94 R O C K Suppression Promotes Neurite Growth on Aggrecan 94 Rho-GTPases Regulate Neurite Growth on Inhibitory Substrates 95 R O C K Suppression Alters Growth Cone Morphology 95 C H A P T E R 4: G E N E R A L DISCUSSION 98 NEURITE GROWTH F R O M BRAINSTEM-SPINAL NEURONS 99 Comparison of Brainstem-Spinal Neurite Growth With In Vivo Models 99 Why are Mature Neurons Less Capable of Axonal Growth? 100 Why are Raphe-Spinal Neurons More Plastic? 103 PROMOTION OF A X O N A L GROWTH B Y T R E A T M E N T TO THE N E U R O N 106 Enhancement of the Intrinsic Axonal Growth Capacity of Neurons 106 Modulating Cyclic Nucleotide Levels to Enhance Regeneration Capacity 107 Model Limitations, Remedies, and Future Studies 109 Significance 114 Concluding Remarks 115 FINIS 116 R E F E R E N C E S 117 VI LIST OF F IGURES Figure 1-1. Schematic drawings of a growth cone characteristics 13 Figure 1-2. Schematic diagram of the Rho-signaling pathway in a growth cone 17 Figure 1-3. Schematic of inhibitory myelin components and their putative receptors 22 Figure 1-4. The chick brainstem-spinal neuron populations 29 Figure 1-5. Schematic diagram of the in vitro brainstem-spinal neuron outgrowth assay.... 32 Figure 1-6. An example of neurite outgrowth from a whole E9 chick dorsal root ganglion. 34 Figure 2-1. Retrograde D i l labeling of chick brainstem-spinal neurons 46 Figure 2-2. Differential neurite outgrowth of young and mature raphe- and vestibulospinal neurons grown in serum-free media on laminin substrates 48 Figure 2-3. Comparison of neurite growth from populations of brainstem-spinal neurons. . 51 Figure 2-4. D i l labeled E l 7 vestibulospinal neurons 54 Figure 2-5. Serotonergic immunoreactivity in raphe-spinal neurites 57 Figure 3-1. R O C K immunoreactivity 72 Figure 3-2. Aggrecan inhibits chick D R G neurite outgrowth 75 Figure 3-3. Y-27632 promoted neurite outgrowth from E9 NGF-treated D R G explants grown on inhibitory aggrecan substrates 77 Figure 3-4. Y-27632 (25 pM) promoted neurite outgrowth from E9 NGF-treated D R G explants grown on substrates of increasing concentrations of aggrecan 80 Figure 3-5. Y-27632 (25 pM) promoted neurite outgrowth from BDNF- and NT-3-treated D R G explants grown on inhibitory aggrecan substrates 82 Figure 3-6. Y-27632 promoted neurite outgrowth from E l 5 NGF-treated D R G explants grown on inhibitory substrates of myelin membrane fragments 85 Figure 3-7. Y-27632 promoted neurite outgrowth from E l 5 NGF-treated D R G explants grown on inhibitory substrates of longitudinal adult rat spinal cord cryo-sections 87 Figure 3-8. High magnification images of growth cone morphology 90 Figure 3-9. Inhibitory aggrecan substrates stimulate actin bundles in growth cones 92 Figure 4-1. R O C K inhibition stimulates the migration of glial cells from brainstem explants grown on aggrecan substrates 112 LIST OF A B B R E V I A T I O N S 5-HT 5 -hydroxy-tryptamine (serotonin) A N O V A analysis of variance ATP adenosine triphosphate BDNF brain derived neurotrophic factor C3 C 3 -exotranferase cAMP adenosine-3',5'-cyclic monophosphate CFDA,SE 5-(and-6)-carboxyfluorescein. diacetate, succinimidyl ester cGMP guanosine-3 ',5 '-cyclic monophosphate CNS central nervous system CSPG chondroitin sulfate proteoglycan db-cAMP dibutyryl-cAMP Dec deleted in colorectal cancer D i l l,l'-dioctadecyl-3,3,3',3'-tetramethylindocarbocyanin e perchlorate D M E M Dulbecco's Modified Eagles Medium DREZ dorsal root entry zone D R G dorsal root ganglia E embryonic day G A G glycosamino-glycan GAP GTPase activating proteins GDP guanosine diphosphate GEF guanosine nucleotide exchange factors GFAP glial fibrillary acidic protein GTP guanosine triphosphate GTPases guanosine triphosphatases L M laminin M A G myelin associated glycoprotein M L C myosin light chain M L C K myosin light chain kinase ML-7 myosin light chain kinase inhibitor M L F medial longitudinal fasciculus mRNA messenger ribonucleic acid NGF nerve growth factor NgR Nogo-66 receptor NT-3 neurotrophin-3 NT-4 neurotrophin-4 OMgp oligodendrocyte-myelin glycoprotein PBS phosphate buffered saline P K A protein kinase A PNS peripheral nervous system R raphe-spinal RAGS regeneration associated genes Red Red nucleus RGC retinal ganglion cell RhoGDI guanosine nucleotide dissociation inhibitor Rm medullary reticulospinal neurons ROKoc Rho-kinase R O C K Rho-kinase Rp pontine reticulospinal neurons SCI spinal cord injury S E M standard error of the mean Sema semaphorin TNF tumor necrosis factor-alpha V vestibulospinal neurons Y-27632 (+)-R-trans-4-(l-Aminoethyl)-N-(4-pyridyl)-cyclohexanecarboxamide Vll l S T A T E M E N T OF O R I G I N A L CONTRIBUTIONS This thesis contains materials that have been previously published or submitted for publication: Borisoff J. F., Pataky D. M . , McBride C. B., Steeves J. D. (2000). Raphe-spinal neurons display an age-dependent differential capacity for neurite outgrowth compared to other brainstem-spinal populations. Exp Neurol 166(1): 16-28. Borisoff J. F., Chan C. C. M . , Hiebert G. W., Oschipok L. , Robertson G. S., Zamboni R., Steeves J. D., Tetzlaff W. (Submitted). Suppression of Rho-Kinase Activity Promotes Axonal Growth on Inhibitory CNS Substrates. The thesis author, Jaimie Borisoff, was the primary researcher responsible for all results presented in the above articles and this thesis. Technical expertise in Western blotting was provided by Carmen Chan. Handling of radioactive materials was performed by Loren Oschipok. Dr. Gordon Hiebert provided cryosections of rat spinal cord tissue. G. Robertson and R. Zamboni are employees of Merck Frosst and kindly provided Y-27632 for use in these experiments. Initial immunohistochemistry expertise was provided by Dr. Chris McBride. Grace Lee, Carmen Chan, Suzanna Chen, and Fengtai Zhang kindly provided technical assistance with computer tracing of neurite outgrowth. A l l work performed by collaborators (other than work done at Merck Frosst) was performed under the supervision and experimental design provided by the thesis author. The above statements and assessment of work performed by the thesis author are correctly stated as above. John D. Steeves, Ph.D. Thesis Supervisor IX A C K N O W L E D G E M E N T S I thank John Steeves for taking the gamble that an engineer would be able to make a meaningful contribution to spinal cord injury research. I have certainly learned a great deal about the pursuit of science under your supervision. Everyone involved in spinal cord research certainly owe you a great debt for your unwavering pursuit in building this research community into the talented force it has become. I thank Wolfram Tetzlaff for his inspirational pursuit of the science of spinal cord injury. I would hope to some day half match your passion for research in whatever field I may find myself. I also thank you for your advice throughout my studies. I would also like to thank the other members of my supervisory committee for their contributions to this work: Drs. Timothy O'Connor, Vanessa Auld, and Tim Murphy. I would like to thank my colleagues at ICORD for making our research environment an enjoyable place to work. Without neglecting the rest of our team, I would especially like to thank Chris McBride, John McGraw, Lowell McPhail, and Ward Plunet for your stimulating conversations. Without you crackers, I would have finished this thesis two years earlier, although at what price to quality of life and philosophical pursuit? I would also like to thank the Natural Sciences and Engineering Research Council of Canada and the Rick Hansen Institute for financial support. Finally, I would especially like to thank my partner, Carrie Linegar, for her love, support, and patience during this odyssey. You are a universal constant in my life, without which any success would seem less meaningful. C H A P T E R 1: G E N E R A L I N T R O D U C T I O N 2 Traumatic injury to the central nervous system (CNS) results in the failure of the affected neurons to regenerate (Schwab and Bartholdi, 1996; Steeves and Tetzlaff, 1998; Fournier and Strittmatter, 2001; Geller and Fawcett, 2002; Plunet et al, 2002). This is dramatically seen in cases of spinal cord injury (SCI), in which paralysis is the usual outcome. As research into SCI progressed, it became increasingly clear that no single factor is solely responsible for the failure of CNS regeneration. After over two decades of research, beginning with the seminal work of Richardson and Aguayo (Richardson et al, 1980), it is apparent that there are two main classes of preventative factors: extrinsic CNS environment factors and intrinsic neuronal factors (Tetzlaff and Steeves, 2000). Intrinsic factors refer to the molecular composition of injured neurons (i.e. pattern of gene expression and activity of proteins), and address the apparent inability of mature CNS neurons to regenerate. The impact of intrinsic factors has been somewhat lessened by the work of Aguayo and colleagues showing that axotomized CNS neurons are able to grow into grafts of peripheral nerve, a permissive environment for axonal growth (Richardson et al, 1980; David and Aguayo, 1981). However, as shown below, the dearth of axonal growth seen under these favorable conditions supports intrinsic propensity as important contributors to regeneration failure. Extrinsic factors address the influence that non-neuronal cells have on regenerating axons. This influence can be found in the scarcity of growth promoting molecules presented to regenerating axons - molecules that are normally expressed in the developing CNS or regenerating peripheral nervous system (PNS); and in the abundance of growth inhibitory molecules associated with both the normal and injured CNS. Of course, these two classes of preventative factors alone do not address all the problems associated with SCI. They only address the current rate-limiting step in most acute, and all chronic, cases of SCI - the need to promote axonal extension through or around the inhibitory environment of the lesion site, or scar, and into the distal spinal cord where appropriate synaptic connections must be made. Another issue in acute SCI injury is the fact that significant neural damage arises from the secondary damage elicited from crush lesions to the CNS (Tator, 1995; Steeves and Tetzlaff, 1998). Much like a bruise to the skin spreads well beyond the initial impact site, axonal damage in SCI spreads further than the initial lesion, resulting in an increase in the functional deficit. Secondary damage is caused by a cascade of inflammation events and cell death (Amar and Levy, 1999). Fortunately, much is known about these pathologies from 3 other experimental systems. As well, significant research into cell death and inflammation is well under way in experimental SCI models, and this research has already led to a clinical application. For instance, the steroid methylprednisolone is now routinely given to SCI patients, although results have been mixed and modest (Bracken and Holford, 2002). Nevertheless, this is an example of the types of treatments that may find their way into a clinical setting before strategies to promote true axonal regeneration arrive. Hopefully, these therapies will result in the sparing of tissue and a reduction in functional deficit, and thus, the promotion of axonal elongation (the focus here) will be an easier task to accomplish. This thesis addresses two separate questions in spinal cord injury research regarding the promotion of axonal extension, both of which deal with intrinsic neuronal factors limiting spinal cord regeneration success. 1) Do mature brainstem-spinal neurons lose the intrinsic ability to regenerate? and 2) By targeting the intrinsic components in axonal growth cones, can multiple inhibitory factors be neutralized with a single, effective treatment? These questions are tackled with the use of in vitro models that directly assay neurite growth extension under varying conditions. To summarize the format of this thesis, I begin by briefly introducing the current knowledge of spinal cord regeneration regarding extrinsic environment factors and intrinsic neuronal factors. This is followed by a broader review of axonal growth mechanisms and intracellular signaling pathways (i.e. Rho-GTPases) that can be interfered with to mitigate inhibitory CNS factors. I then present the rationale behind and hypotheses of the specific questions I addressed experimentally. The introduction is concluded by a description of the model systems that I used to study these problems. Further relevant introductions and the results of my experiments are presented in Chapters 2 and 3 as reproductions of published or submitted articles. Chapter 4 presents a general discussion of my findings and their significance to future studies into spinal cord regeneration. 4 EXTRINSIC CNS E N V I R O N M E N T A L L IMITAT IONS T O R E G E N E R A T I O N In 1980, Aguayo and colleagues (Richardson et al, 1980) repeated landmark experiments first performed in 1911 by F. Tello in the laboratory of Ramon y Cajal (Ramon y Cajal, 1991). These experiments showed that some CNS neurons are capable of regeneration into a peripheral nerve grafted onto the stump of the transected optic nerve. This demonstrated that CNS neurons are capable of regrowth under certain conditions, and that the environment of peripheral nerves is permissive for axonal growth. A further possible implication of this research is that the CNS contains inhibitory molecules that do not support the growth of axons. These early findings have led to significant research into the inhibitory components of the injured CNS, and consequently, into methods of overcoming that inhibition. It is commonly thought that the environment of the injured CNS has the following two broad components: 1) the presence of inhibitory molecules distal to the injury that are normally expressed in CNS myelin; and 2) the formation after injury of a glial scar containing many inhibitory molecules. Myelin in White Matter Tracks Inhibits Axonal Growth The best characterized example of a CNS axonal inhibitor is myelin. It is currently known that five separate factors contribute to myelin inhibition: Nogo, formerly NI-35/250 (Caroni and Schwab, 1988; Bandtlow et al, 1993; Schwab et al, 1993; Spillmann et al, 1998; Chen et al, 2000; GrandPre et al, 2000), myelin associated glycoprotein (MAG) (McKerracher et al, 1994; Filbin, 1995; L i et al, 1996; Tang et al, 1997), arretin (Janani et al, 1998), oligodendrocyte-myelin glycoprotein (OMgp) (Wang et al, 2002), and a chondroitin sulfate proteoglycan (CSPG) (Niederost et al, 1999). Experimental evidence that supports myelin as a major inhibitor of CNS regeneration comes from many studies (for review, see Huber and Schwab, 2000; Qiu et al, 2000; Fournier and Strittmatter, 2001; Grandpre and Strittmatter, 2001), including in vitro neurite outgrowth assays showing growth inhibition by myelin-derived CSPG (Niederost et al, 1999), M A G (McKerracher et al, 1994; Mukhopadhyay et al, 1994), and OMgp (Wang et al, 2002). Axonal growth in vivo has been modestly promoted by the application of monoclonal antibody IN-1 (Schnell and 5 Schwab, 1990, 1993; Bregman et al, 1995; Merkler et al, 2001), an antibody that binds to Nogo, albeit with questionable specificity, resulting in the suppression of myelin inhibition. Very recently, the Strittmatter group reported corticospinal axon regeneration in a rat spinal cord hemisection model by the intrathecal administration of a specific antagonist peptide to the Nogo receptor (GrandPre et al, 2002). Another method, that targets myelin inhibition in its entirety, is the immunological removal of myelin by the application of serum complement and an antibody to galactocerebroside. This method has promoted the growth of descending axons in the injured spinal cords of embryonic chicks (Keirstead et al, 1992) and adult rats (Dyer et al, 1998), as evidenced by retrograde labeling. Along the same lines, a novel vaccine approach has been developed (Huang et al, 1999; David and Ousman, 2002). Mice were immunized to a gross myelin preparation prior to undergoing a spinal cord lesion paradigm. Compared to controls, vaccinated mice were reported to have extensive growth of corticospinal fibres. These examples highlight the significant role that inhibitory components of myelin may play in CNS regeneration failure. The Glial Scar Inhibits Axonal Growth Another major source of inhibitory molecules is found in the glial scar that forms after injury. The cellular composition of the scar comes from activated astrocytes and macrophages, as well as other invading cells (Fitch and Silver, 1997a, b; Stichel and Muller, 1998; Fawcett and Asher, 1999; Shearer and Fawcett, 2001). Putative inhibitory molecules that are expressed in the scar include various CSPGs (Snow et al, 1990; McKeon et al, 1995; Bovolenta et al, 1997; Fitch and Silver, 1997a), collagen-fV (Stichel et al, 1999b; Stichel et al, 1999a), tenascin (McKeon et al, 1991; Zhang et al, 1995), class-3-semaphorins (Pasterkamp et al, 1999; Pasterkamp and Verhaagen, 2001; De Winter et al, 2002), and Eph-3B (Miranda et al, 1999). J. Silver and colleagues gathered perhaps the most significant evidence of the lesion scar as a major growth inhibitor (Davies et al, 1997; Davies et al, 1999). They transplanted dissociated embryonic or adult dorsal root ganglia (DRG) cells into the spinal cords of adult rats. Long distance growth of D R G axons was observed along the myelinated white matter tracks when the transplant procedure was performed with minimal injury to the CNS. Regenerating axons grew until they encountered a CSPG-immunoreactive scar that formed at the site of an applied lesion. Furthermore, in 6 cases where the procedure caused scarring at the transplant site, the D R G axons became inhibited and trapped, unable to extend past the scar. This inhibition also correlated with the expression of CSPG in the scar that formed. Other evidence supporting the scar as inhibitor has been found in studies that aimed to neutralize the synthesis or activity of scar constituents. The suppression of collagen-TV deposition at a CNS lesion has been shown to promote the growth of injured axons through the scar (Stichel et al, 1999b; Stichel et al, 1999a). As well, enzymatic cleavage of the G A G chains on CSPG molecules helps to render those molecules non-inhibitory to growth in vitro (Zuo et al, 1998b; Zuo et al, 1998a), and promotes regeneration in vivo (Krekoski et al, 2001; Bradbury et al, 2002). The lesion scar contains several inhibitory molecules and evidence shows that by targeting those molecules axonal growth can be promoted. In summary, it is well established that a multitude of inhibitory molecules are expressed in the environment of injured axons. Because of the abundant evidence collected regarding environmental limitations to regeneration, a common experimental technique is to manipulate the CNS parenchyma in order to counteract these inhibitory molecules. It appears that efforts to neutralize many inhibitory molecules at once will be a difficult task to accomplish. As well, modifying the CNS environment directly may have unforeseen consequences to uninjured axons, possibly further exacerbating the injury. It is also currently unknown how many and which of the inhibitors may need to be overcome. With these reasons in mind, an ideal way to attack this problem would be to target the neuron directly, rather than attempting to neutralize many extrinsic inhibitory molecules at once. As described below, the axonal growth cone contains intracellular signaling pathways intrinsic to the neuron that mediate the negative effects of inhibitory molecules. By targeting possible convergence points in these intracellular signaling pathways, it may be possible to release the neuron from the effects of several inhibitory factors with a single treatment. The intrinsic neuronal characteristics that both limit regeneration success and provide possible therapeutic targets are described in the following sections. 7 INTRINSIC N E U R O N A L L IMITATIONS T O R E G E N E R A T I O N CNS neurons are capable of axonal growth into the permissive environment of a peripheral nerve graft (Richardson et al, 1980; David and Aguayo, 1981; Kobayashi et al, 1997; Ye and Houle, 1997), demonstrating that CNS neurons are intrinsically able to regenerate. Unfortunately, CNS regenerative capability is more complex. For example, it has been shown that a peripheral nerve grafted into a cervical spinal cord lesion supports the ingrowth of injured rubrospinal axons; however, a peripheral nerve grafted into a thoracic cord injury does not result in the growth of axons (Fernandes et al, 1999). It is thought that this lack of growth is due to the lack of an appropriate cell body response (see below) in rubrospinal neurons that are injured at the thoracic level, because growing fibres into a cervical graft correlate with the upregulation of genes associated with regeneration. This upregulation is absent in thoracically injured rubrospinal neurons, an observation that supports the notion that these neurons are intrinsically incapable of mustering the appropriate programs necessary for successful regeneration. Only with a proximal axotomy do rubrospinal neurons upregulate the appropriate regeneration associated genes (for review, see Plunet et al, 2002). The silver lining of this research is that neurons that normally do not possess the intrinsic capacity for regeneration can be induced to grow under favorable conditions. The elucidation of these favorable conditions is ongoing. However, it is currently known that intrinsic neuronal limitations to regeneration exist within and between species. It is also known that the age of a neuron is a confounding factor in regeneration propensity. Fortunately, as described below, it is also clear that an appropriate neuronal response can be induced in intrinsically incapable neurons, thus enabling regeneration potential. Age-Dependent Limitations to Regeneration It is well established that there exists a critical period in development during which successful functional regeneration is possible. For example, the developing chick spinal cord is capable of supporting fibre regrowth and functional regeneration i f injured prior to embryonic day (E) 13 (Shimizu et al, 1990; Hasan et al, 1993). Similarly, the neonatal opossum has a critical period of plasticity whereby ascending and descending spinal cord 8 tracts are able to grow past lesions (Xu and Martin, 1991; Treherne et al, 1992; Wang et al, 1994; Varga et al, 1995). Although late developing neurons contribute to growth in these models, true regenerating fibres are also observed. It is clear that after the critical period has passed, regeneration of spinal cord projecting axons is not possible, indicating that the age of the injured spinal cord is a factor in regeneration propensity. It is unclear in these models, however, whether regeneration failure is caused by intrinsic neuronal limitations or environmental factors, as the end of the critical period for regrowth correlates with the appearance of the inhibitory molecules found in myelin (Steeves et al, 1994; Varga et al, 1995). More direct evidence for age-dependent limitations come from in vitro co-culture models similar to the models used in this thesis. Embryonic cerebellar Purkinje cells are able to grow axons on adult white matter substrates (i.e. myelin), but post-natal neurons fail to grow even when confronted with the permissive growth environment of a peripheral nerve (Dusart et al, 1997). In the same manner, early post-natal neurons from the entorhino-hippocampal projection (Li et al, 1995; Prang et al, 2001), and embryonic neurons of the retinotectal projection (Chen et al, 1995) also are able to grow in adult 'inhibitory' environments, whereas more mature neurons fail to grow in young 'permissive' environments. As a possible mechanism for age-dependent decline in growth ability, the Barres group have recently shown in vitro that RGC neurons receive an intrinsic developmental cue from their neighboring amacrine cells (Goldberg et al, 2002). This switched R G C process growth from axonal to dendritic. The cue is apparently irreversible, resulting in the failure of axonal regeneration from mature RGC cells regardless of permissive growth environments or trophic stimuli (Goldberg et al, 2002). Thus, there exist in the literature several examples of different neuronal populations that display an age-dependent intrinsic neuronal inability to regenerate in normally permissive environments, exemplifying intrinsic neuronal factors as important to regeneration capability. Cell-Specific Limitations to Regeneration Perhaps the most common example of intrinsic neuronal limitations to regeneration is found in the comparison between PNS or CNS regeneration. Unlike CNS injury, it is well known that anatomical axonal regeneration after an injury to a peripheral nerve is possible 9 (Fu and Gordon, 1997). If the D R G neurons sustain an axotomy to their central processes, regrowth is limited and functional recovery does not occur (reviewed by Bradbury et al, 2000). Here, the cause for regeneration failure seems to be the environment, as axons injured in the dorsal root will grow past the lesion and stop at the dorsal root entry zone (DREZ), a well defined barrier between the PNS and CNS (Bradbury et al, 2000). However, i f exogenous trophic factors are applied in this model, growth past the DREZ into the CNS occurs (Ramer et al, 2000; Ramer et al, 2002). As well, significant regeneration of dorsal column axons occurs i f a pre-conditioning peripheral nerve injury is performed prior to the CNS lesion (Neumann and Woolf, 1999). A similar effect is observed when cAMP agonists are injected into DRG, suggesting a possible mechanism for this conditioning effect (see Chapter 4; Neumann et al, 2002b; Qiu et al, 2002). These models indicate that the intrinsic growth state (or cell body response - see below) of the neuron is an important factor to regeneration failure. Furthermore, they indicate that techniques such as pre-conditioning can enhance the intrinsic neuronal growth state and overcome inhibitory environmental factors, resulting in axonal regeneration in the CNS. Other evidence supporting the importance of intrinsic factors in regulating axonal growth capacity is found in the popular technique of grafting peripheral nerves into the CNS. For example, brainstem-spinal neurons project axons into a peripheral nerve graft; however, corticospinal neurons do not (Richardson et al, 1984; Ye and Houle, 1997). Both groups of neurons subserve motor control functions and project similar distances to the same target areas. However, only brainstem-spinal axons are capable of growth into the permissive environment of a peripheral nerve. This indicates that cell-specific differences in the intrinsic ability to regenerate exist within functionally similar populations of CNS neurons. Spinal cord regeneration occurs in larval tadpole frogs (anurans), although this ability is lost after metamorphosis (Forehand and Farel, 1982; Beattie et al, 1990). Interestingly, only descending fibres are capable of growth in tadpoles - ascending sensory fibres are unable to regenerate. Pre-metamorphic frogs are also capable of optic nerve regeneration, and moreover, adult frogs retain this capacity (Forehand and Farel, 1982; Beattie et al, 1990). As well, functional recovery after spinal cord transection occurs in the goldfish (Bunt and Fi l l -Moebs, 1984; Sharma et al, 1993). However, anatomical tracing has shown that not all spinal projecting populations contribute to this recovery. Only 11 of 17 brainstem nuclei 10 were shown to originate regenerating axons (Bunt and Fill-Moebs, 1984; Sharma et al, 1993). A similar intrinsic difference between neuronal phenotype is seen in the cerebellum. Injured olivocerebellar axons readily grow into a permissive embryonic neural graft or Schwann cell graft, whereas, injured Purkinje axons invariably fail to regrow (Rossi et al, 1997) . This data emphasize that the dissimilar regenerative response of Purkinje cell and olivocerebellar axons is due to different intrinsic properties of these neuronal populations (Rossi et al, 1997). Why these growth differences exist within different CNS populations that grow in similar environments is unknown - it does however, underscore the importance of cell-specific intrinsic neuronal limitations to regeneration and the need to examine each cell population independently. The Cell Body Response The intrinsic state of an injured neuron ultimately refers to its induced pattern of gene expression and protein activity that may or may not be conducive to regeneration (reviewed by Plunet et al, 2002). The cell body response specifically refers to the battery of genes necessary for regrowth, commonly known as 'regeneration associated genes' (RAGS). Examples of known RAGS are: cytoskeletal proteins such as GAP-43 and CAP-23 (Skene, 1989; Bomze et al, 2001); cell adhesion molecules such as L l and N C A M (Becker et al, 1998) ; and neurotrophins and their receptors (Kobayashi et al, 1997). Recently, the product of an epithelial differentiation gene, SPRR1A, was reported to be expressed in axotomized axons 60-fold levels greater than in uninjured axons (Bonilla et al, 2002). These are only some of the known RAGS and undoubtedly many more exist and await elucidation. One of the hallmarks of successful PNS regeneration is the sustained upregulation of several RAGS (reviewed by Fernandes and Tetzlaff, 2000). Unfortunately, this upregulation does not occur in the CNS, where it is at best transitory. However, it is possible to induce sustained R A G expression in injured rubrospinal neurons with the use of exogenously applied trophic factors at the level of the cell bodies in the red nucleus (Kobayashi et al, 1997). In a recent paper investigating the chronically injured spinal cord, Tetzlaff and colleagues have shown that rubrospinal neurons survive, and can upregulate R A G expression and grow into a PNS graft following the cell body application of brain derived neurotrophic factor (BDNF) one year after axotomy (Kwon et al, 2002). This result highlights the 11 importance of the intrinsic neuronal state on regeneration ability, and demonstrates that with the correct therapeutic approach, changing the intrinsic state of neuron is not only feasible but also beneficial to spinal cord regeneration. In summary, neuronal regeneration is dependent on cell age, cell type, and induced cell body response. Fortunately, it seems possible to modulate the cell body response with the use of trophic factors that induce gene expression, thus promoting axonal growth in cells normally incapable. Another putative method of modulating the intrinsic properties of injured neurons is to target the intracellular signaling pathways that mediate growth cone behavior and ultimately axonal growth. A benefit of this approach is that the affected growth cones are at the injury site, and as such are far more accessible to clinical treatments than the cell bodies residing in the brainstem or cortex. Additionally, current clinical practice is to operate on the injury site (i.e. to decompress the spinal cord and stabilize the vertebrae), subsequently enabling the application of appropriate pharmacological agents directly to the lesion. Thus, the growth cone and its signaling pathways offer both viable physical targets for therapeutic intervention, as well as viable molecular targets, as described below. A X O N A L G R O W T H A N D R H O - G T P a s e s Axonal Growth Mechanisms The growth of an axon is driven by its distal tip, the site of the bi-functional growth cone. The growth cone is capable of sensing cues present in the environment and responding to those cues with directed growth. Thus, the growth cone is both navigator and driver, and its actions create the framework for the multitude of axonal connections that form during brain development. The growth cone is primarily composed of filamentous actin (F-actin) in the peripheral and central domains and microtubules that extend from the axon into the central domain (reviewed by Letourneau, 1996; Suter and Forscher, 1998; Gallo and Letourneau, 2000; Suter and Forscher, 2000). The peripheral domain is comprised of two dynamic F-actin based 12 structures - filopodia made from polarized bundles of F-actin, and lamellipodia made from a meshwork of F-actin (Fig. 1-1). These actin-rich structures are thought to be the engines of growth cone extension, providing the necessary force for motility (reviewed by Tanaka and Sabry, 1995; Suter and Forscher, 1998). Growth cone motility may occur by the coupling of the actin cytoskeleton to the extracellular substrate via ligand/receptor interactions and the process known as the 'retrograde flow' model (Lin and Forscher, 1995). Retrograde flow of actin is the sum effect of actin polymerization at the leading edges of filopodia and lamellipodia, translocation towards the central domain via myosin motors, and actin depolymerization at the proximal most portion of the growth cone periphery (Fig. 1-1B; Lin et al, 1997). It is the rate of actin polymerization and retrograde flow that dictate the rate of axonal extension in this model. Differential substrate adhesiveness is thought to be a factor in this model in determining growth cone steering events (Lin and Forscher, 1995). It is also possible that growth cone motility and turning is a result of local differences in second messenger cascades that regulate the actin-myosin force-generating machinery, irrespective of substrate adhesiveness (Isbister and O'Connor, 1999). In any event, to oversimplify the remainder of the axonal growth process, after extension of the actin structures, microtubules invade into the central domain of the growth cone, thus consolidating and extending the length of the axon. Directed navigation of axonal growth occurs through the local extension of filopodia and lamellipodia and microtubule consolidation. The dynamics of actin structures begin with the binding of an extracellular guidance ligand to a corresponding receptor on the growth cone. This sets off an intracellular signaling cascade that regulates the dynamics of the actin cytoskeleton. hi general, these guidance cues are either attractive (e.g. Netrins) or repulsive (e.g. Semaphorins), although the effect often depends on the receptors expressed on the growth cone (for review, see Y u and Bargmann, 2001) and the intrinsic state of intracellular signaling molecules (i.e. cAMP levels, see Fig. 1-2; Song et al, 1998; Cai et al, 1999). Thus, the effect of a guidance ligand on the actin cytoskeleton can either be local extension or retraction. The regulation of these actin dynamics is thought to be under the control of members of the Rho family of small guanosine triphosphatases (GTPases), of which there are at least 12 known members in mammals, the most commonly studied being Rho A, Racl , and Cdc42 (reviewed by Luo, 2000). 13 Figure 1-1. Schematic drawings of a growth cone characteristics. (A) A growth cone has distinct actin filament-based features, including distal, 'finger-like' filopodia and central, fan-shaped lamellipodia. (B) Representation of the cytoskeleton of a typical filopodium, including a theoretical growth mechanism underlying axonal elongation. Actin filaments are assembled in the distal tip of the growth cone and disassembled in the central domain of the growth cone allowing for the advance of microtubules and consolidation of the axon shaft. Growth cone advance occurs by the process of retrograde flow (FLOW) of actin, whereby receptor-ligand binding anchors the growth cone to the substrate and provides traction for the myosin-actin force generating machinery. Retrograde flow of actin is the sum effect of actin polymerization at the leading edges of filopodia and lamellipodia, translocation towards the central domain via myosin motors, and actin depolymerization at the proximal most portion of the growth cone periphery. Thus, the rate of growth cone advance can be modulated at any of the three fundamental actin events: assembly, retrograde flow, or disassembly. Adapted from Lin and Forscher, 1995. 14 15 Actin Cytoskeletal Changes are Mediated by Rho-GTPases Much of the current knowledge of growth cone actin dynamics was first elucidated from research into actin activity during changes in fibroblast morphology (Hall, 1998). Fibroblasts undergo three distinct actin-based cellular events: the formation of filopodia, lamellipodia, and stress fibres. By using constitutively active, mutants, the Hall laboratory showed that these three actin filament structures are respectively formed from the activity of Cdc42, Rac, and Rho (Nobes and Hall, 1995). Conversely, dominant-negative mutants inhibited the formation of these actin structures that normally appeared due to specific extracellular signals. For instance, dominant-negative Rho microinjected into fibroblasts exposed to lysophosphatidic acid inhibited the formation of stress fibres (Nobes and Hall, 1995; Lim et al, 1996). It has also been shown in fibroblasts that the GTPases can affect each other's activities (Nobes and Hall, 1995). Thus, there exists a signaling hierarchy amongst the GTPases - from Cdc42 to Rac to Rho. Unfortunately, the signaling pathways in neurons are not that simple. Similar to fibroblasts, the neuronal growth cone has distinct actin structures that are controlled by specific Rho-GTPases (Luo et al, 1997; Luo, 2000). Of particular interest to this thesis, the control of actin dynamics in the growth cone directly leads to advancement or retraction of an axon - hence, the interest in Rho-GTPases from neural regeneration scientists. In neuronal cells lines, Rac and Cdc42 activity promotes the formation of lamellipodia and filopodia, whereas Rho activity causes neurite retraction and growth cone collapse (Jalink et al, 1994; Tigyi et al, 1996; Leeuwen et al, 1997; Sebok et al, 1999). Several experiments in primary neuron cultures reveal more complex actions of Rho-GTPases on neurite growth. Expression of constitutively active Cdc42 in chick spinal neurons promoted neurite outgrowth and the formation of both filopodia and lamellipodia, whereas a dominant-negative Cdc42 inhibited growth (Brown et al, 2000). In chick retinal neurons, over-expression of wild-type Racl induces neurite growth, whereas dominant-negative Racl inhibits this growth (Albertinazzi et al, 1998). To confound this effect, it was shown that both constitutively active and dominant-negative Racl inhibit the growth of chick spinal neurons on a permissive laminin substrate (Kuhn et al, 1998). This implies that an optimum level of GTPase activity, or cycling between the GTP- and GDP-bound forms, is needed in this model. 16 Regarding neurite retraction and growth cone collapse, inhibition of Rho with C3-exotranferase promoted the elongation of retinal ganglion cell (RGC) axons on inhibitory substrates (Lehmann et al, 1999; Wahl et al, 2000). Similarily, expression of constitutively active RhoA in cerebellar granule cells inhibited the formation of axons, whereas C3-inhibition of Rho promoted axon initiation and elongation (Bito et al, 2000). Racl and Cdc42 activity are necessary for the neurite growth induced by netrin-1 through its deleted in colorectal cancer (Dec) receptor (Li et al, 2002). Moreover, this study showed that Rho and Rho-kinase inhibition also led to DCC induced neurite outgrowth. In general, it is thought that Cdc42 and Rac activity promote neurite growth and Rho activity inhibits neurite growth (Luo, 2000). Thus, Rho-GTPases have profound effects on growth cone actin structures and are important constituents of the pathways that mediate axonal growth or retraction. The activity of Rho-GTPases can be thought of as a molecular switch - on when bound to GTP and off when bound to GDP (Fig. 1-2). The regulation of Rho-GTPases occurs in two ways. First, RhoGDI (guanosine nucleotide dissociation inhibitor) seems to play a role both in shuttling Rho-GTPases between the cytosol and the plasma membrane (where the functions of Rho activity can occur) and stabilization of the inactive GDP-bound Rho-GTPase, although the full function of RhoGDI remains unclear (Lang et al, 1996; Mackay and Hall, 1998; Forget et al, 2002). Second, the regulation of Rho-GTPases between the active and inactive state is through the actions of two families of proteins: guanosine nucleotide exchange factors (GEFs) that factilitate the binding of GTP to turn Rho-GTPases active, and GTPase activating proteins (GAPs) that promote the endogenous GTPase activity of Rho-GTPases to inactivate them. The most studied GEF involved with growth cone dynamics is Trio, which contains a separate GEF domain for both Rac and Rho (Bateman and Van Vactor, 2001). The analysis of C. Elegans and Drosophila trio mutants has shown that Trio functions in axonal elongation and guidance (Steven et al, 1998; Awasaki et al, 2000). Thus, upstream GEFs and GAPs seem to be attractive targets for therapeutic intervention. For instance, ephexin, a Rho-GTPase GEF, can be modulated (by interaction with an Eph receptor) to increase the activity of RhoA while decreasing the activity of Racl and Cdc42, leading to growth cone collapse and neurite retraction (Shamah et al, 2001). The expression of a dominant-negative ephexin abolished growth cone collapse and promoted growth (Shamah et al, 2001). In this manner, a single target can effect the reciprocal actions 17 Figure 1-2. Schematic diagram of the Rho-signaling pathway in a growth cone. Inhibitory extracellular ligands bind growth cone receptors to begin the signaling cascade. Receptor activation somehow alters the balance of Rho-GEF and Rho-GAP activity favoring an increase in activated, GTP-bound Rho. Conversely, Rho activity can be down-regulated by RhoGDI sequestration to the cytoplasm, an effect promoted by intracellular c A M P / P K A elevation. Rho-GTP then activates its downstream effector kinases, of which R O C K is depicted here. R O C K in turn has effects on several substrates that affect actin events that lead to growth cone advance or retraction. R O C K both phosphorylates myosin light chain and myosin light chain phosphatase, causing direct activation of myosin contractility as well as indirect activation via inhibition of M L C phosphatase. Actin-myosin contractility is thought to be the necessary force behind neurite retraction since myosin motors may drive the retrograde flow of actin from the leading edge of the growth cone. R O C K also activates LEVI kinase, which inhibits cofilin activity, leading to a decrease in actin filament depolymerization. The reduction of F-actin depolymerization may serve to stablize the actin cytoskeleton, leading to a decrease in actin treadmilling and the inhibition of growth cone advance. 19 of Rho and Cdc42/Racl to promote axonal growth. Unfortunately, the number of GEFs and GAPs that are currently known is quite large - in Drosophila, there are six known Rho-GTPases and approximately 40 GEFs and GAPs (Luo, 2000). Reasons for this are unknown; however, it may be that specific GEFs/GAPs only interact with specific receptors, limiting their usefulness as therapeutic targets. This knowledge contributes to the idea that Rho may be a convergence point from several inhibitory receptor/ligand interactions to the actin cytoskeleton (see below). Because of this putative convergence, I have focused my attention on Rho and its downstream effectors as possible pharmacological targets. The effects of Rho activity are mediated by its binding with downstream protein kinase effectors (Fig. 1-2). The effectors bind to Rho on a short effector loop domain that becomes exposed during Rho-GTP binding (Bishop and Hall, 2000). The most well characterized (and subject of this thesis) Rho-effector is the serine/threonine kinase, Rho-kinase/ROKot/ROCK-2, hereinafter referred to as ROCK. The signaling of Rho to the actin cytoskeleton is mediated by ROCK, which in turn has several downstream targets (Amano et al, 2000). For example, R O C K regulates the phosphorylation and activity of myosin light chain (MLC), both directly by phosphorylation of M L C (Amano et al, 1996) and indirectly by inactivation of myosin phosphatase (Kimura et al, 1996). As well, R O C K phosphorylates LIM-kinase (Maekawa et al, 1999; Ohashi et al, 2000), which in turn is necessary for Sema3A-induced growth cone collapse mediated via phosphorylation and inhibition of cofilin (Aizawa et al, 2001). The consequence of R O C K activity in the growth cone to cause neurite retraction or growth cone collapse is in part due to two effects. First, R O C K increases actin-myosin contractility through the phosphorylation and activation of M L C . Actin-myosin contractility is thought to be the necessary force behind neurite retraction (Amano et al, 1998) because myosin motors drive the retrograde flow of actin from the leading edge of the growth cone (Lin et al, 1996). Second, R O C K phosphorylates LIM-kinase, which in turn reduces F-actin filament depolymerization through the phosphorylation and inhibition of cofilin (Arber et al, 1998). The reduction of F-actin depolymerization may serve to stablize the actin cytoskeleton, leading to a decrease in actin treadmilling and the inhibition of neurite outgrowth (Meberg and Bamburg, 2000). Accordingly, R O C K may be an attractive target to neutralize in order to interfere with the signaling from Rho to the actin cytoskeleton. 20 Inhibitory Molecules Act Through Rho-GTPases The effect of inhibitory molecules found in the injured CNS is growth cone collapse and/or axonal retraction, events controlled by the Rho-GTPases. Semaphorin-3A, known at the time as collapsin-1, was one of the first purified proteins shown to induce growth cone collapse and inhibit neurite growth (Luo et al, 1993). More recently, it has been shown that Sema-3 A causes growth cone collapse through the activation of Rho-GTPases (reviewed by Liu and Strittmatter, 2001). C3-exotoxin inhibition of Rho promoted axonal elongation from chick D R G cells that were inhibited by Sema-3 A (Jin and Strittmatter, 1997). Two lines of evidence suggest that the activation of Rho in this model is downstream of Racl activation. First, the suppression of Racl signaling blocks Sema-induced growth inhibition (Jin and Strittmatter, 1997; Kuhn et al, 1999; Vastrik et al, 1999). Second, the ectopic expression of Plexin-Bl (a semaphorin receptor) in fibroblasts causes cell contractility and stress fibre formation reminiscent of Rho activation, which is blocked by dominant-negative Racl (Driessens et al, 2001). P. Letourneau, at the 2001 Winter Brain Conterence, reported further evidence in support of the Rho pathway as important in Sema-3A signalling. He reported that the suppression of R O C K signaling (with Y-27632, a specific inhibitor of ROCK; Uehata et al, 1997) promoted the growth of D R G neurites in the presence of Sema-3 A. Although the involvement of more than one Rho-family GTPase in Sema-3A signaling is undoubtedly complex, the suppression of the Rho pathway appears potent. The Eph receptors and their ephrin ligands comprise a family of growth cone guidance molecules that normally inhibit the growth of axons. Recent experiments have revealed that inhibitory signaling through EphA receptors is mediated through Rho-GTPases. Exogenously applied ephrin-A5 causes the activation of RhoA and inactivation of Racl , resulting in the growth cone collapse of retinal ganglion cells (Wahl et al, 2000). This collapse can be overcome by inhibiting Rho activity with C3, or by inhibiting R O C K activity with Y-27632. Similarly, the activation of EphA4 by ephrin-A1 directly interacts with and modulates ephexin, a Rho-GTPase GEF (Shamah et al, 2001). This modulation of GEF activity results in the activation of RhoA and inhibition of Cdc42 and Racl activity, leading to growth cone collapse in RGCs. The expression of a dominant-negative ephexin mutant inhibits this collapse (Shamah et al, 2001), supporting the approach of interfering with Rho pathway molecules to promote axonal growth. 21 Axonal inhibition by myelin may also be mediated by Rho-GTPases. The inhibition of Rho activity by C3 application has been shown to promote the extension of PC 12 neurites and RGC axons grown on myelin and purified M A G substrates in vitro (Lehmann et al, 1999). As well, the expression of dominant-negative RhoA promoted the growth of PC 12 neurites on these substrates. Moreover, in the same report, it was shown that C3 application in vivo promoted axonal regeneration. The optic nerve of adult rats was crushed and C3 applied. Axonal growth from RGCs was observed up to 500 pm from the crush site, although the authors speculate that this limited distance may be due to the poor cellular penetration inherent to the C3 molecule (Lehmann et al, 1999). Very recently, neurite growth inhibition due to M A G was shown to act through the p75 neurotrophin receptor and Rho (Yamashita et al, 2002). Adult D R G neurons with non-functional p75 receptors are insensitive to M A G . As well, M A G activates RhoA and colocalizes with and binds to p75 in neurons (Fig. 1-3; Yamashita et al, 2002). Interestingly, in addition to Nogo, it has also been shown that both M A G (Domeniconi et al, 2002; Liu et al, 2002) and OMgp (Wang et al, 2002) bind to the recently identified (Fournier et al, 2001) Nogo receptor (Fig. 1-3). Thus, three inhibitory components of myelin bind to the same receptor, suggesting a further upstream site of convergence to the Rho-pathway. Also regarding myelin inhibition and Rho, several years ago Strittmatter and colleagues reported a confounding result (Jin and Strittmatter, 1997). They showed that C3 inhibition of Rho promoted robust axonal extension of chick D R G cells grown on inhibitory myelin substrates. However, the growth cones of these axons appeared collapsed. While this result indicates possible theories of axonal growth mechanisms, it also speaks to the need to assay axonal elongation when studying growth inhibition. Regardless of the shape of the growth cone, the extension of injured axons is ultimately the result sought after. Another example of possible Rho involvement in regeneration failure comes from activated glial cells, such as those found at CN$ injury sites, that secrete the pro-inflammatory cytokine, tumor necrosis factor-alpha (TNF) (Hopkins and Rothwell, 1995). TNF has wide-ranging effects on neurons, including cell death (Venters et al, 2000). Recently, TNF has also been shown as a potent inhibitor of hippocampal axon growth in vitro (Neumann et al, 2002a). This growth inhibition was accompanied by activation of Rho. The application of C3 neutralized the Rho activation and promoted axonal extension and 22 Figure 1-3. Schematic of inhibitory myelin components and their putative receptors. Three myelin membrane proteins inhibitory to axonal growth are Nogo, OMgp, and M A G . A l l three may bind to Nogo-66 receptors in the axonal growth cone, although downstream signaling events are less clear. M A G has also been shown to bind both to ganglioside GTlb and to the p75 low-affinity neurotrophin receptor. p75 may regulate the affects of M A G through Rho-GTPase signaling to the actin cytoskeleton. OMgp, oligodendrocyte-myelin glycoprotein; M A G , myelin associated glycoprotein; NgR, Nogo-66 receptor. Myelin membrane Nogo-66 domain Growth cone membrane (C IgG domains | Signal peptide | Leucine rich repeats • Serine/threonine repeats • P75 death domain • Cysteine rich repeats O Oligosaccharide • Leucine rich repeat C terminal domain 24 branching (Neumann et al, 2002a). Thus, spinal cord injury has inhibitory growth influences from both the axonal growth substrates and from molecules secreted from injury-induced activated glial cells. Interestingly, Rho inactivation may be a viable therapeutic approach to overcoming both. In summary, it is well established that members of the Rho family of small GTPases play a central role in the signalling events from extracellular inhibitory molecules to the growth cone actin cytoskeleton. Because of the many inhibitory molecules expressed in the environment of regenerating axons, it is my approach to target a putative convergence point in the intracellular signaling pathways common to many inhibitory molecules. Hence, the targeting of ROCK, Rho's major effector kinase, with possible therapeutic drugs like Y -27632 that specifically suppresses R O C K activity. R A T I O N A L E A N D H Y P O T H E S E S It has been in vogue to dismiss intrinsic neuronal limitations as a possible reason for CNS regeneration failure. For instance, a perusal of recent reviews on spinal cord regeneration (Horner and Gage, 2000; Fournier and Strittmatter, 2001) suggest that, due to the work of Agyuao showing the growth of CNS axons into peripheral nerve grafts (Richardson et al., 1980), CNS neurons are inherently capable of regeneration. Unfortunately, the actual number of axons capable of growth in these permissive environments is low. For example, Tetzlaff and colleagues found a mean of 43 magnocellular neurons from the red nucleus grew into a PNS graft (Kobayashi et al, 1997). Compare this to the approximately 1200 magnocellular neurons contained in the red nucleus. Certainly with less than 4 or 5 percent of CNS neurons able to grow in a permissive environment there are some intrinsic neuronal factors at work in regeneration failure. As well, corticospinal neurons consistently fail to grow into PNS grafts (Richardson et al, 1984; Ye and Houle, 1997), indicating that these neurons may have some further intrinsic limitations that other CNS neurons do not. Recently however, research has been directed at both defining the intrinsic neuronal limitations to regeneration (Goldberg et al, 2002), as well as attempting to enhance the intrinsic capacity for growth (Berry et al, 25 1996; Fischer et al, 2001; Neumann et al, 2002b; Qiu et al, 2002; Snider et al, 2002). Thus, it seems advantageous to further research the possible intrinsic limitations to regeneration and to investigate methods for increasing the neuronal growth capacity directly. Early experiments with the brainstem-spinal neuron model described below showed that young embryonic (E8) brainstem-spinal neurons could be successfully cultured, and are capable of surviving and growing neurites for several days in vitro. After some experience with this model, I speculated whether the time frame for the initiation of successful culture of brainstem-spinal neurons could be extended beyond E8. A fundamental question regarding SCI and traumatic brain injury is whether mature neurons retain the intrinsic capacity for regeneration. As described above, cerebellar Purkinje (Dusart et al, 1997), retinotectal (Chen et al, 1995; Goldberg et al, 2002), and entorhino-hippocampal (Li et al, 1995) neurons all display an age-dependent decline in their propensity to regrow neurites in vitro. With no great intellectual leap, I hypothesized that mature brainstem-spinal neurons lose their ability to extend neurites in vitro. However, with the diversity of anatomy and function displayed by brainstem-spinal populations, I also asked whether there existed phenotypic differences in the abilities of brainstem-spinal neurons to grow. It is a popular belief amongst SCI researchers that serotonergic neurons are inherently more plastic than their non-aminergic neighbours; however, there exists almost nothing in the literature that directly addresses this topic. I suggest that the belief arose because of the many SCI studies showing growth of serotonergic fibres into permissive growth environments. The common observation of sertonergic fibres was most likely due to the availability of phenotypic markers in the form of antibodies to serotonin, a tool not available to most other brainstem-spinal populations. With the aim of directly answering this question in mind, I hypothesized that differences in the propensity for neurite regrowth exist within sub-populations of brainstem-spinal neurons. I addressed these above two hypotheses by culturing neurons from 5 different populations of brainstem-spinal neurons at both a young (E8) and advanced (El 7) age, the results of which have been published (Borisoff et al, 2000) and are reproduced as Chapter 2. The above hypotheses address in vitro the intrinsic limitation for regeneration that is exhibited by higher vertebrate species. I was also interested in the environmental factors that 26 inhibit regeneration, and specifically the question: Is it possible to change the intrinsic state of the neuron from inhibition to growth? Recent advances in intracellular signaling by axonal guidance molecules have led to the study of CNS inhibitory molecule signaling. This limited knowledge is the basis for the idea that axonal inhibition may be overcome by modulating the growth cone response to inhibitory molecules, as opposed to eliminating or masking the inhibitory factor directly. A corollary to this proposition is that intracellular signaling of several inhibitory molecules may converge onto a common effector molecule, thus enabling the suppression of several inhibitory molecules at once. As described above, a candidate target for this convergence is Rho-GTPase and its effector kinase ROCK. I hypothesized that the in vitro application of Y-27632, a specific inhibitor of R O C K signaling, may promote neurite regrowth on different inhibitory substrates. I addressed this hypothesis by culturing D R G explants on substrates comprised of either chondroitin sulfate proteoglycan or myelin, the results of which have been submitted for publication and are reproduced as Chapter 3. IN VITRO M O D E L S OF SPINAL C O R D R E G E N E R A T I O N The bulk of knowledge about CNS regeneration following spinal cord injury has been elucidated through in vivo models, often producing results that are difficult to interpret and that are irreproducible by other labs (see landmark studies by Cheng et al., 1996; Ramon-Cueto et al, 2000). In order to obtain mechanistic information about various problems associated with SCI and traumatic brain injury, several labs have used in vitro models to address pertinent cell biology questions. Unfortunately, most likely due to the ease of use, investigators often utilize less relevant cells such as neuronal-like cell lines, cerebellar granule cells, and retinal ganglion cells to study regeneration. Although many cellular functions are conserved between different cell types, in order to more fully understand the cellular and molecular mechanisms important to SCI, the specific neuronal populations affected must be studied. SCI is characterized by a permanent loss of both motor and sensory function. Thus, a successful treatment for SCI must address the loss of both functions. To achieve such a 27 treatment necessitates the study of both descending and ascending systems in order to gain the specific knowledge required to direct and implement an effective repair strategy. Descending motor systems are comprised of axons from corticospinal neurons and several different populations of brainstem-spinal neurons, including reticulospinal, rubrospinal, vestibulospinal, and raphe-spinal neurons. The sensory system is comprised of axons originating from dorsal root ganglia, neurons with both peripheral and central projections. Both systems are amenable to study in vitro. To study the factors regulating the regeneration of these systems, I chose the developing chick as a model system. The chick is a vertebrate biped with motor and sensory systems that are conserved among humans and other higher vertebrates (Steeves et al, 1987; Webster et al, 1990; O'Donovan et al, 1992). As well, there is a large body of knowledge about axonal regeneration and development collected from the use of chick models. Chick neurons are especially favored in studies of Rho-GTPases (Jin and Strittmatter, 1997; Kuhn et al, 1999), a topic central to this thesis. The chick is also precocial, having a development time of approximately 21 days, where after hatching it is capable of walking almost immediately. This accelerated development occurs entirely in ovo, lending the chick to uncomplicated experimental manipulation during specific developmental windows, a procedure not possible in rodent models. The developing chick also possesses favorable handling and economic benefits, characteristics that further support its use as a model system. The developing chick has a critical period for successful spinal cord regeneration, a characteristic exploitable for studies of CNS repair. Functional regeneration of severed brainstem-spinal pathways occurs in the chick following a spinal cord transection in ovo when the injury occurs prior to embryonic day (E) 12 (Shimizu et al, 1990; Hasan et al, 1993). On E l3 and later, however, repair of these pathways does not occur, as is the case following an adult spinal cord injury. Thus, the developing chick offers two distinct periods of regenerative capacity: a young permissive period in which axons are normally elongating through the spinal cord and are capable of regrowth following injury, and a mature restricted period in which axons have completed their elongation and regeneration fails. Primary neurons from both periods are accessible for in vitro experimentation, enabling the investigation of many relevant SCI questions. 28 Explants of Identified Brainstem-Spinal Neurons In order to answer questions regarding neurons germane to the restoration of motor function following SCI, the Steeves' laboratory, with my contribution, developed a method to culture identified brainstem-spinal neurons (Pataky et al, 2000). The neuronal origin of CNS motor output to the spinal cord is primarily found within the brainstem (Eidelberg, 1981; Grillner and Wallen, 1985; Grillner and Dubuc, 1988). In mammals, the other major contributor to motor function originates in the sensorimotor cortex, where the cell bodies of the often sensationalized corticospinal tract are found. The corticospinal tract primarily mediates fine motor control of distal extremities (Lawrence and Kuypers, 1968), and is not as important to the initiation and control of the motor patterns required for standing and walking which are primarily controlled by brainstem-spinal pathways (Steeves et al, 1987; Valenzuela et al, 1990). Thus, the focus of the model is on brainstem-spinal neurons (Fig. 1-4), although it is not lost on the author that avians do not possess a corticospinal tract, as is found in higher vertebrates (Webster et al, 1990). Reticulospinal neurons are the major contributor to locomotor function (Steeves et al, 1987; Whelan, 1996), the cell bodies of which are found in the reticular formations as bilateral columns in the pons, medulla, and midbrain (Glover, 1993). In the experiments described in Chapter 2, two distinct groups of reticulospinal neurons are excised and studied: medullary (Rm) and pontine (Rp) reticulospinal neurons (Fig. 1-4). Rm explants contain neurons from both the nucleus reticularis caudalis and the nucleus reticularis gigantocellularis (Okado and Oppenheim, 1985). Rp explants contain neurons from the nucleus reticularis pontis parvicellularis and the nucleus reticularis pontiscaudalis pars gigantocellularis (Okado and Oppenheim, 1985). Vestibulospinal neurons control postural balance (Lund and Pompeiano, 1968) and visual stability (Boyle and Pompeiano, 1980). Found near the ponto-medullary junction, they are the most lateral group of brainstem-spinal neurons. Vestibulospinal neurons are comprised of lateral, medial, and descending groups (Glover and Petursdottir, 1991), although in the experiments described in Chapter 2, they are exised together as a single group, notated as vestibulospinal (V) neurons (Fig. 1-4). Raphe-spinal (R) neurons are found at the midline of the brainstem (Cabot et al, 1982), from the caudal pons through the length of the medulla (Fig. 1-4). Raphe neurons are 29 F i g u r e 1-4. The chick brainstem-spinal neuron populations, as revealed through low-magnification fluorescence microscopy of retrograde labeling of D i l from the cervical spinal cord. The dorsal view of an E8 whole-mount brainstem with the cerebellum filleted open clearly shows the vestibulospinal (V) population as well as the M L F fiber tract. The ventral view depicts the four major brainstem-spinal populations studied here that originate from the hindbrain. The fifth population is found in the midbrain, consisting of rubrospinal neurons from the Red nucleus (seen here - upper right panel) as well as the slightly more rostral interstitial nucleus of Cajal. The four right panels are representative slices from an E l5 brainstem from appropriate rostro-caudal levels. These slices would then be micro-dissected to obtain explants of specific brainstem-spinal populations. V = vestibular complex; R = raphe-spinal neurons; Rp = pontine reticulospinal neurons; Rm = medullary reticulospinal neurons; Red = Red nucleus; M L F = medial longitudinal fasciculus. 31 predominantly of a serotonergic phenotype (JJkeda and Goto, 1971; Dube and Parent, 1981) and unmyelinated (Westlund et al, 1992). Functionally, raphe-spinal neurons are considered to be modulators of motor control (Iwamoto et al, 1980; Azmitia, 1999; Schmidt and Jordan, 2000) and sensory transmission (Duggan and Griersmith, 1979), rather than as direct effectors of motor function. Raphe-spinal neurons are bilateral, although in the experiments described in Chapter 2, they are excised as a single population. The Red nucleus is found in the midbrain and participates in the modulation of motor control, as well as the coordination of fine motor control of distal extremities (Arshavsky et al, 1988; Jarratt and Hyland, 1999). The Red nucleus is located near the interstitial nucleus of Cajal, a spinally projecting population important in the coordination of eye and head movement. Because of the close proximity of these two nuclei, both populations are excised together and are indicated as midbrain neurons in Chapter 2 (Fig. 1-4). Because most brainstem-spinal neurons lack known phenotypic markers (an obvious exception is serotonergic raphe neurons), the Steeves laboratory developed an in ovo labeling paradigm (Pataky et al, 2000). We exploited the fact that the one common feature of brainstem-spinal neurons is the extension of an axonal projection to the spinal cord. The detailed methods of this paradigm are provided in Chapter 2. Briefly, the cell bodies of brainstem-spinal neurons were retrogradely labeled by the insertion of a crystal of D i l adjacent to the cervical spinal cords of E5 chicks, a time during the developmental period of axonal growth from the descending pathways. The egg was then resealed and incubated until the appropriate time for start of tissue culture. Dissected brainstems were sliced into coronal sections whereby the individual brainstem-spinal nuclei could be viewed with a fluorescent microscope and micro-dissected into distinct explants. A key to this model is the fact that D i l is incorporated into the regrowing neurites, enabling the analysis of in vitro growth (Fig. 1-5). Thus, explants of identified, isolated brainstem-spinal neurons, compatible for many types of in vitro studies are prepared. A schematic of the entire procedure is provided in Figure 1-5. Explants of Dorsal Root Ganglia As a complement to the brainstem-spinal model described above, I also used explants of E9 or E l 5 dorsal root ganglia (Fig. 1-6). D R G neurons compose the ascending sensory systems that are damaged in a traumatic spinal cord injury. These sensory systems are 32 Figure 1-5. Schematic diagram of the methods of the in vitro brainstem-spinal neuron neurite outgrowth assay. A crystal of D i l is implanted adjacent to the cervical spinal cord of an E5 chick embryo in ovo to retrogradely label the spinally projecting neurons that have their cell bodies in the brainstem (A). Days later, the brainstem is dissected and sliced into sections from which distinct explants of brainstem-spinal neurons can be obtained (B). Explants are plated into culture with appropriate substrates and treatment media. Neurite outgrowth that contains redistributed D i l can then be visualized with fluorescence microscopy (C). Adapted from Pataky et al., 2000. Labeled Brainstem-Spinal Neurons Micro-Dissection Of Brainstem Slice In Vitro Explant Culture Axonal Outgrowth Assay 34 Figure 1-6. An example of neurite outgrowth from a whole E9 chick dorsal root ganglion. Neurite outgrowth can be visualized with several different markers, including the vital dye CFDA,SE and antibodies to axonal proteins. Myosin light chain immunoreactivity is shown here (A). Quantification of neurite outgrowth is completed by manually tracing the growth in Photoshop or NfH Image (B). 35 36 composed of distinct subpopulations identified by their sensory modality, trophic factor sensitivity, and cell soma size. In general, small diameter nociceptors are nerve growth factor (NGF) sensitive; medium diameter mechanoreceptors are brain derived neurotrophic factor (BDNF) sensitive; and large diameter proprioceptors are neurotrophin-3 (NT-3) sensitive (for review, see Lindsay, 1996; Snider and Wright, 1996). In an in vitro model designed to isolate the axonal growth inducing effects of neurotrophins from the survival promoting effects, Snider and colleagues (Lentz et al, 1999) have shown that NGF, BDNF, and NT-3 promote distinct axonal growth morphologies from different populations of D R G neurons. Thus, explants of D R G offer the choice of (at least) three different populations for experimentation, selectable by the application of one of the three neurotrophins mentioned above to the culture medium, although the bulk of the experiments described in this thesis were performed with the NGF sensitive population. As described above, the developing chick provides the opportunity to study neurons during both the permissive and restrictive periods of successful regeneration. Young D R G neurons from E9 chick were used in the majority of the experiments described here. However, at this age, D R G neurons are not sensitive to myelin-derived inhibitors (Mukhopadhyay et al, 1994; Fournier et al, 2001), most likely due to the lack of appropriate receptor expression. For example, the recently identified receptor for Nogo is only upregulated after E12 in the chick (Fournier et al, 2001). Growth cones of sensory neurons younger than E12 do not collapse in response to myelin or purified Nogo (Fournier et al, 2001) . Furthermore, the inhibitory actions of two other myelin proteins, M A G and OMgp have recently been shown to act through the Nogo receptor (Liu et al, 2002; Wang et al, 2002) . Thus, in experiments using myelin or spinal cord as growth substrates, explants from E l 5 D R G were used. The brainstem-spinal model was used specifically for the study of intrinsic limitations to regeneration, as described in Chapter 2. However, after working with the brainstem-spinal neuron model, it became clear that D R G neurons were the ideal choice for my experiments in neurite growth promotion on inhibitory substrates. Nothing is known about the brainstem-spinal neuron protein expression pattern for the receptors to guidance molecules and inhibitory molecules. That is one reason for the use of D R G neurons for the study of growth 37 promotion on substrates of inhibitory molecules described in Chapter 3. Two other reasons for the use of D R G neurons are: they have been used as a model system in previous studies involving Rho-GTPases, thus enabling the comparison of my data to the literature; and they are also amenable to higher through-put screens of putative therapeutic drugs that may promote axonal growth on inhibitory molecules. Therefore, D R G cultures facilitated the efficient study of growth promoting molecules. The rationale is to return to the brainstem-spinal neuron model upon the discovery of effective growth promoting agents in D R G cultures. 38 CHAPTER 2 : RAPHE-SPINAL NEURONS DISPLAY AN AGE-DEPENDENT DIFFERENTIAL CAPACITY FOR NEURITE OUTGROWTH COMPARED TO OTHER BRAINSTEM-SPINAL POPULATIONS 39 This chapter presents an in vitro study of the intrinsic growth capacities of brainstem-spinal neurons. Specific populations of young and mature brainstem-spinal neurons are grown as explants on permissive laminin substrates. The results show that the growth capacity of brainstem-spinal neurons is dependent on both neuronal age and specific neuronal population. Raphe-spinal neurons display an ability to grow at later ages than other brainstem-spinal neurons. Thus, intrinsic neuronal limitations may be an important factor in the failure of CNS regeneration. This study was published (Borisoff et al, 2000) and is reproduced here with formatting consistent with the rest of the thesis. A more thorough and timely discussion of intrinsic axonal growth capacity is provided in the general discussion in Chapter 4. I N T R O D U C T I O N In higher vertebrates, substantial regrowth of injured spinal cord projection neurons is only seen in immature animals. For example, in the embryonic chick (Shimizu et al, 1990; Hasan et al, 1993) and neonatal opossum (Xu and Martin, 1991; Treherne et al, 1992), there is an established permissive period for the successful regeneration of brainstem-spinal neurons. In the chick, anatomical regeneration and functional repair of severed brainstem-spinal pathways occurs following a spinal cord transection in ovo until embryonic day (E) 12 (Shimizu et al, 1990; Hasan et al, 1993). After approximately E13, however, axonal repair of these pathways is restricted and function remains impaired, as occurs after an adult spinal cord injury. There are several, potentially overlapping, reasons for the failure of axonal regeneration within the central nervous system (CNS) of adult or developmentally later stage animals. One hypothesis involves maturational changes in the extrinsic environment encountered by injured axons. In the chick, the onset of the restrictive period for functional axonal repair correlates with the appearance of myelin in spinal pathways. By immunologically delaying the onset of myelination, the developmental age limit for successful regeneration was extended to E15 (Keirstead et al, 1992). A correlation between the appearance of inhibitory myelin proteins and axonal repair has also been shown in the developing opossum CNS 40 (Varga et al, 1995). In mammals, several molecules present in myelin are inhibitory to axon growth, including myelin-associated glycoprotein and NI-35/250/Nogo (McKerracher et al, 1994; Filbin, 1995; L i et al, 1996; Chen et al, 2000; GrandPre et al, 2000). The application of antibodies to NI-220/250 (Schnell and Schwab, 1990, 1993) or the immunological removal of myelin (Dyer et al, 1998) results in only limited regeneration, suggesting other factors, such as those found at the lesion site within the astroglial scar may also be involved. For example, astrocyte-associated molecules such as tenascin (McKeon et al, 1991; Zhang et al, 1995) and chondroitin sulphate proteoglycans (Snow et al, 1990; Dow et al, 1994; McKeon et al, 1995; Bovolenta et al, 1997; Fitch and Silver, 1997b) may contribute to a regeneration barrier (for review, see Fawcett and Asher, 1999). A second hypothesis relates to changes in the intrinsic ability of CNS neurons to regrow axons. A n age-dependant decline in the ability of some neurons to regenerate is evident, regardless of environmental factors. In vitro, embryonic Purkinje cells display extensive axonal regenerative capacities on adult white matter substrates (i.e. myelin), but post-natal neurons are unable to regenerate even when confronted with the permissive growth environment of a peripheral nerve (Dusart et al, 1997). The entorhino-hippocampal projection (Li et al, 1995), and retinotectal projection (Chen et al, 1995) also display an age-dependent intrinsic neuronal inability to regenerate even in permissive environments. In addition, different types of neurons display varying capacities for regeneration. In the adult frog, retinal ganglion cells (RGC) are able to reestablish connections with their tectal targets, resulting in functional recovery after optic nerve injury. However, most other (CNS) neurons, including brainstem-spinal neurons, appear unable to regenerate (Forehand and Farel, 1982; Beattie et al, 1990). In the adult rat, axonal growth of brainstem-spinal, but not corticospinal, axons occurs within a peripheral nerve graft (a permissive environment for axon growth) placed into a cervical spinal cord injury site (Richardson et al, 1984). Thus, the intrinsic ability of neurons to regenerate may vary between different neuronal phenotypes projecting over comparable distances or even to similar target regions. We were interested in examining the intrinsic age-related abilities for axon regeneration in neurons germane to spinal cord injury. The specific neurons mediating many behaviors such as the initiation and control of locomotion are found in multiple regions within the brainstem, including the reticular formation, raphe, vestibular and Red nuclei (Eidelberg, 41 1981; Grillner and Wallen, 1985; Grillner and Dubuc, 1988). In the developing chick brainstem, neurogenesis is complete prior to E3 (McConnell and Sechrist, 1980), with the first descending axons entering the cervical spinal cord on E3-E4 (Okado and Oppenheim, 1985; Glover, 1993). Functional synaptic input from supraspinal projections is detectable within the spinal cord by E6 (Shiga et al, 1991; Sholomenko and O'Donovan, 1995), and the development of axonal projections from the brainstem to all levels of the cord is complete by E l l (Okado and Oppenheim, 1985; Shimizu et al, 1990; Glover and Petursdottir, 1991; Shiga et al, 1991; Glover, 1993). This early developmental period of axon outgrowth and pathfinding correlates with the permissive period for successful regeneration in the chick (reviewed in Keirstead and Steeves, 1998). The onset of the restrictive period for regeneration of brainstem-spinal pathways in the chick (El3) correlates with the onset of spinal myelination (Keirstead and Steeves, 1998), as well as the appearance of mature phenotypic markers, such as glial fibrillary acidic protein (GFAP) in astrocytes (unpublished observations, J. McGraw and J.D. Steeves). By E l 7, chick spinal myelination is relatively complete (Hartman et al, 1979). These features of chick CNS development suggest that at E17, brainstem-spinal neurons are fully differentiated and mature or "adult-like", further supported by the precocial locomotor abilities of chicks upon hatching. We developed a method for studying brainstem-spinal neurons by employing an in vivo labeling technique to identify brainstem-spinal neurons and subsequently follow their survival and neurite regrowth in vitro (Pataky et al, 2000). After labeling the spinal projection neurons on E5, we prepared identified explant cultures from brainstem tissue obtained during either the permissive or restricted periods of axonal repair. In this study of age-dependent neurite regeneration, we chose two time points, E8 and E l 7 , as the experimental ages for young and mature neurons, respectively. Neurite outgrowth from five different populations of brainstem-spinal neurons was examined and compared between explants obtained from E8 and E l 7 chicks. 42 M A T E R I A L S A N D M E T H O D S Reagents were obtained from Canadian Life Technologies (Burlington, ON, Canada) unless otherwise stated. In Ovo Dye Application Prior to culturing, brainstem-spinal neurons were specifically labeled using a retrograde tracing paradigm (Pataky et al., 2000). Fertilized White Leghorn chicken eggs (Gallus gallus domesticus) were obtained from a local supplier and incubated at 37.5°C in a humidified atmosphere. Briefly, on embryonic day (E) 4.5, a temporary opening was made in the eggshell to expose the embryo. An insect pin coated with crystallized fragments of the carbocyanine dye, D i l (l,r-dioctadecyl-3,3,3',3'-tetramethylindocarbocyanine perchlorate, Molecular Probes Inc., Eugene, OR) was used to implant the retrograde tracer. With care being taken not to directly damage the spinal cord, the D i l was positioned in contact with the external surface of the mid-cervical spinal cord producing close apposition of the dye crystals with the ventral and lateral funiculi of the spinal cord. The eggshell was then sealed and incubated until E8 or E l 7, permitting ample time for the retrograde transport of the D i l to the cell bodies of brainstem-spinal neurons. Tissue Culture On the appropriate experimental day, each embryo was removed from the shell and decapitated. The brainstem was then dissected in ice cold C a + + and M g + + free Hank's balanced salt solution supplemented with 0.5% glucose. Coronal sections from E8 (300 um) and E l 7 (350 urn) were sliced with a motorized Mcllwain Tissue Sheer (Brinkmann, Westbury NY) . Single slices from the appropriate rostral-caudal brainstem level were then micro-dissected with a razor blade to obtain explants containing specific brainstem-spinal neurons. Explants were plated individually in 24-well culture plates (Costar) pre-coated for 3 hours at 37°C with 50 Lig/ml laminin. Cultures were grown in a humidified 37°C incubator with a 5%C02/95% air atmosphere in Neurobasal-A medium supplemented with B-27 (for E l 7 explants) or in Neurobasal medium supplemented with N-2 (E8 explants). Both media 43 were also supplemented with glutamine (0.5 mM), and 100 units/ml penicillin/streptomyocin. N-2 and B-27 supplements contain insulin, transferrin, selenium, progesterone, and putrescine, while B-27 also has additional additives. Experiments were also performed with Dulbecco's Modified Eagles Medium (DMEM) culture medium supplemented with 10% Fetal Bovine Serum. Initial results indicated that the Neurobasal formulations produced the most consistent results for both E8 and E17 cultures (data not shown). Although Neurobasal-A and B-27 was formulated for adult rat hippocampal cultures (Brewer et al, 1993; Brewer, 1997), the initial results indicated its appropriateness for late stage embryonic chick CNS neurons that have many adult-like characteristics. After 3 days, E l 7 cultures were fixed with 10%o formalin (Fisher Scientific, Nepean, ON) in 0.1M phosphate buffered saline (PBS) containing 10% sucrose. E8 cultures were fixed one day sooner because of the substantial neurite outgrowth already present after 2 days in vitro. Neurite Outgrowth Assay Images of the Dil-labeled neurites were captured with a 4X objective on an inverted Nikon fluorescence microscope equipped with a Princeton Micromax cooled CDD. Due to the extreme variability in axon number and the extensive neurite branching observed in the E l 7 explants, the number of neurites per explant was classified semi-quantitatively to fall into one of the following four categories: zero neurites (no outgrowth), a single neurite per explant (low outgrowth), from 2 to 10 neurites per explant (moderate outgrowth), and greater than 10 neurites per explant (robust growth). The substantial neurite outgrowth from E8 explants was assessed by first tracing neurites from the captured images using Photoshop 5.0 (Adobe Systems Inc., San Jose, CA). Mean neurite length was calculated from the tracings using NTH Image verl.61 by averaging the 50 longest neurites from each explant. The mean neurite length was then compared between different populations of brainstem-spinal neurons using a Student's t-test. Immunohistochemistry Explants were plated on laminin-coated (50 pg/ml) glass coverslips (12 mm round, Fisher). After 3 days, cultures were fixed with 4% paraformaldehyde (BDH Laboratory Supplies, Poole, England). Prior to processing for 5-hydroxy-tryptamine (5-HT) 44 immunohistochemistry, D i l labeled neurites were photographed with a rhodamine filter set using a Ziess Axioscope (Carl Ziess, Thornwood, NY) and a Spot colour digital camera (Diagnostic Instruments, Sterling Heights, MI). Non-specific binding sites in the cultures were then blocked for 30 minutes in PBS containing 10% normal goat serum (Jackson ImmunoResearch, West Grove, PA) and 0.3% Triton X-100 (Sigma, St. Louis, MO). The primary 5-HT antibody (Eugene Tech, Ridgefield Park, NJ) was applied to the explants at 1:2000 dilution in blocking solution and left overnight at 4°C. Cultures were rinsed in PBS followed by a 1 hr. incubation at room temperature with Rhodol green conjugated goat anti-rabbit secondary antibody (Molecular Probes) diluted 1:200 in PBS containing 1% normal goat serum and 0.3% Triton X-100. Cultures were then rinsed in PBS, and coverslips applied prior to examination with a FITC filter set. Images of the 5-HT immunopositive neurites were prepared from the same region previously examined for D i l labeling. Cell Viability Counts To assess the number of healthy neurons per explant, fixed explants were cryo-protected in 18% sucrose, embedded in Tissue-Tek (Sakura Finetek U.S.A., Inc., C A , USA), and frozen at -21°C. Serial sets of cyro-sections were prepared at a thickness of 20 pm. Viability of D i l labeled cells was confirmed by nuclear staining explant sections with 1 u.g/ml Bisbenzimide H 33258 (Sigma) and observing nuclear morphology. Every second section was scored by counting only Dil-labeled cells viewed under a 40X objective. A measure of cell size was obtained by capturing images with the Spot camera and tracing Dil-labeled cell bodies with the computer program Nothern Eclipse (Empix Imaging, Inc., Mississagua, Ont., Canada). A mean cell diameter was then calculated from these measurements by assuming a spherical shaped cell. A significance difference was checked using a Student's t-test. R E S U L T S Brainstem-Spinal Neuron Labeling To study the competence of brainstem-spinal neurons to regrow neurites in vitro, primary explant cultures were prepared from the brainstems of chicks. Brainstem-spinal neurons were 45 retrogradely labeled prior to culturing with carbocyanine dye crystals implanted in ovo into the cervical spinal cord of E5 chicks. Whole mount fluorescent microscopy of E8 brainstems labeled with D i l displayed all five of the populations examined in this study: the raphe nucleus, the medullary reticular formation, the vestibular complex, the pontine reticular formation, and the midbrain nuclei consisting of the Red nucleus and the interstitial nucleus of Cajal (Fig. 2-1A and data not shown). Specific explants were produced by micro-dissection of 300 pm (E8) or 350 pm (E17) thick brainstem slices. For example, two distinct explants containing E l 7 vestibulospinal neurons were obtained by the micro-dissection of one slice from the pontomedullary junction (Fig. 2-1B). Slices from the medulla contained both raphe-spinal and medullary reticulospinal neurons (Fig. 2-1C). Thus, each micro-dissection produced 2 explants containing reticulospinal neurons and one explant containing raphe-spinal neurons. Similar micro-dissections of tissue slices from the appropriate rostro-caudal level of the brainstem produced explants containing the other brainstem-spinal populations studied, including explants containing pontine reticulospinal neurons and midbrain explants (not shown). Note that the close proximity of the Red nucleus and the interstitial nucleus of Cajal precluded their accurate separation by micro-dissection. Thus, explants containing both of these midbrain-spinal populations were used. Brainstem-Spinal Neurite Outgrowth Explants were grown in serum-free Neurobasal media on laminin substrates for two (E8 explants) or three (E17 explants) days. Substantial neurite outgrowth was consistently obtained from both E8 raphe and vestibular explants (Figs. 2-2A and 2-2B), as well as from E8 explants containing pontine reticulospinal, medullary reticulospinal, and midbrain neurons (data not shown). The observed neurite outgrowth was unbranched (Figs. 2-2A and 2-2B) and dense, typically containing thick bundles of fasciculated neurites which are particularly evident under phase-contrast optics. In one representative experiment, the mean neurite length from raphe and vestibular explants was 860 + 15 pm and 777 ± 1 8 pm, respectively (n = 200; 50 longest neurites or neurite bundles from each of 4 explants per nuclei were averaged). Occasionally, neurites were measured at over 1.5 mm in length. Shorter neurites were not counted due to the difficulty in resolving individual processes or bundles within the dense mat of neurites closest to the explants. Similarly, robust neurite 46 Figure 2-1. Retrograde D i l labeling of chick brainstem-spinal neurons. (A) Raphe-, vestibulo-, and medullary and pontine reticulospinal neurons are visible from the ventral surface of a whole mount E8 brainstem. Transverse sections of E l 7 brainstems reveal (B) vestibulo-, (C) raphe- and medullary reticulospinal neurons. V = vestibular complex; R = raphe-spinal neurons; Rp = pontine reticulospinal neurons; Rm = medullary reticulospinal neurons. Scale bar = 1000 urn. 47 48 Figure 2-2. Differential neurite outgrowth of young and mature raphe- and vestibulospinal neurons grown in serum-free media on laminin substrates. D i l label is incorporated into the growing neurites to reveal robust outgrowth from both E8 raphe (A) and vestibular (B) explants obtained from retrogradely-labeled brainstems. Substantial outgrowth is also obtained from E l 7 raphe explants (C), but not vestibular explants (D). Computer tracings of E l 7 outgrowth from C and D more clearly reveal raphe (E) and vestibulospinal (F) neurite outgrowth. V = vestibular explant; R = raphe explant; Scale bar = 1000 um. E F v F 50 outgrowth was also obtained from both raphe and vestibular explants when initial plating was begun at ages up to E l 2 (data not shown). From all five types of mature E l 7 explants, Dil-labeled neurites were occasionally observed growing on laminin substrates, albeit in numbers far fewer than from E8 explants. Qualitative observations clearly indicated that neurite outgrowth was greatly reduced between E8 explants (Figs. 2-2A and 2-2B) and E l 7 explants (Figs. 2-2C and 2-2D). This is exemplified by the majority of E l 7 explants that produced no neurites at all, with the exception of raphe explants (see below and Fig. 2-3). Morphologically, E l 7 neurites were typically "wiry-looking", meandering, branched, and usually not found in fascicles (Figs. 2-2C and 2-2D). In short, the sparse neurite outgrowth from E l 7 explants was in contrast to the dense, unbranched, and fasciculated growth of E8 neurites. Note, the lack of substantial neurite outgrowth from E l 7 explants was not due to decreased neuronal survival in culture (see below). Because initial observations indicated that E l 7 raphe explants produced substantially more neurite outgrowth than other mature explants, we assessed the growth capabilities of distinct explants of E17 brainstem-spinal neurons. E17 explants were cultured in separate wells for 3 days. The number of Dil-labeled neurites grown from each explant was counted and expressed as one of four categories (Fig. 2-3): no outgrowth (zero neurites), single neurites, moderate outgrowth (between 2 and 10 neurites per explant) and robust outgrowth (greater than 10 neurites per explant). The majority of E17 explants containing midbrain-, pontine reticulo-, vestibulo-, and medullary reticulo-spinal neurons produced no outgrowth (73, 85, 85, and 66% of all explants examined, respectively). Robust outgrowth was seen from these explants in only 2.4, 4.3, 1.5, and 1.3% respectively. In contrast, only a small fraction (6.5%) of E l 7 raphe explants produced no outgrowth, while more than a third (35%) of E17 raphe explants produced robust levels of neurite outgrowth. This measure is likely to have underestimated the differences between brainstem-spinal populations, as the number of neurites produced from robust E l 7 raphe explants was far greater than robust explants containing other E l 7 brainstem-spinal neurons. Many raphe explants produced over 50 discernable neurites, while the best example from any of the other brainstem-spinal populations was one vestibular explant that produced 22 labeled neurites. Neurite length was not considered because of the very limited number of neurites produced 51 Figure 2-3. Comparison of neurite outgrowth from five populations of E l 7 brainstem-spinal neurons. Neurite outgrowth is scored as belonging to one of four categories: robust = >10 neurites per explant; moderate = from 2 to 10 neurites per explant; single neurites; and zero neurites. The raw number of explants in each category is tabled below the chart. Raphe-spinal neurons produced more moderate and robust outgrowth than the other populations. 52 NEURITE O U T G R O W T H F R O M E l 7 B R A I N S T E M - S P I N A L E X P L A N T S *-> c JS CL X LU c CJ Q_ 10% 90% 80% 70% 60% 50% 40% 30% 20% 10% 0% Raphe Medullary Reticular Vest ibular Pontine Reticular Midbrain IB Robust O u t g r o w t h 38 • Moderate O u t g r o w t h 51 21 11 • Single Axons 12 I Zero Growth 51 112 40 30 Brainstem-Spinal Explant 53 by all but the raphe explants. However, a qualitative examination indicated that raphe explants also produced far longer neurites than any of the explants containing other brainstem-spinal neurons (Figs. 2-C and 2-2D, and data not shown). Interestingly, neurite outgrowth from both young and mature brainstem-spinal neurons displayed a clockwise tendency (Fig. 2-2). Clockwise neurite outgrowth from retinal ganglion cells (RGCs) has also been reported in fish (Heacock and Agranoff, 1977) and Xenopus (Grant and Tseng, 1986) when grown on laminin or polylysine, and from the chick and mouse (Halfter et al, 1987) when grown on basal lamina. However, R G C outgrowth is straight when growing on three-dimensional substrata (Halfter et al, 1987), thus leading these authors to propose that neurite growth might be forced into a clockwise pattern on a two-dimensional substrate, perhaps due to the helical cytoskeletal architecture of axons. To our knowledge, this is the first report of clockwise neurite outgrowth from any neuronal population other than RGCs. This may be an inherent feature of long projection CNS axons. Brainstem-Spinal Neuron Survival Because the majority of E l 7 explants produced no neurite outgrowth, we examined the viability of the cultured brainstem-spinal neurons. After 3 days in culture, fixed explants were cryosectioned and stained with the fluorescent D N A binding dye, Bisbenzimide H 33258. Examination of stained sections showed many large, brightly-labeled Dil-containing neurons with nuclear morphology indicative of healthy cells (Fig. 2-4). Even from explants that produced zero neurite outgrowth in culture (e.g. the vestibular explant shown in Fig. 2-4), numerous healthy cells were seen. Dil-labeled neurons with apoptotic nuclear morphologies were rarely detected. The difference in neurite outgrowth from raphe explants when compared to explants of the other brainstem-spinal neurons necessitated an examination of cell survival between the different phenotypes. After 3 days in culture, E l 7 raphe and vestibular explants were serial sectioned and stained with Bisbenzimide. Bright, Dil-containing neurons were counted from every second section. No significant difference in the mean neuron number per explant was found between raphe and vestibular explants (419 ± 48 and 548 ± 95, respectively; n=4 explants each). The mean cell diameters were 18.0 ± 0.3 pm (n=25 cells) and 23.3 ± 0.8 pm (n=24 cells) for raphe-spinal and vestibulospinal neurons, respectively. Because we were 54 F i g u r e 2-4. D i l labeled E l 7 vestibulospinal neurons. After 3 days in culture, many large and healthy neurons with nuclear staining indicative of living cells (asterisks) are seen in 20 pm transverse sections through the explant. These cells are clearly distinct from cells with apoptotic morphologies (arrowhead). Scale bar = 30 urn. 55 56 primarily interested in whether a similar number of healthy cells were present in explants from both strong and weak populations of regenerating brainstem-spinal neurons, and because of the similarity between cell sizes, a cell count correction to more accurately assess the cell number was not performed. To compare the survival of E l 7 neurons with E8 neurons, E8 vestibular explants were grown for two days and healthy cells were counted as described above. The mean number of healthy neurons per E8 vestibular explant was 144 ± 20 (n=4). In some cases, E l 7 explants were grown for longer periods. This was to provide extended time for the upregulation of genes that may be necessary for promoting neurite outgrowth. It has been shown that explants of adult rat RGCs can extend neurites almost immediately onto laminin-coated surfaces after a priming lesion of the optic nerve in vivo 7 days before plating (Meyer and Miotke, 1990). Adult RGCs can also extend neurites after 5 days in culture when explanted without a prior optic nerve lesion (Meyer and Miotke, 1990). To this end, vestibular explants were cultured for 9 days. Once again, neurite outgrowth was minimal; however, numerous healthy neurons were still observed within Bisbenzimide-stained sections (data not shown). 5-HT Immunohistochemistry After previously capturing the outgrowth of Dil-labeled neurites with a digital camera using a rhodamine filter set, the neurotransmitter phenotype of neurites growing from raphe explants was determined in fixed explant cultures by incubation with a 5-HT antibody. Subsequently, images of the Rhodol green conjugated secondary antibody were captured using a FITC filter set. Much of the Dil-label was washed out of the neurites during the antibody processing, which necessitated the prior imaging of neurite outgrowth. The staining patterns revealed a majority of the raphe-spinal neurites as double-labeled for both D i l and 5-HT immunoreactivity (Fig. 2-5). On the rare occasion, neurites only labeled with D i l or 5-HT were observed. Vestibular explants that produced neurite outgrowth were also processed for 5-HT immunohistochemistry; however, no 5-HT staining was detected. Thus, the majority of Dil-labeled neurite outgrowth from raphe explants was also positive for 5-HT, confirming that the raphe-spinal population studied was indeed serotonergic. 57 Figure 2-5. Serotonergic immunoreactivity in raphe-spinal neurites. High (A and B) and low (C and D) power images of reveal co-localization of D i l (A and C) and serotonin immunoreactivity (B and D). Scale bar = 63 pm (A and B), 125 pm (C and D). D 59 DISCUSSION The failure of adult CNS regeneration may be due to inhibitory molecules within the mature CNS (e.g. myelin), astroglial scarring and cyst formation at the lesion site, or the intrinsic inability of mature neurons to regenerate. Most likely, it is a combination of these factors (reviewed by Schwab and Bartholdi, 1996). The present study focused on the age-dependent ability of brainstem-spinal neurons to regenerate, as well as whether distinct brainstem-spinal phenotypes have different regenerative capacities. We present here an example of the differential neurite growth capacities of distinct brainstem-spinal populations. This differential effect is seen between young and mature neurons growing on permissive substrates: most mature (El7) brainstem-spinal neurons fail to extend neurites on laminin, whereas robust outgrowth predominates from explants containing younger (E8) brainstem-spinal neurons. As well, at E l 7 , only serotonergic raphe-spinal neurons were capable of robust neurite outgrowth on laminin, whereas all other E l 7 brainstem-spinal populations showed little or no neurite outgrowth. Development of an In Vitro Brainstem-Spinal Neuron Model of Regeneration We have previously reported that the in ovo application of D i l to the cervical spinal cord of E5 chicks produces labeling of brainstem-spinal projections by E8 (Pataky et al, 2000). hi those experiments, we cultured both dissociated and explanted tissue to study brainstem-spinal neuron survival and neurite outgrowth. Here, we extend these techniques to the more mature age of E l 7 , and note that E l 7 brainstem-spinal neurons survive in culture for at least 9 days. The cellular viability was confirmed by observing the nuclear morphology of brightly Dil-labeled cells. This indicates that regardless of the age of culture, hundreds of healthy, labeled brainstem-spinal neurons survive within each explant. Explants of both young (E8) and mature (El7) brainstem-spinal neurons regrew neurites in culture. We use the term "regrow" here because the method for producing a brainstem-spinal explant culture necessarily severs the projection axons that previously transported the D i l label from the spinal cord. The choice of E8 and E l 7 as the starting points for explant cultures coincides with the permissive and restrictive periods, respectively, of brainstem-60 spinal neuron regeneration in the chick (Keirstead et al, 1992). Thus, we have developed an in vitro model that may help elucidate both the intrinsic and extrinsic mechanisms underlying mature brainstem-spinal neuron regeneration failure. Perhaps equally important, this technique enables the study of neuronal phenotypes that are targets for regenerative interventions after spinal cord injury. The observation of 5-HT immunoreactivity within neurites growing from E l 7 raphe explants confirmed that the Dil-labeled neurite outgrowth was serotonergic. As in all vertebrate animals examined, previous studies in the chick have found that the midline raphe-spinal neurons are serotonergic (Ikeda and Goto, 1971; Dube and Parent, 1981). Although the great majority of neurites from raphe explants in the present study were double-labeled for both D i l and 5-HT, we occasionally observed neurites positive for D i l or 5-HT only. Neurites with 5-HT-positive immunoreactivity without concomitant D i l labeling may be explained by the inclusion of rostrally projecting raphe neurons in the explant (Ikeda and Goto, 1971; Dube and Parent, 1981). Dil-labeled neurites negative for 5-HT likely represented the small population of non-serotonergic raphe-spinal neurons (Cabot et al, 1982). Brainstem-Spinal Neurite Outgrowth is Age-Dependent At E8, during the permissive period for brainstem-spinal regeneration, we observed robust neurite outgrowth in vitro from brainstem-spinal neurons. However, this growth capacity was greatly diminished at more mature ages, as has been reported for other CNS populations. For example, L i and colleagues (1995) co-cultured young and mature entorhinal and hippocampal explants oriented to support the growth of projections from either explant into the other. They found that only neurites from young explants grew into the other explant, regardless of the target explant age. The more mature neurons did not innervate the target explants, even when presented with younger, presumably more permissive tissue. Similar findings of reduced neurite outgrowth from mature tissue were also reported for cerebellar Purkinje cells (Dusart et al, 1997) and retinotectal neurons (Chen et al, 1995). Considered with our observations, these data suggest that factors other than an extrinsic inhibitory growth environment contribute to the regenerative failure of adult CNS neurons. Although E l 7 brainstem-spinal neurons showed a marked overall decrease in neurite outgrowth capacity, this was not due to decreased neuronal survival. In fact, the number of 61 healthy neurons counted in E l 7 vestibular explants was greater than in E8 explants. This survival difference is partly accounted for by the difference in explant thickness (300 pm for E8 vs. 350 pm for E17). Nevertheless, the data supports the notion that diminished growth capacity at mature ages is not related to greater neuronal death (Chen et al, 1995; Dusart et al, 1997). Nor was the decreased number of Dil-labeled neurites from E17 explants due to any technical difficulty in the visualization of Dil-labeled neurites. Comparisons between fluorescence (Dil) images and phase contrast images did not reveal significant differences in the number of neurites exiting the explants. Another variable thought important for determining the relative regenerative capacities of different neuronal phenotypes is the onset and duration of developmental differentiation. Early onset of differentiation is correlated with the early loss of regenerative potential, while late-developing neurons lose regenerative potential similarly late. One example of a late-developing long projection neuron is the corticospinal tract. After neonatal injury, partial regeneration is observed from the corticospinal tract, mediated by both regenerating and late-developing (uninjured) neurons (Tolbert and Der, 1987; Bregman et al, 1989). In contrast, other supraspinal projections injured at the same age do not appear to recover (Tolbert and Der, 1987; Bregman et al, 1989). Another example of late developing supraspinal neurons that demonstrate regenerative potential later in development is the rubrospinal projection in the opossum (Wang et al, 1994). These studies suggest that compared to early developing neurons, late developing populations are capable of regeneration after injuries later in development. In the chick, the first brainstem-spinal projection to extend is the medullary reticulospinal tract, followed closely by the pontine and interstitial reticular projections (Glover, 1993). Subsequently, the raphe-spinal and the lateral vestibulospinal tracts develop. Later projections emerge from other pontine reticular and vestibular groups, followed finally by the rubrospinal projection (Shiga et al, 1991). Therefore, the raphe-spinal projection extends near the middle of the developmental time frame of brainstem-spinal neurons. Thus, our in vitro data suggests that not all late developing neurons retain a prolonged capacity for regrowth as compared to earlier developing neurons. 62 Differential Outgrowth From Mature Brainstem-Spinal Neurons In the present study, only raphe-spinal neurons retained their capacity for neurite outgrowth at more mature ages, even though the cultures of different E l 7 brainstem-spinal populations showed similar neuronal survival. At E l7 , raphe-spinal neurons were able to extend neurites as early as 24 hours after plating (data not shown). Robust neurite outgrowth was seen from explants of these neurons after 3 days. In contrast, the 4 other brainstem-spinal populations tested did not produce any neurite outgrowth in the majority of the cases, even i f the explant cultures were extended for as long as 9 days. Differential plasticity of adult monoaminergic spinal systems has previously been shown in vivo by Wang and colleagues (1991). They measured the sprouting of uninjured descending fibres into areas of the spinal cord denervated by dorsal rhizotomy. Serotonergic raphe-spinal neurons responded with increased sprouting into the denervated region, but no sprouting from noradrenergic neurons was observed (Wang et al, 1991). No trophic factor treatments or permissive substrates were used to stimulate growth, providing strong evidence that serotonergic neurons have an increased propensity to sprout. Several in vivo spinal cord regeneration studies demonstrated regrowth of injured descending fibres when various treatments (typically modifying the substrate and trophic factor environments of regenerating axons) are employed (c.f. Schwab and Bartholdi, 1996; Bregman, 1998; Fawcett and Geller, 1998). Of particular interest to the present study, axotomized neurons from the red nucleus, locus coeruleus, and raphe nucleus have been reported to regrow into and beyond embryonic spinal cord cells transplanted into lesions in neonatal rats (Bernstein-Goral and Bregman, 1993). A l l three populations grew in similar proportional numbers (i.e. approximately 30-35% of neurons in each nuclei examined regenerated axons past the transplant site). Similarly, Xu and colleagues demonstrated axonal regeneration of several populations of brainstem-spinal neurons into Schwann cell guidance channels grafted into the lesioned and excised spinal cord of adult rats, when BDNF and NT-3 are also applied (Xu et al, 1995). Interestingly, neurons from the vestibular nuclei were the most commonly found supraspinal neurons (67% of retrogradely-labeled supraspinal neurons). Note that brainstem labeling was only detected when trophic factors were applied. In a later report by the same group, supraspinal ingrowth into implanted Schwann cell channels was achieved without the use of trophic factors, but care was taken to ensure the 63 restoration of cerebrospinal fluid flow (Xu et al, 1999). Of the 19 populations of brainstem-spinal neurons they studied, raphe-spinal neurons provided the majority of the supraspinal regrowing fibres (Xu et al, 1999). This report compares well with our finding of retained regenerative potential of mature raphe-spinal explants grown in serum- and growth factor-free culture. Perhaps with appropriate changes in the trophic and/or substrate environments, we will be able to identify the conditions that best facilitate the regrowth of numerous populations of brainstem-spinal neurons at equally late stages of development, or after adult spinal cord injury. The mechanisms underlying the differential capacity for axonal regeneration between different CNS populations are unknown, but may include variations in axon diameter, degree of myelination, response to extracellular signals and/or modulation of intrinsic growth programs (c.f. Schwab and Bartholdi, 1996; Keirstead and Steeves, 1998; Fernandes and Tetzlaff, 2000). For example, after spinal cord transection in the adult zebrafish there is a differential capacity for axonal regrowth that correlates with the ability of specific neuronal populations to upregulate growth associated molecules (Becker et al, 1998). One class of cerebrospinal neurons (from the nucleus of the medial longitudinal fascicle, the intermediate reticular formation, and the magnocellular octaval nucleus) retained the ability for regeneration after a distal lesion and also upregulated mRNA for L l analogues and GAP-43. However, a second group of cerebrospinal neurons (from the nucleus ruber, the nucleus of the lateral lemniscus, and the tangential nucleus) was unable to regenerate following a distal lesion and failed to upregulate these molecules (Becker et al, 1998). Thus, the factors influencing the regenerative capacity of severed CNS axons are undoubtedly complex. In conclusion, our in vitro assessment of identified brainstem-spinal neuron regrowth shows that a differential capacity for neurite outgrowth exists at later embryonic ages. Although the growth from older embryonic raphe-spinal explants is less than that of younger embryonic neurons, raphe-spinal neurons retain an enhanced capacity for growth at later ages compared to the four other brainstem-spinal populations examined. We feel that this model will prove useful in dissecting the mechanisms underlying the regenerative failure of mature CNS neurons. 64 C H A P T E R 3: S U P P R E S S I O N O F R H O - K I N A S E A C T I V I T Y P R O M O T E S N E U R I T E G R O W T H O N I N H I B I T O R Y C N S S U B S T R A T E S 65 As evidenced by the data from the previous chapter and reports described in the introduction, it is becoming increasingly clear that the intrinsic neuronal capacity for axonal regrowth may limit regeneration in the CNS. In an effort to overcome intrinsic neuronal limitations, this chapter presents a pharmacological method to enhance the axonal growth capacity of neurons. Explants of D R G were grown in vitro on substrates associated with the inhibitory CNS environment. The intracellular signaling of inhibitory molecules was suppressed with a specific inhibitor of Rho-kinase. This promoted neurite regrowth on aggrecan, myelin, and spinal cord. The data suggest that suppression of Rho-kinase activity may enhance the intrinsic capacity of neurons and promote axonal regrowth. This study was recently submitted for publication. It is presented here with formatting consistent with the rest of the thesis. Similar to Chapter 2, further and more general discussion topics are presented in Chapter 4. INTRODUCTION Axonal regeneration in the adult mammalian CNS is hampered by several growth-inhibitory molecules found both in the injury scar and in the distal myelinated tracts (for review, see Fitch and Silver, 1997b; Fournier and Strittmatter, 2001; McGraw et al, 2001). An important class of inhibitory molecules is the chondroitin sulfate proteoglycans (CSPGs), found both in myelin (Niederost et al, 1999) and within the spinal cord injury scar (Lemons et al, 1999). Aggrecan is one prominent member of the CSPG family and is expressed in the developing, adult, and injured spinal cord (Lemons et al, 2001). Aggrecan is inhibitory to sensory and retinal neurites grown in vitro (Snow and Letourneau, 1992; Challacombe et al, 1997). Similarly, other members of the CSPG family have been shown to be both inhibitory to neurite growth and expressed in the CNS following injury (McKeon et al, 1991; Dow et al, 1994; Gates et al, 1996; Dou and Levine, 1997; Fitch and Silver, 1997a; Davies et al, 1999; Lemons et al, 1999). The extent to which individual inhibitory molecules contribute to axonal regeneration failure is less clear: it is difficult to predict the number and the nature of the inhibitors that should be neutralized in order to achieve functional regeneration and recovery. 66 Axonal growth inhibition is accompanied by distinct changes in growth cone behavior and morphology, mediated primarily through the action of the Rho family of small guanosine triphosphatases (GTPases) on the actin cytoskeleton (Luo, 2000). In non-neuronal cells, activation of Rac and Cdc42 induces the formation of lamellipodia and filopodia respectively, while Rho activation induces stress fiber and focal adhesion formation (Nobes and Hall, 1995). The picture is not as clear in neurons; however, Rho activation has been shown to be at least partly responsible for the effects of several neurite growth inhibitors including Ephrin-A5 induced collapse of retinal axons (Wahl et al, 2000), myelin inhibition of retinal and dorsal root ganglion (DRG) cells (Jin and Strittmatter, 1997; Lehmann et al., 1999), and Semaphorin 3A inhibition of D R G growth (Jin and Strittmatter, 1997). Since the effects of several inhibitors of regeneration and repulsive guidance cues appear to be mediated by Rho, a viable approach to stimulating axonal growth may be to block growth inhibitory molecules simultaneously by interfering with the Rho pathway (McKerracher, 2001). Rho has several downstream effectors (Hall, 1998) among which R O C K may be an attractive target for therapeutic intervention. R O C K (Rho-kinase, R O K a , ROCK-2) is a recently identified serine/threonine kinase that is highly expressed in brain (Matsui et al., 1996; Hashimoto et al, 1999). R O C K mediates the signaling of Rho to the actin cytoskeleton through several downstream targets (Amano et al, 2000). For example, R O C K regulates the phosphorylation of myosin light chain (MLC), both by direct phosphorylation of M L C (Amano et al, 1996) and by the inactivation of myosin phosphatase (Kimura et al, 1996). As well, R O C K phosphorylates LM-kinase (Maekawa et al, 1999; Ohashi et al, 2000), which in turn is necessary for Sema3A-induced growth cone collapse mediated via phosphorylation of cofilin (Aizawa et al, 2001). In order to test whether R O C K signaling is important to inhibiting axonal outgrowth, I have used a cell-permeable pyridine derivative, (+)-R-trans-4-(l-aminoethyl)-N-(4-pyridyl)-cyclohexanecarboxamide (Y-27632) (Ishizaki et al, 2000), to suppress R O C K activity. Whole explants of chick D R G were grown on inhibitory substrates of aggrecan, myelin, and adult spinal cord cryo-sections in the presence or absence of Y-27632 and examined for neurite outgrowth and growth cone morphology. 67 M A T E R I A L S A N D M E T H O D S Reagents were obtained from Canadian Life Technologies (Burlington, ON) unless otherwise indicated. (+)-R-trans-4-(l-aminoethyl)-N-(4-pyridyl) cyclohexane carboxamide (Y-27632) was synthesized at Merck Frosst as the bis trifluroacetic acid (TFA) salt. The majority of experiments were performed using the TFA salt of Y-27632. In some experiments, the activity of Y-27632 obtained commercially from Biomol Research Laboratories (Plymouth Meeting, PA) was examined. This commercial source of Y-27632 was provided as the dihydrochloride salt of Y-27632 (Y-27632-2HC1). Unless otherwise indicated, all experiments performed with Y-27632 were completed using the TFA salt. A direct comparison revealed similar results with both the TFA and 2HC1 salt formulations of Y -27632. A l l surgical procedures and animal care protocols were performed in accordance with the Canadian Council for Animal Care and approval of the local animal care committee. Tissue Culture and Neurite Outgrowth Assay Whole D R G explants were removed from the lumbar spinal cord of embryonic day (E) 9 chicks. D R G were plated individually in 48-well Nunc culture plates pre-coated for 2 hours at 37°C with a solution of 12.5 pg/ml laminin plus 0 to 50 pg/ml aggrecan (Sigma, St. Louis, MO). Cultures were grown in a humidified 37°C incubator with a 5%C02/95% air atmosphere in serum-free Neurobasal medium supplemented with B-27 nutrients, glutamine (0.5 mM), and 100 units/ml penicillin/streptomycin. Media were also supplemented with Nerve Growth Factor (NGF), Brain Derived Neurotrophic Factor (BDNF), or Neurotrophin-3 (NT-3) at 20 ng/ml (kindly provided by Regeneron Pharmaceuticals Inc.). Before incubating, the R O C K inhibitor, Y-27632 (kindly provided by Merck-Frosst, Kirkland, PQ), was added at concentrations of 0-50 pM. After 18-36 hours, cultures were fixed with 10% formalin (Fisher Scientific, Nepean, ON) in 0.1 M phosphate buffered saline (PBS). Phase-contrast images of the D R G neurites were captured with a 4X objective on an inverted Nikon microscope equipped with a Princeton Instruments Micromax cooled CDD. The outgrowth was assessed by tracing neurites from the captured images using NTH Image verl.61, as described previously (Borisoff et al, 2000). Mean neurite length was determined by pooling 68 the lengths of the longest 10 neurites per explant from each treatment group. Only 10 neurites per explant were considered due to the sparse growth of control explants. Total neurite length was determined by the summation of the lengths of all neurites from each explant, and thus considers both length and the number of neurites growing from each explant. Myelin Substrates To challenge regenerating axons with more complex inhibitory environments, myelin membrane fractions and adult rat spinal cord were used as growth substrates for E l 5 chick D R G in identical media conditions as described above. Myelin was prepared from a standard protocol (Poduslo, 1975). Tissue was isolated from adult (200-250 g) Sprague-Dawley rat brains. Rats were anesthetized with a lethal dose of chloral hydrate at 900 mg/kg (BDH Laboratory Supplies, Poole, England) and the brain was removed and ground to a powder with a mortar and pestle on dry ice. The powdered tissue was then homogenized in 0.25 M sucrose in PBS containing a protease inhibitor cocktail (Roche Diagnostics, Laval, PQ). The supernatant was then overlaid on 0.85 M sucrose in PBS and spun at 75000 g for 30 min. Material at the interface was re-suspended in ice cold water and centrifuged for 25 min at 25,000 g. The pellet was collected, re-suspended in ice cold water, and centrifuged for 15 min at 10,000g. This was repeated once more, before the pellet was re-suspended in 0.25 M sucrose and overlaid on 0.85 M sucrose and centrifuged for l h at 75,000 g. The purified myelin was collected at the interface and washed twice with ice cold water. To prepare the culture substrate, 25 ul drops of myelin solution (at a protein concentration of 4 ug/ml) mixed with 12.5 pg/ml laminin were dried overnight onto 48-well culture trays (Nunc), yielding 0.1 u.g of myelin protein deposited. E l 5 D R G explants were plated onto the dried myelin substrates in the same medium described above. Fixed cultures were stained with Alexa-546-conjugated phalloidin (Molecular Probes). Fluorescent-labeled neurite outgrowth was imaged and analyzed as described above. Spinal Cord Cryo-Sections Spinal cord substrates were prepared from adult Sprague-Dawley rats (200-250 g). Rats were anesthetized with a lethal dose of chloral hydrate and perfused with ice-cold 0.1 M 69 PBS. Cervical spinal cord was then removed and frozen at -80 °C. Frozen horizontal sections were cut at 20 pm with a cryostat and mounted onto the bottom of 35 mm plastic tissue culture dishes (Nunc). Approximately 1.5 ml of medium was added to the dishes and D R G explants were placed onto the spinal cord sections. Medium level was adjusted such that the meniscus kept the explants from floating off the sections while maintaining complete coverage of the tissue. After 6-8 hours incubation, medium was added to a final volume of 3 ml and cultures were grown for 2-3 days. To label the neurite outgrowth, the vital dye, 5-(and-6)-carboxyfluorescein diacetate, succinimidyl ester (5(6)-CFDA,SE; Molecular Probes, Eugene, OR) was used. CFDA,SE was diluted 1:10 in PBS and 100 pi was added to 3 ml of medium. Cultures were then incubated for 20 minutes and rinsed 3 times in PBS before fixation in 10% formalin. Images of neurite outgrowth were immediately captured with a Zeiss Axioscope (Carl Zeiss, Thornwood, NY) and a Spot colour digital camera (Diagnostic Instruments, Sterling Heights, MI). Neurite growth was measured in two different ways. First, total growth and length (as described above) was quantified irrespective of whether the growth occurred on grey or white matter. Second, a ratio of total neurite growth on white versus grey matter was calculated. Only explants that straddled both grey and white matter were considered in this calculation. Note: outgrowth from each explant was measured from an equal area of substrate for either grey or white matter. Growth Cone Measurements E9 D R G were explanted onto glass coverslips (12 mm round, Fisher) coated with 12.5 pg/ml laminin plus 6.25 pg/ml aggrecan, or laminin alone. In these experiments, D R G were grown overnight in NGF supplemented media. After 18 hours, 50 pM Y-27632 was added to the media, and cultures were fixed in 4% paraformaldehyde (BDH) after 30, 60, or 90 minutes. Cells were rendered permeable in 0.1 M PBS with 0.1% Triton X-100 (PBS-X) (Sigma) for 30 minutes. Alexa-546-conjugated phalloidin (Molecular Probes) was then added at a concentration of 1 unit per 200 pi and incubated at 37°C for 20 minutes. Fluorescent images of growth cones were captured with a 100X oil immersion objective on the above described microscope/camera setup. To analyze growth cone area, images were traced in Photoshop 6.0 (Adobe Systems Inc., San Jose, CA) and the number of pixels per growth cone was recorded. A growth cone was defined as the enlarged area distal to where the initial 70 splaying of the neurite began, including all lamellipodia and filopodia. To analyze filopodia number and length, all filopodia on each growth cone were traced in NIH Image. Immunohistochemistry D R G grown on glass coverslips were incubated for 1 hr at 37°C with the primary antibody, anti-ROCK-2 (Cat. # H-85, Santa Cruz Biotech, Santa Cruz, CA), at a dilution of 1:100 in PBS-X. Following a 30 minute blocking step in 10% donkey serum, Alexa-488-conjugated donkey anti-rabbit antibody (Santa Cruz) was added at a dilution of 1:500 in PBS-X for 1 hr at 37°C. Western Blots E9 spinal cord (SC) and D R G tissue samples were homogenized in 0.01 M Tris-HCI buffer containing a protease inhibitor cocktail (Boehringer), and aliquots containing 10-20 pg total protein were subjected to conventional 10%> SDS-polyacrylamide gel electrophoresis. The separated proteins were transferred onto Immobilon-P Transfer Membrane (Millipore, Nepean, ON) and blocked with 5% skim milk powder (BLOTTO) in TBST for 1 hour. The membranes were incubated with rabbit anti-ROCK-2 (Santa Cruz, Cat. # H-85) antibodies and visualized by peroxidase-conjugated goat anti-rabbit IgG secondary antibody (Jackson ImmunoResearch, West Grove, PA). ROCK Kinase Assay A commercial kinase assay (S6 Kinase Assay, Upstate Biotechnology, NY) was used to measure the effect of Y-27632 on R O C K activity, following the manufacturer's protocol. Recombinant active ROCK-2 (Upstate) at concentrations of 5 mU or 20 mU were used in the kinase assay. Y-27632 was added to the assay solution to a final concentration ranging from 0 to 15000 nM. R O C K activity was measured by the incorporation of P3 2-labeled ATP into the substrate peptide. Activity was expressed as counts per minute (cpm) minus background cpm (activity without either R O C K or kinase substrate present). 71 R E S U L T S ROCK is Expressed in Chick DRG The expression of R O C K has recently been demonstrated at high levels in brain (Matsui et al, 1996), including bovine pyramidal neurons, Purkinje cells (Hashimoto et al, 1999), and chick retinal ganglion cells (Wahl et al, 2000). To confirm the presence of R O C K protein in chick DRG, cultures grown on glass coverslips were stained with an antibody to ROCK-2. Labeled growth cones were imaged at high magnification and observed to display distinct expression of R O C K immunoreactivity (Fig. 3-1 A). The same growth cone stained with phalloidin to label actin is shown in Figure 3-IB. Western blots of E9 chick spinal cord and D R G using the same antibody confirmed the presence of R O C K protein as a single band of approximately 160 kD (Fig. 3-1C). Thus, R O C K protein is expressed by D R G cells and is present in D R G growth cones. ROCK is Inhibited by Y-27632 Recently, a pyridine-derivative, Y-27632-2HC1 (known as Y-27632), was synthesized and reported as a specific inhibitor of Rho-kinase family enzymes (Uehata et al, 1997). It is well established that Y'-21'632 inhibits the activity of R O C K in a variety of systems (Ishizaki et al, 2000). Y-27632 inhibits R O C K activity by competitive binding with ATP to the catalytic site (Ishizaki et al, 2000). The specificity of Y-27632 for R O C K was reported to be 100 times greater than that for protein kinase A , protein kinase C, or myosin light chain kinase (Uehata et al, 1997), as well as over 20 times greater than that for two other downstream Rho effectors, citron kinase and protein kinase N (Ishizaki et al, 2000). In order to confirm the inhibitory effects of Y-27632 (i.e. the TFA salt) on R O C K activity, a commercially available kinase assay was used to measure R O C K activity in the presence of increasing concentrations of Y-27632. Recombinant active-ROCK was incubated with the S6 Kinase Substrate peptide and P32-labeled ATP. The S6 peptide is an amino-acid sequence that includes the serine phosphorylation site recognized by ROCK. Y-27632 was added to the mixture at concentrations ranging from 0 to 15000 nM. Y-27632 significantly reduced the incorporation of P32-labeled ATP into the substrate peptide in a dose-dependent 72 Figure 3-1. R O C K immunoreactivity. High magnification images of growth cones from cultures stained with .anti-ROCK (A) or phalloidin to label actin (B). C, Western blots revealed R O C K protein present in E9 D R G and spinal cord (SC). D, R O C K kinase activity assay demonstrated the efficacy of Y-27632 treatment to significantly reduce the incorporation of P3 2-labeled-ATP into the S6 substrate peptide. P < 0.0001, A N O V A . Scale bar = 10 urn. log[ Y-27632 Concentration (nM) ] 74 manner (p < 0.0001, A N O V A ) . Y-27632 inhibits the activity of R O C K with an I C 5 0 of 540 nM (non-linear regression analysis, R 2 = 0.98, Fig. 3-ID). Aggrecan Substrates Inhibit the Growth of DRG Neurites Aggrecan is a member of the CSPG family of neurite growth inhibitors and is present in the developing, adult, and injured CNS (Lemons et al, 2001). An established in vitro model to assess axonal growth inhibition by CSPGs employs as a substrate the combination of aggrecan and laminin at various ratios (Snow and Letourneau, 1992; Condic et al, 1999). Whole D R G from E9 chick were explanted onto these substrates in serum-free media supplemented with NGF at 20 ng/ml. After 18-36 hours in culture, a halo of neurite growth formed in a manner inversely proportional to the concentration of aggrecan present (ranging from 0-50 pg/ml) in the substrate while the laminin concentration was held constant at 12.5 Lig/ml (Fig. 3-2). On 3 ixg/ml aggrecan, total neurite outgrowth per explant was 27 ± 8 mm (mean ± SEM, throughout the chapter), while on 25 and 50 u.g/ml aggrecan, total outgrowth was massively reduced to 3.6 ± 0.7 and 1.0 ± 0.3 mm respectively. This inhibition was significant in a dose-dependent manner (p < 0.0001, one-way A N O V A ) . Unless otherwise indicated, an aggrecan concentration of 50 p-g/ml was used in the remainder of the experiments described below, in agreement with previously used concentrations (Condic et al, 1999). ROCK Inhibition Stimulates Neurite Outgrowth on Aggrecan Suppression of the Rho GTPase signaling pathway has been shown to overcome neurite growth inhibition due to myelin, M A G , Collapsin-1 (Sema3A), and A5 Ephrins (Jin and Strittmatter, 1997; Lehmann et al, 1999; Wahl et al, 2000). Y-27632 stimulated neurite growth on aggrecan in a concentration dependent manner (Fig. 3-3). Qualitatively, a long, dense halo of neurite growth was usually observed in treated wells compared to the short, sparse growth of controls (Fig. 3-3A and 3-3B). Total outgrowth significantly increased in a dose-dependent manner, peaking between 25 and 50 uM of R O C K inhibitor (p = 0.007, A N O V A ) . The stimulated outgrowth also resulted in significantly longer neurites (p < 0.0001, A N O V A ) , measured by the length of the longest 10 neurites per explant (Fig. 3-3C 75 Figure 3-2. Aggrecan inhibits chick D R G neurite outgrowth. E9 NGF-treated D R G explants were grown on inhibitory aggrecan/laminin substrates. With laminin concentration constant at 12.5 pg/ml, aggrecan inhibited both total outgrowth (A) and length (B) in a dose-dependent manner, n = 6 or 7 explants per treatment, p < 0.0001, A N O V A . 76 120000 E sz % o CO ,o 80000H 40000 E CD c 0 1200n 800 400 0 3.13 6.25 12.5 25 50 B 0 3.13 6.25 12.5 25 50 Aggrecan Concentration (ug/ml) 77 Figure 3-3. Y-27632 promoted neurite outgrowth from E9 NGF-treated D R G explants grown on inhibitory aggrecan substrates. Low magnification phase contrast images of vehicle treated control (A) and Y-27632 (25 pM) treated (B) D R G neurite outgrowth. Dose-response curves of Y-27632 treatment are shown for total outgrowth (C) and length (D). Y-27632 treatment was significant, measured by A N O V A , p < 0.01. n = 6 or 7 explants per treatment. Scale bar = 200 urn. 2000 D 400 I 0 5 10 25 50 c o £ o O) »fe C O s e CD Q. 300 200-100-0 5 10 25 50 Y-27632 Concentration (uM) 79 and 3-3D). Only 10 neurites per explant were considered due to the sparse growth from control explants. The effect of a single dose of R O C K inhibitor (50 pM) on cultures grown on various concentrations of aggrecan was also tested. When D R G neurites grew on substrates of high aggrecan concentration that are inhibitory to growth, Y-27632 significantly increased both total outgrowth and length (p < 0.005, two-way A N O V A ; Fig. 3-4A and 3-4B). In comparison, growth on permissive substrates of low aggrecan concentration was not affected by Y-27632 (total outgrowth, p > 0.05; Fig. 3-4C and 3-4D). It is important to note that as the ratio of aggrecan to laminin lessened, the relative stimulative effect of Y-27632 also lessened. Thus, under permissive conditions such as a high laminin to aggrecan ratio, the effect of Y-27632 was minimal or completely absent. Y-27632 appears to stimulate growth on inhibitory substrates only, rather than causing the general effect of a basal promotion of neurite extension regardless of substrate. In another experiment, I compared the activities of the Merck Frosst synthesized Y -27632 (TFA salt) with that of Y-27632-2HC1 obtained commercially from Biomol Research Laboratories (Plymouth Meeting, PA). Similar to the TFA salt of Y-27632, Y-27632-2HC1 at a concentration of 25 pM improved the outgrowth of NGF-stimulated D R G grown on the aggrecan (50 pg/ml) substrate by 650 ± 180 % (p < 0.05, post-hoc Dunnett's test). Also comparable to the TFA salt of Y-27632, Y-27632-2HC1 treatment significantly increased both total outgrowth (p = 0.005, A N O V A ) and length (p < 0.0001) in a dose-dependant manner over a concentration range from 5 pM to 50 pM. D R G comprise a heterogeneous population of neurons with distinct trophic factor sensitivities. The previous experiments were all performed on NGF-responsive D R G neurons. To determine i f other populations of D R G are also amenable to the growth promoting effects of Y-27632 treatment, I grew D R G explants in the presence of either 20 ng/ml BDNF or NT-3. In both situations, 25 pM and 50 pM Y-27632 significantly increased both total outgrowth and length (Fig. 3-5). ROCK Inhibition Stimulates Neurite Outgrowth on Myelin and Spinal Cord I tested the ability of Y-27632 to stimulate neurite growth on the more complex inhibitory environments of myelin membrane fractions and adult rat spinal cord cryo-80 Figure 3-4. Y-27632 (25 uM) promoted neurite outgrowth from E9 NGF-treated D R G explants grown on substrates of increasing concentrations of aggrecan. Y-27632 treatment significantly increased both total outgrowth (A) and length (B) of D R G neurites grown on 4 different inhibitory concentrations of aggrecan. The effects of Y-27632 treatment were minimal on both total outgrowth (C) and length (D) for growth on low (permissive) concentrations of aggrecan. Note: the concentration of laminin was fixed at 12.5 pg/ml throughout. As the ratio of aggrecan to laminin decreases to favor a more permissive growth environment, the effect of Y-27632 treatment also decreases. Rather than causing a general effect of increased axonal extension, Y-27632 appears to promote D R G neurite growth on only inhibitory substrates, n = 5 or 6 explants per treatment group, p < 0.005, 2-way A N O V A (for A and B). Aggrecan Concentration (ng/ml) 82 Figure 3-5. Y-27632 (25 pM) promoted neurite outgrowth from BDNF- and NT-3-treated D R G explants grown on inhibitory aggrecan substrates. Increased outgrowth was significant compared to controls, n = 7 explants for each treatment. *, p < 0.05; **, p < 0.0001, t-test. 83 A 1500 g 1000 o S 500-o 1— CD Q. B 300 200H 100-BDNF NT-3 Total Outgrowth BDNF NT-3 Length 84 sections. Crude myelin membranes were mixed with laminin (12.5 pg/ml) and dried overnight onto the bottom of tissue culture plates, as previously reported (McKerracher et al, 1994). For both myelin membrane and spinal cord cryo-section experiments, E l 5 D R G were used instead of E9 because the up-regulation of the Nogo receptor occurs at around E l3 in chick, thus rendering younger neurons non-responsive to Nogo and myelin inhibition (Fournier et al, 2001). D R G explants were grown overnight on the myelin substrates in NGF-supplemented media. As shown in Figure 3-6, Y-27632 at a concentration of 50 pM produced a nearly two-fold increase in total outgrowth (from 15700 ± 2800 pm to 27900 ± 4700 pm, p < 0.05, t-test) and a significant increase in length (from 340 ± 9 pm to 400 ± 8 pm, p< 0.0001, t-test). To better mimic the environment in vivo, adult rat spinal cord was fresh frozen, cryo-sectioned horizontally at 20 pm thickness, and mounted onto tissue culture dishes, hi this experiment, chick D R G explants were grown on top of the cryo-sections for 3 days in NGF-supplemented media in the presence or absence of Y-27632 (Fig. 3-7). Y-27632 treatment at a concentration of 50 pM significantly increased both total outgrowth (from 11000 ± 2600 pm to 34300 ± 7200 pm, p < 0.01, t-test) and length (from 370 ± 13 pm to 540 ± 15 pm, p < 0.0001, t-test). As well, Y-27632 treatment resulted in significantly more growth onto spinal cord white matter compared to controls. This was quantified as the ratio of total outgrowth on white matter over grey matter (Fig. 3-7E). Y-27632 treatment resulted in an increased ratio of white vs. grey matter outgrowth (from 0.43 ± 0.08 to 1.19 ± 0.21, p < 0.01, t-test). ROCK Inhibition Alters Growth Cone Morphology Axonal growth inhibition or retraction is often accompanied by growth cone collapse. To observe the effects of aggrecan and R O C K inhibition on growth cone morphology, DRG were grown on glass coverslips coated with inhibitory aggrecan/laminin or laminin alone. In this experiment, an aggrecan concentration of 6.25 pg/ml was used which was still significantly inhibitory (see Figs. 3-2 and 3-4). This allowed more neurite growth to occur than at 50 pg/ml of aggrecan (as used in the previous experiments), enabling the observation of many isolated growth cones. After overnight culture, Y-27632 or vehicle was added to the medium for 90 minutes and the cultures were fixed and stained with a fluorophore-conjugated phalloidin to label actin. 85 Figure 3 - 6 . Y-27632 promoted neurite outgrowth from E l 5 NGF-treated D R G explants grown on inhibitory substrates of myelin membrane fragments. Images of cultures labeled with phalloidin reveal D R G neurite outgrowth from controls (A) and Y-27632 (50 uM) treatment (B). Y-27632 treatment significantly increased both total outgrowth (C) and length (D). n = 16, 14 explants per treatment. *, p < 0.05; **, p < 0.0001, t-test. Scale bar = 100 Lim. 87 Figure 3-7. Y-27632 promoted neurite outgrowth from E l 5 NGF-treated D R G explants grown on inhibitory substrates of longitudinal adult rat spinal cord cryo-sections. Images of cultures labeled with the vital dye CFDA,SE reveal D R G neurite outgrowth from controls (A) and Y-27632 treatment (B). Y-27632 treatment significantly increased both total outgrowth (C) and length (D). n = 10 and 8 explants per treatment. (E) Y-27632 treatment also resulted in an increased growth of D R G neurites on spinal cord white matter vs. grey matter. Data was expressed as the ratio of total neurite outgrowth on white matter over grey matter. Only explants that straddled both grey and white matter were considered in this experiment, n = 4 and 5 explants respectively. *, p < 0.01; **, p < 0.0001, t-test. Note: cultures were grown for 3 days in this experiment. Scale bar = 200 pm. 89 Neurites grown on permissive laminin substrates had large growth cones with many long filopodia, and the presence of very few actin bundles in the central domain of the growth cone (Fig. 3-8A). Growth cones of D R G plated onto inhibitory aggrecan/laminin substrates were also large with many filopodia, although these were much shorter (Fig. 3-8B). An obvious feature of these growth cones, regardless of size or shape, was the presence of distinct actin bundles in the central domain of the growth cone (Fig. 3-8B and Fig. 3-9). After treatment with Y-27632 for 90 minutes, growth cones appeared much smaller, more branched, with fewer but longer filopodia than control growth cones (Fig. 3-8C). Treated growth cones also featured a noticeable decrease in lamellipodial spreading and possessed fewer actin bundles in the central domain (Fig. 3-8C). The quantification of growth cone area was derived from a random sampling of at least 44 isolated growth cones per treatment, and thus considered growth cones of all sizes and shapes from each treatment. Y-27632 treatment 2 2 at 50 pM resulted in: decreased growth cone area from a mean of 130 ± 7 pm to 95 ± 8 pm (p < 0.05, post-hoc Dunnett's test; Fig. 3-8D); decreased number of filopodia per growth cone from 10.7 ± 1.1 to 6.95 ± 0.9 (p < 0.05, Dunnett's test; Fig. 3-8E); and increased length of growth cone filopodia from 5.8 ± 0.4 pm to 8.9 ± 0.5 pm (p < 0.001, Dunnett's test; Fig. 3-8F). In general, Y-27632 treatment appeared to cause more 'streamlined' growth cones (Fig. 3-8G and 3-8H). Thus, inhibition of the Rho signaling pathway increased the outgrowth of D R G neurites on inhibitory aggrecan substrates and resulted in smaller, 'filopodial' growth cones, lacking lamellipodia. To further characterize the presence of actin bundles in the center of growth cones, 30 growth cones from each treatment group were analyzed by manually counting the number of distinct actin bundles in each growth cone (see arrow heads in Fig. 3-9). Although a clear trend toward fewer actin bundles was evident between control cultures and treatment with Y -27632, the decrease was not significant (p > 0.05, A N O V A , Fig. 3-9). However, the difference in actin bundles between non-treated cultures grown on inhibitory aggrecan/laminin vs. laminin-only substrates was more striking. The number of actin bundles significantly decreased from 6.6 ± 0.9 to 4.1 ± 0.6 respectively (p = 0.014, t-test). This data indicates that aggrecan substrates promote the formation of actin bundles in the central domains of D R G growth cones. 90 Figure 3-8. High magnification images of growth cone morphology. Cultures were stained with fluorophore-conjugated phalloidin to label actin filaments. D R G explants were grown on permissive laminin (A) or inhibitory aggrecan (B and C) substrates. Vehicle (B) or Y -27632 (C) was added to cultures for 90 minutes prior to fixation. Note the presence of actin bundles in the centre of the growth cone in B. Growth cone morphology was quantified by area (D), filopodia per growth cone (E), and filopodial length (F). (G-H) Low magnification images of growth cones grown on inhibitory aggrecan substrates and treated with vehicle (G) or 50 p M Y-27632 (H) for 90 minutes prior to fixation. Note: Y-27632 treatment appears to cause 'stream-lined' growth cones. L M denotes that growth was on permissive laminin-only substrates, n = 130, 45, 44, 49 growth cones for D. n = 20 growth cones for each treatment in E. n = 213, 153, 138, 297 filopodia for F. *, p < 0.05; **, p < 0.001, post-hoc Dunnett's test. Scale bar = 10 um for A-C , 40 pm for G-H. 92 Figure 3-9. Inhibitory aggrecan substrates stimulate the formation of actin bundles in the center of growth cones. Growth cones labeled with fluorophore-conjugated phalloidin from D R G explants grown on aggrecan (A) and laminin (B) substrates. The number of actin bundles (arrowheads) per growth cone grown on aggrecan in the presence of various concentrations of Y-27632 was counted (C). L M denotes that growth was on permissive laminin-only substrates, n = 30 for each group. *, p = 0.014, t-test. Scale bar = 10 pm. 0 10 50 LM Y-27632 Concentration (uM) 94 DISCUSSION Numerous axonal growth inhibitors exist in the injured CNS. It is uncertain how many of these inhibitors will need to be neutralized in order to achieve successful regeneration in adult mammals. The activities of growth cone guidance molecules are mediated through secondary signaling pathways and eventually converge onto the actin cytoskeleton (for review, see Suter and Forscher, 2000). Here, it was my approach to target putative convergence points of multiple signaling pathways in order to neutralize several inhibitory molecules simultaneously. One candidate target is Rho-GTPase and its major effector kinase, ROCK. By inhibiting R O C K with Y-27632, I have reduced the inhibition of D R G neurite growth on substrates consisting of aggrecan, myelin, and adult spinal cord. I have also shown that Y-27632 treatment causes distinct changes to growth cone morphology, resulting in streamlined, filopodial growth cones. ROCK Suppression Promotes Neurite Growth on Aggrecan The application of Y-27632 to D R G neurons promoted neurite growth on aggrecan. Aggrecan is a member of the chondroitin sulfate family of proteoglycans (CSPGs) and is expressed throughout the adult CNS (Schwartz et al, 1996; Lemons et al, 2001). Importantly, CSPGs are found in the scar that forms after injury (McKeon et al, 1991; Gates et al, 1996; Dou and Levine, 1997; Fitch and Silver, 1997a; Davies et al, 1999; Lemons et al, 1999) and in the myelinated CNS white matter (Niederost et al, 1999). Little is known about the receptors and intracellular signaling of CSPGs, although it is probable that CSPGs act through receptor based mechanisms in a manner similar to myelin-derived Nogo inhibition (Fournier et al, 2001). Thus far it is known that CSPGs up-regulate intracellular calcium in D R G growth cones (Snow et al, 1994), and that growth inhibition by the NG2 proteoglycan is mediated by an unknown receptor linked to intracellular second messenger pathways: i.e. NG2 binds a cell surface molecule and growth inhibition is partially reversed by pertussis toxin treatment or the pharmacological increase in intracellular calcium or cAMP (Dou and Levine, 1997). Here, I show that aggrecan causes the formation of distinct bundles of actin in the center of growth cones reminiscent of Rho-induced stress fibre 95 formation observed in fibroblasts (Nobes and Hall, 1995), and aggrecan significantly inhibits the growth of D R G neurites. As well, the suppression of R O C K activity through treatment with Y-27632 promoted the growth of D R G neurites on aggrecan. These results implicate the Rho-pathway as important in growth cone signaling induced by inhibitory aggrecan. Rho-GTPases Regulate Neurite Growth on Inhibitory Substrates The actions of Rho family small GTPases on the actin cytoskeleton have been well demonstrated in fibroblasts (Hall, 1998): Rho activation causes the formation of stress fibres and cell contractility, Rac activation stimulates lamellipodia formation, and Cdc42 activation stimulates filopodia formation (Nobes and Hall, 1995). While the role of small GTPases in the control of the neuronal cytoskeleton is less clear, it is apparent that Rho-GTPases are important to neuronal growth and development (Luo, 2000). The Rho-pathway has been implicated in a number of models of axonal growth inhibitors and repulsive guidance cues. Ephrin-A5 causes activation of Rho and inactivation of Rac, resulting in growth cone collapse of retinal ganglion cells (Wahl et al, 2000). This collapse can be overcome by inhibiting Rho activity with C3, or by inhibiting R O C K activity with Y -27632 (Wahl et al, 2000). The treatment of D R G (Jin and Strittmatter, 1997) and retinal ganglion cells (Lehmann et al, 1999) with C3 promoted neurite outgrowth on myelin and myelin-associated glycoprotein (MAG) substrates. As well, C3-treated D R G were rendered non-responsive to Sema3A-induced growth inhibition (Jin and Strittmatter, 1997). Similarly, I have demonstrated here that R O C K inhibition by Y-27632 increases the growth of D R G neurites on aggrecan, myelin, and spinal cord substrates, thus adding to the list of candidate inhibitory molecules that converge onto the Rho signaling pathway. ROCK Suppression Alters Growth Cone Morphology Interestingly, the promotion of neurite growth on aggrecan by inhibiting R O C K activity resulted in a marked alteration in growth cone morphology. Compared to controls, Y-27632 treated growth cones were smaller, lacking in lamellipodia, and had fewer but longer filopodia. Treated cultures also had fewer actin bundles in the central domain of their growth cones. These features are reminiscent of C3-treated D R G growth cones that grew rapidly in Sema-3 A and myelin environments (Jin and Strittmatter, 1997). In this study, it was reported 96 that C3 eliminated 'lamellipodial spreading' while promoting neurite growth (Jin and Strittmatter, 1997). In contrast, others have found that C3 and Y-27632 prevented retinal ganglia growth cone collapse due to Ephrin-A5, although Wahl et al. (2000) did not report neurite growth rate. Additionally, mouse cerebellar cells grown on permissive matrigel substrates responded to C3 and Y-27632 treatment with an increase in axon number and larger growth cones, while constitutively active RhoA transfection produced smaller growth cones (Bito et al, 2000). In this previous experiment, only C3 caused an increase in neurite length, while Y-27632 (10 pM) did not. This may have been due to the low concentration of Y-27632 used: in agreement I found only a minimal effect on neurite length when 10 p M Y -27632 was used, but observed a robust effect at 25-50 pM. Regardless, it appears that neurite elongation can be increased with either smaller or larger growth cone morphologies. This may be due to differences in cell type and growth substrate and the activity of other actin regulating molecules. For instance, constitutively active Racl promoted the growth of spinal cord motor neurites on myelin substrates (Kuhn et al, 1999). As well, constitutively active Cdc42 has been shown to promote neurite growth while stimulating the formation of filopodia and lamellipodia (Brown et al, 2000). It is possible that optimal growth on myelin and other inhibitors could be achieved by experimentally activating Racl and Cdc42 and simultaneously inhibiting the Rho-pathway. The differences in actin distribution in growth cones exposed to laminin versus aggrecan reported here strengthen this concept. By inhibiting R O C K with Y-27632, growth cones growing on aggrecan substrates adopt two characteristics of growth cones growing on laminin - long filopodia and fewer central actin bundles. It remains to be shown whether the activation of Rac and/or Cdc42 concomitant with Rho/ROCK inhibition results in the formation of more filopodia and larger lamellipodia when grown on aggrecan, as well as a further stimulation of neurite outgrowth. Growth cone contact with positive guidance cues promotes the accumulation of F-actin in the peripheral area of the growth cone at the site of contact (Lin and Forscher, 1993; O'Connor and Bentley, 1993). In the central regions, actin is depleted, a process thought necessary to allow the invasion of microtubules, thus enabling consolidation of the axon (reviewed by Tanaka and Sabry, 1995; Suter and Forscher, 1998). An abnormal presence of F-actin bundles in the center of the growth cone may hamper this invasion of microtubules. This may contribute to the mechanism of growth cone inhibition by aggrecan that correlated 97 with our observation of central actin bundles. A redistribution of actin from the periphery to the center of the growth cone has also been demonstrated for inhibition due to collapsin-1 and myelin (Fan et al, 1993; Kuhn et al, 1999). Perhaps the activation of Rho by inhibitory molecules promotes the formation of actin bundles in the center of growth cones, as is the case for Rho-induced formation of stress fibres in fibroblasts (Nobes and Hall, 1995). Inhibition of the Rho pathway may reduce the formation of central actin bundles and thereby promote axonal growth. The majority of the experiments reported here were performed on NGF-induced D R G neurite growth. In addition, I demonstrated that aggrecan inhibition of BDNF- and NT-3-induced outgrowth also responded to Y-27632 treatment with increased growth. This, along with the documented results of C3 treatment on retinal ganglion cells, DRG, and cerebellar cells mentioned above, demonstrates that several different neuronal populations may respond positively to Rho pathway interference. Thus, cell permeable molecules, such as Y-27632, that inhibit targets in the Rho pathway may be a viable therapeutic approach to spinal cord and other CNS injuries. C H A P T E R 4: G E N E R A L DISCUSSION 99 The results presented in this thesis demonstrate in vitro that the intrinsic properties of neurons are a major determinant in the success or failure of axonal regeneration. The data presented in Chapter 2 validate in brainstem-spinal neurons the well-characterized notion that the neuronal propensity for axonal regeneration declines with age. As well, I have shown that diversity exists within sub-populations of brainstem-spinal neurons with respect to growth propensity - raphe-spinal neurons are inherently more capable of neurite growth at later stages of development compared to other brainstem-spinal neuron populations. In Chapter 3, I examined neurite growth under the unfavorable conditions provided by inhibitory molecules commonly found in the environment of the injured CNS. Most importantly with regards to the intrinsic state of neurons, the data in Chapter 3 demonstrate that with pharmacological treatment to affected neurons, the effect of multiple inhibitory molecules on the behavior of neurons can be modulated to promote robust growth from normally inhibited axons. Thus, the intrinsic growth state of neurons can be modulated and enhanced. This thesis demonstrates in two studies that intrinsic neuronal properties are an important factor in determining the success or failure of axonal regeneration, and that altering the intrinsic state of a neuron can overcome growth inhibition due to extrinsic inhibitory molecules. NEURITE GROWTH FROM BRAINSTEM-SPINAL NEURONS Comparison of Brainstem-Spinal Neurite Growth With In Vivo Models The experiments reported in Chapter 2 were performed in vitro, inevitably leading them to comparisons with more clinically relevant, established in vivo models. It has been shown in vivo that regeneration of brainstem-spinal neurons decreases with age. After E12 in embryonic chick, regeneration of spinal pathways was not observed (Shimizu et al, 1990; Hasan et al, 1993). This finding has helped establish the notion of a critical period of repair, after which regeneration fails (Steeves et al, 1994). Experiments performed in the Steeves' Laboratory have shown that the critical period for repair can be extended by the immunological suppression of the onset of myelination (Keirstead et al, 1992). Embryonic chicks that have undergone myelination suppression and are injured at E l 5 are capable of both anatomical and functional recovery, clearly displayed by their ability to walk upon 100 hatching. This data implicates the environment as the major determinant of regeneration failure. In contrast, the dysmyelination of the hatchling chick spinal cord also results in some anatomical regeneration; however, walking ability is not restored (Keirstead et al, 1995). This data may be an indication of an intrinsic neuronal decline in the propensity for axonal regrowth. Another explanation is that further maturational changes in myelin and other glial components (i.e. astrocytes) of the spinal cord parenchyma that are present in the hatchling chick but are absent at E l5 contribute to this decrease in regeneration success. The confusion in the analysis of experimental findings from in vivo models support the role of the in vitro models described in this thesis for studying the mechanisms behind axonal growth inhibition. In the experiments described in Chapter 2, brainstem-spinal neurons of different ages (E8 or El7) were grown under the same, ostensibly permissive conditions. Mature neurons from most populations of brainstem-spinal neurons did not extend neurites, while raphe-spinal neurons extended neurites to a far lower extent than younger neurons. These experiments are performed without the influence of extrinsic inhibitory molecules on neurite growth. In a similar manner, as described in the Introduction, retinal, cerebellar, and hippocampal neurons (Chen et al, 1995; L i et al, 1995; Dusart et al, 1997; Prang et al, 2001; Goldberg et al, 2002) show an intrinsic age-dependent neuronal inability to regenerate regardless of the permissiveness of the environment. Thus, the data indicate that an intrinsic change in the neuron, rather than the inhibitory environment of the mature CNS, is the major determinant in neurite growth ability in these models. Why are Mature Neurons Less Capable of Axonal Growth? The well-characterized finding that older neurons are less capable of axonal extension than younger neurons begs the question: why? In essence, this question addresses possible reasons for the inability of mature neurons to regenerate. Answers to this question may be speculated upon in two ways. First, there probably exist some functional reasons that favor less growth ability at more mature ages - i.e. stability is favored over plasticity. Second, there must exist some molecular reasons that cause a decline in axonal growth capability, most likely because of differences in gene expression. Functional reasons for inferior growth from mature neurons are purely speculative. However, it seems plausible that given the metabolic investment in neural development to 101 create the complexities of the mature brain, a mechanism to actively inhibit aberrant connections is desirable. This active inhibition is commonly thought of as an effect of myelination (Keirstead and Steeves, 1998). Myelination of spinal pathways coincides with the onset of regeneration failure in several species (Steeves et al, 1994; Varga et al, 1995). As well, interfering with myelin composition often promotes sprouting from different neuronal populations (reviewed by Fouad et al, 2001). More specifically regarding neuronal plasticity, there is evidence that the presence of myelin protein in the mature CNS inhibits aberrant sprouting of uninjured axons. For instance, the application of neutralizing antibodies to the inhibitory myelin No go-A protein induces sprouting from uninjured Purkinje cells (Buffo et al, 2000) and corticospinal neurons (Thallmair et al, 1998). It is also possible that non-neuronal factors other than myelin may play an inhibitory role that no longer facilitates axon growth in the mature CNS, but that now favors axonal conduction functions. Such factors may be found in changes in the target cells of afferent axons and changes in maturing astrocytes. These changes would make sense from a functional view in that the task of the mature CNS is one of action potentials and synaptic transmission, thus needing specialized molecular mechanisms in post-synaptic cells and the surrounding environment. The above functional reasons do not address evolutionary pressures on the development of higher vertebrates. Chances are that, due to the lengthy time needed to effect CNS regeneration under the best circumstances (fish or mammalian PNS regeneration), any higher vertebrate sustaining a spinal cord injury would surely be at the whim of predators before repair could take place. In fact, not until the development of antibiotics were humans living an extended period following spinal injury. Thus, with death the likely outcome of spinal cord injury, an effective CNS repair strategy was not needed. Molecular reasons for a decline in the neuronal ability for axonal growth are perhaps more easily understood, and most likely mediated by changes in gene expression as neurons age. Developmental gene regulation may be switched on by axonal contact with post-mitotic cells, or by cell body contact with interneurons, as is the case for RGCs (Goldberg et al, 2002). It was reported that RGC neurons received a developmental cue from contact with retinal amacrine cells, switching RGC process growth from axonal to dendritic (Goldberg et al, 2002). The cue resulted in the failure of axonal regeneration from mature RGC cells 102 regardless of permissive growth environments or trophic stimuli, a strong indication that the intrinsic axonal growth state of RGC neurons had been changed to another functional state. Undoubtedly, this intrinsic change resulted in changes to a battery of gene expression profiles. One class of gene changes that occur with age is the expression of receptors for inhibitory molecules, thus rendering more mature neurons susceptible to the inhibitory environment endogenous to the mature CNS. For instance, D R G from adult rats are inhibited in vitro by myelin-associated glycoprotein; however, D R G neurons from newborn rats are not (Mukhopadhyay et al, 1994). An explanation for this effect may be found in similar data regarding myelin inhibition due to Nogo protein, as M A G has now been identified as an inhibitory ligand for the Nogo receptor (Liu et al, 2002). The receptor for Nogo is upregulated in chick after E l 2, coinciding with the phenomenon of myelin inhibition in D R G neurons (Fournier et al, 2001). In a confirmatory gain-of-function experiment, the expression of the Nogo receptor in E7 D R G or retinal cells induced myelin inhibition in these previously unresponsive neurons (Fournier et al, 2001). The levels of cAMP have also been implicated in developmental neuronal changes (Cai et al, 2001). Young neurons have high endogenous levels of cAMP and readily grow on M A G . In contrast, mature neurons have low levels of cAMP and are inhibited by M A G (Cai et al, 2001). Thus, developmental changes in the expression of receptors to inhibitory growth molecules or intracellular signaling molecules may contribute to the intrinsic differences in the properties of embryonic vs. adult neurons, and this may contribute to regeneration failure in the adult CNS. In complement to the upregulated expression of receptors to inhibitory molecules, the down-regulation of appropriate receptors to permissive factors in the adult CNS may also be important. Condic and colleagues have shown that embryonic neurons adapt to inhibitory laminin/aggrecan growth substrates by increasing mRNA and protein for integrin receptors (Condic et al, 1999). This endogenous effect may serve to convey an increased adhesiveness and/or positive signaling cascade that improve the growth capacity of embryonic neurons growing in environments with low levels of permissive molecules. In contrast, adult neurons are unable to adapt their integrin expression, rendering them inhibited and unable to grow (Condic, 2001). Interestingly, the transfection of a single a-integrin gene into adult neurons restores their growth potential on aggrecan to that of young neurons (Condic, 2001). Another 103 example is found in an experiment in which adult and embryonic retinal neurons were grown in vitro on permissive laminin substrates (Bates and Meyer, 1997). A blocking antibody to a (31-integrin receptor inhibited growth of embryonic neurons, but had no effect on adult neurons. Interestingly, both adult and embryonic neurons express (31-integrin receptors, suggesting that adult neurons have lost p i signaling function, possibly due to the down-regulation of its co-receptor (Bates and Meyer, 1997). The data also suggest that adult neurons express a laminin growth-promoting receptor not functionally present in embryonic neurons (Bates and Meyer, 1997). Regarding the experiments described in Chapter 2, it is quite possible that E l 7 brainstem-spinal neurons have down-regulated the appropriate integrin receptors for laminin, or have lost the functional ability to respond favorably to laminin, thus contributing to the lack of neurite growth observed. It is also possible that the superior growth displayed by raphe-spinal neurons is due to their continued expression of the appropriate laminin receptors. The lack of appropriate receptors for growth-promoting molecules, and/or the expression of receptors for growth-inhibiting molecules may also help explain the intrinsic differences in axonal growth propensity seen between populations of young and mature neurons. Thus, the composition of the axonal growth environment vis-a-vis the expression of particular growth-promoting and growth-inhibiting molecules may be the major factor in determining which distinct (with regards to receptor expression profile) populations of neurons (if any) are capable of regeneration. Of course, other molecular changes intrinsic to neurons occur as synapses are formed and the neural developmental focus shifts from axonal elongation and pathfinding to axonal conduction. Regardless, it appears that the inhibitory effects of the mature CNS environment may be mediated by the regulation of specific receptors to inhibitory molecules, thus implicating intrinsic neuronal factors as important to regeneration failure. Fortunately, as data described above and presented in this thesis show, the intrinsic capacity for neuronal growth can be therapeutically enhanced, perhaps to the point where successful in vivo regeneration is possible. Why are Raphe-Spinal Neurons More Plastic? As well as a general age-dependent decline in axonal growth, there exist differences in growth propensity amongst different populations of neurons that are poorly understood. Why 104 raphe-spinal neurons in particular are more plastic than other spinally projecting populations is unknown, although the functional characteristics of serotonergic systems may offer some insight. At this point it may be useful to look at the entire brain serotonergic system in order to gain some elucidation into plasticity. A likely candidate for a molecular mechanism of increased plasticity in raphe neurons is the regeneration-associated gene, GAP-43. High levels of GAP-43 mRNA is found in several regions of adult brain associated with increased plasticity, including the substantia nigra, hippocampal neurons, cerebellar granule cells, and raphe neurons (Bendotti et al, 1991; Kruger et al, 1993; McNamara and Lenox, 1997). As well, in descending systems, GAP-43 was detected in serotonergic raphe-spinal neurons, but not in rubrospinal neurons (Zou et al., 1996). Recently, the expression of both GAP-43 and CAP-23 was shown to promote regeneration of D R G axons both in vitro and in vivo, although GAP-43 alone was ineffective (Bomze et al, 2001). The expression of specific regeneration-associated genes may account for the increased growth capacity from raphe-spinal neurons observed here. Serotonergic neurons have diverse functions, including the.modulation of spinal reflexes (Schmidt and Jordan, 2000) and serotonin has a postulated role as a neurotrophic factor (Azmitia, 1999). Accordingly, there are diverse morphological characteristics evident in raphe-spinal neurons. Serotonergic axons range from small to large diameter, and both myelinated and unmyelinated axons exist (Westlund et al, 1992). The degree of myelination of a particular region in the CNS has been postulated to inversely relate to the quantity of axonal spouting in that region (Kapfhammer and Schwab, 1994; Keirstead and Steeves, 1998). For instance, examples of sprouting have been reported in lightly myelinated regions of the CNS such as the olfactory system (Devor, 1976), the hippocampus (Cotman et al, 1981), and the molecular layer of the cerebellum (Rossi et al, 1991). It is possible that the axonal growth from raphe-spinal neurons reported here is from unmyelinated axons characteristic of most raphe-spinal neurons (Westlund et al, 1992). Contrast that with the highly myelinated supraspinal neurons in the ventral funiculus which subserve more direct control of locomotor functions (Anderson et al, 2000), and which have a poor capacity for growth as observed in Chapter 2. This is a tenuous reason however, as the experimental evidence indicates higher amounts of sprouting into regions of sparse myelination (Kapfhammer and Schwab, 1994), not necessarily from unmyelinated axons. At any rate, the 105 degree of myelination is an interesting and compelling theory regarding the axonal growth potential of CNS neurons. Another functional characteristic of serotonergic systems that may explain their increased propensity for axonal sprouting is their role in the development. There are several roles described for serotonin in the developing CNS (reviewed by Azmitia, 1999; Schmidt and Jordan, 2000), including as a trophic factor (Benton et al, 1997) and differentiating factor for both neurons (Mazer et al, 1997; Yan et al, 1997) and non-neuronal cells (Whitaker-Azmitia et al, 1990). Descending raphe-spinal fibres reach the spinal cord fairly early in development in the rat (by E l 5 they reach the lumbar cord; Schmidt and Jordan, 2000). However, the mature pattern of serotonergic innervation is not fully established in the lumbar cord until arguably post-natal day 21 (Rajaofetra et al, 1989) or day 60 (Newton et al, 1989). This lengthy time frame for the establishment of the mature raphe-spinal innervation pattern suggests that an increased window of plasticity is inherent in the raphe-spinal system, perhaps explaining their greater growth observed in this thesis. Descending serotonin systems have also been described as a developmental modulator of spontaneous rhythmic neuronal networks in the spinal cord (Branchereau et al, 2002). This study showed that removal of serotonin input (i.e. the medulla) to the spinal cord in organotypic culture induced the expression and sprouting of serotonergic neurons in the spinal cord and resulted in the early maturation of inhibitory systems in the rhythmic neuronal networks (Branchereau et al, 2002). Interestingly, the results indicate that an autonomous feedback mechanism may exist whereby serotonin denervation may induce serotonergic differentiation and sprouting. This may be a possible mechanism explaining raphe-spinal neurons increased propensity for axonal growth following axotomy. It has been shown that descending serotonergic systems play a role of modulating spinal reflexes, even to the point in some cases, of initiating locomotion in spinalized animals (reviewed by Schmidt and Jordan, 2000). Because of their function and possible ability to regenerate to a greater extent than other supraspinal systems, raphe-spinal neurons may be useful targets in therapeutic strategies to promote spinal cord regeneration. In one rat model of a lateral spinal cord hemisection, sertonergic fibres sprouted from the contralateral side of the cord (Saruhashi et al, 1996). This sprouting correlated with the onset of some degree of functional recovery that is normally seen in this hemisection model, perhaps mediated 106 through a serotonin-induced facilitation of motor output. In an effort to induce functional recovery after a full spinal cord transection in rats, Privat and colleagues transplanted embryonic raphe cells below the level of the lesion, resulting in hindlimb weight support and triggering of locomotor activity on a treadmill (Ribotta et al, 2000). Unfortunately, a contrasting result may exist in paraplegic humans. Barbeau and colleagues have shown that 5-HT receptor antagonists facilitate some locomotor recovery from chronic injury (reviewed by Barbeau and Rossignol, 1994). They propose that improved locomotion is a result of an antagonistic serotonin-induced decrease in the chronic spasticity that interferes with stepping. Spasticity could be a result of increased serotonergic sprouting following injury, an unfortunate possible maladaptation of raphe-spinal plasticity. The factors influencing the regenerative capacity of severed CNS axons are obviously undoubtedly complex. Nevertheless, it seems that an increased understanding of the intrinsic axonal growth potentials of raphe-spinal neurons and other neuronal populations would contribute to the development of effective SCI therapies. PROMOTION OF AXONAL GROWTH BY TREATMENT TO THE NEURON Enhancement of the Intrinsic Axonal Growth Capacity of Neurons An attractive method for inducing axonal regeneration in the adult CNS is to enhance the intrinsic neuronal capacity for regeneration. Several methods have been attempted. For example, the growth of axons from the lesioned central branch of D R G can be increased i f a peripheral lesion is performed concomitantly (Richardson and Issa, 1984). As well, a similar conditioning lesion can promote the regeneration of dorsal column axons past a spinal cord lesion and into the distal cord, although the distance past the lesion was minimal (Neumann and Woolf, 1999). The conditioning lesion paradigm presumably causes a battery of genes to be regulated that is necessary for axonal regeneration, genes that are not normally activated by a central injury only (for review, see Plunet et al, 2002). Interestingly, as described below, the effects of conditioning lesions are mimicked by cAMP modulation (Neumann et al, 2002b; Qiu et al, 2002). A similar mechanism may be behind the promotion of retinal ganglion axon regeneration via manipulations to the affected eye. Either peripheral nerve 107 implants into the eye (Berry et al, 1996) or mechanical injury to the lens (Leon et al, 2000; Fischer et al, 2001) promoted significant axonal growth in the lesioned optic nerve. Another approach is to directly express specific genes. Skene and colleagues have shown that the expression of two growth cone proteins, GAP-43 and CAP-23, promoted regeneration of D R G axons both in vitro and in vivo (Bomze et al, 2001). The application of neurotrophins to the cell bodies of injured axons is also effective in enhancing the intrinsic growth state of neurons (Plunet et al, 2002). BDNF and NT-4 have been shown to prevent atrophy of and promote regeneration into a peripheral nerve of lesioned rubrospinal neurons (Kobayashi et al, 1997). This treatment also promoted the expression of regeneration associated genes such as GAP-43 and T a l tubulin (Kobayashi et al, 1997). Thus, it is feasible to effectively promote axonal regeneration by inducing the expression of regeneration-associated genes through various methods. The approach undertaken in this thesis was to enhance the intrinsic axonal growth capacity of neurons by altering the activity of the Rho-signaling pathway. By suppressing the activity of ROCK, neurite extension was promoted in the inhibitory growth environments provided by aggrecan, myelin, and spinal cord cryo-sections. Presumably, this method is an example of enhancement of intrinsic growth capacity through manipulations to growth cone proteins rather than gene regulation. In these growth enhancements, the intracellular signaling pathways from extrinsic inhibitory molecules to the actin growth machinery are hijacked. The result is either promotion of growth or indifference to inhibition, in which case growth due to positive cues (i.e. laminin) is now possible. Modulating Cyclic Nucleotide Levels to Enhance Regeneration Capacity It is clear that modulating the activity of Rho-GTPases is an effective method of enhancing the intrinsic axonal growth capacity of neurons. An alternative approach shown effective in several labs to promote axonal growth on inhibitory substrates is the use of cell permeable cyclic nucleotide analogs. A finding that sparked renewed interest in the effects of cyclic nucleotides on axonal growth came from Poo and colleagues (reviewed by Song and Poo, 1999). They showed that growth cone repulsion induced by M A G and Sema-3 A can be converted to attraction by cAMP or cGMP agonists, respectively (Song et al, 1997; Song et al, 1998). Similarly, a cAMP antagonist converted netrin-1 affects on growth cones from 108 attraction to repulsion (Ming et al, 1997). Filbin and colleagues have extended these findings to show that the effects of neurotrophin priming to promote neurite growth on inhibitory myelin substrates are mediated by a cAMP-dependent mechanism (Cai et al, 1999). Two very recent papers have used cAMP agonists to enhance the intrinsic growth capacity of injured D R G neurons in vivo: injection of db-cAMP into D R G promoted sprouting of lesioned dorsal column axons into the injury site (Neumann et al, 2002b; Qiu et al, 2002). As summarized by Song and Poo (1999), these results indicate that the response of a growth cone to an external guidance molecule may depend on the internal state of the neuron. Thus, the intrinsic condition of the neuron regarding intracellular signaling mechanisms can be modulated to promote significant axonal growth in an inhibitory environment. The promotion of axonal growth by cAMP may have underlying significance to Rho-GTPase signaling. Protein kinase A (PKA) is activated by cAMP and has been shown to phosphorylate RhoA (Lang et al, 1996). This in turn prevents RhoA from binding to ROCK, thus inactivating Rho signaling (Dong et al, 1998). As well, P K A phosphorylation promotes Rho attachment to RhoGDI (guanosine nucleotide dissociation inhibitor), causing the sequestration of Rho-GTP to the cytoplasm and away from its putative membrane associated effectors, also inhibiting Rho activity (Lang et al, 1996; Forget et al, 2002). P K A also affects other events downstream to Rho such as neurite retraction induced by lysophosphatidic acid (Tigyi et al, 1996), myosin light chain activity (Kreisberg et al, 1997) , and A D F dephosphorylation (Meberg et al, 1998). Filbin and colleagues hypothesize two separate mechanisms of cAMP promotion of regeneration - an early phase requiring sustained high levels of cAMP and a later phase in which a transient increase in cAMP levels induces long acting results (Qiu et al, 2002). The later phase may be a result of P K A -mediated gene transcription (i.e. the cell body response), while the early phase is consistent with immediate local growth cone responses to cAMP as shown previously (Song et al, 1998) . The effects of R O C K inhibition reported here are consistent with the early phase mechanism behind cAMP promotion of regeneration. Thus, it is possible that both cAMP agonists and Rho or R O C K inhibition act on the same intracellular signaling pathway, albeit at different levels of the signaling cascade. It is also possible that Rho-signaling manipulation will only result in profuse sprouting into CNS lesions as is seen with cAMP application that 109 possibly only activates the early phase mechanism in vivo (Neumann et al, 2002b). A more robust response in which axons pass through the lesion and grow into the distal cord may require additional treatments that further enhance the intrinsic neuronal response to injury. Model Limitations, Remedies, and Future Studies The effects of the Rho-pathway on cells are widespread, including cell division, cellular (i.e. muscle) contractility, transformation, cell migration, process outgrowth, and synapse formation (reviewed by Hall and Nobes, 2000; Luo, 2000). Thus, the application of agents that modulate activity in the Rho pathway could have various consequences in animal models and human trials. These concerns are, of course, applicable to any method that interferes with intracellular signaling pathways, especially in one such as the use of cAMP (described above). Fortunately, Y-27632 is well studied in vivo with systemic applications in hypertension (Uehata et al., 1997) and erectile dysfunction models (Chitaley et al, 2001). Y -27632 can be administered orally to rats at 30 mg/kg for 10 days without any reported toxicity (Narumiya et al, 2000). As well, an older, less specific inhibitor of ROCK, HA1077, has produced positive results in human trials of delayed cerebral vasospasm without serious side effects (for review, see Ono-Saito et al, 1999). The current goal of our lab is the in vivo application of Y-27632 directly to spinal cord lesions, a method that should mitigate any widespread effects elsewhere. Unfortunately, there are possible consequences of local application in limiting the drug's effectiveness discussed below. The in vitro models used in Chapter 3 expose both the neuronal cell bodies as well as their growth cones to the R O C K inhibitor drugs. Thus, the cellular location of the drugs effects cannot be distinguished, leading to some confusion in the proposed mechanism of action. As well, unknowns such as this could be one reason for possible difficulties in transferring in vitro research to more pertinent in vivo animal models. For instance, the effects of BDNF on the survival and regeneration of rubro-spinal neurons is only seen when BDNF is applied at the cell bodies in the red nucleus (Kobayashi et al, 1997), not when applied to the spinal cord (W. Tetzlaff, unpublished observations). The data reported in this thesis do not completely distinguish the local mode of action of R O C K inhibitors. However, the drug's quick effects (30-90 minutes) on growth cone morphology support the mode of action as one on the growth cone. Presumably, effects at the cell body (gene regulation) 110 would take much longer to be executed. This question may be definitively answered with the use of compartmented cultures that isolate cell bodies from distal axonal growth cones (Kimpinski et al, 1997), or through positive results from ongoing in vivo studies using localized sources of treatment. The finding in this thesis that neurite growth inhibition induced by both myelin and a chondroitin sulfate proteoglycan can be overcome by suppressing the activity of R O C K is significant. It suggests that there exists a convergence in intracellular signaling pathways from multiple inhibitory molecules to the actin growth machinery through the Rho-GTPase pathway. However, one limitation to this work is that only inhibition from aggrecan, myelin, and spinal cord cryosections were studied with the Y-27632 inhibitor. Nonetheless, research from other labs can expand the list of Rho/ROCK-convergent inhibitory molecules to include: Sema-3A (Jin and Strittmatter, 1997), although this is controversial - see (Kuhn et al., 1999); Eph-receptors and ephrins (Wahl et al., 2000); the purified myelin-derived inhibitor M A G (Lehmann et al, 1999); and tumor necrosis factor-a (Neumann et al, 2002a). As well, I have obtained preliminary data on the effects of Y-27632 on D R G neurite growth on a substrate of the inhibitory molecule, tenascin-R. Y-27632 application increased the total neurite outgrowth of D R G explants grown on tenascin compared to controls from 42 ± 7 mm to 66 ± 9 mm (p < 0.05, t-test). Although this increase was much less than that seen for aggrecan, the substrate was not nearly as inhibitory as aggrecan (compare growth from tenascin controls to typical aggrecan controls: 42 ± 7 mm vs. 3.6 ± 0.7 mm). Other inhibitory molecules remain to be studied, including other semaphorims, CSPG family members such as versican, collagen-IV, and other repulsive guidance molecules that await proof of expression at the site of CNS lesions. However, the data described above are a strong indication that the Rho-pathway is a convergence point for inhibitory molecule signaling. The field would benefit from further studies focused on other points in the Rho-signaling pathway or possible synergistic alternative signaling pathways. This would flesh out the mechanism by which inhibitory molecules act to retard growth cone advance, as well as offering new targets for therapeutic intervention. Unfortunately, the tools available to researchers limit these types of studies. Often drugs affecting signaling molecules are cell impermeable (e.g. C3-exotoxin), or non-specific (many of the kinase inhibitors). As a preliminary attempt, I applied ML-7 , a commercially available specific inhibitor of myosin I l l light chain kinase (MLCK). If the effects of R O C K signaling are mediated primarily through myosin light chain activity (see Introduction and Fig. 1-2), then ML-7 treatment to inhibit M L C K should be similar to Y-27632 treatment to inhibit ROCK. Concentrations of 1 uM and 10 p M were minimally effective compared to Y-27632: a non-significant small increase in total outgrowth from D R G grown on aggrecan (p > 0.05, A N O V A ) and a significant increase in mean length (p = 0.02, A N O V A ; but compare lengths from control cultures to increase induced by ML-7 (10 uM) and Y-27632 (50 uM): 301 ± 22 um vs. 361 ± 20 um vs. 611 ± 33 urn). Interestingly, in light of the high drug concentrations needed in vivo to achieve an effective intracellular concentration, ML-7 at concentrations from 50 p M and higher had disastrous effects. Although the explants adhered to the tissue culture dishes, there was a complete absence of either migrating cells or neurites extending from the explants, an effect previously not observed by myself. Compare this to a plateau effect for Y-27632 application - concentrations as high as 250 uM were effective in promoting neurite growth on aggrecan. This result lends favorably to the potential of applying Y-27632 safely in vivo. In addition, the data confirms that R O C K signaling involves more than regulation of myosin light chain activity, perhaps also involving LIM-kinase activation (Maekawa et al, 1999; Ohashi et al, 2000), a topic requiring further study. As suggested in the Introduction, the natural future in vitro step in this research is to study the effectiveness of Y-27632 on axonal outgrowth from brainstem-spinal neurons grown on inhibitory substrates. Research into neurons responsible for motor output from the brain would nicely complement the work already done on sensory neurons from the DRG. I have attempted some preliminary work in this regard. Unfortunately, the Y-27632 induced outgrowth data from experiments with E8 vestibulospinal neurons grown on aggrecan substrates was inconclusive, partly due to inconsistent and sparse outgrowth associated with brainstem explants compared to outgrowth from DRG. In order to gain knowledge on the effects of R O C K inhibition on CNS cells, I also looked at the migration of glial cells emanating from the brainstem-spinal explants using the nuclear label, Bisbenzimide H 33258 (Fig. 4-1). Interestingly, the area covered by migrating cells in Y-27632 treated cultures was significantly greater than in control cultures (780000 ± 142000 um 2 vs. 189000 ± 48800 urn2, p < 0.0005, t-test). This is effect is readily seen as a large halo of cells surrounding the central brainstem explant (Fig. 4-1). The data indicate that Rho-112 Figure 4-1. R O C K inhibition stimulates the migration of glial cells from brainstem explants grown on aggrecan substrates. Bisbenzamide nuclear labeled glial cells emanating from an E8 vestibulospinal explant grown for 2 days in Y-27632 (50 pM) treated (A) or untreated (B) growth factor-free media. The area covered by migrating cells was much larger from treated cells than control cells (C). *, p < 0.0005, t-test. Scale bar = 250 pm. 113 114 R O C K signaling may be an important pathway in the migration of cells in inhibitory environments. It may be possible to exploit this result in cellular bridging treatments for SCI in which, for example, growth-permissive channels seeded with Schwann cells have been implanted into lesioned spinal cords (reviewed by Jones et al, 2001). This treatment promoted the ingrowth of regenerating axons into the channels (Xu et al, 1997). One major drawback to Schwann cells is they do not migrate out of the channel, thus providing little incentive for regenerating axons to enter the distal (inhibitory) spinal cord. Perhaps by treating the Schwann cell channels with a R O C K inhibitor such as Y-27632, cell migration out of the channel may be induced. The effect of promoting Schwann cell migration may be enhancement of the cellular interface between the host spinal cord and implanted channel, as well as inducing axonal growth into the distal cord, perhaps increasing functional recovery seen in these models. SIGNIF ICANCE The studies presented here have yielded several significant results that are of benefit to spinal cord injury research. I have shown in Chapter 2 that raphe-spinal neurons have a greater capacity for neurite regrowth compared to other brainstem-spinal populations. It is also known that the corticospinal tract is infamous for its reluctance to grow in permissive environments that support of the growth of other neuronal populations (Richardson et al., 1984; Xu et al, 1995). These findings suggest two things. First, there exist favorable neuronal populations as attractive targets for therapeutic intervention. By enhancing the growth of the most capable neurons, partial recovery from injury may be a clinical possibility sooner than i f full regeneration is sought. Interesting results have been reported using new rehabilitation techniques in humans with incomplete spinal cord injuries (Harkema, 2001). Second, methods of enhancing the intrinsic capacity for axonal growth may vary depending on neuronal population. Raphe-spinal neurons may require the expression of fewer regeneration-associated genes than corticospinal neurons. Raphe-spinal neurons may normally express genes associated with a higher level of intrinsic plasticity, thus enabling an easier enhancement to full axonal regeneration capability. 115 I have shown in Chapter 3 that the suppression of R O C K activity promoted neurite regrowth on the inhibitory substrates of aggrecan, myelin, and spinal cord. This significant finding shows that it is possible to overcome the effects of multiple inhibitory molecules with one simple pharmacological treatment. To the best of my knowledge, the data are also the first finding of the involvement of the Rho-signaling pathway in axonal growth inhibition due to chondroitin sulfate proteoglycans. As described in the Introduction, CSPGs are an important component of both myelin and the glial scar that forms after injury. A caveat regarding experimental methodology is also revealed in Chapter 3. The growth cones of Y -27632 treated cultures were smaller than control cultures, perhaps appearing 'collapsed'. A similar finding was reported with C3 treated axons grown on myelin and Semaphorin substrates (Jin and Strittmatter, 1997). Thus, it is important to observe axonal extension in models of growth inhibition. After all, the promotion of axonal extension is the ultimate goal in this research thread. It is popular to remark that regeneration is a recapitulation of development. Thus, therapeutic interventions need to induce in neurons the expression of the same genes that were present in development. Unfortunately, the presence of inhibitory molecules both in the myelin endogenous to the mature CNS and in the glial scar that forms after injury renders the mature injured CNS far different from the developmental CNS. As well, permissive guidance molecules may be lacking in the mature CNS. This necessitates additional approaches such as methods to render axons indifferent to inhibitory molecules. Suppression of R O C K activity may be such a method. Snider and colleagues, in an elegant study, have shown that different signaling pathways mediate regenerative versus developmental sensory axon growth (Liu and Snider, 2001; Markus et al, 2002). Thus, treatments that enhance the intrinsic axonal growth capacity of neurons may need to increase the expression of regeneration-associated genes specifically tailored for a regenerative response, rather than recapitulating developmental growth processes. Concluding Remarks In conclusion, I have demonstrated that mature brainstem-spinal neurons have an intrinsic decline in their capacity for neurite regrowth compared to young neurons. Raphe-spinal neurons are unique among supraspinal projection neurons in that they retain some axonal 116 growth capacity later in their development. These results emphasize that it is necessary to focus research on the affected neuronal populations in addition to studying the inhibitory environment present following CNS injury. I also investigated a method directed at the intracellular signaling pathways in growth cones. Pharmacological inhibition of R O C K overcame the effects of several inhibitory factors that limit neurite regrowth. This is a demonstration of the possibility (and desirability) to enhance the intrinsic axonal growth capability of neurons through pharmacological methods. Future experiments will be to apply these methods to in vivo animal models of SCI in the hopes of one day contributing to a clinical treatment for spinal cord injury. Other than the promotion of axonal elongation through the injury scar and distal spinal cord, there are of course additional factors that will need to be overcome in order to achieve successful functional regeneration (Schwab and Bartholdi, 1996; Steeves and Tetzlaff, 1998; Tetzlaff and Steeves, 2000). Regenerated fibres need to follow existing or re-expressed axonal pathfinding cues and establish functional synapses with target cells. The establishment of the correct topographic map is then critical to functional outcome, perhaps achievable through rehabilitation and activity-dependent mechanisms. Other issues then arising are re-myelination of the regenerated axons and reversal of muscle atrophy in affected areas. Thus, the current common focus in the majority of regeneration laboratories to promote axonal elongation may only be the proverbial tip of the iceberg in the overall goal of restoration of motor and sensory function. Increasingly, it is widely acknowledged that an effective treatment for SCI will require a combinatorial approach, addressing several, i f not all, of these issues described above. FINIS 117 REFERENCES Aizawa H. , Wakatsuki S., Ishii A. , Moriyama K. , Sasaki Y . , Ohashi K. , Sekine-Aizawa Y. , Sehara-Fujisawa A. , Mizuno K. , Goshima Y . , Yahara I. (2001). Phosphorylation of cofilin by LIM-kinase is necessary for semaphorin 3 A-induced growth cone collapse. Nat Neurosci 4(4): 367-373. Albertinazzi C , Gilardelli D., Paris S., Longhi R., de Curtis I. (1998). Overexpression of a neural-specific rho family GTPase, cRaclB, selectively induces enhanced neuritogenesis and neurite branching in primary neurons. Journal of Cell Biology 142(3): 815-825. Amano M . , Fukata Y . , Kaibuchi K. (2000). Regulation and functions of Rho-associated kinase. Exp Cell Res 261(1): 44-51. Amano M . , Ito M . , Kimura K. , Fukata Y . , Chihara K. , Nakano T., Matsuura Y . , Kaibuchi K. (1996). Phosphorylation and activation of myosin by Rho-associated kinase (Rho-kinase). JBiol Chem 271(34): 20246-20249. Amano M . , Chihara K. , Nakamura N . , Fukata Y . , Yano T., Shibata M . , Ikebe M . , Kaibuchi K . (1998). Myosin II activation promotes neurite retraction during the action of Rho and Rho-kinase. Genes to Cells 3(3): 177-188. Amar A. P., Levy M . L. (1999). Pathogenesis and pharmacological strategies for mitigating secondary damage in acute spinal cord injury. Neurosurgery 44(5): 1027-1039; discussion 1039-1040. Anderson E. S., Bjartmar C , Hildebrand C. (2000). Myelination of prospective large fibres in chicken ventral funiculus. JNeurocytol 29(10): 755-764. Arber S., Barbayannis F. A. , Hanser H. , Schneider C , Stanyon C. A. , Bernard O., Caroni P. (1998). Regulation of actin dynamics through phosphorylation of cofilin by LIM-kinase. Nature 393(6687): 805-809. Arshavsky Y . I., Orlovsky G. N . , Perret C. (1988). Activity of rubrospinal neurons during locomotion and scratching in the cat. Behav Brain Res 28(1-2): 193-199. Awasaki T., Saito M . , Sone M . , Suzuki E., Sakai R., Ito K. , Hama C. (2000). The Drosophila trio plays an essential role in patterning of axons by regulating their directional extension. Neuron 26(1): 119-131. 118 Azmitia E. C. (1999). Serotonin neurons, neuroplasticity, and homeostasis of neural tissue. Neuropsychopharmacology 21(2 Suppl): 33S-45S. Bandtlow C. E., Schmidt M . F., Hassinger T. D., Schwab M . E., Kater S. B. (1993). Role of intracellular calcium in NI-35-evoked collapse of neuronal growth cones. Science 259(5091): 80-83. Barbeau H. , Rossignol S. (1994). Enhancement of locomotor recovery following spinal cord injury. Curr Opin Neurol 7(6): 517-524. Bateman J., Van Vactor D. (2001). The Trio family of guanine-nucleotide-exchange factors: regulators of axon guidance. J Cell Sci 114(Pt 11): 1973-1980. Bates C. A. , Meyer R. L. (1997). The neurite-promoting effect of laminin is mediated by different mechanisms in embryonic and adult regenerating mouse optic axons in vitro. Developmental Biology 181(1): 91-101. Beattie M . S., Bresnahan J. C , Lopate G. (1990). Metamorphosis alters the response to spinal cord transection in Xenopus laevis frogs. Journal of Neurobiology 21(7): 1108-1122. Becker T., Bernhardt R. R., Reinhard E., Wullimann M . F., Tongiorgi E., Schachner M . (1998). Readiness of zebrafish brain neurons to regenerate a spinal axon correlates with differential expression of specific cell recognition molecules. Journal of Neuroscience 18(15): 5789-5803. Bendotti C., Servadio A. , Samanin R. (1991). Distribution of GAP-43 mRNA in the brain stem of adult rats as evidenced by in situ hybridization: localization within monoaminergic neurons. J Neurosci 11(3): 600-607. Benton J., Huber R., Ruchhoeft M . , Helluy S., Beltz B. (1997). Serotonin depletion by 5,7-dihydroxytryptamine alters deutocerebral development in the lobster, Homarus americanus. J Neurobiol 33(4): 357-373. Bernstein-Goral FL, Bregman B. S. (1993). Spinal cord transplants support the regeneration of axotomized neurons after spinal cord lesions at birth: a quantitative double-labeling study. Experimental Neurology 123(1): 118-132. Berry M . , Carlile J., Hunter A . (1996). Peripheral nerve explants grafted into the vitreous body of the eye promote the regeneration of retinal ganglion cell axons severed in the optic nerve. Journal of Neurocytology 25(2): 147-170. 119 Bishop A. L., Hall A . (2000). Rho GTPases and their effector proteins. Biochem J 348(Pt 2): 241-255. Bito H. , Furuyashiki T., Ishihara H. , Shibasaki Y . , Ohashi K. , Mizuno K. , Maekawa M . , Ishizaki T., Narumiya S. (2000). A critical role for a Rho-associated kinase, pl60ROCK, in determining axon outgrowth in mammalian CNS neurons. Neuron 26(2): 431-441. Bomze H . M . , Bulsara K . R., Iskandar B. J., Caroni P., Skene J. H . (2001). Spinal axon regeneration evoked by replacing two growth cone proteins in adult neurons. Nature Neuroscience 4(1): 38-43. Bonilla I. E., Tanabe K. , Strittmatter S. M . (2002). Small proline-rich repeat protein 1A is expressed by axotomized neurons and promotes axonal outgrowth. J Neurosci 22(4): 1303-1315. Borisoff J. F., Pataky D. M . , McBride C. B., Steeves J. D. (2000). Raphe-spinal neurons display an age-dependent differential capacity for neurite outgrowth compared to other brainstem-spinal populations. Exp Neurol 166(1): 16-28. Bovolenta P., Fernaud-Espinosa I., Mendez-Ofero R., Nieto-Sampedro M . (1997). Neurite outgrowth inhibitor of gliotic brain tissue. Mode of action and cellular localization, studied with specific monoclonal antibodies. European Journal of Neuroscience 9(5): 977-989. Boyle R., Pompeiano O. (1980). Responses of vestibulospinal neurons to sinusoidal rotation of neck. JNeurophysiol 44(4): 633-649. Bracken M . B., Holford T. R. (2002). Neurological and functional status 1 year after acute spinal cord injury: estimates of functional recovery in National Acute Spinal Cord Injury Study II from results modeled in National Acute Spinal Cord Injury Study 111. J Neurosurg 96(3 Suppl): 259-266. Bradbury E. J., McMahon S. B., Ramer M . S. (2000). Keeping in touch: sensory neurone regeneration in the CNS. Trends Pharmacol Sci 21(10): 389-394. 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 A B C promotes functional recovery after spinal cord injury. Nature 416(6881): 636-640. 120 Branchereau P., Chapron J., Meyrand P. (2002). Descending 5-hydroxytryptamine raphe inputs repress the expression of serotonergic neurons and slow the maturation of inhibitory systems in mouse embryonic spinal cord. J Neurosci 22(7): 2598-2606. Bregman B. S. (1998). Regeneration in the spinal cord. Curr Opin Neurobiol 8(6): 800-807. Bregman B. S., Kunkel-Bagden E., McAtee M . , O'Neill A . (1989). Extension of the critical period for developmental plasticity of the corticospinal pathway. Journal of Comparative Neurology 282(3): 355-370. Bregman B. S., Kunkel-Bagden E., Schnell L. , Dai H. N . , Gao D., Schwab M . E. (1995). Recovery from spinal cord injury mediated by antibodies to neurite growth inhibitors [see comments]. Nature 378(6556): 498-501. Brewer G. J. (1997). Isolation and culture of adult rat hippocampal neurons. Journal of Neuroscience Methods 71(2): 143-155. Brewer G. J., Torricelli J. R., Evege E. K. , Price P. J. (1993). Optimized survival of hippocampal neurons in B27-supplemented Neurobasal, a new serum-free medium combination. Journal of Neuroscience Research 35(5): 567-576. Brown M . D., Cornejo B. J., Kuhn T. B., Bamburg J. R. (2000). Cdc42 stimulates neurite outgrowth and formation of growth cone filopodia and lamellipodia. J Neurobiol 43(4): 352-364. Buffo A. , Zagrebelsky M . , Huber A. B., Skerra A. , Schwab M . E., Strata P., Rossi F. (2000). Application of neutralizing antibodies against NI-35/250 myelinTassociated neurite growth inhibitory proteins to the adult rat cerebellum induces sprouting of uninjured purkinje cell axons. J Neurosci 20(6): 2275-2286. Bunt S. M . , Fill-Moebs P. (1984). Selection of pathways by regenerating spinal cord fiber tracts. Brain Res 318(2): 307-311. Cabot J. B., Reiner A. , Bogan N . (1982). Avian bulbospinal pathways: anterograde and retrograde studies of cells of origin, funicular trajectories and laminar terminations. Progress in Brain Research 57(79-108. Cai D., Qiu J., Cao Z., McAtee M . , Bregman B. S., Filbin M . T. (2001). Neuronal cyclic A M P controls the developmental loss in ability of axons to regenerate. J Neurosci 21(13): 4731-4739. 121 Cai D. M . , Shen Y . J., De Bellard M . , Tang S., Filbin M . T. (1999). Prior exposure to neurotrophins blocks inhibition of axonal regeneration by M A G and myelin via a cAMP-dependent mechanism. Neuron 22(1): 89-101. Caroni P., Schwab M . E. (1988). Antibody against myelin-associated inhibitor of neurite growth neutralizes nonpermissive substrate properties of CNS white matter. Neuron 1(1): 85-96. Challacombe J. F., Snow D. M . , Letourneau P. C. (1997). Dynamic microtubule ends are required for growth cone turning to avoid an inhibitory guidance cue. J Neurosci 17(9): 3085-3095. Chen D. F., Jhaveri S., Schneider G. E. (1995). Intrinsic changes in developing retinal neurons result in regenerative failure of their axons. Proceedings of the National Academy of Sciences of the United States of America 92(16): 7287-7291. Chen M . S., Huber A. B., van der Haar M . E., Frank M . , Schnell L. , Spillmann A . A. , Christ F., Schwab M . E. (2000). Nogo-A is a myelin-associated neurite outgrowth inhibitor and an antigen for monoclonal antibody IN-1. Nature 403(6768): 434-439. Cheng H. , Cao Y . H. , Olson L. (1996). Spinal Cord Repair in Adult Paraplegic Rats - Partial Restoration of Hind Limb Function. Science 273(527'4): 510-513. Chitaley K. , Wingard C. J., Clinton Webb R., Branam H. , Stopper V . S., Lewis R. W., Mills T. M . (2001). Antagonism of Rho-kinase stimulates rat penile erection via a nitric oxide-independent pathway. Nat Med 7(1): 119-122. Condic M . L. (2001). Adult neuronal regeneration induced by transgenic integrin expression. J Neurosci 21(13): 4782-4788. Condic M . L. , Snow D. M . , Letourneau P. C. (1999). Embryonic neurons adapt to the inhibitory proteoglycan aggrecan by increasing integrin expression. J Neurosci 19(22): 10036-10043. Cotman C. W., Nieto-Sampedro M . , Harris E. W. (1981). Synapse replacement in the nervous system of adult vertebrates. Physiol Rev 61(3): 684-784. David S., Aguayo A. J. (1981). Axonal elongation into peripheral nervous system "bridges" after central nervous system injury in adult rats. Science 214(4523): 931-933. David S., Ousman S. S. (2002). Recruiting the immune response to promote axon regeneration in the injured spinal cord. Neurosdentist 8(1): 33-41. 122 Davies S. J., Goucher D. R., Doller C , Silver J. (1999). Robust regeneration of adult sensory axons in degenerating white matter of the adult rat spinal cord. J Neurosci 19(14): 5810-5822. Davies S. J., Fitch M . T., Memberg S. P., Hall A . K. , Raisman G., Silver J. (1997). Regeneration of adult axons in white matter tracts of the central nervous system. Nature 390(6661): 680-683. De Winter F., Oudega M . , Lankhorst A . J., Hamers F. P., Blits B., Ruitenberg M . J., Pasterkamp R. J., Gispen W. H. , Verhaagen J. (2002). Injury-induced class 3 semaphorin expression in the rat spinal cord. Exp Neurol 175(1): 61-75. Devor M . (1976). Neuroplasticity in the rearrangement of olfactory tract fibers after neonatal transection in hamsters. J Comp Neurol 166(1): 49-72. Domeniconi M . , Cao Z., Spencer T., Sivasankaran R., Wang K . C., Nikulina E., Kimura N . , Cai H. , Deng K. , Gao Y. , He Z., Filbin M . T. (2002). Myelin-Associated Glycoprotein Interacts with the Nogo66 Receptor to Inhibit Neurite Outgrowth. Neuron 35(283-290. Dong J. M . , Leung T., Manser E., Lim L. (1998). cAMP-induced morphological changes are counteracted by the activated RhoA small GTPase and the Rho kinase ROKalpha. Journal of Biological Chemistry 273(35): 22554-22562. Dou C. L., Levine J. M . (1997). Identification of a neuronal cell surface receptor for a growth inhibitory chondroitin sulfate proteoglycan (NG2). J Neurochem 68(3): 1021-1030. Dow K. E., Ethell D. W., Steeves J. D., Riopelle R. J. (1994). Molecular correlates of spinal cord repair in the embryonic chick: heparan sulfate and chondroitin sulfate proteoglycans. Exp Neurol 128(2): 233-238. Driessens M . H. , Hu H. , Nobes C. D., Self A. , Jordens I., Goodman C. S., Hall A . (2001). Plexin-B semaphorin receptors interact directly with active Rac and regulate the actin cytoskeleton by activating Rho. Curr Biol 11(5): 339-344. Dube L., Parent A . (1981). The monoamine-containing neurons in avian brain: I. A study of the brain stem of the chicken (Gallus domesticus) by means of fluorescence and acetylcholinesterase histochemistry. Journal of Comparative Neurology 196(4): 695-708. Duggan A. W., Griersmith B. T. (1979). Inhibition of the spinal transmission of nociceptive information by supraspinal stimulation in the cat. Pain 6(2): 149-161. 123 Dusart I., Airaksinen M . S., Sotelo C. (1997). Purkinje cell survival and axonal regeneration are age dependent: an in vitro study. Journal of Neuroscience 17(10): 3710-3726. Dyer J. K. , Bourque J. A. , Steeves J. D. (1998). Regeneration of brainstem-spinal axons after lesion and immunological disruption of myelin in adult rat. Experimental Neurology 154(1): 12-22. Eidelberg E. (1981). Consequences of spinal cord lesions upon motor function, with special reference to locomotor activity. Progress in Neurobiology 17(3): 185-202. Fan J., Mansfield S. G., Redmond T., Gordon-Weeks P. R., Raper J. A . (1993). The organization of F-actin and microtubules in growth cones exposed to a brain-derived collapsing factor. J Cell Biol 121(4): 867-878. Fawcett J. W., Geller H . M . (1998). Regeneration in the CNS: optimism mounts. Trends Neurosci 21(5): 179-180. Fawcett J. W., Asher R. A . (1999). The glial scar and central nervous system repair. Brain Res Bull 49(6): 377-391. Fernandes K . J., Tetzlaff W. (2000) Gene Expression in Axotomized Neurons: Identifying the Intrinsic Determinants of Axonal Growth. In: Regeneration in the Central Nervous System (Ingoglia N , Murray M , eds). New York: Marcel Dekker. Fernandes K . J., Fan D. P., Tsui B. J., Cassar S. L., Tetzlaff W. (1999). Influence of the axotomy to cell body distance in rat rubrospinal and spinal motoneurons: differential regulation of GAP-43, tubulins, and neurofilament-M. J Comp Neurol 414(4): 495-510. Filbin M . T. (1995). Myelin-associated glycoprotein: a role in myelination and in the inhibition of axonal regeneration? Current Opinion in Neurobiology 5(5): 588-595. Fischer D., Heiduschka P., Thanos S. (2001). Lens-injury-stimulated axonal regeneration throughout the optic pathway of adult rats. Exp Neurol 172(2): 257-272. Fitch M . T., Silver J. (1997a). Activated macrophages and the blood-brain barrier: inflammation after CNS injury leads to increases in putative inhibitory molecules. Exp Neurol 148(2): 587-603. Fitch M . T., Silver J. (1997b). Glial cell extracellular matrix: boundaries for axon growth in development and regeneration. Cell Tissue Res 290(2): 379-384. 124 Forehand C. J., Farel P. B. (1982). Anatomical and behavioral recovery from the effects of spinal cord transection: dependence on metamorphosis in anuran larvae. Journal of Neuroscience 2(5): 654-652. Forget M . A. , Desrosiers R. R., Gingras D., Beliveau R. (2002). Phosphorylation states of Cdc42 and RhoA regulate their interactions with Rho GDP dissociation inhibitor and their extraction from biological membranes. Biochem J361(Pt 2): 243-254. Fouad K. , Dietz V. , Schwab M . E. (2001). Improving axonal growth and functional recovery after experimental spinal cord injury by neutralizing myelin associated inhibitors. Brain Res Brain Res Rev 36(2-3): 204-212. Fournier A . E., Strittmatter S. M . (2001). Repulsive factors and axon regeneration in the CNS. Curr Opin Neurobiol 11(1): 89-94. Fournier A. E., GrandPre T., Strittmatter S. M . (2001). Identification of a receptor mediating Nogo-66 inhibition of axonal regeneration. Nature 409(6818): 341-346. Fu S. Y . , Gordon T. (1997). The cellular and molecular basis of peripheral nerve regeneration. Mol Neurobiol 14(1-2): 67-116. Gallo G., Letourneau P. C. (2000). Neurotrophins and the dynamic regulation of the neuronal cytoskeleton [Review]. Journal of Neurobiology 44(2): 159-173. Gates M . A. , Fillmore FL, Steindler D. A . (1996). Chondroitin sulfate proteoglycan and tenascin in the wounded adult mouse neostriatum in vitro: dopamine neuron attachment and process outgrowth. J Neurosci 16(24): 8005-8018. Geller H. M . , Fawcett J. W. (2002). Building a bridge: engineering spinal cord repair. Exp Neurol 174(2): 125-136. Glover J. C. (1993). The development of brain stem projections to the spinal cord in the chicken embryo. Brain Research Bulletin 30(3-4): 265-271. Glover J. C , Petursdottir G. (1991). Regional specificity of developing reticulospinal, vestibulospinal, and vestibulo-ocular projections in the chicken embryo. Journal of Neurobiology 22(4): 353-376. Goldberg J. L. , Klassen M . P., Hua Y . , Barres B. A . (2002). Amacrine-signaled loss of intrinsic axon growth ability by retinal ganglion cells. Science 296(5574): 1860-1864. Grandpre T., Strittmatter S. M . (2001). Nogo: a molecular determinant of axonal growth and regeneration. Neurosdentist 7(5): 377-386. 125 GrandPre T., L i S., Strittmatter S. M . (2002). Nogo-66 receptor antagonist peptide promotes axonal regeneration. Nature 417(6888): 547-551. GrandPre T., Nakamura F., Vartanian T., Strittmatter S. M . (2000). Identification of the Nogo inhibitor of axon regeneration as a Reticulon protein. Nature 403(6768): 439-444. Grant P., Tseng Y . (1986). Embryonic and regenerating Xenopus retinal fibers are intrinsically different. Developmental Biology 114(2): 475-491. Grillner S., Wallen P. (1985). Central pattern generators for locomotion, with special reference to vertebrates. Annual Review of Neuroscience 8(233-261. Grillner S., Dubuc R. (1988). Control of locomotion in vertebrates: spinal and supraspinal mechanisms. Advances in Neurology 47(425-453. Halfter W., Reckhaus W., Kroger S. (1987). Nondirected axonal growth on basal lamina from avian embryonic neural retina. Journal of Neuroscience 7(11): 3712-3722. Hall A . (1998). Rho GTPases and the actin cytoskeleton. Science 279(5350): 509-514. Hall A. , Nobes C. D. (2000). Rho GTPases: molecular switches that control the organization and dynamics of the actin cytoskeleton. Philos Trans R Soc Lond B Biol Sci 355(1399): 965-970. Harkema S. J. (2001). Neural plasticity after human spinal cord injury: application of locomotor training to the rehabilitation of walking. Neurosdentist 7(5): 455-468. Hartman B. K. , Agrawal H . C , Kalmbach S., Shearer W. T. (1979). A comparative study of the immunohistochemical localization of basic protein to myelin and oligodendrocytes in rat and chicken brain. Journal of Comparative Neurology 188(2): 273-290. Hasan S. J., Keirstead H. S., Muir G. D., Steeves J. D. (1993). Axonal regeneration contributes to repair of injured brainstem-spinal neurons in embryonic chick. Journal of Neuroscience 13(2): 492-507. Hashimoto R., Nakamura Y . , Kosako H. , Amano M . , Kaibuchi K. , Inagaki M . , Takeda M . (1999). Distribution of Rho-kinase in the bovine brain. Biochem Biophys Res Commun 263(2): 575-579. Heacock A. M . , Agranoff B. W. (1977). Clockwise growth of neurites from retinal explants. Sconce 198(4312): 64-66. Hopkins S. J., Rothwell N . J. (1995). Cytokines and the nervous system. I: Expression and recognition. Trends Neurosci 18(2): 83-88. 126 Homer P. J., Gage F. H . (2000). Regenerating the damaged central nervous system. Nature 407(6807): 963-970. Huang D. W., McKerracher L., Braun P. E., David S. (1999). A therapeutic vaccine approach to stimulate axon regeneration in the adult mammalian spinal cord. Neuron 24(3): 639-647. Huber A. B., Schwab M . E. (2000). Nogo-A, a potent inhibitor of neurite outgrowth and regeneration. Biol Chem 381(5-6): 407-419. Ikeda H. , Goto J. (1971). Distribution of monoamine-containing cells in the central nervous system of the chicken. Japanese Journal of Pharmacology 21(6): 763-784. Isbister C. M . , O'Connor T. P. (1999). Filopodial adhesion does not predict growth cone steering events in vivo. Journal of Neuroscience 19(7): 2589-2600. Ishizaki T., Uehata M . , Tamechika I., Keel J., Nonomura K. , Maekawa M . , Narumiya S. (2000). Pharmacological properties of Y-27632, a specific inhibitor of rho-associated kinases. Mol Pharmacol 57(5): 976-983. Iwamoto G. A. , Ryu H. , Wagman I. H. (1980). Effects of stimulation of the caudal brain stem on late ventral root reflex discharge elicited by high threshold sural nerve afferents. Brain Res 183(1): 193-199. Jalink K. , van Corven E. J., Hengeveld T., Morii N . , Narumiya S., Moolenaar W. H. (1994). Inhibition of lysophosphatidate- and thrombin-induced neurite retraction and neuronal cell rounding by ADP ribosylation of the small GTP-binding protein Rho. Journal of Cell Biology 126(3): 801-810. Janani R., Xiao Z . - C , Lian J., Kottis V. , Essagain C , Braun P. E., McKerracher L., David S. (1998). Monoclonal antibodies against the myelin-derived axon growth inhibitor arretin. Soc. Neurosci. Abstr. 24(2): 1560. Jarratt H. , Hyland B. (1999). Neuronal activity in rat red nucleus during forelimb reach-to-grasp movements. Neuroscience 88(2): 629-642. Jin Z., Strittmatter S. M . (1997). Racl mediates collapsin-1-induced growth cone collapse. J Neurosci 17(16): 6256-6263. Jones L. L., Oudega M . , Bunge M . B., Tuszynski M . H . (2001). Neurotrophic factors, cellular bridges and gene therapy for spinal cord injury. J Physiol 533(Pt 1): 83-89. 127 Kapfhammer J. P., Schwab M . E. (1994). Inverse patterns of myelination and GAP-43 expression in the adult CNS: neurite growth inhibitors as regulators of neuronal plasticity? J Comp Neurol 340(2): 194-206. Keirstead H . S., Steeves J. D. (1998). CNS Myelin: Does a Stabilizing Role in Neurodevelopment Result in Inhibition of Neuronal Repair after Adult Injury? Neuroscientist 4(273-284. Keirstead H . S., Hasan S. J., Muir G. D., Steeves J. D. (1992). Suppression of the onset of myelination extends the permissive period for the functional repair of embryonic spinal cord. Proceedings of the National Academy of Sciences of the United States of America 89(24): 11664-11668. Keirstead H . S., Dyer J. K. , Sholomenko G. N . , McGraw J., Delaney K . R., Steeves J. D. (1995). Axonal regeneration and physiological activity following transection and immunological disruption of myelin within the hatchling chick spinal cord. J Neurosci 15(10): 6963-6974. Kimpinski K. , Campenot R. B. , Mearow K. (1997). Effects of the neurotrophins nerve growth factor, neurotrophin-3, and brain-derived neurotrophic factor (BDNF) on neurite growth from adult sensory neurons in compartmented cultures. J Neurobiol 33(4): 395-410. Kimura K. , Ito M . , Amano M . , Chihara K. , Fukata Y . , Nakafuku M . , Yamamori B. , Feng J., Nakano T., Okawa K. , Iwamatsu A. , Kaibuchi K . (1996). Regulation of myosin phosphatase by Rho and Rho-associated kinase (Rho-kinase). Science 273(5272): 245-248. 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 Talphal-tubulin mRNA expression, and promote axonal regeneration. J Neurosci 17(24): 9583-9595. Kreisberg J. I., Ghosh-Choudhury N . , Radnik R. A. , Schwartz M . A . (1997). Role of Rho and myosin phosphorylation in actin stress fiber assembly in mesangial cells. American Journal of Physiology 273(2 Pt 2): F283-288. 128 Krekoski C. A. , Neubauer D., Zuo J., Muir D. (2001). Axonal regeneration into acellular nerve grafts is enhanced by degradation of chondroitin sulfate proteoglycan. J Neurosci 21(16): 6206-6213. Kruger L., Bendotti C , Rivolta R., Samanin R. (1993). Distribution of GAP-43 mRNA in the adult rat brain. J Comp Neurol 333(3): 417-434. Kuhn T. B., Brown M . D., Bamburg J. R. (1998). Racl-dependent actin filament organization in growth cones is necessary for betal-integrin-mediated advance but not for growth on poly-D-lysine. J Neurobiol 37(4): 524-540. Kuhn T. B., Brown M . D., Wilcox C. L., Raper J. A. , Bamburg J. R. (1999). Myelin and collapsin-1 induce motor neuron growth cone collapse through different pathways: inhibition of collapse by opposing mutants of racl. J Neurosci 19(6): 1965-1975. Kwon B. K. , Liu J., Messerer C., Kobayashi N . R., McGraw J., Oschipok L., Tetzlaff W. (2002). Survival and regeneration of rubrospinal neurons 1 year after spinal cord injury. Proc Natl Acad Sci U SA 99(5): 3246-3251. Lang P., Gesbert F., Delespine-Carmagnat M . , Stancou R., Pouchelet M . , Bertoglio J. (1996). Protein kinase A phosphorylation of RhoA mediates the morphological and functional effects of cyclic A M P in cytotoxic lymphocytes. EMBO Journal 15(3): 510-519. Lawrence D. G., Kuypers H. G. (1968). The functional organization of the motor system in the monkey. I. The effects of bilateral pyramidal lesions. Brain 91(1): 1-14. Leeuwen F. N . , Kain H . E., Kammen R. A. , Michiels F., Kranenburg O. W., Collard J. G. (1997). The guanine nucleotide exchange factor Tiaml affects neuronal morphology; opposing roles for the small GTPases Rac and Rho. Journal of Cell Biology 139(3): 797-807. Lehmann M . , Fournier A. , Selles-Navarro I., Dergham P., Sebok A. , Leclerc N . , Tigyi G., McKerracher L. (1999). Inactivation of Rho signaling pathway promotes CNS axon regeneration. J Neurosci 19(17): 7537-7547. Lemons M . L. , Howland D. R., Anderson D. K . (1999). Chondroitin sulfate proteoglycan immunoreactivity increases following spinal cord injury and transplantation. Exp Neurol 160(1): 51-65. Lemons M . L., Sandy J. D., Anderson D. K. , Howland D. R. (2001). Intact aggrecan and fragments generated by both aggrecanse and metalloproteinase-like activities are present 129 in the developing and adult rat spinal cord and their relative abundance is altered by injury. J Neurosci 21(13): 4772-4781. Lentz S. I., Knudson C. M . , Korsmeyer S. J., Snider W. D. (1999). Neurotrophins support the development of diverse sensory axon morphologies. J Neurosci 19(3): 1038-1048. Leon S., Y in Y . , Nguyen J., Irwin N . , Benowitz L. I. (2000). Lens injury stimulates axon regeneration in the mature rat optic nerve. Journal of Neuroscience 20(12): 4615-4626. Letourneau P. C. (1996). The Cytoskeleton in Nerve Growth Cone Motility and Axonal Pathfinding. Perspectives on Developmental Neurobiology 4(2-3): 111-123. L i D., Field P. M . , Raisman G. (1995). Failure of axon regeneration in postnatal rat entorhinohippocampal slice coculture is due to maturation of the axon, not that of the pathway or target. European Journal of Neuroscience 7(6): 1164-1171. L i M . , Shibata A. , L i C , Braun P. E., McKerracher L., Roder J., Kater S. B. , David S. (1996). Myelin-associated glycoprotein inhibits neurite/axon growth and causes growth cone collapse. Journal of Neuroscience Research 46(4): 404-414. L i X . , Saint-Cyr-Proulx E., Aktories K. , Lamarche-Vane N . (2002). Racl and Cdc42 but not RhoA or Rho kinase activities are required for neurite outgrowth induced by the Netrin-1 receptor DCC (deleted in colorectal cancer) in N1E-115 neuroblastoma cells. J Biol Chem 111(\7)\ 15207-15214. Lim L., Manser E., Leung T., Hall C. (1996). Regulation of phosphorylation pathways by p21 GTPases. The p21 Ras-related Rho subfamily and its role in phosphorylation signalling pathways. Eur J Biochem 242(2): 171-185. Lin C. H. , Forscher P. (1993). Cytoskeletal remodeling during growth cone-target interactions. J Cell Biol 121(6): 1369-1383. Lin C. H. , Forscher P. (1995). Growth cone advance is inversely proportional to retrograde F-actin flow. Neuron 14(4): 763-771. Lin C. H. , Espreafico E. M . , Mooseker M . S., Forscher P. (1996). Myosin drives retrograde F-actin flow in neuronal growth cones. Neuron 16(4): 769-782. Lin C. H. , Espreafico E. M . , Mooseker M . S., Forscher P. (1997). Myosin drives retrograde F-actin flow in neuronal growth cones. Biological Bulletin 192(1): 183-185. 130 Lindsay R. M . (1996). Role of neurotrophins and trk receptors in the development and maintenance of sensory neurons: an overview. Philos Trans R Soc Lond B Biol Sci 351(1338): 365-373. Liu B. P., Strittmatter S. M . (2001). Semaphorin-mediated axonal guidance via Rho-related G proteins. Curr Opin Cell Biol 13(5): 619-626. Liu B. P., Fournier A. , GrandPre T., Strittmatter S. M . (2002). Myelin-Associated Glycoprotein as a Functional Ligand for the Nogo-66 Receptor. Science 21(21. Liu R. Y. , Snider W. D. (2001). Different signaling pathways mediate regenerative versus developmental sensory axon growth. J Neurosci 21(17): RC164. Lund S., Pompeiano O. (1968). Monosynaptic excitation of alpha motoneurones from supraspinal structures in the cat. Acta Physiol Scand 73(1): 1-21. Luo L. (2000). Rho GTPases in neuronal morphogenesis. Nat Rev Neurosci 1(3): 173-180. Luo L., Jan L. Y . , Jan Y . N . (1997). Rho family GTP-binding proteins in growth cone signalling. Current Opinion in Neurobiology 7(1): 81-86. Luo Y . , Raible D., Raper J. A. (1993). Collapsin: a protein in brain that induces the collapse and paralysis of neuronal growth cones. Cell 75(2): 217-227'. Mackay D. J., Hall A . (1998). Rho GTPases. JBiol Chem 273(33): 20685-20688. Maekawa M . , Ishizaki T., Boku S., Watanabe N., Fujita A. , Iwamatsu A. , Obinata T., Ohashi K. , Mizuno K. , Narumiya S. (1999). Signaling from Rho to the actin cytoskeleton through protein kinases R O C K and LIM-kinase. Science 285(5429): 895-898. Markus A. , Zhong J., Snider W. D. (2002). Raf and Akt Mediate Distinct Aspects of Sensory Axon Growth. Neuron 35(65-76. Matsui T., Amano M . , Yamamoto T., Chihara K. , Nakafuku M . , Ito M . , Nakano T., Okawa K. , Iwamatsu A. , Kaibuchi K. (1996). Rho-associated kinase, a novel serine/threonine kinase, as a putative target for small GTP binding protein Rho. Embo J 15(9): 2208-2216. Mazer C , Muneyyirci J., Taheny K. , Raio N . , Borella A. , Whitaker-Azmitia P. (1997). Serotonin depletion during synaptogenesis leads to decreased synaptic density and learning deficits in the adult rat: a possible model of neurodevelopmental disorders with cognitive deficits. Brain Res 760(1-2): 68-73. 131 McConnell J. A. , Sechrist J. W. (1980). Identification of early neurons in the brainstem and spinal cord: I. An autoradiographic study in the chick. Journal of Comparative Neurology 192(4): 769-783. McGraw J., Hiebert G. W., Steeves J. D. (2001). Modulating astrogliosis after neurotrauma. J Neurosci Res 63(2): 109-115. McKeon R. J., Hoke A. , Silver J. (1995). Injury-induced proteoglycans inhibit the potential for laminin-mediated axon growth on astrocytic scars. Experimental Neurology 136(1): 32-43. McKeon R. J., Schreiber R. C., Rudge J. S., Silver J. (1991). Reduction of neurite outgrowth in a model of glial scarring following CNS injury is correlated with the expression of inhibitory molecules on reactive astrocytes. J Neurosci 11(11): 3398-3411. McKerracher L. (2001). Spinal cord repair: strategies to promote axon regeneration. Neurobiol Dis 8(1): 11-18. McKerracher L., David S., Jackson D. L., Kottis V. , Dunn R. J., Braun P. E. (1994). Identification of myelin-associated glycoprotein as a major myelin-derived inhibitor of neurite growth. Neuron 13(4): 805-811. McNamara R. K. , Lenox R. H. (1997). Comparative distribution of myristoylated alanine-rich C kinase substrate (MARCKS) and Fl/GAP-43 gene expression in the adult rat brain. J Comp Neurol 379(1): 48-71. Meberg P. J., Bamburg J. R. (2000). Increase in neurite outgrowth mediated by overexpression of actin depolymerizing factor. J Neurosci 20(7): 2459-2469. Meberg P. J., Ono S., Minamide L. S., Takahashi M . , Bamburg J. R. (1998). Actin depolymerizing factor and cofilin phosphorylation dynamics: response to signals that regulate neurite extension. Cell Motility & the Cytoskeleton 39(2): 172-190. Merkler D., Metz G. A. , Raineteau O., Dietz V. , Schwab M . E., Fouad K. (2001). Locomotor recovery in spinal cord-injured rats treated with an antibody neutralizing the myelin-associated neurite growth inhibitor Nogo-A. Journal of Neuroscience 21(10): 3665-3673. Meyer R. L., Miotke J. (1990). Rapid initiation of neurite outgrowth onto laminin from explants of adult mouse retina induced by optic nerve crush. Experimental Neurology 107(3): 214-221. 132 Ming G. L., Song H. J., Berninger B., Holt C. E., Tessier-Lavigne M . , Poo M . M . (1997). cAMP-dependent growth cone guidance by netrin-1. Neuron 19(6): 1225-1235. Miranda J. D., White L. A. , Marcillo A. E., Willson C. A. , Jagid J., Whittemore S. R. (1999). Induction of Eph B3 after spinal cord injury. Exp Neurol 156(1): 218-222. Mukhopadhyay G., Doherty P., Walsh F. S., Crocker P. R., Filbin M . T. (1994). A novel role for myelin-associated glycoprotein as an inhibitor of axonal regeneration. Neuron 13(3): 757-767. Narumiya S., Ishizaki T., Uehata M . (2000). Use and properties of ROCK-specific inhibitor Y-27632 [Review]. Regulators And Effectors Of Small Gtpases, Pt D : Rho FamUy 325(273-284. Neumann H. , Schweigreiter R., Yamashita T., Rosenkranz K. , Wekerle H. , Barde Y . A . (2002a). Tumor necrosis factor inhibits neurite outgrowth and branching of hippocampal neurons by a rho-dependent mechanism. J Neurosci 22(3): 854-862. Neumann S., Woolf C. J. (1999). Regeneration of dorsal column fibers into and beyond the lesion site following adult spinal cord injury. Neuron 23(1): 83-91. Neumann S., Bradke F., Tessier-Lavigne M . , Basbaum A. I. (2002b). Regeneration of Sensory Axons within the Injured Spinal Cord Induced by Intraganglionic cAMP Elevation. Neuron 34(6): 885-893. Newton B. W., Burkhart A . B., Hamill R. W. (1989). Immunohistochemical distribution of serotonin in spinal autonomic nuclei: II. Early and late postnatal ontogeny in the rat. J Comp Neurol 279(1): 82-103. Niederost B. P., Zimmermann D. R., Schwab M . E., Bandtlow C. E. (1999). Bovine CNS myelin contains neurite growth-inhibitory activity associated with chondroitin sulfate proteoglycans. J Neurosci 19(20): 8979-8989. Nobes C. D., Hall A . (1995). Rho, rac, and cdc42 GTPases regulate the assembly of multimolecular focal complexes associated with actin stress fibers, lamellipodia, and filopodia. Cell 81(1): 53-62. O'Connor T. P., Bentley D. (1993). Accumulation of actin in subsets of pioneer growth cone filopodia in response to neural and epithelial guidance cues in situ. J Cell Biol 123(4): 935-948. 133 O'Donovan M . , Semagor E., Sholomenko G., Ho S., Antal M . , Yee W. (1992). Development of spinal motor networks in the chick embryo. J Exp Zool 261(3): 261-273. Ohashi K. , Nagata K. , Maekawa M . , Ishizaki T., Narumiya S., Mizuno K . (2000). Rho-associated kinase R O C K activates LIM-kinase 1 by phosphorylation at threonine 508 within the activation loop. JBiol Chem 275(5): 3577-3582. Okado N . , Oppenheim R. W. (1985). The onset and development of descending pathways to the spinal cord in the chick embryo. Journal of Comparative Neurology 232(2): 143-161. Ono-Saito N . , Niki I., Hidaka H. (1999). H-series protein kinase inhibitors and potential clinical applications. Pharmacol Ther 82(2-3): 123-131. Pasterkamp R. J., Verhaagen J. (2001). Emerging roles for semaphorins in neural regeneration. Brain Res Brain Res Rev 35(1): 36-54. Pasterkamp R. J., Giger R. J., Ruitenberg M . J., Holtmaat A . J., De Wit J., De Winter F., Verhaagen J. (1999). Expression of the gene encoding the chemorepellent semaphorin III is induced in the fibroblast component of neural scar tissue formed following injuries of adult but not neonatal CNS. Mol Cell Neurosci 13(2): 143-166. Pataky D. M . , Borisoff J. F., Fernandes K . J., Tetzlaff W., Steeves J. D. (2000). Fibroblast growth factor treatment produces differential effects on survival and neurite outgrowth from identified bulbospinal neurons in vitro. Exp Neurol 163(2): 357-372. Plunet W., Kwon B. K. , Tetzlaff W. (2002). Promoting axonal regeneration in the central nervous system by enhancing the cell body response to axotomy. J Neurosci Res 68(1): 1-6. Poduslo S. E. (1975). The isolation and characterization of a plasma membrane and a myelin fraction derived from oligodendroglia of calf brain. J Neurochem 24(4): 647-654. Prang P., Del Turco D., Kapfhammer J. P. (2001). Regeneration of entorhinal fibers in mouse slice cultures is age dependent and can be stimulated by NT-4, GDNF, and modulators of G-proteins and protein kinase C. Exp Neurol 169(1): 135-147. Qiu J., Cai D., Filbin M . T. (2000). Glial inhibition of nerve regeneration in the mature mammalian CNS. Glia 29(2): 166-174. Qiu J., Cai D., Dai H. , McAtee M . , Hoffman P. N . , Bregman B. S., Filbin M . T. (2002). Spinal Axon Regeneration Induced by Elevation of Cyclic A M P . Neuron 34(6): 895-903. 134 Rajaofetra N . , Sandillon F., Geffard M . , Privat A. (1989). Pre- and post-natal ontogeny of serotonergic projections to the rat spinal cord. J Neurosci Res 22(3): 305-321. Ramer M . S., Priestley J. V. , McMahon S. B. (2000). Functional regeneration of sensory axons into the adult spinal cord. Nature 403(6767): 312-316. 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(2): 239-249. Ramon y Cajal S. (1991) Cajal's degeneration and regeneration of the nervous system . New York :: Oxford University Press. Ramon-Cueto A. , Cordero M . I., Santos-Benito F. F., Avila J. (2000). Functional recovery of paraplegic rats and motor axon regeneration in their spinal cords by olfactory ensheathing glia. Neuron 25(2): 425-435. Ribotta M . G., Provencher J., Feraboli-Lohnherr D., Rossignol S., Privat A. , Orsal D. (2000). Activation of locomotion in adult chronic spinal rats is achieved by transplantation of embryonic raphe cells reinnervating a precise lumbar level. J Neurosci 20(13): 5144-5152. Richardson P. M . , Issa V . M . (1984). Peripheral injury enhances central regeneration of primary sensory neurones. Nature 309(5971): 791-793. Richardson P. M . , McGuinness U . M . , Aguayo A. J. (1980). Axons from CNS neurons regenerate into PNS grafts. Nature 284(5753): 264-265. Richardson P. M . , Issa V. M . , Aguayo A. J. (1984). Regeneration of long spinal axons in the rat. Journal of Neurocytology 13(1): 165-182. Rossi F., Wiklund L. , van der Want J. J., Strata P. (1991). Reinnervation of cerebellar Purkinje cells by climbing fibres surviving a subtotal lesion of the inferior olive in the adult rat. I. Development of new collateral branches and terminal plexuses. J Comp Neurol 308(4): 513-535. Rossi F., Bravin M . , Buffo A. , Fronte M . , Savio T., Strata P. (1997). Intrinsic properties and environmental factors in the regeneration of adult cerebellar axons. Prog Brain Res 114(4): 283-296. 135 Saruhashi Y . , Young W., Perkins R. (1996). The recovery of 5-HT immunoreactivity in lumbosacral spinal cord and locomotor function after thoracic hemisection. Exp Neurol 139(2): 203-213. Schmidt B. J., Jordan L. M . (2000). The role of serotonin in reflex modulation and locomotor rhythm production in the mammalian spinal cord. Brain Res Bull 53(5): 689-710. Schnell L. , Schwab M . E. (1990). Axonal regeneration in the rat spinal cord produced by an antibody against myelin-associated neurite growth inhibitors. Nature 343(6255): 269-272. Schnell L., Schwab M . E. (1993). Sprouting and regeneration of lesioned corticospinal tract fibres in the adult rat spinal cord. European Journal of Neuroscience 5(9): 115 6-1171. Schwab M . E., Bartholdi D. (1996). Degeneration and regeneration of axons in the lesioned spinal cord. Physiological Reviews 76(2): 319-370. Schwab M . E., Kapfhammer J. P., Bandtlow C. E. (1993). Inhibitors of neurite growth. Annual Review of Neuroscience 16(565-595. Schwartz N . B., Domowicz M . , Krueger R. C , Jr., L i H. , Mangoura D. (1996). Brain aggrecan. Perspect Dev Neurobiol 3(4): 291-306. Sebok A. , Nusser N . , Debreceni B., Guo Z., Santos M . F., Szeberenyi J., Tigyi G. (1999). Different roles for RhoA during neurite initiation, elongation, and regeneration in PC 12 cells. Journal of Neurochemistry 73(3): 949-960. Shamah S. M . , Lin M . Z., Goldberg J. L., Estrach S., Sahin M . , Hu L. , Bazalakova M . , Neve R. L., Corfas G., Debant A. , Greenberg M . E. (2001). EphA receptors regulate growth cone dynamics through the novel guanine nucleotide exchange factor ephexin. Cell 105(2): 233-244. Sharma S. C , Jadhao A . G., Rao P. D. (1993). Regeneration of supraspinal projection neurons in the adult goldfish. Brain Res 620(2): 221-228. Shearer M . C , Fawcett J. W. (2001). The astrocyte/meningeal cell interface—a barrier to successful nerve regeneration? Cell Tissue Res 305(2): 267-273. Shiga T., Kunzi R., Oppenheim R. W. (1991). Axonal projections and synaptogenesis by supraspinal descending neurons in the spinal cord of the chick embryo. Journal of Comparative Neurology 305(1): 83-95. 136 Shimizu L, Oppenheim R. W., O'Brien M . , Shneiderman A. (1990). Anatomical and functional recovery following spinal cord transection in the chick embryo. Journal of Neurobiology 21(6): 918-937. Sholomenko G. N . , O'Donovan M . J. (1995). Development and characterization of pathways descending to the spinal cord in the embryonic chick. Journal of Neurophysiology 73(3): 1223-1233. Snider W. D., Wright D. E. (1996). Neurotrophins cause a new sensation. Neuron 16(2): 229-232. Snider W. D., Zhou F.-Q., Zhong J., Markus A. (2002). Signaling the Pathway to Regeneration. Neuron 35(13-16. Snow D. M . , Letourneau P. C. (1992). Neurite outgrowth on a step gradient of chondroitin sulfate proteoglycan (CS-PG). J Neurobiol 23(3): 322-336. Snow D. M . , Lemmon V. , Carrino D. A. , Caplan A. I., Silver J. (1990). Sulfated proteoglycans in astroglial barriers inhibit neurite outgrowth in vitro. Experimental Neurology 109(1): 111-130. Snow D. M . , Atkinson P. B. , Hassinger T. D., Letourneau P. C , Kater S. B. (1994). Chondroitin sulfate proteoglycan elevates cytoplasmic calcium in D R G neurons. Dev Biol 166(1): 87-100. Song FL, Ming G., He Z., Lehmann M . , McKerracher L., Tessier-Lavigne M . , Poo M . (1998). Conversion of neuronal growth cone responses from repulsion to attraction by cyclic nucleotides. Sconce 281(5382): 1515-1518. Song H. J., Poo M . M . (1999). Signal transduction underlying growth cone guidance by diffusible factors. Current Opinion in Neurobiology 9(3): 355-363. Song H. J., Ming G. L., Poo M . M . (1997). cAMP-induced switching in turning direction of nerve growth cones [published erratum appears in Nature 1997 Sep 25;389(6649):412]. Nature 388(6639): 275-279. Spillmann A. A. , Bandtlow C. E., Lottspeich F., Keller F., Schwab M . E. (1998). Identification and characterization of a bovine neurite growth inhibitor (bNI-220). Journal of Biological Chemistry 273(30): 19283-19293. Steeves J. D., Tetzlaff W. (1998). Engines, accelerators, and brakes on functional spinal cord repair. Annals of the New York Academy of Sciences 860(412-424. 137 Steeves J. D., Sholomenko G. N . , Webster D. M . (1987). Stimulation of the pontomedullary reticular formation initiates locomotion in decerebrate birds. Brain Research 401(2): 205-212. Steeves J. D., Keirstead H. S., Ethell D. W., Hasan S. J., Muir G. D., Pataky D. M . , McBride C. B. , Petrausch B., Zwimpfer T. J. (1994). Permissive and restrictive periods for brainstem-spinal regeneration in the chick. Prog Brain Res 103(243-262. Steven R., Kubiseski T. J., Zheng H. , Kulkarni S., Mancillas J., Ruiz Morales A. , Hogue C. W., Pawson T., Culotti J. (1998). UNC-73 activates the Rac GTPase and is required for cell and growth cone migrations in C. elegans. Cell 92(6): 785-795. Stichel C. C., Muller H . W. (1998). The CNS lesion scar: new vistas on an old regeneration barrier. Cell & Tissue Research 294(1): 1-9. Stichel C. C , Niermann H. , D'Urso D., Lausberg F., Hermanns S., Muller H. W. (1999a). Basal membrane-depleted scar in lesioned CNS: characteristics and relationships with regenerating axons. Neuroscience 93(1): 321-333. Stichel C. C , Hermanns S., Luhmann H . J., Lausberg F., Niermann H. , D'Urso D., Servos G., Hartwig H. G., Muller H. W. (1999b). Inhibition of collagen IV deposition promotes regeneration of injured CNS axons. European Journal of Neuroscience 11(2): 632-646. Suter D. M . , Forscher P. (1998). A n emerging link between cytoskeletal dynamics and cell adhesion molecules in growth cone guidance. Curr Opin Neurobiol 8(1): 106-116. Suter D. M . , Forscher P. (2000). Substrate-cytoskeletal coupling as a mechanism for the regulation of growth cone motility and guidance. J Neurobiol 44(2): 97-113. Tanaka E., Sabry J. (1995). Making the connection: cytoskeletal rearrangements during growth cone guidance. Cell 83(2): 171-176. Tang S., Woodhall R. W., Shen Y . J., deBellard M . E., Saffell J. L. , Doherty P., Walsh F. S., Filbin M . T. (1997). Soluble myelin-associated glycoprotein (MAG) found in vivo inhibits axonal regeneration. Molecular & Cellular Neurosciences 9(5-6): 333-346. Tator C. H . (1995). Update on the pathophysiology and pathology of acute spinal cord injury. Brain Pathol 5(4): 407-413. Tetzlaff W., Steeves J. D. (2000) Intrisinc and extrinsic glial determinants of axonal regeneration in the injured spinal cord. In: Degeneration and Regeneration in the Nervous System (Saunders NR, Dziegielewska K M , eds), pp 93-118. Amsterdam: Harwood. 138 Thallmair M . , Metz G. A. , Z'Graggen W. J., Raineteau O., Kartje G. L. , Schwab M . E. (1998). Neurite growth inhibitors restrict plasticity and functional recovery following corticospinal tract lesions. Nat Neurosci 1(2): 124-131. Tigyi G., Fischer D. J., Sebok A , Marshall F., Dyer D. L., Miledi R. (1996). Lysophosphatidic acid-induced neurite retraction in PC12 cells: neurite-protective effects of cyclic A M P signaling. Journal of Neurochemistry 66(2): 549-558. Tolbert D. L., Der T. (1987). Redirected growth of pyramidal tract axons following neonatal pyramidotomy in cats. Journal of Comparative Neurology 260(2): 299-311. Treherne J. M . , Woodward S. K. , Varga,Z. M . , Ritchie J. M . , Nicholls J. G. (1992). Restoration of conduction and growth of axons through injured spinal cord of neonatal opossum in culture. Proceedings of the National Academy of Sciences of the United States of America 89(1): 431-434. Uehata M . , Ishizaki T., Satoh FL, Ono T., Kawahara T., Morishita T., Tamakawa H. , Yamagami K. , Inui J., Maekawa M . , Narumiya S. (1997). Calcium sensitization of smooth muscle mediated by a Rho-associated protein kinase in hypertension. Nature 389(6654): 990-994. Valenzuela J. I., Hasan S. J., Steeves J. D. (1990). Stimulation of the brainstem reticular formation evokes locomotor activity in embryonic chicken (in ovo). Brain Res Dev Brain Res 56(1): 13-18. Varga Z. M . , Bandtlow C. E., Erulkar S. D., Schwab M . E., Nicholls J. G. (1995). The critical period for repair of CNS of neonatal opossum (Monodelphis domestica) in culture: correlation with development of glial cells, myelin and growth-inhibitory molecules. European Journal of Neuroscience 7(10): 2119-2129. Vastrik I., Eickholt B. J., Walsh F. S., Ridley A. , Doherty P. (1999). Sema3A-induced growth-cone collapse is mediated by Racl amino acids 17-32. Curr Biol 9(18): 991-998. Venters H. D., Dantzer R., Kelley K. W. (2000). Tumor necrosis factor-alpha induces neuronal death by silencing survival signals generated by the type I insulin-like growth factor receptor. Ann N Y Acad Sci 917(210-220. Wahl S., Barth H. , Ciossek T., Aktories K. , Mueller B. K . (2000). Ephrin-A5 induces collapse of growth cones by activating Rho and Rho kinase. J Cell Biol 149(2): 263-270. 139 Wang K . C , Koprivica V. , K im J. A. , Sivasankaran R., Guo Y . , Neve R. L. , He Z. (2002). Oligodendrocyte-myelin glycoprotein is a Nogo receptor ligand that inhibits neurite outgrowth. Nature 417(6892): 941-944. Wang S. D., Goldberger M . E., Murray M . (1991). Plasticity of spinal systems after unilateral lumbosacral dorsal rhizotomy in the adult rat. Journal of Comparative Neurology 304(4): 555-568. Wang X . M . , Qin Y . Q., Martin G. F. (1994). Developmental plasticity of reticulospinal and vestibulospinal axons in the North American opossum, Didelphis virginiana. The Journal of Comparative Neurology 349(288-302. Webster D. M . , Rogers L. J., Pettigrew J. D., Steeves J. D. (1990). Origins of descending spinal pathways in prehensile birds: do parrots have a homologue to the corticospinal tract of mammals? Brain, Behavior & Evolution 36(4): 216-226. Westlund K . N . , Lu Y . , Coggeshall R. E., Willis W. D. (1992). Serotonin is found in myelinated axons of the dorsolateral funiculus in monkeys. Neurosci Lett 141(1): 35-38. Whelan P. J. (1996). Control of locomotion in the decerebrate cat. Prog Neurobiol 49(5): 481-515. Whitaker-Azmitia P. M . , Murphy R., Azmitia E. C. (1990). Stimulation of astroglial 5-HT1A receptors releases the serotonergic growth factor, protein S-100, and alters astroglial morphology. Brain Res 528(1): 155-158. Xu X . M . , Martin G. F. (1991). Evidence for new growth and regeneration of cut axons in developmental plasticity of the rubrospinal tract in the North American opossum. Journal of Comparative Neurology 313(1): 103-112. X u X . M . , Guenard V. , Kleitman N . , Aebischer P., Bunge M . B. (1995). A combination of BDNF and NT-3 promotes supraspinal axonal regeneration into Schwann cell grafts in adult rat thoracic spinal cord. Experimental Neurology 134(2): 261-272. X u X . M . , Chen A. , Guenard V. , Kleitman N . , Bunge M . B. (1997). Bridging Schwann cell transplants promote axonal regeneration from both the rostral and caudal stumps of transected adult rat spinal cord. JNeurocytol 26(1): 1-16. X u X . M . , Zhang S. X . , L i H. , Aebischer P., Bunge M . B. (1999). Regrowth of axons into the distal spinal cord through a Schwann-cell-seeded mini-channel implanted into hemisected adult rat spinal cord. Eur J Neurosci 11(5): 1723-1740. 140 Yamashita T., Higuchi FL, Tohyama M . (2002). The p75 receptor transduces the signal from myelin-associated glycoprotein to Rho. J Cell Biol 157(4): 565-570. Yan W., Wilson C . C , Haring J. H . (1997). 5-HTla receptors mediate the neurotrophic effect of serotonin on developing dentate granule cells. Brain Res Dev Brain Res 98(2): 185-190. Ye J. FL, Houle J. D. (1997). Treatment of the chronically injured spinal cord with neurotrophic factors can promote axonal regeneration from supraspinal neurons. Exp Neurol 143(1): 70-81. Y u T. W., Bargmann C. I. (2001). Dynamic regulation of axon guidance. Nat Neurosci 4(Suppl): 1169-1176. Zhang Y . , Anderson P. N . , Campbell G., Mohajeri FL, Schachner M . , Lieberman A. R. (1995). Tenascin-C expression by neurons and glial cells in the rat spinal cord: changes during postnatal development and after dorsal root or sciatic nerve injury. Journal of Neurocytology 24(8): 585-601. Zou X . C , Ho R. H. , Wang X . M . , Martin G. F. (1996). Evidence for GAP-43 within descending spinal axons in the North American opossum, Didelphis virginiana. Brain Behav Evol 47(4): 200-213. Zuo J., Neubauer D., Dyess K. , Ferguson T. A. , Muir D. (1998a). Degradation of chondroitin sulfate proteoglycan enhances the neurite-promoting potential of spinal cord tissue. Exp Neurol 154(2): 654-662. Zuo J., Ferguson T. A. , Hernandez Y . J., Stetler-Stevenson W. G., Muir D. (1998b). Neuronal matrix metalloproteinase-2 degrades and inactivates a neurite-inhibiting chondroitin sulfate proteoglycan. J Neurosci 18(14): 5203-5211. 

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