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UBC Theses and Dissertations

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UBC Theses and Dissertations

Myelinating cells in repair of spinal cord injury Assinck, Peggy Lee 2017

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MYELINATING CELLS IN REPAIR OF SPINAL CORD INJURY by  Peggy Lee Assinck  B.Sc. (Honours), Brock University, 2008  A THESIS SUBMITTED IN PARTIAL FULFILLMENT OF THE REQUIREMENTS FOR THE DEGREE OF  DOCTOR OF PHILOSOPHY in THE FACULTY OF GRADUATE AND POSTDOCTORAL STUDIES (Neuroscience)  THE UNIVERSITY OF BRITISH COLUMBIA (Vancouver)  August 2017  © Peggy Lee Assinck, 2017 ii  Abstract The damage inflicted by spinal cord injury (SCI) occurs in two phases. The primary injury is a mechanical insult to the spinal cord, resulting in permanent loss of cells and tissue structure. Multiple mechanisms of secondary injury extend this damage. One such mechanism is thought to be progressive loss of myelin, such that axons that survive primary injury are rendered dysfunctional by conduction block. As a result, myelin repair has emerged as a major research focus. The goals of this research were: i) to evaluate the extent of spontaneous repair by endogenous glia, and ii) to determine whether transplantation in a clinically relevant scenario can improve the outcome of SCI. In Chapter 2, I characterized the source and extent of spontaneous myelin repair in experimental SCI. I systematically assessed the cellular origin of new myelin and myelinating cells in transgenic mice, by genetically labeling multiple lineages prior to SCI. Contrary to prevailing dogma (that endogenous myelin repair was limited), we found that ~30% of myelinated axons at the injury epicentre were ensheathed de novo (since injury) at three months after SCI. In addition, the majority of myelinating Schwann cells (SCs) in the injured spinal cord were derived from oligodendrocyte precursor cells (OPCs) and infiltration of peripheral myelinating SCs made only a small contribution. In Chapter 3, I investigated the potential for improving spontaneous repair (and the outcome of SCI) through glial transplantation. Skin-derived precursor cells directed to a SC fate (SKP-SCs) were transplanted at the site of chronic SCI. At 21 weeks after transplantation (29 weeks-post SCI), SKP-SCs contained thousands of growing/regenerating axons, which were myelinated by either transplanted or endogenous SCs. The presence of endogenous SCs was increased after SKP-SC transplantation. Rats that received SKP-SCs had higher functional motor scores and displayed less bladder wall thickening (a hallmark of bladder dysfunction following SCI) compared to iii  controls. These data contribute to our understanding of the endogenous glial repair response after SCI, both in the absence of treatment and following a clinically relevant cell transplantation. These endogenous repair mechanisms might be exploited and augmented to develop novel treatments for SCI.    iv  Lay Summary The spinal cord can be considered as a cable of wires connecting the brain to the body, as well as a computer that integrates information from the body and brain. In this analogy, individual nerve fibers represent wires. After spinal cord injury (SCI), the flow of information between brain and body is interrupted, affecting movement, sensation, and bodily functions. Like electrical wires, some nerve fibers are covered in insulation, called myelin, that improves conduction. My research examines the importance of myelin after SCI. I study when and how myelin is lost and repaired after SCI, and whether transplanting cells that form myelin can improve function after SCI. I found that the spinal cord produces two types of new myelin to insulate nerve fibers after injury, and that we can improve this repair process by transplanting cells from skin, which has the potential to be performed in people with SCI.   v  Preface This dissertation is based on data from experiments run primarily in the laboratory of Dr. Wolfram Tetzlaff, but data from national collaborations are included and several transgenic mice lines were obtained on a collaborative basis from international laboratories.   A portion of the text in Chapter 1 and 4 has been published: *P. Assinck, * G.J. Duncan, *B.J. Hilton , *J.R. Plemel, W. Tetzlaff. (2017). Cell transplantation for spinal cord injury. Nature Neuroscience 20 637-647. *Authors contributed equally. I acknowledge that Gregory J Duncan largely wrote, Dr. Brett Hilton largely wrote, Dr. Jason R Plemel largely wrote, and Dr. Wolfram Tetzlaff largely wrote 1.4.1, 4.4.1-3. The figures in Chapter 1 were prepared for this publication in collaboration with the co-authors, but published in a different form after redrawing by the journal’s artist. A version of Chapter 2 has been accepted for publication: *P. Assinck, G.J. Duncan, J.R. Plemel, M.J. Lee, J.S. Stratton, J. Liu, L.M. Ramer, Shin H. Kang, D.E. Bergles, J. Biernaskie, W. Tetzlaff. (2017) Myelinogenic plasticity of oligodendrocyte precursor cells following CNS trauma. Journal of Neuroscience. The in vitro and in vivo portions of Chapter 2 were supervised by Drs. Jeff Biernaskie and Wolfram Tetzlaff, respectively. I planned and organized all aspects of this work and conducted or supervised all aspects of spinal cord injury (SCI) surgery, animal care, necropsy and tissue collection, immunohistochemistry, microscopy, image analysis, statistical analysis and manuscript composition. G.J. Duncan also made substantial contributions to the work, was involved in experimental design, and helped me with spinal cord injury surgical procedures (blinding, preparation, suturing), animal care, necropsy and tissue collection, and manuscript editing. Dr. Plemel contributed substantially to experimental design and manuscript editing. M.J. Lee helped with image analysis. Under Dr. Biernaskie’s supervision, Dr. Stratton vi  was responsible for Figure 2.8. Dr. Liu performed spinal cord injury surgeries. Dr. L.M. Ramer contributed to writing and manuscript edits. Drs. Kang and Bergles contributed to experimental design and made minor edits to the manuscript. Dr. Biernaskie contributed to design of in vitro and in vivo experiments and manuscript edits. Dr. Tetzlaff was involved in all aspects of experimental design, supervised the in vivo portion of the study, and contributed substantially to manuscript composition. The work was conducted with approval of the University of British Columbia Animal Care Committee and the University of Calgary Animal Care Committee. The relevant animal care certificate numbers are A13-0328 (UBC) “Mice Spinal Cord Injury” and A13-0329 (UBC) “Mouse Breeding ICORD Tetzlaff Lab” and AC1200168 (U of Calgary). This work was funded by grants held by Dr. Wolfram Tetzlaff from the Canadian Institutes of Health Research (CIHR). A version of Chapter 3 has been prepared and submitted for publication. *P. Assinck, J. S. Sparling, S. Dworski, G. J. Duncan, D. L. Wu, J. Liu, B.K. Kwon, J. Biernaskie, F. D. Miller, W. Tetzlaff. Transplantation of skin precursor-derived SCs improves outcome after chronic spinal cord injury in rats. The in vitro and in vivo portions of that work were supervised by Drs. Freda Miller and Wolfram Tetzlaff, respectively. I planned and organized all aspects of this work and conducted or supervised all aspects of SCI surgery, cell transplantation, animal care, behavioral testing, necropsy and tissue collection, microscopy, image analysis, and statistical analysis; I also co-wrote the manuscript. Dr. Sparling assisted with experimental design, blinding, and manuscript editing. S. Dworski (under Dr. Freda Miller’s supervision) isolated and expanded SKP-SCs at the University of Toronto, prepared the cells for transplantation in Vancouver, and the data shown in Figure 3.1 b-d and Figure 3.10 a and b. G. J. Duncan helped me conduct the BBB behavioral test, and assisted me animal care and manuscript editing. L.D Wu contributed to vii  image analysis. All surgeries were performed by Dr. Liu. Mr. Y. Jiang contributed to tissue sectioning and image analysis. Dr. Kwon contributed to experimental design and manuscript edits. Dr. Miller supervised in vitro work and contributed to experimental design and manuscript edits. Dr. Tetzlaff worked closely with me throughout the project, supervised the in vivo experiments, contributed to experimental design, and co-wrote the manuscript. The work was conducted with approval of the University of British Columbia Animal Care Committee. The relevant animal care certificate numbers include: A10-0017 “Copy of- Anatomical and functional recovery after spinal cord contusion injury”, and A14-0212 “New Anatomical and functional recovery after spinal cord contusion injury”. This work was funded by grants held by Dr. Wolfram Tetzlaff from the Canadian Stem Cell Network and the Canadian Institutes of Health Research (CIHR).  viii  Table of Contents  Abstract .......................................................................................................................................... ii	Lay Summary ............................................................................................................................... iv	Preface .............................................................................................................................................v	Table of Contents ....................................................................................................................... viii	List of Tables .............................................................................................................................. xiv	List of Figures ...............................................................................................................................xv	List of Abbreviations ................................................................................................................ xvii	Acknowledgements .................................................................................................................. xxiii	Dedication ...................................................................................................................................xxv	Chapter 1: General introduction ..................................................................................................1	1.1	 Introduction overview ..................................................................................................... 1	1.2	 Clinical implications of SCI ............................................................................................ 2	1.2.1	 SCI demographics ................................................................................................... 2	1.2.2	 Priorities of the SCI population .............................................................................. 5	1.3	 Pathology of contusion SCI ............................................................................................ 5	1.3.1	 Primary SCI ............................................................................................................ 6	1.3.2	 Secondary SCI and injury progression ................................................................... 7	 Secondary SCI: vascular effects ......................................................................... 8	 Secondary SCI: immune effects ........................................................................ 10	 Secondary SCI: the glial scar ............................................................................ 12	1.4	 Cell transplantation for repair of SCI ............................................................................ 15	ix  1.4.1	 Historical perspective ............................................................................................ 15	1.4.2	 Candidate cell types .............................................................................................. 16	1.4.3	 Mechanisms of action of transplanted cells .......................................................... 17	 Neuroprotection ................................................................................................ 17	 Molecules secreted by transplanted cells .......................................................... 18	 Immunomodulation ........................................................................................... 19	 Preventing blood vessel loss or improving angiogenesis ................................. 20	 Axonal growth .................................................................................................. 21	 Myelin regeneration .......................................................................................... 23	1.5	 Myelin biology .............................................................................................................. 25	1.5.1	 Oligodendrocyte lineage cells ............................................................................... 27	1.5.2	 Oligodendrocyte lineage cells in response to local CNS injury ........................... 31	1.5.3	 SC development .................................................................................................... 33	1.5.4	 The SC response to PNS injury ............................................................................ 36	1.5.5	 The SC response to CNS injury ............................................................................ 38	1.6	 Transplantation of SCs as a treatment for CNS injury ................................................. 41	1.6.1	 Peripheral nerve grafts .......................................................................................... 43	1.6.2	 Transplantation of SCs after CNS injury .............................................................. 44	1.6.3	 Moving SC transplantation towards the clinic ...................................................... 46	1.6.4	 Alternative sources of SCs .................................................................................... 47	 SKP-SCs ........................................................................................................... 48	1.7	 Overview of experiments and hypothesis ..................................................................... 49	x  Chapter 2: Myelinogenic plasticity of oligodendrocyte precursor cells following spinal cord contusion injury ...........................................................................................................................55	2.1	 Introduction ................................................................................................................... 55	2.2	 Materials and methods .................................................................................................. 57	2.2.1	 Transgenic mice and cre induction ....................................................................... 57	2.2.2	 Surgical procedures ............................................................................................... 58	 SCI .................................................................................................................... 59	 Root and nerve injury ........................................................................................ 60	2.2.3	 Tissue preparation and immunohistochemistry .................................................... 60	2.2.4	 FACS and immunocytochemistry ......................................................................... 61	2.2.5	 Cell counting and analysis .................................................................................... 62	2.3	 Results ........................................................................................................................... 65	2.3.1	 Genetic fate mapping identifies PDGFRa progeny in the adult spinal cord ........ 65	2.3.2	 PDGFRa+ cells, recombined prior to injury, contribute to new oligodendrocyte ensheathment/myelination after SCI ..................................................................................... 68	2.3.3	 The majority of myelinating SCs in the injured spinal cord are derived from PDGFRa+ cells .................................................................................................................... 71	2.3.4	 Olig2+ cells give rise to P0+ SCs in response to contusion SCI. ......................... 73	2.3.5	 Recombined PDGFRa-expressing cells from the PNS do not give rise to myelinating SCs in vitro or in vivo ....................................................................................... 74	2.3.6	 Peripheral myelinating SCs migrate into the spinal cord after injury and contribute to myelination ....................................................................................................................... 76	2.4	 Discussion ..................................................................................................................... 76	xi  2.4.1	 OPC derived ensheathment/myelination is substantial after SCI ......................... 77	2.4.2	 The majority of myelinating SCs in the contused spinal cord are derived from PDGFRa+ CNS progenitors ................................................................................................. 79	2.5	 Conclusion .................................................................................................................... 82	Chapter 3: Transplantation of skin precursor-derived SCs yields better locomotor outcomes and reduced bladder pathology in rats with chronic spinal cord injury .............105	3.1	 Introduction ................................................................................................................. 105	3.2	 Methods....................................................................................................................... 108	3.2.1	 Animals ............................................................................................................... 108	3.2.2	 Spinal cord contusion injury ............................................................................... 108	3.2.3	 SKP isolation and differentiation into SKP-SCs ................................................ 110	3.2.4	 Immunocytochemistry ........................................................................................ 111	3.2.5	 Cell transplantation ............................................................................................. 111	3.2.6	 Behavioural assessments ..................................................................................... 112	 Open field locomotion (BBB) ......................................................................... 113	 CatWalk .......................................................................................................... 113	 Irregular horizontal ladder .............................................................................. 113	3.2.7	 Tract tracing and tissue processing ..................................................................... 114	3.2.8	 Immunohistochemistry ....................................................................................... 115	3.2.9	 Histological quantifications ................................................................................ 115	 Determining lesion volume, average intact tissue, and tissue width .............. 116	 SKP-SC transplant volumes, counts, orientation, and proliferation ............... 117	 GFAP intensity analysis .................................................................................. 118	xii	 SC myelination and non-myelinating SC analysis .......................................... 119	 Axon counts .................................................................................................... 119	 Bladder analysis .............................................................................................. 120	3.2.10	 Statistical analyses .............................................................................................. 121	3.3	 Results ......................................................................................................................... 121	3.3.1	 Skin Derived Precursors (SKPs) generate high purity SCs (SKP-SCs) in vitro ……..................................................................................................................... 121	3.3.2	 Transplanted SKP-SCs survive and bridge the lesion in chronic SCI ................ 122	3.3.3	 SKP-SCs prompted an increase in intact tissue surrounding the site of chronic SCI ………………………......................................................................................... 123	3.3.4	 SKP-SCs integrate into spinal cord tissue, mitigate the formation of the glial scar, and provide a permissive axon growth substrate ................................................................ 123	3.3.5	 SKP-SCs promote growth/regeneration of host axons ....................................... 125	 SKP-SCs myelinate axons in chronic SCI ...................................................... 126	3.3.6	 SKP-SC transplantation augments spontaneous CNS repair by endogenous SCs……. ............................................................................................................................. 127	3.3.7	 SKP-SCs from adult skin show similar behaviour to neonatal-derived SKP-SCs….. ............................................................................................................................. 129	3.3.8	 Transplantation of SKP-SCs eight weeks following SCI resulted in better locomotor outcomes ............................................................................................................ 129	3.3.9	 SKP-SC transplantation reduces bladder pathology observed after SCI ............ 131	3.4	 Discussion ................................................................................................................... 132	Chapter 4: General discussion ..................................................................................................161	xiii  4.1	 Summary of thesis ....................................................................................................... 161	4.2	 Endogenous myelin repair after SCI ........................................................................... 161	4.3	 Exogenous myelin repair after SCI ............................................................................. 165	4.4	 Future and clinical perspectives .................................................................................. 167	4.4.1	 Improved risk assessment ................................................................................... 167	4.4.2	 Improved pre-clinical models ............................................................................. 169	4.4.3	 Assessing efficacy in the chronic stage of injury ................................................ 170	4.4.4	 Defining recovery ............................................................................................... 170	4.4.5	 Aligning outcome measures with the priorities of the SCI community .............. 171	4.4.6	 Managing expectations ....................................................................................... 172	Bibliography ...............................................................................................................................174	 xiv  List of Tables  Table 2-1. Overview of transgenic mouse lines ............................................................................ 83	Table 2-2. Overview of specific animal numbers that underwent qualitative or quantitative analysis .......................................................................................................................................... 84	Table 2-3. Primary antibody table ................................................................................................ 85	Table 3-1. List of primary antibodies used ................................................................................. 138	Table 3-2. List of relevant compared significant correlations .................................................... 139	 xv  List of Figures Figure 1.1 The pathophysiology of rat contusion SCI .................................................................. 51	Figure 1.2 Restoring neuronal connectivity following SCI via cell transplantation ..................... 53	Figure 1.3 Regeneration of oligodendrocyte-derived myelin with cell transplantation ............... 54	Figure 2.1 Genetic labeling of NG2 glia in tamoxifen-inducible PDGFRαCreER uninjured control mice .................................................................................................................................. 86	Figure 2.2 Recombination in central canal-associated cells, pericytes and a subset of PNS endoneurial cells in PDGFRaCreER uninjured control mice ...................................................... 88	Figure 2.3 PDGFRa+ progenitors proliferate and contribute to oligodendrocyte lineage cells in response to SCI ............................................................................................................................. 90	Figure 2.4 Extensive new ensheathment/myelination by oligodendrocytes derived from PDGFRa+ progenitors 12 weeks after SCI .................................................................................. 92	Figure 2.5 PDGFRa+ progenitor-derived SCs express typical hallmarks of SC myelination in PDGFRaCreER:mGFP mice ....................................................................................................... 94	Figure 2.6 The majority of myelinating SCs in the injured spinal cord are derived from PDGFRa+ progenitors .................................................................................................................. 95	Figure 2.7 Olig2+ cells give rise to myelinating SCs after SCI ................................................... 97	Figure 2.8 PDGFRα+ cells from the adult dorsal root ganglion and spinal root of PDGFRα:H2BGFP mice do not exhibit SC fate in vitro .............................................................. 99	Figure 2.9 Recombined PDGFRa+cells in the PNS do not give rise to P0+ cells in response to peripheral injury .......................................................................................................................... 101	Figure 2.10 P0+ SCs give rise to a small number of P0+ SCs after SCI ..................................... 103	xvi  Figure 3.1 Neonatal rat SKP-SCs express typical SC markers in vitro ...................................... 140	Figure 3.2 SKP-SC grafts survive in vivo in the chronically injured spinal cord ....................... 141	Figure 3.3 SKP-SCs show a predominantly rostral-caudal orientation and a low incidence of proliferation at 21 weeks post transplantation ............................................................................ 143	Figure 3.4 High SKP-SC survival results in significantly more intact tissue than in matched control animals ............................................................................................................................ 145	Figure 3.5 SKP-SCs mitigate the formation of the chronic glial scar ........................................ 146	Figure 3.6 SKP-SCs promote axonal growth/regeneration into the chronic lesion .................... 148	Figure 3.7 Significantly more TH+ and SERT+ axons at the rostral and middle levels of the SKP-SC bridge compared to the caudal levels ........................................................................... 150	Figure 3.8 SKP-SCs myelinate axons ......................................................................................... 151	Figure 3.9 The SKP-SC group contains significantly more SC myelin and non-myelinating SCs than the control groups ................................................................................................................ 152	Figure 3.10 Rat SKP-SCs harvested from adult skin express typical SC markers in vitro ........ 154	Figure 3.11 SKP-SCs isolated from adult skin survived well and demonstrated similar properties as neonatal isolated SKP-SCs ..................................................................................................... 155	Figure 3.12 Chronically contused animals transplanted with SKP-SCs demonstrate greater locomotor outcomes and decreased pathological thickening of the bladder wall ...................... 157	Figure 3.13 SKP-SCs elicited functional improvements on paw angle and step sequence parameters on the CatWalk ......................................................................................................... 159	xvii  List of Abbreviations + = -positive  aSMA= a smooth muscle actin ANOVA= analysis of variance APC= adenomatous polyposis coli (APC; also known as CC1) ASIA= American Spinal Injury Association  ATP= adenosine triphosphate BDNF= brain derived neurotrophic factor BBB= Basso, Beattie and Bresnahan locomotor rating scale BMP= bone morphogenetic protein BMSC(s)= MSC(s) from bone marrow  Brn2= brain 2 BSA= bovine serum albumin CAD= Canadian dollars CAD19= cadherin-19 caspr= contactin -associated protein  CC1= adenomatous polyposis coli (also know as APC) CD= cluster of differentiation (ex. CD13: cluster of differentiation-13) CGRP= calcitonin gene-related peptide CIDS= CNS injury-induced immune deficiency syndrome  CIHR= Canadian Institutes of Health Research Cldn11= claudin11 CNP= cyclic nucleotide phosphodiesterase  xviii  CNS = central nervous system CNTF= ciliary neurotrophic factor CreER= Cre recombinase  CS-56= chondrotin sulfate-56 CsA= cyclosporine A CSPG(s)= Chondroitin sulfate proteoglycan(s) CST= cortical spinal tract dpi= days post-injury DMEM= Dulbecco’s modified eagle medium DNA= deoxyribonucleic acid dP6= dorsal progenitor domain DRG= dorsal root ganglia  E= embryonic day  ECM= extracellular matrix molecule EDTA= ethylenediaminetetraacetic acid EdU= 5-ethynyl-2’-deoxyuridine EGF= epidermal growth factor EGR2= early growth response protein 2 (also known as Krox20) Epb4= band 4.1-like protein G EPI-NCSCs= epidermal neural crest stem cells  erbB= erythroblastic leukemia viral oncogene homolog ES= embryonic stem FACS= fluorescence activated cell sorting xix  Fasn= fatty acid synthase FBS= fetal bovine serum FGF= fibroblast growth factor FRET= fluorescence resonance energy transfer GDNF= glial cell line-derived neurotrophic factor GFAP= glial fibrillary acidic protein GFP= green fluorescent protein Glut1= glucose transporter-1 GRP17= glycine rich protein-17 hpi= hours post-injury HSPG= heparin sulfate proteoglycan heparin sulfate proteoglycans (HSPGs) i.p. = intraperitoneal injection  ICC= immunocytochemistry  ICORD= International Collaboration on Repair Discoveries IGF1= insulin growth factor-1 IH= Infinite Horizon (impactor) IHC= immunohistochemistry  lL= interleukin (ex. IL-1: interlukin-1) iPS= induced pluripotent stem Krox20= early growth response protein 2 (EGR2) KSPG(s)= keratin sulfate proteoglycan(s) KW= Kruskal-Wallis one-way analysis of variance Kv1.2= voltage gated potassium channel 1.2 xx  LIF= leukemia inhibitory factor LINGO= neurite outgrowth inhibitor receptor interacting protein  LSD= least significant difference M1= conventionally activated macrophage phenotype M2= alternatively active macrophage phenotype  MAG= Myelin associated glycoprotein MBP= myelin basic protein MCP-1= macrophage chemoattractant protein-1 mGFP= membrane tethered green fluorescent protein  MOG= myelin oligodendrocyte glycoprotein MRF= myelin gene regulatory factor MS= Multiple Sclerosis  MSC(s)= mesenchymal stem cell(s) MWU=Mann-Whitney U NCAM= neural cell adhesion molecule NEG= -negative NF-200= neurofilament 200 NG2= neuron-glia antigen-2 NGF= nerve growth factor NRG1= neuregulin 1 NSPC(s)= neural stem and progenitor cell(s) NT-3= neurotrophin-3 NT-4/5= neurotrophin-4/5  xxi  O4= oligodendrocyte marker 4 Oct6= octamer-binding transcription factor 6 (also known as SCIP; Tst-1, POU3fl) OEC(s)= olfactory ensheathing cell(s) Olig= oligodendrocyte transcription factor (ex. Olig2) OPC(s) = oligodendrocyte precursor cell(s) P0= myelin protein zero p75-NTR= low-affinity nerve growth factor receptor PBS= phosphate buffer solution  PDGFRa= platelet-derived growth factor receptor a PDGFRb = platelet-derived growth factor receptor b PLP= proteolipid protein pMN= ventral progenitor domain  PMP22 = peripheral myelin protein 22 PNS= peripheral nervous system PDL= poly-d-lysine  PTPs= protein tyrosine phosphate sigma RM= repeated measures ROS= reactive oxygen species RST= rubrospinal tract s.c. = Subcutaneous injection   SC(s)= Schwann cell(s) SCI= spinal cord injury SCIP= octamer-binding transcription factor (also known as Oct6) xxii  SCN= Stem Cell Network SEM= standard error of the mean SERT= serotonin transporter Shh= sonic hedgehog Sirt= Sirtuin SKP(s)= skin-derived precursor(s) SKP-SC(s)= skin-derived precursor Schwann cell(s) SMI312= Sternberger monoclonal incorporated antibody 312 Sox10= sex determining region Y-related high mobility group-box10 TGFb= transforming growth factor beta TH= tyrosine hydroxylase  TNFa= tumour necrosis factor- a tSCI= Traumatic spinal cord injury UBC = University of British Columbia USA= United States of America wpi= weeks post-injury YFP= yellow fluorescent protein   xxiii  Acknowledgements Firstly, I would like to thank my supervisor and mentor Dr. Wolfram Tetzlaff. I have learned a tremendous amount from Dr. Tetzlaff over the years, including both invaluable scientific training and the odd fantastic pun. I cannot thank him enough for enriching my scientific environment with unwavering excitement about discovery, valuable advice, and for shaping me into the critical thinker I am today.   I would like to thank my committee members, Dr. Tim O’Connor, Dr. Brian Kwon, and Dr. Fabio Rossi, who guided me through my PhD with patience, encouragement, and support.   I am extremely grateful to the funding agencies that supported this work: the Canadian Institutes for Health Research (CIHR), the CIHR Transplantation Training Program, the Stem Cell Network (SCN), and the University of British Columbia (UBC). I would also like to thank the International Collaboration on Repair Discoveries (ICORD), the SCN, and the Neuroscience Program at UBC for the many travel awards over the years that allowed me to travel to conferences to present my work, and learn from my international colleagues. I would like to acknowledge the contributions of the animal models; without them, we could not continue our important research.  I would like to thank Drs. Mary Bunge, Lyn Jakeman, and Naomi Kleitman for being incredible female scientific role models; Drs. Freda Miller and Jeff Biernaskie for expert assistance and for hosting a portion of the research; Drs. William Richardson, Hiroshi Takebayashi (via Dr. Scott Whittemore), and Ueli Suter who shared transgenic mice; and Drs. Michael Wegner, Matt Ramer, and J. Trimmer for gifting antibodies.   I would like to thank the members of the Tetzlaff lab past and present who supported me along the way. Of important note, Gregory J. Duncan motivated me via his tireless dedication to research and learning, his kind gestures, and his caring contributions to our collaborative work over the years. I learned a lot from him and enjoyed all of our conversations, intellectually stimulating and otherwise. I also want to highlight my two mentors and senior students in the laboratory, Drs. Joseph Sparling and Jason Plemel, who made the most substantial impact on both my experience in graduate school and the enclosed data. I could not have done it without their friendships, support, feedback, and advice. Working with Michael Lee was a highlight of my time in the lab. Although I was supposed to be his supervisor, he taught and supported me a ton over the years and I would have been ‘at a loss without him'. Dr. Brett Hilton, I thank you for the great conversations, enjoyable collaborations, important feedback, and tireless editing. Jie Liu is an all-star surgeon and contributed so much to this work. Clarrie Lam and Nicole Janzen were outstanding lab managers who kept our scientific environment on the rails and supported xxiv  me when I needed it the most. I also want to acknowledge the ever-growing group of Tetzlaff trainees whose support, assistance, feedback, and critical discussions made my PhD years manageable and enjoyable: Dr. Ward Plunet, Dr. Femke Streijger, Dr. Oscar Seira, Yuan Jiang, Doug Brown, Sohrab Manesh, Aaron Moulson, Maia Blomberg, Peter Fan, Kenny Wu, Nathan Micheals, Ailbish Skinner, Lisa Anderson, Leo Wu, Jasdeep Grewal, Danielle Chung, Daniel Sykora, Bas L Fransen, Tim Beernink, Saba Fatemi, Steven Sidhu and others.  I would like to thank the greater ICORD network that has enriched my training environment over the years. Dr. Leanne Ramer has been an incredible mentor whose help and editing was invaluable in my thesis writing process. Dr. Tom Oxland and Dr. Lowell McPhail have provided outstanding leadership and support over the years. Dr. Jacquelyn Cragg has provided advice, friendship, and coffee dates that were more valuable than she likely knows. In addition, Drs. Andrei Krassioukov, Matt Ramer, Jessica Inskip, John Kramer, Chris West, Jaimie Borisoff, Tim Bhatnagar, and Mark Crawford, as well as Diana Hunter, Steve Mattucci, Jie Liu, and Cameron Lam provided encouragement and help over the years. Finally and importantly, Cheryl Niamath, Matt Sahl, Jeremy green, Simon Liem, and Yasaman Best provided invaluable guidance and assistance.  My non-academic support network has also provided unfailing support and encouragement. My involvement in my many sporting endeavors has helped me maintain some semblance of work-life balance. To my many diverse teammates, coaches, and managers, thank you for motivating me to stay healthy and always pushing me to always be better. To my friend, Russell Winkelaar, thank you for your support and accompanying me on so many fun adventures over the years. My friends Jamie, Aly, Darryl, Kate, Scott and the rest of the Ontario crew have accepted and supported me, and always making my visits home eventful and enjoyable. Thanks to Anand for helping me get through the end of this lengthy process and special thanks to Monica for supporting me through this PhD process even though she has already done her share of supporting.   I thank my whole family for their love and support over the years. Lastly and most importantly, I thank my Mom, Dad, Brother and Grandma, for telling me that I can do anything, for supporting me while I try, and for understanding when I moved across the country to follow my dreams. xxv  Dedication  To the SCI community, in the hopes that research will result in meaningful advances and tangible change.   1 Chapter 1: General introduction 1.1 Introduction overview The loss of myelin after spinal cord injury (SCI) is thought to contribute to motor, sensory, and autonomic dysfunction. Re-establishing myelin after SCI occurs via both new myelin formation on a pre-existing axon that survived the injury (remyelination in response to demyelination) or new myelin formation on a newly grown axon segment (myelination). The degree to which myelin loss is progressive after injury, and to which endogenous myelination contributes to spontaneous repair after SCI, remain points of contention amongst researchers. Research to date has focused on two strategies to repair or replace myelin after SCI: 1) manipulating the speed or extent of endogenous myelin repair or 2) introducing exogenous myelinating cells via cell transplantation. This dissertation examines spontaneous endogenous myelin repair in the wake of experimental contusion SCI and the efficacy of transplanting exogenous myelinating cells as a candidate treatment.  In the introduction, I discuss the clinical implications of SCI, particularly as they affect the direction of SCI research. I describe the relevant cascades of secondary damage that contribute to the pathophysiology of contusion SCI, which is the most widely-used pre-clinical model of SCI. I review the mechanisms and rationale underlying cell transplantation as a therapy for SCI and then focus on reviewing the biology of myelinating glia (oligodendrocytes and Schwann cells [SCs]), and the literature on the behavior of glial lineage cells in response to injury. Finally, I focus on the literature on experimental SC transplantation, discussing treatment options tested pre-clinically and clinically to date.   2 1.2 Clinical implications of SCI The central nervous system (CNS) governs our motor, sensory and autonomic functions and the spinal cord is mainly responsible for transmitting neural signals to and from the peripheral nervous system (PNS) and ultimately to the effector organs. SCI disrupts the transmission of these signals between the brain and the body. Due to the limited intrinsic repair capacity of the CNS, damage to the spinal cord (a product of both primary and secondary mechanisms of injury, described below) often results in permanent and profound functional deficits. The extent of neurological impairments in motor, sensory and autonomic function correspond to the level and severity of the injury (Kirshblum et al., 2002), and are a product of damage to both white matter ascending and descending tracts and interneurons and motor neurons of the grey matter.  The care costs associated with SCI are substantial: the lifetime health care costs per person average $1.5-$3 million Canadian dollars (CAD) depending on the level of injury and the yearly economic burden associated with the ~1300 people with traumatic SCI surviving their injury each year is estimated at 2.5 billion CAD (Krueger et al., 2013). For patients and their families, SCI is a life-changing event, demanding adaptions that are tolerated to different degrees by different people. To conduct research that will positively impact the SCI community, we must first understand more about this diverse group of individuals and their priorities for recovery and repair.   1.2.1 SCI demographics SCI can be the repercussion of either a traumatic (e.g., motor vehicle accident or fall) or a non-traumatic (e.g., birth defect, tumors etc.) incident (Norenberg et al., 2004).   3 The worldwide prevalence of traumatic SCI (tSCI) is estimated to be between 223 and 755 per million (Wyndaele and Wyndaele, 2006). When estimated per country, the annual incidence rates in traumatic SCI varied from 12.1 per million in the Netherlands to 57.8 per million in Portugal (van den Berg et al., 2010; Nijendijk et al., 2014; Jazayeri et al., 2015). The incidence in Canada falls at the high end of this wide range: the discharge incidence of tSCI in Canada is estimated at 41 per million.  An estimated ~85,000 Canadians are currently living with a SCI of which 51% (43,974) are caused by trauma  (Noonan et al., 2012) indicating that the worldwide estimates cited above are likely too low. Recent data reveal that degenerative cervical myelopathies are common and could increase the latter figures (Davies et al., 2017) . Historically, tSCI was prominent in the younger population while non-traumatic SCI was more typical in older people (Noonan et al., 2012); however, this distribution is changing due to falls in an aging, yet still active, population (van den Berg et al., 2010). Males are 2-4 times more likely to sustain a tSCI (Wyndaele and Wyndaele, 2006; Couris et al., 2010). The focus of most research, including my dissertation, is tSCI; therefore, for the purposes of this dissertation, SCI is synonymous with tSCI. However, we should acknowledge the need for developing more models of non-traumatic SCI to facilitate research to address this patient population. To date, most work in rodent models of non-traumatic SCI involves intramedullary spinal cord tumours (Ex. Caplan et al., 2006; Zhuang et al., 2017). However, recent advances include models of chronic syringomyelia (Lee et al., 2017), degenerative cervical myelopathy (Davies et al., 2017), and genetic models of neural tube defects (Mohd-Zin et al., 2017).    4 The functional outcome of SCI, is dictated by severity and level of injury, and is described using the American Spinal Injury Association (ASIA) impairment scale. The AIS A scale is based on the highest detectable level of preserved sensory and motor function; a diagnosis of AIS A indicates complete loss of motor and sensory function below the level of SCI while AIS E indicates normal neurological function. It is important to note that functional completeness does not correspond with anatomical completeness: even among patients with a diagnosis of AIS A SCI, anatomical transection is rarely observed (Kakulas, 1984) and over one hundred thousands axons may be spared (Kakulas and Kaelan, 2015). This is important from a research perspective, since it indicates that spared axons that span the site of SCI are not conducting information across the injury site or if they do their signals/connections are not functionally effective.   Historically, many patients who sustained complete cervical SCI died early after injury or within the first year post-SCI, succumbing to SCI-related complications (Donovan, 2007). Advances in acute care and management have dramatically improved both survival rate and functional outcome of SCI over the past 30 decades. The estimated prevalence amongst Canadians suggest that 56% of people with a SCI are paraplegic (due to thoracic, lumbar or sacral injures) and that 44% are tetraplegic (cervical SCI) (Noonan et al., 2012). Regarding the incidence of SCI, recent data reveal that 66% of acute traumatic SCIs occur at the cervical level of the spinal cord and the majority of them are functionally incomplete (i.e., with an AIS score of B, C, or D (Dvorak et al., 2014).    5 1.2.2 Priorities of the SCI population Neurological impairments resulting from SCI include muscle weakness and paralysis, loss of sensation, muscle stiffness and spasticity, neuropathic pain, cardiovascular deficits, respiratory deficits, impaired sexual and reproductive function, and impairments of bowel and bladder function. These neurological deficits give rise to a host of secondary health complications, with the most problematic including urinary tract infections, pressure sores/ulcers, and respiratory infections (Levi et al., 1995; Winslow and Rozovsky, 2003; Dryden et al., 2004). Importantly, the SCI population has expressed their research priorities and emphasized the importance of improvements in hand and arm function for tetraplegics and sexual function for paraplegics. Improved bladder and bowel function were a priority for both populations (Anderson, 2004). A recent systematic review of similar studies confirmed these findings, and emphasized the critical role of managing secondary complications of SCI in achieving quality of life (Simpson et al., 2012). Based on these studies, researchers have placed more emphasis on understanding pathological mechanisms underlying autonomic dysfunction and hand impairment, and on developing outcome measures specific to hand function and autonomic consequences of SCI.  1.3 Pathology of contusion SCI Primary injury to the spinal cord is usually produced by a burst fracture of a vertebral body or by fracture-dislocation of the spinal column resulting in incomplete anatomical damage. Full histological laceration of the spinal cord is a less common event and typically seen as a result of full fracture-dislocation of the spinal column in high   6 speed accidents or the result of violence (gunshots, knifes) (Bunge et al., 1993; Dumont et al., 2001b; Norenberg et al., 2004). Here, I review the pathophysiology of contusion SCI in both humans and animal models and the progression of changes over days, weeks, and months, using the widely-accepted concepts of primary injury followed by secondary injury cascades.   1.3.1 Primary SCI Primary injury involves a series of near-immediate pathological events after SCI resulting in the loss of neurons and CNS glia (oligodendrocytes and astrocytes) (Bunge et al., 1993; Norenberg et al., 2004; Boldin et al., 2006; Choo et al., 2007). It is characterized by direct damage to the vasculature, causing hemorrhage and impaired tissue perfusion at the injury site (Tator and Fehlings, 1991). This coincides with the severing of axons and the disruption of cell membranes at injury epicenter, resulting in the rapid necrotic death of spinal cord cells and associated damage to cells, axons, and myelin in the areas adjacent to the injury epicenter (Hausmann, 2003; Choo et al., 2007; Mothe and Tator, 2012). Unlike in humans, in the case of contusion SCI in our animal models, the cell damage and loss is most severe in the dorsal and central aspects of the spinal cord (due to the dorsal impact after laminectomy); cell loss is apparent in the grey matter in the early stages of injury. Importantly, differences in the primary injury mechanisms in animal models of contusion, dislocation or distraction cause differences in the pattern and extent of hemorrhage and tissue strain, and lead to differences in secondary pathology (Choo et al., 2008; Chen et al., 2016).  In agreement, in humans a   7 high degree of acute hemorrhage is a predictor of poor functional outcomes (Noyes, 1987; Boldin et al., 2006). Our understanding of pathophysiology of acute SCI in humans is limited to data from imaging and post-mortem autopsies. Although human SCI can be very heterogeneous (see demographics, above), the most common injury involves a blunt (closed) trauma that causes vertebral fracture(s) which in turn produces a ventral contusion of the underlying spinal cord (Bunge et al., 1993; Schwab and Bartholdi, 1996; Dumont et al., 2001b; Norenberg et al., 2004). Animal models of contusion SCI attempt to mimic the human pathology and have become the standard for pre-clinical SCI (Kwon et al., 2002a; Dietz and Curt, 2006). There are several commercially available impactor devices; of these, the Infinite Horizon (IH) impactor was developed specifically to control the force applied to the spinal cord and was used for the experiments included in this dissertation. Of note, the animal model differs from the clinical scenario in that the vertebral laminae and spinous processes (dorsal bony structures overlying the spinal cord) are surgically removed, and the spinal cord is hit directly from the dorsal side to produce the contusion. This dramatically improves reproducibility of the injury, but clearly reduces clinical relevance. Therefore, the laboratory of Dr. Oxland works on the development of animal models for closed injuries as they occur in fracture-dislocations or distractions (Choo et al., 2007; Choo et al., 2008; Chen et al., 2016).  1.3.2 Secondary SCI and injury progression Primary SCI initiates a series of pathological cascades that act over days, weeks, and months to determine the ultimate extent of the injury. The secondary phase of SCI is   8 the result of multiple mechanisms, including hemorrhage, edema, ischemia, free radical generation, vascular dysregulation, immune cell infiltration, ion dysregulation and excitotoxicity (Kwon et al., 2002a; Hausmann, 2003; Kwon et al., 2004; Norenberg et al., 2004; Oyinbo, 2011; Ahuja et al., 2017). After contusion SCI, local inflammation contributes to the formation of a large fluid filled cyst in rats and humans, which is much larger than the primary injury area. Astrocytes become hypertrophic and interact with other types of cells to make up the glial/fibrotic scar, which forms a border around the injury site (Cregg et al., 2014). Our understanding of the cells that make up the SCI lesion site continues to grow; we now appreciate that the chronic injury site is a complex arrangement of interacting cells (Horn et al., 2008; Meletis et al., 2008; Barnabe-Heider et al., 2010; Busch et al., 2010; Goritz et al., 2011; Sabelstrom et al., 2013; Soderblom et al., 2013). The mature lesion site consists of both an outer region, made up largely of hypertrophic astrocytes, and an inner core containing neuron-glial antigen-2 (NG2)-positive (+) cells, meningeal/vascular- derived fibroblasts, pericytes, ependymal cells and phagocytic macrophages (simplified in Figure1.1) (Cregg et al., 2014). The mature lesion is a formidable barrier to axonal regeneration: here, I describe the mechanisms of secondary injury relevant to my work in this dissertation. Secondary SCI: vascular effects Primary SCI creates an area of hemorrhage, but both the affected area and the associated hemorrhagic necrosis get larger over time post-injury (Simard et al., 2007). Hemorrhagic necrosis is thought to result primarily from hemoglobin neurotoxicity (Regan et al., 2008) and the associated accumulation of iron (Hua et al., 2006), and seems   9 to preferentially affect neurons and OPCs (Thorburne and Juurlink, 1996; Regan and Guo, 1998). SCI is also associated with extensive edema and ischemia (Dumont et al., 2001b) resulting from blood brain barrier breakdown (Bartanusz et al., 2011) and local vascular damage and associated effects (Dumont et al., 2001b), respectively. These processes are succeeded by the release of vasoactive and/or endothelial destructive compounds that are responding to damaged neural and endothelial cellular membranes, the activation of glial cells, and the arrival of blood-borne inflammatory cells (Noble and Wrathall, 1989; Tator and Fehlings, 1991; McKenzie et al., 1995; Popovich et al., 1996; Schnell et al., 1999). Blood brain barrier breakdown expands with time, peaking days after SCI, but appears to persist in chronic injury (Cohen et al., 2009).  In addition to blood entering the parenchyma of the spinal cord, vascular damage disrupts blood supply. The resulting ischemia creates a zone of oxygen and glucose deprivation, and causes energy failure (Saikumar et al., 1998; Harris and Attwell, 2012). Neurons and oligodendrocytes are particularly vulnerable to ischemic conditions. Neuronal consequences include, edema, ongoing membrane depolarization, release of excitatory neurotransmitters (mainly glutamate), and decreased neurotransmitter reuptake (Liang et al., 2007; Harris and Attwell, 2012). White matter is also vulnerable to ischemic damage (Pantoni et al., 1996) presumably because oligodendrocyte lineage cells are particularly susceptible to oxidative stress (Noble et al., 1988; Merrill et al., 1993; Oka et al., 1993; Husain and Juurlink, 1995). As metabolic by-products and cellular contents accumulate at the site of SCI, cells that survived the primary injury are damaged and die. A major cause of oligodendrocyte cell death after injury is excitotoxicity; elevated glutamate preferentially kills oligodendrocyte cells compared to astrocytes (Matute et al.,   10 1997; McDonald et al., 1998; Li and Stys, 2000). Glutamate accumulation (Panter et al., 1990; Wrathall et al., 1996) has a net effect of increased intracellular calcium (Choi, 1988; Matute et al., 2007) which is associated with many harmful downstream events, such as activation of proteases, lipases, phosphatases, and endonucleases as well as disruption of ion transporters, endoplasmic reticulum and mitochondria, and increased production of reactive oxygen species (ROS) (Kroemer et al., 1998; Dumont et al., 2001a; Szydlowska and Tymianski, 2010). The resulting ROS buildup results in extensive neuronal and glial membrane damage by peroxidation near the site of SCI (Jia et al., 2012). Release of extracellular ATP also contributes to oligodendrocyte cell death (Wang et al., 2004). The dramatic loss of oligodendrocytes after SCI is central to this dissertation; in animal models 93% of oligodendrocytes die at the lesion epicenter within 7 days of SCI (McTigue et al., 2001). Secondary SCI: immune effects  Breakdown of the blood brain barrier and leakage of blood into the CNS tissue instigates a profound immune response (Donnelly and Popovich, 2008; Hawthorne and Popovich, 2011). CNS-derived microglia (Ajami et al., 2007) respond to local injury within seconds (Davalos et al., 2005; Nimmerjahn et al., 2005; Hines et al., 2009) and their production of cytokines, chemokines, and eicosanoids attract other inflammatory cells within the first day of injury. Proinflammatory cytokines (e.g., tumour necrosis factor alpha[TNF-a], interleukin [IL]-1B, IL-2 and interferon -g) may be a major cause of oligodendrocyte death after SCI; these have been demonstrated to induce cell death in   11 culture (Curatolo et al., 1997; Sherwin and Fern, 2005; Li et al., 2008) and be present in higher levels after SCI (Kwon et al., 2010b; Stammers et al., 2012).   Neutrophils are the first blood-borne inflammatory cells to arrive at the injury site, peaking in number at 12-24 hours post-injury (hpi) and then disappearing in the weeks post-injury (wpi) (Taoka et al., 1997; Carlson et al., 1998; Fleming et al., 2006; Beck et al., 2010). In open wounds neutrophils are thought to sterilize the injury site and may be required for promoting functional recovery but they are a rich source of free oxygen radicals and myeloperoxidase that can contribute to damage of uninjured cells in the spared tissue of the spinal cord (Taoka et al., 1997; Neirinckx et al., 2014). In rats, lymphocytes infiltrate the lesion early on, with maximum accumulation 3-7 days post-injury (dpi); in contrast lymphocyte accumulation in both mice and humans peaks months after injury (Sroga et al., 2003; Kigerl et al., 2006).  Microglia undergo proliferation and contribute to phagocytosis, becoming indistinguishable from blood-derived macrophages (Hausmann, 2003; David and Kroner, 2011); accumulation of these immune cells peaks 5-7 days post SCI (Popovich et al., 1997; Carlson et al., 1998; Fleming et al., 2006; Ajami et al., 2011). These accumulated activated macrophages/microglia persist at the injury site for weeks to months (Popovich et al., 1997; Carlson et al., 1998; Fleming et al., 2006) and drive progression of secondary injury (Cregg et al., 2014); for example, they play a significant role in lesion clearance and protracted axonal dieback (Horn et al., 2008). Macrophages at the site of SCI are generally pro-inflammatory or “conventionally activated” (M1) in phenotype; without intervention, these tend to dominate the lesion indefinitely, without transitioning to an anti-inflammatory or “alternatively activated” (M2) phenotype (Mosser and Edwards, 2008; Kigerl et al., 2009). T-lymphocytes also   12 play a role after SCI: their accumulation peaks 6-10 weeks post-injury, and their contribution is relatively small compared to the other leukocytes involved (Popovich et al., 1997; Schnell et al., 1999; Fleming et al., 2006; Goldmann et al., 2016; Jin and Yamashita, 2016). Secondary SCI: the glial scar After CNS injuries, including SCI, the lesion becomes surrounded by reactive astrocytes, which align and interact with other cells to form a glial scar. The scar serves to protect the remaining parenchymal tissue and cordon off the injured area and intense inflammatory zone (Silver and Miller, 2004; Cregg et al., 2014; Liddelow and Barres, 2016). While the scar does serve a protective function by protecting normal tissue from an inflammatory lesion core (Burda and Sofroniew, 2014; Cregg et al., 2014; Filous and Silver, 2016), the scar is generally thought to be both a physical and chemical inhibitor of axon growth. A caveat to this is that, recent work using transgenic mouse lines suggests that preventing the formation of the astrocytic scar significantly reduced the ability of axons to regrow (Anderson et al., 2016), which demonstrates the complexity of the glial scar-axon interactions after injury. However, this prevention of scar formation will increase inflammation which becomes a confounding factor in this experiment. The scarring process starts within days of injury when inflammatory cells accumulate in the lesion and initiate the migration and some proliferation of astrocytes to the outermost lesion border (Fitch and Silver, 1997).  A thin layer of reactive astrocytes accumulates on the spared rim (in the case of contusion SCI) and proliferates (Bush et al., 1999; Faulkner et al., 2004; Wanner et al., 2013; Sofroniew, 2015). Primary SCI causes   13 an initial reduction of ~50% of the astrocytes, depending on the severity of injury (Rosenberg and Wrathall, 1997; Grossman et al., 2001), but this loss is replenished by injury-induced proliferation. However, the majority of the reactive response is due to astrocyte hypertrophy rather than glial cell division: reactive astrocytes produce and accumulate intermediate filament proteins, including glial fibrillary acidic protein (GFAP), vimentin, and nestin (Pekny et al., 1999; Xu et al., 1999a; Wilhelmsson et al., 2004; Bardehle et al., 2013). Over time, there is a restructuring of the astrocytic lesion border such that it becomes a physical barrier that is particularly obstructive to axons growing along the long axis of the spinal cord (Bardehle et al., 2013; Wanner et al., 2013).  Reactive astrocytes produce a host of molecules that are potent chemical inhibitors of axon growth including chondroitin sulfate proteoglycans (CSPGs) (McKeon et al., 1991; McKeon et al., 1995; McKeon et al., 1999; Yamaguchi, 2000; Jones et al., 2003; Tang et al., 2003; Busch and Silver, 2007; Alilain et al., 2011; Brown et al., 2012; Kawano et al., 2012; Li et al., 2013; Takeuchi et al., 2013) and karatan sulfate proteoglycans (KSPGs) (Jones et al., 2002; Imagama et al., 2011). CSPGs are present within 24 hours of SCI and increased expression has been noted for months post-injury (McKeon et al., 1999; Jones et al., 2003; Tang et al., 2003). The CSPGs are a family of extracellular matrix molecules (ECM) proteins, including neurocan, aggrecan, brevican, phosohacan, versican and NG2 (Margolis and Margolis, 1993; Cregg et al., 2014). After injury, CSPGs may inhibit axonal growth by binding to the axonal receptor protein tyrosine phosphate-s (PTPs); this interaction converts the distal axon from an active growth cone to an immobilized, dystrophic axon tip (Lang et al., 2015).   14 In addition to mature astrocytes, several other cell types contribute to the glial scar. For example, dividing ependymal cells also take on reactive astrocyte phenotypes and can contribute to scar formation (Johansson et al., 1999; Meletis et al., 2008; Barnabe-Heider et al., 2010). There is also evidence that fibroblast-like-stromal cells can divide and add to the scar. These cells appear to be derived from meninges, pericytes, or other vascular-associated cells and form a fibrotic-like layer that interacts with scar astrocytes (Decimo et al., 2011; Goritz et al., 2011; Fernandez-Klett et al., 2013; Sabelstrom et al., 2013; Soderblom et al., 2013). For example, Type A pericytes (as defined by Goritz et al., 2011) delaminate from blood vessels in response to injury and give rise to a stromal cell population which contribute to the fibrotic scar (Goritz et al., 2011). In addition, a fate mapping study demonstrated that collagen-1a1/platelet derived growth factor receptor b (PDGFRb)+/cluster of differentiation (CD)-13+ cells proliferate and contribute to the fibrotic scar after injury (Soderblom et al., 2013). This fibrotic component of the scar is replete with extracellular fibronectin, collagen, and laminins (Shearer and Fawcett, 2001). Analogous to astrocytes and CSPGs, these stromal cells and their associated ECM molecules are thought to help wall off the lesion site but may also block axon regrowth (Davies et al., 1999; Pasterkamp et al., 1999; Stichel et al., 1999; Kawano et al., 2012; Sabelstrom et al., 2013; Soderblom et al., 2013; Cregg et al., 2014).  Neuron glial factor 2 (NG2)+ cells, including oligodendrocyte precursor cells (OPCs), are also associated with scarring. NG2 is the most heavily up-regulated CSPG after CNS injury (Levine, 1994) and accordingly was thought to play a role in inhibiting regeneration after injury (Dou and Levine, 1994; Fidler et al., 1999; Chen et al., 2002; Tan et al., 2006). Recently, NG2+ cells have been observed in close proximity with   15 dystrophic tips of severed axons (Zhang et al., 2001; McTigue et al., 2006; Busch et al., 2010) and may actually support dystrophic axons persisting at the lesion-parenchymal border (Di Maio et al., 2011; Hughes et al., 2013; Filous et al., 2014) thus decreasing axonal dieback. Further research is required to better understand this interaction which may involve neurons forming synapses with these NG2+ OPCs, potentially limiting the advancement of a dystrophic axon tip (Silver et al., 2015).   1.4 Cell transplantation for repair of SCI 1.4.1 Historical perspective Cell transplantation at the site of CNS injury has a long and distinguished history of researchers attempting to replace the environment of the CNS with a permissive substrate for growth. In the early 1900s, in the laboratory of Santiago Ramon y Cajal and Jorge Francisco Tello, transplanted pre-degenerated peripheral nerve segments onto the proximal stumps of severed optic nerves and observed retinal axons growing into the nerve grafts (Cajal, 1991). This seminal work was revisited in 1981, when experiments by Richardson, David and Aguayo demonstrated that adult spinal axons can grow long distances into peripheral nerve grafts (Richardson et al., 1980; David and Aguayo, 1981). In parallel and applying a similar rationale, Richard Bunge envisioned treating SCI with SCs, and the pioneering work by Bunge, Bunge and Wood on in vitro cultures of SCs for spinal cord repair has led to recent clinical trials (Bunge, 2016). The notion that fetal tissue can promote CNS repair was established by Anders Björklund and colleagues, who replaced lost cell types in models of Parkinson’s and Huntington’s disease (Bjorklund and Lindvall, 2000). Transplantation of fetal spinal cord tissue into injured spinal cord   16 resulted in successful graft survival, differentiation of transplanted cells, host–graft integration, and connectivity (Bregman et al., 1993). Intraspinal grafting of human fetal spinal cord tissue is both feasible and safe in humans (Wirth et al., 2001). Current approaches owe a great deal to these researchers, who established the capacity of transplanted cells to differentiate, promote axon growth and myelinate within the injured spinal cord.  1.4.2 Candidate cell types While numerous cells and tissues have been assessed for their capacity to treat SCI with transplantation, the most widely studied are SCs (see 1.6), neural stem and progenitor cells (NSPCs), OPCs, olfactory ensheathing cells (OECs) and mesenchymal stem cells (MSCs) (Tetzlaff et al., 2011). Cells for transplantation can be generated from adult and embryonic sources, induced pluripotent stem cells (Lu et al., 2014) and, potentially, via direct conversion technology (Yang et al., 2013). Transplanted cells vary with respect to the species, age, and source of the donor, as well as culture conditions and co-treatments. As a result, comparisons between studies, even those that focus on a particular cell type, are challenging, as the cell name (for example, NSPC or MSC) can be an umbrella term for an assortment of cells that differ due to these variables. The timing after SCI and location of cell transplantation can also influence transplanted cell fate; for example, NSPCs transplanted into host parenchyma predominantly differentiate into oligodendrocytes, while lesion-site transplantation leads to more astrocyte differentiation (Piltti et al., 2013a). Transplanted cells may improve the functional   17 outcome of SCI through a variety of mechanisms, including neuroprotection, immunomodulation, axon sprouting and/or regeneration and myelin regeneration.  1.4.3 Mechanisms of action of transplanted cells Neuroprotection Many cell types are thought to mitigate the secondary damage after SCI, including SCs, NSPCs, MSCs, OECs and OPCs (Raisman, 2001; Tetzlaff et al., 2011). An increase in spared tissue adjacent to the injury site is generally correlated with improvements in sensorimotor function in preclinical models of injury (Basso et al., 1996; Schucht et al., 2002; Plemel et al., 2008) and in humans with SCI (Kakulas, 2004). However, there are several caveats to studies that propose neuroprotection as a mechanism of action. First, classical neuroprotective agents that are known to reduce secondary damage are only effective within hours of injury (Kwon et al., 2011), yet cell transplantation is often performed 1–2 weeks after SCI to improve graft survival (Tetzlaff et al., 2011). Even at these later time points of transplantation—when known mediators of secondary damage have already been released—transplantation has been associated with increased tissue sparing (Pearse et al., 2004b; Biernaskie et al., 2007; Barbour et al., 2013). Second, many cell transplantation candidates might contribute to the appearance of increased spared white matter by producing myelin in the adjacent parenchyma on spared axons. SC myelination is particularly relevant, as it requires more space than oligodendrocyte myelin and might enlarge the cross-sectional area of white matter tracts (Lankford et al., 2002). Third, cell transplantation could enhance axon regeneration and sprouting,   18 followed by myelination in the parenchyma surrounding the injury site. If there are more myelinated axons or regenerated fibers, this could in itself increase the cross-sectional area adjacent to the injury site. Thus, an increase in spared tissue adjacent to the injury site might reflect bona fide protection, a regenerative response that increases the amount of white matter adjacent to the injury site, or a combination of both. Molecules secreted by transplanted cells One widely proposed mechanism underlying neuroprotection mediated by transplanted cells is the secretion of bioactive molecules such as trophic factors and cytokines. Many cell types, including MSCs, NSPCs and SCs, secrete trophic factors and cytokines in vitro and can increase the presence of these factors after transplantation (Sharp et al.; Hawryluk et al., 2012; Cantinieaux et al., 2013; Zhang et al., 2013a). While these factors may enhance host cell survival, regulate gliosis, modulate inflammation and/or improve the regeneration of blood vessels, the increase in these factors and their neuroprotective properties is largely correlative. Only a few studies have systemically employed cell-type specific deletion to investigate which cell-derived factor(s) is (are) necessary for neuroprotection. These studies require the transplantation of cells that are modified and diminished in their capacity to express/secrete specific factors (Ritfeld et al., 2015). For example, the neuroprotective effects of transplanted MSCs from bone marrow (BMSCs) after SCI are reduced when secretion of brain-derived neurotrophic factor (BDNF) is impaired (Ritfeld et al., 2015). However, BDNF knockdown also reduces BMSC survival, leaving it unclear whether BDNF directly promotes tissue sparing or whether it is secondary to some other BMSC-mediated neuroprotective effect.   19 Moving forward, it will be important to understand the secretome of transplanted cells and to use factor-specific loss-of-function experiments to determine what factors are beneficial after transplantation. Immunomodulation While some aspects of inflammation are related to tissue damage, many repair mechanisms rely on beneficial aspects of inflammation (Jones et al., 2005; Plemel et al., 2014a; Gadani et al., 2015). The complex nature of inflammation is best exemplified by the phenotypic separation of macrophages and potentially microglia into neurotoxic, proinflammatory (M1) and immunoregulatory (M2) subsets, the latter of which secretes factors that promote axon growth (Kigerl et al., 2009; Gensel and Zhang, 2015) and enhance remyelination (Miron et al., 2013). While the M1 or M2 designation reflects an oversimplification and there are other macrophage activation states (Xue et al., 2014), they are yet to be described in vivo after SCI (Gensel and Zhang, 2015). Cell transplantation can provide benefits through immunomodulation, by attenuating detrimental inflammation or stimulating beneficial inflammation. For example, NSPC transplantation improves ankle movement and hindlimb placement in association with a slight increase in the abundance of T cells, a decrease in B cells, and reduced M1-like macrophages following SCI (Cusimano et al., 2012). NSPCs express and secrete many factors that could regulate inflammation, yet demonstrating a causal role would require transplanting cells with specific genetic modifications of candidate genes of interest. MSCs can also modify the immune response after injury by elevating levels of anti-  20 inflammatory cytokines and reducing levels of proinflammatory cytokines when transplanted 3 or 7 days after SCI (Abrams et al., 2009; Nakajima et al., 2012). Although MSCs do not enter the spinal cord when delivered intravenously 1 day after contusive SCI, their injection is associated with increased forelimb–hindlimb coordination and improved micturition, which correlates with increased spared tissue and an elevation in M2 markers (DePaul et al., 2015). There is also decreased proinflammatory cytokine production in the spleen and blood, indicating a systemic change in the inflammatory state. Importantly, conditioned medium from MSCs can improve motor function after SCI, suggesting that cell transplantation may not be required to achieve functional benefits (Cantinieaux et al., 2013). Preventing blood vessel loss or improving angiogenesis There is widespread blood vessel loss after SCI, which induces local hypoxia within and adjacent to the injury site (Tator and Fehlings, 1991; Okon et al., 2013). The proliferation of endothelial cells, or angiogenesis, occurs as early as 3 days after SCI, but only after 14 days does vessel perfusion recover to baseline levels (Figley et al., 2014). Restoration of oxygen and nutrients within spared tissue is rarely discussed but might be a general mechanism by which transplanted cells influence outcome; transplant-derived secreted molecules might promote blood vessel protection and regeneration. For example, fetal tissue grafts re-establish oxygen levels within the graft and adjacent to the injury site (Stokes and Reier, 1991), which is correlated with elevated blood vessel density in and around the transplant site (Horner et al., 1996). Fetal tissue grafts also elevate glucose   21 utilization, suggesting an overall increase in metabolism compared to SCI injured controls (Horner and Stokes, 1995). In addition, transplanted OECs increase vasculature within the graft and align vasculature along the length of the spinal cord (Lopez-Vales et al., 2004; Ramer et al., 2004; Richter et al., 2005), but whether they, or any other transplanted cell type, accelerate tissue re-oxygenation and nutrient access after injury is unknown. The use of in vivo live imaging techniques with vital dyes or genetically encoded fluorescence resonance energy transfer (FRET) sensors to measure dynamic changes in tissue oxygen (Quaranta et al., 2012), glucose (Bittner et al., 2010) and/or other tissue metabolites (Plemel et al., 2014a), coupled with injury and transplantation, will improve our understanding of the dynamic metabolic changes after SCI and how transplantation might regulate these processes. In these investigations of tissue oxidation and metabolites, large animal models such as pigs may be crucial, since they permit the placement of dialysis probes or pressure sensors into the spinal cord (Okon et al., 2013). Axonal growth A major goal of cell transplantation after SCI is to promote axon regeneration and plasticity. One well-established mechanism of transplantation-induced axon growth is formation of growth-permitting bridges across the lesion site (Figure 1.2a). In addition to the grafted material spanning the lesion site, the molecules transplanted cells produce, such as laminin, provide a substrate for axon growth. SCs (Xu et al., 1995a), OECs (Takeoka et al., 2011), NSPCs (Lu et al., 2012) and MSCs (Lu et al., 2004) can form bridges following SCI leading to axon regeneration, which can generally be accentuated   22 by combining cells with biomaterials (Fouad et al., 2005; Lu et al., 2012; Krishna et al., 2013) or trophic factors (Lu et al., 2012; Bunge, 2016). There is considerable heterogeneity in the capacity of different axon populations to grow onto bridges: while there are many documented cases of brainstem, propriospinal, and sensory ascending axons on bridges of various sources, there are limited examples of robust corticospinal regeneration onto a transplanted cell bridge (Kadoya et al., 2016). However, stimulating large numbers of host axons to exit these bridges at the distal interface remains a ubiquitous challenge (Xu et al., 1997). While long-distance regeneration of host axons remains elusive, an alternative approach to reconnecting segments following SCI is through the formation of neuronal relays. When transplanted cells are capable of neuronal differentiation, survival, axon outgrowth and synapse formation with host neurons, a relay circuit can form between host axons and newly-differentiated transplanted neurons (Figure 1.2 b). Early studies demonstrated that axons could project into and arise from embryonic day 14 rat spinal cord tissue grafts following their transplantation into SCI cavities (Reier, 1985; Bregman and Reier, 1986; Jakeman and Reier, 1991). More recent findings reveal that NSPCs can differentiate into neurons with a capacity for relay formation (Lu et al., 2012). In this study grafted cells were embedded within a fibrin matrix to aid in their retention within a complete transection site, and the matrix contained a cocktail of trophic factors, plus a protease inhibitor to support cell survival and vascular ingrowth. When this cell-embedded matrix was delivered two weeks after complete SCI, host axons synapsed onto newly-differentiated neurons, which extended large numbers of axons long distances rostral or caudal to injury. This approach resulted in significant hind limb functional   23 improvement: grafted rats had movement around each joint of the hind limb, in contrast to minimal-to-no movement in controls, and this was abolished following re-transection at the rostral graft–host interface. An additional mechanism through which cell transplantation might promote axon growth is by modifying the astrocyte response to injury. For example, after SC transplantation, regenerating axons can be found in direct association with GFAP+ astrocyte processes that elongate in association with transplanted SC processes (Williams et al., 2015). Regenerating axons grow within a basal lamina tunnel alongside transplanted cells and GFAP+ astrocyte processes, and the extent of axon regeneration correlates with the number of astrocyte processes found within the bridge (Williams et al., 2015). This unusual pattern of growth, featuring a close association between axon sprouts and astrocytic processes, suggests that cell transplantation can modify astrogliosis in a way that promotes or permits axon regeneration; however, the molecular mechanisms of this modification remain unknown. Myelin regeneration Loss of myelin is observed early after SCI in both rodents (Totoiu and Keirstead, 2005; Lasiene et al., 2008) and humans (Kakulas, 2004; Norenberg et al., 2004). In addition, oligodendrocyte apoptosis continues for weeks after contusive SCI in rodents, nonhuman primates, and humans (Crowe et al., 1997; Emery et al., 1998). Demyelination impairs conduction and severs an important metabolic shuttle between oligodendrocytes and axons (Funfschilling et al., 2012; Lee et al., 2012c). Replacing lost oligodendrocytes   24 and myelin improves conduction (Smith et al., 1979; Blight and Young, 1989) and may protect axons from degeneration. As a result, using transplanted cells to facilitate myelin regeneration, the process by which cells produce myelin around axons that have lost their myelin sheaths, has been identified as a treatment goal to improve the outcome of SCI (Figure 1.3).  While transplantation of SCs (reviewed in 1.6, below), OPCs, or NSPCs capable of producing oligodendrocytes can be used to enhance myelin regeneration (Tetzlaff et al., 2011; Plemel et al., 2014b; Myers et al., 2016), the specific role of grafted cells remains unclear. Extensive endogenous remyelination occurs following contusion SCI in rodents (Powers et al., 2012). Transplanted OPCs or NSPCs are likely to compete with endogenous remyelinating cells for denuded axons; several studies have found an increase in the number of myelinated axons at endpoint, which is correlated with improved forelimb–hind limb coordination (Keirstead et al., 2005; Cao et al., 2010; All et al., 2015) or enhanced frequency of plantar stepping (Karimi-Abdolrezaee et al., 2006). However, the differentiation of transplanted cells into oligodendrocytes does not always result in improved motor recovery, particularly if the level of remyelination is not sufficient to promote an overall increase in the number of myelinated fibers (Plemel et al., 2011). Improving the differentiation of transplanted cells to oligodendrocytes with exogenous factors or genetic modification may be necessary to increase remyelination above the level attained by host precursors. However, given the efficacy of endogenous remyelination, it is possible that enhancing this process may promote functional improvement without the need for cell transplantation. This is an emerging theme in the field, and is supported by the work in this dissertation.   25 1.5 Myelin biology   Myelin is the extension of the glial plasma membrane duplication that wraps larger axons in both the PNS and CNS. The most widely appreciated role of myelin is increasing the speed of propagation of the action potential by changing the electrical properties of the axon. Myelinated segments (internodes) create regions of high resistance and low capacitance, and restrict action potentials to the nodes of Ranvier. Saltatory conduction – the process by which action potentials travel from node to node – increases the speed of signal propagation 20-100-fold compared to a (theoretical) unmyelinated axon of the same diameter (Nave and Werner, 2014). In addition to optimizing the efficiency of neurotransmission, myelination has other important consequences; for example, the presence of myelin eliminates the need for adenosine triphosphate (ATP)-dependent Na+/K+ exchange along the entire axon and allows for the maintenance of the resting membrane potential (Wang et al., 2008). In addition, the myelin sheath provides crucial trophic support to the axon (Nave and Werner, 2014): when myelin is damaged in trauma or disease, neuronal function and survival are compromised, resulting in axonal degeneration and associated loss of neurological function (Crawford et al., 2013).  The structure of myelin has been well characterized in both the PNS and the CNS and provides information about its development (i.e., newly-established versus mature). The critical microscopic features are the major period lines, which indicate the compacted intracellular membranes, and the intraperiod lines at the extracellular membrane. Mature myelin consists of up to 160 layers of glial membrane, with internodes extending up to 1.7mm in the CNS and 2mm in the PNS (Hildebrand et al., 1993; Hildebrand et al., 1994). SCs are the myelinating cells of the PNS and oligodendrocytes are the myelinating   26 cells of the CNS; they produce structurally similar but distinguishable myelin (Stolt and Wegner, 2016). The most obvious difference is in the number of myelin sheaths observed per cell:  SCs myelinate one axon, while oligodendrocytes myelinate axons in much lower ratios (e.g., up to 1:60) (Nave and Werner, 2014). CNS and PNS myelin in the spinal cord also have unique myelin periodicity (averaging 17nm for SC myelin compared to 15.5nm for oligodendrocyte myelin) and have distinct developmental origins (neural crest for SCs and ventral and dorsal neural tube for oligodendrocytes) (Nave and Werner, 2014). In the PNS, SCs typically myelinate axons with a diameter larger than 1µm. In the CNS, oligodendrocytes myelinate axons over a larger range, starting with diameters from 0.2 to 1.2µm (Lee et al., 2012b; Nave and Werner, 2014).  Myelin in the CNS and the PNS contains both overlapping and distinctive proteins, but can be distinguished by characteristic dominant protein expression. For example, SC myelin contains significant amounts of myelin protein zero (P0; 21%), Periaxin (16%), myelin basic protein (MBP; 8%) and relatively scant amounts (<1%) of proteolipid protein (PLP)1, cyclic nucleotide phosphodiesterase (CNP), myelin-associated glycoprotein (MAG), fatty acid synthase (Fasn), and band 41.-like protein G (Epb4). Oligodendrocyte myelin contains significant amounts of PLP1 (17%), MBP (8%), CNP (4%) and scant amounts (<1%) of myelin oligodendrocyte glycoprotein (MOG), MAG, sirtuin 2 (Sirt2), and claudin 11 (Cldn11) (Nave and Werner, 2014). The cellular origin of myelin is central to this dissertation; I will therefore review relevant features of both oligodendrocytes and SCs in development and as they respond to injury.    27 1.5.1 Oligodendrocyte lineage cells Oligodendrocyte lineage cells are derived from two specific regions of the embryonic neural tube, the ventral progenitor domain (pMN) and the dorsal progenitor domain (dP6). Sonic hedgehog (Shh) signaling is essential for ventral-derived OPCs (acting through the transcription factor Olig2) (Lu et al., 2002) while dorsal bone morphogenic proteins (BMPs) and Wnts suppress dorsal OPC development (Agius et al., 2004; Shimizu et al., 2005). Oligodendrocyte lineage cell production from the pMN occurs first, between embryonic day (E)12.5 and E14.5, when early progenitors give rise to OPCs, which proliferate and migrate dorsally and laterally throughout the spinal cord, and are thought to produce ~80% of oligodendrocytes (Rowitch et al., 2002; Rowitch and Kriegstein, 2010; Fancy et al., 2011; Tripathi et al., 2011). A second wave OPC production from dP6 occurs at E15.5 (Cai et al., 2005; Fogarty et al., 2005; Vallstedt et al., 2005); these progenitors are shh independent (Cai et al., 2005), are thought to account for ~20% of the OPCs in the adult, and preferentially populate the dorsal aspects of the cord. Despite these distinctive origins, these populations of cells produce oligodendrocytes with similar electrical properties, and myelin with similar range and number of internodes (Tripathi et al., 2011).  The myelinating oligodendrocyte is a post-mitotic cell that moves through specification, migration, and proliferation steps from its original OPC state. The oligodendrocyte lineage includes multiple distinguishable developmental stages, including OPCs, immature oligodendrocytes, mature oligodendrocytes and myelinating oligodendrocytes (Emery, 2010). While single proteins that distinguish cells along the oligodendrocyte lineage have proven vague there are cell specific marker combinations.   28 OPCs can be identified due to their expression of platelet derived growth factor receptor a (PDGFRa) and NG2, immature oligodendrocytes by their expression of CNP and oligodendrocyte marker 4 (O4), and mature oligodendrocyte express markers of adenomatous polyposis coli (APC; also known as CC1) and the compact myelin marker MBP. Myelinating oligodendrocytes are often identified phenotypically, as they begin to wrap multiple axons; they also express CC1 and MBP (de Castro and Bribian, 2005). OPCs are the most crucial to this dissertation and therefore will be prioritized within this introduction. In development, OPC proliferation and migration populate the CNS (Kessaris et al., 2006). The cues that direct migration of OPCs are obviously complex but there is evidence indicating that chemotactic cues, such as semaphorins, ephrins, and netrins, play an important role in attracting and repelling OPCs (Jarjour and Kennedy, 2004; Cohen, 2005). Live imaging of OPCs revealed that OPCs migrate by extending many large and highly-branched filopodium-like process in a stochastic manner, until they halt upon making contact with an axon (Kirby et al., 2006). OPCs appear to be responsive to both environmental chemotactic cues but also to each other, as their processes retract upon contact with other OPC processes and therefore occupy a mutually exclusive domain (Hughes et al., 2013). In the adult, PDGFRa/NG2+ OPCs are thought to be fate restricted to the oligodendrocyte lineage in the uninjured situation (Rivers et al., 2008; Kang et al., 2010). However, OPCs in the mature CNS are not quiescent: they play an ongoing role in oligodendrocyte turnover (Rivers et al., 2008; Rosenberg et al., 2008; Lasiene et al., 2009; Psachoulia et al., 2009; Kang et al., 2010). Multiple mitogens facilitate optimal   29 OPC proliferation; for example, blocking neurotrophin-3 (NT-3) or PDGFRa in vivo drastically decreases OPC proliferation (Barres and Raff, 1993; Barres et al., 1994; Fruttiger et al., 1999). In vitro, OPCs also respond to other mitogens, such as basic fibroblast growth factor (FGF), which is sufficient for inducing proliferation (Eccleston and Silberberg, 1985; Bogler et al., 1990; Barres et al., 1993); bFGF is also associated with an increase in PDGFRa expression, which in turn inhibits spontaneous differentiation (Bogler et al., 1990; McKinnon et al., 1990; Tang et al., 2000).  Insulin growth factor-1 (IGF1) also induces proliferation (McMorris and Dubois-Dalcq, 1988; Barres et al., 1993), particularly when accompanied by NT-3 or another OPC mitogen (Barres and Raff, 1994). After OPC specification and proliferation, differentiation, axon ensheathment, and axon wrapping precede the formation of mature compact myelin. Given that differentiation appears to happen spontaneously in vitro (Barres and Raff, 1999), it is thought that differentiation is the default state and that a combination of transcriptional and environmental factors maintain OPCs in their proliferative and non-differentiative state (Li et al., 2009). In other words, it is de-repression of negative regulators that allows for oligodendrocyte differentiation. In development, we know that many differentiation and myelination inhibitors exist (Barres and Raff, 1999; Baumann and Pham-Dinh, 2001; Emery, 2010) and become down-regulated around the time of myelination onset, including Notch signaling (Genoud et al., 2002; Givogri et al., 2002) and Wnt3A (Shimizu et al., 2005; Feigenson et al., 2009). When levels of these inhibitors decrease, transcription factors are systematically recruited to facilitate oligodendrocyte- specific gene expression (Gokhan et al., 2005). Olig1 and Olig2 are necessary for the   30 differentiation and maturation of oligodendrocytes (Lu et al., 2002) and Olig 1 is responsible for the upregulation of myelin proteins such as MBP and PLP (Xin et al., 2005). The transcription factor, myelin-gene regulator factor (MRF), is a major regulator of oligodendrocyte maturation and removing MRF can completely block myelination (Emery et al., 2009). In addition, neural cell adhesion molecule (NCAM) (Charles et al., 2000), Wnt (Fancy et al., 2009), glycine rich protein 17 (GPR17) (Chen et al., 2009) and neurite outgrowth inhibitor receptor interacting protein (LINGO) (Mi et al., 2005) play roles in regulating the timing of myelination onset.  In addition to threshold axonal diameter (Nave and Werner, 2014), oligodendrocyte myelination is regulated by electrical activity (Wake et al., 2011; Gibson et al., 2014), the spatial density of OPCs (Rosenberg et al., 2008), signals from local astrocytes (Back et al., 2005; Hammond et al., 2014) and immune-based cells (Ruckh et al., 2012; Miron et al., 2013; Yuen et al., 2013). Importantly though, myelin sheath formation can be initiated even on appropriately sized synthetic fibers suggesting that oligodendrocytes do not require dynamic signaling from the axon for myelination to begin (Rosenberg et al., 2008). Recent work suggests that oligodendrocyte myelination and myelin remodeling can continue well into adult life (Miller et al., 2012; Young et al., 2013) and the process appears to be regulated by experience both during development and adulthood (Liu et al., 2012). Crucially (and often under-appreciated), PLP knockout induces axonal degeneration, despite normal appearing oligodendrocyte myelination; this highlights the important role that oligodendrocyte myelination plays in maintaining axonal integrity (Griffiths et al., 1998).     31 1.5.2 Oligodendrocyte lineage cells in response to local CNS injury The most striking difference in injury-induced behaviour of myelinating cells in the PNS and CNS is their survival: in response to a local axonal injury in an adult CNS, 30-40% of oligodendrocytes die, in contrast to the near-complete survival of SCs in the PNS (Ludwin, 1990). As described above, oligodendrocyte loss after injury likely results from multiple events, including excitotoxicity, inflammatory cytokines released by microglia and infiltrating neutrophils, and vulnerability to oxidative stress resulting from ischemia (Almad et al., 2011). The cells that survive after injury do not have a major role in CNS repair; they are thought to be inactive or quiescent and cannot contribute to remyelination of axons (Blakemore and Keirstead, 1999; Almad et al., 2011; Crawford et al., 2016b). Therefore, the death of oligodendrocytes is thought to result in demyelination of local axons and associated damage to local axons. Neuronal/axonal damage following demyelination likely results from many factors, but insufficient oligodendrocyte-derived trophic support is thought to contribute (Goldberg and Barres, 2000).  Considerable debate remains regarding whether demyelinated axons persist long after injury (Totoiu and Keirstead, 2005; Lasiene et al., 2008; Powers et al., 2012; Powers et al., 2013; Plemel et al., 2014b). However, the data supporting persistent demyelination are often derived from rodent studies, where the spinal cord was examined in cross section; using this approach, it is difficult to differentiate between severed axons that persist near the lesion without myelin and axons that were spared but have become demyelinated. Experiments that involved tracing spared axons through the lesion found no evidence that a population of intact, chronically demyelinated axons remains following SCI, suggesting that endogenous myelin regeneration is highly efficient in   32 rodents (Lasiene et al., 2008; Powers et al., 2012). The persistence of chronically demyelinated axons after human SCI has only been examined in a handful of studies, in which it is generally noted that few demyelinated axons persist (Kakulas, 2004; Norenberg et al., 2004; Guest et al., 2005) except in cases of persistent cord compression, which may result in ongoing demyelination (Bunge et al., 1993; Guest et al., 2005). In contrast to the PNS, where myelin debris is removed within weeks of injury, myelin debris can be observed months and even years after CNS injury (Miklossy et al., 1991; George and Griffin, 1994; Becerra et al., 1995; Buss et al., 2004) and myelin-associated inhibitors can restrict axon growth (Grados-Munro and Fournier, 2003; Nash et al., 2009). Unlike SCs, oligodendrocytes maintain a modest upregulation of myelin proteins after injury (Bartholdi and Schwab, 1998; Jessen and Mirsky, 2008) and express very low levels of proteins in phagocytic pathways, suggesting they do not contribute to myelin uptake (Cahoy et al., 2008). In addition, because of the blood brain barrier in the CNS, macrophages may only access the immediate area of injury (i.e., where the blood brain barrier is disrupted) and may not have access to distal degenerating axon to assist with myelin clearance (Popovich and Hickey, 2001; Vargas and Barres, 2007). Unlike mature oligodendrocytes, OPCs that are resident in the CNS or derived from the subventricular zone do contribute to repair: they differentiate into oligodendrocytes, which are in turn capable of contributing to myelination after injury (Blakemore and Keirstead, 1999; McTigue et al., 2001; Zai and Wrathall, 2005; Horky et al., 2006; Lytle and Wrathall, 2007; Tripathi and McTigue, 2007; Lytle et al., 2009; Barnabe-Heider et al., 2010; Almad et al., 2011; Hesp et al., 2015). For example, local OPCs can proliferate, migrate, and replace oligodendrocytes destroyed by laser   33 microsurgery (Kirby et al., 2006).  Despite their capacity for differentiation and myelination, the repair derived from OPCs is incomplete, and does not reconstitute pre-injury levels of myelin, likely resulting in part from a general loss of axons.  It is also know that the efficiency of myelination after injury declines with age after injury (Crawford et al., 2013). It is still debated whether OPCs are strictly fate restricted to the oligodendrocyte lineage in vivo, as there is some evidence suggesting that these cells can become astrocytes and SCs following injury (Skoff, 1990; Fulton et al., 1992; Rivers et al., 2008; Kang et al., 2010; Zawadzka et al., 2010; Richardson et al., 2011; Tripathi et al., 2011). For more, see specific introduction to Chapter 2.   1.5.3 SC development SCs in the PNS arise from the neural crest, emerging from the dorsal aspect of the neural tube as it closes (Dupin et al., 2006). In embryonic development (E14-15 in mice), neural crest cells differentiate into SC precursors, which in turn give rise to immature SCs between E15 and E17. These immature SCs produce either myelinating or non-myelinating cells in the adult (Jessen and Mirsky, 2005; Woodhoo and Sommer, 2008). Importantly, the so-called SC precursors also produce a variety of other cell types, including endoneurial fibroblasts, melanocytes, parasympathetic neurons, and mesenchymal stem cells (Joseph et al., 2004; Adameyko et al., 2009; Nitzan et al., 2013; Dyachuk et al., 2014; Espinosa-Medina et al., 2014; Kaukua et al., 2014). Ensheathment of large groups of PNS axons can be observed at E18; true myelination is observed at birth.    34 The control of these differentiation points is governed by specific survival factors, mitogens, and differentiation signals from axons (Jessen and Mirsky, 1999). Using a combination of specific lineage progression markers and morphological characteristics, we can differentiate between SC stages (Jessen and Mirsky, 2005).  For example, SC precursors exclusively express cadherin-19 (CAD-19), myelinating SCs highly express P0 (amongst many other myelin markers), and non-myelinating SCs upregulate low-affinity nerve growth factor receptor (p75-NTR) (Jessen and Mirsky, 2005). Neuregulin1 (NRG1) - erythroblastic leukemia viral oncogene homolog (erbB) signaling an is an important regulator of myelin sheath thickness but also plays an important role in SC precursors survival and myelinating vs non-myelinating SC fate-decisions (Matthews, 1968; Michailov et al., 2004; Jessen and Mirsky, 2005; Taveggia et al., 2005). Migration through radial sorting is regulated by many different signaling including NRG1, IGF, neurotrophin-3 (NT-3) and BDNF (Cheng et al., 2000; Meintanis et al., 2001; Yamauchi et al., 2004) while SC proliferation appears to be dependent on axonal mitogens such as NRG1 (Komiyama and Suzuki, 1992), and transforming growth factor b (TGFb) (Jessen and Mirsky, 2005) or interactions with laminin (Yang et al., 2005; Yu et al., 2005). Sex determining region Y-related high mobility group-box10 (Sox10), octamer-binding transcription factor (Oct6) and early growth response protein 2 (EGR2; also known as Krox20) are essential components of the regulatory network in SCs (Stolt and Wegner, 2016). In addition, two death signals have been identified including p75-NTR and TGFb (Syroid et al., 2000; Jessen and Mirsky, 2005). There is also evidence inhibitory signaling pathway may be active in immature SCs to avoid the onset of myelination  (Jessen and Mirsky, 2005). The mechanisms that underlie the transition to a myelinating   35 SC are not entirely clear; it seems to require laminin as the knockout laminin and/or its receptors produces a deficit in myelination (Xu et al., 1994; Feltri et al., 2002; Pietri et al., 2004; Yang et al., 2005; Yu et al., 2005). In the adult PNS, more than 90% of the cell bodies in the endoneurial space are SCs (Campana, 2007). While they are classified as myelinating SCs and non-myelinating SCs (Jessen and Mirsky, 2005), all SCs have the ability to myelinate local axons (Aguayo et al., 1976) if they receive the correct axonal signals (Taveggia et al., 2005). In addition, SCs can take on diverse phenotypes depending on the axons they associate with: for example, SCs can have a modality-based phenotype (either motor or sensory), and preferentially facilitate regeneration of axons with a matching modality (Hoke et al., 2006).  Myelinating SCs wrap axons at a 1:1 ratio and undergo radial sorting prior to birth with some evidence that reorganization continues postnatally (Woodhoo and Sommer, 2008); SCs are spaced at every 1mm along the length of the axon (Campana, 2007). Under the control of several transcription factors, including Sox10, Oct6, brain 2 (Brn2), and Krox20, myelinating SCs upregulate membrane lipids and myelin proteins such as MAG, peripheral myelin protein 22 (PMP-22), MBP, and P0 (Nagarajan et al., 2002; Svaren and Meijer, 2008; Pereira et al., 2012). Krox20 is thought to be the master regulator for SC myelination and is the transcription factor required for SC myelination and ongoing myelin maintenance. Krox20 expression stimulates precursor cells to exit the cell cycle, activates myelin genes (PMP-22, P0, MBP), and suppresses myelination inhibitors (Topilko et al., 1994; Decker et al., 2006; Jessen and Mirsky, 2008; Mirsky et al., 2008).   36  1.5.4 The SC response to PNS injury The SC response to injury is thought to be critical for efficient regeneration in most contexts and is likely critical to functional recovery after PNS injury (Gaudet et al.; Vargas and Barres, 2007). SCs respond to nerve injury rapidly: the myelinating SCs of the distal injured axon survive (Ludwin, 1990) and even prior to the onset of axon degeneration, they proliferate (Jessen and Mirsky, 2005; Vargas and Barres, 2007), and undergo gene expression changes that facilitate their detachment from the axon (Liu et al., 1995; Guertin et al., 2005). Myelinating SCs stop production of myelin proteins and both non-myelinating and myelinating SCs upregulate genes that are pro-regenerative and promote axonal growth (Pellegrino et al., 1986; Trapp et al., 1988; White et al., 1989; Funakoshi et al., 1993). The SCs align along the preexisting basal lamina tubes, known as bands of Büngner, and support growing axons through both substrate-based guidance and trophic cues (Stoll et al., 1989; Griffin and Thompson, 2008). For example, SCs in the injured nerve upregulate many neurotrophins, including glial cell line-derived neurotrophic factor (GDNF), artemin, BDNF, neurtrophin-4/5 (NT-4/5) and nerve growth factor (NGF) (Funakoshi et al., 1993; Jessen and Mirsky, 2008; Arthur-Farraj et al., 2012) while ciliary neurotrophic factor (CNTF) and NT-3 are decreased (Funakoshi et al., 1993; Anand et al., 1997; Jessen and Mirsky, 2008). They also secrete basal lamina components, consisting of laminin, type IV collagen and heparin sulfate proteoglycans (HSPGs), which are crucial for myelination (Bunge et al., 1990). The clearance of myelin debris is an important aspect of successful PNS regeneration (Brosius Lutz and Barres, 2014). SCs have the ability to phagocytose their own myelin debris early after injury   37 (Perry et al., 1995) in an autophagic process recently described as myelinophagy . In addition, SCs release cytokines/chemokines, including TNF-a, leukemia inhibitory factor (LIF), IL-1 a, IL-1b, IL-6 and macrophage chemoattractant protein 1 (MCP-1) as well as galectin-3 (Reichert et al., 1994; Rotshenker, 2011). These factors play a role in the recruitment of monocytes/macrophages from the blood (Subang and Richardson, 2001; Tofaris et al., 2002; Shubayev et al., 2006; Vargas et al., 2010; Gaudet et al., 2011) into the basal lamina tubes, which facilitates debris clearance (Dailey et al., 1998; Vargas et al., 2010). Further research is required to better understand these mechanisms of myelin clearance which are crucial for peripheral nerve regeneration and why these fail in the CNS.  Initial investigations of the injury response suggested that SCs dedifferentiated from their mature state to contribute to repair (Jessen and Mirsky, 2005) . However, more recent research suggests that mature SCs adopt a pro-reparative phenotype in response to injury, and this cell is similar to but distinct from an immature SC (Vargas and Barres, 2007; Jessen and Mirsky, 2010). This transformation is regulated by c-jun activation and Raf/Erk signaling (Arthur-Farraj et al., 2012; Napoli et al., 2012; Jessen and Mirsky, 2016); when c-jun was inactivated in SCs, functional recovery was impaired after nerve injury, and there was a dramatic increase in neuronal death (Arthur-Farraj et al., 2012). There is evidence that the repair state of SCs might be different amongst sensory and motor SCs (He et al., 2012). After SCs contribute to this positive repair environment by providing trophic support and substrate for axons to re-grow along, they can re-differentiate by reinstating mature SC gene expression and contribute to   38 ensheathment/myelination of regenerating axons (Fawcett and Keynes, 1990; LeBlanc and Poduslo, 1990; Stoll and Muller, 1999; Mirsky et al., 2008). Despite the pro-regenerative response of SCs, spontaneous axonal regeneration, while effective in the PNS compared to the CNS, is far from ideal (Hoke, 2006). Predictably, PNS injuries have poorer outcomes when there is a large gap between the injury site and the innervation target (Fu and Gordon, 1995a; Hood et al., 2009) which can result in incomplete or misdirected regeneration which is often accompanied by neuropathic pain (Sunderland, 1947). Long distance regeneration is slow with a growth rate of 1mm/day (Sunderland, 1947) and often results in chronic prolonged denervation (Weinberg and Spencer, 1978; Pellegrino and Spencer, 1985; Fu and Gordon, 1997) resulting in failure of the PNS regenerative process (Fu and Gordon, 1995b). Over time post-injury, SCs lose their ability to proliferate, migrate and support axonal regeneration and eventually atrophy (Fu and Gordon, 1997; Li et al., 1997; You et al., 1997; Hall, 1999) when not in contact with axons.  1.5.5 The SC response to CNS injury SCs are not observed in the CNS under normal (i.e., non-pathological) conditions. The boundaries of the CNS are defined by the glia limitans, which consists of astrocyte processes covered by basal lamina that lies below the pia and around blood vessels (Fraher, 1992). The glia limitans delineates the PNS from the CNS where nerve roots enter and exit the spinal cord and it has been argued that the presence of astrocytes on the central side of the transitional zone is what prevents SCs from invading the CNS (Fraher, 1992). Following SCI, a substantial process of remodeling occurs (Beattie et al., 1997),   39 resulting in a lesion site that lacks normal CNS cytoarchitecture. In regions that lack GFAP+ astrocytes, SCs have been observed in both in animal models and human SCI (Bresnahan, 1978; Bunge, 1994a; Bunge et al., 1994; Hill et al., 2001; Guest et al., 2005). In addition, SCs have been found in the spinal cord following the injection of chemical demyelinating compounds (i.e., ethidium bromide or lysolecithin) (Blakemore, 1975, 1976; Graca and Blakemore, 1986), in regions of demyelination in experimental autoimmune encephalomyelitis mice (Raine et al., 1978), and in multiple sclerosis (MS) plaques in patients (Ghatak et al., 1973; Itoyama et al., 1983; Itoyama et al., 1985). In general, SCs are uncovered in the CNS whenever there is a loss of both astrocytes and oligodendrocytes in a particular region (Blakemore, 1975; Shields et al., 2000) and this process appears to be a natural spontaneous response in the injured and demyelinated CNS. SCs have been located in the spinal cord as early as a week or two post-injury in animal models of SCI (Beattie et al., 1997; Brook et al., 2000; Bruce et al., 2000) but are generally not observed in human tissue until several week or even months post-injury (Kakulas, 1984; Buss et al., 2007). In human cadaveric tissue, approximately 85% of samples from humans more than 4 months post SCI contained SCs (Bruce et al., 2000; Norenberg et al., 2004) and SCs have been observed years and even decades after SCI (Kakulas, 1984; Bunge et al., 1993; Bruce et al., 2000; Guest et al., 2005; Buss et al., 2007).  For many years it was assumed that SCs found in the CNS must have migrated in via perivascular nerves, meningeal nerves, and/or the spinal roots (Franklin and Blakemore, 1993). In the spinal cord, this notion is particularly attractive, as trauma often results in the disruption of the glial limitans at nerve roots, thus providing an opening for   40 SC migration into the CNS (Franklin and Blakemore, 1993). In support of this, it was noted that SCs in the CNS are commonly observed near the dorsal roots and adjacent to blood vessels (Duncan et al., 1988; Fraher, 1992). However, this idea was challenged by evidence that SCs can be derived from CNS precursors in culture (Mujtaba et al., 1998; Crang et al., 2004) and following transplantation of those cells into demyelinated lesions in the CNS (Keirstead et al., 1999; Crang et al., 2004; Talbott et al., 2006). The latter findings led Blakemore and colleagues to suggest that the SCs that remyelinate demyelinated CNS axons may be generated from CNS precursors, specifically oligodendrocyte precursor cells (OPCs), the same cells that give rise to myelinating oligodendrocytes during development and under non-pathological conditions in the adult CNS (Blakemore, 2005).  Until very recently, it was impossible to definitively test Blakemore’s theory, but advances in genetic fate mapping of cells have made it possible to trace the origin of the SCs found in the CNS. Franklin and colleagues (Zawadzka et al., 2010) used inducible reporter expression to label OPCs by administering tamoxifen to transgenic mice in which Cre recombinase is expressed under the PDGFRa promoter and drives the expression of yellow fluorescent protein (YFP). In this study, cells were defined as SCs based on the expression of the SC-specific transcription factor SCIP/OCT6, and/or 1:1 myelination of axons with the expression of the SC-specific protein, periaxin. The authors concluded that after focal demyelination, endogenous SCs are derived primarily from OPCs and not from peripheral myelinating SCs, ependymal cells, or parenchymal astrocytes (Zawadzka et al., 2010). The extent to which this process occurs following SCI   41 has never been addressed, and neither has the potential role of these OPC derived SCs in spontaneous and treatment related repair following SCI.  1.6 Transplantation of SCs as a treatment for CNS injury   In terms of the functional outcome of SCI, it is unclear whether endogenous SCs found in and near the injury site have beneficial roles or detrimental effects. While they may contribute to myelination and axon growth and guidance, SCs in the injured spinal cord can cause axons growing in meandering paths within the injury site, where they may make aberrant connections, which may contribute to pain and spasticity rather than useful function (Norenberg et al., 2004) a process coined “Schwannosis”. In animal models of SCI and MS, spontaneous remyelination by SCs has been suggested to improve conduction through axons, promote the growth of some CNS axons, and correlate with varying degrees on locomotor recovery (Blight and Young, 1989; Pender, 1989; Beattie et al., 1997; Jeffery and Blakemore, 1997; Brook et al., 1998; Jasmin et al., 2000; Murray and Fischer, 2001; Felts et al., 2005; James et al., 2011; Bartus et al., 2016). Compared to the CNS, the peripheral nerve environment in adult mammals shows far better axonal regeneration and functional recovery following injury (Johnson et al., 2005; Yiu and He, 2006). The inhibitory nature of the CNS appears to play an important role in the failure of CNS axonal regeneration, as CNS axons can grow long distances in peripheral nerve grafts, an environment known to be permissive to axonal regeneration (as described above) (Richardson et al., 1980; Aguayo et al., 1981). The presence of SCs is a major factor in the efficient axonal regeneration and functional recovery that occurs   42 in the PNS (Hall, 1986, 1989). In addition to promoting the regeneration of axons, SCs are also responsible for remyelinating regenerated axons, thereby facilitating functional recovery post-injury (Bixby et al., 1988).  One of the major potential advantages of using SCs, as opposed to other cell types, for transplantation to repair SCI relates to their suitability for autologous transplantation (in which patients can donate their own cells for transplantation). This approach circumvents any immune rejection of transplanted cells, and obviates the usual need for long-term (and harmful) immunosuppressive drug therapy following transplantation. Although it is possible to generate SCs from peripheral nerve for autologous transplantation, the invasive harvesting procedure for SCs involves permanent nerve damage, which is obviously an undesirable outcome given that the overall goal of this treatment is to return lost function following SCI. SCs have been considered for decades as a potential treatment for CNS injury (Bunge, 1975; Bunge, 2008)  and promising work focused on the use of peripheral nerve grafts and suspended nerve-derived Schwann after injury has led to some excitement specific to SCs as a potential treatment for SCI. In this section, I review the literature dedicated to testing peripheral cell grafts and nerve-derived SC transplantations into the CNS that has led to several SC-based clinical trials in the USA and discuss alternative sources of SCs including skin derived precursors differentiated into SCs (SKP-SCs).     43 1.6.1 Peripheral nerve grafts Peripheral nerve grafts transplanted into the CNS after injury can both promote the growth of spinal cord axons and contribute to ensheathment and myelination of host CNS axons (Richardson et al., 1980; David and Aguayo, 1981). Importantly, peripheral nerve grafts can support the growth of local sensory and propriospinal axons but they only seem to support the growth of supraspinal axons when the graft is located relatively close to the cell body of the growing axon (Aguayo et al., 1981; Richardson et al., 1982; Richardson et al., 1984; Oudega et al., 1994). Generally, of the axons that do grow into and across peripheral nerve grafts, many do not reach the distal graft-host interface and rarely do they cross the graft-host boundary to re-enter intact GFAP+ parenchymal tissue (Richardson et al., 1982; Oudega et al., 1994; Oudega and Hagg, 1996). Similarly, SCs from peripheral grafts ensheathe/myelinate growing axons within the grafts (David and Aguayo, 1981) but graft cell ensheathment/myelination is rarely observed near the distal graft-host interface, largely because SCs are typically reluctant to migrate into GFAP+ host parenchymal tissue (Fishman et al., 1983).  It has been repeatedly demonstrated that the peripheral nerve environment is conducive to CNS axonal growth, and that SCs are essential for these growth-supporting properties. This is evidenced in acellular nerve grafts: when resident SCs are killed but the growth supportive SC basal lamina remains, host CNS axon growth and ensheathment/myelination are compromised (Berry et al., 1988; Smith and Stevenson, 1988).     44 1.6.2 Transplantation of SCs after CNS injury Recognition of the importance of SCs to the regenerative capacity of peripheral nerves naturally led to the study of the effects of transplanting dissociated SCs into a variety of models of demyelinating diseases (Blakemore et al., 1986; Honmou et al., 1996; Iwashita and Blakemore, 2000; Kohama et al., 2001; Bachelin et al., 2005; Zujovic et al., 2012). SCs are typically harvested from the nerve (termed nerve-derived SCs) using an invasive surgical excision of peripheral nerve (Shields et al., 2000) followed by SC isolation and expansion in vitro prior to cell transplantation (Morrissey et al., 1991). Much of the work in SC transplantation work after SCI was spearheaded by Richard and Mary Bunge at the Miami Project for Cure Paralysis (Oudega and Xu, 2006; Fortun et al., 2009; Bunge and Wood, 2012; Wiliams and Bunge, 2012; Yang et al., 2015; Hosseini et al., 2016). Early work using SCs as a treatment for SCI focused on implanting synthetic guidance channels filled with SCs directly into a full transection site after injury in rodents (Xu et al., 1995b; Xu et al., 1995a; Chen et al., 1996; Xu et al., 1997; Xu et al., 1999b; Pinzon et al., 2001). Testing Schwan cell-seeded conduits in a full transection SCI model allowed researchers to reliably observe the growth of axons without the confound of differentiating between regenerating and spared axons. These studies yielded similar results to those observed with peripheral nerve grafting: sensory and propriospinal axons grew efficiently through these SC seeded channels but largely failed to re-enter host tissue at the distal interface (Xu et al., 1995b; Xu et al., 1995a; Chen et al., 1996; Xu et al., 1997; Fortun et al., 2009).    45 From this point, pre-clinical research focused on more clinically-relevant models of SCI, where researchers could test the histological and behavioral outcomes of different delayed transplantation regimes involving the transplantation of SCs into the contusion injury site of rodents. SC survival was improved when transplantation was delayed to 7 days post-injury and with the addition of immunosuppression; however, most of the transplanted cells were still lost within the first week through a combination of apoptosis and necrosis (Hill et al., 2006; Hill et al., 2007). As reviewed by Fortun and colleagues (2009), these studies have shown that SCs transplanted into the injured spinal cord consistently bridge lesion cavities, promote axonal regeneration and remyelination, and improve functional recovery over control treatments (Martin et al., 1991; Paino and Bunge, 1991; Martin et al., 1993; Li and Raisman, 1994; Paino et al., 1994; Martin et al., 1996; Xu et al., 1997; Imaizumi et al., 2000; Takami et al., 2002; Pearse et al., 2004a; Pearse et al., 2004b; Barakat et al., 2005; Fouad et al., 2005; Hill et al., 2006; Oudega and Xu, 2006; Pearse et al., 2007; Schaal et al., 2007; Fortun et al., 2009; Sharp et al., 2012b). Transplanted nerve-derived SCs can restore conduction across transection injuries localized to the dorsal columns (Imaizumi et al., 2000), facilitate subtle changes in locomotor recovery (Takami et al., 2002; Schaal et al., 2007; Sharp et al., 2012a), and may even be effective after delayed transplantation at 8 weeks post SCI (Barakat et al., 2005). Lastly, SCI transplantation work using genetically-labeled rat nerve-derived SCs demonstrated that the level of endogenous SCs in the injured spinal cord is increased substantially by nerve-derived SCs transplantation; this was the first indication that the endogenous SC response might play a substantial role in the repair/recovery observed after exogenous SC transplantation (Hill et al., 2006).    46 Although SC transplantation after SCI is thought to facilitate elements of neuroprotection, tissue preservation, remyelination, and axonal regeneration, when tested in pre-clinical animal models, only modest improvements in locomotor function have been observed (Fouad et al., 2005; Fortun et al., 2009; Tetzlaff et al., 2011; Bunge and Wood, 2012). Therefore, researchers have started to turn their attention towards using co-treatments that focus on improving cell survival after transplantation, increasing integration of transplanted SCs into the host parenchyma, augmenting trophic support to encourage supraspinal axon growth along SCs, and improving the ability of axons to grow back into host parenchyma (Oudega and Xu, 2006; Fortun et al., 2009; Bunge and Wood, 2012; Wiliams and Bunge, 2012).   1.6.3 Moving SC transplantation towards the clinic Importantly, transplantation of nerve-derived SCs derived from human nerves have similar benefits in terms of repair and functional recovery after transplantation into rodent models of SCI (Guest et al., 1997a; Pearse et al., 2007; Bastidas et al., 2017). These human cells can be harvested from patients via sural nerve biopsy for purification and expansion in vitro prior to transplantation (Morrissey et al., 1991; Casella et al., 1996; Guest et al., 1997b). The sural nerve is ideal, both because it is accessible and permits a large portion of nerve to be excised to provide an adequate number of cells for efficient expansion (Akiyama et al., 2002). Sural nerve harvesting is associated with decreased sensation for 16-34 years post-surgery in 76% of patients (Ijpma et al., 2006); in addition, it can result in abnormal or spontaneous sensations, and in rare situations, neuropathic pain (Schoeller et al., 2004). This invasive surgical excision of segments of   47 peripheral nerve, causing peripheral nerve injury, is a drawback to using nerve-derived SCs. Nerve-derived SC transplantation as a treatment for SCI is currently being tested in clinical trials. Work out of Iran suggests that nerve-derived SCs transplanted 2 years post SCI appear to be safe and are associated with modest improvements in light touch sensation, motor function (depending on whether the patients group sustained a cervical or thoracic SCI), and individual reports of improved bladder and bowel function (Saberi et al., 2008; Saberi et al., 2011). In the USA, the Miami Project to Cure Paralysis completed a Phase 1 clinical trial (NCT01739023; clinical where 6 patients received autologous nerve-derived SCs 1-2 months following a complete thoracic traumatic SCI. The group reported no surgical, medical, or neurological complications and no adverse events resulting from the SC transplantation, suggesting that the transplantation of autologous SCs in subacute SCI is generally safe (Anderson et al., 2017). The same group is currently enrolling patients in a Phase 1 safety trial specific to patients with chronic (>1-year post-injury) SCI (NCT02354625; clinical  1.6.4 Alternative sources of SCs Given that harvesting SCs from a peripheral nerve is invasive and risk-associated, researchers have investigated other potential sources of SCs. Embryonic stem (ES) cells, induced pluripotent cells (iPS), and reprogrammed somatic cells are potential sources of SCs but are high-risk options given that undifferentiated cells could contaminate cultures. In addition, it has been suggested that MSCs could differentiate into a SC like state in   48 culture (Kuroda et al., 2011). However, a more attractive alternative tissue source is adult mammalian skin, from which we can isolate 1) SKP-SCs; 2) epidermal neural crest stem cells (EPI-NCSCs) (Sieber-Blum et al., 2004); and 3) hair follicle pluripotent stem cells (Amoh et al., 2005). For this thesis, we will review the literature specific to skin derived precursors (SKPs) and our ability to differentiate them into a pure SC population, as they represent the safest and most well defined sources of SCs compared to above mentioned cells. SKP-SCs Our collaborators in Toronto (laboratory of Dr. Freda Miller) discovered SKPs in the dermis of both rodent and human skin (Toma et al., 2001; Toma et al., 2005) and demonstrated that these cells can be differentiated into SCs (SKP-SCs) in vitro (Toma et al., 2001; Fernandes et al., 2004; Toma et al., 2005; McKenzie et al., 2006). SKPs can be isolated from the whisker pads, as well as the dorsal and ventral trunk of rodents; in humans, SKPs have been isolated from the scalp, and the glabrous neonatal and juvenile foreskin (Toma et al., 2001; Toma et al., 2005; Hunt et al., 2008; Jinno et al., 2010; Kuroda et al., 2011). SKP-SCs can enhance PNS regeneration after chronic denervation (Walsh et al., 2010) and previous work in our laboratory demonstrated that SKP-SCs could survive transplantation and promote repair and functional recovery when delivered immediately following contusion SCI in rat (Biernaskie et al., 2007; Sparling et al., 2015). Transplantation of these cells showed similar benefits to nerve-derived SCs (based on studies conducted by other groups); however, the SKP-SCs appear to integrate better with host tissue (Fortun et al., 2009), raising the possibility that the SKP-SCs may   49 provide some benefits as a treatment for SCI that exceed their nerve-derived counterparts. In contrast to peripheral nerve biopsy, the harvest of SKPs involves a simple skin biopsy, which is much less invasive and carries little-to-no risk of causing deficits. Given the potential advantage of SKP-SCs over nerve-derived SCs in terms of integration with the injured CNS, and the reduced risk associated with the harvest of SKPs, it appears that SKPs may be a better source of SCs for clinical application in SCI than peripheral nerve. Therefore, I examined the potential for SKP-SC-mediated repair of chronic rodent SCI in Chapter 3 of this dissertation.  1.7 Overview of experiments and hypothesis  Aim 1: To determine the origin and extent of endogenous ensheathment/myelin based repair in response to contusion SCI (Chapter 2). Using transgenic mice, I labeled OPCs (PDGFRa+ and Olig2+ cell populations) and peripheral myelinating SCs (P0+ cell populations) prior to injury, and investigated their contribution to oligodendrocyte ensheathment/myelination and SC ensheathment/myelination in response to contusion SCI. Hypothesis 1: I hypothesized that PDGFRa+ cells, specifically OPCs, contribute to the majority of myelination observed in response to contusion SCI giving rise to both oligodendrocytes and SCs.     50 Aim 2: To investigate the therapeutic potential of transplanting exogenous SKP-SCs, in a clinically relevant scenario involving chronic contusion SCI in rats (Chapter 3). I transplanted labelled SKP-SCs into the 8 weeks post-injury contusion site and assessed functional outcome measures throughout and histological outcome measures at 21 weeks post-injury comparing SKP-SC treated rodents to controls.  Hypothesis 2: I hypothesized that SKP-SCs transplanted eight weeks after contusion SCI would survive, bridge the lesion site making it permissive for axon growth, contribute to remyelination, and promote functional recovery.                   51         Figure 1.1 The pathophysiology of rat contusion SCI These schematics depict the normal rat spinal cord and the rat spinal cord at subacute (defined as 1-2 wpi) and chronic (defined as 6-8 wpi) stages of contusion SCI. (a) The white matter of the normal spinal cord contains axons wrapped in myelin, while neuronal somata populate the grey matter. Glial cells, including OPCs, microglia, and astrocytes, are found throughout white and grey matter. (b) Spinal cord contusion in rat induces secondary injury cascades that cause progressive damage over time. At the subacute stage, considerable loss of neurons, axons, oligodendrocytes and myelin is apparent. Axons die back from the lesion site and myelin debris accumulates at the rostral and caudal borders of the lesion. Activated microglia, hematogenous macrophages, and other inflammatory cells are recruited to the lesion site. In addition, astrocytes become reactive, extending processes, proliferating, and (in collaboration with fibrotic cells) forming a scar. OPCs and other cells that express the NG2 also proliferate and are recruited to the scar. (c) In chronic stages of SCI, astrocytes and other cells thicken the scar, and a fluid-filled cavity typically forms at the lesion center. Activated microglia and macrophages persist within the lesion, though at reduced numbers compared to the subacute period. Some OPCs differentiate into new oligodendrocytes that produce myelin, and SCs are often found myelinating axons near or in the lesion epicenter.    52    53  Figure 1.2 Restoring neuronal connectivity following SCI via cell transplantation (a) Transplanted cells, such as SCs, can support axon growth by providing a bridge across the injury site. However, growing axons, which frequently have a tortuous appearance in the bridge, are often unable to re-enter host tissue at the distal graft-host interface. (b) Transplanted cells may improve functional connectivity through the formation of a relay. Such relays occur when host axons synapse onto transplant-derived neurons, which harbor a capacity to extend axons long distances from the injury and synapse with host neurons.      54 Figure 1.3 Regeneration of oligodendrocyte-derived myelin with cell transplantation Transplanting precursors of myelinating glia can result in the formation of new myelin sheaths. Transplanted OPCs or NSPCs compete with endogenous OPCs to remyelinate axons.           55 Chapter 2: Myelinogenic plasticity of oligodendrocyte precursor cells following spinal cord contusion injury  2.1 Introduction SCI causes both primary neural injury and a spreading wave of secondary tissue damage (Kwon et al., 2002a; Kwon et al., 2004; Norenberg et al., 2004). This secondary injury leads to oligodendrocyte death and axon demyelination, which may leave axons vulnerable to degeneration (Bresnahan et al., 1976; Blight, 1983b; Crowe et al., 1997; Totoiu and Keirstead, 2005; McTigue and Tripathi, 2008; Plemel et al., 2014b). Remyelination of axons in and around the SCI site is viewed as an important element of regeneration (Bunge et al., 1960; Bresnahan, 1978; Totoiu and Keirstead, 2005; Almad et al., 2011; James et al., 2011; Powers et al., 2012; Plemel et al., 2014b) and has provided a rationale for clinical trials involving transplantation of neural precursor cells (Cummings et al., 2005; Keirstead et al., 2005; Priest et al., 2015). To avoid the costs and safety concerns of transplantation, an alternative strategy may be to augment spontaneous remyelination after SCI. However, to do this, it is critical to determine the contributions of various endogenous progenitors to this regenerative process (Barnabe-Heider et al., 2010; Gregoire et al., 2015; Stenudd et al., 2015). The extent of oligodendrocyte remyelination following SCI has been quantified using indirect measures, based on the assumption that new myelin sheaths are thinner relative to the size of their associated axon (Blakemore, 1974). However, using internodal length to identify new myelin sheaths and axonal tracing to identify a specific tract after   56 SCI it was discovered that new myelin has only marginally reduced thickness in the injured rodent spinal cord (Lasiene et al., 2008; Powers et al., 2012; Powers et al., 2013). Thus, in the context of SCI, evaluating the extent of spontaneous remyelination based on thin myelin likely represents a considerable underestimation of repair; as such the true extent of spontaneous oligodendrocyte myelination following SCI remains unknown.  Myelinating oligodendrocytes are derived from OPCs, characterized by the expression of NG2 and PDGFRa (Nishiyama et al., 1996; Rivers et al., 2008; Kang et al., 2010; Young et al., 2013). Mice expressing inducible Cre recombinase (CreER) under control of PDGFRα promoter/enhancers have allowed for in vivo tracking of oligodendrocyte lineage cells (Rivers et al., 2008; Kang et al., 2010). For example, PDGFRa-expressing cells generate new myelinating oligodendrocytes as late as three months after SCI (Hesp et al., 2015). Given the persistence of OPC differentiation, it is particularly important to determine the magnitude of their contribution to remyelination after SCI.   In addition to oligodendrocytes, SCs have been shown to contribute to the myelination of axons after CNS damage, both in SCI (Bresnahan, 1978; Bunge et al., 1993; Guest et al., 2005), and in demyelinating lesions of the spinal cord (Blakemore, 1975). In these settings, SC myelination of spinal axons is predominately localized to areas of significant astrocyte loss (Itoyama et al., 1985). The prevailing view has been that SCs migrate into the damaged spinal cord from the PNS via spinal nerve roots, meningeal fibers, or autonomic nerves following breakdown of the glia limitans (Franklin and Blakemore, 1993). However, PDGFRa+ cells can also give rise to SCs following demyelinating chemical lesions (Zawadzka et al., 2010). The contribution of OPCs to   57 oligodendrocyte and SC myelination after a clinically relevant contusion SCI has not yet been determined using in vivo fate mapping techniques. Here, we systematically assessed the capacity of multiple cell types to form myelinating oligodendrocytes and SCs following contusion SCI. We demonstrate that PDGFRa+ OPCs contribute to approximately 30% of myelin sheaths surrounding axons in the vicinity of the lesion site 12 wpi. We further show that PDGFRa+ OPCs give rise to the majority of myelinating SCs found in the spinal cord after injury, with only a small contribution stemming from the P0+ peripheral SC population. These data reveal the diverse behavior of endogenous PDGFRa+ cells in response to SCI and reveal that they contribute substantially to myelin regeneration.   2.2 Materials and methods  2.2.1 Transgenic mice and cre induction Two lines of PDGFRα-CreERT2 mice, PDGFRα-CreERTM (I; Kang et al., 2010; #018280, Jackson Laboratories) and PDGFRα-CreERT2(II; Rivers et al., 2008), were crossed with Rosa26-eYFP (#006148; Jackson Laboratories) or the membrane tethered Rosa26-membrane tethered green fluorescent protein (mGFP)mT/mG) (#007576; Jackson Laboratories) reporter mice. In addition, Olig2-CreERTM (Takebayashi et al., 2002) and P0-CreERT2 (Leone et al., 2003) mouse lines were individually crossed with the Rosa26-eYFP reporter mouse (#006148; Jackson Laboratories). PDGFRa+ cells for in vitro experiments were isolated from PDGFRa:H2B-GFP mice (Hamilton et al., 2003)   58 (#007669, Jackson Laboratories) via flow cytometry (fluorescence activated cell sorting; FACS). An overview of the transgenic mice used is shown in Table 2-1 and Table 2-2. All experiments were carried out in accordance with protocols approved by the UBC and University of Calgary animal care committees and the Canadian Council on Animal Care. Mice of either sex were group housed (2-6 mice/cage) in secure conventional rodent facilities on a 12-hour light/dark cycle with constant access to food and water. Cre-mediated recombination was induced at 8-10 weeks of age via intraperitoneal tamoxifen (Sigma, T5648) dissolved in corn oil at 20mg/mL. PDGFRα-CreERTM(I):Rosa26-eYFP, PDGFRα-CreERTM(I):Rosa26-mGFP(mT/mG), PDGFRα-CreERT2(II):Rosa26-eYFP, Olig2-CreERTM:Rosa26-eYFP, and P0-CreERT2:Rosa26-eYFP mice received 3mg of tamoxifen per day for 5 consecutive days; PDGFRα-CreERT2(II):Rosa26-mGFP(mT/mG) received only 0.5mg of tamoxifen for 2 consecutive days. Tamoxifen free controls (corn oil only injections) were run in all mouse lines both in injured and non-injured mice.  2.2.2 Surgical procedures After the final day of tamoxifen injection, we allowed two weeks for tamoxifen clearance following the final tamoxifen injection before mice received a SCI. Both 1 and 2 week clearances in the PDGFRα-CreERTM (I):Rosa26e-YFP mice were tested; results were qualitatively similar for both clearing intervals. All spinal cord, dorsal root, and sciatic nerve injuries, as well as the harvesting of dorsal roots and sciatic nerves from the PDGFRα:H2B-GFP mice, were performed at 10-12 weeks of age.   59 SCI Thoracic contusion SCI was delivered with the Infinite Horizons Impactor (Precision Systems Instrumentations). Animals were anaesthetized using isofluorane (4% induction, 1.5% maintenance) and received buprenorphine (Temgesic®; 0.02 mg/kg, s.c., McGill University) pre-operatively. After the skin at the surgical site was shaved, cleaned, and disinfected, the animal was secured in a stereotaxic frame on a warming blanket; body temperature was maintained at 36.5°C. The spinal cord was exposed via a midline incision in the skin and superficial muscles, and blunt dissection of the muscles over the T8–T11 vertebrae. The spinal cord was exposed via a T9-T10 laminectomy and the vertebrae were stabilized with clamps. The impactor delivered a 70 kdyne midline contusion injury. After injury, the muscle and skin were closed with continuous, 5-0 Vicryl sutures, and interrupted 4-0 Prolene sutures, respectively. Animals received buprenorphine and lactated Ringers (1ml, s.c.) every 12 hours for 48 hours following SCI. Bladders were expressed twice daily until spontaneous micturition returned (approximately 1-2 weeks after SCI). A subset of animals, received 5-ethynyl-2’-deoxyuridine (EdU) (Invitrogen, A10044) daily (50mg/kg, i.p.) for the first two weeks following SCI. Injured mice were perfused at 1 wpi, 3 wpi, and 12 wpi.  In addition, ‘uninjured controls’ were perfused 2 weeks post tamoxifen when the injury would have been sustained and ‘uninjured age matched controls’ were perfused alongside the 12 wpi group to assess cell formation at these time points in the absence of injury.    60 Root and nerve injury We performed a dorsal root crush or a sciatic nerve crush in PDGFRα-CreERTM(I):Rosa26-mGFP(mT/mG) mice. As for SCI, animals were anaesthetized using isofluorane and received buprenorphine pre-operatively. The skin at the surgical site was shaved, cleaned, and disinfected. For sciatic nerve crush injury, a small proximal-to-distal incision was made in the skin immediately posterior to the left femur. The sciatic nerve was exposed and crushed twice 3mm distal to the obturator tendon for 15 seconds with #5 fine Dumont forceps. Similarly, for cervical dorsal root crush injury, the left C5 and C6 roots were exposed via a lateral hemilaminectomy and durotomy, and crushed twice for 15 seconds. Mice were perfused 2 weeks post tamoxifen and termed ‘uninjured controls’ and injured mice were perfused at either 4wpi or 12 wpi.  2.2.3 Tissue preparation and immunohistochemistry Mice were deeply anaesthetized and transcardially perfused with PBS followed by cold 4% paraformaldehyde. The spinal cord, dorsal root, and sciatic nerves were harvested and post-fixed in 4% paraformaldehyde (2 hours post-fixation for dorsal and sciatic nerves; 6 hours for spinal cord) before cryoprotection in a series of increasingly concentrated sucrose solutions. Tissue was frozen in OCT Embedding Compound (Tissue-Tek; Electoron Microscopy Sciences, PA USA), sectioned in either the longitudinal or cross-sectional plane (as stated in figure legends) at 20µm thickness on a cryostat, and stored at -80 °C. Prior to immunohistochemistry, frozen sections were thawed, then rehydrated in phosphate buffer solution (PBS), and incubated in 10% donkey serum dissolved in PBS with 0.1% Triton. Prior to immunohistochemistry   61 targeting myelin, delipidization was performed using ascending and descending ethanol washes.  Primary antibodies (Table 2-3) were applied overnight at room temperature, followed by application of the appropriate Dylight or Alexa Fluor secondary antibodies (Jackson ImmunoResearch Labratories Inc.) for two hours. In some sections, nuclei were labeled with Hoechst 33342 (1:1500). To visualize EdU, the Click-iT® EdU Alexa Fluor® 647 (Invitrogen, C10340) Imaging Kit was used according to the manufacturer’s instructions.  2.2.4 FACS and immunocytochemistry Sciatic nerve segments (5mm distal to the sciatic notch) and dorsal root/dorsal root ganglia (DRGs; distal to dorsal root entry) were harvested from PDGFRα:H2B-GFP mice (4 independent experiments with 2-5 mice pooled per experiment) at 10-12 weeks of age. Tissue was finely chopped then incubated at 37oC for 30 min in collagenase Type-IV (Sigma, 1mg/ml) followed by trituration (4x every 10 min) until a single cell suspension was obtained. Cells were resuspended in 0.1% bovine serum albumin (BSA)/Hank’s balanced salt solution and passed through a 40µm filter, and GFP+ and GFP-negative (NEG) cells were sorted using FACS (BD FACS Aria III, pressure 20 PSI, nozzle 100µm). Sorted cells were collected and expanded for 1-2 weeks on coated poly-D-lysine/laminin (20µg/ml) plates/slides, in SC proliferation/ differentiation media (1% horse serum, 1% penicillin/streptomycin, 2% N2, bovine pituitary extract [20ng/ml], neuregulin [10ng/ml], forskolin [5mM], L-glutamine in Dulbecco’s modified Eagle’s medium (DMEM)/ F12). For immunocytochemistry, cells were fixed in 2% paraformaldehyde for 5 minutes, then incubated with 0.5% TritonX and 5% BSA for 1   62 hour. Primary antibodies (Table 2-3) were applied overnight at 4°C, and appropriate Alexa Fluor secondary antibodies (Jackson ImmunoResearch Laboratories, Inc) were applied for 2 hours at room temperature followed by Hoechst deoxyribonucleic acid (DNA) stain (1:2000, 5 minutes). Images were captured on an Axio Observer inverted light microscope (Zeiss) or an Observer Fluorescence microscope (40x objective, Zeiss).   2.2.5 Cell counting and analysis  Imaging and cell counts were performed on a Zeiss AxioObserver.Z1 inverted confocal microscope equipped with a Yokogawa spinning disk and Zen 2012 software. Investigators were blinded to the animal identity (i.e., uninjured control or post-injury time point). For all analyses, quantifications were completed on cross-sectional tissue and the significance was measured using the Kruskal-Wallis (KW) test with a follow up Mann-Whitney U (MWU) test and considered significant if p values= < 0.05. All data are reported as means +/- standard error (SEM). For assessing recombination efficiency in OPCs in the uninjured spinal cord, the PDGFRα-CreERTM(I):Rosa26-eYFP and PDGFRαCreERT2(II):Rosa26mGFP(mT/mG) T9/10 mouse spinal cord sections were processed immunhistochemically for GFP, PDGFRα, and Olig2 (n=3/group). Recombination efficiency percentages were reported as follows (% = [# of non-vascular associated GFP+ PDGFRa+ Olig2+/ # of non-vascular associated PDGFRa+ Olig2+] *100). An average of 150 cells was counted per mouse.  For assessing oligodendrocyte lineage cell counts in PDGFRα-CreERTM(I):Rosa26-eYFP mice, we investigated either uninjured mice (uninjured 12 wpi age matched control; n=3) or mice post SCI (3 wpi, n=4; 12 wpi, n=6) using sections   63 stained for Olig2 (nuclear marker specific to the oligodendrocyte lineage), PDGFRa (OPCs), CC1 (post-mitotic oligodendrocytes) and YFP (recombined cells). A blinded observer determined the lesion epicenter by selecting the section with the least spared tissue based on axonal staining on another slide. Systematic random sampling was conducted in a single section (20x) at the epicenter or an equivalent T9/10 section for uninjured 12 wpi age matched controls. This systematic random sampling included placing a large grid over the entirety of the cross-section at injury epicenter and randomly selecting a coordinate in each 3 X 3 sub-gridded area and imaging all the appropriate boxes within these coordinates. Boxes were equally spaced apart with 400µm between boxes in any direction to ensure adequate sampling. All counts were conducted within optical dissector boxes with an area of 20000 µm2 each and cells were required to be Olig2+ to be included in the analysis. We then went on to count the number of Olig2+ cells that were PDGFRa+, CC1+, and/or YFP+. Percentages were calculated by taking the density of the cell of interest and dividing it by the density of the total population (clearly demonstrated in y-axis of graphs).  For examining oligodendrocyte myelin in PDGFRα-CreERT2(II): Rosa26-mGFP(mT/mG) mice, we investigated either uninjured mice (uninjured 12 wpi age matched control; n=3) or mice post SCI (3 wpi, n=4 and 12 wpi, n=5) using sections stained for β3-Tubulin, neurofilament-200 (NF-200), and Sternberger monoclonal incorporated antibody-312 (SMI312) collectively (to visualize axons), MBP (myelin), P0 (peripheral myelin) and GFP (recombined cells). A blinded rater determined the lesion epicenter as described above. High-power (63x primary magnification) systematic random sampling was conducted in a single section at the epicenter or an equivalent   64 T9/10 section for uninjured 12 wpi age matched controls. This systematic random sampling (as described above) used optical dissector boxes equally spaced apart with a distance of 230µm between boxes in any direction to ensure adequate sampling. The number of MBP+ axons were counted and then of those axons, the proportion of GFP+/NEG sheaths around axons were counted. Additionally, we counted axons that were surrounded by GFP but were MBPNEG. All counts were conducted within optical dissector boxes with an area of 400µm2 each and extrapolated to the entire cross section of the cord. Ensheathed/myelinated axons were only counted when a mGFP+ tube (new ensheathment/myelin) or MBP+ tube (myelin marker) entirely surrounded an axon profile (collectively: β3-Tubulin+, NF-200+, and SMI312+). Total MBP+P0NEG axons at epicentre were counted. Percentages of new mGFP+MBP+ axons at epicentre were calculated as follows: (mGFP+MBP+P0NEG/MBP+P0NEG)*100. We then calculated the number of GFP+ tubes surrounding axons (split into the MBP+ or MBPNEG ratios) as follows: (GFP+MBP+P0NEG) + (GFP+MBPNEGP0NEG). In order to examine the contribution of PDGFRα+ cells to SC remyelination after SCI, spinal cord sections from PDGFRα-CreERTM(I):Rosa26-eYFP were processed immunohistochemically for β3-Tubulin, NF-200, and SMI312 collectively (to visualize axons), P0 (peripheral myelin) and YFP (recombined cells). High-power (63x primary magnification) cross-sectional images at lesion epicenter (as defined above) were captured (Week 12 uninjured; n=4; 3wpi, n=7; 12wpi, n=7), and P0+/YFP+/NEG myelin sheaths were counted.   To examine the contribution of P0+ cells to SC remyelination after SCI, spinal cord sections from P0-CreERT2:Rosa26-eYFP mice were processed   65 immunohistocchemically for β3-Tubulin, NF-200, and SMI312 collectively, P0 and YFP. High-power (63x) images were captured to encompass the spinal cord section at the lesion epicenter (Week 12 uninjured control, n=4; 1wpi, n=5; 3wpi, n=6; 12wpi, n=7), and P0+/YFP+/NEG myelin sheaths were counted.  In addition, recombination efficiency was assessed by counting the number of P0+ tubes that were co-labeled with YFP in three consecutive T9/T10 dorsal and ventral root sections in uninjured control P0-CreERT2:Rosa26-eYFP mice (n=4). An average of 600 cells were counted per uninjured mouse.   2.3 Results 2.3.1 Genetic fate mapping identifies PDGFRa progeny in the adult spinal cord                                             We systematically assessed the ability of different cell types to form myelinating cells in response to SCI using six transgenic mouse lines. Due to the inducible nature of Cre recombinase by tamoxifen in all mouse lines used, cells were labelled prior to injury with no additional recombination taking place after SCI. Animals were allowed to survive for three or twelve weeks after SCI. The first goal of our study was to obtain an estimate of the amount of new myelin formation following SCI and determine its cellular origin. As it is widely accepted that oligodendrocytes are produced via the differentiation of OPCs, we performed OPC fate mapping with two independent mouse lines driving the expression of a tamoxifen-inducible Cre recombinase under the PDGFRa promoter: PDGFRα-CreERTM mice, ((Kang et al., 2010) denoted throughout as PDGFRα-CreER[I]) and PDGFRα-CreERT2 mice ((Rivers et al., 2008) denoted throughout as PDGFRα-CreER[II]).    66  We examined the extent and identity of recombined cells (defined as cells that were PDGFRa promoter active cells at the time of tamoxifen dosing and hence positive for YFP or GFP) in the adult uninjured control PDGFRα-Cre:YFP or mGFP mice. Tamoxifen was administered at 8-10 weeks of age, and recombination in the spinal cord with the attached dorsal roots was examined 14 days later (Figures 2.1 and 2.2). In all four mouse lines, tamoxifen induced Cre activation induced abundant and robust expression of YFP or mGFP. The PDGFRα-CreER(I):Rosa26-mGFP(mT/mG) and PDGFRα CreER(II):Rosa26-eYFP mouse lines were investigated to confirm observations made in separate mouse lines but no quantification was performed in these mice. Oligodendrocyte lineage cells are identifiable by characteristic protein expression: Olig2 is expressed in both OPCs and mature oligodendrocytes, while PDGFRα and NG2 are expressed in OPCs, and CC1 and MYRF are specific to oligodendrocytes (Kitada and Rowitch, 2006; Emery et al., 2009; Bujalka et al., 2013). Example images taken two weeks after tamoxifen induction—at the time when the injury would have been inflicted—demonstrate GFP co-labeling with OPC markers PDGFRa/olig2 (Figure 2.1 a-c) and NG2 (Figure 2.1 d), in cells that have a typical OPC morphology visualized with the membrane-tethered reporter (Figure 2.1 e, f, g). The recombination efficiency at the time of injury (number of non-vascular GFP+ PDGFRa+ cells divided by the total number of non-vascular PDGFRa+ cells) in the spinal cord of PDGFRα-CreER(I):YFP and PDGFRα-CreER(II):mGFP mice was 85 ± 2% and 69 ± 1%, respectively.   In addition to cells in the oligodendrocyte lineage, tamoxifen-induced recombination was also observed to a lesser extent in other PDGFRa-expressing cells in   67 both PDGFRa-CreER:YFP or mGFP mice lines. Within the CNS, we encountered recombination in vascular-associated cells (Figure 2.2 a-e). Blood vessel-associated cells expressing YFP were located on the outside of the endothelial layer (delineated by RECA and glucose transporter-1 (Glut1; Figure 2.2 a & b) and on the inside of the outer basal lamina (Figure 2.2 c), consistent with the location of pericytes. The majority of YFP+ cells in this region expressed PDGFRa and PDGFRb (Figure 2.2 d) which classifies them as type A pericytes (Goritz et al., 2011). A second small subset of YFP+ vascular-associated cells appeared to also express the vascular-associated cell marker asmooth muscle actin (aSMA; Figure 2.2 e; defined as type B pericytes by Goritz et al., 2011). We occasionally observed recombination in some central canal-associated cells, whose cell bodies were located immediately outside of the ependymal layer with a process extended towards the lumen of the canal (Figure 2.2 f, g).   In the PNS, recombined cells in PDGFRa-CreER:YFP or mGFP mice were found in the dorsal root (Figure 2.2 h), DRG (Figure 2.2 i), and sciatic nerve (Figure 2.2 j). These GFP+ cells in the PNS are endoneurial cells and expressed fibronectin (Fig. 2.2 k). YFP+ vascular-associated cell bodies were also encountered in the PNS (e.g., Figure 2.2 l), with associated processes encircling blood vessels in the dorsal root. Importantly, recombined cells in the dorsal root did not co-express the myelinating SC marker P0, or p75-NTR which is typically expressed in non-myelinating SCs (Figure 2.2; i, j, l, m).     68 2.3.2 PDGFRa+ cells, recombined prior to injury, contribute to new oligodendrocyte ensheathment/myelination after SCI                                   To estimate the amount of new ensheathments/myelin formed by endogenous oligodendrocytes after SCI, we performed lineage tracing of OPCs in PDGFRα-CreER:YFP or mGFP mice recombined prior to injury (Figure 2.3 a). Twelve weeks after SCI, YFP+ cells had retained EdU (Figure 2.3 b), indicating that they proliferated in response to the trauma. In contrast to the uninjured scenario, where the majority of recombined cells co-expressed PDGFRa and NG2 (indicating they are OPCs; Figure 2.1), most of the recombined cells at 12 weeks after SCI co-expressed CC1 (identifying them as oligodendrocytes; Figure 2.3 c & d). In mGFP mice (i.e., carrying the membrane-tethered reporter), recombined cells with the morphological features of both OPCs and mature oligodendrocytes were highly enriched around the injury site, and these cells were also encountered at great distances from the injury epicenter. These morphological cellular features of recombined cells correlated with immunohistochemical profiling: OPCs characterized by ramified processes and the expression of PDGFRa (Figure 2.3 e) and a larger percent of mature oligodendrocytes with tube formation and rostro-caudally aligned processes also expressed the transcription factor MYRF which is essential for myelination (Figure 2.3 f) (Emery et al., 2009; Koenning et al., 2012). Crucially, the expression of the contactin-associated protein (Caspr) in the axonal-glial contacts of the paranodes could be easily co-labelled with mGFP+ tubes, suggestive of oligodendrocyte ensheathment and ongoing myelination (Figure 2.3 g).  To quantify the extent of new oligodendrocytes formed after SCI, we  performed analysis in PDGFRα-CreER(I):YFP mice and co-immunostained Olig2+ cells for either,   69 PDGFRa (OPCs) or CC1 (oligodendrocytes), and counted their total number and their number also staining for  YFP+ (recombined cells). Recombined YFP+ cells co-labelling  with Olig2 and PDGFRa represent OPCs and co-labelling with Olig2 and CC1 is indicative of differentiation into oligodendrocytes. There was no difference observed following injury in the density of total OPCs (PDGFRa+Olig2+ cells) nor in the recombined subpopulation of OPCs (YFP+PDGFRa+Olig2+; Figure 2.3h). In contrast, there were significantly more oligodendrocytes at 12 wpi compared to the 3 wpi (Figure 2.3 i). Presumably this increase in new oligodendrocytes was due to OPC differentiation into new oligodendrocytes as there were more new oligodendrocytes (YFP+CC1+Olig2+) at 12 wpi compared to the 12-week uninjured control group (Figure 2.3 i). We found that oligodendrocytes continued to differentiate between 3 and 12 wpi as there was a higher percentage of recombined new oligodendrocytes (83%) compared to 3 wpi (65%) and the week 12 uninjured groups (49%; Figure 2.3 j). Reciprocally, the 12 wpi group showed the lowest percentage of recombined OPCs (17%) compared to the 3 wpi (35%) and week 12 uninjured groups (51%; Figure 3j) in the recombined oligodendrocyte lineage (YFP+Olig2+). Further, our data reveal that there is a 5-fold increase in the number of new oligodendrocytes at 12 wpi (13929+/-1356) compared to the 12 week uninjured control group (2978+/- 1031) and that 53% of the epicenter oligodendrocytes at 12 wpi are from new YFP+ oligodendrocytes (Figure 2.3 i). Collectively, these data demonstrate that following SCI there is considerable oligodendrogenesis from PDGFRα+ OPCs recombined prior to injury, and this is prolonged after SCI.   70 In order to determine the extent of ensheathment/myelination by recombined PDGFRa+ cells, we performed quantitative analyses in PDGFRα-CreER(II):mGFP mice using cross-sections of the spinal cord from animals 3 or 12 weeks after SCI and uninjured age-matched controls (Figure 2.4). At 3 and 12 wpi, mGFP+ cells could be visualized extending processes that surrounded nearby axons but only some of these mGFP+ tubes co-labelled with MBP (Figure 2.4 a & b).  Following a moderate-severe contusion SCI, there was a ~75% reduction in the number of MBP+ myelin sheaths remaining at 3 and 12 weeks after SCI compared to the uninjured age matched spinal cord (Figure 2.4 c). By 3 weeks post-SCI there were newly generated oligodendrocyte-derived myelin sheaths (i.e., in mGFP+, MBP+, P0NEG cells); the production of new oligodendrocyte-derived myelin sheaths continued as the percentage of these newly generated sheaths increased by 12 wpi (Figure 2.4 d). The percentage of new myelin produced by oligodendrocytes accounted for about 20% of the total myelinated axons at the epicenter (i.e. mGFP+) by 12 wpi. This percentage of newly myelinated axons represents an underestimate, because recombination efficiency in this mouse line was only 68% (calculated during initial mouse characterization and discussed above). When we account for this recombination efficiency we estimate that de novo myelin generation at injury epicenter by 12 weeks after SCI approaches 30%. To our surprise, even at 12 wpi about one third of these mGFP+ rings surrounding axons did not reveal detectable MBP expression, suggesting that mGFP+ tubes label either OPC processes engaging with axons or early oligodendrocyte ensheathment (MBPNEG; Figure 2.4 e).  De novo myelin production in the uninjured age-matched spinal cord was much slower and only 2.16% of the MBP+P0NEGmyelin sheaths were mGFP+, i.e. newly-generated over a comparable   71 period of 12 weeks (Figure 2.4 d). Taken together, these data demonstrate that there is a 6-10 fold increase in de novo myelination at the lesion epicenter in the three months following SCI compared to uninjured age-matched controls.  2.3.3 The majority of myelinating SCs in the injured spinal cord are derived from PDGFRa+ cells  SC myelination is prominent following contusive SCI, and PDGFRa+ cells have been show to produce myelinating SCs following chemical demyelination (Zawadzka et al., 2010). To determine if PDGFRa+ cells are also responsible for SC myelination, we examined the expression of myelinating SC markers in recombined cells of PDGFRa-CreER:YFP or mGFP mice. In PDGFRa-CreER:mGFP mice the membrane-tethered reporter allowed us to readily visualize co-expression of mGFP and P0 in myelin tubes after SCI, suggesting that PDGFRa+ cells gave rise to myelinating SCs after contusion SCI (Figure 2.5 a-d; arrows). Twelve weeks after SCI, PDGFRa+ progenitor-derived SC tubes exhibited many characteristics of mature SC myelin including the expression of Caspr in the axonal-glial contacts of the paranodes (Figure 2.5 a) as well as the formation of Schmidt-Lantermann incisures (Figure 2.5 b). Although this was not quantified, example images revealed that recombined myelinating SCs expressed the transcription factor Krox20 (Figure 2.5 c) and were surrounded by a basal lamina staining for laminin (Figure 2.5 d), both hallmarks of myelinating SCs. Importantly, P0+, PDGFRa-derived myelin was abundant in both independent PDGFRα-CreER mouse   72 lines (i.e., in lines I and II) after SCI and PDGFRa-fate mapping did not label SCs in the peripheral nerve (Figure 2.2 h-m).  We next examined the distribution and contribution of OPC-derived SCs in the PDGFRα-CreER:YFP or mGFP mice to the myelination of axons at the SCI  epicenter (Figure 26 a-d,f). We observed that the P0+ SC myelin was most concentrated in the dorsal regions of the cord near the injury epicenter. Due to the nature of the dorsal contusion injury, the cytoarchitecture of the spinal cord was most disrupted within the lesion core and in the dorsal column regions, indicated by sparse GFAP staining (Figure 2.6 a’). Twelve weeks after SCI, these dorsal areas of substantial astrocyte loss were populated by YFP+ myelin sheaths that also expressed the SC-specific myelin marker P0 (Figure 2.6 a, a’, a’’). Because SC myelin is surrounded by a thicker cytoplasmic outer wrapping compared to oligodendrocyte myelin, we were able to observe distinct recombined rings around SC-myelinated axons in both the YFP (Figure 2.6 a-d) and mGFP (Figure 2.6 e) reporter lines. SC myelin sheaths in the vicinity of the dorsal root entry zone were typically not derived from PDGFRa+ cells (i.e. were YFPNEG; Figure 2.6 c’); in contrast, the medial dorsal columns contained many P0+ and YFP+ myelin sheaths (Figure 2.6 c’’). Recombined P0+ myelin sheaths possessed morphological characteristics indicative of SCs. For example, they myelinate axons with a one-to-one sheath-to-cell ratio (Figure 2.6 d; arrowheads) and expressed a basal lamina (Figure 2.5 d). Recombined myelinating SCs were also observed in mGFP reporter mice (Figure 2.6 e, e’). The number of SCs profiles derived from recombined PDGFRa progenitors in the injured spinal cord increased over time; by 12 weeks post-SCI, 67 ± 7.4% of the P0+ myelin co-expressed YFP suggesting an ongoing production of SCs by PDGFRa   73 progenitors after injury (Figure 2.6 f). In contrast, there was no change in the extent of ensheathment/myelination by peripherally-derived (YFPNEG) SCs between three and twelve wpi. The percentage of recombined SC tubes represents a slight underestimate considering that the recombination efficiency in this mouse line was 84%. After accounting for this recombination efficiency, we estimate that approximately 70-80% of the SC profiles at injury epicenter 12 weeks after SCI were derived from PDGFRa cells and their progeny. Keeping in mind that PDGFRa labels several populations of cells in the spinal cord and dorsal root, these data are consistent with an early appearance of SCs derived from PDGFRaNEG cells—presumably migratory SCs (see below)—from the dorsal root entry zone coupled with an ongoing increase of SCs derived from PDGFRa+ cells and their progeny.    2.3.4 Olig2+ cells give rise to P0+ SCs in response to contusion SCI.  Considering that fate mapping using PDGFR-aCreER:YFP or mGFP mice labels other cell types in addition to OPCs, we wanted to test whether cells specific to the oligodendroglial lineage can give rise to P0+ myelinating SCs after SCI. We lineage traced the Olig2+ cells using Olig2-CreER:YFP mice.  In the spinal cord of uninjured control Olig2-CreER:YFP mice, we observed tamoxifen-induced recombination in oligodendrocytes and OPCs (Figure 2.7 a), as well as in a subset of gray matter astrocytes, as previously described (Dimou et al., 2008). Importantly, unlike the PDGFRa-CreER:YFP or mGFP lines, there was no recombination in vascular-associated cells, central canal-associated cells, or cells in the dorsal roots (Figure 2.7 b),   74 suggesting that the only cellular overlap between Olig2-CreER and PDGFRa-CreER  is with OPCs. Despite having a low recombination efficiency in OPCs at time of injury compared to the PDGFRaCreER mice, Olig2CreER:YFP+ cells were encountered in the dorsal columns (Figure 2.7 c) and adjacent to the lesion epicenter cavity, at twelve weeks after SCI (Figure 2.7 d). A subset of these recombined cells expressed P0 and exhibited the morphology of typical myelinating SCs (Figure 2.7 d). This confirms that cells of the oligodendroglial lineage give rise to myelinating SCs after contusion SCI.  2.3.5 Recombined PDGFRa-expressing cells from the PNS do not give rise to myelinating SCs in vitro or in vivo Migration of SCs from the periphery into the CNS parenchyma was originally considered the primary source of myelinating SCs after SCI (Franklin and Blakemore, 1993). To examine the potential of peripheral PDGFRa-expressing cells to give rise to myelinating SCs, we FACS-isolated GFP+ (and GFPNEG) cells from dorsal root/DRG and from sciatic nerves of adult PDGFRa:H2B-GFP mice and characterized the fate of these cells in vitro (Figure 2.8). When grown in SC proliferation/differentiation media for one week, GFPNEG cells exhibited a bipolar morphology consistent with SC differentiation (Figure 2.8 d & f), while GFP+ cells assumed a flattened morphology (Figure 2.8 c & e). Cells from the GFPNEG cell fraction (Figure 2.6 o-v) expressed proteins characteristic of SC precursors, such as p75-NTR (o, p), nestin (q, r) and Sox2 (s, t). In contrast, GFP+ cells (Figure 2.8 g-n) did not express SC lineage markers, but expressed αSMA, consistent with a fibroblast-like fate.    75 Considering that progenitor populations can behave differently in their quiescent state compared to in the wake of injury, where they have been noted to proliferate and contribute to repair (Joe et al., 2010; Almad et al., 2011), we wanted to specifically look at the responses of the peripheral PDGFRa recombined precursors in response to local injury. To determine whether injury-activated PDGFRa+ cells in the dorsal root or sciatic nerve produce myelinating SCs, PDFGRa-CreER:mGFP mice were given tamoxifen two weeks before dorsal root (Figure 2.9 a, b, d) and sciatic nerve (Figure 2.9 c) crush injuries.  After PNS injury, recombined  cells with a flattened and branched morphology were numerous in the endoneurial spaces; Figure 2.9 a-c). Twelve weeks after dorsal root crush (Figure 2.9 d), there was no evidence of mGFP+/P0+ SCs in the dorsal root (Figure 2.9 d’, d’’) or the DRG; Figure 2.9 d’’’). In the PNS, neither dorsal root injury nor sciatic nerve crush stimulated PDGFRa cells to express the SC marker P0; PDGFRα derived cells also did not possess SC morphology or close associations with axons (Figure 2.9 a-d). Consistent with our findings above, inadvertent spinal cord damage during dorsal root injury did stimulate recombined cells to express P0, indicative of SC production, but these cells were restricted within the injured spinal cord (Figure 2.9 d below dotted line).  These findings indicate that PDGFRa-expressing cells residing in the PNS do not give rise to SCs in vitro nor after local PNS injury in vivo.     76 2.3.6 Peripheral myelinating SCs migrate into the spinal cord after injury and contribute to myelination In order to examine the contribution of mature, myelinating SCs in the PNS to myelination after SCI, we used P0Cre-ER:YFP mice. In uninjured control mice, recombination occurred in only 6.9 ± 1.18% and 24.1± 2.37% of the dorsal and ventral myelinating SCs, respectively; no recombination was observed within the CNS (Figure 2.10 a-d), as expected from specificity of P0 for PNS myelin. Twelve weeks after SCI, a small population of myelinating SCs was observed at the lesion epicenter (Figure 2.10 e & e’). Importantly, the relative contribution of the recombined YFP+/P0+ myelin sheaths was small relative to the number of P0+ myelin sheaths both three and twelve weeks after SCI (Figure 2.10 f). When taking the low recombination efficacy into consideration, the percentage of P0+ cell-derived SCs is less than 10% of the total P0+ tubes in the injured spinal cord, supporting the notion that OPCs generate the majority of SCs in the cord after SCI.  2.4 Discussion  Here we determined which cell types contribute to the regeneration of myelin following contusion SCI, by systematically assessing the contributions of PDGFRα+, olig2+, and P0+ cells to the generation of myelinating glia and axonal ensheathment/myelination following SCI. Using in vivo genetic fate mapping methods, we show that OPCs generate the vast majority of myelin produced by both oligodendrocytes and SCs following SCI, indicating that these endogenous progenitors are capable of restoring myelin sheaths in the absence of treatment.    77 The extent and source of spontaneous remyelination in the injured spinal cord has been the focus of considerable interest and debate for more than a decade, with important implications for treatment development after SCI. Previous fate mapping experiments of PDGFRa progeny using various recombination time points after SCI revealed that oligodendrogenesis and new myelin formation is an ongoing process in chronic SCI (Hesp et al., 2015). We used genetic fate mapping of the PDGFRa progeny recombined prior to injury to characterize the amount of de novo oligodendrocyte- and SC-derived myelination over time. At 12 wpi, we found extensive increases in new oligodendrocyte formation and in ensheathment/myelination by oligodendrocytes derived from PDGFRα+ progenitors labelled prior to injury. Importantly, the vast majority of myelinating SC tubes in the injured spinal cord are centrally-derived:  both PDGFRα+ progenitors and Olig2-expressing cells gave rise to myelinating SCs, while only a minority were derived from the P0+ peripheral population.   2.4.1 OPC derived ensheathment/myelination is substantial after SCI The notion of ongoing demyelination after SCI as a pathological process limiting functional recovery (Blight, 1983b; Totoiu and Keirstead, 2005) has fuelled both preclinical and clinical research (Plemel et al., 2014b). There is some evidence in pre-clinical work that transplanting myelinating cells can increase or accelerate myelin repair compared to spontaneous repair (Keirstead et al., 2005; Karimi-Abdolrezaee et al., 2006; Cao et al., 2010; All et al., 2015). Consistent with previous findings, we observed extensive oligodendrocyte production from OPCs in the astrocyte-rich parenchyma surrounding the injury (McTigue et al., 2001; Tripathi and McTigue, 2007; Sellers et al.,   78 2009; Hesp et al., 2015) and the contribution of new oligodendrocytes to the myelination of axons in the chronic injury site (Hesp et al., 2015). By tracking cumulative myelination of spinal axons over time after injury (Figure 2.4), we found that spontaneous myelination is progressive over time: at 3 weeks after SCI, de novo ensheathment/myelination accounted for ~15% of all MBP+ myelin sheaths, and by 12 weeks post-SCI this number approximated 30% at injury epicenter. Considering that the overall myelin (Figure 2.4 c) did not demonstrate a significant change between 3wpi and 12wpi yet there is a significant increase in the amount of new myelin (Figure 2.4 d) it suggests that myelin is being turned over and replaced between 3wpi and 12wpi.  To our surprise, about one-third of the 6,000 de novo ensheathments did not reveal MBP expression at 12 wpi indicative of OPC processes or early oligodendrocyte ensheathments. Taken together, there is considerable new myelin production after SCI and even in the chronic setting there is ongoing myelin turnover.  These findings of extensive new myelin production after SCI suggest that this regenerative response is an important contributor for functional recovery after SCI. Still, it is possible that new myelin production is not a regenerative process for the sole purpose of regenerating lost myelin segments. For example, even in the fully myelinated optic nerve and in the absence of demyelination there is de novo myelination by intercalation between existing myelin sheaths (Young et al., 2013). It is therefore possible that injury is stimulating such de novo myelination that is not linked with active or ongoing demyelination. In such a scenario, this non-regenerative myelin production might not contribute to functional recovery. Even if new myelin production is regenerative (i.e. replacing lost myelin), we cannot differentiate between myelination of spared axons, cut   79 axons that have been severed outside of the plane of section, or newly-sprouted axons. This is important because we do not yet know whether newly-ensheathed/myelinated axons are functional. Oligodendrocytes can myelinate even dead axons and/or artificial axons (Lee et al., 2012b; Lee et al., 2013; Bechler et al., 2015) and thus some of the new myelin produced after SCI might be of severed axons, which is unlikely to provide a functional benefit. These considerations underscore the importance of addressing the functional importance of new myelin production and how it is associated with spontaneous recovery after SCI.   2.4.2 The majority of myelinating SCs in the contused spinal cord are derived from PDGFRa+ CNS progenitors For many decades and in diverse animal models, SCs have been observed in the injured spinal cord, including in humans (Bunge et al., 1961; Bresnahan, 1978; Bunge et al., 1993; Beattie et al., 1997; Guest et al., 2005; James et al., 2011). The prevailing interpretation has been that these SCs migrated in from the PNS to contribute to myelination within the CNS (Franklin and Blakemore, 1993; Sims et al., 1998; Jasmin et al., 2000). Here, we demonstrate that, when we consider recombination efficiency, approximately 70-80% of the myelinating SCs in the spinal cord 12 weeks post contusion SCI are derived from resident PDGFRa+ cells. The CNS origin of SCs is supported by our findings that Olig2-recombined cells also give rise to SCs after SCI. We conclude that PDGFRa+ OPCs thus are the primary contributor to the myelinating SC population observed in the injured spinal cord. We also estimate that less than 10% of the SCs in the cord after injury are derived from peripheral myelinating SCs. Therefore, via these   80 indirect measurements, we can only account for the cellular origin of about 80-90% of the myelinating SCs encountered in the contused spinal cord suggesting the existence of one or more other contributing cell populations. It is possible that non-myelinating SCs or other DRG progenitors (Vidal et al., 2015) could be producing SCs in response to injury. The generation of SCs from PDGFRa+ OPCs after CNS injury—and not from migration and/or differentiation of recombined PDGFRa+ precursors residing in the dorsal or ventral roots—is consistent with previous observations after focal chemical demyelination of the spinal cord (Zawadzka et al., 2010). In agreement, OPCs transplanted into demyelination lesions are able to generate SCs (Talbott et al., 2005; Talbott et al., 2006). A recent study proposed a CNS origin of SCs after SCI on the basis of dorsal rhizotomies (Bartus et al., 2016). Here we provide conclusive evidence using fate mapping of various candidate cells expressing Cre from the PDGFRa and Olig2 promoters after contusion injury. The production of SCs from OPCs is not observed in culture or after co-transplantation with astrocytes and thus differentiation is BMP-dependent (Talbott et al., 2006). De Castro et al. recently reported that cell specific deletion of STAT3 in astrocytes decreases remyelination by oligodendrocytes in favour of SCs using a chemical demyelination model (Monteiro de Castro et al., 2015). Taken together, these findings suggest that an astrocyte-derived signal is required for OPC differentiation into oligodendrocytes. In agreement with this hypothesis, OPC-derived SC differentiation was observed after contusion SCI mainly in the dorsal columns where astrocytes are rare. Astrocytes within these regions may also be phenotypically different from astrocytes in other regions of the cord (Tsai et al., 2012). A better understanding of the regenerative potential of OPCs and their molecular regulation may provide new   81 avenues in CNS repair. SC transplantation has been shown to elicit moderate functional improvements in pre-clinical models (Pearse et al., 2004b; Biernaskie et al., 2007) and is currently being studied in clinical trials (Clinical NCT01739023, NCT0235425). SC transplantation can result in an even greater endogenous SC response after SCI (Hill et al., 2006; Biernaskie et al., 2007; Sparling et al., 2015). Therefore, gaining a better understanding of the endogenous SC response observed after injury in both rodents and humans could help to develop future treatment strategies. Importantly, the finding of Schwannosis in the chronic clinically injured SCI population (Bruce et al., 2000; Norenberg et al., 2004) has been discussed as a cause of functional decline pointing to the need for a better understanding of the biological processes regulating SC production after SCI. Transgenic deletion of NRG1 in all cells prevented myelination by SCs in the injured spinal cord of mice and these mice showed worse functional outcomes in an open field locomotor test as early as one week after SCI (Bartus et al., 2016). However, this recovery could also be due to other neuroprotective effects of NRG1 after SCI (Gauthier et al., 2013; Alizadeh et al., 2017). Hence, the functional significance of endogenous SC myelination still remains to be shown. This formation of SCs in the areas of most extensive spinal cord damage, where astrocytes become sparse, might represent an endogenous repair mechanism and a target for future therapeutic interventions; bridging severe injuries by stimulating OPCs to produce SCs might allow for a more conducive environment for axonal growth and myelin formation.    82 2.5 Conclusion Our findings demonstrate that endogenous OPCs are capable of extensive myelination after SCI. The majority of OPCs remain lineage-restricted and produce myelinating oligodendrocytes that ensheath axons. We also demonstrate that PDGFRa+ cells (likely OPCs) are the source of the majority of SCs present in the spinal cord after clinically-relevant (contusion) SCI. This repair mechanism by endogenous, CNS-derived SCs may represent a novel therapeutic target for repairing the injured spinal cord in areas of grossly disrupted cytoarchitecture.                  83    Table 2-1. Overview of transgenic mouse lines           Transgenic Mice Labelled Cell Populations  Cellular Label Tamoxifen  Reference PDGFRαCreERTM(I): Rosa26eYFP  Short Form:  PDGFRαCreER(I): YFP PDGFRa+ Cells Before Injury:  -OPCs -vascular associated cells -central canal associated cells -PNS endoneurial cells (fibroblast-like) Cytoplasmic YFP  Short Form: YFP 3mg/day  for 5 days Kang et al., 2010; Jackson Labs; 006148 PDGFRαCreERTM(I): Rosa26mGFP(mT/mG)  Short Form:   PDGFRαCreER(I): mGFP PDGFRa+ Cells Before Injury:  -OPCs -vascular associated cells -central canal associated cells -PNS endoneurial cells (fibroblast-like) Membrane-tethered  GFP   Short Form: mGFP 3mg/day  for 5 days Kang et al., 2010;  Jackson Labs; 007576 PDGFRαCreERT2(II): Rosa26eYFP  Short Form:   PDGFRαCreER(II): YFP PDGFRa+ Cells Before Injury:  OPCs -vascular associated cells -central canal associated cells -PNS endoneurial cells (fibroblast-like) Cytoplasmic YFP  Short Form: YFP 3mg/day  for 5 days Rivers et al., 2008;   Jackson Labs; 006148 PDGFRαCreERT2(II):  Rosa26mGFP(mT/mG)  Short Form:   PDGFRαCreER(II): mGFP PDGFRa+ Cells Before Injury:  OPCs -vascular associated cell -central canal associated cells -PNS endoneurial cells (fibroblast-like) Membrane-tethered  GFP   Short Form: mGFP 0.5mg/day  for 2 days Rivers et al., 2008;  Jackson Labs; 007576 Olig2CreERTM: Rosa26eYFP  Short Form:   Olig2CreER: YFP Olig2+ Cells Before Injury:  OPCs -oligodendrocytes -population of  -grey matter astrocytes Cytoplasmic YFP  Short Form: YFP 3mg/day  for 5 days Takebayashi et al., 2002;  Jackson Labs; 006148 P0CreERT2: Rosa26eYFP  Short Form:   P0CreER: YFP P0+ Cells Before Injury:  -myelinating peripheral SCs Cytoplasmic YFP  Short Form: YFP 3mg/day  for 5 days Leone et al., 2003;   Jackson Labs; 006148 PDGFRa:H2B-GFP  Cells currently expressing PDGFRa: -PDGFRa+ cells in CNS and PNS  Cytoplasmic YFP  Short Form: YFP n/a Hamilton et al., 2003,  Jackson Labs; 007669   84 Table 2-2. Overview of specific animal numbers that underwent qualitative or quantitative analysis    Group Qualitative Analysis Quantitative Analysis PDGFRαCreERTM(I): Rosa26eYFP    Time of Injury (Uninjured Spinal Cord) 6 3 3 wpi (Injured Spinal Cord) 10 7 12 wpi (Injured Spinal Cord) 10 7 Week 12 Uninjured (Uninjured Spinal Cord) 6 4 Time of Injury (Uninjured Sciatic Nerve) 3  No TA Controls (Injured Spinal Cord) 3  No TA Controls (Uninjured Spinal Cord) 3  PDGFRαCreERTM(I): Rosa26mGFP(mT/mG)    Time of Injury (Uninjured Spinal Cord) 3  3 wpi (Injured Spinal Cord) 3  12 wpi (Injured Spinal Cord) 10  Week 12 Uninjured (Uninjured Spinal Cord) 6  Time of Injury (Uninjured Sciatic Nerve) 3  4 wpi (Inured Sciatic Nerve) 3  12 wpi (Injured Sciatic Nerve) 3  4 wpi (Injured Dorsal Root 3  12 wpi (Injured Dorsal Root) 6  No TA Controls (Injured Spinal Cord) 2  No TA Controls (Uninjured Spinal Cord) 2  PDGFRαCreERT2(II): Rosa26eYFP     12 wpi (Injured Spinal Cord) 6  Week 12 Uninjured (Uninjured Spinal Cord) 6  No TA Controls (Injured Spinal Cord) 2  No TA Controls (Uninjured Spinal Cord) 2  PDGFRαCreERT2(II): Rosa26mGFP(mT/mG)    Time of Injury (Uninjured Spinal Cord) 6 3 3 wpi (Injured Spinal Cord) 6 4 12 wpi (Injured Spinal Cord) 6 5 Week 12 Uninjured (Uninjured Spinal Cord) 6 3 No TA Controls (Injured Spinal Cord) 2  No TA Controls (Uninjured Spinal Cord) 2  Olig2CreERTM: Rosa26eYFP    Time of Injury (Uninjured Spinal Cord) 3  12 wpi (Injured Spinal Cord) 10  Week 12 Uninjured (Uninjured Spinal Cord) 6  No TA Controls (Injured Spinal Cord) 4  No TA Controls (Uninjured Spinal Cord) 4  P0CreERT2: Rosa26eYFP    Time of Injury (Uninjured Spinal Cord) 6  1 wpi (Injured Spinal Cord) 6 5 3 wpi (Injured Spinal Cord) 6 6 12 wpi (Injured Spinal Cord) 10 7 Week 12 Uninjured (Uninjured Spinal Cord) 6 4 Time of Injury (Uninjured Sciatic Nerve) 3  No TA Controls (Injured Spinal Cord) 2  No TA Controls (Uninjured Spinal Cord) 2    85 Table 2-3. Primary antibody table   Primary Antibody Used as a Marker For Host Source, Catalogue # Ratio; IHC/ICC APC (CC1 Clone) Oligodendrocytes Mouse,  IgG2bκ Calbiochem, OP80 1:300; IHC αSMA clone  EPR5368 Fibroblast-like Phenotype Rabbit  Millipore, MABT381 1:500; ICC αSMA Vascular-associated Cells (Type B  Pericytes as defined by Goritz et al., 2011) Rabbit Abcam, ab5694 1:500; IHC BIII-Tubulin  (Tuj1) Neural-Specific Tubulin Mouse, IgG2bk Sigma, T8660 1:1000; IHC Contactin  Associated  Protein (Caspr) Paranodes Rabbit Abcam, ab34151 1:500; IHC Fibronectin  Fibroblasts and Extracellular Matrix Mouse Sigma, F7387 1:400; IHC GFAP Reactive Astrocytes Rabbit DAKO, Z0334 1:1000; IHC GFP Reporter+ Cells (YFP+ or GFP+) Chicken Abcam, ab13970 1:1000; IHC GFP Reporter+ Cells (YFP+ or GFP+) Goat Rockland, 600-101-215 1:500; IHC GFP Reporter+ Cells (YFP+ or GFP+) Rabbit Abcam, ab290 1:6000; IHC GFP Reporter+ Cells (YFP+ or GFP+) Rat Nacalai tesque, 04404-84 1:1000; IHC GFP Reporter+ Cells (YFP+ or GFP+) Mouse,  IgG1 Millipore, mab3580 1:500; IHC Glut1 Endothelial Cells Goat Santa Cruz, sc-1605 1:200; IHC Krox 20 (Egr2) Myelinating SCs Rabbit BioLegend (Covance) PRB-236P 1:500; IHC Laminin Basal Lamina, Extracellular Matrix Rabbit Sigma, L9393 1:200; IHC MBP Myelin Chicken Aves, MBP 1:200; IHC MBP Myelin Goat Santa Cruz, sc-13914 1:500; IHC MYRF, GM98 (N terminus) Differentiated Oligodendrocytes Rabbit Gift from Dr. Wegner 1:500; IHC Nestin,  clone 2Q178 Consistent with an Immature/  Proliferative SC Phenotype Rabbit  Santa Cruz, sc-58813 1:500; ICC Neurofilament  200 Heavy Chain Neurofilaments Mouse,  IgG1 Sigma, N0142 1:1000; IHC NG2 Oligodendrocyte Precursors Rabbit Millipore, AB5320 1:200; IHC Olig2 Oligodendrocyte Precursors and  Oligodendrocytes Rabbit Millipore, AB9610 1:300; IHC P0 Myelinating SCs,  Peripheral Myelin Chicken Aves, PZ0 1:100; IHC P75-NTR Consistent with an Immature/ Proliferative SC Phenotype Rabbit Millipore, AB1554  1:200; ICC P75-NTR Non Myelinating SCs Rabbit Sigma, N3908 1:100; IHC PDGFRα Oligodendrocyte Precursors,  Type A Pericytes (as defined by  Goritz et al., 2011) Goat RNDsytems, af1062 1:100; IHC   PDGFRβ Type A Pericytes  (as defined by Goritz et al., 2011) Rabbit Abcam, ab32570 1:100; IHC Reca 1, clone HIS52 Endothelial Cells Mouse,  IgG1 Serotec, MCA970R 1:500; IHC SMI312 Pan Neurofilaments Mouse Covance, SMI-312R-100 1:1000; IHC Sox2 Consistent with an Immature/  Proliferative SC Phenotype Rabbit Stemgent, 09-0024 1:200; ICC        86       Figure 2.1 Genetic labeling of NG2 glia in tamoxifen-inducible PDGFRαCreER uninjured control mice (a, b) After the 2 week tamoxifen wash-out period, Tamoxifen-induced YFP expression in the cytoplasm was observed in two independent PDGFRαCreER mouse lines (line I and II) crossed to the YFP reporter line. The majority of PDGFRα+ (red)/Olig2+ (blue) cells exhibited recombination (green) in PDGFRαCreER(I):YFP mice (a, a’); recombination was more modest in PDGFRαCreER(II):YFP mice (b, b’) when observed in uninjured control mice at 14 days post tamoxifen treatment. (c-d) In both lines, YFP (green), Olig2 (red), and PDGFRα (blue) were co-expressed (c), and the recombined population of cells was also co-expressed with the NG2+ (red) population. Arrows point to rare examples of GFP+ cells not overlapping with NG2+ cell (d). (e) PDGFRαCreER mice were crossed with membrane-tethered (mGFP; green). (f) 3D rendering at fourteen days after tamoxifen treatment, the majority of PDGFRa+ cells are recombined (mGFP+; green; arrows) with a small subset of PDGFRa+ cells not recombined (arrowhead). It was rare to find recombined cells that had matured into an oligodendrocyte (CC1+; blue) at time of injury. (g) Confocal image including Z plane from outlined box in 1f demonstrating that PDGFRa does not overlap with CC1 (blue) cells but it is two independent cells on top of one another. All images were taken in spinal cord cross sections. Scale bars, 200µm (a, b, e); 50µm (d); 10µm (a’, b’, c, f).   87        88     Figure 2.2 Recombination in central canal-associated cells, pericytes and a subset of PNS endoneurial cells in PDGFRaCreER uninjured control mice  (a, b) In the uninjured spinal cord of PDGFRaCreER:YFP or mGFP, recombination (green) was observed in a subset of blood vessel-associated cells (arrow) located on the outside of the endothelial layer (RECA, red; Glut1, blue) but inside the outer basal lamina, (red; c) consistent with the location of pericytes. The majority of the YFP-expressing vascular-associated cells expressed PDGFRa (blue; d) and PDGFRb (red; d), referred to previously as type A pericytes. A small subset of the YFP+ vascular associated cells appeared to co-express of YFP (green) and aSMA (red; e; referred to previously as a type B pericytes marker). (f, g) Recombination can also be seen in a small number of cells located peripherally in the wall of the central canal, each with a process extending into the lumen of the canal (f); these cells were also NG2+ (red; g). (h-l) A subset of endoneurial/perineurial cells and pericytes (green) that exhibited recombination (i.e., YFP+ or mGFP+) were found in the dorsal root (h), dorsal root ganglion (i), and sciatic nerve (j) in the uninjured PNS in both lines of PDGFRaCreER. Many recombined cells in the dorsal root co-expressed the fibroblast marker fibronectin (blue; k). Rarely were YFP+ pericyte cell bodies encountered (e.g., arrow, nucleus, 3D rendering, l), with associated processes encircling blood vessels in the dorsal root. (m) Importantly, recombined cells in the dorsal root did not co-express the non-myelinating SC marker p75 (cyan; m), or the myelinating SC marker P0 (red; i,j,l,m). Images were taken in spinal cord cross section in a-g and longitudinal sections in h-k. Scale bars, 100µm (a); 50µm (i); 10µm (b, c, f, g, h, j, k, m); 5µm (d, e, l).     89       90     Figure 2.3 PDGFRa+ progenitors proliferate and contribute to oligodendrocyte lineage cells in response to SCI (a) Timeline for SCI experiment. Tamoxifen was administered to 8-10 week old mice; mice were given a 2-week tamoxifen washout period prior to a T9/T10 contusion SCI. Twelve weeks after SCI, recombined YFP+ cells incorporated EdU (red; b) and many recombined cells differentiated into CC1-expressing (red) mature oligodendrocytes (arrow; c and d). Note: schematic in top right corner of images indicates approximate location where image was taken based on spinal cord cross section. Using the mGFP reporter mice, a small subset of recombined cells continue to express PDGFRα (blue; arrow; e) while the majority of recombined cells are now oligodendrocytes with extended processes ensheathed/myelinated axons and express MYRF (red; arrow; f).  (g) mGFP+ oligodendrocytes expressed the paranodal marker Caspr (arrows) with split channels (g’). There were no differences observed in the overall number of OPCs (PDGFRa+Olig2+) or recombined OPCs (YFP+PDGFRa+Olig2+) across the groups (h). There was a decrease in the overall oligodendrocytes (CC1+Olig2+) at 3 wpi and 12 wpi compared to the week 12 uninjured group (p <0.039; i). There were more overall oligodendrocytes at 12 wpi compared to 3 wpi (p=0.033; i). There were more new oligodendrocytes (YFP+CC1+Olig2+) at 12 wpi compared to the week 12 uninjured group (p=0.020; i) but the difference observed between the week 12 uninjured group and 3wpi did not reach significance (p=0.07; i). Amongst the total YFP+Olig2+ population, there was a higher percentage of oligodendrocytes (YFP+CC1+Olig2+) at 12 wpi compared to both 3 wpi (p=0.033) and the week 12 uninjured control group (p=0.02; j). Reciprocally, there was a lower percentage of OPCs (YFP+PDGFRa+Olig2+) at 12 wpi compared to the 3 wpi (p=0.033) and the week 12 uninjured control group (p=0.02; j).  Images were taken in both epicenter spinal cord cross sections (b, c, d, e) and longitudinal sections (f, g) near epicenter.  *= p<0.05; += p< 0.1. Scale bars, 50µm (c); 20µm (b, d); 15µm (f); 10µm (e, g); 2µm (g’).    91        92     Figure 2.4 Extensive new ensheathment/myelination by oligodendrocytes derived from PDGFRa+ progenitors 12 weeks after SCI Twelve weeks after contusion injury, large numbers of membrane-bound mGFP+ tubes (green) were observed in PDGFRαCreER (II):mGFP mice indicating new ensheathment/myelination (Figure a, b). Slides were stained for axons (blue), MBP (red or purple), mGFP (green), and P0 (not shown). Note: Some images are displayed as flattened images (combining large numbers of z-stacks into one image; a, b, b’) while others are a single z-stack image (a’, b;). A proportion of sheaths co-expressed MBP (e.g., a’ and b’’; single optical plane at higher magnification; arrows denote clear mGFP+ [green] and MBP+ [purple or red] tubes).  (c-e) Quantification of axons myelinated by oligodendrocytes after SCI (i.e., MBP+, P0NEG processes) at the lesion epicenter demonstrated significantly less myelinated axons at 3wpi and 12wpi compared to uninjured age matched controls (c). The percentage of newly myelinated axons (surrounded by both mGFP+ and MBP+ tubes) to total myelinated axons (surrounded by just MBP+ tube) was significantly higher at 12 wpi compared to 3wpi and uninjured age matched controls (d). Quantification of MBP expression in mGFP+ sheaths indicative of new myelin (open portion of bar) and of MBPNEG mGFP+ sheaths indicative of either OPC process wrapping or merely ensheathing oligodendrocytes (closed portion of bar; e). All images were taken in epicenter spinal cord cross sections. *= p<0.05. Scale bars, 200µm (b); 100µµ (a); 20µm (b’); 5µm (a’, b’’).    93    94    Figure 2.5 PDGFRa+ progenitor-derived SCs express typical hallmarks of SC myelination in PDGFRaCreER:mGFP mice (a-b) Arrows point to paranodal marker Caspr (white; a) and to Schmidt-Lanterman incisures (area where myelin is less compact allowing a accumulation of GFP antibody and a decrease in P0; arrows; b); both indicative of mature Nodes of Ranvier and myelination. (c-d) 12 weeks after injury, recombined myelinating SCs expressed the transcription factor Krox20 (arrow, c) and were surrounded by a basal lamina (d; arrows point to cell body), both hallmarks of myelinating SCs. All images taken in spinal cord longitudinal sections near epicenter. Scale bars, 5µm (b, c, d); 3µm (a).        95    Figure 2.6 The majority of myelinating SCs in the injured spinal cord are derived from PDGFRa+ progenitors (a) Twelve weeks after spinal cord contusion in PDGFRaCreER:YFP or mGFP mice, P0+ (red) SC myelin was abundant within the dorsal columns in areas of substantial astrocyte loss. There were two distinct populations (a’’) of P0+ myelin sheaths, a YFPNEG population (c’) and a YFP+ population (b, c’’); most YFPNEG P0+ myelin sheaths were found closer to the dorsal root entry zone whereas the YFP+ P0-postive sheaths were found mainly medially in the dorsal column (c’’). (b) Arrowheads points to YFP+/P0+ myelin sheaths with the cell bodies of a SCs in the image plane. The arrow denotes an oligodendrocyte ensheathed YFP+ nerve fiber. These nerve fibers revealed the one-to-one sheath-to-cell ratio typical for SCs (arrowheads in d). (e) SC myelination was also apparent using the membrane-tethered reporter mGFP and horizontal sections through the lesion site (e, e’; dorsal to the left). (f) The number of YFP+/P0+ SC profiles (green portion of bar) increased between 3 and 12 weeks after SCI (p=0.001). There were significantly more overall P0+ profiles (grey + green portion of bar) at 12wpi compared to 3 wpi and the majority of P0+ tubes were also YFP+. Images were taken in both epicenter spinal cord cross sections (a, b, c, d) and longitudinal sections near epicenter (e). *p= <0.05. Scale bars, 200µm (a, e); 20µm (c’, c’’, a’, a’’); 10um (e’); 5µm (b, d).                         96               97      Figure 2.7 Olig2+ cells give rise to myelinating SCs after SCI Olig2creER:YFP mice were used to fate-map oligodendroglial lineage cells.  (a) In the uninjured thoracic spinal cord 2 weeks after tamoxifen dosing, recombination (YFP; green) occurred in Olig2+ cells (white) across the oligodendrocyte lineage (preferentially observed in CC1+ oligodendrocytes [red] and to a lesser extent PDGFRα+ OPCs [blue]) and a subset of grey matter astrocytes (b). There was no recombination observed in cells associated with the central canal (blue outline surrounds central canal; split channels with axons [white] and YFP [green]; b’’), or the PNS (dorsal root; b’’’). (c) 12 weeks after SCI, YFP+ cells were abundant at the lesion epicenter and a subset of the YFP+ cells demonstrated typical SC markers and morphology in the dorsal columns (c’, c’’) and in close proximity to a cavitation at the epicenter (d, d’). All images were taken in spinal cord cross sections. Scale bars, 200µm (a, b, c); 50µm (d); 20µm (a’); 10µm (b’, b’’, b’’’, c’, c’’, d’).            98    99     Figure 2.8 PDGFRα+ cells from the adult dorsal root ganglion and spinal root of PDGFRα:H2BGFP mice do not exhibit SC fate in vitro Sciatic nerve (a) and dorsal root ganglion/spinal root (DRG/root; b) -derived cell suspensions from adult PDGFRα:H2BGFP mice were sorted for GFP-expression by FACS. (c-f) PDGFRα:H2BGFP+ and PDGFRα:H2BGFPNEG cells were grown in SC proliferation/differentiation media for one week. The bipolar morphology of PDGFRαH2BGFPNEG cells was consistent with SC differentiation (arrow, d, f). PDGFRαH2BGFP+ cells derived from both peripheral sources exhibited a flattened morphology under the same conditions. (g-n) Consistent with morphological findings, PDGFRαH2BGFP+ cells derived from the sciatic nerve (g, i, k, m) or DRG/roots (h, j, l, n) did not express SC lineage markers such as p75-NTR (g, h), nestin (i, j) and Sox2 (k, l) but expressed αSMA, consistent with a fibroblast-like phenotype. (o-v) In contrast, isolated PDGFRα:H2BGFPNEG cells derived from the sciatic nerve (o, q, s, u) or DRG/roots (p, r, t, v) expressed markers of SC precursors such as p75-NTR (o, p), nestin (q, r) and Sox2 (s, t). Some αSMANEG cells were found in the GFPNEG fraction (u, v, arrows). Scale bars, 100µm (c-f); 20µm (g-v).          100       101     Figure 2.9 Recombined PDGFRa+cells in the PNS do not give rise to P0+ cells in response to peripheral injury (a-c) Four weeks after dorsal root crush (a, b) or sciatic nerve crush injury in PDGFRaCreER:mGFP mice (c), mGFP+ cells had branched and flattened processes (fibroblast-like) extending  in the endoneurium between clusters of P0+ myelinating SCs. 12 weeks after a severe dorsal root crush injury (d), there was no evidence of mGFP+/P0+ SCs in the dorsal root (d’, d’’) or the DRG (d’’’). Only the injured spinal cord harbored recombined cells expressing both mGFP and P0. (arrows; d’). Images were taken in root or sciatic nerve longitudinal sections (a, b, c) or spinal cord cross sections (d).  Scale bars, 200µm (d); 50µm (d’); 20µm (d’’’); 10µm (a, b, c, d’’).                 102    103     Figure 2.10 P0+ SCs give rise to a small number of P0+ SCs after SCI In uninjured controls, no recombination was observed within the thoracic spinal cord P0creER:YFP mice (a); recombination (YFP; green) was only observed in the PNS surrounding P0+ tubes consistent with SC morphology (dorsal root: a’, sciatic nerve: c).  Assessment of recombination efficiency in the uninjured roots revealed more YFP+ (green) P0+ (red) tubes in the ventral roots compared to the dorsal roots (d). (e-f) Twelve weeks after SCI, YFP+ myelinating SCs were observed at the injury epicenter of P0creERT2:YFP mice treated with tamoxifen 2 weeks prior to injury (e, e’). The relative contribution of YFP+/P0+ myelin sheaths was low relative to the total number of P0+ myelin sheaths at both 3wpi and 12wpi (<5 SC sheaths per section). There were significantly more overall P0+ tubes (grey + green bar) and more P0+/YFPNEG tubes (grey portion of bar) at 12wpi compared to 3wpi. Images were taken in spinal cord cross sections (a), dorsal root cross sections (b), sciatic nerve longitudinal sections (c), or spinal cord cross sections at epicenter (e). *p=<0.05. Scale bars, 200µm (a, e); 20µm (c); 10µm (a’), 2µm (b, e’).            104 	     105 Chapter 3: Transplantation of skin precursor-derived SCs yields better locomotor outcomes and reduced bladder pathology in rats with chronic spinal cord injury  3.1  Introduction SCI is often devastating, and at present there are limited treatments to improve neurologic function beyond surgical decompression of the spinal cord and rehabilitation therapy. Cell transplantation has long been considered a promising potential treatment for SCI and a variety of candidate cell types are currently under intense investigation in pre-clinical models and clinical trials. Neural cells and their precursors, transplanted in the injured spinal cord may elicit a range of beneficial effects: they can replace lost myelin (Cummings et al., 2005; Keirstead et al., 2005; Karimi-Abdolrezaee et al., 2006; Cao et al., 2010; Plemel et al., 2012), facilitate axonal growth across the injury site (Xu et al., 1995a; Ramon-Cueto et al., 2000; Fouad et al., 2005; Biernaskie et al., 2007; Kanno et al., 2014), stimulate plasticity in spared host circuits (reviewed by Ruff et al., 2012) and exert anti-inflammatory or neuroprotective effects (Ankeny et al., 2004; Mitsui et al., 2005; Biernaskie et al., 2007; Neuhuber et al., 2008; Cusimano et al., 2012; Nakajima et al., 2012; Hilton et al., 2016). In addition, some transplanted cells may differentiate into neurons that can relay impulses across the site of injury to replace lost neural connectivity (Cummings et al., 2005; Bonner et al., 2010; Bonner et al., 2011; Lu et al., 2012).  Most cell types tested to date only fulfill a subset of these reparative roles (for reviews see Ruff et al., 2011; Tetzlaff et al., 2011; Sandner et al., 2012). As one potential   106 candidate cell for the treatment of SCI, SCs possess many desirable properties. SCs secrete trophic factors (Hoke et al., 2006; Krause et al., 2014) and form cellular conduits, similar to the bands of von Büngner found in a regenerating nerve (Funakoshi et al., 1993; Bunge, 1994b; Fu and Gordon, 1997). These cellular conduits align rostro-caudally, and attract and guide the growth of axons into and across the site of SCI (Paino et al., 1994; Xu et al., 1995a; Xu et al., 1999b; Oudega et al., 2001; Hill et al., 2006). In addition, SCs myelinate regenerating and spared axons, provide neuroprotection, and promote axonal growth/plasticity and concomitant functional recovery (Pearse et al., 2004b). Finally, because SCs can be harvested from peripheral nerves (Duncan et al., 1981) or from skin (McKenzie et al., 2006), and expanded in vitro, they are well suited for autologous transplantation, which circumvents the need for immunosuppression and eliminates many ethical issues related to cell acquisition.  To generate sufficient cells for clinical application after SCI, peripheral nerve-derived SCs are typically harvested via excision of 12-15cm of sural nerve (Saberi et al., 2008; Saberi et al., 2011), a procedure that is known to result in sensory deficits and carries the risk of painful neuroma formation (Hood et al., 2009). Those deficits/risks can be avoided entirely by using an accessible minimally invasive source of SCs, those generated from SKPs. SKPs are resident multi-potent stem cells found in the dermis of adult mammalian skin (Biernaskie et al., 2009). SKPs have properties similar to neural crest stem cells, and can be isolated from rodent and human skin to produce functional, myelinating SCs known as SKP-SCs (Toma et al., 2005; McKenzie et al., 2006).  Previously, we demonstrated that SKP-SCs isolated from mouse tissue and transplanted sub-acutely (1 wpi) into the site of rat SCI survived and promoted repair and   107 functional recovery (Biernaskie et al., 2007). In addition, we found that acute (directly post-injury) transplantation of SCs isolated from either neonatal rat nerve or skin-derived precursors promote repair and functional recovery after cervical crush injury (Sparling et al., 2015). SKP-SCs integrated with and promoted sparing of host tissue, facilitated the growth and remyelination of host axons, and increased the presence of endogenous SCs. While these findings are promising, practical issues need to be considered for the clinical translation of SKP-SCs for autotransplantation in sub-acute SCI. Notably, time (in vitro) is required to produce SKP-SCs in sufficient numbers for autologous transplantation. In addition, neurologic function stabilizes over time after SCI, making it considerably easier to detect true efficacy in a clinical trial when the transplantation can be delayed by many mpi (Fawcett et al., 2007).  Given the invasive nature of cell transplantation, a chronic transplantation approach gives patients more time to make informed decisions regarding participation in a trial (Illes et al., 2011). Finally, and importantly, the overwhelming majority of individuals with SCI are currently living with injuries that would be considered ‘chronic’, and so developing therapies that are effective in this setting would have broad applicability. This is indeed a significant unmet need, as very few SCI transplantation pre-clinical studies have been undertaken in a setting resembling chronic injury, and to date, only a few cell treatments have demonstrated any efficacy in such studies (Houle and Tessler, 2003; Barakat et al., 2005 for review see Tetzlaff et al., 2011; Granger et al., 2014).  To address these issues, we transplanted SKP-SCs isolated from rat skin directly into thoracic spinal cord contusion lesions in adult rats at 8 weeks after SCI and examined the long-term anatomical repair (at 29 wpi) and functional outcome (up to 27 wpi).  We   108 found that conspecific SKP-SCs survived long-term at the site of chronic SCI, integrated with spared host tissue, mitigated astroglial scar formation, promoted axonal growth and myelination, and stimulated participation of endogenous SCs in the repair process.  Most importantly, chronic SKP-SC transplantation elicited better functional outcomes and improved bladder pathology in the chronic injury setting.  3.2 Methods 3.2.1 Animals Forty-nine adult female Sprague Dawley rats (295+10g; Charles River Laboratories, Wilmington WA) were used in this study. All procedures were approved by the Hospital for Sick Children Research Institute and the UBC Animal Care Committee in accordance with the guidelines of the Canadian Council on Animal Care. Animals were housed in a room with a reverse light/dark cycle with free access to food and water throughout the study. 3.2.2 Spinal cord contusion injury  Rats received buprenorphine (0.03mg/kg, s.c.) pre-operatively and were anaesthetized with isofluorane (4% induction, ~1.5% maintenance); body temperature was maintained at 36.5±0.5°C. Lidocaine (0.5ml) with 2% epinephrine was injected at the surgical site for additional analgesia and vasoconstriction. The spinal cord was exposed via a thoracic midline incision between T6 and L1 and a laminectomy at vertebra T9 under strictly aseptic conditions. The T8 and T10 dorsal vertebral processes were   109 stabilized with Allen clamps and a 200 Kdyne force-controlled contusion was delivered with an IH impactor (Precision Systems, Lexington, KY; Scheff et al., 2003). Following injury, muscle and skin were sutured in layers. Lactated Ringers solution (10ml, s.c.) was administered every 12 hours for two days to prevent dehydration. Bladders were manually expressed three times daily until spontaneous micturition returned. Antibiotics (Baytril; 10mg/kg, s.c.) were administered as needed to treat minor bladder infections.  Following injury, three rats were euthanized due to injury-related complications and three rats were excluded from the study prior to treatment, based on spontaneous recovery and injury parameters. We excluded animals that were outliers in terms of injury severity; specifically, we removed animals with the Basso, Beattie, and Bresnahan (BBB) score or subscore at 7 wpi that was >2 standard deviations discrepant from the mean. We also removed animals in which area under the force curve (measured by the IH impactor) was >2 standard deviations from the mean. Injured rats (n=31) were divided into two groups that were matched based on peak force of injury, area under the force curve, maximum impactor displacement, pre-injury weight, weight prior to transplant, and BBB scores and BBB subscores at 2 dpi and for each subsequent week prior to transplantation. One group was randomly chosen to receive cell treatment (n=15) and the other medium only (n=16) and investigators were blinded to which animals were given either a cell treatment or medium injection. Two additional rats received adult-derived SKP-SCs; behavioural and histological assessment of these animals was performed alongside the main experiment as investigators were blinded. An additional group of rats received the same 200 Kdyne contusion without treatment to form a time-of-transplant control group (n=10); two animals were excluded from this group because the area under the force   110 curve or BBB scores were >2 standard deviations from the pre-treatment group mean used previously.   3.2.3 SKP isolation and differentiation into SKP-SCs  As previously described (Toma et al., 2001; Fernandes et al., 2004; Toma et al., 2005; Biernaskie et al., 2009), neonatal or adult primary rat skin-derived precursors (SKPs) were prepared from the back skin of either neonatal (P0-P3) or 2 month old transgenic Sprague-Dawley rats that expressed GFP in all cells (SLC, Japan).  Secondary spheres were generated by digesting SKPs with collagenase (1mg/ml) then mechanically dissociating to liberate single cells, which were sub-cultured at a density of 35,000-50,000 cells/ml in flasks. Cells were grown at 37ºC and 5% CO2, fed with SKP proliferation medium (DMEM:F-12; 3:1; Invitrogen, Carlsbad, CA) containing 1% penicillin/streptomycin (Cambrex, East Rutherford, NJ), 2% B27 supplement (Invitrogen), 20 ng/ml epidermal growth factor (EGF; BD Biosciences, Bedford, MA), and 40 ng/ml FGF2 (BD Biosciences) every 5 days, and passaged every 10 days. SCs were differentiated from passage three neonatal or adult SKPs (SKP-SCs) as previously described (McKenzie et al., 2006; Biernaskie et al., 2007). After two or three passages, purified SKP-SCs were frozen in 90% fetal bovine serum (FBS)/ 10% dimethyl sulfoxide at -80ºC for long-term storage. Approximately 2 weeks prior to transplantation, the SKP-SCs were thawed, plated, and expanded under SC proliferation medium [DMEM/F12 (3:1), 1% penicillin/streptomycin, 2% N2 supplement (Invitrogen), 25 ng/ml NRG1β (R&D Systems, Minneapolis, MN), and 5 µM forskolin (Sigma-Aldrich, St. Louis, MO)]    111  3.2.4 Immunocytochemistry  To investigate the expression of typical SCs markers in the SKP-SC cultures, the cells were fixed for 15 min in 4% paraformaldehyde. For a list of primary antibodies used, see Table 3-1. The secondary antibodies were generated in goat, conjugated to Alexa 488, 555 or 647, and used at a dilution of 1:1000 (Invitrogen). Hoechst 33258 (1:1000, Sigma-Aldrich) was used to visualize nuclei. Images were captured via a Zeiss Axiovert 200 spinning disk confocal microscope (Yokogawa, Sugar Land, TX) and C9100-13 EM-CCD camera (Hamamatsu, San Jose, CA), with Volocity acquisition software. SC purity was expressed as the percentage of cells positive for S100β, and the total cell number was determined by Hoechst staining. Four different culture samples were used to determine the purity of neonatal and adult SKP-SCs and at least three fields of view were selected randomly (i.e., while visualizing Hoechst) for each sample.    3.2.5 Cell transplantation  For transplantation at 8 wpi, SKP-SCs were removed from laminin/poly-d-lysine (PDL)-coated plates/flasks by gentle agitation and spraying after 3-5 minute incubation in 0.25% Trypsin/ethylenediaminetetraacetic acid (EDTA). Trypsin was inactivated with 10% FBS and cells were triturated gently to produce a single cell suspension, which was then centrifuged (1000 rpm, 5 min.) and re-suspended at 200,000 cells/µl in fresh DMEM:F12 (3:1). Prior to transplantation, rats were anaesthetized and prepared for surgery as outlined above. The laminectomy at T9 was re-exposed and scar tissue was removed to allow access to the site of SCI. One million neonatal or adult SKP-SCs in 5 µl   112 of medium was stereotaxically injected directly into the epicenter of the contusion site using a 10 µl Hamilton syringe fitted with a glass micropipette (~80 µm tip size). Cyclosporine A (CsA) was delivered to all animals in homecage drinking water (150 mg/l of water; Neoral; Novartis) beginning 4 days before transplantation and continuing for the duration of the study. Oral administration was replaced with injectable CsA for 4 days post-transplantation (Sandimmune, Novartis; 15mg/kg, s.c.). One medium injected animal was euthanized in the 5 days following the transplantation surgery due to bladder complications (final n=15).   3.2.6 Behavioural assessments All behavioural raters were blind to the treatment groups. Functional locomotor abilities were assessed bi-weekly using the open-field BBB score and subscore; (Basso et al., 1995; Basso, 2004), footprint analysis (CatWalk, Noldus, Netherlands; (Hamers et al., 2001), and the irregular horizontal ladder (Metz and Whishaw, 2002). All animals were acclimatized to the testing environment/equipment and trained prior to collection of baseline behavioural data, which were collected during the week prior to SCI. The animals were given two days to recover from SCI before open field-testing resumed, and seven weeks to regain consistent weight supported stepping prior to resuming Catwalk and ladder testing. Animals were further acclimatized to CatWalk and ladder apparatuses prior to the collection of 7 wpi pre-treatment baseline data. The animals were given one week to recover from transplantation surgery before open field-testing resumed and two weeks before CatWalk and ladder testing resumed. From that point onward, all locomotor tests were conducted bi-weekly at the same time of day by the same investigator until 27   113 wpi, when the animals underwent axonal tracing procedures, which precluded any further behavioural testing. Open field locomotion (BBB) Open field locomotion was scored on the BBB scale (Basso et al., 1995), and BBB subscale (Basso, 2004). Two raters assigned a score at the time of testing and all animals scored a 21 on the BBB and a 13 on the BBB subscore during pre-injury baseline assessments. CatWalk  The CatWalk system (Noldus, Netherlands), enables objective assessment of locomotion parameters based on quantitative footprint data (Hamers et al., 2001; Vrinten and Hamers, 2003). Unusable runs were identified based on the following criteria: 1) running at an inconsistent speed, 2) stopping in the middle of the run, 3) running exceptionally fast or slow. At least five runs were recorded for each rat during each testing session, and three runs were selected by a rater based on consistency of the run and the inclusion of three complete uninterrupted step cycles. The following parameters were examined: forelimb and hindlimb stride length, hindlimb paw angle, hindlimb paw width, hindlimb paw print intensity, and overall step sequence patterns. Irregular horizontal ladder Hindlimb stepping was also assessed using the irregular horizontal ladder (Metz and Whishaw, 2002). Animals were trained to cross the ladder toward their home cage   114 and each crossing was recorded using a high definition digital camera (Sony, Toronto, Canada) for subsequent scoring. The ladder always included the same number of overall rungs and spaces (ranging from one-five missing rungs per space), but the rung positions were changed for each testing session to avoid any training effects. A frame-by-frame analysis of video recordings of hindlimb stepping yielded error scores (averaged over five trials per session per animal) for the number of overall steps and errors for each hind paw (Metz and Whishaw, 2002).  3.2.7  Tract tracing and tissue processing At 27 wpi (two weeks prior to endpoint), all animals underwent one of two different tracing procedures, corticospinal tract (CST) or rubrospinal tract (RST) tracing as previously described (Hiebert et al., 2002; Kwon et al., 2002b); data not shown. Although there was no overt morbidity, axon tracing precluded any further behavioural testing.   Prior to euthanasia and transcardial perfusion, the bladder of each rat was emptied by manual expression. The time-of-transplant control group was euthanized at 8 wpi, whereas the SKP-SC- and medium-treated rats were euthanized at 29 wpi. All rats were overdosed with ketamine (210 mg/kg, i.p.) and xylazine (30 mg/kg, i.p.) and transcardially perfused with 0.12M PBS followed by 4% paraformaldehyde in PBS (0.1M). The thoracic spinal cord encompassing the injury site and the bladder were removed, post-fixed overnight in 4% paraformaldehyde, cryoprotected overnight in 12, 18, and then 24% sucrose in 0.12M PBS, frozen on dry ice and stored frozen at -80°C. The thoracic spinal cord was sectioned longitudinally in the sagittal plane and the   115 bladders were cut transversely, both at a thickness of 20 µm using a Microm cryostat (Heidelberg, Germany) onto Superfrost Plus slides (Fisher, Houston, TX) and stored at -80°C. The sections of the thoracic spinal cord from each rat were mounted as a series of 10 slides containing 10 spinal cord sections each, so that each slide held a section from every 200 µm through the spinal cord.   3.2.8 Immunohistochemistry Tissue sections were permeabilized with 0.1% Triton-X-100 and treated with 10% donkey serum for 30 min to prevent non-specific binding. For immunolabeling of myelin proteins, brief delipidation was also performed, using graded ethanol solutions prior to the blocking step. For a list of the primary antibodies used, see Table 3-1.  The secondary antibodies were generated in donkey or goat, conjugated with Dylight fluorochromes 405, 488, 594 or 649, and used at a concentration of 1:200 (Jackson ImmunoResearch Laboratories, West Grove, PA). In some sections, nuclei were stained with Hoechst 33342 (1:5000).   3.2.9 Histological quantifications A Zeiss (Oberkochen, Germany) Axioplan 2 microscope fitted with image acquisition software (Northern Eclipse; Empix, Mississauga, Canada) was used for low magnification images. For higher magnification images, including those used for cell counts, axon counts, and confirmation of co-localization, images were captured on a Zeiss AxioObserver Z1 (Zeiss, Germany) confocal microscope fitted with a CSU-X1 Yokogawa spinning disc and solid state lasers with 405, 488, 565 and 639 wavelengths.   116 All image analysis was completed by individuals blind to treatment. Associated with this, the green channel (with GFP+ cells) was not provided to the individuals assessing histological outcomes except when necessary; e.g., for GFP+ cell counts and volume analyses. Images were merged using Photoshop CS2 or CS4 (Adobe, San Jose, CA). All measurements (e.g., distances, areas, and intensities) except cell/axon counts and vector analysis were performed with Sigma Scan Pro 5 (Systat, Chicago, IL).  Among SKP-SC-treated rats, we defined a subgroup of five animals with very successful survival of transplanted cells (defined by more than 70,000 cells present; see Figure 3.2 e; Figure 3.3) and compared them to a matched (by injury force/displacement and pre-transplant behavioural parameters) medium-treated subgroup (n=5) and a matched time-of-transplant subgroup (n=5).  This subgroup analysis was conducted to see whether long-term survival of grafted cells is required for specific observed benefits (e.g., lesion volume and intact tissue width; Figure 3.4) or when performing counts on all 38 animals was not practical (axon and axon sub-type counts: Figure 3.6 g-i, Figure 3.7; SKP-SC orientation analysis: Figure 3.3). Determining lesion volume, average intact tissue, and tissue width All three analyses were done on tissue processed with antibodies to GFAP (reactive astrocyte marker) prior to imaging at 5X.  Lesion area was defined as GFAPNEG area or GFAP+ area with disrupted/abnormal cytoarchitecture and was manually outlined in each spinal cord section. Cavalieri’s principle was applied for volume calculations, i.e., V= S [area x section thickness x number of sections in each sampling block]. The same sections were used to estimate the amount of intact tissue; we measured the thinnest   117 combination of both 1) the spared rim representing the narrowest width of the rim with GFAP+ tissue with normal cytoarchitecture on either side of the lesion (as described above); 2) any additional normal-appearing cytoarchitecture resulting from spared tissue bridges. The narrowest width of spared tissue was summed within a section and the average sum from all sections of one animal yielded the mean intact tissue value for that animal. A subsequent width of spinal cord at the narrowest point was measured by drawing a one pixel thick line across the cord at its narrowest point. SKP-SC transplant volumes, counts, orientation, and proliferation Transplant volumes were examined in tissue sections processed with antibodies against GFP, P0, and GFAP. Images were taken at 5X and thresholds were set by the observer to yield an area occupied by the GFP+ SKP-SCs using Sigma Scan software. These GFP+ transplant areas were further subdivided into regions inside the lesion (defined above) versus outside the lesion and converted into transplant volumes using Cavalieri’s principle. The high-survival transplant subgroup (n=5) was used to estimate densities of the SKP-SCs within these transplants. Confocal z-stacks imaged at 63X were collected in random locations every 320µm apart through the transplants and the Hoechst stained nuclei embedded within GFP+ SKP-SCs were counted within the volume of the z-stacks using the optical dissector technique. The resulting GFP+ SKP-SC density was used to estimate the total number of SKP-SCs in the previously measured GFP+ volumes in SKP-SC-treated animals. The volume of SKP-SCs ranged from 0.0006 to 0.23 mm3 and the average density of cells was 5.325x10-4 cells/µm3.  In the same set of images, we   118 determined the percentage of GFP+ SKP-SCs that were also P0+, to yield the percentage of myelinating SKP-SCs.  The average orientation of the transplanted SKP-SCs was measured in the SKP-SC subgroup (n=5): three serial sections were selected and single plane images of the GFP+ area were analyzed with ImageJ software (National Institute for Health, USA). Each cell was assigned a vector representing the orientation of the cell and the deviation away from zero (rostral-caudal orientation) was calculated and compared between the rostral, middle and caudal sections of the cord. Cells that were observed in cross section (i.e. orthogonal to the plane of sections) were given a 90° value. The orientation values were compared to a value of 45 degrees (representing a random orientation of the cells in a 2D plane). An additional quantification of SKP-SC proliferation was performed in these three animals (with transplantation estimates >70,000 cells), by counting the GFP+ cells that co-expressed the proliferation marker Ki67 at 21 wpi. GFAP intensity analysis Intensity profiles of the GFAP immunoreactivity were generated along six (one pixel wide and 320 µm long; 5X primary magnification; Figure 3.5 a-c) lines perpendicular to the lesion edge. The animal averages were then averaged for each group and further broken down according to whether or not this drawn line (extended into the lesion) was directly on SKP-SCs (GFP+), directly on endogenous SCs (GFPNEG, P0+) or directly on cavity (with no SKP-SCs or P0+ endogenous SCs in the vicinity). In each situation, the intensity data was normalized to the far distant rostral intact cord for each given animal to correct for inter-animal variations.   119 SC myelination and non-myelinating SC analysis  The overall volume of SC myelin (P0+) was measured using a thresholding procedure as described for the GFP+ transplant volume in 5X images. The volume of P0+ myelin associated with transplant-derived SKP-SCs was estimated by measuring the overlap between GFP+ and P0+ areas on each section and converting those areas to volumes (as described above). All P0+ structures not closely associated with GFP+ SKP-SCs were assumed to be of endogenous SC origin. The myelination state was corroborated with antibodies, including Caspr and the voltage gated potassium channel Kv1.2., known markers surrounding the nodes of Ranvier. To examine the non-myelinating SC content of the 29 wpi spinal cord tissue, we used p75-NTR immunohistochemistry and conducted a volume estimate using the same methods as for GFP+ and P0+ volume analyses described above on an adjacent set of sections. Axon counts  Two axon analyses were conducted in the three subgroups (n=5 each). In the first analysis comparing the transplantation groups, for each animal, we counted the number of axons in a single 63X optical plane in five sections spaced 200µm apart, whereby the middle section contained the central canal. Axons were counted when crossing a line spanning the lesion (as defined previously) at the narrowest point of the spinal cord (blue lines; Figure 3.6 a, c, e).  In addition, we counted the number of axons, axon subsets, and/or P0+ myelinated axons within the GFP+ SKP-SC bridges at three lines drawn at the rostral (100µm from rostral interface), middle, and caudal (100µm from caudal interface)   120 portions of the GFP+ grafts (yellow lines; Figure 3.6 e) to further determine whether any gradients of axon growth exist, e.g. due to ingrowing supraspinal axons. Axons that intersected with the drawn lines were counted throughout the 20 µm section using the section with the largest SKP-SC transplant area which happened to be within 200 µm ventral to the central canal. Images for all axon counts were captured at 63X, including tyrosine hydroxylase (TH)+ axons, serotonin transporter (SERT)+ axons, calcitonin gene-related peptide (CGRP)+ axons, axons (bIII tubulin/NF-200), and P0+ axons (bIII tubulin/NF-200/P0+). Axons surrounded by P0+ sheaths were counted to calculate the percentage of myelinated axons within the SKP-SC bridges. Bladder analysis Wet weight of the dissected bladders was recorded. Bladders were then dabbed dry and a small 3 mm ring-shaped band was cut from the bladder starting at the rostro-caudal midline moving caudally and frozen in Tissue-Tek O.C.T. (Sakura, Netherlands) in an orientation that allowed for transverse sectioning of the bladder wall. Bladders were handled carefully in dissection and preparation to avoid stretching the tissue; also, tissue preparation was performed by blinded experimenters. Four animals were excluded from the bladder analysis due to severe damage to the bladder during tissue processing. Sections were stained for 5 minutes in 0.1% Cresyl Violet and subsequently imaged at 10X. Two sections were measured per animal (2000µm apart), where a one pixel thick line was drawn through the thickest section of the bladder wall to obtain a mean wall thickness.   121  3.2.10 Statistical analyses Statistics were calculated using SPSS (IBM; Markam ON, CA). If data met assumptions of normality and homogeneity of variance, groups were compared using the appropriate independent t-test, paired t-test, or a repeated measures (RM) analysis of variance (ANOVA) for two groups and using a one-way ANOVA with a Least Significant Difference post hoc test for three groups with data presented as mean ± SEM.  If the assumptions of normality and homogeneity of variance were not met: groups were compared using the KW one-way analysis of variance with follow up MWU-test for three independent groups and just the MWU-test for two paired groups or a Friedman test with follow up Wilcoxon test for three paired groups and just the Wilcoxon test for two paired groups with data presented as raw values with the median indicated. Correlation analysis was conducted using Pearson’s correlation coefficient (parametric) or Spearman’s rank correlation coefficient tests (non-parametric). The significance level for all two-tailed tests was p<0.05. Trends were reported for two-tailed tests when p<0.1 indicative of a significant one-tailed test.  3.3 Results  3.3.1 Skin Derived Precursors (SKPs) generate high purity SCs (SKP-SCs) in vitro The experimental timeline is summarized in Figure 3.1 a. Prior to transplantation we confirmed the identity and purity of putative neonatal SKP-SCs by triple-label immunocytochemistry for GFP and SC markers p75-NTR and P0 (Figure 3.1 b) or S100b   122 and GFAP (Figure 3.1 c). The clear majority of cells in these cultures were positive for these markers (Figure 3.1 b & c) and cell counts indicated that 98.4% of the GFP+ neonatal cells expressed S100b (Figure 3.1 d).    3.3.2 Transplanted SKP-SCs survive and bridge the lesion in chronic SCI   Eight weeks after SCI, the lesion site was prominent, and filled with macrophages and GFAP+ tissue strands indicative of reactive astrocytes (Figure 3.2 a).  At 29 wpi, lesion sites in medium-injected rats were characterized by large empty cavities that were walled off by a sharp border of GFAP+ astrocytes (Figure 3.2 b). In contrast, in animals that received SKP-SCs, the lesion site was spanned by bridges of GFP+ cells (Figure 3.2 c). In sections immunostained with GFP enhancer (e.g., Figure 3.2 d) and with nuclear staining (Hoechst), cell counts revealed that GFP+ cells were present in the spinal cord in all animals, although there was a high degree of variability in cell numbers. Eight of 15 animals that received SKP-SCs contained 70-120,000 GFP+ cells, whereas four animals had 20-70,000 cells, and three rats had fewer than 20,000 cells at 21 weeks post-transplantation (Figure 3.2 e). The majority of the SKP-SCs displayed predominant rostral-caudal orientation deviating (on average) not more than 25-30 degrees from the rostro-caudal axis of the spinal cord (Figure 3.3 g, h).  Immunostaining for Ki67 indicated minimal proliferation of the transplanted cells (less than 0.1%; Figure 3.3 i-k).    123 3.3.3 SKP-SCs prompted an increase in intact tissue surrounding the site of chronic SCI  We defined the lesion site as tissue with absent or disrupted cytoarchitecture as revealed by GFAP-immunostaining of the host astrocytes. Hence, the lesion volume measures the destroyed and/or grossly abnormal spinal cord tissue, including the cavity volume. To assess the amount of intact tissue surrounding the SCI site, we measured residual GFAP+ spinal cord tissue with intact-appearing cytoarchitecture in the spared rim as well as within the lesion. At 29 wpi, both SKP-SC- and medium-treated animals had significantly more intact tissue at the lesion epicentre than time-of-transplant controls euthanized at 8 wpi (Figure 3.2 g), suggesting an expansion of intact tissue between 8 and 29 wpi. This spontaneous repair was augmented in animals with high survival of SKP-SCs (Figure 3.4 b).   3.3.4 SKP-SCs integrate into spinal cord tissue, mitigate the formation of the glial scar, and provide a permissive axon growth substrate  A major contributor to regeneration failure after SCI is the glial scar (Silver and Miller, 2004; Filous and Silver, 2016). At 8 wpi, a loose mesh of astrocytic processes bordered and extended into the site of SCI, and the lesions were commonly filled with macrophages and microglia (Figure 3.5 a). By 29 wpi, in the medium control group, most macrophage/microglia had cleared, and the lesions were characterized by “empty” cavities, presumably fluid-filled in vivo, with sharply demarcated borders exhibiting elevated GFAP-expression (Figure 3.5 b, k, l). In contrast, at 29 wpi in the SKP-SC   124 group, host astrocytes extended multiple fine processes into the SKP-SC transplant, particularly at the rostro-caudal boundaries of the lesion site (Figure 3.5 c-f, k and l).  Densitometric analysis (along the white line spanning the cavity-parenchyma interface in Figure 3.5a-c) revealed significantly higher GFAP expression in both 29 wpi groups compared to the time-of-transplant control group, indicating that reactive gliosis increases over time following SCI (Figure 3.5 g, h). Progressive gliosis was mitigated in SKP-SC-treated animals; GFAP expression was reduced compared to medium-only controls (Figure 3.5 g, h). In SKP-SC-treated rats, GFAP-intensity was high adjacent to regions of cavitation and close to endogenous SCs, and reduced in astrocytes neighbouring SKP-SCs (Figure 3.5 i; quantified in Figure 3.5 j). These effects of SKP-SCs delivered in chronic SCI are reminiscent of our observations following subacute transplantation of SKP-SCs at 7 dpi (Biernaskie et al., 2007).  We then asked whether SKP-SCs provided a more permissive growth substrate. In medium-treated controls, laminin, which is expressed by SCs in peripheral nerves, was typically restricted to blood vessels in the spinal cord and to the host SCs present in the spared tissue margin of the lesion site (Figure 3.5 k; top row). In contrast, in SKP-SC-treated animals, the spinal cord tissue displayed extensive laminin immunoreactivity. Laminin was expressed both within the transplants and in the host tissue bordering the SKP-SC bridges where many of the laminin+ cells were GFPNEG endogenous SCs expressing p75-NTR (Figure 3.5 k; bottom row).  Integration of SKP-SCs into host spinal cord tissue and reduced GFAP-immunoreactivity led us to ask whether these differences reflected a change in the inhibitory nature of the glial scar. When we examined other inhibitory proteins associated   125 with scar formation, including the proteoglycan neurocan as well as the chondroitin sulphate proteoglycan epitope recognized by CS-56, both proteins were expressed in a sharply demarcated boundary around lesion cavities in the medium controls (Figure 3 l; top row), but not at the interface of SKP-SCs and host tissue (Figure 3 l; bottom row).   3.3.5 SKP-SCs promote growth/regeneration of host axons   To determine whether SKP-SC transplantation enhances axon growth, we first examined spinal cord sections immuno-labeled with antibodies against large- and small- calibre axons (NF-200/βIII-tubulin) in animals with high cell survival (above the grey line in Fig 2.3 e). At 8 wpi, some spared host tissue bridged the lesion site and contained small bundles of axons running in a rostro-caudal orientation (Figure 3.6 a, b). By 29 wpi, lesions in medium-treated controls were characterized by tightly demarcated cavities with only rare host tissue septae still containing some axons (Figure 3.6 c, d). In contrast, SKP-SC-treated rats with high numbers of surviving SKP-SCs contained many axons extending through SKP-SC bridges, most of which also maintained a rostro-caudal orientation (Figure 3.6 e, f, and m). The average number of axons within a single confocal plane at the lesion-epicentre of the spinal cord (counted along the blue stippled lined in Figs. 3.6 a, c, e; i.e., with the spared rim included) was less than 140 in medium control animals at 29 wpi. There were twice as many axons present after SKP-SC transplantation, mostly in the cell bridges crossing the lesion (Figure 3.6 g).   When we examined specific populations of axons in animals with high cell survival, more TH+ and SERT+ axons were found at the lesion site in the SKP-SC-treated animals compared to both control groups (Figure 3.6 h-l, n). Predictably, there   126 were more TH+ and SERT+ axons at the rostral and middle levels of the SKP-SC bridges than the caudal level (counted at the yellow lines shown in Figure 3.6 e), reflecting their origin from the brainstem. Numerous SERT+ and TH+ axons had advanced to the caudal host interface and a small number crossed this interface and entered the caudal spinal cord where they appeared to stop within several hundred microns (Figure 3.6 j, l). The tortuous appearance of those axons, taken together with the rostro-caudal gradient in their density, indicates that these monaminergic axons had grown/regenerated through the SKP-SC bridges. There were no differences in the total number of axons (NF-200/ βIII-tubulin) or CGRP+ axons across these rostro-caudal levels (Figure 3.7). Of note, peptidergic sensory axons expressing Substance P or CGRP represented < 3% of all the axons in these grafts (Figure 3.7). SKP-SCs myelinate axons in chronic SCI  To determine whether transplanted SKP-SCs myelinated axons that had grown into the grafts, we performed triple labeling for GFP, P0, and axons (NF-200/βIII-tubulin; Figure 3.8 a-c). We found clear evidence of myelination by SKP-SCs. At 29 wpi, 73% of the SKP-SCs were myelinating, as defined by the formation of a thin GFP+ cytoplasmic layer surrounding P0+ myelin ensheathing a NF-200+ or βIII-tubulin+ axon.  Myelination was further confirmed by immuno-labeling for Kv1.2 (Figure 3.8 d, e), the main potassium channel in the juxtaparanodal axon membrane (Rasband et al., 1998) and Caspr (Figure 3.8 f, g), which contributes to the septate junctions between the axon and the paranodal loops of myelin-forming SCs (Einheber et al., 1997). Moreover, 52% of the axons that had grown into the SKP-SC bridges were wrapped in P0+ myelin.    127  3.3.6 SKP-SC transplantation augments spontaneous CNS repair by endogenous SCs                                                                                                                                       In our previous studies (Biernaskie et al., 2007; Sparling et al., 2015), SKP-SC transplantation prompted an increase in endogenous myelinating SCs occupying the spinal cord. To address the endogenous SC response in chronic SCI, we examined spinal cord sections for GFPNEG (i.e., host) SCs, both P0+ cells (myelinating) and p75-NTR+ (non-myelinating).  P0+ SC myelin was often encountered in the lesion walls and the residual tissue strands in and around the lesions in the time-of-transplant (Figure 3.9 a) and medium (Figure 3.9 b & f) control groups. However, more P0+ myelin sheaths were found in animals that received SKP-SCs (Figure 3.9 c, d, g). In SKP-SC-treated animals, myelin sheaths were SKP-SC-derived (GFP+) or host SC-derived (P0+ GFPNEG); there were more host SC-derived myelin sheaths in SKP-SC-treated animals, even when SKP-SC survival was limited (Figure 3.9 d). There was no correlation between the GFP+ volume and the total P0+ volume (r=0.030, p=0.916) in the SKP-SC-treated group. This suggests that even the transient presence of SKP-SCs within the injured spinal cord enhanced the endogenous SC repair response. GFP+ SKP-SCs accounted for a relatively small proportion of the P0 myelin by volume, as only 9.6% of the overall P0 volume in animals that received SKP-SCs overlapped with GFP+ cells (green portion of bar in Figure 3.9 i). This indicates that endogenous SCs generated the clear majority of P0 myelin found 29 weeks after SKP-SC transplantation (black portion of bar in Figure 3.9 i).    128 We also analyzed the location of SC-derived myelin in all groups, and found that both the time-of-transplant and medium control groups had more endogenous SC myelin in the spared tissue than in the lesion site (Figure 3.9 j). In contrast, the SKP-SC group exhibited similar amounts of SC myelin in both locations. SKP-SC-treated animals had a larger P0 volume inside the lesion site than either control group, and a larger P0 volume outside the lesion compared to the time-of-transplant group (Figure 3.9 j). Interestingly, the amount of intact tissue positively correlated with overall P0 volume (r=0.502, p<0.001), P0 volume inside the lesion (r=0.429, p=0.009), and P0 volume outside the lesion (r=0.557, p<0.001), suggesting that sparing or survival of host tissue is associated with greater SC myelin content in the injured spinal cord (Table 3-2). P75-NTR is highly expressed in non-myelinating SCs. As seen for P0, there were many bright p75-NTR+ cells with spindle shaped SC morphology in the spared host tissue of medium-injected and SKP-SC transplanted animals (29 wpi; data not shown), particularly in tissue bridging the lesion site in animals that received SKP-SCs (Figure 3.9 e & h). However, most of the p75-NTR+ cells in the tissue rim were GFPNEG endogenous SCs, whereas those in the lesion bridges were often GFP+ transplanted cells (Figure 3.9 h). Approximately 9% of the total P75-NTR+ volume was GFP+. Quantitative examination of p75-NTR expression bore results like those for P0; the p75-NTR volume was correlated with the amount of intact tissue (r=0.388, p= 0.042); suggesting that increased intact host tissue is associated with enhanced SC content in general, not just increased SC myelination.     129 3.3.7 SKP-SCs from adult skin show similar behaviour to neonatal-derived SKP-SCs                                               To extend the clinical relevance of our findings, we performed pilot transplants (n=2) in rats with chronic SCI using adult-derived SKP-SCs to determine whether the outcome would be qualitatively different from our neonatal-derived transplanted animals. Prior to transplantation, we confirmed the identity and purity of the adult SKP-SCs by immunocytochemistry for GFP and the SC markers p75-NTR and P0 (Figure 3.10 a) or S100b and GFAP (Figure 3.10 b). Cell counts revealed that 99% of the GFP+ neonatal cells expressed S100b. Like the SKP-SCs of neonatal rats, approximately 10% of transplanted adult-derived SKP-SCs persisted at 21 weeks post-transplant. These integrated into host tissue, provoked little reactivity in host astrocytes, stimulated substantial growth/regeneration of axons, including TH+ fibers, and produced SC myelin while stimulating the endogenous SC response (Figure 3.11 a-g).  In these pilot experiments, the behaviour of adult-derived SKP-SCs was like those derived from neonates.  3.3.8 Transplantation of SKP-SCs eight weeks following SCI resulted in better locomotor outcomes   Having established that SKP-SC transplants promote histological repair in chronic SCI, we asked whether these cells also enhanced functional locomotor recovery. The BBB scores in all groups showed spontaneous locomotor recovery from the time of contusion SCI to the time -of-transplant, and no functional decline was observed in the weeks following SKP-SC transplantation or medium injection. A RM-ANOVA indicated   130 that the difference between the medium and SKP-SC groups changed over the last 16 weeks of the study with a significant group difference at week 19 and 21 (Figure 3.12). The amount of P0 expression outside the lesion was correlated with the BBB scores at 27 wpi (r=0.648, p=0.009).  To complement the open field measurements, we performed gait analysis. Average forelimb stride length was ~140 mm pre-injury, dropped to 100 mm post-injury, and remained at that value in the medium group at 26 wpi. In contrast, in rats receiving SKP-SC, stride length recovered partially to 108 mm at 26 wpi. RM-ANOVA revealed that the difference between the medium and SKP-SC groups changed for both forelimb and hindlimb stride length (normalized to pre-treatment values) over the last 16 weeks of the study with a significant group difference indicated at each time point denoted with an asterisk. This indicates that the SKP-SC group could maintain longer forelimb and hindlimb stride lengths compared to the medium only group, which showed a further decline in the hindlimbs. SKP-SCs treatment also resulted in a reduced angle of hindlimb paw rotation and a decreased frequency of abnormal gait patterns compared to the medium only group (Figure 3.13 a & b). No significant differences were detected on hindlimb print width or intensity parameters, the BBB subscore, or the horizontal ladder analysis (Figure 3.12 c-f; respectively). In a subsequent study with a similar experimental timeline, we asked whether SKP-SCs transplantation had any effect on the development of neuropathic pain. We demonstrated no difference in either mechanical or thermal sensitivity between the transplanted and medium-injected control groups (data not shown) suggesting that transplanted SKP-SCs had no impact on sensitivity to sensory stimuli.    131 3.3.9 SKP-SC transplantation reduces bladder pathology observed after SCI  Autonomic dysfunction is a major concern following SCI and improving bladder function has been ranked amongst the most important priorities for people living with SCI (Anderson, 2004). Although all rats regained spontaneous micturition, we conducted post-hoc histological analysis of the bladder wall to examine the effects of SKP-SC transplantation on bladder pathology. Qualitative examination revealed bladder wall “thickenings” in the medium treated animals (Figure 3.12 d) that were absent in animals transplanted with SKP-SCs (Figure 3.12 e). Measurements of the thickest point revealed that bladder walls of medium-treated rats were almost twice as thick as SKP-SC-treated rats (Figure 3.12 f). These bladder wall thickenings are typical of bladder-sphincter dyssynergia and/or a lack of inhibitory supraspinal input causing detrusor over-activity (de Groat et al., 1990; Pikov and Wrathall, 2001; Tai et al., 2006). Here, SKP-SC treatment prevented the formation of pathological thickenings of the bladder wall after contusion SCI and the bladder wall thickness was negatively correlated with both BBB and BBB subscore (Spearman’s rho= -0.414 and -0.425, p=0.036 and 0.030, respectively Figure 3.12 g & h). For a full list of relevant correlations, see Table 3-2.         132 3.4 Discussion  We demonstrate that SKP-SCs transplanted 8 weeks after a thoracic spinal cord contusion survive for more than 5 months at the site of chronic SCI, integrate within spared host tissue, and promote several types of neural repair, including axonal growth/regeneration and SC myelination. Endogenous SCs exhibited a substantial reparative response, which was enhanced by transplanted SKP-SCs. Importantly, SKP-SC transplantation at chronic stages of SCI spurred growth/regeneration of supraspinal axons into and through the bridges of grafted cells, improved locomotor function, and reduced bladder pathology. Together, our findings suggest that SKP-SCs represent a promising intervention for SCI, with the potential to confer benefit in the chronic setting.  At 21 weeks post transplantation, SKP-SCs survived and integrated into the injured spinal cord. Our measurements indicating that up to 12% SKP-SCs survived long term likely represents an underestimate, as we did not account for cells remaining in the injection needle or lost due to reflux; previous estimates suggest that these reduced the total number of injected cells by 20% (Hill et al., 2006). The extensive integration of SKP-SCs in our 8-week delayed transplantation was unexpected and prompted our analysis of an 8 wpi time-of-transplant group to better understand the environment encountered by the transplanted SKP-SCs. At 8 wpi the lesion was full of mononuclear cells, lacked a dense glial scar, and contained numerous GFAP+ processes as well as small bundles of axons that ran rostro-caudal through the site of injury on either site of the ventral fissure. Our 8-week-old lesions differ from those described by other authors who reported a more sharply demarcated cavity (Wrathall et al., 1985; Bresnahan et al., 1991). This variability across studies may be due to methodological differences as the speed and severity of injury, animal gender, and type of impactor used differed across laboratories. The   133 substantial number of rostro-caudal oriented spared axons, together with astrocytic processes, may have provided a bridging scaffold along which the transplanted SKP-SCs aligned. SKP-SCs cellular bridges, also with a rostral-caudal orientation, likely facilitated both the growth/regeneration of local axons and the accumulation of endogenous SCs. Supporting this notion, the axons within the lesions containing transplanted SKP-SCs were frequently myelinated by endogenous SCs as well as transplanted SKP-SCs, and those axons could be found in either GFAP+ or GFAPNEG regions of the lesion environment.   Axons are well known to grow into and within SC grafts but the ‘off-ramp’ issue is well documented where axons struggle with leaving the graft and growing back into the astrocyte rich host parenchyma. Despite the disruption of normal GFAP cytoarchitecture at the lesion site, SKP-SC processes and astrocyte processes were frequently found near one another, particularly at the transplant host interface, and this arrangement appeared to provide a permissive growth environment for axons entering and exiting the SKP-SC grafts. These findings are reminiscent of the permissive SC graft/spinal cord interfaces described previously after SC transplantations in combination with MatrigelR (Williams et al., 2015) or with expression of NG2 (Deng et al., 2011; Deng et al., 2013). The interactions between the GFAP+ astrocyte processes and SKP-SCs observed in this study suggest the formation of a permissive growth environment that may alleviate some of the ‘off-ramp’ associated issues observed after SCI. Compared to endogenous SCs, transplanted SKP-SCs induced lower expression of GFAP in adjacent astrocytes, and the low GFAP expression in the walls of the lesion sites plus the extensive expression of laminin beyond the confines of SKP-SC transplants likely contributed to a favourable axon-growth environment. The permissive nature of the transplant environment was illustrated by the thousands of axons found in our SKP-SC bridges, including axons of brainstem   134 origin based on TH and SERT immunolabeling. There was no evidence of CST axon regeneration which was not surprising in the light of previous studies reporting the reluctance of CST axons to grow into SC environments in the absence of co-treatments (Takami et al., 2002; Biernaskie et al., 2007; Pearse et al., 2007; Schaal et al., 2007; Hill et al., 2012); (for review Fortun et al., 2009; Bunge and Wood, 2012).  The exit of TH+ axons at the caudal transplant-host interface was encouraging, considering that very few studies have shown growth/regeneration of supraspinal axons without the addition of co-treatments such as olfactory ensheathing cells, Rolipram, trophic factor cocktails or their overexpression by the transplanted cells (Takami et al., 2002; Pearse et al., 2004b; Kanno et al., 2014; Kadoya et al., 2016). However, the connectivity of the TH+ axons in our study and their contribution to functional recovery remains unclear.  The increase in the amount of intact tissue between week 8 and 29 was unexpected, and this increase was even more pronounced in animals with high SKP-SC survival. At present, it remains unclear whether this finding is a result of neuroprotection or neural repair mechanisms. In light of the protracted oligodendrocyte death observed (Crowe et al., 1997) and the increase in P0 myelination between week 8 and 29  (Figure 3.9 i), we hypothesize that the increased thickness of the spared rim is due in part to myelination by SCs as SC myelin takes up significantly more space than oligodendrocyte myelin (Kocsis and Waxman, 2007). Moreover, the addition of axons by axonal growth/regeneration may also contribute to increased thickness of the spared rim which is supported by the higher number of axons found in the SKP-SC (29 wpi) compared to medium control (29 wpi) and the time-of-transplant control (8 wpi) groups (Figure 3.6 g).  In addition, we cannot rule out the possibility that the second surgery (transplantation) at 8wpi triggered a secondary wave of injury factors and cytokines that   135 triggered SC responses and axonal sprouting leading to larger intact rims in medium controls at 29wpi and even larger intact rims in SKP-SC transplanted rats at 29 wpi (Figure 3.2 g, Figure 3.4).  The changes in GFAP astrocyte reactivity, as well as the ongoing increase in SCs over time suggest that an 8-week-old contusion site in the rat may not have matured to a ‘chronic’ status as defined by lesion stability. These long-term changes following SCI warrant further investigation to accurately define the post-injury scenario and better understand ongoing endogenous repair responses. These findings are also highly pertinent to the ongoing debate regarding the appropriate timing of ‘chronic’ interventions in animal models that aim to identify appropriate treatments for chronically injured humans (Kwon et al., 2010a).  Few pre-clinical cell transplantation studies have been performed in chronic spinal cord contusion or compression injuries (Tetzlaff et al., 2011); and arguably these lesion types mimic the clinically occurring spinal cord injuries (Kwon et al., 2010a). No behavioural improvements have been observed following transplantation of neural stem cells (Karimi-Abdolrezaee et al., 2006; Parr et al., 2007; Karimi-Abdolrezaee et al., 2010; Nutt et al., 2013; Jin et al., 2016), olfactory ensheathing cells (Barakat et al., 2005; Zhang et al., 2011) or human embryonic stem cell derived oligodendrocyte precursors (Keirstead et al., 2005) when those cells have been used without co-treatments in ‘chronic’ pre-clinical settings (i.e. close to 2 months or more after injury). There are reports of improved locomotion with a human neural stem cell line transplanted into NOD-scid mice at 30 days after contusion (Salazar et al., 2010) but only discrete effects on gait (base of support) were observed at 60 days post-contusion (Piltti et al., 2013b); this neural stem cell line entered human clinical trial but this was recently stopped due to a combination of small effect sizes and financial considerations (;   136 NCT01321333).  In contrast, mouse neural stem/progenitor cells transplanted 7 weeks after clip compression injury in rats required combinatorial administration of chondroitinaseABC, minocycline, plus a cocktail of EGF, FGF-2, and PDGF to achieve notable functional improvements (Karimi-Abdolrezaee et al., 2010). Taken together, the available data indicate that realizing functional improvements after long treatment delays is extremely challenging, particularly for cellular therapies delivered without co-treatments. Considering this, the recovery seen here with a single cell therapy, SKP-SCs, applied at 8 wpi is remarkable. To our knowledge, only one other chronic (approximately 8 wpi) spinal cord contusion study has demonstrated convincing efficacy in terms of open field locomotor recovery using a single cell therapy in the absence of co-treatments, and that was the study, and subsequent replication study, using SCs generated from peripheral nerve (Barakat et al., 2005; Sharp et al., 2012b). In addition to effects in the open field, our study found benefits in gait performance and improvements in bladder pathology.  The present study indicates that SKP-SCs are a promising candidate cell for SCI repair and in addition have many practical advantages. Most individuals undergo surgery to stabilize the spinal column within a day or two after a SCI (Fehlings et al., 2012). This procedure offers a convenient opportunity to harvest skin from the edge of the incision, which provides a source of SKPs that could generate sufficient SKP-SCs for autologous therapeutic application within 6-8 weeks. Delaying treatment by 6-8 weeks would also allow patients more time to contemplate consent for clinical trials (Illes et al., 2011) and enable more accurate predictions of long-term functional outcomes, which would reduce the number of patients required to test the efficacy of those cells in clinical trials (Fawcett et al., 2007).    137 The efficacy observed here following SKP-SC transplantation at 8 wpi suggests that SKP-SCs represent a promising treatment option for the growing group of people who are living with chronic SCI. Furthermore, the clinical safety and feasibility of autologous SCs as a therapy for chronic SCI has already been established using nerve-derived SCs (Saberi et al., 2008; Saberi et al., 2011) and a clinical trial for those cells transplanted sub-acutely has been completed by Bunge and colleagues (; NCT01739023) and the group has now initiated a chronic trial in combination with rehabilitation (; NCT02354625).  Recently, nerve-derived SCs and SKP-SCs were compared side-by-side at the transcriptome level and only discrete differences between these cell types were detected (Krause et al., 2014). Here, we also reported preliminary findings from a small number of animals transplanted with SKP-SCs generated from adult rodent tissue. Adult and neonatal SKP-SCs were indistinguishable in our hands, which suggests that adult SKP-SCs are likely to provide similar therapeutic efficacy as that seen here with neonatal SKP-SCs. However, future experiments that fully address the safety and efficacy of adult rat and human SKP-SCs will be required to determine whether autologous SKP-SC transplantation merits a clinical trial for the treatment of SCI. The present work represents an important milestone toward the development of an autologous transplantation protocol to treat chronic SCI with SCs generated from the skin, a highly accessible and available alternative to peripheral nerve.       138 Table 3-1. List of primary antibodies used Antibody host, source and concentration specific to immunocytochemistry (ICC) or immunohistochemistry (IHC) are shown.  Antibody Source Concentration mouse anti-S100β Sigma-Aldrich, St. Louis, MO 1:500 rabbit anti-GFAP Dako, Glostrup, Denmark 1:1000 IHC 500 ICC goat anti-GFAP Santa Cruz Biotechnology, Santa Cruz, CA 1:50 mouse anti-GFAP Sigma-Aldrich, St. Louis, MO 1:500 chicken anti-P0 Aves, Tigard OR 1:100 IHC 1:500 ICC chicken anti-GFP Millipore, Billerica, MA 1:1000 rabbit anti-GFP Millipore, Billerica, MA 1:100 goat anti-GFP Rockland Immunochemicals, Gilbertsville, PA 1:200 mouse anti-GFP Millipore, Billerica, MA 1:500 mouse anti-neurofilament 200 (NF-200) Sigma-Aldrich, St. Louis, MO 1:500 rabbit anti-NF-200 AbD Serotec, Releigh, NC 1:1000 mouse anti-beta-3 tubulin (ßIII tubulin) Sigma-Aldrich, St. Louis, MO 1:500 rabbit anti-bIII tubulin Covance, Princeton, NJ 1:500 rabbit anti-serotonin transporter (SERT) Immunostar, Hudson, WI 1:500 sheep anti-tyrosine hydroxylase (TH) Millipore, Billerica, MA 1:200 rabbit anti-calcitonin gene-related peptide (CGRP) Sigma-Aldrich, St. Louis, MO 1:500 rabbit anti-substance P Millipore, Billerica, MA 1:500 mouse anti-p75-NTR Millipore, Billerica, MA 1:500 IHC 1:500 ICC rabbit anti-laminin Sigma-Aldrich, St. Louis, MO 1:200 mouse anti-neurocan Developmental Studies Hybridoma Bank, Iowa City, IO 1:100 mouse anti-chondrontin sulfate proteoglycan (CS56) Sigma-Aldrich, St. Louis, MO 1:200 mouse anti-KV1.2 potassium channels Generous gift from Dr. J. Trimmer, University of California, Davis, CA 1:200 mouse anti-contactin-associated protein (Caspr) Generous gift from Dr. J. Trimmer, University of California, Davis, CA 1:300 mouse anti-Ki67 BD Biosciences, Mississauga, ON 1:20          139  Table 3-2. List of relevant compared significant correlations  Compared Parameters Correlation Coefficient P Value Intact Tissue vs. P0+ve Volume r=0.502 * p<0.001 Intact Tissue vs. P0+ve Volume Inside Lesion r=0.429 * p=0.009 Intact Tissue vs. P0+ve Volume Outside Lesion r=0.557 * p<0.001 Intact Tissue vs. Total Axons r=0.782 * p=0.001 ϕ Intact Tissue vs. P75NTR+ve Volume r=0.388 * p=0.042 Intact Tissue vs. SERT+ Axons r=0.804 * p=0.000 ϕ Intact Tissue vs. TH+ Axons r=0.720 * p=0.004 ϕ Intact Tissue vs. Lesion Volume r=-0.469 * p=0.004 P0+ve Volume Outside Lesion vs. 27wpi BBB ρ =0.648 * p=0.009 P0+ve Volume vs. Total Axons r=0.492 + p=0.063 ϕ P0+ve Volume vs. SERT+ve Axons r=0.512 + p=0.051 ϕ P0+ve Volume vs. TH+ve Axons r=0.463 + p=0.080 ϕ P75-NTR+ve Volume vs. SERT+ve Axons r=0.677 * p=0.031 ϕ P75-NTR+ve Volume vs. TH+ve Axons r=0.739 * p=0.015 ϕ BBB 27wpi vs. Ladder % Error 26wpi ρ =-0.358 + p=0.052 GFAP Max Value vs. Cavity Volume r=0.448 * p=0.006 Bladder Wet Weight vs. 27wpi BBB ρ =-0.501 * p=0.004 Bladder Wet Weight vs 27wpi BBB subscore ρ =-0.582 * p<0.001                140  Figure 3.1 Neonatal rat SKP-SCs express typical SC markers in vitro  (a) Experimental timeline of injury, transplantation, pre- and post-injury behavioural testing, and endpoints. Photomicrograph of cultured GFP-labelled (green) neonatal rat SKP-SCs immunostained for the SC markers p75-NTR and P0 (b), or S100b and GFAP (c), as well as Hoechst (nuclear marker; turquoise). (d) In vitro low power photomicrographs of GFP+ (green) SKP-SCs co-labelled with S100b and Hoechst. Scale bars: 20µm in c and c, and 100µm in d.      141     Figure 3.2 SKP-SC grafts survive in vivo in the chronically injured spinal cord (a-c) Photomicrographs of GFAP-immunostained contused spinal cords from a time-of-transplant control animal (a; 8wpi), a medium injected control animal (b; 29wpi), and a SKP-SC transplanted animal (c; 29wpi). (a) The lesion site is not an empty cavity at the 8wpi time of transplantation but rather filled with mononuclear cells and some residual GFAP+ host tissue (arrows). (b) A large fluid-filled cavity surrounded by GFAP+ host parenchyma is typical in the medium control group at 29 wpi. (c) GFP+ SKP-SCs integrate into the lesion sites and bridge the cavity.  (d) High-magnification image of GFP+ SKP-SCs with Hoechst staining representative for our images used for cell counts. (e) SKP-SC counts at 29 wpi for all transplanted rats. (f) The mean lesion volume was similar among the three main groups (ANOVA). (g) Mean thickness of intact GFAP+ tissue per group. Note that the time-of-transplant control group (8 wpi) had significantly less intact tissue than the medium (* p=0.028; ANOVA with least significant difference post-hoc test[LSD]) and SKP-SC (*p=0.003; ANOVA with LSD) groups (29 wpi). For f and g: SKP-SC n=15; Medium n=13; Time-of-transplant n=8; data represented as mean +/- SEM.  Scale bars: 200 µm in a-c; 10 µm in d.            142    143     Figure 3.3 SKP-SCs show a predominantly rostral-caudal orientation and a low incidence of proliferation at 21 weeks post transplantation Photomicrographs of GFAP-immunostained contused spinal cords from the five animals that make up the large transplant SKP-SC ‘subgroup’ (see methods) at 29 wpi used for quantification in Figure 3.3, 3.4, 3.6, and 3.7. Note that the SKP-SCs integrate into the lesion sites and bridge the cavity. (f) SKP-SC counts at 29 wpi for the rats in the large transplant subgroup. Each bar corresponds to the appropriate image shown above (a-e) and is a subset (>70000 cells) of those shown in Figure 3.2 e. (g) Photomicrograph of box shown in (a) demonstrating the associated vectors assigned to each transplanted GFP+ SKP-SC in the orientation analysis. Note that ‘x’ marks cells that were given a 90-degree value signifying they were coming out of the plane. (h) Quantification demonstrating that the majority of transplanted SKP-SCs displayed a strong rostral-caudal orientation at the rostral, middle and caudal regions of the graph as demonstrated by an average deviation from rostral–caudal axis (0 degrees) of 26 degrees. The rostral, middle and caudle deviations were significantly less than 45 degrees (representative of an average random orientation; *p<0.027; t-test; n=5). (i) Photomicrograph demonstrating the low number of transplanted GFP+ SKP-SCs that co-localized with the proliferative marker Ki67 at 29 wpi. (j) Confocal image taken at the location of the box in I demonstrating clear co-localization of Hoechst and Ki67 in the nucleus of a GFP+ SKP-SC. (k) To assess proliferation of the transplanted SKP-SC at 29 wpi, we quantified the percentage of GFP+ cells that expressed the marker Ki67. Only 0.05% of the cells were Ki67+, suggesting minimal proliferation of SKP-SCs at 21 weeks after transplant.  This Ki67 proliferative index is lower than in benign Schwannomas (average of 1.2% Ki67+ cells) and malignant peripheral nerve sheath tumours (average of 23% Ki67-positive cells; Ghilusi et al., 2009). Accordingly, there was no indication of SKP-SC derived malignancy in our grafts. Scale bars: 200 µm in a-e, 50 µm in g & i, and 20 µm in j. Data are presented as individual data for f (n=5) and k (n=3) and group means +/- SEM for h (n=5).     144            145     Figure 3.4 High SKP-SC survival results in significantly more intact tissue than in matched control animals  (a) The mean lesion volume for animals with high transplant cell survival and their appropriate controls is similar among the three groups. (b) Mean intact GFAP+ tissue (spared rim and tissue bridges included) showed significant differences among the subgroups. Note the significantly larger intact tissue in the SKP-SC subgroup with high transplant survival compared to the medium (*p=0.029; ANOVA with LSD) and time-of-transplant control (*p=0.001; ANOVA with LSD) subgroups. Also, the amount of intact tissue in the medium injected subgroup compared to the time-of-transplant control subgroup approached statistical significance (+p=0.051; ANOVA with LSD). Subgroup analysis: SKP-SC n=5; Medium n=5; Time-of-transplant n=5. All data presented as mean +/- SEM.          146    Figure 3.5 SKP-SCs mitigate the formation of the chronic glial scar  (a-c) Photomicrographs showing the differences in astrocyte organization and reactivity in the (a) time-of-transplant, (b) medium injected and (c) SKP-SC groups. (d) Same as 3 c together with the GFP+ SKP-SCs (green). Note that the lesion area to the left of the graft (star) was filled with endogenous myelinating and non-myelinating SCs (SC markers not shown).  (e & f) Astrocyte processes overlapping in close proximity with SKP-SCs (arrowheads) at the rostral (e) and caudal (f) host-graft interfaces. Note the lack of a well-defined border of reactive astrocytes. (g) Average GFAP (astrocyte) intensity traces (depicted as line drawn in a-c) representing the transition from the lesion to the host parenchyma for the three groups. (h) Bar graphs representing the mean area under the curve for the waveforms shown in g. Note the lower GFAP immunoreactivity in the time-of-transplant group versus the medium (*p<0.001; ANOVA with LSD) and SKP-SC (*p=0.019; ANOVA with LSD) groups, and in the SKP-SC group compared to the medium group (*p=0.030); ANOVA with LSD). (i) GFAP intensity traces in proximity to SKP-SCs, endogenous SCs, or no SCs (cavity) in SKP-SC transplanted spinal cords. (j) Bar graphs representing the mean normalized GFAP pixel intensity in each line used to generate the waveforms shown in i. Note the lower GFAP intensity in locations next to SKP-SCs compared to areas next to the cavity (no-SCs; *p=0.01; RM ANOVA with paired t-test) or next to endogenous SCs (+p=0.051; RM ANOVA with paired t-test). (k) Photomicrographs of laminin immunoreactivity at the rostral lesion-host spinal cord interface at 29 wpi. In the medium treated group, laminin immunoreactivity is largely associated with blood vessels whereas in the SKP-SC transplanted group laminin is widely expressed throughout the graft site containing SKP-SCs and in the surrounding parenchyma. (l) Lesion-host spinal cord interface at 29wpi in the SKP-SC and medium groups, immunolabeled for GFAP, Neurocan, the pan-CSPG marker CS-56, and GFP of the same region of interest; note the minimal glial scar formation with very little CSPG expression in the SKP-SC transplanted spinal cord. For g-j: SKP-SC n=15; Medium n=15; Time-of-transplant n=8. Data represented as mean +/- SEM in h & j. Scale bars 200 µm for a-c; 20 µm in d & e; 100 µm for g; and 50 µm in l.      147    148      Figure 3.6 SKP-SCs promote axonal growth/regeneration into the chronic lesion  (a-f) Representative photomicrographs of axons (NF-200/βIII-tubulin) at low (a, c, e; red) and high (b, d, f; boxed regions from a, c, e; white) magnification for the time-of-transplant (8wpi; a & b), medium (29 wpi; c & d), and SKP-SC (29 wpi; e & f) groups. Blue dotted blue lines in a, c, and e represent the narrowest point through the lesion where axon counts were conducted shown in g-i; yellow dashed lines indicate the regions used for axon counts in the rostral, middle and caudal region of the lesion (Figure 3.7 f-i). (g-i) Average NF-200-/βIII-tubulin+ (g), TH+ (h), and SERT+ (i) axon numbers were counted in animals with high graft survival (above the grey line in Fig 2 e) within a single confocal plane and compared to controls. (g) Note more than twice as many NF-200-/βIII-tubulin+ axons in the SKP-SC group compared to the medium (*p=0.008; KW with MWU) or time-of-transplant groups (*p=0.008; KW with MWU), and significantly more axons in the medium versus the time-of-transplant group (*p=0.032; KW with MWU). (h) More TH+ axons were counted in the SKP-SC group compared to the medium (*p=0.008; KW with MWU) and time-of-transplant groups (*p=0.008; KW with MWU). (i) There were also significantly more SERT+ axons running through the narrowest part of the cord in the SKP-SC group compared to the medium (*p=0.008; KW with MWU) and time-of-transplant groups (*p=0.008; KW with MWU), and more SERT+ axons in the medium group compared to the time-of-transplant group (*p=0.008; KW with MWU). (j-l) Photomicrographs of TH+ axon (red) growth through the SKP-SC grafted lesions at 29 wpi. Boxed regions in j are shown at higher magnification in k & l. (k & l) A large number of descending TH+ axons enter the graft (j & k) and a small number of those axons leave the caudal end of same graft and re-enter the host (arrows; j & l). (m) Representative photomicrograph demonstrating the large number of axons within the SKP-SC graft at 29 wpi. (n) Representative photomicrograph showing TH+ axons and SERT+ axons near the rostral host-graph interface. See also Figure 3.7. For g-i: SKP-SC n=5; Medium n=5; Time-of-transplant n=5; see supplemental methods. Scale bars:  200 µm for a, c, e; 100 µm for j; 50 µm for b, d, f, k, l; 20 µm for m, n. Individual data points for each animal are presented with group medians indicated by solid black lines in g-i.    149        150    Figure 3.7 Significantly more TH+ and SERT+ axons at the rostral and middle levels of the SKP-SC bridge compared to the caudal levels Representative photomicrographs depicting NF200/bIII-tubulin+ (a), TH+ (b), SERT+ (c), CGRP+ (d), and Subtance P+ (e) axons growing through GFP+ SKP-SC bridges. (f-i) Quantifications of NF200/bIII-tubulin+ (f), TH+ (g), SERT+ (h) and CGRP+ (i) axon counts at rostral, middle, and caudal levels (yellow lines in Figure 3.6 e) of the spinal cord in five animals from the SKP-SC subgroup. There were significantly more TH+ and SERT+ axons at the rostral and middle levels of the SKP-SC bridges than at the caudal level (all *p<0.05; Friedman test with follow up Wilcoxon) supporting the interpretation of rostral to caudal regeneration of these brainstem-derived axons. Subgroup analysis: SKP-SC n=5; Medium n=5; Time-of-transplant n=5. Scale bars: 50 µm for e; 20 µm for b 10 µm for a, c, and d. Individual data points for each animal are presented with group medians indicated by solid black lines in f-i.           151 Figure 3.8 SKP-SCs myelinate axons  (a & b) Close-up micrograph of a SKP-SC bridge demonstrating the formation of SC myelin (P0) around host axons by GFP+ SKP-SCs. (c) High magnification confocal optical section showing a GFP+ SKP-SC ensheathing a NF200-/bIII-tubulin+ axon with P0+ myelin. Note the cell body of the SKP-SC (arrow) and the layers of P0 (arrowhead) sandwiched between the axon and the GFP+ cytoplasm in the outer layer of the SKP-SC. (d & e) Photomicrographs of axons ensheathed by GFP+ SKP-SCs (with P0+ myelin in d) and immunostained for Kv1.2+ potassium channels. (f & g) Images of GFP+ SKP-SCs forming tubes (f) or ensheathing axons (NF200/bIII-tubulin; blue in g) with P0+ myelin (red in g) and immunostained for Caspr (red in f; white in g). Note that SKP-SC-ensheathed axons showed appropriate nodal structures, including nodes of Ranvier (arrows; d-g) with ribbon-like labeling of the paranode by Caspr (arrowheads; f & g) and the juxtaparanode by Kv1.2 (arrowheads; d & e) on either side of the node. Scale bars: 20 µm for a, b, g; 10 µm for c &f; 5 µm for d & e.         152  Figure 3.9 The SKP-SC group contains significantly more SC myelin and non-myelinating SCs than the control groups (a-c) Overview micrographs of the spinal cords from the time-of-transplant (8 wpi; a), medium-injected (29 wpi; b, and the GFP+ SKP-SC transplanted (29 wpi; c) group immunostained for P0 and GFAP.  (c-d) Note the extensive presence of P0+/GFPNEG myelin produced by endogenous SCs in the spinal cord of animals with both high (c; greater than 70,000 SKP-SCs at 29wpi) or low (d; less than 50,000 SKP-SCs at 29wpi) survival of transplanted SKP-SCs. (e) The presence of abundant endogenous SCs is further supported by their expression of p75-NTR (expressed highly in non-myelinating SCs) in the absence of GFP, often observed in close proximity to GFP+ SKP-SC grafts. (f) Note the presence of P0+ myelin generated by endogenous (GFPNEG) SCs in a medium-injected spinal cord in close proximity to GFAP+ astrocytes. (g) High magnification photomicrograph of axons (NF200/bIII-tubulin) ensheathed by either GFP+ SKP-SCs (arrow) or GFPNEG endogenous SCs (arrowhead). (h) Non-myelinating p75-NTR+ SCs derived from transplanted GFP+ SKP-SCs (arrow) within the lesioned spinal cord at 29 wpi. (i) Quantification of P0+ volumes by treatment groups; the green segment denotes the mean volume of P0+/GFP+ SKP-SCs. Note that the SKP-SC group had significantly greater SC myelin volume (black and green bar combined) than both the medium (*p=0.005; ANOVA with LSD) and the time-of-transplant control (*p<0.001; ANOVA with LSD) groups. Importantly the mean volume of P0+/GFPNEG endogenous SCs was higher in the SKP-SC group (black portion of bar) than in the medium group (gray bar; *p=0.038; ANOVA with LSD), indicating that SKP-SC transplantation enhances the presence of endogenous SCs in the injured spinal cord. There was no difference observed between the time-of-transplant and medium control groups (+p=0.068; ANOVA with LSD). (j) Distribution of P0+ myelin inside the lesion versus outside the lesion, i.e. in the spared host rim. The SKP-SC group had significantly greater P0+ volume than the time-of-transplant control group both inside (*p=0.001; ANOVA with LSD) and outside (*p=0.006; ANOVA with LSD) of the lesion, but only showed significantly more P0 than the medium group inside the lesion (*p=0.005; ANOVA with LSD). The time-of-transplant control group showed significantly greater P0+ volumes outside the lesion as compared to inside (*p=0.002; paired t-test) and the medium group showed no difference with the same effect (+p=0.067; paired t-test). (k) SKP-SC group approached statistical significance toward having larger p75-NTR+ volumes compared to the medium group (*p=0.074; MWU). (l) Both the SKP-SC and medium groups showed significantly greater p75-NTR+ volume inside the lesion compared to outside (*p=0.002 and *p=0.014; paired t-test, respectively), and the SKP-SC group approached statistical significance toward a higher p75-NTR+ volume compared to the medium group inside the lesion only (+p=0.057; t-test). For i-l: SKP-SC n=15; Medium n=15; Time-of-transplant n=8. Scale bars: 200 µm in a-f; 10 µm in g & h. Data presented as means +/- SEM in i, j, l; Individual data points for each animal presented with group medians indicated by solid black lines in k.     153         154       Figure 3.10 Rat SKP-SCs harvested from adult skin express typical SC markers in vitro Photomicrograph of cultured GFP-labelled rat adult SKP-SCs immunostained for the SC markers p75 and P0 (a) or S100b and GFAP (b) as well as Hoechst (nuclear marker). Scale bars represent 20µm.           155  Figure 3.11 SKP-SCs isolated from adult skin survived well and demonstrated similar properties as neonatal isolated SKP-SCs (a) Photomicrograph of a GFAP-immunostained contused spinal cord from an animal transplanted with adult GFP+ SKP-SCs. At 29 wpi adult SKP-SCs integrate into the lesion sites and bridge the cavity in precisely the same manner observed with transplanted neonatal SKP-SCs (see Fig 3.2 c).  (b) SKP-SC counts at 29 wpi for animals that received an adult SKP-SC transplantation; note that these values were higher than many of those found after neonatal SKP-SC transplantation (see Figure 3.2 e). (c) Overlapping GFAP+ astrocyte processes in close proximity to GFP+ adult SKP-SCs at the caudal graft interface.  Note the lack of a well-defined border of reactive astrocytes, as well as the intermingling of transplanted GFP+ adult SKP-SCs and astrocytic processes. (d) High-magnification photomicrograph of GFP+ SKP-SCs with axons (NF-200/βIII-tubulin) demonstrating the rostral-caudal orientation and extensive number of axons found in adult SKP-SC grafts. (e) Representative photomicrograph showing TH+ axons growing through the middle of the GFP+ adult SKP-SC bridge. (f) Photomicrograph of an injured spinal cord from a rat that received an adult SKP-SC transplantation immunolabeled for GFP, GFAP, and P0; Note the clear co-labelling in yellow between the green GFP+ adult SKP-SCs and red P0 as well as the large endogenous SC response (GFPNEG/P0+; red only) observed. (g) High magnification confocal optical slice showing an adult GFP+ SKP-SC ensheathing a NF200/bIII-tubulin+ axon with P0+ myelin.  Scale bars represent 200 µm for a & f; 40µm for c; 20 µm for d & e; 10 µm for g.     156             157  Figure 3.12 Chronically contused animals transplanted with SKP-SCs demonstrate greater locomotor outcomes and decreased pathological thickening of the bladder wall (a) SKP-SC transplanted animals had higher mean BBB scores than the medium-injected group during much of the open field locomotion testing conducted in this study. A repeated-measures ANOVA examining weeks 12-27 (the last 4 months of the study) showed a significant interaction (*p=0.032); indicating that the difference between those two groups changed over time during that period. Follow-up testing revealed that SKP-SCs had significantly higher BBB scores than the medium group at weeks 19 (*p=0.012; t-test) and 21 (*p=0.026; t-test). (b & c) CatWalk footprint analysis assessing forelimb and hindlimb stride length normalized to pre-transplant values. A repeated-measures ANOVA over the last 16 weeks yielded a significant interaction in normalized forelimb (*p=0.046; b) and hindlimb stride length (*p=0.032; c). (B) Normalized forelimb stride length was significantly greater for SKP-SC treated animals (compared to medium control) at 16, 18, 20, and 24 wpi (p<0.013; t-test) and showed a trend toward significance at 22 wpi (+p=0.069); t-test. (c) Normalized hindlimb stride length showed significant differences at every time point between 16 and 26 wpi (*p<0.039). (d & e) Representative (near-median values) photomicrographs of bladder wall stained with cresyl violet from the medium control group (d) and the SKP-SC transplanted group (e). (f) Quantification of average bladder wall thickness at the thickest point yielded that the SKP-SC group had significantly thinner bladder walls than the medium treated group (*p=0.001; MWU). (g & h) Scatterplots of bladder wall thickness at the thickest point versus average BBB score (g) or average BBB subscore (h) from week 17-27 for both medium (gray squares) and SKP-SC (black dots) groups. Bladder wall thickness showed a significant negative correlation to both BBB and BBB subscore (Spearman’s rho = –0.414 and –0.425, p = 0.036 and 0.030, respectively), indicating that animals with better locomotor function also had thinner bladder walls. See also Figure 3.13. For a-c: SKP-SC n=15; Medium n=15. Data presented as group means +/- SEM. For f-h: SKP-SC n=12; Medium n=14. Scale bars represent 100µm for d and e. Individual data points for each animal are presented with group medians indicated by solid black lines in f and the line of best fit indicated by the solid black lines in g & h.    158    159         Figure 3.13 SKP-SCs elicited functional improvements on paw angle and step sequence parameters on the CatWalk (a) SKP-SC transplanted animals showed a significantly lower paw angle (less outward rotation) over the last 10 weeks of behaviour, as indicated by a significant interaction (*p=0.041; repeated measures ANOVA). Follow-up analysis indicated that the SKP-SC group had significantly lower paw angles at 20 and 22 wpi (*p<0.047; t-test) and a trend at 24 wpi (+p=0.072; t-test) but this difference was not maintained at 26 wpi. (b) The SKP-SC transplanted group showed a significantly smaller percentage of the abnormal stepping pattern LF-RF-RH-LH as compared to the medium control group at week 18 and 26 (*p<0.003; MWU) and approached significance at week 24 (+p=0.076). There were no significant interactions found on the repeated measures ANOVA in mean hindlimb print width (c) or mean hindlimb intensity (d) CatWalk parameters or the mean BBB subscore (e) or the irregular horizontal ladder (f). SKP-SC n=15; Medium n=15. Data is presented as means +/- SEM.      160        161 Chapter 4: General discussion  4.1 Summary of thesis In this dissertation, I addressed two major gaps in our knowledge of myelin repair after SCI. In Chapter 2, I used genetic lineage tracing to evaluate the sources and extent of spontaneous myelin repair by endogenous cells. We found that OPCs (PDGFRa-expressing cells) contribute to approximately 30% of the oligodendrocyte myelin sheaths at epicenter 12 wpi, suggesting that there are effective myelin repair mechanisms in place even in the absence of treatment. Furthermore, we demonstrated that the majority of the SCs found at injury epicenter are derived from oligodendrocye lineage cells in the spinal cord and not from the periphery as previously thought. In Chapter 3, I assessed the ability of skin-derived cells to facilitate myelin repair at clinically-relevant time points after contusion SCI in rodents. We found that transplanting SKP-SCs can facilitate myelin repair and improve functional outcomes of SCI (Chapter 3). In this chapter, I will address the original hypotheses underlying the work (introduced in Chapter 1), discuss the significance of the enclosed findings, and discuss future directions of this work such that it might truly be able to positively affect the lives of people with SCIs.  4.2 Endogenous myelin repair after SCI The dogma that demyelination underpins the failure of conduction across the site of SCI was derived implicitly, based on the findings that some demyelinated axons persist in the vicinity of chronic SCI (Blight, 1983b, 1985) and that axons that traverse the chronic SCI injury site often fail to conduct impulses (Blight, 1983a). Addressing the hypothesis that PDGFRa+ cells   162 contribute to the majority of new oligodendrocyte and SC ensheathment/myelination observed in response to contusion SCI, I made use of several different transgenic mouse lines that allowed me to track OPCs (PDGFRa+), oligodendrocyte lineage cells (olig2+), and myelinating SCs (P0+) to see how each of these cell populations contribute to myelin repair after SCI. Fate mapping studies characterizing the behaviour of endogenous cells after SCI are crucial to our understanding of spontaneous repair. In the wake of SCI, transgenic labeling facilitates direct and reliable quantification of new oligodendrocytes and new myelin tubes (i.e., arising post-injury), information that was previously elusive given that g-ratios do not appear to be an accurate indicator of myelin formed de novo after SCI (Lasiene et al., 2009; Powers et al., 2012; Powers et al., 2013). Prior to fate mapping, SCs infiltrating from the PNS were thought to contribute substantially to remyelination after SCI. This seemed reasonable, given that after SCI, there is extensive damage to the glia limitans, and that SC myelination occurs more prominently in areas located close to peripheral nerves (Snyder et al., 1975; Duncan and Hoffman, 1997). Despite evidence that endogenous OPCs produce SCs after chemical demyelination (Zawadzka et al., 2010; Crawford et al., 2016a) and that OPCs transplanted into demyelination lesions can generate SCs (Talbott et al., 2005; Talbott et al., 2006), many SCI researchers remain skeptical of their central origin. We found in the mouse, after taking into account recombination efficiency, that 70-80 % of the myelinating SCs at the site of SCI are derived from central OPCs and less than 10% are derived from peripheral myelinating SCs. The ability of OPCs, with a neuroepithelial origin, to become SCs, of neural crest origin, fuels some controversy in the field (Bielecki et al., 2016) and helps consolidate OPCs on the growing list of stem cell-like populations throughout the body that demonstrate plasticity beyond that originally described.   163 We also found that endogenous OPCs contribute substantially to new ensheathment/myelin after SCI, which agrees with recent data demonstrating that new oligodendrocytes arise for months post SCI and contribute to new myelin (Hesp et al., 2015). These (and related) data presented in Chapter 2 suggest that endogenous oligodendrocyte myelin repair after SCI may be more efficient than we once suspected. It is well established that demyelination takes place early after SCI; however, it is becoming more widely accepted that spontaneous remyelination occurs in both animal models and human SCI (Bunge et al., 1961; Gledhill et al., 1973; Kakulas, 1999; Totoiu et al., 2004; Smith and Jeffery, 2006; Lasiene et al., 2008; Powers et al., 2012) although the degree of remyelination is still debated. Even though I have determined that PDGFRa-expressing OPCs generate many new myelinating oligodendrocytes and associated ensheathment/myelin after SCI, I cannot conclude that OPCs are the only source of new myelin. For example, we know that ependymal cells can show multi-lineage differentiation and contribute to 4% of the new oligodendrocytes after SCI (Meletis et al., 2008; Gregoire et al., 2015). A potential corollary of our findings on spontaneous myelin repair after SCI, along with data from other groups demonstrating extensive remyelination post SCI in both rodents (Lasiene et al., 2008; Powers et al., 2012) and canines (Smith and Jeffery, 2006), is that the myelination status of spared or re-growing axons may not be the primary reason for conduction failure at the site of SCI (Arvanian et al., 2009). However, it is important to acknowledge that we did not determine whether ensheathment/myelination observed after SCI augmented axonal conduction across the injury site or facilitated the associated functional recovery. Ongoing work in our laboratory is investigating the importance of new oligodendrocyte myelin after SCI, by   164 genetically blocking new oligodendrocyte myelin formation in PDGFRa+ cells to determine whether this influences spontaneous functional recovery (Duncan et al., 2017; in preparation). While the work in this dissertation reveals the dramatic contribution of OPCs in both SC production and myelin repair, it raises a number of important questions for future investigation.  One priority is directly assessing the contribution of non-myelinating SCs to myelinating SCs found in the spinal cord after SCI. To date, determining this has been challenging: one of the most robust SC markers, NTR-P75, is not unique to non-myelinating SCs. Evaluating the importance of OPC-derived myelinating SCs in functional repair and recovery after SCI may be more immediately feasible: conditional knockout of Krox20 (required for SC myelination) (Topilko et al., 1994) in OPCs would allow us to assess the importance of this SC population in myelination and the associated spontaneous functional recovery after SCI. Selectively manipulating Krox20 expression would also provide information on the importance of SC myelin versus the mere presence of SCs in the spinal cord after injury; as discussed in Chapter 1, SCs can contribute to repair in many ways, only one of which is myelinating axons. In addition, recent work in demyelination lesions suggests that OPC that give rise to SCs are derived from predominately the dorsal OPC population (Crawford et al., 2016a). Since transgenic mouse lines are now available that permit independent labeling of dorsally- or ventrally-derived OPCs (Tripathi et al., 2011; Zhu et al., 2011; Crawford et al., 2016), these could be leveraged to (first) understand the response of the ventral and dorsal OPCs after SCI, and (second) determine the relative importance of dorsal and ventral OPCs in spontaneous repair after SCI. Finally, I want to acknowledge that endogenous SCs in the spinal cord could be detrimental to recovery after SCI. For example, we know that Schwannosis (Bruce et al., 2000; Norenberg et al., 2004), the aberrant accumulation of SCs and axons, is observed in > 80% of human SCI cases when   165 examined years after injury (Bruce et al., 2000). Using similar transgenic mouse models to those used in Chapter 2 would allow us to genetically label centrally- and/or peripherally-derived SCs prior to SCI and to follow these cells months and even years after injury, which could reveal important information about the origin of Schwannosis-forming cells.  4.3 Exogenous myelin repair after SCI Addressing the hypothesis that SKP-SCs transplanted eight weeks after contusion SCI would survive, bridge the lesion site making it permissive for axon growth, and contribute to remyelination as well as functional recovery, I transplanted SKP-SCs at the site of chronic SCI, and assessed histological repair, behavioural recovery, and changes in bladder thickness at x weeks after SCI. I found that SKP-SCs bridged the site of SCI, and that growing/regenerating axons grew along the grafts. In addition, SKP-SCs myelinated axons and increased the presence of endogenous SCs. Rats that received SKP-SCs showed improved gate perimeters and open field locomotor scores, and exhibited less bladder wall thickening, a pathological correlate of detrusor-sphincter dyssynergia.  Although we saw modest functional improvements in animals that received SKP-SCs many questions remain regarding the mechanisms underlying these improvements. For example, it is unclear whether the axons being myelinated by either the exogenous or endogenous SCs are making meaningful connections, and therefore contributing to improved conduction across the injury site. In addition, the increased number of axons traversing the lesion site in the SKP-SC group may result from 1) an increase in axons surviving secondary damage after SCI; 2) an increase in newly grown/regenerated axons; and/or 3) an increase in newly grown peripheral axons.  One way to tease out the importance of axonal myelination by transplanted cells would   166 be to transplant SKP-SCs that are incapable of producing myelin, such as SKP-SCs derived from trembler (Pmp22Tr) mutant mice (Henry and Sidman, 1988) or isolated from a Krox20 knockout mouse (Topilko et al., 1994). As we work to uncover the mechanisms underlying recovery, we may discover that we can achieve similar effects via less invasive treatments, to achieve the benefits of transplantation without the inherent risks (see 4.4.1, below).  In addition, more work is required to determine whether SKP-SCs have advantages over nerve-derived cells in terms of promoting repair after SCI. Some work from our laboratory suggests that SKP-SCs have advantages over nerve-SCs, including reduced astroglyosis and improved integration into the host parenchyma (Biernaskie et al., 2007). However, more recent work directly comparing neonatal SKP-SCs and nerve-derived SCs found the functional outcomes (electrophysiological and behavioral) were indistinguishable (Sparling et al., 2015). Recent work from Freda Millers group, directly comparing adult rat SKP-SCs with adult rat nerve-derived SCs also found that the two populations are very similar (Krause et al., 2014). Given the considerable body of literature on transplantation of nerve-derived SCs, more data on SKP-SC transplantation is required. If the two cells types do indeed promote similar levels of repair, SKP-SCs are in theory more feasible for clinical use.  Importantly, the findings in Chapter 3 of this dissertation suggest that it is the process of transplanting SKP-SCs after SCI, rather than direct effects of the transplanted cells, that results in higher locomotor scores compared to control rats. Although we lack the direct evidence, it is tempting to speculate that endogenous SCs, accumulating as a result of SKP-SC transplantation, mediate the beneficial effects. The finding that the number of endogenous SCs at the site of SCI increases with exogenous cell transplantation is in line with data from experiments transplanting nerve-derived SCs (Hill et al., 2006; Biernaskie et al., 2007), OECs (Ramer et al., 2004; Au et   167 al., 2007; Zhang et al., 2011), BMSCs (Lu et al., 2005, 2007) and fibroblasts (Sparling et al., 2015) at the site of SCI. My findings from Chapter 2 suggest that most SCs found in the spinal cord after injury are derived from PDGFRa+ OPCs.  Together, all of these data suggest that exogenous cell transplantation can alter the injury environment and change the balance of OPC-derived oligodendrocytes versus OPC-derived SCs. In the big picture, these findings also highlight two important points: first, more work is required to understand the relationship between transplanted cells and their effects on endogenous precursors; second, it may be possible to manipulate endogenous cells to promote repair after SCI, bypassing the need for invasive cell transplantation. For example, growth factor and ferritin treatments increase OPC proliferation and stimulate oligodendrogenesis (Schonberg et al., 2012; Whittaker et al., 2012).   4.4 Future and clinical perspectives The work presented in this dissertation examines endogenous glial repair after SCI and exogenous cell transplantation as a treatment for SCI. The results highlight the importance of future research focusing on understanding and augmenting spontaneous repair mechanisms. In this final section, I provide my perspective on crucial considerations for moving this work (and other treatments for SCI) toward clinical translation.  4.4.1 Improved risk assessment When considering experimental interventions, people who have sustained a SCI are contemplating treatment for a non-life threatening condition. The risk of treatment is therefore a   168 paramount consideration, and this should be reflected in pre-clinical work. Detrimental outcomes in pre-clinical experiments are rarely documented or reported; however, this is valuable information that could be used in clinical trials, to help people with SCI to effectively weigh the risks and benefits of an experimental treatment. Ideally, people with SCI would require the risk of treatment-based detrimental outcomes to be <1%; however, risk assessment decisions vary from person to person and 15-30% of people with SCI would participate in an experimental treatment regardless of the risk (Kwon et al., 2012). Risk assessment likely varies with time post-SCI; people with SCI may be more risk tolerant in the acute stages, and less so as they adjust to living with SCI. As a result, the acceptable risk of detrimental outcomes should vary depending on the timing of treatment post-SCI.  Investigating the potential for detrimental outcomes is particularly important for transplantation-based therapies, as each transplantation candidate cell carries specific risks during translation that are often poorly understood (Hofstetter et al., 2005). In preclinical work, researchers should perform transplantation into large animals (Kwon et al., 2015) that live longer than rodents, allowing long-term assessment of the tumourigenic potential of human cells that may not thrive in a rodent environment (Salegio et al., 2016). Tumour formation is a major risk that needs to be monitored and reported more regularly, particularly in cases where cells are derived from a pluripotent stage (Nori et al., 2015). Neuropathic pain is another risk that needs to be monitored more closely, particularly in cases of undifferentiated neural stem cells (Hofstetter et al., 2005). Another risk of transplantation is increasing the likelihood of infections. With injuries at T3 or above, there is a substantial systemic depression of immune function, termed CNS injury-induced immune deficiency syndrome (CIDS) (Meisel et al., 2005; Zhang et al., 2013b). The use of immunosuppressants might further weaken immune function; these should be   169 tested and monitored to assess the increased risk of infection after high SCI (Brommer et al., 2016). There might be an upside of CIDS, in that these patients may be less likely to reject allogeneic transplants.   4.4.2 Improved pre-clinical models  Most preclinical data are based on mild to moderately severe thoracic contusion SCI, with substantial spontaneous functional recovery. Testing treatments in these injuries does not provide sufficient evidence for clinical translation of a cell transplantation (Kwon et al., 2013). Functionally incomplete thoracic injuries occur infrequently in humans (approximately 5% of cases in Canada) (Dvorak et al., 2014). In contrast, more than 60% of human SCIs occur in cervical spinal segments, and many lead to complete or pronounced loss of sensorimotor function (Dvorak et al., 2014). Patients with cervical injuries are expected to benefit the most from cell therapies, as regaining two segments of function could mean regaining hand function, conferring independence and greatly improved quality of life (Steeves et al., 2012). In order to align our pre-clinical models with what is seen in the clinic, making use of new rodent cervical injury models, testing treatments in combination with training, and using relevant outcome measures is crucial (Girgis et al., 2007; Lee et al., 2012a; Lam et al., 2014; Chen et al., 2016). In addition, many studies have been undertaken in immunodeficient animals that have vastly different immune responses to SCI (Kobayashi et al., 2012). Such alterations of the immune system can overcome rejection of transplanted cells and may better reveal the tumorigenic potential of a human cell in a rodent. However, these immunodeficient models may not accurately model the human situation.   170  4.4.3 Assessing efficacy in the chronic stage of injury  Most preclinical transplantation studies are performed ~1–2 weeks after SCI. While this practice has provided a model for several clinical trials, a later transplantation time point is highly desirable. Translation in the chronic stages of SCI is more logistically feasible and less ethically contentious. In addition, chronic transplantation will dramatically reduce the number of subjects required for a clinical trial, since the trajectory of spontaneous recovery is more predictable (Steeves et al., 2011). However, the SCI microenvironment changes drastically from the acute to the chronic setting, and this can affect cell survival and differentiation (Nishimura et al., 2013). Unfortunately, few previous preclinical studies have been performed with treatment initiation beyond 2 mpi, and those that have been conducted demonstrate modest or no functional benefits (Barakat et al., 2005; Keirstead et al., 2005; Karimi-Abdolrezaee et al., 2006). The results presented in Chapter 3 add to this body of work; however, only modest functional improvements were observed after SKP-SC transplantation. Despite this, trials using nerve-derived SC transplantation in humans with chronic injuries have been initiated ( Whatever the functional outcome of patients in the treatment group, at least this trial sets a precedent for cell transplantation in the chronic setting.  4.4.4 Defining recovery ‘Recovery’ is defined in the Oxford English Dictionary as “a return to a normal state of health, mind, or strength.” In most animal models of SCI there are significant spontaneous improvements. For example, after the commonly used moderate thoracic contusion, there is   171 initially flaccid paralysis of the hind limbs, but within weeks animals spontaneously regain weight-supported stepping. However, recovery to a normal state, comparable to an uninjured animal, does not occur. As a field, SCI researchers use the term ‘functional recovery’ to refer to a statistically significant functional improvement in a treatment group versus control, although true recovery as such is rarely observed. Hence, we should avoid using this general term and refer to specific improvements. Importantly, since no effective treatment for SCI has been translated to clinical use, it is unclear what degree of functional improvement in preclinical animal models is clinically relevant. Although we focus on motor improvements, treatment can also promote sensory or autonomic functional improvement, which can be more highly prioritized by those with injury (Anderson, 2004).  4.4.5 Aligning outcome measures with the priorities of the SCI community Historically, the goal of SCI physicians and researchers has been to restore walking. However, a survey of the SCI community clearly demonstrated different priorities. The survey identified arm and hand function as most important to people with tetraplegia and sexual function as most important to people with paraplegia; in addition, bladder and bowel function were identified as important by both groups (Anderson, 2004). To some extent, the research community has taken these findings into consideration, and incorporated outcomes on sensation, autonomic function, sexual function and pain in preclinical research. While this was an important step for the field, there is much more we can do: I still feel the need to emphasize the requirement for incorporating questions and hypotheses specific to improving the quality of life for people with SCI into every pre-clinical and clinical study. One way to help researchers to   172 keep this focus is to facilitate ongoing dialogue between the SCI community and SCI researchers; this might be achieved by including more consumers in decision-making processes, perhaps on Editorial Boards of journals and (as already occurs in some instances) in adjudicating funding decisions.   4.4.6 Managing expectations For many people who have sustained a SCI, managing your own expectations for recovery, and the expectations of people around you, can be difficult. Often, predictions are made about the potential of neurological outcome/recovery using the information at hand early after injury. Ongoing research focused on improved predictive tools is important, both to give patients more information early after injury, and to improve clinical trials for SCI (Tigchelaar et al., 2017). At present, predictions vary depending on many factors, but generally always include the message that life will be different now, and recovery will likely not meet anyone’s expectations.  This message is tolerated to different degrees by different individuals. Many people with SCI do not feel as though they have access to options to aid in their recovery. Standard rehabilitation is designed to promote independent living and a return to the community, rather than achieve substantial functional recovery. People with SCI hear about potential medical advancements all the time, as discoveries coming out of laboratories all over the word; when these findings are translated through the media, there is a tendency to focus on the hype. In the face of this complex new reality, people seek out something to augment their recovery, and may search for alternative treatments without the information about relevant risks and/or the expected level of risk aversion. Many people affected by SCI find treatments that appear promising in   173 other countries and feel as though such treatments are being withheld from them in Canada. Some people decide to travel to clinics in other countries to receive treatments that are not available at home, typically because the treatments are unproven and potentially unsafe. In the case of stem/progenitor cell transplantation, this practice has been dubbed Stem Cell Tourism.  Researchers have an obligation both to listen to the priorities and expectations (however unfeasible) of the SCI community, and to educate the SCI community about the potential and the limitations surrounding their research. Researchers and clinicians need to realize that there is a subset of people with SCI who are willing to participate in high-risk clinical trials; an interesting ethical question is whether we have an obligation to provide this group of people with an experimental treatment option within Canada. Teaching the SCI community is demanding, but an important challenge for researchers to take on, such that researchers with both knowledge and perspective are presenting the details and combatting the hype that accompanies media coverage of treatments for SCI. From a pre-clinical research standpoint, researchers need to learn how to break down their work such that it can be understood by a wide audience; they need to explain the significance of the work but also highlight the ongoing concerns, and then be very realistic about expectations for clinical translation. This involves teaching the community about why answering basic science questions is important, why we need to test potential treatments in pre-clinical models, why regulatory systems are important, why phase 1 and 2 clinical trials are important, and the probability of clinical translation succeeding. Although this seems like it should be left to someone else, we (as researchers) are in the best position to inform our consumers and help them make choices about their health. For most of us, our scientific discoveries will not directly improve lives. 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