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Characterization of oligodendrocyte lineage cell responses remote to the lesion site in a murine model… Moulson, Aaron Jack Taylor 2017

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CHARACTERIZATION OF OLIGODENDROCYTE LINEAGE CELL RESPONSES REMOTE TO THE LESION SITE IN A MURINE MODEL OF THORACIC CONTUSION SPINAL CORD INJURY (SCI) by  Aaron Jack Taylor Moulson  B.Sc. (Honours), The University of British Columbia, 2014  A THESIS SUBMITTED IN PARTIAL FULFILLMENT OF THE REQUIREMENTS FOR THE DEGREE OF  MASTER OF SCIENCE in THE FACULTY OF GRADUATE AND POSTDOCTORAL STUDIES (Zoology)  THE UNIVERSITY OF BRITISH COLUMBIA (Vancouver)  January 2017  © Aaron Jack Taylor Moulson, 2017 ii  Abstract Traumatic injury to the adult mammalian central nervous system (CNS) commonly results in permanent functional impairment due to the inability of injured CNS neurons to mount an effective regenerative response.  Injury to the spinal cord is associated with a range of sensory, motor, and autonomic deficits, the most severe of which is complete paralysis.  At a histological level, injury-induced pathophysiological processes have been thoroughly characterized for the tissue area immediately surrounding the lesion epicentre, however remote to the lesion these changes are less well described.  Previous studies have demonstrated that various perturbations, including traumatic injury, demyelination, artificial neural stimulation, neurodegeneration, and social experience, among others, induce robust oligodendrocyte precursor cells (OPC) responses, which function as endogenous precursors for myelinating oligodendrocytes.  De novo myelination in the adult CNS has been implicated in motor learning, memory, and optimization of neural network function, thought to represent a potent form of neural plasticity.  The demonstration of robust OPC proliferation and oligodendrogenesis in models of dorsal rhizotomy, axotomy, and axon degeneration, combined with the robust OPC responses characteristic of SCI lesion epicentres, lead us to hypothesize that contusion SCI would induce significant responses in resident OPC populations remote to the lesion site (specifically comprised of OPC proliferation, oligodendrogenesis, and new myelination).  This may be functionally relevant to myelin plasticity on spared motor and sensory tracts remote to the lesion.  To test this hypothesis, we conducted an in vivo study employing transgenic mouse lines (i.e. PDGFRα-CreERT:ROSA26-YFP and PDGFRα-CreERT:ROSA26-mGFP) that enabled the visualization and fate-mapping of OPCs and their progeny in the adult murine spinal cord following a moderate (70 Kdyne) T9-T10 thoracic contusion injury.  Consistent with our predictions, we observed robust OPC proliferation and iii  oligodendrogenesis remote to the lesion in both the cervical and lumbar spinal cord.  Surprisingly, this did not manifest as increased new myelination, attributed (at least in part) to an observed maturation impairment of newly-formed oligodendrocytes.    iv  Preface The original idea for the fate-mapping of oligodendrocyte precursor cells (OPCs) following murine contusion spinal cord injury (SCI) was developed by Peggy Assinck (PhD student) and Dr. Wolfram Tetzlaff, who supervised the in vivo and histological components of this study.  PDGFRα-CreERT2 transgenic mice were obtained from Dr. Dwight Bergles (John Hopkins University; Baltimore, MD, U.S.A.) and were cross-bred with commercially available ROSA26-YFP and ROSA26-mGFP reporter mouse lines to generate the PDGFRα-CreERT2:ROSA26-YFP and PDGFRα-CreERT2:ROSA26-mGFP transgenic mice used in the current thesis.  Peggy Assinck, with assistance from Gregory Duncan (PhD student), conducted the animal portion of the experiment with a larger cohort than used in the current study.  Lesion epicentre tissue was used for a study performed by Peggy Assinck regarding the myelinogenic plasticity of OPCs following contusion SCI (currently in preparation for publication).  All surgical procedures were completed by Dr. Jie Liu at the Blusson Spinal Cord Centre (B.S.C.C.) on the Vancouver General Hospital (V.G.H.) campus (Vancouver, B.C., Canada).  Behavioural assessment (data not presented here), post-operative animal care, animal sacrifice, and tissue collection were performed by Peggy Assinck and Gregory Duncan.  Cervical, thoracic, and lumbar spinal cord sections were stored in the -80 °C freezer.  I designed all of the histological assessments presented in the current thesis, as well as performed all of the cryostat tissue sectioning, immunohistochemical staining of tissue sections, imaging of stained tissue, image analysis (i.e. cell and myelin sheath counting, immunoreactivity thresholding analyses, etc.), statistical analysis of the data, and created all of the figures and graphs presented in the current thesis.  Critical feedback on experimental progress, proposed assessments, and preliminary data was provided throughout by Dr. Tetzlaff, Dr. Oscar Siera, Gregory Duncan, Peggy Assinck, Doug Brown, Sohrab Manesh, and Brett Hilton.  All of v  the work presented in the current thesis was approved by the University of British Columbia (U.B.C.) Animal Care Committee (A.C.C.) in accordance with the guidelines established by the Canadian Council on Animal Care (C.C.A.C.).  The relevant animal care certificate number for this work is A03-0139 – ‘Anatomical and functional recovery after spinal cord contusion injury.’ vi  Table of Contents  Abstract .......................................................................................................................................... ii Preface ........................................................................................................................................... iv Table of Contents ......................................................................................................................... vi List of Tables .............................................................................................................................. xiii List of Figures ............................................................................................................................. xiv List of Abbreviations ................................................................................................................. xvi Glossary .................................................................................................................................... xxiv Acknowledgements ................................................................................................................. xxvii Dedication ................................................................................................................................. xxix Chapter 1: Introduction ................................................................................................................1 1.1 Spinal cord injury (SCI): clinical presentation ............................................................... 1 1.2 Pre-clinical models of spinal cord injury (SCI) .............................................................. 3 1.2.1 Murine model of contusion spinal cord injury (SCI) using the Infinite Horizons (IH) impactor .................................................................................................................................. 4 1.2.2 Cre-lox biochemistry .................................................................................................. 5 1.2.3 Inducible-conditional estrogen-receptor/Cre-recombinase transgenic mice .............. 6 1.3 Spinal cord injury (SCI): pathophysiology ..................................................................... 9 1.3.1 Primary injury ............................................................................................................. 9 1.3.2 Secondary injury ....................................................................................................... 10 1.3.2.1 Hemorrhage....................................................................................................... 10 1.3.2.2 Ischemia ............................................................................................................ 11 vii  1.3.2.3 Excitotoxicity .................................................................................................... 12 1.3.2.4 Oxidative stress ................................................................................................. 13 1.3.2.5 Blood-borne inflammatory cells ....................................................................... 14 1.3.2.6 Microgliosis ...................................................................................................... 14 1.3.2.7 Progressive cell death ....................................................................................... 15 1.3.3 Pathophysiological effects remote to the lesion site ................................................. 16 1.4 Endogenous cell responses to spinal cord injury (SCI) ................................................ 17 1.4.1 Neural stem/progenitor cells (NSPCs) ...................................................................... 18 1.4.2 Astrocytes ................................................................................................................. 18 1.4.3 Oligodendrocytes ...................................................................................................... 20 1.4.3.1 Oligodendrocyte death ...................................................................................... 20 1.4.3.2 Oligodendrogenesis........................................................................................... 22 1.4.3.3 Remyelination ................................................................................................... 23 1.4.4 Neurons ..................................................................................................................... 24 1.4.4.1 Neuronal death .................................................................................................. 24 1.4.4.2 Axonal transection and regenerative failure ..................................................... 24 1.4.4.3 Axonal plasticity ............................................................................................... 25 1.4.5 Oligodendrocyte precursor cells (OPCs) .................................................................. 26 1.4.5.1 Proliferative response........................................................................................ 26 1.4.5.2 Lineage plasticity .............................................................................................. 27 1.4.6 Microglia ................................................................................................................... 28 1.5 Oligodendrocyte lineage cells ....................................................................................... 29 1.5.1 Development of the oligodendrocyte lineage ........................................................... 29 viii  1.5.2 Extrinsic regulation of oligodendrocyte lineage development ................................. 33 1.5.2.1 PDGF-signaling ................................................................................................ 33 1.5.2.2 FGF-signaling ................................................................................................... 34 1.5.2.3 EGF-signaling ................................................................................................... 34 1.5.2.4 Model for regulation of OPC proliferation ....................................................... 35 1.5.2.5 OPC migration .................................................................................................. 36 1.5.3 Intrinsic regulation of oligodendrocyte lineage development .................................. 37 1.5.3.1 Specification ..................................................................................................... 37 1.5.3.2 Differentiation ................................................................................................... 38 1.5.3.2 Maturation/myelination..................................................................................... 41 1.5.4 Oligodendrocyte precursor cells (OPCs) in the adult CNS ...................................... 43 1.5.4.1 Characteristics ................................................................................................... 43 1.5.4.2 Axo-glial synapses ............................................................................................ 43 1.5.4.3 Lineage potential ............................................................................................... 46 1.5.4.4 Injury response .................................................................................................. 47 1.5.4.5 OPCs in the adult human CNS ......................................................................... 48 1.6 Central nervous system (CNS) myelination ................................................................. 49 1.6.1 Target selection ......................................................................................................... 50 1.6.2 Models of myelin wrapping mechanics .................................................................... 51 1.6.3 Growth of the myelin sheath ..................................................................................... 53 1.6.4 Myelin sheath compaction ........................................................................................ 54 1.6.4.1 Intracellular compaction ................................................................................... 55 1.6.4.2 Extracellular compaction .................................................................................. 56 ix  1.6.5 Myelinating glial support of axons ........................................................................... 57 1.6.6 Myelin plasticity/adaptive myelination in the adult CNS ......................................... 58 1.6.7 Oligodendrocyte turnover in the human CNS: implications for myelin plasticity ... 62 1.6.8 Inhibitors of remyelination ....................................................................................... 63 1.6.8.1 Myelin debris .................................................................................................... 63 1.6.8.2 LINGO-1 ........................................................................................................... 64 1.6.8.3 Wnt-signaling .................................................................................................... 64 1.6.8.4 Extracellular matrix (ECM) components .......................................................... 64 1.6.8.5 Inflammation ..................................................................................................... 65 1.7 Mouse spinal cord anatomy .......................................................................................... 66 1.7.1 Main descending tracts ............................................................................................. 66 1.7.2 Main ascending tracts ............................................................................................... 67 1.7.3 Cervical dorsal column: fasciculus gracilis and fasciculus cuneatus ....................... 69  Chapter 2: Isolated thoracic contusion injury induces robust responses in oligodendrocyte lineage cells remote to the lesion site ..........................................................................................71 2.1 Introduction ................................................................................................................... 71 2.2 Materials and methods .................................................................................................. 72 2.2.1 Animals ..................................................................................................................... 72 2.2.2 Tamoxifen (TMX) administration ............................................................................ 73 2.2.3 Spinal cord injury (SCI) ............................................................................................ 74 2.2.4 Animal sacrifice and tissue collection ...................................................................... 75 2.2.5 Immunohistochemical staining ................................................................................. 76 x  2.2.5.1 Primary antibodies ............................................................................................ 77 2.2.5.2 Secondary antibodies ........................................................................................ 79 2.2.6 Oil Red O staining .................................................................................................... 80 2.2.7 Imaging ..................................................................................................................... 81 2.2.8 Cervical and lumbar spinal cord section sampling protocol ..................................... 81 2.2.9 Image position sampling: whole-cord section analyses ............................................ 82 2.2.10 Image position sampling: fasciculus gracilis analyses ............................................. 83 2.2.11 Cell counting protocol............................................................................................... 84 2.2.12 Myelin sheath counting protocol .............................................................................. 85 2.2.13 Immunoreactivity (IR) thresholding protocol ........................................................... 85 2.2.14 Axonal number indices ............................................................................................. 87 2.2.15 White matter area indices.......................................................................................... 88 2.2.16 Statistical analyses .................................................................................................... 88 2.2.16.1 Whole-cord section analyses ............................................................................. 89 2.2.16.2 Fasciculus gracilis analyses ............................................................................. 89 2.3 Results ........................................................................................................................... 89 2.3.1 Injury parameters ...................................................................................................... 89 2.3.2 Transgene recombination .......................................................................................... 91 2.3.3 Histopathology: whole-cord section analyses ........................................................... 95 2.3.4 OPC proliferation: whole-cord section analyses..................................................... 103 2.3.5 Oligodendrocyte lineage cell densities: whole-cord section analyses .................... 106 2.3.6 Oligodendrogenesis: whole-cord section analyses ................................................. 108 2.3.7 De novo myelination: whole-cord section analyses ................................................ 110 xi  2.3.8 Newly-formed oligodendrocyte maturation: whole-cord section analyses ............ 112 2.3.9 Oligodendrocyte apoptosis...................................................................................... 114 2.3.10 Pathological changes: fasciculus gracilis analyses ................................................. 116 2.3.11 Oligodendrocyte lineage cells responses: fasciculus gracilis analyses .................. 123 2.3.12 OPC lineage plasticity............................................................................................. 127  Chapter 3: Discussion ................................................................................................................131 3.1 Introduction ................................................................................................................. 131 3.2 Overview of experimental findings ............................................................................ 132 3.3 Injury-induced remote OPC proliferation ................................................................... 134 3.4 Injury-induced remote oligodendrogenesis................................................................. 138 3.5 Maturation impairment of newly-formed oligodendrocytes ....................................... 141 3.5.1 Extrinsic factors ...................................................................................................... 142 3.5.1.1 Altered neural activity..................................................................................... 142 3.5.1.2 Microgliosis/inflammation .............................................................................. 142 3.5.1.3 Astrogliosis ..................................................................................................... 144 3.5.1.4 Oxidative stress ............................................................................................... 145 3.5.1.5 Reduced substrate permissiveness for myelination ........................................ 146 3.5.2 Intrinsic factors ....................................................................................................... 147 3.5.2.1 Transcriptional dysregulation ......................................................................... 147 3.5.2.2 Signaling dysregulation .................................................................................. 148 3.5.3 Absence of maturation impairment at lesion epicentre? ......................................... 150 3.6 Oligodendrocyte apoptosis remote to the lesion site .................................................. 150 xii  3.7 OPC lineage plasticity: reliability of immunohistochemical markers ........................ 152 3.8 Murine dorsal column: somatotopic vs. modality-based organization? ..................... 154 3.9 Oligodendrogenesis in degenerating axonal tracts: maladaptive repair response? ..... 154 3.9.1 Activity-independent pathological stimulation of oligodendrogenesis .................. 155 3.9.2 Detrimental effects of continual oligodendrogenesis? ............................................ 155 3.9.3 Alternative non-myelin related function of the OPC response? ............................. 157 3.9.4 Pathological-induced decoupling of oligodendrocyte developmental regulation? . 158 3.10 Remote effects of  isolated SCI on oligodendrocyte lineage cells: potential relation to cognitive deficits observed in SCI patients? ........................................................................... 158 3.11 Proposed model of experimental observations ........................................................... 159 3.15 Conclusion .................................................................................................................. 160 References ...................................................................................................................................162  xiii  List of Tables  Table 1.1 Primary antibodies ........................................................................................................ 78 Table 1.2 Secondary antibodies .................................................................................................... 80  xiv  List of Figures  Chapter 1 Figure 1.1 OPC lineage potential .................................................................................................. 47 Figure 1.2 Mouse spinal cord anatomy ......................................................................................... 68 Figure 1.3 Effect of injury on cervical fasciculus gracilis vs. fasciculus cuneatus ...................... 70  Chapter 2 Figure 2.1 Transgenic mouse lines and experimental timeline..................................................... 73 Figure 2.2 Tissue sampling approach ........................................................................................... 82 Figure 2.3 Lesion epicentre histopathology and injury parameters .............................................. 91 Figure 2.4 Transgene recombination in oligodendrocyte lineage cells ........................................ 94 Figure 2.5 Transgene recombination in non-oligodendrocyte lineage cells ................................. 95 Figure 2.6 Microgliosis in the cervical and lumbar spinal cord ................................................... 99 Figure 2.7 Axonal pathology in the cervical and lumbar spinal cord ......................................... 100 Figure 2.8 Absence of whole-cord section astrogliosis .............................................................. 101 Figure 2.9 Cell proliferation in the cervical and lumbar spinal cord .......................................... 102 Figure 2.10 OPC proliferation in the cervical and lumbar spinal cord ....................................... 105 Figure 2.11 Oligodendrocyte lineage cell densities in the cervical and lumbar spinal cord ...... 107 Figure 2.12 Oligodendrogenesis in the cervical and lumbar spinal cord .................................... 109 Figure 2.13 New myelination in the cervical and lumbar spinal cord ........................................ 112 Figure 2.14 Oligodendrocyte maturation in the cervical and lumbar spinal cord ...................... 114 Figure 2.15 Oligodendrocyte apoptosis in the cervical and lumbar spinal cord ......................... 116 xv  Figure 2.16 Microgliosis in the cervical fasciculus gracilis ....................................................... 119 Figure 2.17 Axonal pathology in the cervical fasciculus gracilis .............................................. 120 Figure 2.18 Astrogliosis in the cervical fasciculus gracilis ........................................................ 121 Figure 2.19 Myelin debris in the cervical fasciculus gracilis ..................................................... 122 Figure 2.20 OPC proliferation in the cervical fasciculus gracilis .............................................. 125 Figure 2.21 Oligodendrogenesis in the cervical fasciculus gracilis ........................................... 126 Figure 2.22 New myelination in the cervical fasciculus gracilis ................................................ 127 Figure 2.23 Absence of Schwann cell production from OPCs in the cervical and lumbar spinal cord ............................................................................................................................................. 129 Figure 2.24 Absence of OPC astrogenesis in the cervical and lumbar spinal cord .................... 130     xvi  List of Abbreviations NeuN – neuronal nuclei specific antigen GFAP – glial fibrillary acidic protein Aldh1L1 – aldehyde dehydrogenase 1 family member L1 OPC – oligodendrocyte precursor cell EdU – 5-ethynyl-2’-deoxyuridine BrdU – 5-bromo-2’-deoxyuridine CNS – central nervous system PNS – peripheral nervous system GFP – green fluorescent protein YFP – yellow fluorescent protein mGFP – membrane-tethered green fluorescent protein PDGFRα – platelet-derived growth factor receptor α PDGF – platelet-derived growth factor FGF – fibroblast growth factor EGF – epidermal growth factor SCI – spinal cord injury wpi – weeks post-injury dpi – days post-injury mpi – months post-injury Gst-π – glutathione S-transferase Pi SC – Schwann cell MS – multiple sclerosis xvii  MBP – myelin basic protein MOG – myelin oligodendrocyte glycoprotein MAG – myelin-associated glycoprotein PLP – proteolipid protein Olig2 – oligodendrocyte transcription factor 2 Olig1 – oligodendrocyte transcription factor 1 DNA – deoxyribonucleic acid RNA – ribonucleic acid mRNA – messenger ribonucleic acid β-APP – Beta amyloid precursor protein T6 – 6th thoracic spinal cord segment T7 – 7th thoracic spinal cord segment T9 – 9th thoracic spinal cord segment T10 – 10th thoracic spinal cord segment L4 – 4th lumbar spinal cord segment L5 – 5th lumbar spinal cord segment C4 – 4th cervical spinal cord segment C5 – 5th cervical spinal cord segment P0 – protein-zero Cre – Cre recombinase CreERT2 – tamoxifen-inducible estrogen receptor/Cre recombinase fusion protein FG – fasciculus gracilis FC – fasciculus cuneatus xviii  TMX – tamoxifen 4-OHT – 4-hydroxytamoxifen Iba-1 – ionized calcium-binding adaptor molecule 1 SMA – smooth muscle actin TTX – tetrodotoxin Kdyne – kilodyne (force measurement) CC1 – clone binding to APC APC – adenomatous polyposis coli CC-3 – cleaved-caspase 3 SMI-312 – Pan axonal neurofilament marker NF-200 – neurofilament 200 MyRF – myelin regulatory factor (a.k.a. MRF) TNF-α – tumor necrosis factor α IL-6 – interleukin-6 IL-1β – interleukin-1β IL-2 – interleukin-2 IL-11 – interleukin-11 IFN-γ – interferon-γ IHC – immunohistochemistry ICC – immunocytochemistry nm – nanometer mm – millimeter µm – micrometer xix  cm – centimeter ChABC – Chondroitinase ABC CSPG – Chondroitin sulphate proteoglycan NG2 – nerve glial antigen 2 (aka CSPG4) SMI-32 – non-phosphorylated neurofilament marker WM – white matter GM – gray matter Glut-1 – glucose transporter 1 NDS – normal donkey serum FSP1 – fibroblast-specific protein 1 ALS – amyotrophic lateral sclerosis NSPCs – neural stem/precursor cells bFGF – basic fibroblast growth factor (a.k.a. FGF-2) IGF-1 – insulin-like growth factor 1 CNTF – ciliary neurotrophic factor Brg1 – transcription activator Brg1 (a.k.a. ATP-dependent helicase SMARCA4) ER – estrogen receptor bp – base pair IH – Infinite Horizons CPG – central pattern generator HBD – hormone binding domain Hsp-90 – heat shock protein 90 TAF-2 – transcription initiation factor TFIID subunit 2 xx  Gpr17 – g-protein coupled receptor 17 Id2 – inhibitor of DNA binding 2 Id4 – inhibitor of DNA binding 4 TH – thyroid hormone BDNF – brain-derived neurotrophic factor  ECM – extracellular matrix CST – corticospinal tract dCST – dorsal corticospinal tract ATP – adenosine triphosphate EM – electron microscopy EAE – experimental autoimmune encephalomyelitis BM – bone marrow G1 – gap 1 phase ROS – reactive oxygen species BSCB – blood-spinal cord-barrier VZ – ventricular zone SVZ – sub-ventricular zone PNS – peripheral nervous system NO – nitric oxide MN – motor neuron Ngn-2 – neurogenin-2 pMN – motor neural progenitor domain PTEN – phosphatase and tensin homolog xxi  CSPG – chondroitin sulphate proteoglycan GABA – gamma amino butyric acid TTX – tetrodotoxin VM – membrane potential EPSP – excitatory post-synaptic potential NMDA – N-methyl-D-aspartate AMPA – α-amino-3-hydroxy-5-methyl-4-isoxazolepropionic acid AAF-1 – apoptosis activating factor 1 Ascl1 – achaete-scute homolog 1 TLR2 – toll-like receptor 2 Fgfr1 – fibroblast growth factor receptor 1 Fgfr2 – fibroblast growth factor receptor 2 MAG – myelin-associated glycoprotein MCT-1 – monocarboxylate transporter 1 MAPK – mitogen-activated protein kinase BMP – bone morphogenetic protein Shh – sonic hedgehog E – embryonic day TNFR1 – tumor necrosis factor 1 TNFR2 – tumor necrosis factor 2 DCC – deleted in colorectal carcinoma Sox5 – sex-determining region Y box 5 Sox6 – sex-determining region Y box 6 xxii  Sox9 – sex-determining region Y box 9 Sox10 – sex-determining region Y box 10 CNPase – 2’,3’-cyclic-nucleotide 3’-phosphodiesterase PDGF-B – platelet-derived growth factor B PDGF-C – platelet-derived growth factor C MMP-2 – matrix metalloproteinase 2 VEGF-A – vascular endothelial growth factor A NF-κB – nuclear factor κB CCL2 – chemokine (C-C motif) ligand 2 CXCL10 – C-X-C motif chemokine 10 (a.k.a. interferon-γ-induced protein 10) Olig1 – oligodendrocyte transcription factor 1 Olig2 – oligodendrocyte transcription factor 2 LINGO-1 – leucine rich repeat and immunoglobin-like domain-containing protein 1 TGF-β – transforming growth factor β ChIP-Seq – chromatin immunoprecipitation sequencing CYP2D6 – cytochrome P450 2D6 (hepatic enzyme) CD4 – cluster of differentiation 4 CD8 – cluster of differentiation 8 NF-1 – neurofibromatosis type 1 MRI – magnetic resonance imaging p53 – transformation-related protein 53 (a.k.a. phosphoprotein p53, cellular tumor antigen p53) Tcf712 – transcription factor 7-like 2 (T-cell specific, HMG-box) NFIA – nuclear factor 1 A-type xxiii  DAPT – N-[N-(3,5-difluorophenacetyl)-L-alanyl]-S-phenylglycine t-butyl ester (Notch inhibitor) Nkx2-2 – Nk 2 homeobox 2 Nkx6-1 – Nk 6 homeobox 1 Nkx6-2 – Nk 6 homeobox 2 Hes5 – Hes family basic helix-loop-helix transcription factor 5 bHLH – basic helix-loop-helix Nrg1-III – neuregulin-1 type III VSV-G – vesicular stomatitis virus G protein GluR2 – glutamate ionotropic receptor AMPA type subunit 2 ErbB – epidermal growth factor receptor (a.k.a. EGFR) Sirt2 – sirtuin 2 Zfhx1b – zinc finger E-box binding homeobox 2 IP – intraperitoneal injection SQ – subcutaneous injection IM – intramuscular injection DWMI – diffuse white matter injury WMI – white matter injury   xxiv  Glossary  Immunohistochemistry – the process of detecting antigens (e.g. proteins) in tissue samples utilizing the specificity of antibody-antigen binding interactions. In vitro – (‘within the glass’) studies performed on biological materials (e.g. cells, proteins, micro-organisms, etc.) outside of their natural biological context (e.g. cell cultures). In vivo – (‘within the living’) studies performed on biological materials (e.g. cells, proteins, etc.) in whole living organisms (e.g. animals, humans, plants, etc.). Optogenetics – a technique that enables light stimuli to control cellular processes (via genetic modification producing expression of light-sensitive ion channels). Slice culture – studies performed on living slices of tissue (e.g. hippocampus, etc.) that enable preservation of tissue architecture.  Cryostat – an apparatus for cutting frozen tissue samples into very fine slices (e.g. 20 µm thick), maintaining a low temperature (~-25 °C) to preserve tissue samples. Knockout – a genetic technique that enables the inactivation of genes within an organism or within a specific cell type. Reporter gene – a gene attached to a regulatory sequence of a gene of interest (hence expressed concurrently), chosen based on characteristics that are easily identified and measured (e.g. fluorescence). Promoter – a DNA sequence that intiates the transcription of a specific gene (e.g. PDGFRα promoter sequences initiate transcription of the PDGFRα gene). STOP codon – (a.k.a. termination codon) is a nucleotide triplet that terminates protein translation. Floxed – a sequence of DNA flanked by LoxP sites (i.e. Cre-recombinase recognition sites). xxv  Recombination – rearrangement of genetic material (e.g. Cre-mediated removal of floxed DNA sequences). Transgenic mouse – a mouse containing gene sequences from another organism (i.e. Cre-recombinase coding region, GFP gene, etc.) introduced via genetic engineering techniques. Excitotoxicity – a pathological process by which cells are damaged and/or killed by overactivation of glutamate receptors (e.g. AMPA or NMDA receptors). Oligodendrocyte – myelinating glial cell of the central nervous system (CNS) Astrocyte – a star-shaped glial cell of the central nervous system (CNS), involved in many functions, including biochemical support of endothelial cells, nutrient provision, maintenance of extracellular ion balance, and repair/scarring following injury.  Microglia – resident immune surveillance cells of the central nervous system (CNS), extremely sensitive to perturbations of CNS environment. Schwann cell – myelinating glial cell of the peripheral nervous system (PNS) Oligodendrocyte precursor cell – endogenous precursor cell of myelinating oligodendrocytes Neuron – electrically-excitable cell forming the basis of neural transmission in the CNS Central nervous system (CNS) – part of the nervous system consisting of the brain and spinal cord. Peripheral nervous system (PNS) – part of the nervous system consisting of the nerves and ganglia outside of the brain and spinal cord. Synapse – specialized connections between neurons and other cells, including other neurons, glia (i.e. OPCs), muscles, etc. Inducible – (genetics) the rapid activation of gene expression in response to an applied stimulus (e.g. tamoxifen administration induces Cre-mediated recombination of transgenes). xxvi  Conditional – (genetics) a knockout that is specific to a tissue or cell type, achieved via genetic engineering techniques that employ specific promoters. Electroporation – (a.k.a. electropermeabilization) is a microbiological technique in which an electrical field is used to increase cellular membrane permeability to allow for the introduction of chemicals, drugs, or DNA molecules. Stem cell – an undifferentiated cell capable of indefinite self-renewal and generation of cells of other lineages (via differentiation). Progenitor cell – an immature cell with a more restricted potential than a stem cell. Oligodendrogenesis – production of new oligodendrocytes from oligodendrocyte precursor cells (OPCs) via differentiation.  xxvii  Acknowledgements  I recognize my graduate supervisor Dr. Wolfram Tetzlaff for allowing me to conduct research in his laboratory.    I offer gratitude to my graduate committee members, Dr. Fabio Rossi and Dr. Matthew Ramer.  Appreciation is extended to members of the Tetzlaff laboratory for critical commentary, including Dr. Oscar Siera, Gregory Duncan, Peggy Assinck, Doug Brown, Sohrab Manesh, and Brett Hilton.  I acknowledge ICORD, Department of Zoology, and U.B.C. staff/faculty for assistance provided, both directly and through maintenance of facilities and equipment.  I am grateful for grant support provided by the NSERC Alexander Graham Bell Canada Graduate Scholarship (Master’s Level) and the CIHR Transplant Training Program administered by the U.B.C. Department of Surgery.    Sincere appreciation is extended to Nicole Janzen for all her help and support throughout.  Special thanks to Doug Brown, Sohrab Manesh, Fraser Muir, Peter Tan, and Oscar Siera for making the laboratory an enjoyable environment.   xxviii  Finally, as in all things, an immense debt of gratitude is owed to my parents, Dawn and Brian, for their unwavering support of my endeavors, both financially and personally. xxix  Dedication  This thesis is dedicated to my parents, Dawn and Brian.1  Chapter 1: Introduction The current thesis aims to characterize the responses of oligodendrocyte lineage cells remote to an isolated thoracic contusion spinal cord injury (SCI), specifically the assessment of OPC proliferation, oligodendrogenesis, and new myelination in the cervical and lumbar spinal cord.  To this end, we employed two separate transgenic mouse lines that enabled fate-mapping of oligodendrocyte precursor cells (OPCs) and their progeny (i.e. PDGFRα-CreERT2:ROSA26-mGFP and PDGFRα-CreERT2:ROSA26-YFP), subjecting them to a 70 Kdyne thoracic contusion injury, followed by immunohistochemical assessment of spinal cord tissue harvested at multiple post-injury time-points.  Based on previous demonstration of OPC responsiveness to various CNS pathological conditions, the influence of axonal degeneration on oligodendrocyte lineage cells, and remote effects of isolated SCI, we hypothesized that an isolated thoracic contusion injury would induce robust and rapid responses in remote oligodendrocyte lineage cell populations, manifesting as increased OPC proliferation, oligodendrogenesis, and increased new myelination.       1.1 Spinal cord injury (SCI): clinical presentation Dependent upon the injury level and severity, the neurological deficits accompanying SCI include paralysis, skeletal muscle spasticity, abnormalities in sensory modalities (e.g. somatosensory function, thermo-sensing, vibration sensing, etc.), neuropathic pain, as well as dysfunctions in various body systems (i.e. cardiovascular, respiratory, bowel, bladder, and sexual organs) (Levi et al. 1996; Blight 2002; Zeilig et al. 2012; Teasell et al. 2010).  The specific pattern of functional deficits is directly related to spinal cord anatomy, specifically the arrangement of long ascending and descending tracts (running in the white matter) and local interneurons and cell bodies (occupying the gray matter) (Blight 2002, Watson & Harrison 2012).  Damage to the white matter 2  disrupts communication between the brain and/or supraspinal centres and innervated peripheral features below the injury level (Blight 2002; Norenberg 2004).  The manifestation of functional deficits is a direct correlate of specific tract damage (e.g. corticospinal tract, rubrospinal tract, etc.).  This anatomical arrangement means that peripheral organs and musculature innervated by spinal nerves above the level of injury remain functional, as communication is not disrupted (Blight 2002; Norenberg 2004).  In addition to white matter, traumatic SCI typically incurs gray matter damage as well, resulting in the loss of interneurons involved in local reflex pathways, central pattern generator (CPG) activity, as well as propriospinal connections between spinal cord levels (Flynn et al. 2011).  This interneuron loss presents as functional deficits in reflexive movements, stereotypic motor responses, and/or motor coordination (Flynn et al. 2011).  Furthermore, preserved intra-spinal circuitry below the injury level may be rendered non-functional if reliant on supra-spinal input.  For example, the loss of supra-spinal stimulation obviates voluntary control of CPG-mediated locomotion, despite preservation of CPG circuitry (common in higher injuries) (Dietz & Michel 2009; Guertin 2013).  In addition to direct neurological impairments, SCI is often accompanied by a host of complications related to autonomic dysfunction, including deep vein thrombosis, urinary tract infections, pressure ulcers, and respiratory infections (Levi et al. 1996; Winslow & Rozovsky 2003; Dryden et al. 2004; DeVivo & Farris 2011), which are common causes of mortality for SCI patients (Dryden et al. 2004; Strauss et al. 2006; Christie et al. 2010).  Human SCI is highly heterogeneous, both in terms of injury mechanism (e.g. contusion, compression, dislocation, laceration, etc.) and functional deficits incurred (dependent upon injury location and severity of tissue damage) (Kirshblum et al. 2002; Norenberg et al. 2004).  Furthermore, SCI is generally classified as either traumatic (e.g. motor vehicle accidents, sports injuries, etc.) or non-traumatic (e.g. tumors, infections, etc.) (Norenberg et al. 2004).  Interestingly, 3  males are over-represented amongst SCI patients, experiencing ~3-fold higher incidence than females (Dryden et al. 2004; Wyndaele & Wyndaele 2006; Couris et al. 2009).  The most common cause of SCI in developed nations is motor vehicle accidents, with a notable increase in falls (correlated with an older population), as well as significant contributions from sports accidents and violent acts (i.e. gun or knife injuries) (Tator 1995; Dryden et al. 2004; Pickett et al. 2006; Couris et al. 2009; Cripps et al. 2011).  Histologically, vertebral dislocations and spine fractures often result in spinal cord contusion, accompanied by varying degrees of compression and/or dislocation (Bunge et al. 1993; Dumont et al. 2001; Norenberg et al. 2004).  Indeed, contusion is the most common mechanism of human SCI (Dumont et al. 2001; Norenberg et al. 2004).   In contrast, lacerations and full cord transections represent a small minority of cases, caused by severe dislocations of the vertebrae, bone splinters, or foreign bodies (i.e. bullets or knife blades) (Bunge et al. 1993; Dumont et al. 2001; Norenberg et al. 2004).  It is important to note that in contrast to pre-clinical animal models, human SCI is typically a combination of multiple injury mechanisms (Kwon et al. 2002; Cheriyan et al. 2014), presenting challenges for clinical translation.  1.2 Pre-clinical models of spinal cord injury (SCI) The purpose of pre-clinical animal models is to recreate features of human SCI as accurately as possible, enabling the assessment of pathophysiological changes and testing of putative therapeutics (Metz et al. 2000; Choo et al. 2008; Kwon et al. 2002).  Rat (Rattus norvegicus) models of SCI are commonly used, as they display similar functional, electrophysiological, histopathological, and morphological outcomes as human SCI (Metz et al. 2000; Dunham et al. 2010).  Studies requiring genetic modifications (e.g. fate-mapping, knockout studies, etc.) typically employ mice (Mus musculus) due to the extensive development of transgenic mouse strains.  Non-4  human primates (i.e. marmosets, macaques, and squirrel monkeys) have also been utilized, predominately in anatomical or therapeutic assessments as they more closely approximate human SCI than murine models (Rosenzweig et al. 2010; Iwanami et al. 2005).  Most pre-clinical SCI studies employ animal models of mid-thoracic injuries (e.g. the current study), however interest has begun to shift to cervical injuries, due to prevalence amongst the human SCI population (particularly resulting from non-penetrating trauma, such as sports injury or motor vehicle accidents) (Dunham et al. 2010).  Pre-clinical SCI mechanisms are generally classified as contusion, compression, distraction, dislocation, laceration, or transection (Cheriyan et al. 2014; Kwon et al. 2002).  Focus here will be on contusion injury, as that it the mechanism employed by the current study.  1.2.1 Murine model of contusion spinal cord injury (SCI) using the Infinite Horizons (IH) impactor Contusion injury devices are designed to inflict a transient, acute injury to the exposed murine spinal cord tissue, including weight-drop apparatuses, electromagnetic impactors, and pressurized air impactors (Allen 1911; Kwon et al. 2002; Cheriyan et al. 2014; Choo et al. 2008).  The most widely employed contusion devices are the New York University (NYU)/Multicentre Animal Spinal Cord Injury Study (MASCIS) impactor, the Infinite Horizons (IH) impactor, and the Ohio State University (OSU) impactor (Kwon et al. 2002; Cheriyan et al. 2014; Basso et al. 1996; Gruner 1992).  The IH impactor is used in the current study, therefore will be the focus of further discussion.  Instead of weight drop height, the IH impactor employs a force-controlled impact, achieved by interfacing a stepping motor with an external computer (Scheff et al. 2003; Choo et al. 2008; Cheriyan et al. 2014).  A sensor attached to the impactor head directly measures the force 5  exerted on the spinal cord tissue, withdrawing the head once the force is met, thus minimizing injury variability.  Furthermore, automatic withdrawal of the impactor head prevents the ‘weight-bounce’ phenomenon observed with other contusion devices (Cheriyan et al. 2014).  The attached sensor measures force of impact, velocity of impactor head at impact, and displacement of the spinal cord tissue.  Force and velocity are typically consistent across animals as they are pre-set, whereas displacement varies according to tissue characteristics (i.e. resistance), which can be heterogeneous.  A commonly identified limitation of the IH impactor is the variability in securing the spinal column using the provided clamps (Kwon et al. 2002; Scheff et al. 2003; Cheriyan et al. 2014) .  To address this, some researchers have developed custom clamps that prove more reliable (Lam et al. 2014; Choo et al. 2008).  However, as inconsistent parenchymal injury and variability in functional deficits following injury with the IH impactor have been reported, this may require further consideration (Scheff et al. 2003; Cheriyan et al. 2014).  1.2.2 Cre-lox biochemistry Cre-recombinase is a 38 kD protein of the integrase family that is derived from the P1 bacteriophage (Nagy et al. 2000; Guo et al. 1997).  Its native role is two-fold: (i) cyclization of the linear phage genome following host cell injection, and (ii) separation of P1 dimers formed through homologous recombination to ensure continual infection of daughter cells following division (Ghosh & Van Duyne 2002).  Cre-recombinase catalyzes recombination between two of its recognition sites, called loxP sites (Hamilton & Abremski 1984).  LoxP sites consist of a 34-bp consensus sequence, containing a core spacer sequence of 8 bp flanked by two 13 bp palindromic sequences (Hamilton & Abremski 1984; Nagy et al. 2000).  Asymmetry of the core sequence defines loxP site orientation (Hamilton & Abremski 1984).  The molecular mechanism is as 6  follows: (i) Cre-recombinase proteins bind to the flanking palindromic sequences of each loxP site, (ii) subsequent tetramer formation between Cre proteins brings loxP sites into close apposition, (iii) Cre-recombinase catalyses recombination between the sites (Pathania et al. 1999; Ghosh & Van Duyne 2002).  The outcome of the recombination depends upon the location and orientation of the loxP sites.  For example, Cre-mediated recombination can result in excision or inversion of a floxed sequence (e.g. cis loxP sites) or insertion/translocation between chromosomes (e.g. trans loxP sites) (Ghosh & Van Duyne 2002; Nagy et al. 2000).  Large distance between loxP sites requires Cre-recombinase to be expressed under a strong promoter for effective recombination.  For example, excision of a ~400 kb genomic region was achieved with 50% efficiency with Cre-recombinase expressed under the control of the remarkably strong CMV enhancer/chicken β-actin promoter combination (Hitoshi et al. 1991; Nagy et al. 1998).  Critical to the efficacy of the Cre-lox system, random occurrence of a 34-bp sequence matching the loxP sequence requires a 1018 bp genome.  As the mouse genome is 2.8 x 109 bp in length, the probability of a loxP sequence occurring randomly is extremely low (Nagy et al. 2000).  Therefore, Cre-mediated recombination in transgenic mice is assumed to be restricted to the exogenously introduced loxP sites.  1.2.3 Inducible-conditional estrogen-receptor/Cre-recombinase transgenic mice The development of transgenic mouse models in which gene function can be modulated has provided immense insight into the pathogenesis of various diseases (e.g. cancer, genetic diseases, traumatic injury, autoimmune disorders, etc.) (Whitfield et al. 2015; Heintz 2001; Hanahan 1988).  Of these models, the tamoxifen/estrogen receptor system is one of the best characterized, enabling the temporal control of gene expression (Danielian et al. 1998; Hayashi & McMahon 2002; Littlewood et al. 1995; R. Feil et al. 1996).  The basis of this system is the acquisition of hormone-7  dependence by previously non-hormone responsive intracellular proteins (Mattioni et al. 1994; Littlewood et al. 1995).  Initial in vitro studies reported that fusion of intracellular proteins (e.g. kinases, transcription factors, etc.) with the hormone-binding domain (HBD) of steroid receptors could convey hormone-dependence on the proteins (Mattioni et al. 1994; Picard 1993).  HBD-fusion proteins are inactive when the ligand is not bound, complexed with a variety of intracellular peptides (e.g. Hsp90) (Pratt 1990; Smith & Toft 1993).  Release from the inhibitory complexes is induced by binding of the cognate ligand to the HBD.  Five vertebrate steroid receptors (i.e. glucocorticoid, mineralocorticoid, androgen, progesterone, and estrogen receptors) have been characterized (Whitfield et al. 2015).  Of these, the estrogen receptor (ER) has been most widely used as a heterologous regulatory domain, however its application has important limitations.  For example, it possesses inherent ligand-dependent transactivation activity (TAF-2 or AF-2) (Berry et al. 1990; Littlewood et al. 1995), therefore utilization in combination with a heterologous partner with weak transcriptional activity or with a transcriptional repressor complicates interpretation of the results.  Furthermore, endogenous 17β-estradiol (ligand for ER) precludes the use of this system in vivo, due to non-specific activation of the receptor (Whitfield et al. 2015).  However, the development of a mutant ER (G525R) with ~1000-fold lower affinity for 17β-estradiol and no TAF-2 transactivation activity enabled application of this system to in vivo models (Danielian et al. 1993).  Critically, the G525R mutant ER retains responsiveness to 4-hydroxytamoxifen (4-OHT, a synthetic estrogen analog), therefore enabling activation (Danielian et al. 1993).  As 4-OHT is expensive, animals are typically dosed with the precursor tamoxifen (as is done in the current study), which is subsequently converted by the hepatic enzyme CYP2D6 into 4-OHT (Bijl et al. 2009; Hoskins et al. 2009).  Further improvement of the ER/HBD system was accomplished by the creation of two ER/HBD variants, each containing a triplet mutation.  The first mutant, 8  known as ERT2 (G400V/M543A/L544A) has 4-fold higher induction efficiency by 4-OHT compared to the original ERTAM (Feil et al. 1997).  The second variant (G400V/L539A/L540A) responds to the synthetic anti-estrogen ICI 182,780 (ICI), but remains non-responsive to 17β-estradiol (Feil et al. 1997).  In vivo application of the inducible ER system in mouse models was revolutionary for the control of gene expression.  Specifically, production of a tamoxifen-dependent Cre-recombinase (Cre-ERTAM) enabled the specific, inducible modification to the mouse genome at a chosen time during the lifespan of the organism (as determined by time of tamoxifen administration) (Feil et al. 1996; Danielian et al. 1998; Hayashi & McMahon 2002).  The generation of transgenic mouse lines expressing Cre-ERTAM under the control of specific promoters enable the excision of chromosomally integrated gene sequences that are flanked by loxP sites upon tamoxifen administration (Danielian et al. 1993; Feil et al. 1997; Hayashi & McMahon 2002).  Characterization of the Cre-ERT2 system revealed a ~10-fold greater sensitivity to 4-OHT as compared to Cre-ERTAM (Indra et al. 1999).  Fate-mapping approaches utilize the Cre-ERTAM system cross-bred to transgenic mice expressing a conditional transgene for a reporter protein (e.g. mGFP, YFP, etc.) (Rivers et al. 2008; Young et al. 2013; Zawadzka et al. 2010).  The conditional transgene consists of promoter and coding region of the gene of interest separated by a floxed STOP codon (i.e. STOP codon flanked by loxP sites) (Nagy et al. 2000).  Therefore, without Cre-recombinase activity the STOP codon prevents gene expression.  Upon tamoxifen administration, Cre-mediated excision of the genomic region containing the STOP codon enables expression of the gene sequence.  Typically, the conditional transgene is driven by a ubiquitous promoter (e.g. ROSA26) while the Cre-ERT2 is expressed under the control of a lineage- or cell type-specific promoter (e.g. PDGFRα) (Nagy et al. 2000).  The current study utilizes PDGFRα-CreERT2:ROSA26-mGFP and PDGFRα-CreERT2:ROSA26-YFP transgenic reporter mouse lines.     9   1.3 Spinal cord injury (SCI): pathophysiology SCI pathophysiology is generally separated into two distinct, but temporally subsequent, processes: (i) the primary injury and (ii) the secondary injury.  The primary injury consists of tissue damage directly attributed to the initial mechanical impact (e.g. bone fragment impingement), whereas the secondary injury is a compilation of protracted pathophysiological processes (e.g. ischemia, inflammation, oxidative stress, apoptosis, etc.) triggered by the primary injury that propagate tissue damage (both spatially and temporally) (Tator & Fehlings 1991; Tator 1995).   1.3.1 Primary injury The primary injury consists of the initial damage due to the mechanical impact to the spinal cord.  Various mechanisms of primary injury have been identified, including contusion, compression, distraction, dislocation, and laceration (Dumont et al. 2001).  Most pre-clinical studies employ animal models of contusion SCI (as does this study), as they are generally thought to most accurately replicate human pathophysiology (Kwon et al. 2002; Dietz & Curt 2006), and contusion is the most common among SCI patients (Bunge et al. 1993; Dumont et al. 2001; Norenberg et al. 2004).  The foremost pathological consequences of the primary injury are (i) vasculature disruption (leading to hemorrhage, edema, and reduced perfusion of injury site) and (ii) necrotic cell death (encompassing neurons, astrocytes, oligodendrocytes, and endothelial cells) through physical disruption of cellular membranes (Choo et al. 2008; Mothe & Tator 2012).  Gray matter is more affected than adjacent white matter regions by the primary injury, as it is generally considered to be less resilient to physical damage and is heavily vascularized (Norenberg et al. 2004; Dumont et al. 2001; Tator & Koyanagi 1997). 10   1.3.2 Secondary injury In addition to the immediate damage, aspects of the primary injury (specifically necrosis and vascular disruption) trigger secondary injury processes (e.g. hemorrhaging, ischemia, excitotoxicity, oxidative stress, infiltration of blood-borne inflammatory cells, activation of microglia, and progressive cell death) that collectively spread damage to adjacent tissue regions that were initially unaffected or mildly affected by the primary impact (Tator & Fehlings 1991; Tator 1995).    1.3.2.1 Hemorrhage Initially, hemorrhaging is localized to the lesion epicentre, but within 12-18 hrs it spreads to adjacent spared tissue regions, inducing substantial cell loss (in part due to iron accumulation resulting from hemoglobin degradation) (Simard et al. 2007; Tator & Koyanagi 1997; Regan et al. 2008; Hua et al. 2006).  Loss of blood-spinal cord-barrier (BSCB) integrity resulting from physical damage to cellular components (i.e. endothelial cells or astrocytes, which maintain BSCB endothelial junctions through their end-feet), serves to propagate the secondary injury (Saadoun & Papadopoulos 2010; Bartanusz et al. 2011).      1.3.2.2 Ischemia Ischemia following SCI is largely attributed to localized vascular damage and subsequent responses (i.e. vasospasm, thrombosis, loss of homeostatic vascular regulation), as well as 11  systemic hypo-perfusion due to neurogenic shock (characterized by bradycardia, reduced cardiac output, and decreased peripheral resistance) (Dumont et al. 2001).  Over time, vascular damage and BSCB breakdown spread to adjacent spared tissue regions, due to the continued production/release of various compounds (e.g. endothelin-1, neurotransmitters, ROS, bradykinins, and histamines) from damaged cells, activated glia (i.e. microglia, astrocytes), and infiltrating blood-borne inflammatory cells (e.g. monocytes, macrophages) (Noble & Wrathall 1989; Tator & Fehlings 1991; Bartanusz et al. 2011; Popovich et al. 1996).  This spreading results in exposure of initially spared tissue components to ischemia, edema, ionic perturbations, as well as blood components not found in CNS under physiological conditions (e.g. serum proteins, macrophages, etc.), representing a significant contributor to the continual damage following injury (Noble & Wrathall 1989; Tator & Fehlings 1991; Popovich et al. 1996).  Serum components have been implicated in resident microglial activation following SCI (Ransohoff & Perry 2009; Takigawa et al. 2009).  BSCB breakdown peaks within days of the injury, gradually declining over a period of several weeks (Noble & Wrathall 1989; Popovich et al. 1996; Bilgen et al. 2001).  Swelling (due to edema) compresses the spinal cord tissue against the overlying dura mater and vertebrae, inducing additional damage and worsening ischemia (through constriction of superficial blood vessels) (Saadoun & Papadopoulos 2010; Saadoun et al. 2008).  Continued necrotic death worsens secondary injury due to the uncontrolled release of ions, excitatory neurotransmitters, and reactive oxygen species (ROS) (Tator & Fehlings 1991; Dumont et al. 2001).  Moreover, ischemia propagates cellular death as it deprives cells of sufficient oxygen and glucose to support metabolic functions (Harris & Attwell 2012), leading to reduced ATP synthesis, rapid depletion of cellular ATP stores, and subsequent catastrophic failure of ionic gradient maintenance, culminating in cellular edema, swelling, and oncotic cellular death (Liang et al. 2007; Szydlowska & Tymianski 12  2010).  Indeed, sustained ischemia is implicated in the progressive and sustained neuronal and glial cell loss over time (Liang et al. 2007; Saadoun & Papadopoulos 2010).    1.3.2.3 Excitotoxicity Propagation of damage to spared tissue regions is also attributed to the release of excitatory neurotransmitters, a consequence of ischemia, necrotic cell death, and astrocytic dysfunction (Panter et al. 1990; Wrathall et al. 1996).  Glutamate levels rapidly rise following SCI, leading to NMDA, AMPA, and kainate receptor activation, the release of more glutamate, and culminating in elevated intracellular Ca2+ concentration (Zhou et al. 2013; Matute et al. 2007).  Intracellular Ca2+ accumulation induces activation of Ca2+-dependent proteases, lipases, phosphatases, and endonucleases that collectively incur acidosis, cellular edema, reduced ATP synthesis, perturbation of ionic transporters, oxidative stress, damage to intracellular components (e.g. membranes, DNA, cytoskeleton, etc.), and organelle dysfunction (e.g. endoplasmic reticulum, mitochondria) (Dumont et al. 2001; Szydlowska & Tymianski 2010; Lewén et al. 2000; Higuchi et al. 2005).  The outcome of these responses is dependent on the relative severity of the excitotoxic insult.  For example, in regions with more severe secondary injury (i.e. injury epicentre) these processes typically lead to necrotic cell death, whereas away from the epicentre the affected cells predominately undergo apoptosis (Dumont et al. 2001; Szydlowska & Tymianski 2010).  Ca2+-induced apoptosis is mediated by the release of mitochondrial cytochrome c, which together with AAF-1 (apoptosis activating factor-1) activates caspase-9, which in turn stimulates caspase-3 and caspase-6 activity (the effectors of programmed cell death) (Budd et al. 2000; Eldadah & Faden 2000).    13  1.3.2.4 Oxidative stress In addition to excitotoxicity, oxidative stress is also a major contributor to cell loss in the injured spinal cord, specifically heightened during tissue reperfusion (Hamann & Shi 2009; Hamann et al. 2008; Yune et al. 2008).  Elevated ROS production leads to oxidation of various cellular components (e.g. proteins, lipids, and nucleic acids) and damages key cellular constituents (e.g. mitochondria, cellular membrane) (Barja 2004; Jia et al. 2012).  In particular, oxidative stress-induced mitochondrial dysfunction compromises cellular energy production, as well as increases ROS production, thus propagating oxidative damage via a positive-feedback loop (Jia et al. 2012).  As compared to other tissues, oxidative stress is particularly prominent in the injured CNS due to several factors, including: (i) concentration of polyunsaturated fatty acids (vulnerable to lipid peroxidation) in glial and neuronal membranes, (ii) high rate of oxidative metabolism, (iii) large yield of oxygen and nitrogen metabolites, and (iv) reduced efficacy of anti-oxidant defence mechanisms (Vaziri et al. 2004; Logan et al. 2005; Hamann et al. 2008).  In particular, lipid peroxidation after SCI contributes extensively to membrane damage, as well as generates by-products that propagate tissue damage (Jia et al. 2012).  For example, acrolein (a non-radical oxidant) has been implicated in membrane damage, mitochondrial dysfunction, and myelin disruption following SCI, particularly detrimental due to its prolonged half-life (compared to ROS) (Hamann et al. 2008; Hamann & Shi 2009; Shi et al. 2011).     1.3.2.5 Blood-borne inflammatory cells As a consequence of BSCB breakdown, blood-borne inflammatory cells enter the CNS parenchyma, starting with neutrophils (beginning within hours, clearing by 5-10 dpi) (Taoka et al. 14  1997; Carlson et al. 1998; Fleming et al. 2006; Beck et al. 2010).  Infiltrating neutrophils release various oxidative and proteolytic enzymes that sterilize the lesion site, however excessive release damages spared tissue (‘bystander effect’).  Indeed, neutrophil-derived elastase (a protease) damages endothelial cells in spared tissue regions, compromising vascular integrity and propagating injury (Taoka et al. 1997; Carlos & Harlan 1994).  In addition to neutrophils and macrophages, T-lymphocytes also infiltrate the CNS parenchyma during the first week post-injury, predominately at the lesion epicentre, where they persist for 6-10 wpi (although at substantially lower numbers than macrophages or neutrophils) (Popovich et al. 1997; Fleming et al. 2006).  The contribution of T-lymphocytes to the secondary damage is controversial, however they can directly lyse oligodendrocytes (Antel et al. 1994), as well as induce apoptosis through death receptor activation (Almad et al. 2011).   1.3.2.6 Microgliosis In addition to infiltrating inflammatory cells, endogenous microglia respond rapidly (within seconds) to the injury (Davalos et al. 2005; Nimmerjahn et al. 2005; Hines et al. 2009), partially attributed to P2Y12 purinergic receptor activation by released nucleotides (e.g. ATP, from damaged cells and/or reactive astrocytes), as well as serum components and/or glutamate (released from damaged endothelial and/or neural cells) (Davalos et al. 2005; Haynes et al. 2006; Ransohoff & Perry 2009).  Activated microglia proliferate, acquire phagocytic morphology, and upregulate a host of cytokines, chemokines, and eicosanoids (David & Kroner 2011).  The density of activated microglia peaks at 1 wpi (Popovich et al. 1997), coincident with peak density of hematogenous macrophages (5-7 dpi), but persist long term in the injured spinal cord (weeks to months post-injury) (Popovich et al. 1997; Carlson et al. 1998; Fleming et al. 2006).  Activated microglia 15  contribute to repair processes through removal of tissue debris, however can also propagate damage to spared tissue through release of proteolytic enzymes, free radicals (e.g. NO), and pro-inflammatory cytokines (e.g. IL-1β, TNF-α) (Chao et al. 1992; Gensel et al. 2009; David & Kroner 2011).  In the injured spinal cord, the majority of microglia/macrophages display the pro-inflammatory M1 phenotype (Mosser & Edwards 2008; Kigerl et al. 2009).  In other injured tissues, M1 macrophages decline and are largely replaced by anti-inflammatory M2 macrophages that promote tissue repair (Mosser & Edwards 2008), a transition recapitulated in vitro by myelin debris phagocytosis by M1 macrophages (Boven et al. 2006; Liu et al. 2006).  However, this transition does not seem to occur following SCI (despite myelin debris phagocytosis) (Kigerl et al. 2009), attributed to TNF-α production by iron-containing macrophages at the lesion epicentre (resulting from hemoglobin degradation) (Kroner et al. 2014).  The persistence of M1 phenotypes in the injured spinal cord is implicated in the propagation of damage to spared tissue (Kigerl et al. 2009; Kroner et al. 2014; David & Kroner 2011).    1.3.2.7 Progressive cell death In addition to acute cell death at the injury epicentre (including both necrosis and apoptosis), progressive cell loss occurs in spared tissue regions for weeks following SCI, consisting mainly of apoptosis (Tator & Fehlings 1991; Tator 1995).  For example, oligodendrocyte apoptosis continues up to 2-3 wpi, contributing to the extensive demyelination of spared axons in white matter regions following SCI (Crowe et al. 1997; Rosenberg & Wrathall 1997; Grossman et al. 2001; Blight 1985; Totoiu & Keirstead 2005; Almad et al. 2011).  In addition to oligodendrocytes, apoptotic microglia, neurons, and astrocytes (in severe injuries) have been observed, in both white and gray matter (Crowe et al. 1997; Yong et al. 1998; Casha et al. 2001).  Reduction of apoptosis following 16  SCI (via administration of a caspase inhibitor) substantially reduced lesion size and improved functional recovery, implicating apoptosis as a key contributor to secondary damage (Kanno et al. 2009; Kanno et al. 2011).  Oligodendrocyte apoptosis occurs along degenerating axonal tracts up to 3-4 wpi in rodent, non-human primate, as well as human SCI (Crowe et al. 1997; Casha et al. 2001; Warden et al. 2001).  This protracted apoptosis is attributed to slow axonal degeneration in the CNS, as compared to the PNS, meaning that the continual loss of axons will correlate with progressive oligodendrocyte loss (Vargas & Barres 2007; Perry et al. 1987; Becerra et al. 1995; Buss et al. 2004).  However, this is not likely to incur additional functional deficits, as the axons are already transected (Vargas & Barres 2007).  In addition to apoptotic and necrotic cell death, autophagic cell death has also been reported following SCI (Kanno et al. 2009; Kanno et al. 2011).  Autophagy can be a protective cellular response in injury environments, as it enables cells to recycle damaged cellular constituents (e.g. aggregated proteins, dysfunctional organelles, etc.) (Mao & Reddy 2010; Xu & Zhang 2011), however excessive levels following SCI have been implicated in axonal degeneration, cellular atrophy, and cell death, thus contributing to secondary tissue damage (Cherra & Chu 2008; Xu & Zhang 2011).  1.3.3 Pathophysiological effects remote to the lesion site The pathophysiology of SCI has been well characterized at injury epicentre and adjacent tissue regions (Tator 1995; Tator & Fehlings 1991), but remote to the lesion it is less well characterized.  Previous findings however suggest that isolated spinal cord injuries induce wide-spread pathology across the neuraxis.  Indeed, isolated thoracic contusion injury in rats induces extensive microgliosis across the brain and upregulation of inflammatory markers, correlated with chronically impaired cognitive function (Wu et al. 2014a).  This effect was replicated in mouse 17  contusion models, with additional revelation of substantial hippocampal neuronal loss, proposed to partially underlie the cognitive deficits (Wu et al. 2014b).  Moreover, isolated thoracic contusion injury in rats induces remote gliogenesis (including microglial, astroglial, and OPC proliferation) in the cervical spinal cord, but does not increase either SVZ or hippocampal neurogenesis (Franz et al. 2014).  OPC proliferation and oligodendrogenesis occur along degenerating axonal tracts in dorsal rhizotomy (Sun et al. 2010), axotomy (Nielsen et al. 2006), and ALS models (Kang et al. 2013).   Taken together, these studies suggest isolated spinal contusion injuries may induce substantive changes in remote endogenous cell populations (e.g. oligodendrocyte lineage cells and microglia), specifically along degenerating axonal tracts.    1.4 Endogenous cell responses to spinal cord injury (SCI) Endogenous cell populations in the adult mammalian spinal cord (e.g. neural stem/progenitor cells, astrocytes, neurons, oligodendrocytes, oligodendrocyte precursor cells, microglia) respond to injury in characteristic ways, dependent upon the location, severity, and type of injury inflicted to the spinal cord, as well as cell intrinsic properties (i.e. proliferative capacity, differentiation ability, vulnerability to damage, lineage plasticity, etc.) (Barnabé-Heider et al. 2010; Meletis et al. 2008; David & Kroner 2011).  Detailed characterization of these responses is critical for understanding pathological processes initiated by the injury, but also for the design of potential therapeutic interventions aimed at manipulating endogenous responses (e.g. in vivo reprogramming to replace lost cells, promoting remyelination, mitigating secondary damage, etc.) (Guo et al. 2014; Plemel et al. 2014).  18  1.4.1 Neural stem/progenitor cells (NSPCs) Neural stem/progenitor cells (NSPCs) are proposed to reside in multiple niches in the adult mammalian spinal cord, including ependymal, sub-pial, and intra-parenchymal regions (Horner et al. 2000; Yamamoto et al. 2001; Barnabé-Heider et al. 2010).  The designation of these cell populations as NSPCs is based on the ability to produce neurons, astrocytes, and oligodendrocytes (i.e. 3 main neural cell types) under appropriate in vitro conditions (Weiss et al. 1996; Yamamoto et al. 2001; Mothe et al. 2011).  Despite the presence of these NSPCs, the adult mammalian spinal cord is considered non-neurogenic (i.e. does not produce new neurons), even under pathological conditions (Horky et al. 2006; Horner et al. 2000; Yamamoto et al. 2001; Zai & Wrathall 2005).  Therefore, the injury-induced proliferation of NSPCs following SCI generates glial cells, restoring glial cell densities in the spared tissue rim to pre-injury levels by 2-6 wpi (Wrathall et al. 1996; Rosenberg et al. 2005; Tripathi & McTigue 2007), despite the fact that ~50% of oligodendrocytes and astrocytes are lost in the tissue rim by 1 dpi following contusion SCI (Grossman et al. 2001).  1.4.2 Astrocytes Astrocyte death following SCI has been noted, resulting from primary and secondary injury processes (Jaeger & Blight 1997; Yong et al. 1998; Grossman et al. 2001).  Reactive astrogliosis is a common feature of several CNS pathologies, characterized by increased GFAP expression, astrocytic cell hypertrophy, and increased deposition of chondroitin sulfate proteoglycans (CSPGs) (Fitch & Silver, 1997; Popovich et al. 1997; Fawcett & Asher 1999).  Around the SCI lesion site, astrogliosis is a critical component of the glial scar, which serves as a physical barrier to limit spread of the necrotic damage to adjacent spared tissue regions (Faulkner et al. 2004; Okada et al. 2006; Wanner et al. 2013).  Indeed, selective ablation of glial scar astrocytes 19  substantially increases lesion size and worsens functional outcomes (Bush et al. 1999; Faulkner et al. 2004).  The astrocytic component of the glial scar consists of pre-existing astrocytes spared by the injury, as well as new astrocytes generated from NSPCs (Meletis et al. 2008; Barnabé-Heider et al. 2010).  Reactive astrocytes also uptake extracellular glutamate, thus mitigating excitotoxic damage to spared cellular components (Swanson et al. 2004; Farina et al. 2007).  Despite these beneficial functions, astrogliosis also contributes to secondary damage after SCI (Swanson et al. 2004; Farina et al. 2007).  For example, reactive astrocytes release ATP (which induces microgliosis) (Davalos et al. 2005), as well as a number of chemokines (e.g. CXCL10, CCL2) and cytokines (e.g. TNF-α, IL-1β, & IL-6) that promote inflammatory cell infiltration and proliferation (Swanson et al. 2004).  Indeed, knockout of astrocytic NF-κB following SCI (thus preventing activation) reduces inflammation, has a neuroprotective effect, and improves functional recovery (Brambilla et al. 2005; Brambilla et al. 2009).  Moreover, reactive astrocytes produce factors that promote BSCB breakdown (e.g. VEGF-A, MMP-2), propagating edema (Swanson et al. 2004; Argaw et al. 2009).  Reactive astrocytes also produce NO in the presence of TGF-β (Hamby et al. 2006) and release glutamate (instead of uptaking glutamate) when exposed to elevated TNF-α levels (Bezzi et al. 2001).  Notably, both TGF-β and TNF-α levels are elevated after SCI (Argaw et al. 2009).  CSPGs and astrocytic scar regions are also thought to potently inhibit axonal regeneration (Popovich et al. 1997; Fawcett & Asher 1999; Silver & Miller 2004; Bradbury et al. 2002).  However, a recent study proposed that reactive astrocytes in the glial scar aid axonal regeneration (Anderson et al. 2016).  Clearly, the influence of astrogliosis may be more complicated than previously appreciated.  20  1.4.3 Oligodendrocytes 1.4.3.1 Oligodendrocyte death  Extensive oligodendrocyte death occurs following SCI (~50% at 1 dpi; ~93% at 1 wpi), due to both necrosis (e.g. lesion epicentre, adjacent tissue regions) and apoptosis (e.g. spared tissue rim, distances from lesion) (McTigue et al. 2001; Crowe et al. 1997; Casha et al. 2001; Almad et al. 2011).  Furthermore, oligodendrocyte apoptosis is evident in white matter tracts for 3-6 wpi in rodent and non-human primate contusion models (Crowe et al. 1997; Shuman et al. 1997), and at least 60 dpi in human SCI (Emery et al. 1998).  Oligodendrocytes are particularly vulnerable to oxidative damage due to several factors: (i) high metabolic rate (required for myelin sheath membrane synthesis), (ii) elevated intracellular iron stores (serving as a co-factor for myelinogenic enzymes), and (iii) low levels of anti-oxidant proteins (e.g. glutathione) (Matute et al. 2007; Almad et al. 2011).  Furthermore, oligodendrocytes are highly sensitive to elevated extracellular ATP and pro-inflammatory cytokines (e.g. TNF-α, IL-1β) (Steelman & Li 2011).  Oligodendrocyte apoptosis contributes to the extensive demyelination of spared axons following SCI, although primary and secondary injury processes can also directly damage myelin sheaths (Crowe et al. 1997; Shuman et al. 1997; Casha et al. 2001).  As individual oligodendrocytes can myelinate 30-80 different axons (Chong et al. 2012), significant loss manifests as widespread demyelination.  Many features of the injured spinal cord may damage oligodendrocytes.  For example, widespread necrotic cell death releases a host of proteolytic enzymes that can damage cells (Juliet et al. 2009).  Blood components have been shown to induce OPC apoptosis in vitro (Juliet et al. 2009; Almad et al. 2011).  Furthermore, even low concentrations of blood components inhibit OPC proliferation and migration (Juliet et al. 2009).  Tissue reperfusion is a major contributor to the formation of oxygen and nitrogen reactive species, which can damage cellular membranes, proteins, and DNA 21  (Almad et al. 2011).  Excitotoxicity stemming from elevated extracellular glutamate also contributes to oligodendrocyte loss following SCI (Almad et al. 2011; Plemel et al. 2014).  Oligodendrocytes express several glutamate receptors (i.e. AMPA, kainate, NMDA) rendering them vulnerable to extracellular glutamate elevation (Káradóttir et al. 2005).  Indeed, exogenous glutamate administration (at comparable levels as the injured spinal cord) induces oligodendrocyte death in the intact rat spinal cord (McAdoo et al. 1999; Xu et al. 2004).  Interestingly, AMPA receptor stimulation promotes OPC migration, proposed to be a homeostatic mechanism to preferentially recruit OPCs to the regions of most severe pathology where oligodendrocyte losses are most likely heaviest (Gudz et al. 2006).  Furthermore, excitotoxicity can be induced by ATP-mediated stimulation of oligodendrocyte P2X7 receptors (Wang et al. 2004; Matute et al. 2007).  Extracellular ATP levels are elevated by 2 hrs post-injury (Wang et al. 2004; Matute et al. 2007).  Furthermore, excitotoxic insult to oligodendrocytes can be exacerbated by pro-inflammatory cytokines (i.e. TNF-α, IL-1β), which are elevated within minutes of injury (Takahashi et al. 2003).  Another contributor to oligodendrocyte loss after SCI are compounds released by infiltrating neutrophils and activated microglia, including free radicals, pro-inflammatory cytokines, glutamate, and proteolytic enzymes (Donnelly & Popovich 2008).  Furthermore, lymphocytes can activate death receptors on oligodendrocytes via the release of various ligands (i.e. TNF-α, IL-2, & IFN-γ), resulting in apoptosis (Antel et al. 1994; Almad et al. 2011).  Activation of the death receptors Fas and p75 on oligodendrocytes can also induce oligodendrocyte death (D’Souza et al. 1996; Demjen et al. 2004; Casaccia-Bonnefil et al. 1996).  In addition to necrosis and apoptosis, autophagic cell death contributes to oligodendrocyte loss following SCI (Kanno et al. 2009; Kanno et al. 2011).  Oligodendrocyte loss is observed several segments away from the lesion site following rodent SCI, localized to degenerating axonal tracts (Warden et al. 2001; Sun et al. 2010; 22  Almad et al. 2011).  In contrast, oligodendrocyte apoptosis was not observed along degenerating axonal tracts following dorsal rhizotomy, suggested to be due to a lack of oxidative stress (Sun et al. 2010; Almad et al. 2011).  Therefore, the protracted oligodendrocyte apoptosis following SCI might be due to a combination of axonal degeneration and secondary injury processes (e.g. oxidative stress) initiated by the primary mechanical impact.    1.4.3.2   Oligodendrogenesis  Committed oligodendrocytes are considered post-mitotic (Keirstead & Blakemore 1997), therefore restoration of pre-injury oligodendrocyte densities in the injured spinal cord is achieved by the generation of new oligodendrocytes (i.e. oligodendrogenesis) from endogenous precursors, the oligodendrocyte precursor cells (OPCs) (Nishiyama et al. 2009; Zai & Wrathall 2005; Barnabé-Heider et al. 2010).  Concentrated oligodendrogenesis in the lesion rim is evident following contusion SCI, resulting in increased densities above pre-injury levels by 2 wpi (Tripathi & McTigue 2007; Rabchevsky et al. 2007).  These newly-formed oligodendrocytes participate in the spontaneous remyelination response that occurs following SCI, forming new myelin sheaths on denuded axons in adjacent tissue regions (see below) (Lasiene et al. 2008; Powers et al. 2013; Plemel et al. 2014).  Oligodendrogenesis following SCI has been demonstrated in spared tissue close to the meningeal border as well as 2-3 mm away from the lesion epicentre (Zai & Wrathall 2005; Rabchevsky et al. 2007; Tripathi & McTigue 2007).  Furthermore, oligodendrogenesis was observed to continue in the injured murine spinal cord for up to 9-12 wpi (Hesp et al. 2015), revealing the robustness of this response.  Oligodendrogenesis is also noted in primate models of SCI (Yang et al. 2006), suggesting commonality of this response.      23  1.4.3.3 Remyelination  Similar to the large scale replacement of glial cells, there is robust spontaneous remyelination following SCI (starting ~2-3 wpi in murine models) that replaces myelin sheaths lost to injury processes (Bunge et al. 1962; Harrison & McDonald 1977; Totoiu & Keirstead 2005; Lasiene et al. 2008; Powers et al. 2012).  Despite characteristically thinner myelin sheaths (as revealed by g-ratios under EM imaging) (Bunge et al. 1962), remyelination is associated with restoration of action potential conduction and properties (James et al. 2011).  Interestingly, the newly formed sheaths subsequently thicken, approaching g-ratios of developmentally-formed myelin by 6 mpi (Powers et al. 2013).  Despite robust remyelination, the persistence of potentially demyelinated axonal profiles in animal models, as well as human tissue, has lead to the contested hypothesis that endogenous remyelination is not complete following SCI (Guest et al. 2005; Totoiu & Keirstead 2005; Plemel et al. 2014).  However, many of these studies suffer from the inability to distinguish between intact demyelinated axons and severed demyelinated axonal segments (Plemel et al. 2014).  More recently it was demonstrated that all intact axons are remyelinated by 12 wpi in murine SCI models (Lasiene et al. 2008; Powers et al. 2012), therefore questioning whether promotion of remyelination is a viable therapeutic target for SCI (Plemel et al. 2014).  As mentioned previously, committed oligodendrocytes are post-mitotic (Keirstead & Blakemore 1997), therefore remyelination is carried out by newly-formed oligodendrocytes produced from OPCs (Nishiyama et al. 2009; Zai & Wrathall 2005; Barnabé-Heider et al. 2010).    24  1.4.4 Neurons 1.4.4.1 Neuronal death Similar to the loss of glial cells, neuronal loss following SCI is attributed to necrosis, apoptosis, and autophagic cell death, due to primary and secondary injury processes (Crowe et al. 1997; Yong et al. 1998; Kanno et al. 2009; Kanno et al. 2011; Chen et al. 2012).  However, in contrast to the robust gliogenesis that restores pre-injury glial densities following injury, lost neurons are not replaced (as the adult mammalian spinal cord is non-neurogenic) (Horky et al. 2006; Horner et al. 2000; Barnabé-Heider et al. 2010).    1.4.4.2 Axonal transection and regenerative failure In addition to neuronal death, the primary injury physically disrupts axons traversing the injury epicentre, whereas secondary injury processes damage spared axons in the adjacent tissue regions (Tator 1995; Blight 2002).  Severed axons retract from the lesion site (Kerschensteiner et al. 2005), induced by infiltrating ED1+ macrophages (Busch et al. 2010), and become stabilized in the parenchyma through the formation of synaptic-like interactions with NG2+ glial cells (Filous et al. 2014).  These proximal axonal segments form dystrophic end-bulbs, typified by disorganized microtubule arrays, thought to represent abortive regenerative attempts (Ertürk et al. 2007; Witte & Bradke 2008).  Failure of CNS regeneration is contrasted with successful, robust, and often complete regeneration in the injured PNS (Lutz & Barres 2014; Silver et al. 2015).  CNS regeneration failure is generally attributed to two main factors: (i) inhibitory post-injury environment and (ii) reduced growth capacity of adult mammalian CNS neurons (Silver et al. 2015).  Various features of the post-injury environment are thought to contribute to regeneration failure, including lack of permissive substrate spanning the lesion (McDonald et al. 1999), 25  inhibitory glial scar components (i.e. CSPGs, fibroblasts) (Silver & Miller 2004; Silver et al. 2015; Busch & Silver 2007), persistence of inhibitory myelin debris (Cafferty et al. 2010; Liu et al. 2006), insufficient trophic support for axonal growth (Kobayashi et al. 1997; Hains et al. 2003; Zhou & Snider 2006), and the absence of growth factors and guidance molecules to guide axons to appropriate targets (Alto et al. 2009; Blesch & Tuszynski 2009).  Furthermore, atrophic neurons whose axons are severed display attenuated responsiveness to growth factor treatments aimed at stimulating regenerative growth/sprouting (Kobayashi et al. 1997; Kwon et al. 2002), limiting therapeutic options (Kwon et al. 2004).    1.4.4.3 Axonal plasticity  The absence of ‘true’ regeneration (i.e. growth from transected axons) is contrasted with extensive axonal plasticity in models of incomplete SCI (Steward et al. 2008; Raineteau et al. 2002; Bareyre et al. 2004; Rosenzweig et al. 2010).  This intra-spinal plasticity is responsible, at least in part, for the spontaneous recovery following rodent and human SCI, attributed to by-pass circuit formation that restores supra-spinal input to circuitry below the lesion site (e.g. lumbar CPGs) (Bareyre et al. 2004; Rosenzweig et al. 2010).  Moreover, substantial cortical rearrangements following SCI (Bareyre et al. 2004; Raineteau & Schwab 2001) and myelin plasticity (i.e. de novo myelination, altered sheath thickness/length) (Fields 2015; Pajevic et al. 2014; Chang et al. 2016) might also be important facets of this spontaneous recovery.    26  1.4.5 Oligodendrocyte precursor cells (OPCs) 1.4.5.1 Proliferative response  Following SCI, OPCs proliferate robustly, attributed to injury-induced elevations in several OPC mitogens (Horky et al. 2006; Lytle et al. 2009; McTigue et al. 2001; Zai & Wrathall 2005).  For example, bFGF levels rise rapidly after SCI (Mocchetti et al. 1996; Tripathi & McTigue 2008) and bFGF induces OPC proliferation in vitro ( Barres et al. 1993; Bögler et al. 1990).  Furthermore, IGF-1 stimulates OPC proliferation (in combination with other factors) (Barres et al. 1993; McMorris & Dubois-Dalcq 1988).  Similarly, CNTF levels are elevated after SCI (Tripathi & McTigue 2008; Zai et al. 2005) and induce OPC proliferation in vitro and during development (Barres et al. 1996).  Proliferation of cultured OPCs is also stimulated by GGF (glial growth factor) (Canoll et al. 1996; Shi et al. 1998), which is also elevated in the injured spinal cord (Zai et al. 2005).  Exogenous administration of mitogens (e.g. GGF, FGF) following SCI increases OPC proliferation (McTigue et al. 1998), with corresponding increases in oligodendrogenesis and functional improvement (Lytle et al. 2009).  OPC proliferation following SCI is robust and rapid, beginning as early as 1 dpi, peaking 3-5 dpi, and continuing through 4 wpi, albeit at a reduced level (McTigue et al. 2001; Zai & Wrathall 2005; Horky et al. 2006; Tripathi & McTigue 2007).  Proliferation occurs close to the lesion site, concentrated in the penumbra, but also in spared white matter at later time-points (Hesp et al. 2015).  In addition to injury-induced elevations of mitogens, OPC proliferation following SCI may be stimulated by homeostatic mechanisms that control OPC density and distribution.  For example, OPC proliferation is reduced via contact inhibition as cell density increase in vitro (Zhang & Miller 1996).  Similar homeostatic mechanisms are thought to regulate OPC density in vivo, as has been demonstrated for cortical OPCs (Hughes et al. 2013).  27  Therefore, extensive death of OPCs at the lesion site following SCI would provide a strong stimulus for local proliferation to restore pre-injury densities.     1.4.5.2 Lineage plasticity In certain pathological conditions (e.g. SCI, MS, demyelination) Schwann cells myelinate central axons (Biernaskie et al. 2007; Bunge et al. 1993; Norenberg et al. 2004; Hampton et al. 2012; Zawadzka et al. 2010; Itoyama et al. 1983).  Genetic fate-mapping studies revealed that in chemical demyelination (Zawadzka et al. 2010) and contusion SCI (Assinck et al, in preparation; Bartus et al. 2016) the majority of these Schwann cells are derived from OPCs.  This fate-transition is not well characterized, but extracellular factors are likely to be involved.  Indeed, OPC production of Schwann cells may be suppressed by astrocytes, as transplanted OPCs following chemical demyelination form Schwann cells in astrocyte-free regions (Talbott et al. 2005; Talbott et al. 2006).  Furthermore, Schwann cell production from transplanted OPCs in the demyelinated CNS is inhibited by noggin administration (produced by astrocytes in vivo) (Talbott et al. 2006) and astrocytic knockout of STAT3 promotes Schwann cell remyelination (at the expense of oligodendroglial remyelination) in chemical demyelination models (Monteiro de Castro et al. 2015).  Schwann cells have a critical role in peripheral regeneration, which is robust and often complete (Lutz & Barres 2014).  Furthermore, Schwann cells support axonal growth in the injured spinal cord (Biernaskie et al. 2007; Fouad et al. 2005; Takami et al. 2002).  Therefore, promoting Schwann cell generation from OPCs may be beneficial following SCI.  There is also speculation that OPCs generate astrocytes under certain pathological conditions (Barnabé-Heider et al. 2010; Komitova et al. 2011; Zawadzka et al. 2010; Dimou et al. 2008; Kang et al. 2010; Simon et al. 2011), however it remains controversial. 28    1.4.6 Microglia Microgliosis (i.e. microglia activation) is characteristic of many CNS pathologies (e.g. traumatic injury, neurodegeneration, epilepsy, prion diseases, etc.), typically accompanied by microglial proliferation (Schwab et al. 2001; Ip et al. 2008; Ip et al. 2006; Ip et al. 2007; Schonrock et al. 1998; Imamoto et al. 1977).  Resident microglia perform immune surveillance to compensate for the occlusion of blood-borne immune cells from the CNS parenchyma under physiological conditions (David & Kroner 2011).  These ‘resting’ microglia possess a ramified morphology, characterized by thin, highly branched processes that continuously extend/retract (Davalos et al. 2005; David & Kroner 2011).  Upon CNS insult (e.g. stab wound, laser ablation), activated microglia adopt an amoeboid morphology with thickened and shorter processes (Ransohoff & Perry 2009) and rapidly reorient towards the lesion site (at a rate of 1.25 µm/min), forming a dense border within 30 min (Davalos et al. 2005), mediated by ATP and NO gradients (Dibaj et al. 2010).  This microglial respond is rapid.  For example, microglial expression of IL-1β mRNA is elevated <12 hrs after contusion SCI (Pineau & Lacroix 2007).  Furthermore, IL-1β protein levels are increased in microglia as early as 5 hours after human SCI (Yang et al. 2004).  Similarly, microglial TNF-α mRNA expression surges at ~1 hr post-injury in murine SCI (Pineau & Lacroix 2007).  Importantly, microglial activation following SCI depends upon primary injury mechanism (i.e. contusion, dislocation, distraction) (Choo et al. 2008; David & Kroner 2011), with contusion injury producing concentrated microgliosis at lesion epicenter and symmetrical activation in the dorsal and ventral columns (Choo et al. 2008).          29  1.5 Oligodendrocyte lineage cells 1.5.1 Developmental of the oligodendrocyte lineage Oligodendrocytes, the myelinating cells of the CNS, originate from a progenitor cell population called oligodendrocyte precursor cells (OPCs) that arise from ventricular germinal zones during development (Takebayashi & Ikenaka 2015; Richardson et al. 2006; Bergles & Richardson 2015).  In contrast to other developmental progenitors, OPCs persist into the adult CNS and maintain the capacity to generate myelinating oligodendrocytes (Nishiyama et al. 2009; Bergles & Richardson 2015).  Therefore, adult OPCs represent a reservoir that can be rapidly mobilized to replace myelin lost through injury or age-related processes (Hesp et al. 2015; Nishiyama et al. 2009), or in response to physiological demand for new myelination (e.g. experience or learning) (Fields 2015; Chang et al. 2016).  OPCs can be defined antigenically by expression of NG2 and PDGFRα (Pringle et al. 1992), as well as Olig2 and Sox10 (oligodendrocyte lineage markers) (Nishiyama et al. 2009; Bergles & Richardson 2015).  Upon differentiation, OPCs downregulate NG2 and PDGFRα expression (but maintain Olig2 and Sox10) (Richardson et al. 2006).  During CNS development, OPCs are first observed in the ventricular germinal zones (VZs) around mid-gestation (E12.5 in mice, E14 in rats, or ~E45 in humans) (Pringle & Richardson 1993; Timsit et al. 1995; Hajihosseini et al. 1996; Orentas & Miller 1996), probably generated by asymmetric division of radial stem/glia cells (i.e. one daughter maintains contact with ventricular and pial surfaces and remains a radial glial cell, whereas the other daughter losses contact and becomes an OPC) (Bergles & Richardson 2015), similar to the generation of migratory neuronal progenitors (Noctor et al. 2001).  Once generated, OPCs proliferate and migrate extensively, becoming evenly distributed throughout white and gray matter by ~E15 (Pringle & Richardson 1993).  In the spinal cord, OPCs are derived from the ventral pMN progenitor domain that previously generated motor 30  neurons (Richardson et al. 2006; Timsit et al. 1995; Takebayashi & Ikenaka 2015).  Following the initial wave of pMN-derived OPCs, a second wave of OPC production begins in the dorsal spinal cord at ~E15.5 (Cai et al. 2005; Fogarty et al. 2005; Vallstedt et al. 2005; Tripathi et al. 2011).  These dorsal OPCs are formed through trans-differentiation, as radial stem/glia cells retract their processes, lose contact with the central canal, and retract towards the pial surface (Fogarty et al. 2005).  Most of these cells become astrocytes, but a subset becomes the dorsally-derived OPC population.  Interestingly, radial glia cells maintain attachment to the pial surface, potentially related to the limited migration of dorsal OPCs (as compared to their ventral counterparts) (Tripathi et al. 2011).  Dorsally-derived OPCs remaining restricted to the dorsal half of the spinal cord, gradually replacing ventrally-derived OPCs in dorsal axonal tracts (Richardson et al. 2006; Tripathi et al. 2011).  In the mouse spinal cord, ~20% of all OPCs are dorsally-derived, whereas the other ~80% are ventrally-derived (Richardson et al. 2006; Tripathi et al. 2011; Takebayashi & Ikenaka 2015).  Generation of oligodendrocytes commences at ~E18.5 in mice, as OPCs differentiate into cells expressing myelin genes (i.e. MBP, PLP) and begin associating with axons in the ventral and dorsal funiculi (Tripathi et al. 2011; Richardson et al. 2006).  Oligodendrogenesis increases following birth, peaking ~2-4 weeks post-natal in mice, and continuing through 8 months post-natal (albeit at a decreased rate) (Rivers et al. 2008).  Development of the pMN domain is instructed by diffusible gradients of Shh (Sonic Hedgehog) (produced by the notochord and floor plate), BMPs, and Wnts (from the roof plate) (Orentas & Miller 1996; Poncet et al. 1996; Pringle et al. 1996; Mekki-Dauriac et al. 2002; Agius et al. 2004; Miller et al. 2004).  In the pMN domain, Shh induces Olig2 expression, required for both motor neuron and OPC production (Richardson et al. 2006; Takebayashi & Ikenaka 2015).  Indeed, the pMN domain fails to form in Olig2-/- mice, with no motor neuron or OPC production, leading to death at birth (Zhou & Anderson 2002; 31  Takebayashi et al. 2002; Lu et al. 2002).  During CNS development, Olig2 is phosphorylated at the serine-147 residue during MN production, which is rapidly dephosphorylated upon the switch to OPC production (Li et al. 2011).  Olig2 proteins phosphorylated at serine-147 form homodimers, but when dephosphorylated they form heterodimers with other bHLH transcription factors (Li et al. 2011).  Heterodimer formation alters Olig2 activity (e.g. DNA binding), thought to contribute to the MN-OPC fate switch, potentially through sequestration of pro-neural transcription factors (e.g. Ngn-2) (Li et al. 2011; Bergles & Richardson 2015).  In addition to Shh, Notch signaling is also involved in OPC specification.  For example, during zebrafish development Notch signaling between neuronal precursors and radial glial cells maintains the later population in a stem-like state, preserving them for subsequent OPC production (Park & Appel 2003; Kim et al. 2008).  Pharmacological inhibition of Notch signaling (via DAPT administration) leads to motor neuron overproduction at the expense of OPCs (Kim et al. 2008).  This model applies to mammals as well, as Jagged2 (a Notch ligand) has been demonstrated to be critical in the restraint of precocious OPC production (Rabadán et al. 2012).  Notch signaling is thought to regulate the expression and/or activity of the phosphatase that dephosphorylates Olig2 at serine-147, potentially mediated through Hes5 modulation (a known oligodendrocyte differentiation inhibitor) (Rabadán et al. 2012; Bergles & Richardson 2015).  In addition to initial specification, Notch signaling is also involved in oligodendrocyte differentiation and myelination (Wang et al. 1998; Hu et al. 2003).  Olig1 mRNA is present before E11 in the VZ, but is not translated until ~E18.5 (Fu et al. 2009), therefore does not contribute to OPC specification in the spinal cord (Bergles and Richardson 2015).  Moreover, some OPCs are produced in Olig2-/- mice, but none are produced in Olig1-/-/Olig2-/- mice (Lu et al. 2002; Zhou & Anderson 2002).  Despite a trivial role in OPC specification (Paes de Faria et al. 2014), Olig1 appears critical for remyelination in the adult CNS (Arnett et al. 32  2004; Emery & Lu 2015).  In contrast to the ventrally-derived OPCs, specification of the dorsally-derived OPCs is not well characterized, although it appears to be Shh-independent (Chandran et al. 2003; Kessaris et al. 2006).  Indeed, dorsal OPC production continues despite disrupted Shh signaling in Nkx6.1-/-/Nkx6.2-/- mice  (Cai et al. 2005; Vallstedt et al. 2005).  Furthermore, OPCs are generated in vitro from Shh-/- embryonic mouse spinal cord cultures (Cai et al. 2005).  Independent OPC populations, specified by different signals, suggests that intrinsic functional differences may exist based on developmental origin (highly relevant for demyelinating disease therapeutics) (Bergles & Richardson 2015).  Different morphological classes of oligodendrocytes have been described (Bjartmar et al. 1994; Butt et al. 1998; Anderson et al. 1999), as have subclasses based on molecular differences (Butt et al. 1998; Kleopa et al. 2004).  Furthermore, dorsal OPCs displace their ventral counterparts from dorsal axons (Tripathi et al. 2011) and display a greater propensity for remyelination (Zhu et al. 2011).  It remains unclear, however, whether this heterogeneity reflects developmental origin (Bergles & Richardson 2015).  Alternatively, as dorsal OPCs are generated later during development (i.e. are ‘younger’) they may have a greater capacity to compete for resources (e.g. growth factors, etc.) (Bergles & Richardson 2015).  Moreover, specific ablation of either OPC population during development (via diphtheria toxin) does not affect the adult OPC complement or phenotype, as the spared OPCs invade and re-populate the depleted regions (Kessaris et al. 2006).  The inability to distinguish dorsal and ventral OPCs by antigenic or electrophysiological properties complicates further investigation of intrinsic differences (Tripathi et al. 2011; Clarke et al. 2012), however genome-wide expression analyses (performed on purified cultures) should be able to address this quandary.    33  1.5.2 Extrinsic regulation of oligodendrocyte lineage development 1.5.2.1 PDGF-signaling PDGFRα-signaling regulates OPC proliferation and survival, mediated by PDGF-A secreted by neurons and astrocytes (Noble et al. 1988; Raff et al. 1988; Richardson et al. 1988; Hart et al. 1989; Pringle et al. 1992).  The mitogenic activity of PDGF-A requires co-participation of ECM molecules, including NG2 (Nishiyama et al. 1996), tenascin-C (Garcion et al. 2001), and αvβ3 integrin (Baron et al. 2002).  Interestingly, OPCs display regional heterogeneity in PDGF-A responsiveness.  For example, genetic deletion of PDGF-A differentially reduces the OPC population of the optic nerve (>99% reduction at birth), spinal cord (~88%), cerebral cortex (~80%), and brainstem (~60%) (Fruttiger et al. 1999).  However, compensation by PDGF-B and/or PDGF-C is feasible, as PDGF-C is expressed in the developing cortex (Hamada et al. 2002) and can activate PDGFRα-signaling, however the role of these factors in OPC development has not been thoroughly characterized (Bergles & Richardson 2015).  In vitro, removal of PDGF halts OPC proliferation and triggers differentiation (Durand & Raff 2000), accompanied by rapid downregulation of PDGFRα expression (seen as a consequence of differentiation, not the cause) (Hart et al. 1989).  Furthermore, reduced PDGF-signaling is implicated in the decline of OPC proliferation rate over CNS development.  Indeed, the prolongation of the OPC cell cycle (~24 hrs at E12.5 to >100 hr at E17 in the mouse spinal cord) can be partially reversed by PDGF supplementation in vitro (van Heyningen et al. 2001).  Moreover, astrocytic and neuronal PDGF-A overexpression stimulates OPC proliferation in both white and gray matter (Calver et al. 1998; Fruttiger et al. 1999; van Heyningen et al. 2001), and OPC density is reduced by ~50% in PDGF+/- mouse embryos (van Heyningen et al. 2001).     34  1.5.2.2 FGF-signaling FGF-signaling promotes OPC mitogenic activity, specifically in combination with PDGF.  For example, PDGF-A induces OPC proliferation and differentiation in vitro, but maintains a reservoir of OPCs (similar to the in vivo situation) (Raff et al. 1988; Tang et al. 2000).  By contrast, combination of PDGF-A and bFGF extends proliferation but inhibits differentiation (Bögler et al. 1990; McKinnon et al. 1990), significantly expanding the OPC population.  Interestingly, this effect is not recapitulated in vivo, as conditional deletion of FGF receptors has minimal effect on OPC proliferation in the murine CNS (Furusho et al. 2011).  In addition to OPC proliferation, FGF-signaling also regulates oligodendrocyte differentiation.  Indeed, committed oligodendrocyte numbers are increased in the spinal cord of bFGF-/- mice (Murtie et al. 2005).  Furthermore, Fgfr1/Fgfr2 knockout in differentiated oligodendrocytes (via CNP-Cre or Olig1-Cre) inhibits myelin protein synthesis and reduced myelin sheath thickness, correlated with attenuated MAP-kinase activity (Furusho et al. 2012).  Therefore, the effect of FGF signaling on OPCs in vivo is complex, and likely to be even more complicated in pathological situations (Ford-Perriss et al. 2001; Bergles & Richardson 2015).    1.5.2.3 EGF-signaling EGF-signaling also regulates oligodendrocyte development.  EGF administration drastically increases OPC number in the adult CNS (Cantarella et al. 2008).  Furthermore, EGF treatment promotes remyelination following gliotoxin-mediated demyelination or perinatal ischemic insult (Scafidi et al. 2014).  Instead of a direct effect on OPCs, EGF is thought to bias NSPCs towards gliogenesis (including OPC production) (Sun et al. 2005; Cantarella et al. 2008; Gonzalez-Perez & Alvarez-Buylla 2011).  Indeed, SVZ NSPCs highly express erbB (i.e. EGF receptor), whereas 35  OPCs do not (Chojnacki & Weiss 2004; Sun et al. 2005).  Moreover, EGF-induced OPC proliferation is not observed outside of the SVZ in slice culture studies (Hill et al. 2013).    1.5.2.4 Model for regulation of OPC proliferation A large number of other polypeptide factors have been implicated in oligodendrocyte lineage development, including members of the TGF-β/BMP family (Dutta et al. 2014; Palazuelos et al. 2014), IGF-1 (McMorris & Dubois-Dalcq 1988; D’Ercole & Ye 2008), neuregulins (Canoll et al. 1996; Vartanian et al. 1999; Fernandez et al. 2000; Carteron et al. 2006; Brinkmann et al. 2008; Ortega et al. 2012), Wnts (Shimizu et al. 2005; Fancy et al. 2009; Dai et al. 2014), neurotrophin-3 (Barres & Raff 1993; Cohen et al. 1996), chemokines (Robinson et al. 1998; Dziembowska et al. 2005; Göttle et al. 2010), CNTF (Barres et al. 1993), BDNF (Vondran et al. 2010; Wong et al. 2013), transferrin (Silvestroff et al. 2013), erythropoietin (Sugawa et al. 2002), thyroid hormone (TH; triiodothyronine) (Barres et al. 1994) , and retinoic acid (Barres et al. 1994).  Cultured OPCs treated with PDGF, but lacking TH will continuously proliferate without differentiation (Barres et al. 1994), whereas OPC proliferation and differentiation are concurrent when TH is present (Tang et al. 2000).  It has been proposed that OPCs possess an intrinsic timer that regulates proliferation and differentiation.  Indeed, individual OPCs cultured with TH and PDGF will divide a predictable number of times (younger OPCs divide more) before all of the progeny synchronously differentiate (Temple & Raff 1985).  However, clonal analysis of mixed cell cultures reveals that the progeny of a single OPC do not necessarily differentiate synchronously (Zhang & Miller 1996; Ibarrola et al. 1996), and clones of oligodendrocyte lineages often contain both myelinating oligodendrocytes and proliferating OPCs (Bergles & Richardson 2015).  It is feasible that this represents external modulation of an intrinsic timer, but regardless it is clear that the regulation of OPC dynamics is 36  complex in vivo.  By contrast, maintenance of stable OPC density is thought to be determined by a balance between the rate of PDGF production (from astrocytes and neurons) and consumption by OPCs (van Heyningen et al. 2001; Calver et al. 1998).  This ‘supply and demand’ model is consistent with declining OPC proliferation rates during CNS development, but is inconsistent with the continual decline through post-natal life despite constant OPC numbers (Rivers et al. 2008; Young et al. 2013).  For example, OPC cell cycles in the corpus callosum increase from ~3 days at P21, to ~10 days at P60, to >100 days at P240 (Psachoulia et al. 2009; Young et al. 2013).  The ‘supply and demand’ model predicts that a reduction of PDGF should increase OPC density, but not affect proliferation rate.  Therefore, a more complete model of OPC regulation would include: (i) decreased mitogenic responsiveness to PDGF, (ii) reduced proliferation, (iii) an intrinsic timer that controls transition between cell cycle stages (instead of differentiation), (iv) steady accumulation of cell cycle inhibitors (e.g. p27Kip1/p57Kip2) (Durand & Raff 2000; Dugas et al. 2007), and (v) reduced activity in key pro-differentiation pathways (Tokumoto et al. 2001; Billon et al. 2004; Bergles & Richardson 2015).    1.5.2.5 OPC migration Regulation of OPC migration is not well characterized, but is likely similar to neuronal guidance, including netrin-1/DCC/Unc5, semaphorin/neuropilin, and ephrin/eph signaling interactions (de Castro & Bribián 2005).  Indeed, OPCs express DCC/Unc5 and are repelled by netrin-1 in vitro (Tsai et al. 2003).  Furthermore, dorsoventral OPC migration is impaired in the spinal cord of netrin-1-/- mice (Jarjour et al. 2003).  OPCs also express neuropilin-1/2, are repelled by Sema3A, but attracted by Sema3F in vitro (Sugimoto et al. 2001; Spassky et al. 2002; Syed et al. 2011).  Interestingly, individual knockouts of these factors do not induce persistent OPC migratory 37  deficits, suggesting significant functional redundancy (Bergles & Richardson 2015).  OPC migration is also influenced by growth factors.  For example, OPC dispersion from germinal zones is reduced in PDGFRα-/- mice, leading to myelination deficits (Fruttiger et al. 1999).  This effect is thought to be indirect (i.e. PDGF stimulates OPC proliferation, increasing local density, forcing self-repulsion, leading to dispersion) (Bergles & Richardson 2015).  Interestingly, OPCs migrate along blood vessels during CNS development (Tsai et al. 2016).  Similar movement along vasculature may be involved in OPC recruitment to lesions in the adult CNS (i.e. SCI, demyelination) (Tsai et al. 2016).   1.5.3 Intrinsic regulation of oligodendrocyte lineage development 1.5.3.1 Specification OPC specification from neuroepithelial cells is largely accomplished by opposing gradients of Shh (ventral) and BMPs (dorsal).  In the spinal cord, OPCs arise from the pMN domain, which produces motor neurons (MN) prior to a neuron-to-glia switch (Zhou & Anderson 2002; Lu et al. 2002; Novitch et al. 2001; Zhou et al. 2001).  In addition to Shh-induced Olig2 (Valstedt et al. 2005; Kessaris et al. 2006), the transcription factors Nkx6-1 (Liu et al. 2003) and Gli2 (Qi et al. 2003) define the pMN borders and influence OPC specification.  Indeed, genetic deletion of Nkx6-1 and Gli2 impairs OPC specification and reduces pMN domain size (Liu et al. 2003; Qi et al. 2003).  Several other transcription factors are also involved in OPC specification.  For example, knockout of Ascl1 (a.k.a. Mash1) substantially reduces OPC numbers (Parras et al. 2007; Sugimori et al. 2008).  Ascl1 overexpression promotes OPC specification at the expense of MNs early in development (Petryniak et al. 2007) and astrocytes at later developmental stages (Nakatani et al. 2013).  The gliogenic transcription factor Sox9 is also involved in OPC specification.  Indeed, 38  Sox9 conditional knockout substantially delays developmental OPC production (Stolt et al. 2003).  Sox9 induces NFIA (pro-glial/anti-neuronal transcription factor) expression (Kang et al. 2012), knockout of which impairs OPC specification (Deneen et al. 2006).  OPCs express a host of transcription factors, including Olig2, Olig1, Nkx2-2, and Sox10 (Emery & Lu 2015).  Olig2 directly regulates Nkx2-2 and Sox10 expression.  For example, electroporation of Olig2 expression vectors into the developing chick spinal cord induces Nkx2-2 (Liu et al. 2007) and Sox10 expression (Zhou et al. 2001; Liu et al. 2007).  Furthermore, Olig2 directly targets Sox10 enhancer sequences (Yu et al. 2013; Küspert et al. 2011).  While not required for OPC maintenance, Nkx2-2 is involved in differentiation (Qi et al. 2001).  Indeed, Nkx2-2 protein levels transiently increase during differentiation (Fu et al. 2002).  Furthermore, Nkx2-2-/- mice display impaired oligodendrocyte differentiation (Qi et al. 2001).  Similarly, Sox10 deletion minimally effects developmental OPC production (Stolt et al. 2002), however oligodendrocyte lineage-specific Sox10/Sox9 co-deletion substantially reduces OPC number in the developing spinal cord (Finzsch et al. 2008), suggesting an important, although partially redundant role for Sox10 in OPC specification (Emery & Lu 2015).  Furthermore, Sox9/Sox10 promote PDGFRα expression in OPCs (important for survival and proliferation), a function antagonized by Sox5/Sox6 activity (Stolt et al. 2006).  Indeed, Sox9/Sox10 co-deletion significantly reduces PDGFRα expression in OPCs (Finzsch et al. 2008).   1.5.3.2 Differentiation Oligodendrocyte differentiation in vivo is considered terminal, despite demonstration of bFGF-induced oligodendrocyte de-differentiation in vitro (Grinspan et al. 1996).  Therefore, tight regulation is required to ensure proper developmental myelination and to maintain a sufficient 39  post-natal OPC reservoir (Emery & Lu 2015).  Various extracellular factors inhibit differentiation, mediated by the transcription factors Sox5, Sox6, Hes5, Id2, and Id4, which are highly expressed in OPCs (Emery & Lu 2015).  For example, Id2/4 function downstream of BMP- (Samanta & Kessler 2004) and Gpr17-signaling (Chen et al. 2009), both known differentiation inhibitors.  Furthermore, Id2 overexpression inhibits differentiation, whereas Id2 knockout promotes differentiation in vitro (even in proliferative conditions) (Wang et al. 2001).  Id2/4 are thought to physically interact with and block nuclear Olig1/2 activity (Samanta & Kessler 2004).  Upon differentiation induction, Id2/4 mRNA levels are robustly decreased, accompanied by a nuclear-to-cytoplasmic shift of Id2 protein (Wang et al. 2001).  Notch-signaling also inhibits oligodendrocyte differentiation (Louvi & Artavanis-Tsakonas 2006).  Axonal expression of Jagged2 (a notch ligand) is substantially reduced in temporally coincidence with myelination onset (Wang et al. 1998).  Furthermore, Delta and Notch (i.e. notch ligands) addition significantly reduces differentiation in vitro (Wang et al. 1998).  Moreover, Notch-1+/- mice display precocious CNS myelination (Givogri et al. 2002).  The inhibitory effect of Notch-signaling is mediated by Hes5, which prevents MBP expression through physical sequestration of Sox10 (Liu et al. 2006).  Indeed, Hes5 deletion accelerates myelin protein expression (e.g. MBP, PLP), while Hes5 over-expression inhibits differentiation in vitro (Liu et al. 2006).  Wnt-signaling also inhibits differentiation, both developmentally and in pathological conditions (Fancy et al. 2009; Ye et al. 2009).  Expression of Tcf712 (complexes with β-catenin) is restricted to the late-OPC/pre-myelinating oligodendrocyte transition (Fancy et al. 2009; Fu et al. 2009; Ye et al. 2009).  The Tcf712/β-catenin complex directly targets Id2, potently inhibiting differentiation (Ye et al. 2009).  However, the role of the Tcf712/β-catenin complex appears more complex, as oligodendrocyte differentiation is impaired in Tcf712-/- mice (Fu et al. 2009; Ye et al. 2009), and inactivation of β-40  catenin specifically in the oligodendrocyte lineage delays differentiation (Dai et al. 2014).  In addition to β-catenin, Tcf712 interacts with HDAC1/2.  As HDAC1/2 represses differentiation inhibiting genes (Ye et al. 2009), compromised function may explain these findings (Bergles & Richardson 2015).  Therefore, Wnt-signaling may exert stage-specific effects (i.e. inhibit specification, partially promote aspects of differentiation, but downregulation is required for differentiation completion) (Dai et al. 2014).  In addition to Id2/4, Sox5/6 also inhibit differentiation.  Sox5/6 expression is downregulated upon differentiation (Stolt et al. 2006), mediated (at least in part) by several microRNAs that target OPC specific genes (Dugas et al. 2010; Zhao et al. 2010).  Oligodendrocyte lineage-specific Sox5/6 co-deletion induces precocious myelin protein expression (e.g. MBP, PLP) during spinal cord development (Stolt et al. 2006), suggesting accelerated differentiation.  Sox5/6 antagonize Sox10 activity (similar to Hes5) by blocking access to promoter elements of Sox10-targeted genes (e.g. MBP) (Stolt et al. 2006).  In contrast, several pro-differentiation factors have been described.  In addition to specification, Olig2 is also a pro-differentiation factor.  Indeed, Olig2 deletion from late-stage OPCs significantly reduces mature oligodendrocyte number (Cai et al. 2007; Mei et al. 2013).  Olig2 complexed with Brg1 (a chromatin remodeling enzyme) promotes differentiation by acting on oligodendrocyte gene enhancers (Yu et al. 2013).  Another pro-differentiation factor is the zinc finger homeobox transcription factor Zfhx1b (a.k.a. Sip1), which is directly targeted by Olig2 and is downstream of Olig1 (Weng et al. 2012).  Zfhx1b conditional knockout results in CNS myelination failure without reducing OPCs (Weng et al. 2012).  Furthermore, Zfhx1b overexpression promotes differentiation in vitro (Weng et al. 2012).  Zfhx1b antagonizes p300/SMAD complex activity at Id2, Id4, and Hes5 gene promoter sequences (downstream of BMP-, Notch-, and Wnt-signaling) and induces Smad7 expression (a negative regulator of BMP-signaling) (Weng et al. 2012).  The Wnt-signaling 41  inhibitor Sox17 is another pro-differentiation factor, relieving the inhibition of the Wnt/β-catenin/Tcf712 complex through promotion of β-catenin degradation (Sohn et al. 2006; Chew et al. 2011).   1.5.3.3 Maturation/myelination Newly differentiated oligodendrocytes undergo dramatic morphological changes, cease migration, and extend networks of processes during maturation to myelinating oligodendrocytes.  This involves upregulation of numerous genes, including myelin proteins (e.g. MBP, PLP), cytoskeletal proteins, lipid metabolism, and axo-glial junction components (Dugas et al. 2006; Cahoy et al. 2008).  Oligodendrocyte development is halted at the differentiation or early myelination stages by deletion of several transcription factors, including Nkx2-2, Olig1, Ascl1, YY1, Zfhx1b, Sox10, and MyRF (Qi et al. 2001; Stolt et al. 2002; Xin et al. 2005; He et al. 2007; Sugimori et al. 2008; Emery et al. 2009; Weng et al. 2012), which appear to act non-redundantly.  Despite early expression in development, Olig1 mainly affects post-mitotic oligodendrocytes.  Indeed, Olig1-/- mice display delayed expression or loss of mature oligodendrocyte markers (e.g. MBP) (Lu et al. 2002).  Luciferase assays revealed that Olig1 directly promotes MBP expression (Xin et al. 2005; Li et al. 2007) and exogenous Olig1 is sufficient to drive MBP expression (Niu et al. 2012).  Paradoxically, Olig1 is cytoplasmic during myelination (shifting from the nucleus), seemingly incompatible with the promotion of myelin gene expression (Arnett et al. 2004).  Nevertheless, cytoplasmic shifting of Olig1 proteins is required for oligodendrocyte membrane expansion and myelination (Niu et al. 2012).  Nkx2-2 deletion significantly reduces developmental MBP and PLP expression (Qi et al. 2001).  Furthermore, Nkx2-2 promotes GFP expression from the PLP promoter in heterologous cell expression systems (Qi et al. 2001).  However, Nkx2-2 also 42  suppresses MBP (Gokhan et al. 2005; Wei et al. 2005) and Sirt2 promoter activity (Ji et al. 2011), suggesting a complex role.  Unlike Olig1, Sox10, and Nkx2-2, MyRF (myelin regulatory factor, a.k.a. GM98, C11Orf9, or MRF) is not expressed by OPCs, but is robustly induced during early differentiation (Emery et al. 2009; Emery & Lu 2015).  Both Sox8/10 (Hornig et al. 2013) and the Olig2/Brg1 chromatin remodeling complex (Yu et al. 2013) directly target a first-intron promoter sequence in the MyRF gene, meaning it is a common target of both Sox10 and Olig2.  Conditional knockout of MyRF stalls differentiation at pre-myelinating oligodendrocytes, culminating in CNS myelination failure (Emery et al. 2009), similar to Olig1-/- mice (Xin et al. 2005) and Sox10-/- zebrafish (Takada et al. 2010), suggesting that all 3 factors are essential for myelination.  MyRF is also required for adult myelin maintenance.  Indeed, MyRF deletion from mature oligodendrocytes reduces myelin gene expression and leads to sheath degeneration (Koenning et al. 2012).  In contrast, Olig2 deletion from mature oligodendrocytes increases both Olig1 and myelin gene expression (Mei et al. 2013).  As predicted, MyRF targets (as revealed by ChIP-Seq) include genes important in maturation/myelination, such as cytoskeletal proteins, lipid metabolic enzymes, transcription factors (e.g. Smad7, Nkx6-2), and myelin proteins (e.g. MBP, PLP) (Bujalka et al. 2013).  Significantly, there is substantial overlap between Olig2, MyRF, and Sox10 targets, suggesting a close functional relationship (Bujalka et al. 2013; Yu et al. 2013).  Indeed, luciferase assays revealed synergistic action of MyRF and Sox10 at a subset of myelin gene regulatory regions (Hornig et al. 2013).  It has been proposed that transient action of the Olig2/Brg1 chromatin remodeling complex makes key gene regulatory regions accessible for subsequent action by Sox10 and MyRF during myelination (Emery & Lu 2015).  43  1.5.4 Oligodendrocyte precursor cells (OPCs) in the adult CNS In contrast to other progenitor cells, OPCs persist into the adult CNS, where they comprise 5-8% of all neural cells (regionally variable), are uniformly distributed across both white and gray matter, and are the primary proliferative cell type (Pringle et al. 1992; Chang et al. 2000; Dawson et al. 2003; Nishiyama et al. 2009).  Despite this, OPC number and density is maintained throughout post-natal life (Rivers et al. 2008), suggesting that excess progeny either differentiates or dies .  1.5.4.1 Characteristics Adult OPCs retain several progenitor characteristics (e.g. proliferation, migration), critical for effective reparative responses following demyelination, but may predispose OPCs to uncontrolled growth (Bergles & Richardson 2015).  Certain human gliomas (i.e. glial cell tumors) express oligodendrocyte lineage genes and display OPC-like migratory behaviour (Robinson et al. 1998; Dougherty et al. 2012; Tsai et al. 2016), suggesting a potential OPC origin.  Indeed, OPC specific co-deletion of p53 and NF-1 (tumor-suppressors) or overexpression of constituently active erbB induces tumorigenesis (Persson et al. 2010).  Moreover, over-stimulation of PDGF-signaling leads to formation of gliomas containing OPC-like cells (Assanah et al. 2006; Lindberg et al. 2009; Persson et al. 2010; Nazarenko et al. 2011).  OPCs express receptors for neurotransmitters, including glutamate (AMPA, NMDA), GABA, and acetylcholine (Barres et al. 1990; Wyllie et al. 1991), confirmed by slice culture (Steinhäser et al. 1994) and gene expression profiling (Seifert et al. 1997; Cahoy et al. 2008).  In addition, OPCs express a variety of voltage-gated ion channels, including those for Na+, K+, and Ca2+ (Bergles & Richardson 2015).  Early in development (when membrane resistance is high), OPCs display Na+-dependent spikes (Chittajallu et al. 2004; Biase et al. 2011). 44   1.5.4.2 Axo-glial synapses OPCs form direct synaptic connections with neurons, making them unique among glial cells (Bergles et al. 2000; Jabs et al. 2005).  OPC-glutamatergic neuron synapses are observed in all CNS regions examined, including the hippocampus (Bergles et al. 2000; Jabs et al. 2005), brainstem (Müller et al. 2009), cortex (Chittajallu et al. 2004), cerebellum (Lin et al. 2005), and white matter tracts (Káradóttir et al. 2005; Kukley et al. 2007).  Similarly, OPC-GABAergic neuron synapses have also been reported (Lin & Bergles 2004; Jabs et al. 2005; Mangin et al. 2008; Tanaka et al. 2009; Vélez-Fort et al. 2010).  The highly conserved nature of these axo-glial synapses suggests a functional importance.  OPC-glutamatergic neuron synapses are formed early during development and undergo maturation in parallel with neuronal synapses (i.e. increased currents, more inputs, etc.) (Bergles & Richardson 2015).  In contrast, cortical OPC-GABAergic neuron synapses transition from synaptic to extra-synaptic during development (Vélez-Fort et al. 2010).  Unlike neurons, OPCs appear to only be post-synaptic, suggesting one-way electrical communication (Bergles & Richardson 2015).  However, axoglial synapses may mediate retrograde non-electrical signaling (e.g. growth factors [e.g. BDNF] or neuromodulators [e.g. endocannabinoids]), which can modulate various neuronal properties (e.g. axon excitability, conduction velocity, myelination, etc.) (Bergles & Richardson 2015).  Both morphological and electrophysiological evidence supports OPC-neuron synaptic functionality.  Indeed, these axo-glial connections display structural hallmarks of functional synapses under EM imaging (i.e. rigidly aligned membranes, electron-dense regions, clusters of pre-synaptic vesicles, etc.) (Bergles et al. 2000; Kukley et al. 2007; Haberlandt et al. 2011).  Furthermore, stimulation of axons induces currents in local OPCs (with similar properties as neuronal EPSP-induced currents), abolished by 45  AMPA receptor antagonism (e.g. cyclothiazide treatment) (Lin & Bergles 2004; Tanaka et al. 2009).  Similar to immature neurons, GABA receptor activation induces OPC depolarization, as intracellular [Cl-] is elevated (Lin & Bergles 2004; Tanaka et al. 2009).  The presence of synapses on highly proliferative cells seems contradictory.  Indeed, these structures are likely transient (De Paola et al. 2006), as the highly dynamic nature of OPCs dictates a need for continual formation and dissolution (Hughes et al. 2013).  Surprisingly, OPC-neuron synapses are maintained during cell division, accomplished via a fission-like process by which each daughter cell receives half of the synaptic connections (Hughes et al. 2013).    Axo-glial synapses are restricted to the OPC stage of oligodendrocyte lineage development.  Upon induction of differentiation, AMPA and NMDA receptor expression is rapidly decreased, followed by dismantling of synaptic structures (Kukely et al. 2010).  Despite their prevalence, the function of these synapses remains unclear.  OPCs are not considered excitable cells, and do not possess an analog of a neuronal axon for signal propagation.  Furthermore, adult OPCs are relatively hyperpolarized (VM ~-95 mV, neuronal VM ~-70 mV), meaning that single synaptic events elicit minimal depolarization (Bergles & Richardson 2015).  However, OPC expression of GluR2-containing AMPA receptors (Lin et al. 2005) (i.e. increased Ca2+ permeability) (Donevan & Rogawski 1995) and NMDA receptors with reduced Mg2+ sensitivity, enables robust Ca2+ influx at more negative VM values (Káradóttir et al. 2005), which could be a signaling mechanism.  AMPA receptor stimulation reduces OPC proliferation, whereas antagonism reduces differentiation in vitro (Yuan et al. 1998; Gudz et al. 2006).  However, as these studies were performed in slice culture or dissociated cell culture a direct effect on neuronal AMPA receptors cannot be excluded (Bergles & Richardson 2015).  Oligodendrocyte lineage-specific GluN1 subunit deletion (required for NMDA receptor function), does not affect OPC proliferation, morphology, distribution, or differentiation ability (Biase et al. 46  2011; Guo et al. 2012).  However, AMPA receptor Ca2+ permeability is slightly increased in OPCs lacking functional NMDA-signaling (Biase et al. 2011).  Despite the energetic costs of dissolution/reformation of synaptic structures to enable continuous OPC movement, axo-glial synaptic signaling conveys key benefits, including: (i) highly specific signaling (unlike neuromodulatory transmitters) and (ii) surveillance of local neural activity (enabling rapid response to a physiological demand for new myelination) (Bergles & Richardson 2015).  Indeed, tetrodotoxin (TTX) administration inhibits developmental myelination (Barres & Raff 1993; Demerens et al. 1996), whereas artificial and/or physiological stimulation promotes OPC proliferation and differentiation in the adult CNS (Simon et al. 2011; Gibson et al. 2014; McKenzie et al. 2014).  Moreover, neural activity induces myelin protein synthesis in associated oligodendrocytes, thus preferentially thickening myelin sheaths on active axons (Wake et al. 2011; Gibson et al. 2014).  It remains to be demonstrated if this activity-dependent regulation is mediated by OPC-neuron synapses, but speculation would predict a central role.   1.5.4.3 Lineage potential OPCs were initially proposed to be multipotent, however genetic fate-mapping revealed progeny restriction to the oligodendrocyte lineage under physiological conditions (Dimou et al. 2008; Rivers et al. 2008; Kang et al. 2010; Zhu et al. 2011).  However, previous studies have reported OPC-derived astrocytes and Schwann cells in certain pathologies (Dimou et al. 2008; Barnabé-Heider et al. 2010; Zawadzka et al. 2010; Komitova et al. 2011) (fig. 1.1), although the majority of progeny are oligodendrocytes.  A notable exception is exclusive OPC astrogenesis following freeze-thaw cortical injury (Tatsumi et al. 2008), although this may be due to cold-induced Olig2 inactivation, as inactivation of Olig2 in OPCs promotes OPC astrogenesis (Zhu et al. 2012).  47  Current consensus is that OPCs do not produce neurons, even under pathological conditions, as previous reports are not considered convincing (Bergles & Richardson 2015).  For example, OPCs were proposed to form piriform cortex neurons (Rivers et al. 2008; Guo et al. 2010), but the labeled cells did not increase over time (which you would expect with continual production) and did not incorporate thymidine analogs (which they would if they were derived from proliferative OPCs) (Kang et al. 2010; Clarke et al. 2012).                         Figure 1.1 Lineage potential of OPCs.  Summary of cell types reported to be generated from oligodendrocyte precursor cells (OPCs) in the heathly (green) and pathological CNS (red).  Adpated from Dimou & Gallo (2015).  1.5.4.4 Injury response Maintenance of the adult OPC population relies predominately on local self-renewal, although new OPCs can be generated from SVZ progenitors in some pathologies (e.g. demyelination of 48  adjacent regions) (Jablonska et al. 2010).  If administered for a sufficient period, 100% of adult OPCs will incorporate thymidine analogs (Kang et al. 2010; Young et al. 2013), suggesting that all adult OPCs retain proliferative capacity.  Focal loss of an individual OPC (e.g. through differentiation, apoptosis, or laser ablation) is compensated for by rapid division of an adjacent OPC (Hughes et al. 2013).  Which of the surrounding OPCs respond is unpredictable at present, but several candidate indicators have been proposed (i.e. cell size, polarization, or access to growth factors) (van Heyningen et al. 2001; Bergles & Richardson 2015).  OPC surveillance behaviour, tiled distribution, and axo-glial synapses enable close observation of local myelination states, while retention of proliferative capacity enables a rapid response to demyelination or physiological stimuli (Bergles & Richardson 2015).  This homeostatic regulation may be disrupted in certain MS lesions, as OPC density is reduced despite a strong demand for new myelination (Lucchinetti et al. 1999; Chang et al. 2000; Boyd et al. 2013).  Pathological changes to ECM composition or cytokine profile may also compromise OPC density regulation (Bergles & Richardson 2015).    1.5.4.5 OPCs in the adult human CNS In contrast to their murine counterparts, adult human OPCs are much less characterized as there is limited material for analysis (i.e. post-mortem tissue or biopsies) (Dimou & Gallo 2015).  Human OPCs display several similarities to rodent OPCs, including: (i) in vitro expression of O4, PDGFRα, and MyT1 (Armstrong et al. 1992; Gogate et al. 1994), (ii) have been shown to proliferate in vivo (Geha et al. 2010; Rhee et al. 2009), and (iii) demonstrate a prolonged cell cycle, which is attributed to either an extended G1-phase or stalling at different G1 check-points (similar to that postulated for murine OPCs) (Dimou et al. 2008; Simon et al. 2011).  By contrast, several features of human OPCs deviate from their murine counterparts, including: (i) differential growth 49  factor responsiveness (e.g. PDGF/bFGF does not elicit proliferation of human OPCs, but rather morphological alterations) (Armstrong et al. 1992; Gogate et al. 1994; Yong et al. 1988), (ii) a lower relative population size in the human CNS (1-3% of all neural cells, as compared to 5-8% in rodents) (Scolding et al. 1998), and (iii) relatively reduced production of oligodendrocytes in the adult CNS (as only ~1/300 oligodendrocytes are replaced per year in humans) (Yeung et al. 2014).  These deviations are highly relevant to clinical trials attempting to promote myelination of the adult CNS based on pre-clinical studies in rodent models.  For example, human-derived neural stem cells (NSCs) transplanted into Pelizaeus-Merzbacher disease patients (characterized by failed CNS myelination) were suggested to form myelin (by MRI), but produced minimal functional improvement (Gupta et al. 2012).  Further characterization of adult human OPCs (specifically in their deviations from rodent OPCs) may be critical for understanding demyelinating diseases (e.g. MS), learning/memory (i.e. the role of myelin plasticity), and other CNS pathologies (Dimou & Gallo 2015; Franklin 2002; Bergles & Richardson 2015).   1.6 Central nervous system (CNS) myelination The evolution of CNS myelination enabled jawed vertebrates to transcend evolutionary pressures constraining nervous system size (i.e. head size, etc.) and develop a more computationally complex, functionally efficient, and powerful nervous system (Hartline et al. 2007; Zalc et al. 2008).  Myelin sheaths are multi-lamellar spiral wraps of glial cell membrane deposited around axons that function to increase membrane resistance and decrease axonal capacitance, thus enabling rapid action potential conduction (Geren 1954; Bunge et al. 1962; Chang et al. 2016).  Furthermore, myelinating glial cells also actively promote voltage-gated Na+-channel clustering at nodes of Ranvier (i.e. gaps between adjacent myelin sheaths), critical for saltatory (i.e. ‘jumping’) 50  conduction (Hartline et al. 2007; Eshed-Eisenbach & Peles 2013; Normand & Rasband 2015).  Multiple properties of myelin sheaths affect conduction velocity, including thickness, length, and axonal coverage pattern (Waxman 1980; Waxman 1997; Babbs & Shi 2013).  Moreover, nodal length and channel density influence velocity and efficiency of action potential propagation (Chang et al. 2016).  It is tempting to speculate that dynamic activity-dependent regulation of CNS myelination parameters (i.e. length, thickness, etc.) may modulate synchronicity of neural circuits (affecting network coordination and oscillatory activity), representing an alternative form of neural plasticity (Fields 2008; Fields 2015; Pajevic et al. 2014).  1.6.1 Target selection  Myelination status and axonal diameter are closely correlated, as CNS axons above a size threshold (≥0.2-0.4 µm) are more likely to be myelinated (Hildebrand et al. 1993; Wang et al. 2008).  This raises two questions: (i) it is simply geometry that promotes myelination? and (ii) what signaling mechanism conveys the size threshold? (Chang et al. 2016).  Studies on peripheral myelination (i.e. Schwann cells) has revealed partial answers.  Schwann cell myelination is responsive to axonal expression of Nrg1-III (mediated by ErbB receptors) with above-threshold levels promoting myelination (even when faced with inhibitory biophysical cues) (Salzer 2015; Mei & Nave 2014).  How axonal Nrg1-III levels are regulated in proportion with diameter remains unknown.  Surprisingly, Nrg1-ErbB signaling appears largely dispensable for CNS myelination (Brinkmann et al. 2008).  Furthermore, several studies suggest that oligodendrocytes may not even require an instructive signal to initiate myelination.  Oligodendrocytes extend membrane sheets containing myelin proteins in the absence of neurons in vitro (Mirsky et al. 1980; Aggarwal et al. 2011).  Furthermore, oligodendrocytes myelinate paraformaldehyde-fixed axons (Rosenberg et al. 51  2008), as well as electrospun nanofibers above a threshold size of 0.3-0.4 µm (Lee et al. 2012).  This threshold may represent a biophysical barrier for myelination, as very few axons <0.3 µm in diameter are myelinated in vivo (Hildebrand et al. 1993; Wang et al. 2008).  Therefore, oligodendrocytes appear to display broad target selection, mediated (at least in part) by molecular sensors that assess biophysical properties (i.e. diameter or curvature) (Chang et al. 2016).  The presence of stereotypical developmental myelination patterns implies that local environmental cues may modulate myelination (Foran & Peterson 1992; Brody et al. 1987; Kinney et al. 1988).  Indeed, myelin does not form on non-axonal structures of sufficient threshold diameter (i.e. dendrites, neuronal somata, etc.) (Chang et al. 2016).  The demonstration of long unmyelinated stretches of otherwise myelinated cortical axons, suggests that myelination can also be differentially regulated even along the same axon (Tomassy et al. 2014).  This might be accomplished by localization of myelination cues along the axon, overlain on an overall graded expression (which may be activity-dependent) (Piaton et al. 2010; Taveggia et al. 2010).  1.6.2 Models of myelin wrapping mechanics Myelin sheath formation involves a dramatic transformation of the corresponding glial cell.  For example, a myelin sheath of 10 layers, 100 µm long, formed on a 1 µm diameter axon requires the production of >6400 µm2 of membrane (Chang et al. 2016).  Formulation of a comprehensive model of myelin wrapping mechanics has been hindered by the delicacy of myelin sheaths and technical complications of nm resolution imaging.  Most of our current understanding is derived from in vitro Schwann cell studies (Geren 1954; Robertson 1955; Bunge et al. 1989).  The initial ‘carpet crawler’ model postulated that the inner membrane tongue ‘crawls’ around the axon, leaving behind a spiral myelin sheath (Geren & Raskind 1953; Geren 1954; Bunge et al. 1989).  52  This model predicts that the inner tongue advances as a straight line, thus there should be a homogenous number of wraps along the axon.  This was contradicted by early EM studies that revealed more wraps in the middle of a forming myelin sheath (Webster 1971; Fraher 1973; Fraher 1978), suggesting that the inner tongue instead advances as a convex curve.  Furthermore, during myelin sheath formation, radial growth (i.e. increase in wraps) and longitudinal growth (i.e. widening of the sheath) must be concurrent.  Development of cell culture systems, fluorescent reporter proteins (e.g. GFP), and electron microscopy led to the ‘ofiomosiac’ (Ioannidou et al. 2012) and ‘liquid croissant’ models (Sobottka et al. 2011), which differ fundamentally on membrane extension.  For example, the ‘ofiomosaic’ model postulates that membrane extension is accomplished through lateral expansion (Ioannidou et al. 2012), whereas in the ‘liquid crossant’ model the membrane extends from the outermost layer (Sobottka et al. 2011).  Limitations in spatial resolution of light microscopy, cell culture conditions, specific protein labeling, and artefactual-prone EM imaging prevented reconciliation of these opposing models (Chang et al. 2016).  The combination of high-pressure freezing techniques (which enhanced preservation of tissue architecture and reduced artefacts introduced by EM processing) (Mӧbius et al. 2010) with high resolution in vivo imaging have led to the current, and purportedly most comprehensive, model of myelin wrapping mechanics (Czopka et al. 2013; Snaidero et al. 2014).  The model is as follows: (i) Exploratory oligodendrocyte processes transform into short, elongating myelin sheaths (Czopka et al. 2013).  (ii) The number of wraps is highest at the point of initial axonal contact, and gradually decrease towards the ends (Snaidero et al. 2014).  (iii) Longitudinal length is shortest for the innermost layer and longest for the outermost layer, with length gradually increasing for the in-between layers (i.e. outer layers completely cover inner layers) (Chang et al. 2016).  (iv) The inner membrane tongue spirally wraps the axon (consistent with helical coil patterns observed by 53  light microscopy) (Chang et al. 2016).  (v) Each wrap has cytoplasm-containing edges forming part of a continuous collar, which remain attached to the axon and eventually form the para-nodal loops as they are moved to the edge of the sheath during longitudinal expansion (Eshed-Eisenbach & Peles 2013; Normand & Rasband 2015).  1.6.3   Growth of the myelin sheath The inner tongue of the forming myelin sheath is considered to be the primary growth zone, revealed by accumulation of newly-synthesized vesicular stomatitis glycoprotein G (VSV-G) (Snaidero et al. 2014).  Interestingly, transient cytoplasmic channels flanked by compact myelin are observed in forming myelin sheaths (Snaidero et al. 2014), thought to mediate the delivery of new membrane (in vesicles) to the inner tongue growth zone (Chang et al. 2016).  The channels appear to gradually close as compaction progresses, but are re-formed during the addition of new wraps to pre-existing myelin sheaths (Snaidero et al. 2014).  Actin cytoskeletal dynamics appear critical to inner tongue advancement during myelination (Nawas et al. 2015; Zuchero et al. 2015).  Indeed, filamentous actin is observed in actively growing sheaths, but absent from compacted myelin (Snaidero et al. 2014; Nawas et al. 2015; Zuchero et al. 2015).  Furthermore, actin filament depolymerisation relieves oligodendrocyte membrane tension in vitro, resulting in membrane spreading (Nawas et al. 2015; Zuchero et al. 2015).  Moreover, pharmacological-induced actin depolymerisation promotes myelin wrapping in the mouse spinal cord in vivo (Zuchero et al. 2015).  Finally, knockout of actin disassembly factors reduces the number of wraps per myelin sheath (Nawas et al. 2015).  To accommodate a variable growth rate across the advancing growth zone, new membrane delivery needs to be tightly coordinated with localized modulation of actin dynamics.  Radial growth is gradually reduced when the final number of wraps is reached, whereas 54  longitudinal growth is maintained as the shorter inner layers need to elongate to match the more mature, and longer, outer layers (Chang et al. 2016).  Indeed, most myelinated axons in the CNS display close to optimal g-ratios (for action potential conduction) (Chomiak & Hu 2009), larger diameter axons have more wraps (Hildebrand et al. 1993), and near integral numbers of wraps are formed (Peters 1964; Waxman & Swadlow 1976).  Interestingly, myelin sheaths formed on nanofibers do not display typical g-ratios (Lee et al. 2012), implicating axonal-derived cues in the diameter-dependent regulation of sheath thickness.  Axonal levels of Nrg1-III are directly correlated with the number of myelin wraps formed by Schwann cells (Nave & Salver 2006), however as conditional deletion of Nrg1 does not affect CNS myelination (Mei & Nave 2014; Brinkmann et al. 2008), alternative cues are likely involved that converge on the regulation of vesicle delivery and actin dynamics in the inner tongue (Chang et al. 2016).  Myelin sheath retraction by oligodendrocytes occurs during zebrafish CNS development (Czopka et al. 2013).  Membrane growth zones may also be regions of endocytosis and membrane removal, combined with retrograde vesicular transport through re-opened cytoplasmic channels (Chang et al. 2016).  Modulation of membrane growth and/or removal mechanisms may partially be responsible for myelination changes in response to physiological stimulation or experience.    1.6.4 Myelin sheath compaction Myelin functionality requires extensive compaction (extruding most cytoplasm and extracellular space) to acquire a low dielectric permittivity, crucial for reducing axolemmal capacitance (Eshed-Eisenbach & Peles 2013; Normand & Rasband 2015; Chang et al. 2016).  Compact myelin is characterized by the extraordinary close apposition (~1-3 mm) of plasma membranes over large surface areas (Bunge et al. 1962).  This is achieved by two types of compaction: (i) intracellular 55  compaction (i.e. between two cytoplasmic membrane leaflets, forming the major dense line, as seen under EM), and (ii) extracellular compaction (i.e. between two extracellular leaflets, forming the intra-period lines).  Initially, the compaction was attributed to the myelin proteins MBP and PLP (thought at the time to comprise >80% of myelin proteins), however the process is more complicated (Chang et al. 2016).    1.6.4.1 Intracellular compaction The cytoplasmic protein MBP (myelin basic protein) comprises ~8% of myelin proteins (Jahn et al. 2009), and is implicated in compaction of the two cytoplasmic leaflets.  Indeed, the major dense line is absent in the few uncompacted myelin sheaths that form in shiverer mice (i.e. MBP mutants) (Bird et al. 1978; Privat et al. 1979; Rosenbluth 1980).  The MBP protein can carry up to 19 net positive charges (at neutral pH), as well as bind to negatively charged lipid head groups (Harauz et al. 2009; Harauz & Libich 2009; Nawas et al. 2009).  Furthermore, the protein contains amphipathic α-helical regions that enable partial membrane embedment (Harauz et al 2009).  Therefore, MBP is proposed to neutralize negative charges on the membranes, thus enabling the two cytoplasmic leaflets to come into close apposition by overcoming electrostatic repulsion (Chang et al. 2016).  Interestingly, MBP proteins self-associate into higher-order structures upon membrane interaction and are thought to organize into a hydrogel-like cohesive network during compaction (Aggarwal et al. 2012; Kattnig et al. 2012; Aggarwal et al. 2013), potentially akin to β-amyloid aggregation (Aggarwal et al. 2013).  Membrane interaction also induces formation of α-helices and β-sheets in MBP proteins (Aggarwal et al. 2011; Harauz et al. 2009).  Furthermore, charge neutralization and pH elevation induces concentration-dependent µm-scale droplet formation in purified MBP solutions (Aggarwal et al. 2013; Lee et al. 2014).  Findings from 56  heterologous cell lines, oligodendrocyte cultures, and cell-free membrane systems revealed the presence of a diffusion barrier formed by the MBP network, that effective excludes proteins of a certain size (>30 amino acid residues) from MBP-occupied domains (Aggarwal et al. 2011; Aggarwal et al. 2013).  Therefore, MBP promotes intracellular compaction through charge neutralization, self-assembly into a mesh-like network, and formation of a diffusion barrier.    1.6.4.2 Extracellular compaction The transmembrane protein PLP comprises ~17% of myelin proteins (Palaniyar et al. 1998; Bizzozero & Howard 2002; Bakhti et al. 2013).  Surprisingly, PLP-/- mice display normal intra-period lines and myelination (Klugman et al. 1997; Mӧbius et al. 2008), suggesting either a minimal role in compaction or functional redundancy.  In contrast to intracellular compaction, extracellular compaction is thought to be mediated by non-specific van der Waals forces and hydrogen bonding (Bakhti et al. 2013; Coetzee et al. 1999).  Therefore, the main barriers are electrostatic repulsion between opposing membranes and steric hindrance caused by the glycocalyx (Bakhti et al. 2013).  Exclusion of transmembrane glycoproteins from the compacting region during intracellular compaction (i.e. by the MBP diffusion barrier) may aid extracellular compaction (Aggarwal et al. 2013), however as shiverer mice (i.e. MBP mutants) retain intra-period lines it does not appear to be a requirement (Privat et al. 1979; Inoue et al. 1981).  Increased membrane polysialic acids (via oligodendrocyte-specific polysialytransferase overexpression) is accompanied by split myelin lamellae, attributed to increased steric hindrance (Bakhti et al. 2013; Schnaar et al. 2014).  Furthermore, membrane surface sugar residues are decreased during oligodendrocyte differentiation.  Therefore, removal of the glycocalyx may be a prerequisite for effective extracellular compaction (Bakhti et al. 2013).  Early EM studies revealed initiation of 57  compaction prior to the formation of three layers (Hildebrand et al. 1993).  Furthermore, intra-period lines are observed between cytoplasm-containing layers early in development (Snaidero et al. 2014).  Therefore, reduction of the glycocalyx during oligodendrocyte differentiation uncovers weak interactions that drive extracellular compaction early during myelination.  Intracellular compaction proceeds from the outermost layer inwards (Snaidero et al. 2014), the progression of which is affected by the relative abundance of MBP and CNP (Jahn et al. 2009).  Indeed, intracellular compaction is accelerated in CNP+/- mice and delayed in MBP+/- mice (Snaidero et al. 2014).  Furthermore, CNP overexpression inhibits intracellular compaction, CNP levels far exceed MBP in uncompacted myelin, whereas MBP far exceeds CNP in compacted myelin (Gravel et al. 1996; Yin et al. 1997).  Therefore, intracellular compaction may be regulated by mutual exclusivity between MBP and components of uncompacted myelin (e.g. CNP), similar to the establishment of the axon initial segment, which involves mutual exclusion of various cytoskeletal proteins (Normand & Rasband 2015; Galiano et al. 2012).  Activity-dependent modulation of CNP-MBP dynamics may underlie the reopening of cytoplasmic channels to allow for modulation of myelin sheath thickness, length, or even sheath retraction (Chang et al. 2016).  1.6.5 Myelinating glial support of axons In addition to increasing conduction velocity, myelination may provide axonal support (Nave 2010; Nave & Trapp 2008).  Indeed, as axons can be remarkably long (e.g. >1 m in humans) and axonal transport of metabolic enzymes is slow (0.2-6 mm/day) (Fünfschilling et al. 2012; Lee et al. 2012; Nave 2010; Nave & Trapp 2008; Oblinger et al. 1988), complete metabolic reliance on neuronal somata is not feasible (Fünfschilling et al. 2012; Lee et al. 2012).  Glia-derived trophic support may be derived from two primary sources: (i) astrocytic provision of glucose via nodes of 58  Ranvier (Butt et al. 1994), and/or (ii) oligodendrocyte-derived lactate released through MCT-1 (monocarboxylate transporter-1) (Fünfschilling et al. 2012; Lee et al. 2012; Rinholm et al. 2011).  As myelin sheaths have substantially larger axonal interface area, oligodendrocyte-derived metabolic support is likely more substantive (Brown et al. 2004).  Indeed, reduction of oligodendrocyte MCT-1 expression leads to axonal damage (Lee et al. 2012).  Furthermore, loss of oligodendrocyte-derived metabolic support is implicated in the increased susceptibility of persistently demyelinated axons to degeneration in various white matter pathologies (Irvine & Blakemore 2008).  One argument for promoting remyelination following SCI is to restore metabolic support and reduce vulnerability of the denuded axons to the injury environment (Plemel et al. 2014).  Indeed, oligodendrocyte ablation with diphtheria toxin (Ghosh et al. 2011; Pohl et al. 2011; Traka et al. 2010) or by caspase-9 activation (Caprariello et al. 2012) leads to demyelination and axonal damage (accompanied by reactive gliosis).  Furthermore, mutations in myelin proteins (e.g. PLP, CNP, MAG) lead to axonal damage/degeneration, despite no overt myelin deficits (Griffiths et al. 1998; Lappe-Siefke et al. 2003; Nguyen et al. 2009; Petzold 2005).  Interestingly, animal models that fail to form compact myelin (MBP mutants) do not show axonal degeneration.  For example, neither shiverer mice (up to 2-3 months old) (Griffiths et al. 1998; Loers et al. 2004) nor Long-Evans shaker rats (up to 9 months old) (Smith et al. 2013) show axonal damage/degeneration despite extended demyelination.  Therefore, oligodendrocyte presence may be sufficient for axonal support, whether or not compacted myelin is formed (Plemel et al. 2014).    1.6.6 Myelin plasticity/adaptive myelination in the adult CNS Functional optimization is not achieved through maximization of conduction velocity, as synchronicity of activity can be critical to network function (e.g. visual system) (Waxman 1997; 59  Seidl 2014).  Therefore, not all CNS myelin parameters in vivo are optimized to achieve the fastest action potential conduction (Chang et al. 2016).  Oligodendrocytes appear to respond to individual axon activity through modulation of myelin parameters, however underlying mechanisms remain largely uncharacterized (Richardson et al. 2011; Zatorre et al. 2012; Wang & Young 2014; de Hoz & Simons 2015).  In theory, myelination can modulate neural circuit function by several mechanisms: (i) de novo myelination of unmyelinated axons and/or axonal segments, (ii) remyelination of denuded axons, (iii) alteration of thickness and/or length of existing sheaths, or (iv) retraction of myelin sheaths (Chang et al. 2016; Fields 2015).  Abundant evidence exists for myelin plasticity in the murine CNS.  For example, social isolation of juvenile mice reduces the number and thickness of myelin sheaths formed by individual oligodendrocytes in the prefrontal cortex, associated with deficits in sociability and working memory (Makinodan et al. 2012).  Furthermore, 8-week social deprivation of adult mice reduces myelin gene expression in the pre-frontal cortex, correlated with thinner myelin sheaths, which was reversed by a 4-week social reintegration regime (Liu et al. 2012).  There is ample substrate for de novo myelination in the adult CNS.  Early EM studies revealed numerous unmyelinated axons of sufficient size (i.e. 0.4-0.8 µm in diameter) (Hildebrand et al. 1993), as well as long unmyelinated segments of otherwise myelinated cortical axons (Tomassy et al. 2014).  Consistent with this, oligodendrogenesis continues throughout post-natal life (Richardson et al. 2011; Zatorre et al. 2012; Wang & Young 2014; de Hoz & Simons 2015).  Indeed, tracing of 14C concentration in human oligodendrocyte DNA suggested that the population expands through the fourth decade of life, followed by an annual turnover rate of 0.3% thereafter (Yeung et al. 2014).  Oligodendrogenesis is also observed in the intact adult rodent spinal cord, with a 3-4% turnover over a 12-week period (Barnabe-Heider et al. 2012).  Furthermore, learning to run on an irregularly spaced wheel (i.e. a complex motor 60  task) stimulates OPC proliferation and oligodendrogenesis in adult mice (McKenzie et al. 2014).  Prevention of oligodendrogenesis (via MyRF deletion) impairs motor performance, implicating adult de novo myelination in motor learning (McKenzie et al. 2014).  These studies, and others, demonstrate that environmental stimuli can modulate myelination, which is a decades-old proposal (Fields 2008), presumably mediated through alteration of neural activity.  Several studies have addressed the direct relationship between neural activity and myelination.  For example, unilateral corticospinal tract stimulation induces OPC proliferation and oligodendrogenesis on the stimulated side (Li et al. 2010).  Furthermore, optogenetic-mediated stimulation of the M2 motor cortex was associated with increased oligodendrogenesis and thicker myelin sheaths (Gibson et al. 2014).  The mechanistic basis of activity-dependent modulation of myelination remain largely uncharacterized.  Modulation of existing myelin sheath thickness has been demonstrated in the adult CNS.  For example, PTEN knockout in mature oligodendrocytes leads to myelin sheath thickening via reopening of cytoplasmic channels to shuttle membrane to the inner tongue growth zone (Snaidero et al. 2014; Goebbels et al. 2010).  In contrast, sheath thinning is less well characterized, however it may be a variation of sheath retraction, as revealed in zebrafish larvae (Czopka et al. 2013).  Myelin sheath retraction implies a degree of uncertainty in oligodendrocyte target selection in vivo, which may be rectified by dynamic editing processes to ensure myelination of appropriate structures (Chang et al. 2016).  Selection of targets may not require neuronal activity, but can be significantly affected by it (Hines et al. 2015; Mensch et al. 2015).  For example, inhibition of neuronal synaptic vesicle release (via tetanus neurotoxin) reduces the number of sheaths formed by individual oligodendrocytes, attributed to decreased stability of nascent sheaths (Hines et al. 2015; Mensch et al. 2015).  The demonstration of de novo myelination in heavily myelinated adult white matter regions (e.g. optic nerve, where >98% of axons are 61  myelinated) (Young et al. 2013), suggests that continual oligodendrogenesis and myelination may be required to replace myelin lost due to natural processes throughout post-natal life.  Neither the normal lifespan of oligodendrocytes nor whether internodes can be resorbed over the long term is known, however newly-differentiated oligodendrocytes appear to form all of their internodes within a narrow time frame, after which they do not undergo significant alteration (Watkins et al. 2008; Czopka et al. 2013; Young et al. 2013).  Adult-born oligodendrocytes form more, but shorter, internodes than their developmentally-formed counterparts (Young et al. 2013).  Indeed, in the murine CNS the average internodal length decreases with age (Lasiene et al. 2009), while the number of nodes per unit volume increases (Peters & Kemper 2012).  This may enable the internodes of adult-born oligodendrocytes to intercalate amongst pre-existing internodes, enabling minute adjustments in conduction velocity, particularly important for neural systems reliant on synchronous network activity (e.g. visual and auditory systems) (Fields 2015; Chang et al. 2016).  Indeed, dynamic alteration of internode length and/or thickness is thought responsible for microsecond timing adjustments in the auditory system (Seidl 2014).  Despite the majority of processes being shorter than their developmental counterparts, many adult-born oligodendrocytes in the optic nerve produce one or two internodes that are much longer (>200 µm) (Young et al. 2013), probably formed on the small fraction of previously unmyelinated axons.  In adult CNS regions that contain a much larger fraction of unmyelinated axons, de novo myelination could significant alter neural circuit properties (Chang et al. 2016).  MRI imaging revealed structural changes in human white matter regions in response to specific motor learning (e.g. playing a musical instrument or juggling), proposed to represent myelin plasticity (Bengtsson et al. 2005; Scholz et al. 2009; Zatorre et al. 2012) in response to repetitive activity in nascent or preformed neural circuits.  Similarly, performance of a unilateral reaching task induces contralateral changes 62  in subcortical white matter and overlying gray matter in rats, as revealed by MRI, correlated with increased MBP immunoreactivity (Sampaio-Baptista et al. 2013).  Wheel running also alters OPC proliferation and differentiation dynamics (Ehninger et al. 2011; Simon et al. 2011; McKenzie et al. 2014), although this might be due to exercise rather than learning per se.  Interestingly, OPC proliferation and differentiation may follow circadian rhythms, consistent with the importance of sleep for memory consolidation and motor learning (Matsumoto et al. 2011; Bellesi et al. 2013).  In addition to motor learning, CNS myelination has been shown to be responsive to various stimuli, including social experience (Liu et al. 2012; Makinodan et al. 2012), enriched environmental exposure (Zhao et al. 2012), exercise (Ehninger et al. 2011; Simon et al. 2011), and pregnancy (Gregg et al. 2007), amongst others (Fields 2015; Fields 2008; Richardson et al. 2011; Wang & Young 2014; Chang et al. 2016).  Taken together, adult oligodendrogenesis and myelination is a highly dynamic, tightly regulated process capable of adapting and refining neural circuit function in response to physiological demand (Bergles & Richardson 2015).  1.6.7 Oligodendrocyte turnover in the human CNS: implications for myelin plasticity In contrast to rodents, oligodendrocyte turnover in the adult human CNS is limited, estimated at ~1/300 oligodendrocytes per year, following the fourth decade of life (Yeung et al. 2014).  MRI studies have revealed white matter structural changes in people performing complex motor tasks (i.e. juggling or musical instrument playing) (Bengtsson et al. 2005; Scholz et al. 2009; Zatorre et al. 2012), proposed to reflect activity-dependent myelin plasticity (Chang et al. 2016).  Similar changes in rodent models have been attributed to increased oligodendrogenesis and de novo myelination (Gibson et al. 2014; McKenzie et al. 2014).  By contrast, the alteration of pre-existing myelin sheaths (i.e. alteration of length, thickness, etc.) probably has a more important role in adult 63  human myelin plasticity than de novo myelination, due to the limited oligodendrocyte turnover in the adult human CNS (Yeung et al. 2014; Chang et al. 2016).     1.6.8 Inhibitors of remyelination Remyelination following SCI is considered complete, albeit it is delayed (Lasiene et al. 2008; Powers et al. 2013; Plemel et al. 2014).  Studies of CNS demyelination have revealed numerous extracellular inhibitors of remyelination, including myelin debris, LINGO-1, Wnt-signaling, ECM molecules, and inflammatory components, among others (Plemel et al. 2014; Bergles & Richardson 2015).     1.6.8.1 Myelin debris Myelin debris inhibits remyelination.  For example, extracellular myelin inhibits myelin gene expression in vitro (Plemel et al. 2013), injection of myelin into the demyelinated CNS impairs remyelination (Kotter et al. 2006), and myelin debris is observed in close apposition to demyelination axons (Kotter et al. 2001; Kotter et al. 2005).  Indeed, inefficient myelin clearance in the CNS is implicated in remyelination failure.  For example, depletion of circulating monocytes attenuates myelin phagocytosis and impairs remyelination, with a heightened effect in aged animals (Kotter et al. 2001; Kotter et al. 2005).  Furthermore, heterochronic parabiosis (surgical joining of the circulation of two animals) increases remyelination in aged animals, correlated with increased myelin phagocytosis, as well as enhanced oligodendrogenesis (Ruckh et al. 2012).  Myelin debris persistence is observed in SCI tissue for up to 8 wpi in rodents  (Buss & Schwab 2003) and 3 years in humans (Buss et al. 2005).  Furthermore, the highest concentration of 64  oligodendrogenesis in the tissue rim following SCI is spatially correlated with the highest density of macrophages, known to phagocytose myelin debris (Tripathi & McTigue 2007).    1.6.8.2 LINGO-1 LINGO-1 has also been demonstrated to be an inhibitor of remyelination.  For example, LINGO-1 antagonism increases myelination in vitro (Mi et al. 2009), promotes remyelination following demyelination (Mi et al. 2007; Mi et al. 2009), as well as improves functional recovery following SCI (Ji et al. 2006).  It should be noted that potential influence on axonal plasticity may complicate the interpretation of these findings (Ji et al. 2006).    1.6.8.3 Wnt-signaling Another inhibitor of remyelination is Wnt-signaling, which also inhibits developmental myelination (Fancy et al. 2009; Fancy et al. 2011).  For example, expression of a constitutively-active β-catenin in oligodendrocyte lineage cells inhibits differentiation as well as remyelination (Fancy et al. 2009).  Furthermore, Axin2 stabilization promotes oligodendrocyte differentiation in vitro (Huang et al. 2009), remyelination in cerebellar slice cultures (Fancy et al. 2011), and remyelination following CNS demyelination in vivo (Fancy et al. 2011).    1.6.8.4 Extracellular matrix (ECM) components ECM components also inhibit remyelination.  For example, hyaluronan reduces myelin gene expression in vitro, and accumulates in MS lesions (Jiang et al. 2011; Back et al. 2005).  Furthermore, hyaluronan administration inhibits remyelination, an effect abolished in TLR2-/- mice (Sloane et al. 2010).  CSPGs inhibit oligodendrocyte development in vitro, reversed by ChABC-65  mediated digestion (Keough & Yong 2012; Lau et al. 2013; Siebert & Osterhout 2011).  Furthermore, digestion of lesion-derived CSPGs improves remyelination following lysolecithin-mediated demyelination (Lau et al. 2013).  Finally, CSPGs have also been shown to limit OPC migration (Siebert et al. 2011).  Semaphorin-3A impairs maturation of oligodendrocytes in vitro (Syed et al. 2011), however the in vivo situation is more complicated.  For example, semaphorin-3A infusion impairs remyelination (Boyd et al. 2013; Syed et al. 2011), but overexpression does not enhance remyelination (Piaton et al. 2011).  Following SCI, semaphorin-3A levels peak at 1 wpi in regions of gliosis (i.e. near the lesion) (Kaneko et al. 2006).    1.6.8.5 Inflammation In addition to ECM components, various inflammatory processes significantly influence remyelination.  For example, inhibition of microgliosis (via minocycline) or depletion of macrophages impairs remyelination (Li et al. 2005).  Both M1 (pro-inflammatory) and M2 (anti-inflammatory) macrophages are involved in promoting remyelination (Miron et al. 2013).  For example, inhibition of M1 macrophage recruitment to demyelinated lesions is associated with impaired OPC proliferation, whereas M2 depletion impairs oligodendrocyte differentiation (Miron et al. 2013).  M2 macrophages are important for myelin debris phagocytosis (Miron et al. 2013; Ruckh et al. 2012) and produce factors that promote oligodendrocyte differentiation (e.g. activin-A) (Miron et al. 2013).  Interestingly, depletion of CD4+ and/or CD8+ T-cells impairs remyelination (Bieber et al. 2003), potentially related to the role of innate immune cells in myelin phagocytosis (Ghasemlou et al. 2007).  Stimulation of monocyte and lymphocyte influx in a rodent model of progressive demyelination without enhanced inflammation at 1-year-old (i.e. taiep rats) promotes remyelination, specifically in regions of leukocyte infiltration (Foote & Blakemore 2005; 66  Wilkins et al. 2010; Duncan et al. 1992), suggesting that inflammatory cues are sufficient to promote remyelination of chronically demyelinated axons (Plemel et al. 2014).  Several inflammatory cytokines are implicated in remyelination, including TNF-α (Arnett et al. 2001), IL-1β (Mason et al. 2001), IL-11 (Zhang et al. 2006), lymphotoxin-β receptor (Plant et al. 2007), and endothelin-2 (Yuen et al. 2013).  In addition, several other factors produced by inflammatory cells may also enhance remyelination, including IGF-1 (Mason et al. 2001), BDNF (Skihar et al. 2009; Xiao et al. 2010), and CNTF (Skripuletz et al. 2011).  1.7 Mouse spinal cord anatomy As the current study assesses injury-induced effects remote to the lesion site, it is important to identify the expected position of specific tracts and how they are affected by the injury (i.e. severed vs. spared).  The adult mammalian spinal cord white matter contains ascending sensory afferents and descending motor efferents.  The anatomy of specific spinal tracts is highly conserved across mammals, with the notable exception of the dorsal corticospinal tract (dCST) (Watson 2009; Watson & Harrison 2012).  The specific composition/arrangement of spinal tracts varies across spinal levels (fig. 1.1).       1.7.1 Main descending tracts The main descending tracts of the mouse spinal cord arise from neuron cell bodies residing in the cerebral cortex, red nucleus, hindbrain reticular nuclei, and vestibular nuclei, among others (Liang et al. 2011).  The mouse corticospinal tract arises from the cerebral cortex, decussates in the hindbrain, and descends in the ventral portion of the dorsal funiculus (Watson & Harvey 2009; Watson 2012).  Axonal size can be used to distinguish the dCST (small caliber axons) from the 67  fasciculus gracilis and fasciculus cuneatus (medium caliber axons) (Watson & Harrison 2012).  The position of the dorsal CST is clearly illustrated in Thy1-eYFP transgenic mice, which labels neocortical projection axons (Porrero et al. 2010).  The mouse rubrospinal tract arises from the red nucleus, decussates in the caudal midbrain, and descends in the dorsolateral corner of the lateral funiculus (marked by calretinin staining) (Nyberg-Hansen 1965; Watson et al. 2009; Liang et al. 2012).  Axonal projections from the vestibular nuclei descend as the lateral and medial vestibulospinal tracts.  The un-crossed lateral vestibulospinal tract descends in the ventral funiculus and shifts laterally as it descends, whereas the crossed medial vestibulospinal tract descends in the medial longitudinal fasciculus (adjacent to the ventral median fissure), and terminates prior to lumbar spinal levels (Nyberg-Hansen & Mascitti 1964; Watson & Harrison 2012).  The rostral reticulospinal tract descends in the ventral funiculus, specifically around the ventral and ventromedial periphery.  The caudal reticulospinal tract descends in the lateral funiculus in close proximity to the ventral horn lateral boundary.  The two reticulospinal tracts maintain the same relative position during their descent (Nyberg-Hansen 1965; Nathan et al. 1996).   See fig.1.1 for overview of spinal tracts in the (a) cervical and (b) lumbar spinal cord.                    1.7.2 Main ascending tracts  The majority of the mammalian dorsal funiculus is composed of the fasciculus gracilis and fasciculus cuneatus, which contain ascending sensory afferents of ipsilateral primary neurons (Watson & Harrison 2012).  Fasciculus gracilis and fasciculus cuneatus anatomy is highly consistent across mammalian species (Watson 2012).  A subset of axons in the dorsal column arise from spinal dorsal horn neurons, called the post-synaptic dorsal column (Giesler et al. 1984).  The ventral spinocerebellar and dorsal spinocerebellar tract ascend in the periphery of the lateral 68  funiculus, in contact with the surface of the lateral and ventrolateral funiculi (Terman et al. 1998; Xu & Grant 1994).  As they ascend from lumbar to cervical spinal levels, the dorsal spinocerebellar tract shifts dorsally and the ventral spinocerebellar tract shifts laterodorsally.  The dorsal spinocerebellar tract is ipsilateral to its origin in dorsal root ganglia (DRG) neurons, whereas ventral spinocerebellar tract axons decussate via the ventral white commissure close to their level of spinal cord entry (Watson & Harrison 2012).  Human spinothalamic afferents are typically divided into lateral spinothalamic tract and ventral spinothalamic tract, while in the murine spinal cord there is an additional dorsolateral spinothalamic tract (Apkarian & Hodge 1989; Stevens et al. 1991; Zhang et al. 2000; Friehs et al. 1995).  The dorsolateral and lateral spinothalamic tracts are located medial to the spinocerebellar tracts, whereas the ventral spinothalamic tract ascends in the ventral funiculus (Watson & Harrison 2012).  See fig.1.1 for overview of spinal tracts in the (a) cervical and (b) lumbar spinal cord.            Figure 1.2 Mouse spinal cord anatomy.  Expected anatomical locations of the main ascending (green) and descending (blue) axonal tracts in the (a) cervical and (b) lumbar mouse spinal cord.  Ascending tracts: fasciculus gracilis, gr; fasciculus cuneatus, cu; dorsal spinocerebellar tract, dsc; dorsolateral spinothalamic tract, dlst; ventral spinocerebellar tract, vsc; lateral spinothalamic tract, lst; ventral spinothalamic tract, vst.  Descending tracts: dorsal 69  corticospinal tract, dcs; rubrospinal tract, rs; caudal reticulospinal tract, crts; rostral reticulospinal tract, rrts; lateral vestibulospinal tract, lvs; medial vestibulospinal tract, mvs.  Adapted from Watson & Harrison (2012).  1.7.3 Cervical dorsal column: fasciculus gracilis and fasciculus cuneatus As mentioned, the dorsal column of the mouse spinal cord contains three main axonal groupings: (i) the medial fasciculus gracilis (a.k.a. Tract of Goll), (ii) the lateral fasciculus cuneatus (a.k.a. Tract of Burdach), and (iii) the dorsal corticospinal tract (dCST), located ventrally (Watson & Harrison 2012; Watson 2009).  The fasciculus gracilis and fasciculus cuneatus contain sensory afferents carrying proprioception, light touch, and vibration information that are running rostrally to the nucleus gracilis & nucleus cuneatus in the medulla oblongata (Watson & Harrison 2012).  Assuming a somatotopic organization of the dorsal column, the fasciculus gracilis carries ipsilateral sensory afferents from T7 and below, while the fasciculus cuneatus carries ipsilateral sensory afferents from T6 and above (Nord 1967; Johnson et al. 1968; Smith & Deacon 1984; Maslany et al. 1991).  Importantly, this anatomical arrangement means that these axonal tracts are differentially impacted by the T9-T10 contusion injury.  As they originate caudal to the injury, axons in the fasciculus gracilis are injured, while axons in the fasciculus cuneatus are spared as they originate rostral to the injury.  Furthermore, of practical importance is that these tracts can be reliably differentiated in cervical spinal cord tissue sections stained with broad-spectrum axonal markers (i.e. NF-200, SMI-312) (Watson & Harrison 2012).  In the current study we exploit this anatomical arrangement to assess the influence of axonal injury on oligodendrocyte lineage cells remote to the lesion site in the cervical spinal cord by comparing injury-induced changes between the fasciculus gracilis (i.e. injured axons) and the fasciculus cuneatus (i.e. spared axons).  70   Figure 1.3 Effect of injury on cervical fasciculus gracilis vs. fasciculus cuneatus.  Thoracic contusion injury differentially impacts the axons running in the cervical fasciculus gracilis (severed axons – red) and fasciculus cuneatus (spared axons – green).  This dichotomy was exploited to assess the influence of axonal degeneration on oligodendrocyte lineage cells remote to the lesion site (specifically OPC proliferation, oligodendrogenesis, and new myelination).        71  Chapter 2: Isolated thoracic contusion injury induces robust oligodendrocyte lineage cell responses remote to the lesion site in the murine spinal cord  2.1  Introduction At a histological level, injury-induced pathophysiological processes have been thoroughly characterized for the tissue area immediately surrounding the lesion epicentre (Tator 1995; Tator & Fehlings 1991), however remote to the lesion these changes are less well described.  Previous studies have demonstrated that various perturbations, including traumatic injury (Zai & Wrathall 2005; Buffo et al. 2005; Hesp et al. 2015), demyelination (Franklin 2002), artificial neural stimulation (Li et al. 2010; Gibson et al. 2014), neurodegeneration (Kang et al. 2013; Behrendt et al. 2013; Philips et al. 2013; Magnus et al. 2008), and social experience (Makinodan et al. 2012), among others, induce robust oligodendrocyte precursor cells (OPC) responses, which function as endogenous precursors for myelinating oligodendrocytes.  De novo myelination in the adult CNS has been implicated in motor learning, memory, and optimization of neural network function, thought to represent a potent form of neural plasticity (Chang et al. 2016; Fields 2015).  The demonstration of robust OPC proliferation and oligodendrogenesis in models of dorsal rhizotomy (Sun et al. 2010), axotomy (Neilson et al. 2006), and axon degeneration (Kang et al. 2013), combined with the robust OPC responses characteristic of SCI lesion epicentres (McTigue et al. 2001), lead us to hypothesize that contusion SCI would induce significant responses in resident OPC populations remote to the lesion site (specifically comprised of OPC proliferation, oligodendrogenesis, and new myelination).  This may be functionally relevant to myelin plasticity on spared motor and sensory tracts remote to the lesion (Chang et al. 2016; Fields 2015).  To test 72  this hypothesis, we conducted an in vivo study employing transgenic mouse lines (i.e. PDGFRα-CreERT2:ROSA26-YFP and PDGFRα-CreERT2:ROSA26-mGFP) that enabled the visualization and fate-mapping of OPCs and their progeny in the adult murine spinal cord following a moderate (70 Kdyne) T9-T10 thoracic contusion injury.       2.2 Materials and methods 2.2.1 Animals All procedures were approved by the University of British Columbia (Vancouver, B.C., Canada) Animal Care Committee (A.C.C.) in accordance with the guidelines established by the Canadian Council on Animal Care (C.C.A.C.).  Mice were generated through selective breeding of transgenic reporter mouse lines expressing either yellow fluorescent protein (YFP) or membrane-bound green fluorescent protein (mGFP) under the constitutively-active ROSA26 promoter to transgenic mouse lines expressing a Cre-recombinase/modified estrogen receptor fusion protein (CreERT2) under the control of the PDGFRα promoter.  Once generated, male and female PDGFRα-CreERT2:ROSA26-YFP (n=22) (fig. 2.1a) and PDGFRα-CreERT2:ROSA26-mGFP (n=9) transgenic mice (fig. 2.2b) were utilized to enable the conditional and inducible labeling of PDGFRα-expressing cells with the reporter proteins (i.e. YFP or mGFP), thus allowing visualization and fate-mapping of their progeny following SCI.  Animals were group-housed in standard mouse housing cages (up to 4 animals/cage) in reverse-lighting housing rooms (21 °C; 12 h:12h light:dark cycle) at the Blusson Spinal Cord Centre (B.S.C.C.; Vancouver, B.C., Canada) and were provided ad libitum access to standard mouse diet and water.        73   Figure 2.1 Transgenic mouse lines and experimental timeline.  Transgenic mouse lines used in the current study.  a) PDGFRα-CreERT2:ROSA26-YFP mice (n=22).  Tamoxifen administration induces the expression of cytoplasmic YFP in PDGFRα-expressing cells.  b) PDGFRα-CreERT2:ROSA26-mGFP mice (n=9).  Tamoxifen administration induces the expression of membrane-bound GFP in PDGFRα-expressing cells.  c) Experimental timeline.  Animals were dosed with tamoxifen (3 mg in 0.15 ml corn oil) for a period of 5 days, followed by a 2 week clearing period prior to a T9-T10 thoracic contusion SCI (70 Kdyne).  Animals were sacrificed at 1 dpi, 5 dpi, 2 wpi, and 12 wpi.  Mouse and syringe images obtained from the Servier® medical art database (accessed through http://www.servier.com/Powerpoint-image-bank).       2.2.2 Tamoxifen (TMX) administration To induce Cre-mediated recombination of the transgene construct (thus labeling PDGFRα-expressing cells with the reporter proteins), mice were administered tamoxifen (3 mg dissolved in 0.15 ml of corn oil; i.p.) for a period of 5-days, followed by a 2-week period prior to injury to allow for clearance of residual tamoxifen from the animals (to avoid any potential confounding influence on injury processes) (fig. 2.1c).  Non-tamoxifen treated control animals (to assess 74  spontaneous recombination of the transgene construct) were administered vehicle injections (0.15 ml corn oil; i.p.) for the same 5-day period, ending 2 weeks prior to injury.  2.2.3 Spinal cord injury (SCI) Following the 2-week clearance period, animals were subjected to a moderate (70 Kdyne) T9-T10 thoracic contusion spinal cord injury (SCI), as previously described (Choo et al. 2008; Sparling et al. 2015).  Animals were anesthetized with isoflurane (~4% for induction of anesthesia, ~1.5% for maintenance during surgical procedures) in oxygen (1 L/min).  During surgical preparation, animals were placed onto an electric heating pad to maintain body temperature.  Once anesthetized, Lacri-lube® eye lubricating solution (Allergan, Plc; Parsippany-Troy Hills, NJ, U.S.A.) was applied to prevent orbital desiccation during surgery.  Once the animals had achieved a surgical plane of anesthesia (confirmed by failure to elicit hind-limb reflexive contraction in response to robust pinching of the toes on both hind-limbs), animals were prepared for surgery.  Briefly, an area of skin (~2” x 2”) overlying the lower thoracic and lumbar spinal cord region (centred on ~T9-T10) was shaved, followed by sterilization with alternating washes of povidone-iodine (Betadine®, Purdue Products L.P.; Stamford, CT, U.S.A.) and 70% ethanol (3 in total).  Finally, animals were administered bilateral injections of lactated Ringer’s solution (1 ml/kg; s.q.) into the thoracic region above the hind-limbs (to prevent dehydration during the surgical procedure), as well as a single injection of buprenorphine (a semi-synthetic opioid derivative of thebaine) (1 mg/kg; s.q.) as a pre-emptive pain control measure.  Once prepared, animals were transferred under continuous isoflurane anesthesia to the operating table.  Using a sterile scalpel blade, an incision was made in the skin overlying the thoracic spinal cord, the back musculature overlying the spine was split at the midline, and a T9-T10 dorsal laminectomy was completed to expose the 9th and 75  10th thoracic spinal cord segment.  The exposed spinal cord was then impacted with the Infinite Horizons (IH) spinal cord injury impactor (Precision Systems & Instrumentation; Fairfax Station, VA, U.S.A.) set to generate a moderately severe spinal cord injury (70 Kdyne).  Once the wound was closed by suturing the back musculature and overlying skin back together, animals were removed from isoflurane and placed in an incubator (37 °C) before being transferred to housing cages (pending sufficient recovery).  Standard mouse diet and HydroGel™ (ClearH2O; Westbrook, ME, U.S.A.) were placed on the cage bottom to ensure ease of access.  To minimize post-surgical pain, buprenorphrine (1 mg/kg; s.q.) was administered twice daily until 3 dpi, then once daily through 1 wpi.  Animals were also administered lactated Ringer’s solution (1 ml; s.q.) twice daily through 1 wpi to ensure adequate hydration (continued if weight loss or other overt pathological signs were evident).  Monitoring was continued through the post-injury period (i.e. daily weighing, visual inspection, etc.) for signs of unexpected surgical complications (e.g. urinary tract infections, hind-limb autonomy, etc.), accompanied by appropriate intervention when necessary, in accordance with established animal care guidelines.  2.2.4 Animal sacrifice and tissue collection At five time-points during the post-injury period (i.e. 1 dpi, 5 dpi, 2 wpi, and 12 wpi) (fig. 2.1c), animals were sacrificed via an overdose of chloral hydrate (100 mg/kg, i.p.) and trans-cardially perfused with PBS, followed by 4% paraformaldehyde fixative solution.  Entire spinal cords (beginning at C1 and extending caudally to the conus medullaris, including the majority of the cauda equina) were collected, post-fixed in 4% paraformaldehyde overnight, prior to being cryoprotected in subsequent 12%, 18%, and 24% sucrose solutions (each for 24 hours).  Spinal cords were then separated (using a scalpel) into cervical (C1 to C8), thoracic (T1 to 8 mm rostral 76  to the lesion epicentre), lesion epicentre (8 mm rostral to 8 mm caudal to the lesion epicentre), and lumbar segments (8mm caudal to the lesion epicentre to the conus medullaris), with each segment then frozen over dry ice in Tissue-Tek® optimal cutting temperature (O.C.T.) embedding medium (Sakura Finetek USA Inc.; Torrance, CA, U.S.A.) and stored at -80 °C until further processing.  The cervical and lumbar spinal cord segments were then sectioned into 20 µm cross-sections on a cryostat (~-25 °C), collected onto Superfrost Plus® microscope slides (Fisher Scientific; Hampton, NH, U.S.A.) in a serial manner and stored at -80 °C until processing for immunohistochemical analysis.  2.2.5 Immunohistochemical staining  To label specific antigens, spinal cord tissue sections were stained using established immunohistochemical methods.  Frozen spinal cord tissue sections were removed from -80 °C storage and thawed for 1-2 hrs at room temperature.  Once thawed, slides were outlined with ImmunoEdge™ pen (Vector Laboratories; Burlingame, CA, U.S.A.), creating a hydrophobic barrier around the tissue sections to minimize solution run-off in subsequent incubation steps.  Slides were then dried for 20 minutes at room temp (to ensure drying of pen), followed by a 10 min rehydration period in 0.01M PBS.  As many of the targeted antigens in the current study are intracellular (e.g. Olig2) or localized in compact myelin (e.g. MBP), delipidation (i.e. partial removal of membrane lipids) was required to ensure adequate staining.  Delipidation was accomplished by a step-wise series of ethanol solutions of increasing concentration (series: 50%, 70%, 90%, 95%, 100%, 100%, 95%, 90%, 70%, 50%; 2 min each), followed by washing in 0.01M PBS with agitation to remove residual ethanol (10 min; x3).  Slides were then incubated with 10% normal Donkey serum (NDS) (in 0.01M PBS with 0.1% Triton X-100) for 30 min, followed by 77  overnight application (~15-17 hrs) of primary antibodies at room temperature.  The next day, following washing (0.01M PBS; 10 min; x3), secondary antibodies raised in donkey and conjugated to Alexa fluorophores (405 nm, 488 nm, 594 nm, and/or 647 nm) were applied to the tissue sections for 2 hrs at room temperature.  Following washing (0.01M PBS; 10 min; x3), tissue sections were cover-slipped in Fluoromount-G® (Southern Biotech; Birmingham, AL, U.S.A.) and stored at 4 °C until imaging.  To minimize the effect of variation between staining sessions on measured outcomes, all slides used in a given analysis were stained concurrently using the same NDS blocker, primary antibody, and secondary antibody solutions.  2.2.5.1 Primary antibodies The following primary antibodies were used in the study (also see table 1): rabbit anti-Aldh1L1 (Abcam; Cambridge, U.K.; 1:1000), chicken anti-GFP (Abcam; Cambridge, U.K.; 1:400), mouse anti-CD11b (Serotec; Kidlington, Oxford, U.K.; 1:500), mouse anti-CC1 (EMD; Darmstadt, Germany; 1:300), goat anti-PDGFRα (R&D Systems; Minneapolis, MN, U.S.A.; 1:200), rabbit anti-Olig2 (Millipore; Billerica, MA, USA; 1:500), rabbit anti-GFP (Abcam; Cambridge, U.K.; 1:4000), chicken anti-MBP (Aves Labs; Tigard, OR, U.S.A.; 1:200), mouse anti-NF-200 (Sigma-Aldrich; St. Louis, MO, U.S.A.; 1:500), mouse anti-SMI-312 (Covance; Dedham, MA, U.S.A.; 1:1000), rabbit anti-Iba-1 (Wako Chemicals; Cape Charles, VA, U.S.A.; 1:2000); rabbit anti-GFAP (Wako Chemicals; Cape Charles, VA, U.S.A.; 1:500); chicken anti-P0 (Aves Labs; Tigard, OR, U.S.A.; 1:1000), guinea pig anti-NeuN (Millipore; Billerica, MA, USA; 1:1000); rabbit anti-Gst-π (MBL International; Woburn, MA, U.S.A.; 1:300), rabbit anti-cleaved caspase-3 (BD Pharmingen; San Diego, CA, U.S.A.; 1:500), rabbit anti-Ki-67 (Abcam; Cambridge, U.K.; 1:400), goat anti-Glut-1 (Santa Cruz Biotechnology; Dallas, TX, U.S.A.; 1:500), mouse anti-SMI-32 78  (BioLegend; San Diego, CA, U.S.A.; 1:1000), rabbit α-SMA (Abcam; Cambridge, U.K.; 1:500), rat anti-GFP (Nacalai Tesque; Kyoto, Japan; 1:1000), rabbit anti-FSP-1 (Millipore; Billerica, MA, USA; 1:500), rabbit anti-degraded MBP (Chemicon; Temecula, CA, U.S.A.; 1:2000); rabbit anti-SCG-10 (Novus Biologicals; Littleton, CO, U.S.A.; 1:1000).  Table 1. Primary antibodies.  (Columns L-R) Antibody target, structure/cell type of interest, host species of the antibody, source company, catalog number (#), and working concentration.  79  2.2.5.2 Secondary antibodies The following fluorescent secondary antibodies were used in the study (also see table 2): Dylight® 405 nm-conjugated donkey anti-goat (Jackson Immunoresearch Laboratory; West Grove, PA, U.S.A.; 1:200), Dylight®  405 nm-conjugated donkey anti-chicken (Jackson Immunoresearch Laboratory; West Grove, PA, U.S.A.; 1:200), Dylight® 405 nm-conjugated donkey anti-rabbit (Jackson Immunoresearch Laboratory; West Grove, PA, U.S.A.; 1:200), Dylight® 405 nm-conjugated donkey anti-mouse IgG2B (Jackson Immunoresearch Laboratory; West Grove, PA, U.S.A.; 1:200), Alexa Fluoro® 488 nm-conjugated donkey anti-chicken (Jackson Immunoresearch Laboratory; West Grove, PA, U.S.A.; 1:200), Alexa Fluoro® 488 nm-conjugated donkey anti-rabbit (Jackson Immunoresearch Laboratory; West Grove, PA, U.S.A.; 1:200), Alexa Fluoro® 488 nm-conjugated donkey anti-mouse (Jackson Immunoresearch Laboratory; West Grove, PA, U.S.A.; 1:200), Alexa Fluoro® 488-conjugated donkey anti-mouse IgG1 (Jackson Immunoresearch Laboratory; West Grove, PA, U.S.A.; 1:200), Alexa Fluoro® 488 nm-conjugated donkey anti-rat (Jackson Immunoresearch Laboratory; West Grove, PA, U.S.A.; 1:200), Alexa Fluoro® 594 nm-conjugated donkey anti-rabbit (Jackson Immunoresearch Laboratory; West Grove, PA, U.S.A.; 1:200), Alexa Fluoro®  594 nm-conjugated donkey anti-mouse (Jackson Immunoresearch Laboratory; West Grove, PA, U.S.A.; 1:200), Alexa Fluoro® 594 nm-conjugated donkey anti-goat (Jackson Immunoresearch Laboratory; West Grove, PA, U.S.A.; 1:200), Alexa Fluoro® 647-conjugated donkey anti-rabbit (Jackson Immunoresearch Laboratory; West Grove, PA, U.S.A.; 1:200), Alexa Fluoro®  647 nm-conjugated donkey anti-mouse (Jackson Immunoresearch Laboratory; West Grove, PA, U.S.A.; 1:200), Alexa Fluoro® 647 nm-conjugated donkey anti-goat (Jackson Immunoresearch Laboratory; West Grove, PA, U.S.A.; 1:200), Alexa Fluoro® 647 nm-conjugated donkey anti-guinea pig (Jackson Immunoresearch Laboratory; West Grove, PA, 80  U.S.A.; 1:200), Alexa Fluoro® 647 nm-conjugated donkey anti-chicken (Jackson Immunoresearch Laboratory; West Grove, PA, U.S.A.; 1:200).  Table 2. Secondary antibodies. (Columns L-R) emission wavelength of attached fluorophore, host species of the antibody, species specificity (i.e. primary antibody recognition), source company, catalog number (#), and working concentration.   2.2.6 Oil Red O staining To aid in the visualization of myelin debris, cervical spinal cord sections were stained with Oil Red O stain (a.k.a. Solvent Red 27, Sudan Red 5B; C26H24N4O), a lysochrome diazo dye used to stain neutral triglycerides and lipids (Proescher 1927; Cholewiak et al. 1968), following an established protocol (Ramirez-Zacarias et al. 1992; Prophet 1992).  Importantly, Oil Red O 81  differentially stains intact myelin sheaths on axons (seen as a pale pink/red colour) and myelin debris (dark purple), thus allowing for the identification of myelin debris in tissue sections (Perry et al. 1987; Prineas et al. 1993; Mickel et al. 2016).  Briefly, tissue slides were removed from -80 °C storage and thawed for at least 1 hr at room temperature.  Slides were then washed in 0.01M PBS with 0.25% Triton X-100 for 15 min, followed by incubation in the Oil Red O working solution (60% Oil Red O stock, 40% dH2O, then filtered) for 30 min, followed by washing in running tap water for 10 min (note: ions in the tap water are required for the reaction).  Following washes in 0.01M PBS (3x; 1 min), slides were cover-slipped with Fluoromount-G® (Southern Biotech; Birmingham, AL, U.S.A.) and stored at 4 °C until imaging.  2.2.7 Imaging Immunofluorescence of antigen-bound secondary antibodies on spinal cord tissue sections was visualized using a Zeiss Axio Observer Z1 confocal microscope (Carl Zeiss AG; Oberkochen, Germany) equipped with a Yokogawa X-1 Spinning Disk (Yokogawa; Tokyo, Japan), controlled using the Zen® Blue software package (Carl Zeiss AG, Oberkochen, Germany), which enabled the assessment of co-localization between 4 channels, corresponding to 4 different emission wavelengths of secondary antibodies (i.e. 405 nm, 488 nm, 594 nm, and 647 nm) at up to 63x optical magnification.  2.2.8 Cervical and lumbar spinal cord section sampling protocol For all analyses, three sequential sections spaced 400 µm apart from the cervical spinal cord (centred on C4) and from the lumbar spinal cord (centred on L4) per animal were analyzed.  To ensure consistency in sampling location between animals, the Allen Brain Institute’s (Seattle, WA, 82  U.S.A.) atlas of the mouse spinal cord (accessed through mousespinal.brain-map.org) was utilized to determine the section in each animal that corresponded to the desired spinal cord level (i.e. either C4 or L4) based on absolute distances (possible due to serial sectioning of the spinal cord segments).  Analysis was then carried out on this section, as well as the two sections located 400 µm rostral and caudal.        Figure 2.2 Tissue sampling approach.  a) 3 sections per animal, spaced 400 µm apart were sampled from the cervical (centred on ~C4) and lumbar (centred on ~L4) spinal cord (injured at T9/T10).  b) Subsequent analyses (outlined in the text) were performed on these sections to address the research question of remote changes in oligodendrocyte lineage cells following thoracic contusion injury.                     2.2.9 Image position sampling: whole-cord section analyses Image positions for whole-cord section analyses were sampled from spinal cord sections according to established unbiased stereological procedures (West, 2012).  First, a preview image of the cord section was taken at 40x magnification.  Once the whole cord section could be visualized, the tissue was outlined using the free-form tool in the Zen® Blue software package (Carl Zeiss AG; 83  Oberkochen, Germany), followed by the overlaying of a grid across the outlined tissue section (providing a template to ensure regularly spaced sampling).  A random number generator (Randomness and Integrity Services Ltd.; Dublin, Ireland; accessed through www.random.org) was utilized to generate random starting coordinates from which image positions were then assigned in a systematically random manner (using the overlain grid as a template).  30-40 image positions were sampled from each cord section for cell counting analyses (i.e. OPC proliferation, oligodendrocyte cell densities, oligodendrogenesis, and oligodendrocyte maturation) (at 40x magnification) and 60-70 images per cord section for the new myelination analysis (at 63x magnification).  Z-stack depth was set to 20 µm (i.e. the thickness of the tissue section) using multiple channels as a reference (to compensate for the variable tissue penetration of different antibodies).  The interval between subsequent z-stack planes was set according to the feature of interest and the required resolution (e.g. for cell counting it was set at 1.0 µm, for myelin sheath counting it was set at 0.5 µm to provide additional resolution).  2.2.10 Image position sampling: fasciculus gracilis analyses Image positions for fasciculus gracilis vs. fasciculus cuneatus analyses were sampled from cervical spinal cord sections using a modified protocol as that used for whole cord section analyses.  Axonal stains (i.e. NF-200/SMI-312) were used to differentiate between the fasciculus gracilis and fasciculus cuneatus in cervical spinal cord sections (3 sections per animal centred on C4) based on expected murine spinal cord anatomy (Watson & Harrison 2012).  Once delineated, the fasciculus gracilis and fasciculus cuneatus were outlined using the free-form drawing tool in the Zen® Blue software package (Carl Zeiss AG; Oberkochen, Germany).  A grid was then overlaid on the respective regions and 2-3 positions in the fasciculus gracilis (along the spinal cord midline) 84  and 4 positions in the fasciculus cuneatus (2 on each side of the midline, placed roughly in the middle of the fasciculus cuneatus, well away from the fasciculus gracilis border) were systematically assigned.  Z-stack depth and interval were set as described above.                           2.2.11 Cell counting protocol For analyses that involved the counting of desired cell types within the sample images (i.e. OPC recombination efficiency, OPC proliferation, oligodendrocyte lineage cell densities, oligodendrogenesis, oligodendrocyte maturation, and Iba-1+ cell density analyses), unbiased stereological procedure was followed (West, 2012).  Briefly, using the Zen® Blue software package (Carl Zeiss AG; Oberkochen, Germany) desired cell types were quantified in sample images.  A square counting frame of consistent area (comprising ~85% of the total image area) was overlain on the image, delineating the counting area.  All objects meeting the requirements for designation as the desired cell type (i.e. morphology, antigenic expression, etc.) completely within the counting frame were included, as were objects transected by the upper and right borders of the counting area (objects transected by the lower and left border were excluded).  Furthermore, objects transected by the upper z-stack plane were included, whereas those transected by the lower z-stack plane were excluded from the analysis.  Co-localization between immunofluorescent stains of interest was confirmed by movement through the z-stack planes, as well as orthogonal projection images (where required).  Cell count data is either expressed as a density (i.e. cells/mm3) or as a percentage of a specific cell population (e.g. % of all committed oligodendrocytes that are newly-formed).  85  2.2.12 Myelin sheath counting protocol For analyses that involved the counting of myelin sheaths (i.e. new myelination analysis), the same unbiased stereological procedures as outlined above were followed (West, 2012), but with subtle modification.  As myelin sheaths are smaller than cell bodies (in cross-section), images were taken at 63x magnification with z-stack intervals set to 0.5 µm.  Furthermore, as individual myelin sheaths are not expected to be wholly contained within a single tissue section (as they can be >200 µm in length) (Young et al. 2013), exclusion criteria based on the transection of the object by the top or bottom z-stack plane were not followed.  Instead, starting at the centre z-stack plane, objects potentially representing newly formed sheaths (i.e. mGFP+/MBP+ sheaths around axons) were identified.  Movement through the z-stack and/or orthogonal projection images confirmed or rejected this tentative designation, based on morphology, co-localization between mGFP and MBP, and anatomical relation to axons.   Myelin sheath counting data is presented as a density (sheaths/mm2).   2.2.13 Immunoreactivity (IR) thresholding protocol For analyses that involved immunoreactivity thresholding (i.e. Iba-1 and GFAP immunoreactivity analyses), established protocols were followed, as described previously (Cardona et al. 2006).  For the whole-cord section analyses, a preview image of the whole cord section was first taken at 20x magnification.  Once the whole cord section could be visualized, the tissue was outlined using the free-form tool in the Zen® Blue software package (Carl Zeiss AG; Oberkochen, Germany), ensuring that the entirety of the spinal cord tissue was included.  A tiled image (10% overlap between adjacent tiles) with z-stacks (depth: 20 µm, interval: 1 µm) was then taken of the whole cord section.  For the fasciculus gracilis vs. fasciculus cuneatus analyses, images were sampled 86  using the same method as outlined above for the fasciculus gracilis vs. fasciculus cuneatus cell counting analyses.  Images were taken at 40x magnification with z-stacks (depth: 20 µm, interval: 1 µm) at the sampled positions.  All images for a particular immunoreactivity thresholding analysis were taken with the same camera settings (i.e. exposure, EM gain, etc.), microscope settings, and were post-processed in identical manners.  Images for both the whole-cord section and fasciculus gracilis vs. fasciculus cuneatus analyses were identically processed.  First, maximum projection images were generated from the original image files (this collapses all of the z-planes into a single plane) in the Zen® Blue software package (Carl Zeiss AG; Oberkochen, Germany), followed by image export (uncompressed TIFF).  Images were then further processed in Microsoft® Paint (Microsoft; Seattle, WA, U.S.A.) to ensure inclusion of only the region of interest in the thresholding (for whole-cord section analyses this was the whole cord, excluding the peripheral roots, dura mater, etc.; for fasciculus gracilis vs. fasciculus cuneatus analyses this was the image area) (using the free-form tool and crop function).  Processed images were then opened in ImageJ (NIH; Washington, D.C., U.S.A.) for thresholding analysis.  Thresholding in ImageJ was performed as outlined previously (Cardona et al. 2006).  In brief, images were converted to 8-bit (‘Image’ » ‘Type’ » ‘8-bit’), followed by the manual thresholding of an overlain masking layer (‘Adjust’ » ‘Threshold’).  Threshold levels were manually set based on morphological features of the elements of interest (e.g. microglial cell processes).  Once the threshold level was set, the total area in pixels of the overlain layer was calculated (‘Analyze’ » ‘Analyze Particles’), which is a measure of the area covered by the stain of interest (e.g. Iba1 or GFAP).  The total area of the cord section or image was then calculated, to compensate for variations in cord size between animals.  Immunoreactivity thresholding data is presented as a percentage of total cord/image area (e.g. % of total area that is Iba1+). 87   2.2.14 Axonal number indices As counting individual axons is highly time consuming, labour intensive, and potentially lacks consistency and/or accuracy, axon numbers were estimated using an automated approach described previously (Sathyanesan et al. 2012; Tobin et al. 2014), with subtle modification.  In brief, images at 63x magnification with z-stacks (depth: 20 µm, interval: 1.0 µm) were sampled from ventrolateral and dorsal white matter regions of spinal cord sections stained for a combination of NF-200 and SMI-312 (pan-axonal stains), using the same systemic random sampling procedure as outlined above.  Images were taken using the same camera settings (i.e. exposure, EM gain, etc.), microscope settings, and were post-processed in identical manners.  First, maximum projection images were generated from the original image files using the Zen® Blue software package (Carl Zeiss AG; Oberkochen, Germany), followed by image export (uncompressed TIFFs).  Exported images were then processed further using ImageJ (NIH; Richmond, VA, U.S.A.).  In brief, images were converted to 8-bit (‘Image’ » ‘Type’ » ‘8-bit’), followed by the manual thresholding of an overlain masking layer (‘Adjust’ » ‘Threshold’) based upon expected axonal morphology.  Once the masking layer was overlain, the number of particles in the image area was calculated (‘Analyze’ » ‘Analyze Particles’).  To limit inclusion of particles to the appropriate diameter for axons, the size range was set to 0.5-3 µm, inclusively, based on axonal diameters in the mouse spinal cord (Ong et al. 2008).  This generated an axonal density index (i.e. number of particles per image area).  To estimate the total number of axons in each white matter area (i.e. ventrolateral and dorsal WM), axonal density indices were multiplied by white matter area indices (described below).  Axon number index data is presented as an estimated total number of axons.  88   2.2.15 White matter area indices For the assessment of white matter atrophy, tiled images (10% overlap between adjacent tiles) with z-stacks (depth: 20 µm, interval: 1.0 µm) of whole cord sections immunostained for MBP and a combination of NF-200 and SMI-312 were taken at 20x magnification.  Maximum projection images were then generated from the original image files in the Zen® Blue software package (Carl Zeiss AG; Oberkochen, Germany), followed by image export (uncompressed TIFFs).  Exported images were processed in Microsoft® Paint (Microsoft; Seattle, WA, U.S.A.) to include only the white matter regions of interest (i.e. ventrolateral and dorsal WM), guided by MBP and axonal stains (using the free-form tool and crop function).  Processed image files were then opened in ImageJ (NIH; Richmond, VA, USA) and the area of the WM regions and total cord section was measured.  To assess for injury-induced white matter atrophy, the area of specific white matter regions was normalized to total cord area, presented as a white matter area index.  2.2.16 Statistical analysis All data are presented as mean ± SEM.  For all analyses statistical significance was set at p<0.05.  As the sample sizes used in the current study were of insufficient number to perform the Shapiro-Wilk, Kolmogorov-Smirnov, or the D’Agostino and Pearson Omnibus normality tests (i.e. n<5 for all groups), we exclusively utilized non-parametric tests (i.e. the Mann Whitney-U test) to assess statistical significance.  All statistical analyses were performed using the GraphPad® Prism 5.01 software package (GraphPad; San Diego, CA, U.S.A.).    89  2.2.16.1 Whole-cord section analyses OPC recombination efficiency and Olig2+ recombined cell percentage data was assessed using the Mann Whitney-U test (12 wpi vs. uninjured).  Iba-1 immunoreactivity and Ki-67+ cell density data was assessed using the Mann Whitney-U test (1 dpi vs. uninjured, 5 dpi vs. uninjured, 2 wpi vs. uninjured, 12 wpi vs. uninjured).  β-APP+ profile number, GFAP immunoreactivity, axonal number index, and white matter area index data was assessed using the Mann Whitney-U test (12 wpi vs. uninjured).  OPC proliferation and oligodendrogenesis data was assessed using the Mann Whitney-U test (1 dpi vs. uninjured, 5 dpi vs. uninjured, 2 wpi vs. uninjured, 12 wpi vs. uninjured).  Oligodendrocyte lineage cell density, new myelination, and oligodendrocyte maturation data was assessed using the Mann Whitney-U test (12 wpi vs. uninjured).    2.2.16.2 Fasciculus gracilis analyses OPC proliferation, Iba-1 immunoreactivity, and Iba-1+ cell density data was assessed using the Mann Whitney-U test (1 dpi vs. uninjured, 5 dpi vs. uninjured, 2 wpi vs. uninjured, 12 wpi vs. uninjured).  Oligodendrogenesis, new myelination, β-APP+ profile number, GFAP immunoreactivity data was assessed using the Mann Whitney-U test (12 wpi vs. uninjured; fasciculus gracilis and fasciculus cuneatus).  2.3 Results 2.3.1 Injury parameters Consistency of injury parameters in pre-clinical SCI models is critical to ensure uniformity of resultant histopathology (Choo et al. 2008; Lam et al. 2014; Chen et al. 2016), particular relevant for the study of endogenous cell responses (e.g. oligodendrocyte lineage cells) as heterogeneity in 90  primary injury (e.g. axonal tract specific damage, cell loss, etc.) or secondary injury processes (e.g. inflammation) may confound injury-induced responses.  Previous utilization of the injury model employed here (i.e. moderate T9-T10 thoracic contusion injury using the IH impactor) reported a high degree of consistency (Choo et al. 2008; Lam et al. 2014; Chen et al. 2016).  Comparison of injury parameters between animals of a given genotype revealed minimal variation in injury parameters, as measured by the IH impactor sensor.  For example, PDGFRα-CreERT2:ROSA26-mGFP animals received injuries with largely consistent force (71.3 ± 0.67 Kdynes) (fig. 2.3c), impactor head velocity at impact (126.5 ±2.13 m/s) (fig. 2.3d), and displacement of the impactor head on impact (540 ± 29.4 µm) (fig. 2.3e).  Similarly, injury parameter variation was minimal for PDGFRα-CreERT2:ROSA26-YFP animals as well, with consistent force (70.5 ± 0.22 Kdynes) (fig. 2.3c), velocity (124.7 ± 0.80 m/s) (fig. 2.3d), and displacement (625 ± 79.0 µm) (fig. 2.3e).  Importantly, no significant differences in injury force (p=0.493, Mann-Whitney U test), impactor head velocity (p=0.683, Mann-Whitney U test), or impactor head displacement (p=0.335, Mann-Whitney U test) were observed between PDGFRα-CreERT:ROSA26-YFP and PDGFRα-CreERT:ROSA26-mGFP animals.  To confirm that injury consistency correlated into similar resultant histopathology, tissue sections at lesion epicentre (located at T9-T10) were visually assessed for gross histopathological appearance, using markers that enabled visualization of injury-induced changes.  GFAP immunoreactivity delineates the glial scar bordering the lesion site (Tator 1995; Sofroniew 2009), whereas MBP immunoreactivity visualises the extent of demyelination (Lasiene et al. 2008; Powers et al. 2013).  As expected, the extent of tissue loss, lesion size, and lesion location were decidedly comparable between PDGFRα-CreERT:ROSA26-YFP (fig. 2.3a) and PDGFRα-CreERT:ROSA26-mGFP animals (fig. 2.3b), suggesting grossly similar histopathology. 91          Figure 2.3 Lesion epicentre histopathology and injury parameters.  Lesion epicentre (T9-T10) tissue from (a) PDGFRα-CreERT2:ROSA26-YFP (YFP, green; GFAP, red) and (b) PDGFRα-CreERT2:ROSA26-mGFP (mGFP, green; MBP, blue) transgenic mice.  Lesion is outlined.  Injury parameters measured by the IH impactor, including (c) force, (d) velocity, and (e) displacement.  2.3.2 Transgene recombination Transgenic fate-mapping approaches rely on the assumption of effective, but restricted, transgene recombination in the desired cell population (Whitfield 2015).  The transgene constructs used in the current study are designed to restrict YFP and mGFP expression to PDGFRα-expressing cells 92  upon tamoxifen administration.  In the adult murine spinal cord, PDGFRα is expressed by OPCs (Pringle et al. 1992), a subpopulation of vasculature-associated pericytes, and a population of fibroblastic cells in the peripheral roots (Rivers et al. 2008; Kang et al. 2010).  We therefore sought to characterize the efficiency and cell-type restriction of transgene recombination in PDGFRα-CreERT2:ROSA26-YFP (fig. 2.4a) and PDGFRα-CreERT2:ROSA26-mGFP transgenic mouse lines (fig. 2.4b).  These specific transgenic mouse lines, and similar ones, have been previously characterized by other research groups (Rivers et al. 2008; Zawadzka et al. 2010; Kang et al. 2010), as well as employed by our laboratory for fate-mapping studies of OPCs following SCI (Assinck et al. in prep) and toxin-induced CNS demyelination (i.e. cuprizone- and lysolecithin-induced demyelination) (Duncan et al. in prep).  Consistent with previous findings, transgene recombination was highly efficient in spinal cord OPC populations (defined as Olig2+/PDGFRα+ cells) (fig. 2.4c-d) in both injured (12 wpi) (95.4 ± 0.4%; n=4) and uninjured (96.2 ± 0.6%; n=4) PDGFRα-CreERT2:ROSA26-YFP animals, with no significant difference between them (p=0.678, Mann-Whitney U test) (fig. 2.3e).  Furthermore, the majority of recombined cells (i.e. YFP+ cells) belonged to the oligodendrocyte lineage (defined as Olig2+ cells) in injured (12 wpi) (90.1 ± 0.2%; n=4) and uninjured (90.3 ± 0.8%; n=4) PDGFRα-CreERT2:ROSA26-YFP animals, with no significant difference between them (p=0.988, Mann-Whitney U test) (fig. 2.4f).  As expected, the ~10% of recombined cells not belonging to the oligodendrocyte lineage (i.e. Olig2- cells) were comprised of vascular-associated pericytes (defined as recombined cells with tubular morphology in close relation to α-SMA+ mural cells and Glut-1+ blood vessels) (fig. 2.5a-b) and a population of fibroblastic cells in the peripheral roots (defined as recombined cells with fibroblast-like morphology amongst P0+ myelin sheaths) (fig. 2.5c).  Transgene recombination within these non-oligodendrocyte lineage cell populations has been previously reported by our research group and 93  others (Kang et al. 2010; Assinck et al. in prep).  In theory, recombination only occurs upon tamoxifen administration (as the CreERT2 fusion protein should be restricted to the cytoplasm in the absence of ligand binding).  However, many CreERT2-based systems display ‘leaky’ recombination (i.e. transgene recombination in the absence of tamoxifen) (Kemp et al. 2004).  This is particularly relevant for fate-mapping studies that assume the temporal restriction of transgene recombination to the tamoxifen dosing period.  Therefore, we sought to assess the degree of spontaneous recombination in non-tamoxifen treated PDGFRα-CreERT2:ROSA26-YFP and PDGFRα-CreERT2:ROSA26-mGFP mice.  As expected, based on previous characterization of these transgenic mouse lines (Kang et al. 2010; Assinck et al. in prep), there was minimal spontaneous recombination in the spinal cord, as revealed by the absence of recombined cells (i.e. YFP+ or mGFP+) (data not shown).  Collectively, these findings confirm the efficacy of the PDGFRα-CreERT2:ROSA26-YFP and PDGFRα-CreERT2:ROSA26-mGFP transgenic mouse lines as a reliable system in which to assess oligodendrocyte lineage cell responses to SCI. 94                       Figure 2.4 Transgene recombination in oligodendrocyte lineage cells.  Recombined cells (i.e. YFP+ or mGFP+) in spinal cord tissue from (a) PDGFRα-CreERT2:ROSA26-YFP (YFP, green) and (b) PDGFRα-CreERT2:ROSA26-mGFP (mGFP, green) transgenic mice.  c) Recombined oligodendrocyte precursor cell (OPC) in PDGFRα-CreERT2:ROSA26-YFP tissue (i.e. a YFP+/Olig2+/PDGFRα+ cell) (YFP, green; Olig2, white; PDGFRα, blue).  d) 3-D rendering of a recombined cell in PDGFRα-CreERT2:ROSA26-mGFP tissue with OPC morphology (mGFP, green; Hoechst, blue).  e) OPC recombination efficiency in injured (12 wpi) (orange) and uninjured (blue) PDGFRα-95  CreERT2:ROSA26-YFP tissue.  f) Percentage (%) of recombined cells that belong to the oligodendrocyte lineage (defined as Olig2+) in injured (12 wpi) (orange) and uninjured (blue) PDGFRα-CreERT2:ROSA26-YFP tissue.               Figure 2.5 Transgene recombination in non-oligodendrocyte lineage cells.  a) Recombined vasculature-associated pericytes in PDGFRα-CreERT2:ROSA-YFP tissue (defined as recombined cells in close-association with Glut-1+ blood vessels and α-SMA+ mural cells) (YFP, green; α-SMA, red; Glut-1, white).  b) 3D-rendering of a putative recombined vasculature-associated pericyte in PDGFRα-CreERT2:ROSA-mGFP tissue (mGFP, green; Hoechst, blue).  c) Recombined peripheral root cells in PDGFRα-CreERT2:ROSA-mGFP tissue (mGFP, green; FSP-1, blue; P0, red).  2.3.3 Histopathology: whole-cord section analyses As pathophysiological responses remote to a SCI lesion site are less well characterized than for the lesion epicenter (Tator 1995; Tator & Fehlings 1991), and since OPCs are highly responsive to perturbations in extracellular conditions (Buffo et al. 2005; McTigue et al. 2001; Zhang et al. 96  2013), we sought to characterize injury-induced pathological changes in the cervical and lumbar spinal cord (i.e. sampling locations).  We focused here on common CNS pathological features, specifically microgliosis, astrogliosis, axonal damage/loss, and tissue atrophy (Tator 1995; David & Kroner 2011).  Initially, we performed whole-cord section analyses with the notable caveat of sampling from both spared and injured axonal tracts.  Evident cervical microgliosis, as revealed by increased Iba-1 immunoreactivity above uninjured levels (expressed as % of total cord area) was observed as early as 1 dpi (p=0.0286, Mann Whitney-U test; n=4), continuing through 5 dpi (p=0.0286, Mann Whitney-U test; n=4), 2 wpi (p=0.0286, Mann Whitney-U test; n=4), and 12 wpi (p=0.0286, Mann Whitney-U test; n=4) (fig. 2.6a-c).  In contrast, lumbar Iba-1 immunoreactivity did not increase above uninjured levels until 5 dpi (p=0.0286, Mann Whitney-U test; n=4), but remained elevated through 2 wpi (p=0.0286, Mann Whitney-U test; n=4) and 12 wpi (p=0.0286, Mann Whitney-U test; n=4) (fig. 2.6d-f).  Clusters of cells with increased Iba1-immunoreactivity and activated microglial morphology (i.e. amoeboid-like soma with thick, short processes) were observed in both the injured cervical and lumbar spinal cord (fig. 2.6g-h).  Cervical microgliosis was largely constricted to the fasciculus gracilis, as well as peripheral edges of the dorsolateral white matter, which anatomically corresponds to the dorsal spinocerebellar tract and dorsolateral spinothalamic tract (Watson & Harrison 2012).  In contrast, lumbar microgliosis was concentrated around the peripheral rim of the ventrolateral white matter, corresponding to the anatomical location of the lateral spinothalamic tract, ventral spinocerebellar tract, lateral vestibulospinal tract, rostral reticulospinal tract, and ventral spinothalamic tract (Watson & Harrison 2012).  To assess for axonal damage, tissue sections were stained for β-amyloid precursor protein (β-APP) (fig. 2.7a-d).  Accumulation of β-APP in axons is commonly interpreted as being representative of axonal transport impairment resulting from damage (Cornish et al. 2000; Li et al. 1995).  Indeed, punctate 97  β-APP staining is widely used to demonstrate diffuse axonal damage in models of traumatic brain injury (TBI) and other CNS pathologies (Choo et al. 2008).  β-APP+ axonal profiles were substantially elevated at 12 wpi in both the cervical (p=0.0238, Mann Whitney-U test; n=4) and lumbar spinal cord (p=0.0238, Mann Whitney-U test; n=4) (as compared to uninjured levels) (fig. 2.7e), consistent with significant axonal damage.  β-APP+ staining displayed similar distribution as Iba-1 immunoreactivity.  For example, in the cervical spinal cord β-APP+ axonal profiles were concentrated in the fasciculus gracilis, as well as peripheral edges of the dorsolateral white matter (fig. 2.7a), whereas in the lumbar spinal cord they were concentrated around the peripheral rim of the ventrolateral white matter (fig. 2.7c).  The overlapping distributions of β-APP and Iba-1 in both the cervical and lumbar spinal cord is consistent with microglial activation along degenerating/damaged axonal tracts, as observed in axotomy (Neilson et al. 2006), SCI (Rabchevsky et al. 2005), dorsal rhizotomy (Sun et al. 2010), and ALS (Kang et al. 2013) models.  An important caveat is that punctate β-APP+ profiles may be alternatively interpreted as axonal debris (either extracellular or internalized by phagocytic microglia).  However, regardless of interpretation as intact axons or debris, β-APP accumulation is consistent with axonal injury.  Distal axon fragments undergo degeneration (albeit slowly in the CNS) (Vargas & Barres 2007), however estimation of dorsal and ventrolateral white matter axonal numbers did not reveal overt axonal loss in either the cervical or lumbar spinal cord (fig. 2.7f).  Similarly, atrophy of dorsal and ventrolateral white matter regions was not evident (fig. 2.7g), although the possibility of overall cord atrophy is not excluded.  Assessment of GFAP immunoreactivity, as a measure of astrogliosis, did not reveal any whole-cord increases at 12 wpi as compared to uninjured levels (fig. 2.8a-f).  Traumatic injury to the adult murine spinal cord elicits proliferation in multiple endogenous cell populations (e.g. NSPCs, microglia, astrocytes, OPCs) (Horky et al 2006; Horner et al. 2000).  98  Quantification of Ki-67+ nuclei density (representing actively proliferating cells) revealed increased cervical proliferation above uninjured levels (471 ± 40.7 cells/mm3; n=4) at 1 dpi (1065 ± 101 cells/mm3; n=4; p=0.0286, Mann Whitney-U test), 5 dpi (2267 ± 516 cells/mm3; n=4; p=0.0286, Mann Whitney-U test) and 2 wpi (1747 ± 102 cells/mm3; n=4; p=0.0286, Mann Whitney-U test), before returning to uninjured levels by 12 wpi (366 ± 28 cells/mm3; n=4; p=0.0286, Mann Whitney-U test) (fig. 2.9a-c).  Conversely, in the lumbar spinal cord a spike in cell proliferation well above uninjured levels (427 ± 102 cells/mm3; n=4) was observed at 1 dpi (8250 ± 889 cells/mm3; n=4; p=0.0286, Mann Whitney-U test), with reduced, but still elevated, proliferation continuing through 5 dpi (1994 ± 377 cells/mm3; n=4; p=0.0286, Mann Whitney-U test) and 2 wpi (2375 ± 503 cells/mm3; n=4; p=0.0286, Mann Whitney-U test), before returning to uninjured levels at 12 wpi (892 ± 149 cells/mm3; n=4; p=0.0571, Mann Whitney-U test) (fig. 2.9d-f). 99      Figure 2.6 Microgliosis in the cervical and lumbar spinal cord.  Cervical microglial activation in (a) injured (12 wpi) and (b) uninjured PDGFRα-CreERT2:ROSA26-YFP tissue (Iba-1, red).  Outlined areas represent regions of microgliosis.  c) Quantification of Iba-1 immunoreactivity in the cervical spinal cord at 1 dpi, 5 dpi, 2 wpi, and 12 wpi (orange), compared to uninjured controls (blue).    Lumbar microglial activation in (d) injured (12 wpi) and (e) uninjured PDGFRα-CreERT2:ROSA26-YFP tissue (Iba-1, red).  Outlined areas represent regions of microgliosis.  f) Quantification of Iba-1 immunoreactivity in the lumbar spinal cord at 1 dpi, 5 dpi, 2 wpi, and 12 wpi (orange), compared to uninjured controls (blue).  (g) Microglia in the injured spinal cord display typical characteristics of activated microglia, including short, thick processes and higher density (h) whereas microglia in the uninjured spinal cord have long slender processes and lower density, characteristic of ‘resting’ ramified microglia (Iba-1, red; Hoechst, blue).  100   Figure 2.7 Axonal pathology in the cervical and lumbar spinal cord.  β-APP immunoreactivity in the (a, b) cervical and (c, d) lumbar spinal cord of PDGFRα-CreERT2:ROSA26-YFP (β-APP, turquoise).  Outlined areas represent regions of axonal damage.  Note close spatial correlation of axonal damage patterns with microglial activation (fig. 2.6).  e)  Quantification of β-APP+ axonal profiles in the cervical and lumbar spinal cord at 12 wpi (orange) compared to uninjured controls (blue).  Estimation of ventrolateral and dorsal white matter (f) axonal number and (g) area (as a percentage of cord cross-sectional area) at 12 wpi (orange) and uninjured controls (blue).    101                    Figure 2.8 No significant astrogliosis across the whole-cord section in the cervical and lumbar spinal cord.  Cervical astrogliosis in (a) injured (12 wpi) and (b) uninjured PDGFRα-CreERT2:ROSA26-mGFP tissue (GFAP, white).  c)  Quantification of GFAP immunoreactivity in the cervical spinal cord at 12 wpi (orange) and uninjured controls (blue).  Lumbar astrogliosis in (d) injured (12 wpi) and (e) uninjured PDGFRα-CreERT2:ROSA26-mGFP tissue (GFAP, white).  f) Quantification of GFAP immunoreactivity in the lumbar spinal cord at 12 wpi (orange) and uninjured controls (blue).   102                      Figure 2.9 Cell proliferation in the cervical and lumbar spinal cord.  Cervical cell proliferation in (a) injured (5 dpi) and (b) uninjured PDGFRα-CreERT2:ROSA26-YFP tissue (Ki-67, red; Hoechst, blue).  c) Quantification of Ki-67+ nuclei density in the cervical spinal cord at 1 dpi, 5 dpi, 2 wpi, and 12 wpi (orange) and uninjured controls (blue).  Lumbar cell proliferation in (d) injured (1 dpi) and (e) uninjured PDGFRα-CreERT2:ROSA26-YFP tissue (Ki-67, red; Hoechst, blue).  f) Quantification of Ki-67+ nuclei density in the lumbar spinal cord at 1 dpi, 5 dpi, 2 wpi, and 12 wpi (orange) and uninjured controls (blue).               103  2.3.4 OPC proliferation: whole-cord section analyses Robust and rapid OPC proliferation is characteristic of a variety of CNS pathologies (e.g. SCI, neurodegeneration, cortical stab injury, ischemic stroke, etc.) (Buffo et al. 2005; McTigue et al. 2001; Zhang et al. 2013).  The OPC proliferative response following contusion SCI has been thoroughly characterized at lesion epicentre (McTigue et al. 2001), but whether remote OPC populations respond similarly is largely unknown.  To assess OPC proliferation remote to the lesion site, cervical and lumbar spinal cord sections were stained for the proliferation marker Ki-67.  Ki-67 expression is restricted to the cell cycle (i.e. G1, S, G2, and M phases) (Scholzen & Gerdes 2000), thus is commonly used as a marker that labels cells actively proliferating at the time of tissue collection, as opposed to thymidine analogs (e.g. BrdU, EdU) that label all cells that proliferated during the administration period.  Due to incompatibility between available Olig2 and Ki-67 antibodies we were unable to assess co-localization, precluding the definitive identification of the proliferating cells as members of the oligodendrocyte lineage (defined by Olig2 expression).  Therefore, we relied upon the demonstration that recombined cells are predominately of the oligodendrocyte lineage and that recombined pericytes are associated with Glut-1+ vasculature (Krueger & Bechmann 2009).  Quantification of the percentage of YFP+/PDGFRα+ cells with OPC morphology (i.e. extensive, branched arbours of processes extending into the parenchyma) that were Ki-67+ (fig. 2.10a) revealed significant injury-induced proliferation above uninjured levels (6.0 ± 0.7%; n=4) in the cervical spinal cord starting at 5 dpi (17.2 ± 1.8%; p=0.0286, Mann Whitney-U test; n=4), continuing through 2 wpi (17.9 ± 4.1%; p=0.0286, Mann Whitney-U test; n=4), but returning to uninjured levels by 12 wpi (6.5 ± 1.1%; p=0.886, Mann Whitney-U test; n=4) (fig. 2.10b).  By contrast, in the lumbar spinal cord there was increased proliferation of YFP+/PDGFRα+ cells above uninjured levels (7.1 ± 1.5%; n=4) at 1 dpi (24.9 ± 3.9%; p=0.0286, 104  Mann Whitney-U test; n=4), remaining elevated through 5 dpi (21.6 ± 4.1%; p=0.0286, Mann Whitney-U test; n=4) and 2 wpi (27.9 ± 4.3%; p=0.0286, Mann Whitney-U test; n=4), prior to returning to uninjured levels by 12 wpi (12.9 ± 1.9%; p=0.114, Mann Whitney-U test; n=4) (fig. 2.10c).  Interestingly, there was a non-statistical trend for a more robust proliferative response in the lumbar spinal cord as compared to the cervical spinal cord at 1 dpi, 5 dpi, and 12 wpi.  Despite ~90% of recombined cells belonging to the oligodendrocyte lineage (i.e. Olig2+), and the demonstration of OPC morphology among the proliferating cells, it is feasible that this injury-induced increase in Ki-67+/PDGFRα+/YFP+ cells represents pericyte proliferation instead of OPC proliferation.  To address this possibility, we assessed the proliferation of vascular-associated recombined cells (i.e. pericytes) remote to the lesion.  Cervical and lumbar spinal cord sections were stained for Ki-67 (proliferating cells) and Glut-1 (endothelial cells of blood vessels).  Quantification of the density of Ki-67+ cells either associated with Glut-1+ blood vessels (representing pericytes) or not associated with blood vessels (representing OPCs) revealed no examples of Ki-67+ recombined cells associated with Glut-1+ blood vessels with pericyte morphology (i.e. tubular) at 1 dpi, 5 dpi, 2 wpi, 12 wpi, or in uninjured animals in either the cervical or lumbar spinal cord (fig. 2.10d-f).  Of particular importance was the absence of Ki-67+ vasculature-associated recombined cells at time-points shown to have robust proliferation of PDGFRα+/YFP+ cells (i.e. 5 dpi and 2 wpi in the cervical spinal cord, and 1 dpi, 5 dpi, and 2 wpi in the lumbar spinal cord) (fig. 2.10e-f).  Therefore, the recombined cells observed to proliferate remote to the lesion site represent non-vasculature-associated PDGFRα+ recombined cells with OPC morphology, which are most likely OPCs.  In summary, thoracic contusion injury induces robust OPC proliferation remote to the lesion site in both the cervical and lumbar spinal cord.  105                        Figure 2.10 OPC proliferation in the cervical and lumbar spinal cord.  a) PDGFRα+ recombined cell proliferation in PDGFRα-CreERT2:ROSA26-YFP tissue (YFP, green; Ki-67, red; Hoechst, blue; PDGFRα, white).  Quantification of PDGFRα+ recombined cell proliferation in the (b) cervical and (c) lumbar spinal cord at 1 dpi, 5 dpi, 2 wpi, and 12 wpi (orange) compared to uninjured controls (blue).  d) Non-proliferative vasculature-associated recombined cells (i.e. pericytes) in PDGFRα-CreERT2:ROSA26-YFP tissue (YFP, green; Ki-67, red; Hoechst, blue; Glut-1, white).  Quantification of non-vasculature-associated (NV) and vasculature-associated (VAS) recombined cell proliferation in the e) cervical and f) lumbar spinal cord at 1 dpi, 5 dpi, 2 wpi, and 12 wpi (orange) and uninjured controls (blue).  106   2.3.5 Oligodendrocyte lineage cell densities: whole-cord section analyses OPCs in the adult CNS serve as precursors for myelinating oligodendrocytes (Dimou et al. 2008; Rivers et al. 2008; Kang et al. 2010; Zhu et al. 2011).  Robust OPC proliferation and subsequent differentiation has been shown in murine models of CNS demyelination, SCI, as well as several other CNS pathologies (Buffo et al. 2005; McTigue et al. 2001; Zhang et al. 2013).  Therefore, changes in oligodendrocyte lineage cell densities may be an initial indication of an injury response.  The robust OPC proliferation observed in both the cervical and lumbar spinal cord would be predicted to manifest as an increase in total oligodendrocyte lineage cells.  We therefore sought to characterize the densities of total oligodendrocyte lineage cells (Olig2+ cells) (fig. 2.11a), committed oligodendrocytes (Olig2+/CC1+/PDGFRα- cells) (fig. 2.11b), and OPCs (Olig2+/PDGFRα+/YFP+/CC1- cells) (fig. 2.11c) at 12 wpi in cervical and lumbar spinal cord sections.  Designation of cell types was aided by visual assessment for expected morphology.  There was no change in cervical densities of total oligodendrocyte lineage cells (52388 ± 1607 cells/mm3; p=0.486, Mann Whitney-U test; n=4), committed oligodendrocytes (42674 ± 2433 cells/mm3; p=0.486, Mann Whitney-U test; n=4), or OPCs (3712 ± 175 cells/mm3; p=0.343, Mann Whitney-U test; n=4) at 12 wpi compared to uninjured controls (54455 ± 2210 Olig2+ cells/mm3; 45188 ± 2323 Olig2+/CC1+/PDGFRα- cells/mm3; 3394 ± 228 Olig2+/PDGFRα+/YFP+/CC1- cells/mm3; n=4 for all groups) (fig. 2.11d).  By contrast, there was a significant increase in the lumbar spinal cord densities of total oligodendrocyte lineage cells (68812   ± 2426 cells/mm3; p=0.0286, Mann Whitney-U test; n=4) and committed oligodendrocytes (59734 ± 1547 cells/mm3; p=0.0286, Mann Whitney-U test; n=4) at 12 wpi as compared to uninjured controls (57629 ± 1347 Olig2+ cells/mm3; 49037 ± 2128 Olig2+/CC1+/PDGFRα- cells/mm3; n=4 for all groups), but no 107  change in lumbar OPC density at 12 wpi (4397 ± 331 cells/mm3; n=4; p=0.686, Mann Whitney-U test) as compared to uninjured controls (4089 ± 445 cells/mm3; n=4) (fig. 2.11e).  The difference between the cervical and lumbar spinal cord may reflect a more robust lumbar proliferative response (corroborated by OPC proliferation data) or a greater degree of whole cord tissue atrophy.  Alternatively, increased committed oligodendrocyte density may reflect enhanced oligodendrogenesis, as has been reported previously in murine models of SCI (Rabchevsky et al. 2007), axotomy (Neilson et al. 2006), dorsal rhizotomy (Sun et al. 2010), and ALS (Kang et al. 2013).      Figure 2.11 Oligodendrocyte lineage cell densities in the cervical and lumbar spinal cord.  a) Oligodendrocyte lineage cells (defined as Olig2+ cells) (Olig2, white).  b) Committed oligodendrocytes (defined as CC1+/Olig2+ cells) (Olig2, white; CC1, red).  c) Oligodendrocyte precursor cells (OPCs) (defined as YFP+/Olig2+/PDGFRα+ cells) (YFP, green; Olig2, white; PDGFRα, white).  Densities of oligodendrocyte lineage cells, committed oligodendrocytes, 108  and OPCs in (d) cervical and (e) lumbar PDGFRα-CreERT2:ROSA26-YFP tissue at 12 wpi (orange) and uninjured controls (blue).  2.3.6 Oligodendrogenesis: whole-cord section analyses The demonstration of robust OPC proliferation in both the cervical and lumbar spinal cord, but no resultant change in OPC density at 12 wpi suggests that either the excess progeny died or differentiated into oligodendrocytes.  Enhanced oligodendrogenesis associated with robust OPC proliferation is characteristic of several CNS pathologies, including SCI (Rabchevsky et al. 2007), dorsal rhizotomy (Sun et al. 2010), and ALS (Kang et al. 2013).  In particular, robust oligodendrogenesis is concentrated in the spared tissue rim around murine contusion SCI lesion sites (Tripathi & McTigue 2007).  Furthermore, pronounced oligodendrogenesis extends through 9-12 wpi following contusion SCI (Hesp et al. 2015), suggesting robustness of the response (at least temporally).  Therefore, we sought to assess oligodendrogenesis remote to the lesion site in both the cervical and lumbar spinal cord.  To do this we exploited the fact the progeny of recombined (i.e. YFP+) OPCs will maintain expression of the reporter protein, as the STOP codon has been excised from the transgene construct (thus enabling the constitutive ROSA26 promoter to drive expression of the YFP gene).  Therefore, oligodendrocytes differentiated from recombined OPCs since the time of tamoxifen administration (i.e. 2 weeks prior to the injury) will maintain expression of the reporter protein, thus enabling their identification as ‘newly-formed’ (fig. 2.12a-b).  Quantification of newly-formed oligodendrocytes (i.e. YFP+/CC1+/Olig2+/PDGFRα- cells) as a percentage of total oligodendrocytes (i.e. CC1+/Olig2+/PDGFRα- cells) revealed significantly increased oligodendrogenesis in the cervical spinal cord at 12 wpi (13.3 ± 1.9%; p=0.0286, Mann Whitney-U test: n=4) as compared to temporally-matched (i.e. 14 weeks post-tamoxifen) 109  uninjured controls (6.7 ± 0.6%; n=4) (fig. 2.12c-e).  Similarly, increased oligodendrogenesis was observed in the lumbar spinal cord at 12 wpi (13.9 ± 0.8%; p=0.0286, Mann Whitney-U test; n=4) as compared to temporally-matched uninjured controls (7.9 ± 0.9%; n=4) (fig. 2.12f-h).  Assessment of oligodendrogenesis at earlier time-points post-injury (i.e. 1 dpi, 5 dpi, and 2 wpi) revealed a steady (albeit non-statistically significant) accumulation of new oligodendrocytes over time, as might be expected of a continual process (data not shown).        Figure 2.12 Oligodendrogenesis in the cervical and lumbar spinal cord is increased after thoracic SCI.  a) Newly-formed oligodendrocyte in PDGFRα-CreERT2:ROSA26-YFP cervical spinal cord tissue at 12 wpi (defined as 110  YFP+/CC1+/Olig2+/PDGFRα- cells) (YFP, green; CC1, red; Olig2, white; PDGFRα, blue).  b) 3D-rendering of a newly-formed oligodendrocyte in PDGFRα-CreERT2:ROSA26-mGFP tissue at 12 wpi (mGFP, green; Hoechst, blue).  Cervical newly-formed oligodendrocytes at (c) 12 wpi and in (d) uninjured PDGFRα-CreERT2:ROSA26-YFP tissue (YFP, green; CC1, red; Olig2, white; PDGFRα, blue).  (e) Quantification of newly-formed oligodendrocytes (YFP+/CC1+/Olig2+ cells) as a percentage of total committed oligodendrocytes (CC1+/Olig2+ cells) in the cervical spinal cord at 12 wpi (orange) and temporally-matched uninjured controls (blue).  Lumbar newly-formed oligodendrocytes at (f) 12 wpi and in (g) uninjured PDGFRα-CreERT2:ROSA26-YFP tissue (YFP, green; CC1, red; Olig2, white; PDGFRα, blue).  (h) Quantification of newly-formed oligodendrocytes (YFP+/CC1+/Olig2+ cells) as a percentage of total committed oligodendrocytes (CC1+/Olig2+ cells) in the cervical spinal cord at 12 wpi (orange) and temporally-matched uninjured controls (blue).                           2.3.7 De novo myelination: whole-cord section analyses Oligodendrocytes are the myelinating glial cells of the mammalian CNS, functioning to form myelin sheaths around axons (Zalc et al. 2008; Chang et al. 2016).  Logically, an increased production of new oligodendrocytes should correlate with an increase in new myelination.  We therefore sought to assess the extent of new myelination in the cervical and lumbar spinal cord.  Similar to the oligodendrogenesis analysis, we exploited the fact that progeny of recombined precursors will maintain reporter protein expression.  As the majority of cytoplasm is extruded during myelin sheath compaction (Chang et al. 2016) we utilized a transgenic mouse line expressing a membrane-bound GFP protein (i.e. PDGFRα-CreERT2:ROSA-mGFP).  Therefore, myelin sheaths produced by newly formed oligodendrocytes (i.e. differentiated from recombined OPCs post-tamoxifen) will retain expression of the mGFP protein, enabling their identification (fig. 2.13a/c).  As remyelination in the spared tissue rim at lesion epicentre is largely complete by 12 wpi in murine models of contusion SCI (Lasiene et al. 2008; Powers et al. 2013), we quantified 111  new myelin sheath density (i.e. mGFP+/MBP+ sheaths around axons) at 12 wpi in the cervical and lumbar spinal cord to ensure completion of the response.  Surprisingly, there was no difference in new myelination in the cervical spinal cord at 12 wpi (153.2 ± 19.0 sheaths/mm2; p=1.00, Mann Whitney-U test; n=4) as compared to temporally-matched uninjured controls (169.3 ± 62 sheaths/mm2; n=3) (fig. 2.13a-b).  Similarly, in the lumbar spinal cord there was no difference in new myelination at 12 wpi (177.7 ± 29 sheaths/mm2; p=0.0952, Mann Whitney-U test; n=6) as compared to temporally-matched uninjured controls (255.8 ± 30 sheaths/mm2; n=3) (fig. 2.13c-d).  Furthermore, a notable non-statistical trend for reduced new myelination in the injured lumbar spinal cord was apparent (fig. 2.13d), which was maintained even when new myelin sheath density was normalized to axon density to compensate for axonal loss (data not shown).   Figure 2.13 De novo myelination in the cervical and lumbar spinal cord is not significantly changed after SCI.  a) Cervical newly-formed myelin sheaths (defined as mGFP+/MBP+ sheaths around axons) in PDGFRα-112  CreERT2:ROSA26-mGFP tissue (mGFP, green; MBP, red; axons, white).  b) Quantification of cervical new myelin sheath density at 12 wpi (orange) and temporally-matched uninjured controls (blue).  c) Lumbar newly-formed myelin sheaths in PDGFRα-CreERT2:ROSA26-mGFP tissue (mGFP, green; MBP, red; axons, white).  d) Quantification of lumbar new myelin sheath density at 12 wpi (orange) and temporally-matched uninjured controls (blue).     2.3.8 Newly-formed oligodendrocyte maturation: whole-cord section analyses The demonstration of increased oligodendrogenesis but no increase in new myelination in either the cervical or lumbar spinal cord (with a trend for reduced lumbar new myelination) may be explained by a maturation impairment of newly-formed oligodendrocytes.  To investigate this possibility, we assessed the expression of a late-stage oligodendrocyte maturation marker (Gst-π) among newly-formed oligodendrocytes, used as a proxy of successful maturation (fig. 2.14a-b).  Gst-π is expressed by mature oligodendrocytes just prior to myelination, thus can be utilized to distinguish committed, but immature, oligodendrocytes (i.e. CC1+/Olig2+/Gst-π- cells) from mature oligodendrocytes (i.e. CC1+/Olig2+/Gst-π+ cells).  Consistent with a maturation impairment, we observed a significantly reduced proportion of newly-formed oligodendrocytes (i.e. YFP+/CC1+/Olig2+ cells) that expressed Gst-π in the lumbar spinal cord at 12 wpi (82.5 ± 1.5%; p=0.019, Mann Whitney-U test; n=6) as compared to temporally-matched uninjured controls (91.2 ± 0.9%; n=3) (fig. 2.14d).  Similarly, in the cervical spinal cord we observed a strong, but non-statistical, trend towards reduced proportion of newly-formed oligodendrocytes expressing Gst-π at 12 wpi (76.7 ± 2.3%; p=0.067, Mann Whitney-U test; n=6) as compared to temporally-matched uninjured controls (86.9 ± 2.4%; n=3) (fig. 2.14c).  Therefore, the absence of increased new myelination despite robust injury-induced oligodendrogenesis may, at least in part, be explained by a maturation impairment of newly-formed oligodendrocytes. 113    Figure 2.14 Oligodendrocyte maturation in the cervical and lumbar spinal cord is reduced after SCI.  Newly-formed oligodendrocytes in (a) injured (12 wpi) and (b) uninjured PDGFRα-CreERT2:ROSA26-YFP tissue (YFP, green; CC1, red; Gst-π, white; Hoechst, blue).  Quantification of mature newly-formed oligodendrocytes (YFP+/CC1+/Gst-π+ cells) as a percentage of total newly-formed oligodendrocytes (YFP+/CC1+ cells) in the (c) cervical and (d) lumbar spinal cord at 12 wpi (orange) and uninjured controls (blue).  2.3.9 Oligodendrocyte apoptosis If newly-formed oligodendrocytes suffer from impaired maturation in the cervical and lumbar spinal cord, indications of cell death may be evident.  Committed oligodendrocytes that fail to form myelin sheaths presumably have no function in the CNS, therefore would be expected to undergo cell death (including apoptosis), as has been demonstrated previously for genetically-induced maturation failure (via MyRF knockout) (Emery et al. 2009).  Therefore, we investigated 114  the occurrence of newly-formed oligodendrocytes (i.e. YFP+/CC1+ cells) undergoing apoptosis in the cervical and lumbar spinal cord at several time-points post-injury (i.e. 1 dpi, 5 dpi, 2 wpi, and 12 wpi).  Immunostaining for cleaved caspase-3 (a marker of cells actively undergoing apoptosis) (Porter & Jänicke 1999) revealed examples of apoptotic newly-formed oligodendrocytes (i.e. YFP+/CC1+/Casp3+ cells) in both the cervical and lumbar spinal cord (fig. 2.15a-b).  In addition to newly-formed oligodendrocytes, we also observed examples of apoptotic pre-existing oligodendrocytes (i.e. YFP-/CC1+/Casp3+ cells) in both the cervical and lumbar spinal cord (fig. 2.15c-d), consistent with previous description of prolonged apoptosis after contusion SCI (Shumann et al. 1997; Casha et al. 2001).  Unfortunately, the scarcity of Casp3+ nuclei (~2-3 per cord section) precluded effective quantification of apoptosis remote to the lesion site, probably related to the restricted expression of cleaved-caspase 3 to the transient apoptotic process.  TUNEL staining (Gorczyca et al. 1993) or more rigorous sampling protocols may address this limitation in future.  Regardless, these findings are consistent with the proposed model of oligodendrocyte maturation impairment remote to the lesion site. 115   Figure 2.15 Oligodendrocyte apoptosis in the cervical and lumbar spinal cord.  Apoptotic newly-formed oligodendrocytes (defined as YFP+/CC1+/Casp3+ cells) in (a) lumbar (2 wpi) and (b) cervical (2 wpi) injured PDGFRα-CreERT2:ROSA26-YFP tissue (YFP, green; CC1, red; cleaved caspase-3, white; Hoechst, blue).  Apoptotic pre-existing oligodendrocytes (defined as YFP-/CC1+/Casp3+ cells) in (a) lumbar (2 wpi) and (b) cervical (5 dpi) injured PDGFRα-CreERT2:ROSA26-YFP tissue (YFP, green; CC1, red; cleaved caspase-3, white; Hoechst, blue).    2.3.10 Histopathology: fasciculus gracilis analyses Since our whole spinal cord analysis included a random mixture of fibre tracts that were spared, proximal or distal to injury we decided to assess the influence of axonal injury/degeneration on observed changes to the oligodendrocyte lineage and focused the subsequent analyses on the cervical fasciculus gracilis (severed by the injury), compared to the adjacent fasciculus cuneatus (spared by the injury).  As OPCs are highly responsive to perturbations in extracellular conditions (Buffo et al. 2005; McTigue et al. 2001; Zhang et al. 2013), we first sought to characterize injury-116  induced pathological changes localized to the fasciculus gracilis.  As for the whole-cord section analyses, we selected here common CNS pathological features, specifically microgliosis, astrogliosis, axonal damage/loss, as well as myelin debris (Tator 1995; David & Kroner 2011; Plemel et al. 2013).  Substantially elevated microgliosis, as revealed by Iba-1 immunoreactivity, was observed in the fasciculus gracilis as compared to the fasciculus cuneatus at 1dpi (p=0.0286, Mann Whitney-U test; n=4), 5 dpi (p=0.0286, Mann Whitney-U test; n=4), 2 wpi (p=0.0286, Mann Whitney-U test; n=4), and 12 wpi (p=0.0286, Mann Whitney-U test; n=4) (fig. 2.16a-c).  Furthermore, quantification of Iba-1+ cell density revealed localized increase in the fasciculus gracilis as compared to the fasciculus cuneatus at 1 dpi (p=0.0286, Mann Whitney-U test; n=4), 5 dpi (p=0.0286, Mann-Whitney-U test; n=4), 2 wpi (p=0.0286, Mann Whitney-U test; n=4), and 12 wpi (p=0.0286, Mann Whitney-U test; n=4) (fig. 2.16d-f), suggesting that the microgliosis response includes microglial proliferation and/or migration in addition to activation of local microglia.  It is feasible that this effect might be due to tissue atrophy and/or axonal loss resulting in a reduced size of the fasciculus gracilis, which could be addressed in future with proliferative markers (e.g. Ki-67 or EdU/BrdU) to probe microglial proliferation.  Interestingly, both the Iba-1 immunoreactivity and Iba-1+ cell density quantifications revealed a steady increase through 12 wpi, consistent with a continually growing response.  In addition to reactive microglia, OPC behaviour can be modulated by reactive astrocytes (common feature of CNS histopathology).  Assessment of GFAP immunoreactivity (expressed as % of image area) revealed localized robust astrogliosis in the fasciculus gracilis at 12 wpi (55.6 ± 2.9%; p=0.0238, Mann Whitney-U test; n=4) as compared to temporally-matched uninjured controls (14.8 ± 1.1%; n=4) (fig. 2.18a-c).  By contrast, no elevation of GFAP immunoreactivity was observed in the fasciculus cuneatus at 12 wpi (24.6 ± 2.9%; p=0.548, Mann Whitney-U test; n=4) as compared to temporally-matched 117  uninjured controls (19.2 ± 0.7%; n=4) (fig. 2.18a-c), consistent with localization of astrogliosis to damaged/degenerating axonal tracts.  Quantification of β-APP+ profiles (interpreted as damaged axons) revealed substantial increases in the fasciculus gracilis at 12 wpi (106.9 ± 7.1; p=0.0238, Mann Whitney-U test; n=4) as compared to temporally-matched uninjured controls (2.7 ± 0.8; n=4), whereas no elevation was observed in the fasciculus cuneatus at 12 wpi (4.0 ± 0.6; p=0.437, Mann Whitney-U test; n=4) as compared to uninjured levels (3.3 ± 0.5; n=4) (fig. 2.17a-c).  Axonal loss was also observed in the fasciculus gracilis at 12 wpi, as revealed by absence of NF-200+/SMI-312+ profiles (fig. 2.17d-e).  To confirm that remaining axonal profiles represent transected axons rather than spared axons, we stained for SCG-10 (a.k.a. stathmin-2), a neuronal-specific growth-associated protein that accumulates in regenerating axons and the proximal segments of severed axons (Okazaki et al. 1995), implicated in lesion-induced sprouting and structural plasticity (Callahan et al. 1994; Viallet et al. 2002).  A large majority of the remaining axonal profiles in the fasciculus gracilis were SCG-10+, suggesting that they are severed stumps of axons not spared axons (fig. 2.17f-g).  The localized accumulation of β-APP+ profiles, decreased axonal profiles, and SCG-10+ axonal profiles in the fasciculus gracilis is in agreement with the predicted pattern of injury based on the injury model used (i.e. T9-T10 contusion).  Myelin debris is a known inhibitor of oligodendrocyte differentiation (Kotter et al. 2005; Plemel et al. 2013).  Persistence of myelin debris in the parenchyma is a common feature of SCI histopathology (Buss & Schwabb 2003; Buss et al. 2005), attributed to reduced phagocytic activity in the CNS (Ruckh et al. 2012).  MBP+ clusters were observed in the fasciculus gracilis at 12 wpi that were not associated with axons, consistent with myelin debris (fig. 2.19a-b).  To confirm, we stained cervical spinal cord sections with Oil Red O (a lipid dye which preferentially stains myelin debris dark purple, in contrast to staining normal myelin a pale red/purple) (Perry et al. 1987; Prineas et 118  al. 1993; Mickel et al. 2016).  Obvious myelin debris was observed specifically in the fasciculus gracilis (fig. 2.19e-f).  Finally, we also stained cervical spinal cord sections for degraded MBP (dMBP), revealing immunostaining localized specifically in the fasciculus gracilis (fig. 2.19c-d).  Collectively, these findings demonstrate the presence of myelin debris in the fasciculus gracilis at 12 wpi.  It is possible that this represents myelin debris internalized in phagocytic cells (e.g. microglia/macrophages) and not extracellular.  This has relevance for the effect of the myelin debris on oligodendrocyte lineage cells, as they may not be directly exposed.    Figure 2.16 Microgliosis is increased in the cervical fasciculus gracilis after thoracic contusion.  Overview of microglia in the cervical dorsal column of (a) injured (12 wpi) and (b) uninjured PDGFRα-CreERT2:ROSA26-YFP tissue.  Outlined area represents a region of microgliosis (Iba-1, red).  c) Quantification of Iba-1 immunoreactivity in the fasciculus gracilis (FG) and fasciculus cuneatus (FC) at 1 dpi, 5 dpi, 2 wpi, and 12 wpi (orange) and uninjured controls (blue).  Microglial morphologies in the (d) injured (12 wpi) and (e) uninjured fasciculus gracilis (Iba-1, red; 119  Hoechst, blue).  f) Quantification of Iba-1+ cell density in the fasciculus gracilis (FG) and fasciculus cuneatus (FC) at 1 dpi, 5 dpi, 2 wpi, and 12 wpi (orange) and uninjured controls (blue).              Figure 2.17 Axonal pathology and axonal sprouting in the cervical fasciculus gracilis. Axonal damage in the cervical dorsal column of (a) injured (12 wpi) and (b) uninjured PDGFRα-CreERT2:ROSA-mGFP tissue (β-APP, turquoise).  c) Quantification of β-APP+ axonal profiles in the cervical fasciculus gracilis and fasciculus cuneatus at 12 wpi (orange) and uninjured controls (blue).  Overview of dorsal column axonal complement in (d) injured (12 wpi) and (e) uninjured cervical spinal cord (NF-200/SMI-312, white).  Outlined area represents a region of axonal loss.  120  SCG-10 (a.k.a. stathmin-2) expression in fasciculus gracilis axons in (f) injured (12 wpi) and (g) uninjured spinal cord tissue (SCG-10, red; NF-200/SMI-312, white).   Figure 2.18 Astrogliosis in the cervical fasciculus gracilis.  Overview of GFAP expression in the cervical dorsal column of (a) injured (12 wpi) and (b) uninjured PDGFRα-CreERT2:ROSA26-mGFP tissue (GFAP, white).  Outlined area represents region of putative astrogliosis.  c) Quantification of GFAP immunoreactivity in the fasciculus gracilis and fasciculus cuneatus at 12 wpi (orange) and uninjured controls (blue).       121       Figure 2.19 Myelin debris in the cervical fasciculus gracilis.  Overview of MBP staining in the cervical dorsal column in (a) injured (12 wpi) and (b) uninjured PDGFRα-CreERT2:ROSA26-mGFP (MBP, blue; NF-200/SMI-312, white).  Inset images show putative myelin debris (i.e. MBP clusters not associated with axons – denoted by *) in the fasciculus gracilis.  Degraded MBP staining in the cervical dorsal column of (c) injured (12 wpi) and (d) uninjured PDGFRα-CreERT2:ROSA26-mGFP tissue (dMBP, red; MBP, blue; NF-200/SMI-312, white).  Inset images show dMBP+ myelin debris in the fasciculus gracilis.  Overview of Oil Red O stained cervical dorsal column in (e) injured (12 wpi) and (f) uninjured PDGFRα-CreERT2:ROSA26-mGFP tissue.  Inset images show myelin debris (i.e. darker staining regions) in the fasciculus gracilis.  122  2.3.11 Oligodendrocyte lineage cell responses: fasciculus gracilis analyses Analysis of whole-cord sections (as done for previous analyses) inherently involves inclusion of both spared and the proximal and distal parts of injured axonal tracts in the assessment.  Previous studies have demonstrated that axonal injury/degeneration is associated with microgliosis, OPC proliferation, and oligodendrogenesis in axotomy (Neilson et al. 2006), dorsal rhizotomy (Sun et al. 2010), ALS (Kang et al. 2013), and SCI models.  Therefore, we sought to assess the influence of axonal injury/degeneration on local oligodendrocyte lineage cells remote to the lesion in the cervical and lumbar spinal cord.  Ideally, identification of axonal tracts would be aided by retrograde and/or anterograde tracers (e.g. BDA, microruby, etc.) to provide unambiguous designation of specific axonal populations for analysis, however tracers were not utilized in the current study.  Instead, we exploited the anatomical relation of the fasciculus gracilis and fasciculus cuneatus in the dorsal column of the cervical spinal cord (Watson & Harrison 2012; Watson 2009).  As the fasciculus gracilis contains sensory afferents originating caudal to the lesion, the axons are severed by the injury (i.e. damaged/degenerating axonal tract), whereas the sensory afferents of the fasciculus cuneatus originate rostral to the lesion site, thus are spared (i.e. spared axonal tract).  This provides a useful system in which to assess the influence of axonal damage/degeneration on oligodendrocyte lineage cell responses remote to the lesion.  Quantification of the percentage of YFP+/PDGFRα+ cells with OPC morphology (i.e. extensive, branched arbours of processes extending into the parenchyma) that were Ki-67+ revealed substantially elevated proliferation in the fasciculus gracilis as compared to the fasciculus cuneatus at 5 dpi (p=0.0286, Mann Whitney-U test; n=4), 2 wpi (p=0.0294, Mann Whitney-U test; n=4), and 12 wpi (p=0.0286, Mann Whitney-U test; n=4) (fig. 2.20a-d).  Furthermore, concentrated oligodendrogenesis was observed in the fasciculus gracilis, as revealed by an increased percentage 123  of newly-formed oligodendrocytes (i.e. YFP+/CC1+/Olig2+/PDGFRα- cells) at 12 wpi (14.7 ± 1.7%; p=0.0286, Mann Whitney-U test; n=4) as compared to temporally-matched uninjured controls (6.1 ± 0.5%; n=4) (fig. 2.21a-d).  In contrast, oligodendrogenesis was not elevated in the spared fasciculus cuneatus, as the percentage of newly-formed oligodendrocytes at 12 wpi (7.5 ± 1.1%; p=0.0571, Mann Whitney-U test; n=4) was not increased above temporally-matched uninjured controls (4.1 ± 1.0%; n=4) (fig. 2.21a-d), however there was a notable trend for increased oligodendrogenesis (consistent with the whole-cord analyses) (fig. 2.21d).  Interestingly, quantification of new myelination (i.e. density of mGFP+/MBP+ sheaths) in the fasciculus gracilis revealed a significant decrease at 12 wpi (66.6 ± 10.2 sheaths/mm2; p=0.0275, Mann Whitney-U test; n=6) as compared to temporally-matched uninjured controls (248.9 ± 54.4 sheaths/mm2; n=3) (fig. 2.22a-d).  Consistent with a specific effect of the damaged axons, there was no difference in new myelination in the fasciculus cuneatus at 12 wpi (277 ± 60.6 sheaths/mm2; p=0.714, Mann Whitney-U test; n=6) as compared to temporally-matched uninjured controls (199 ± 98.8 sheaths/mm2; n=3) (fig. 2.22a-d).  124   Figure 2.20 OPC proliferation in the cervical fasciculus gracilis.  Overview of PDGFRα+ cell proliferation in the cervical dorsal column of (a) injured (2 wpi) and (b) uninjured PDGFRα-CreERT2:ROSA26-YFP tissue (YFP, green; Ki-67, red; Hoechst, blue; PDGFRα, white).  c) Proliferative PDGFRα+ cell with OPC morphology in the fasciculus gracilis at 2 wpi (YFP, green; Ki-67, red; Hoechst, blue; PDGFRα, white).   d) Quantification of PDGFRα+ cell proliferation (as a percentage of total PDGFRα+ cells in the fasciculus gracilis (FG) and fasciculus cuneatus (FC) at 1 dpi, 5 dpi, 2 wpi, and 12 wpi (orange) and uninjured controls (blue).   125                   Figure 2.21 Increased oligodendrogenesis in the cervical fasciculus gracilis after spinal cord injury.  Overview of oligodendrogenesis in the cervical dorsal column of (a) injured (12 wpi) and (b) uninjured PDGFRα-CreERT2:ROSA26-YFP tissue (YFP, green; CC1, red; PDGFRα, blue; Olig2, white).  c) Newly-formed oligodendrocytes (defined as YFP+/CC1+/Olig2+/PDGFRα- cells) in the cervical fasciculus gracilis at 12 wpi.  d) Quantification of newly-formed oligodendrocytes (i.e. YFP+/CC1+/Olig2+ cells) as a percentage of total committed oligodendrocytes (i.e. CC1+/Olig2+ cells) in the fasciculus gracilis and fasciculus cuneatus at 12 wpi (orange) and uninjured controls (blue).      126                        Figure 2.22 Reduced new myelination in the cervical fasciculus gracilis after spinal cord injury.  a) Overview of new myelination in the cervical dorsal column in injured (12 wpi) PDGFRα-CreERT2:ROSA26-mGFP tissue (mGFP, green; MBP, blue).  b) 3D-rendering of a newly-formed myelinating oligodendrocyte in PDGFRα-CreERT2:ROSA26-mGFP tissue (mGFP, green).  c) Newly-formed myelin sheaths (defined as mGFP+/MBP+ sheaths around axons) in the cervical fasciculus gracilis at 12 wpi (mGFP, green; MBP, blue; NF-200/SMI-312, white).  d) Quantification of new myelin sheath density in the cervical fasciculus gracilis and fasciculus cuneatus at 12 wpi (orange) and uninjured controls (blue).   2.3.12 OPC lineage plasticity Despite restriction to the oligodendrocyte lineage under physiological conditions, OPCs may generate cells of other lineages (specifically astrocytes and/or Schwann cells) in certain CNS pathologies (Dimou et al. 2008; Barnabé-Heider et al. 2010; Zawadzka et al. 2010; Komitova et al. 2011).  We therefore sought to investigate injury-induced changes in OPC lineage potential 127  remote to the lesion site.  Similar to the oligodendrogenesis and new myelination analyses, we relied upon the fact that progeny of recombined cells will maintain expression of the reporter proteins (as the STOP codon has been excised from the genome).  Therefore, if OPC-derived astrocytes and/or Schwann cells are generated in the period following tamoxifen administration (i.e. 2 weeks prior to the injury) they will express the reporter proteins (i.e. YFP or mGFP).  Cervical and lumbar spinal cord tissue from PDGFRα-CreERT2:ROSA26-mGFP mice was stained for GFAP (a prototypical astrocyte marker) (Sofroniew 2009).  As mGFP is membrane-bound and GFAP is a cytoskeletal protein we searched for GFAP+ filaments inside mGFP+ cell processes, instead of marker co-localization.  Examination of 12 wpi spinal cord sections did not reveal any cervical or lumbar mGFP+/GFAP+ cells (i.e. putative OPC-derived astrocytes) (fig. 2.24b/d).   As GFAP does not label all non-reactive astrocytes (Sofroniew & Vinters 2010), we sought a complementary assessment of OPC astrogenesis.  Cervical and lumbar spinal cord tissue from PDGFRα-CreERT2:ROSA26-YFP mice was stained for Aldh1L1 (a cytoplasmic astrocyte marker).  Consistent with GFAP staining results, no YFP+/Aldh1L1+ cells were observed at 12 wpi in either the cervical or lumbar spinal cord (fig. 2.24a/c).  Significant pathological changes may be required to stimulate OPC-astrogenesis (Zawadska et al. 2010; Komitova et al. 2011).  Therefore, we focused subsequent assessment on the cervical fasciculus gracilis (i.e. an area of concentrated pathology).  Despite focal astrogliosis, microgliosis, and dense accumulation of mGFP+ processes, no mGFP+/GFAP+ cells were observed in the 12 wpi cervical fasciculus gracilis (fig. 2.24e-f).  To assess for OPC Schwann cell generation, cervical and lumbar PDGFRα-CreERT2:ROSA26-mGFP spinal cord tissue was stained for P0 (a peripheral myelin protein).  As expected, P0 immunostaining was restricted to the cervical and lumbar peripheral roots at 12 wpi, and completely absent from central spinal cord tissue (fig. 2.23a-b).  Furthermore, in the peripheral 128  roots mGFP+/P0+ cells were not observed, suggesting the absence of Schwann cell production from the peripheral recombined cell population.  These findings are consistent with an absence of myelinating Schwann cell production from recombined precursors remote to the lesion site. Collectively, OPC progeny appears to remain restricted to the oligodendrocyte lineage remote to the lesion site in both the cervical and lumbar spinal cord.                            Figure 2.23 Absence of Schwann cell production from OPCs in the cervical and lumbar spinal cord.  Overview of P0 immunostaining in injured (12 wpi) (a) cervical and (b) lumbar PDGFRα-CreERT2:ROSA26-mGFP tissue (mGFP, green; P0, red).  Note restriction of P0 immunostaining to the peripheral roots.                         129        Figure 2.24 Absence of OPC astrogenesis in the cervical and lumbar spinal cord.  a) Injured (12 wpi) (a) cervical and (c) lumbar PDGFRα-CreERT2:ROSA26-YFP tissue stained for Aldh1L1 (an astrocyte marker) (YFP, green; Aldh1L1, red; Hoechst, blue).  Note absence of marker co-localization (i.e. YFP+/Aldh1L1+ cells).  Injured (12 wpi) (b) cervical and (d) lumbar PDGFRα-CreERT2:ROSA26-mGFP tissue stained for GFAP (an astrocyte marker) (mGFP, green; GFAP, red; Hoechst, blue).  Note absence of marker co-localization (i.e. mGFP+/GFAP+ cells).  e) Overview of cervical dorsal column of injured (12 wpi) PDGFRα-CreERT2:ROSA-mGFP stained for GFAP (mGFP, green; GFAP, red; Hoechst, blue).  f) Inset image from (e) demonstrating absence of marker co-localization in the injured fasciculus gracilis at 12 wpi.           130  Chapter 3: Discussion  3.1 Introduction As the myelinating glia of the CNS, the impact of pathological insult on oligodendrocytes is of intense interest for both pathophysiological characterization and therapeutic targeting (Bergles & Richardson 2015; Emery & Lu 2015).  Murine contusion SCI elicits a robust oligodendrocyte lineage response at lesion epicentre, consisting of OPC proliferation, oligodendrogenesis, and remyelination of denuded axons (McTigue et al. 2001; Rabchevsky et al. 2007; Lasiene et al. 2008; Powers et al. 2013).  As OPCs are highly responsive to extracellular perturbations (Buffo et al. 2005; McTigue et al. 2001; Zhang et al. 2013) a similar response may be initiated remote to the lesion, potentially triggered by axonal damage/degeneration.  Indeed, robust OPC proliferation and oligodendrogenesis occurs along damaged/degenerating axonal tracts in murine models of dorsal rhizotomy (Sun et al. 2010), axotomy (Neilson et al. 2006), and ALS (Kang et al. 2013).  Moreover, isolated thoracic SCI induces remote gliogenesis (including OPC proliferation) (Franz et al. 2014).  Microglia-derived factors may play a role in the initiation/propagation of this response.  Indeed, chemical-activation of microglia stimulates OPC proliferation in vivo (Schonberg et al. 2007) and microgliosis is coincident with OPC proliferation and oligodendrogenesis along damaged/degenerating axonal tracts in dorsal rhizotomy (Sun et al. 2010), axotomy (Neilson et al. 2006), and ALS (Kang et al. 2013) models.  As isolated murine thoracic SCI induces robust microgliosis remote to the lesion site (Wu et al. 2014a; Wu et al. 2014b), remote OPC populations are exposed to microglia-derived factors.  The hypothesis addressed by the current thesis is that isolated thoracic contusion spinal cord injury (SCI) will induce a robust response in remote oligodendrocyte lineage cell populations (predicted to consist of OPC proliferation, 131  oligodendrogenesis, and new myelination), potentially attributed to axonal damage-induced microgliosis.  3.2 Overview of experimental findings To address the stated hypothesis, we conducted an in vivo study utilizing a well-characterized murine model of thoracic (T9-T10) contusion SCI and two distinct transgenic reporter mouse lines (PDGFRα-CreERT2:ROSA26-YFP and PDGFRα-CreERT2:ROSA26-mGFP) that enabled detailed assessment of OPC proliferation, oligodendrogenesis, and new myelination remote to the lesion site at various time-points spanning the sub-acute and acute post-injury periods (i.e. 1 dpi, 5 dpi, 2 wpi, and 12 wpi).  We initially performed whole-cord section analyses, with the notable caveat of sampling from injured and uninjured tracts indiscriminately.  As OPCs are highly responsive to the local environment, we first sought to characterize injury-induced histopathological features remote to the lesion site.  Using established quantification methods, we revealed robust microgliosis, axonal damage, and cell proliferation, but no whole-cord astrogliosis, white matter atrophy, nor overt axonal loss.  Ki-67 immunostaining revealed rapid and robust OPC proliferation in both the cervical and lumbar spinal cord, coincident with microgliosis (both spatially and temporally).  Despite this proliferative response, neither cervical nor lumbar OPC density was altered at 12 wpi.  Importantly, lumbar committed oligodendrocyte density was increased at 12 wpi, suggesting that excess OPC progeny may differentiate into oligodendrocytes.  Indeed, we observed robust cervical and lumbar oligodendrogenesis, reflected in a ~2-fold increase in the percentage of newly-formed oligodendrocytes at 12 wpi (as compared to uninjured controls).  It might be expected that increased oligodendrogenesis should be accompanied by increased new myelination.  Surprisingly however, there was no difference in either cervical or lumbar new 132  myelination (as revealed by new myelin sheath density) at 12 wpi as compared to temporally-matched uninjured controls.  We hypothesized two potential explanations: (i) newly-formed oligodendrocytes in the injured spinal cord form fewer myelin sheaths per cell and/or (ii) a reduced proportion of newly-formed oligodendrocytes mature to the myelinating stage in the injured spinal cord (i.e. display impaired maturation).  As adult-born oligodendrocytes are known to produce more myelin sheaths than their developmental counterparts (Young et al. 2013), the later explanation is more likely.  Indeed, we observed reduced expression of a late-stage maturation marker (Gst-π) among newly-formed oligodendrocytes at 12 wpi.  If maturation is disrupted, signs of cell death should be evident.  Immunostaining for cleaved caspase-3 revealed apoptotic newly-formed oligodendrocytes remote to the lesion site.  We also observed apoptotic pre-existing oligodendrocytes, consistent with the prolonged oligodendrocyte apoptosis that follows SCI (Casha et al. 2001; Crowe et al. 1997).  As axonal injury/degeneration is correlated with robust oligodendrocyte lineage cells responses across CNS pathologies (e.g. dorsal rhizotomy, axotomy, and ALS) (Sun et al. 2010; Neilson et al. 2006; Kang et al. 2013), we sought to constrain assessment of observed responses to an injured axonal tract.  Without the aid of tracers (e.g. BDA), we relied upon the anatomical relation of sensory afferents in the cervical dorsal column (assuming a somatotopic organization) (Watson & Harrison 2012; Niu et al. 2013), which are differentially affected by the thoracic injury.  We focused specifically on fasciculus gracilis afferents, as they are injured by the contusion.  Utilizing established quantification methods (same as above), we revealed concentrated OPC proliferation and oligodendrogenesis in the fasciculus gracilis, accompanied by focal microgliosis and astrogliosis.  In agreement with noted whole-cord trends, we observed reduced new myelination in the fasciculus gracilis.  This may partially be attributed to a lack of permissive substrate for myelination (i.e. spared/uninjured axons).  Indeed, β-APP 133  immunostaining confirmed concentrated axonal pathology in the fasciculus gracilis.  Furthermore, persistent myelin debris (as revealed by dMBP immunostaining and Oil Red O staining) was observed in the fasciculus gracilis at 12 wpi.  Despite restriction to the oligodendrocyte lineage under physiological conditions, OPCs are proposed to generate cells of other lineages (specifically astrocytes and/or Schwann cells) in certain CNS pathologies (Dimou et al. 2008; Barnabé-Heider et al. 2010; Zawadzka et al. 2010; Komitova et al. 2011).  Immunostaining for markers of astrocytes (i.e. GFAP and Aldh1L1) and Schwann cells (i.e. P0) did not reveal any deviation from the oligodendrocyte lineage by recombined OPC progeny.  3.3 Injury-induced OPC proliferation remote to the lesion site OPCs respond rapidly (<1 dpi) (Horky et al. 2006; Simon et al. 2011) to several CNS insults, including stab wound (Buffo et al. 2005; Dimou et al. 2008; Levine 1994), ischemia (Zhang et al. 2013), and SCI (McTigue et al. 2001).  These ‘reactive’ OPCs are characterized by several features, including: (i) cell body hypertrophy, (ii) numerous short and thick processes, (iii) increased proliferation rate, and (iv) upregulation of the NG2 proteoglycan (Levine 1994).  In vivo imaging revealed dynamic OPC responses to demyelination or focal laser ablation, comprised of enhanced proliferation, migration, and accumulation at the lesion (Hughes et al. 2013; Hill et al. 2013).  Similarly, contusion SCI triggers robust OPC proliferation at the lesion epicentre as early as 1 dpi, peaking at 3-5 dpi, and continuing through 2 wpi (McTigue et al. 2001; Zai & Wrathall 2005).  We observed a largely similar response caudal to the lesion, with increased lumbar OPC proliferation starting at 1 dpi, peaking at 5 dpi, and continuing through 2 wpi.  By contrast, the cervical proliferative response was more delayed, beginning at 5 dpi, but persisting through 2 wpi.  However, the sparse sampling across the post-injury period (i.e. 1 dpi, 5 dpi, 2 wpi and 12 wpi) 134  may be ignorant of the actual proliferation onset, as well the termination of the response.  A more thorough assessment may therefore be warranted.  The rapid onset of lumbar OPC proliferation (i.e. beginning at 1 dpi) is incompatible with a diffusible trigger originating from the lesion (e.g. blood serum, cell debris, etc.).  As it enables rapid communcation to remote spinal locations, axonal injury/degeneration may elicit the observed response.  Indeed, we observed concentrated OPC proliferation in the cervical fasciculus gracilis (i.e. a damaged/degenerating axonal tract).  Similarly, OPC proliferation occurs along degenerating axonal tracts in dorsal rhizotomy (Sun et al. 2010), axotomy (Neilson et al. 2006), Alzheimer’s disease (AD) (Behrendt et al. 2013), and ALS models (Kang et al. 2013; Philips et al. 2013; Magnus et al. 2008).  Demyelination may also contribute, as OPC proliferation in response to demyelinating insult is well-known (Keirstead et al. 1998; Levine & Reynolds 1999; Franklin et al. 2002; Komitova et al. 2011) and at least a degree of demyelination is expected in dorsal rhizotomy, SCI, ALS, and AD models (Sun et al. 2010; Neilson et al. 2006; Kang et al. 2013).  Indeed demyelination may be a key trigger for OPC proliferation, as neither p25-induced cortical neuronal death (Cruz et al. 2003; Cruz et al. 2006; Sirko et al. 2013), loss of nigral dopaminergic neurons in a murine model of Parkinson’s disease (PD) (Steiner et al. 2006), nor non-demyelinating inflammatory insult (Bello et al. 1999) elicits OPC proliferation.  While not directly assessed, at least a degree of demyelination may be inferred from several observations, including: (i) presence of myelin debris, (ii) apoptosis of pre-existing oligodendrocytes (which presumably formed myelin sheaths), and (iii) axonal loss (which should be accompanied by loss of associated oligodendrocytes).  In addition to demyelination directly, oligodendrocyte damage/loss may also trigger OPC proliferation, potentially as a homeostatic mechanism to replace oligodendrocytes.  Prolonged oligodendrocyte apoptosis following injury would provide sustained signals for OPC proliferation.  Indeed, we observed both cervical and 135  lumbar oligodendrocyte apoptosis through at least 2 wpi.  OPC proliferation may also be induced by microgliosis.  Indeed, chemical activation of microglia is sufficient to induce OPC proliferation (Schonberg et al. 2007) and microglia-derived cytokines are implicated in OPC proliferation following CNS demyelination (Almad et al. 2011; Dimou & Gallo 2015).  Furthermore, M1 macrophage/microglia ablation following focal demyelination attenuates OPC proliferation (Miron et al. 2013).  Microglial activation along degenerating axonal tracts is known, attributed to axonal fragmentation (Neumann et al. 2009) and release of glutamate (de Jong et al. 2005).  For example, microgliosis occurs along degenerating axonal tracts in dorsal rhizotomy (Sun et al. 2010), contusion SCI (McTigue et al. 2001), and ALS (Kang et al. 2013; Philips et al. 2013) models, associated with OPC proliferation.  Indeed, we observed substantial overlap between Iba-1 and β-APP immunostaining (consistent with microgliosis along degenerating axons), as well as focal microgliosis in the cervical fasciculus gracilis concomitant with concentrated OPC proliferation.  Therefore, the effect of axonal damage on OPC proliferation may be mediated through microglial activation along degenerating tracts.  Manipulation of microglial responses (i.e. blocking activation, etc.) and characterization of the cytokine composition of the post-injury environment may help elucidate a causal relationship.  In addition to microglia-derived cytokines, a host of other potential OPC mitogens are upregulated in the pathological CNS (e.g. bFGF, PDGF, IGF-1, CNTF, IL-6, IL-8, CCL5, CXCL1, prolactin, adenosine, and ATP) (Robinson et al. 1998; Wu et al. 2000; Kadi et al. 2006; Miller 2002), which may contribute, at least partially, to the observed OPC proliferation.  Even subtle changes might have substantive effects, as OPC responsiveness may be altered following injury, as demonstrated previously (Barres et al. 1993; Cohen et al. 1996; McTigue et al. 1998).  In addition to extracellular factors, altered neural activity can also elicit OPC proliferation.  For example, optogenetic-mediated stimulation of layer V 136  projection neurons induces robust OPC proliferation in the premotor cortex (Gibson et al. 2014).  Furthermore, OPC proliferation is stimulated by novel locomotor paradigms (e.g. wheel running) (Ehninger et al. 2011; Simon et al. 2011; McKenzie et al. 2014) and enriched environment exposure (Zhao et al. 2012), presumably underlain by altered neural activity.  It is feasible that novel forelimb gait patterns compensating for lost hind-limb locomotor function, underlain by novel activity patterns, may stimulate OPC proliferation along corresponding spared motor and sensory tracts.  Indeed, cervical OPC proliferation was observed within the cervical enlargement (i.e. containing the motor and sensory innervation of the forelimbs) and was not constrained to regions of axonal damage (i.e. OPC proliferation occured in proximity to spared axons).  Interestingly, inhibiting neural activity may also promote OPC proliferation.  For example, robust OPC proliferation is observed following conduction blockage, inhibition of vesicular release, or AMPA receptor antagonism in demyelinated lesions (Gautier et al. 2015) as well as following deprivation of whisker sensory input (Mangin et al. 2012).  It is feasible that reduced activity of transiting axons due to inflammation, CSPG and/or NG2 deposition (discussed below) may elicit proliferative responses from associated OPC populations.  Further interrogation employing electrophysiological and/or optogenetic approaches (combined with tracers to unambiguously identify specific tracts) may address these speculations.  In summary, injury-induced remote OPC proliferation may represent a cumulative effect of several factors, including: (i) microgliosis (triggered by axonal injury/degeneration), (ii) altered neural activity, (iii) oligodendrocyte damage/demyelination, (iv) altered extracellular levels of OPC mitogens, and/or (v) injury-induced changes in OPC responsiveness.  137  3.4 Injury-induced oligodendrogenesis remote to the lesion site The best characterized function of adult OPCs is as precursors of myelinating oligodendrocytes (Nishiyama et al. 2009; Emery & Lu 2015).  Robust oligodendrogenesis is observed in several CNS pathologies (e.g. SCI, ALS, demyelination, axotomy, dorsal rhizotomy, etc.) (Sun et al. 2010; Kang et al. 2013; Neilson et al. 2006), operating as a homeostatic mechanism to replace lost oligodendrocytes and/or myelin (Almad et al. 2011; Plemel et al. 2014).  Indeed, oligodendrogenesis from local OPCs compensate for focal oligodendrocyte loss in chemical demyelination models (e.g. lysolecithin, cuprizone) (Franklin & ffrench-Constant 2008).  Similarly, concentrated oligodendrogenesis in the contusion SCI lesion rim restores pre-injury densities despite extensive oligodendrocyte loss (~93% at 1 wpi) (Zai & Wrathall 2005; Tripathi & McTigue 2007).  The cervical and lumbar oligodendrogenesis observed here may represent a similar homeostatic response, which would assume a substantial oligodendrocytes loss (discussed below), which was not quantitatively assessed.  Alternatively, oligodendrogenesis may be triggered by axonal damage/degeneration.  Indeed, robust oligodendrogenesis occurs along degenerating axonal tracts in dorsal rhizotomy (Sun et al. 2010), ALS (Kang et al. 2013; Philips et al. 2013), axotomy (Nielsen et al. 2006), and contusion SCI models (Zai & Wrathall 2005; Tripathi & McTigue 2007).  Consistent with this, we observed concentrated oligodendrogenesis in the cervical fasciculus gracilis.  Similar to OPC proliferation, activated microglia may mediate this effect, as microglial-derived factors are implicated in oligodendrogenesis and myelination (Almad et al. 2011; Plemel et al. 2014; Foote & Blakemore 2005; Miron et al. 2013; Morell et al. 1998).  Indeed, minocycline-mediated inhibition of microglial activation impairs remyelination (Li et al. 2005).  Furthermore, M2 macrophages/microglia ablation reduces oligodendrocyte differentiation (Miron et al. 2013), related to reduced myelin debris phagocytosis (Miron et al. 138  2013; Ruckh et al. 2012) and/or reduced production of pro-differentiation factors (e.g. activin-A) (Miron et al. 2013).  It has been proposed that microglia/macrophages (as well as astrocytes) are a requirement for adult oligodendrogenesis (Franklin et al. 1991; Bello et al. 1999; Kotter et al. 2001; Talbott et al. 2005).  Indeed, oligodendrogenesis around ischemic (Mandai et al. 1997; Mabuchi et al. 2000; Komitova et al. 2006) and contusion SCI lesions (Tripathi & McTigue 2007) occurs in regions of high microglia/macrophage density.  Furthermore, TLR4-mediated intra-spinal activation of microglia stimulates robust oligodendrocyte differentiation (Schonberg et al. 2007).  Indeed, elevated oligodendrogenesis observed in the current study was concurrent with microgliosis (both cervical and lumbar), particularly evident in the cervical fasciculus gracilis.  In addition to microglia, reactive astrocytes may also promote oligodendrocyte differentiation.  Indeed, oligodendrogenesis is concentrated in astrocytic scar regions following SCI (Tripathi & McTigue 2007) and reactive astrocytes produce a host of pro-differentiation factors (e.g. PDGF, bFGF, IGF, and CNTF) (Zai & Wrathall 2005; Almad et al. 2011).  Supporting a potential role, we observed elevated oligodendrogenesis in the cervical fasciculus gracilis, coincident with focal astrogliosis.  In addition to reactive glia-derived factors, oligodendrogenesis is regulated by  various extracellular components, including TGF-β/BMP family members (Dutta et al. 2014; Palazuelos et al. 2014), IGF-1 (McMorris & Dubois-Dalcq 1988; D’Ercole & Ye 2008), neuregulins (Canoll et al. 1996; Vartanian et al. 1999; Carteron et al. 2006), NT-3 (Barres et al. 1993; Cohen et al. 1996), Wnts (Shimizu et al. 2005; Fancy et al. 2009; Dai et al. 2014), chemokines (Robinson et al. 1998; Dziembowska et al. 2005; Göttle et al. 2010), CNTF (Barres et al. 1993), BDNF (Vondran et al. 2010; Wong et al. 2013), transferrin (Silvestroff et al. 2013), erythropoietin (Sugawa et al. 2002), thyroid hormone (Barres et al. 1994), and retinoic acid (Barres et al. 1994; Miller 2002).  As extracellular levels of these factors are altered in the injured spinal 139  cord, the observed oligodendrogenesis may reflect, at least in part, injury-induced changes in the extracellular milieu remote to the lesion (Pineau & Lacroix 2007; Tripathi & McTigue 2007; Zai & Wrathall 2005; Oyesiku et al. 1997; Semple-Rowland et al. 1995; McTigue et al. 2000; Buss et al. 2008).  As with OPC proliferaiton, neural activity can stimulate oligodendrogenesis (Chang et al. 2016; Fields 2015; Bergles & Richardson 2015).  Indeed, robust oligodendrocyte production in the premotor cortex, accompanied by de novo myelination, follows optogenetic-mediated stimulation of layer V cortical neurons (Gibson et al. 2014).  Conversely, reduction of activity (via conduction blockage, inhibition of vesicular release, or NMDA receptor antagonism) inhibits oligodendrogenesis in demyelinating lesions (Gautier et al. 2015).  Alteration of activity in spared axons may stimulate oligodendrocyte production remote to the lesion.  Indeed, clusters of newly-formed oligodendrocytes were observed outside of regions of axonal damage (confirmed by absence of β-APP immunostaining).  Activation of homeostatic mechanisms may also prompt oligodendrogenesis.  Adult OPCs possess robust self-repulsive mechanisms (Hughes et al. 2013), involved in maintenance of cell density, a tiled-like arrangement, and process morphology (Dimou & Gallo 2015; Levine 2016).  It is plausible that local increases in OPC density resulting from robust proliferation may enact homeostatic mechanisms promoting differentiation to restore pre-injury OPC densities.  Indeed, oligodendrocyte differentiation has been shown to be density- and spatial-dependent, as well as influenced by local biophysical properties (Rosenberg et al. 2008).  This might be particularly important in regions of concentrated microgliosis and astrogliosis (e.g. the cervical fasciculus gracilis), as increased cell density (due to microglia, astrocyte, and OPC proliferation) and deposition of extracellular factors by reactive glia (e.g. astrocyte-derived CSPGs, OPC-derived NG2) may significantly alter local biophysical properties.  Indeed, we observed elevated microglia densities and dense clustering of mGFP+ OPCs specifically localized 140  to the fasciculus gracilis, along with robust astrogliosis.  In summary, injury-induced oligodendrogenesis remote to the lesion may be attributed to a multitude of factors, including (i) microgliosis, (ii) altered neural activity, (iii) changes in extracellular levels of regulatory factors, and/or (iv) injury-induced alteration in local biophysical properties.  Notably, the level of oligodendrogenesis observed in the uninjured spinal cord (5-7% of all CC1+ oligodendrocytes over a 14-week period) is in close agreement with previous studies (4-5% turnover over a 12-week period) (Barnabé-Heider et al. 2010; Young et al. 2013).  Moreover, cervical and lumbar turnover rates were highly similar, interesting considering the regional heterogeneity among adult OPCs (including cell cycle time) (Dimou & Gallo 2015; Nishiyama et al. 2009; Clarke et al. 2012; Kang et al. 2010).  Interestingly, myelin debris was observed in the cervical fasciculus gracilis at 12 wpi, a region of concentrated oligodendrogenesis.  Myelin debris potently inhibits oligodendrocyte differentiation and myelin gene expression, partially through the induction of Id2/4 expression (Kotter et al. 2006; Plemel et al. 2013).  Several theories may explain this seemingly paradoxical observation, including: (i) internalization of myelin debris by phagocytic microglia (thus obviating any direct effect on local OPCs), (ii) myelin debris and oligodendrogenesis may occupy mutually exclusive regions of the fasciculus gracilis, and/or (iii) oligodendrogenesis may precede myelin debris deposition due to the protracted nature of axonal degeneration in the CNS (Vargas & Barres 2007) (i.e. they are not temporally concurrent).  Further investigation may be warranted.  3.5 Maturation impairment of newly-formed oligodendrocytes Oligodendrocyte maturation impairment has been observed in post-mortem tissue of pre-term neonates (Billiards et al. 2008), animal models of diffuse white matter injury (DWMI) (Buser et al. 2012; Verney et al. 2012), and murine models of ALS (Kang et al. 2013).  Across these 141  pathologies, robust oligodendrogenesis normalizes committed oligodendrocyte numbers (i.e. CC1+ cells), but arresting of subsequent maturation leads to impaired myelination (Rousset et al. 2006; Kang et al. 2013), similar to what we observed remote to the thoracic contusion SCI lesion site.  Several plausible explanations can be proposed, divided into (i) extrinsic and (ii) intrinsic factors.  3.5.1 Extrinsic factors 3.5.1.1 Altered neural activity Neural activity can influence oligodendrocyte maturation and myelination, however as oligodendrocytes myelinate fixed axons and inert nanofibers in vitro (Lee et al. 2012; Rosenberg et al. 2008; Hildebrand et al. 1993; Wang et al. 2008) it may not be a requirement.  Indeed, social isolation impairs oligodendrocyte maturation in the medial prefrontal cortex (mediated by erbB3-NRG1-III signaling) (Makinodan et al. 2012), axonal glutamate release induces myelin protein synthesis in associated oligodendrocytes (Wake et al. 2011), neural activity biases myelination towards active axons (through stabilization of nascent sheaths) (Hines et al. 2015; Mensch et al. 2015), and stimulation of layer V projection neurons thickens myelin sheaths and stimulates de novo myelination in the premotor cortex (Gibson et al. 2014).  Therefore, injury-induced activity changes remote to lesion (as discussed previously) may contribute to the maturation impairment.    3.5.1.2 Microgliosis/inflammation Various activated microglia-derived compounds are implicated in glial and neuronal damage, including MMPs, lipases, reactive species (e.g. O2, H2O2, OH-, NO, ONOO-), excitotoxins (e.g. glutamate, quinolonic acid), chemokines, and cytokines (Pang et al. 2003; Chew et al. 2013; 142  Jurewicz et al. 2005).  Maturing oligodendrocytes are particularly vulnerable to inflammatory insult due to high metabolic demand (Back et al. 2001; Criag et al. 2003; Plemel et al. 2014; Almad et al. 2011).  Indeed, intracerebral LPS injection in young rats induces microglial activation, hypomyelination, and myelin abnormalities (Pang et al. 2003; Chew et al. 2013), microglial-derived TNF-α and IL-1β contribute to myelin damage in models of cerebral hypoxia (Deng et al. 2008; Kaur et al. 2013), and a host of microglia-derived pro-inflammatory cytokines (e.g. IL-1/2/3, IFN-α/β/γ, TNF-α, and lymphotoxin) are found in demyelinating MS lesions (Benveniste 1997).  Of these factors, IFN-γ may be particularly detrimental.  Indeed, IFN-γ exposure in vitro induces oligodendrocyte apoptosis (Durafourt et al. 2012) and increases vulnerability to microglial-derived cytokines (via induction of caspase-1/4/7/8, Fas, and TNFR1 expression)  (Buntinx et al. 2004).  Moreover, IFN-γ-induced ER stress has been implicated in impaired oligodendrocyte maturation and myelination (Lin et al. 2006).  As we observed concurrent oligodendrogenesis and microgliosis, newly-formed oligodendrocytes are born into a potentially injurious inflammatory environment, correlated with maturation impairment.  Amplification of this effect in the cervical fasciculus gracilis may explain, at least in part, the occurrence of robust oligodendrogenesis but substantially decreased new myelination (combined with a lack of permissive substrate – see below).  Conversely, several pro-inflammatory cytokines (e.g. IL-1β, lymphotoxin, and TNF-α) are reported to facilitate remyelination (Arnett et al. 2001; Mason et al. 2001), inflammation is critical for remyelination in toxin-mediated demyelination models (Franklin 2002; Franklin & Goldman 2015), and a lack of inflammation in chronic MS lesion has been proposed as a cause of remyelination failure (Franklin 2002).  This dichotomy may reflect polarization of microglial/macrophages into M1 (pro-inflammatory) and M2 (anti-inflammatory) phenotypes (Miron et al. 2013; Franklin & Goldman 2015).  The persistence of the M1 phenotype is 143  characteristic of SCI pathology (Kigerl et al. 2009), attributed to elevated intracellular iron levels (due to hemorrhaging) and TNF-α signaling at the lesion epicentre (Kroner et al. 2014).  Activated M1 microglia-derived pro-inflammatory cytokines (e.g. TNF-α, IL-1β, IL-2, and IL-17) are detrimental to immature oligodendrocytes both in vivo and in vitro (Favrais et al. 2011; Paintlia et al. 2011; Pang et al. 2010; Steelman & Li 2011; Taylor et al. 2010).  Indeed, TNF-α induces mitochondrial dysfunction in maturing oligodendrocytes (Bonora et al. 2014) and promotes astrogliosis (J. Li et al. 2008).  Activated microglia may also impair oligodendrocyte maturation via cytokine-independent mechanisms.  For example, activated microglia reduce production of key trophic factors for oligodendrocyte lineage development and/or maintenance (e.g. IGF-1, CNTF) (Albrecht et al. 2007; Pang et al. 2010; Talbott et al. 2007) and M1 microglia-derived free radicals (e.g. NO) induce mitochondrial dysfunction in oligodendrocytes (Haynes et al. 2003; Rousset et al. 2006).  In contrast to the M1 phenotype, anti-inflammatory M2 microglia promote oligodendrocyte differentiation in vitro and stimulate remyelination in vivo (Miron et al. 2013).    Attribution of impaired oligodendrocyte maturation to detrimental inflammatory effects would predict M1 persistence remote to lesion, which may warrant further research.    3.5.1.3 Astrogliosis Astrogliosis is also implicated in oligodendrocyte maturation impairment (Buser et al. 2012; Haynes et al. 2003; Verney et al. 2012).  Reactive astrocyte-derived factors are known to impair oligodendrocyte maturation (including BMPs, endothelin-1, TNF-α, adenosine, hyaluronan, IL-1β) (Deng et al. 2014; Hammond et al. 2014; Su et al. 2011; Wang et al. 2011).  In particular, the glycosaminoglycan hyaluronan (observed to accumulate in MS lesions) potently inhibits oligodendrocyte maturation (Back et al. 2005; Dean et al. 2011; Sloane et al. 2010) and adenosine-144  mediated AR1 receptor activation induces white matter damage and ventriculomegaly in neonatal rat models of hypoxia (Ma et al. 2011; Turner et al. 2003; Turner et al. 2002).  As AR1 is highly expressed in immature oligodendrocytes (Turner et al. 2003; Turner et al. 2002), activation may selective impair the maturation process.  Reactive astrocytes also reduce glutamate uptake.  Indeed, astrocytic expression of GLT-1 and GLAST (i.e. glutamate transporters) is reduced in several CNS pathologies (Desilva et al. 2008; Loeliger et al. 2003; Raymond et al. 2011).  Sustained extracellular glutamate levels would propagate excitotoxic insult, particularly detrimental to oligodendrocyte maturation.  Indeed, glutamate-induced excitotoxicity (due to reduced GABAergic axo-glial signaling) is implicated in oligodendrocyte maturation impairment in a murine model of hypoxia-induced WMI (Zonouzi et al. 2015).  We observed robust astrogliosis localized to the cervical fasciculus gracilis correlated with substantially reduced new myelination, consistent with impaired maturation of newly-formed oligodendrocytes.    3.5.1.4 Oxidative stress In vitro exposure to oxidative stress reduces the expression of key pro-differentiation genes (e.g. Sox10, Olig1, Olig2) while increasing the expression of inhibitory genes (e.g. Id2/4), inducing oligodendrocyte maturation failure (French et al. 2009).  Furthermore, in a neonatal rat model of intrauterine growth retardation, oxidative stress inhibits oligodendrocyte maturation and delays myelination (Reid et al. 2012).  Oxidative stress may therefore contribute to the observed maturation impairment remote to the lesion.    145  3.5.1.5 Reduced substrate permissiveness for myelination Axonal expression of GGF-2, Jagged-2, PSA-NCAM, vitronectin and activity-dependent effects on oligodendrocyte maturation imply that the axon is unlikely to be a passive participant in myelination (Charles et al. 2000; Canoll et al. 1996; Franklin et al. 2002; Chang et al. 2016).  Indeed, axonal expression of inhibitory factors (e.g. Jagged2 and PSA-NCAM) are thought to control developmental myelination onset (Wang et al. 1998; Charles et al. 2000; Emery & Lu 2015) as well as inhibit remyelination in MS (Franklin et al. 2002).  It is plausible that pathology may render axons less permissive for myelination, contributing to oligodendrocyte maturation impairment.  We observed extensive axonal damage (as revealed by β-APP immunostaining) and overt axonal loss remote to the lesion.  Indeed, new myelination was substantially reduced in the fasciculus gracilis (region of concentrated axonal pathology) despite robust oligodendrogenesis.  In addition to damage/loss, reduced neural activity of spared axons may impair myelination, as stabilization of nascent myelin sheaths is thought to be activity-dependent (Hines et al. 2015; Mensch et al. 2015).  Reduced activity may decrease successful stabilization (thus subsequent myelination), interpreted as maturation impairment.  Indeed, maturation impairment of newly-formed oligodendrocytes in a murine ALS model is associated with retraction of nascent sheaths from spared axons (consistent with unsuccessful stabilization on non-permissive substrates) (Kang et al. 2013).  Reactive astrocyte-derived CSPGs and inflammatory environments can induce conduction blockages of transiting (but intact) axons (Arvanian et al. 2009; Hunanyan et al. 2010; Redford et al. 1997).  Moreover, OPCs and extracellular NG2 also alter physiological properties of axons.  OPC processes contact nodes of Ranvier (Butt et al. 1999), both exogenous (Hunanyan et al. 2010) and endogenous NG2 (Martin et al. 2001) accumulates at nodes of Ranvier, and  NG2 administration reduces EPSP amplitude in the ventrolateral fasciculus of the uninjured spinal cord 146  (reversed by anti-NG2 antibodies) (Hunanyan et al. 2010; Petrosyan et al. 2013).  As activated OPCs increase NG2 production (Levine et al. 1994), extracellular NG2 levels are likely elevated in regions of OPC proliferation remote to the lesion.  Therefore, the combinatorial effect of robust inflammation (i.e. microgliosis), focal astrogliosis (presumably correlated with CSPGs), and increased NG2 deposition may reduce action potential conduction in transiting (but spared) axons.  As Iba-1 immunoreactivity was increased across the whole-cord section, and microglia-derived factors are diffusible (i.e. cytokines, growth factors, etc.), microglia-induced OPC proliferation and differentiation would not be restricted to damaged axonal tracts, thus newly-formed oligodendrocytes would be born in proximity to spared axons.  Indeed, we observed robust OPC proliferation and oligodendrogenesis across regions containing both spared and intact axons (based on expected murine spinal cord anatomy and β-APP immunostaining).  As oligodendrocytes have been shown to myelinate paraformaldehyde-fixed axons as well as nanofibers in vitro (Lee et al. 2012; Rosenberg et al. 2008; Hildebrand et al. 1993; Wang et al. 2008), a degree of new myelination might be expected even in damaged axonal tracts.  Indeed, we did observe a minimal amount of new myelination in the cervical fasciculus gracilis.  3.5.2 Intrinsic factors 3.5.2.1 Transcriptional dysregulation Oligodendrocyte maturation requires the induction of several distinct ‘waves’ of genes (e.g. MBP, PLP, MAG) (Dugas et al. 2007; Cahoy et al. 2008), reliant on central transcriptional regulators.  Indeed, deletion of several key transcription factors (e.g. Nkx2-2, Olig1, Ascl1, YY1, Zfhx1b, Sox10, and MyRF) stalls oligodendrocyte differentiation and/or maturation prior to myelination onset (Qi et al. 2001; Stolt et al. 2002; Xin et al. 2005; He et al. 2007; Sugimori et al. 2008; Emery 147  et al. 2009; Weng et al. 2012).  In particular, MyRF is critical for maturation, inducing the expression of cytoskeletal genes, lipid metabolism genes, transcription factors (e.g. Smad7, Nkx6-2), and myelin proteins (e.g. MBP, PLP) (Emery & Lu 2015; Yu et al. 2013).  MyRF induction during early differentiation depends on both Sox10 and Olig2 activity (Emery et al. 2009; Hornig et al. 2013).  Indeed, Sox10, Olig2, and MyRF display a close functional relationship, displaying substantial overlap in target sequences (Emery & Lu 2015; Hornig et al. 2013).  It is therefore feasible that pathologically-induced desynchrony of Sox10, Olig2, and MyRF activity in newly-formed oligodendrocytes may impair maturation.  Indeed, oxidative stress reduces Sox10, Olig1, and Olig2 expression in vitro (French et al. 2009).  Furthermore, oligodendrocyte expression of MOG, CNPase, MBP and PLP in vitro is significantly attenuated by exposure to IL-1β and TNF-α, as well as H2O2 and diamide (reactive oxidants) (Jana et al. 2005; Jana et al. 2013).  Moreover, systemic injection of IL-1β disrupts key regulatory factors in oligodendrocyte maturation, including Olig2, Olig1, Sox10, Tcf4, Axin2, HDAC1/3, Nkx2-2, Sox8, and p27Kip1 (Favrais et al. 2011).  Pathologically-induced Sox10/Olig2 disruption leading to failed MyRF induction would explain, at least in part, the observed maturation impairment.    3.5.2.2 Signaling dysregulation Injury-induced dysregulation of key signaling pathways, leading to imbalance, may contribute to the impaired maturation.  Wnt ligands act through Axin-1 and β-catenin to activate Tcf-4, which promotes differentiation of OPCs into pre-oligodendrocytes (Guo et al. 2015; Fancy et al. 2009; Fancy et al. 2011), but potently inhibits subsequent maturation into myelinating oligodendrocytes (Guo et al. 2015).  Indeed, elevated Wnt7a/7b levels are implicated in white matter damage in murine models of hypoxic injury (Yuen et al. 2014).  In addition to Wnts, BMP4-signaling inhibits 148  oligodendrocyte maturation in multiple ways, including (i) downregulation of key maturation-promoting factors (e.g. Olig1/2), (ii) upregulation of inhibitory factors (e.g. Id2/4), (iii) inhibition of myelin protein expression and (iv) increased accessibility of Wnt and Notch target genes via attenuated HDAC activity (Feigenson et al. 2011; See et al. 2004; Cheng et al. 2007; Samanta & Kessler 2004; Wu et al. 2012).  Extracellular BMP levels are increased in several CNS pathologies, including SCI (produced by reactive astrocytes) (Wang et al. 2011) and intrauterine growth retardation (attributed to oxidative stress) (Reid et al. 2012).  Moreover, BMP antagonism (via Noggin) protects white matter from perinatal hypoxic injury (Dizon et al. 2011).  Another potential pathway involved is mammalian target of rapamycin (mTOR) signaling, which regulates protein synthesis, cell proliferation, differentiation, and survival (Laplante & Sabatini 2012).  mTOR signaling (particularly mTORC1) is an important regulator of oligodendrocyte development, specifically the transition from immature oligodendrocyte to mature myelinating oligodendrocyte, as well as myelination extent (Dai et al. 2014; Guardiola-Diaz et al. 2012; Tyler et al. 2011; Wahl et al. 2014; Bercury et al. 2014).  Reduced IGF-1 signaling (mediated through mTOR) is implicated in oligodendrocyte maturation arrest in diffuse WMI (Hansen-Pupp et al. 2011; Hansen-Pupp et al. 2013), IGF-1 protects oligodendrocytes from inflammatory injury in vitro (Paula et al. 2014; Pang et al. 2007), and IGF-1 administration attenuates inflammatory-induced hypomyelination in murine models (Cai et al. 2011; Pang et al. 2010; Ye et al. 2007).  Thyroid hormone (TH) signaling also regulates oligodendrocyte maturation, promoting the expression of myelination-associated genes (both in vitro and in vivo) (Berbel et al. 1994; Billon et al. 2002; Bernal 2007).    149  3.5.3 Absence of maturation impairment at lesion epicentre? As newly-formed oligodendrocytes at lesion epicentre are born into a more severe pathological environment (Tator 1995; Tator & Fehlings 1991; Almad et al. 2011), an interesting question arises: why doesn’t a similar maturation impairment occur at the lesion epicentre?  Firstly, it may, but has not been reported.  If more oligodendrocytes are produced than needed to complete remyelination (Powers et al. 2013; Lasiene et al. 2008; Plemel et al. 2014), impaired maturation might have been overlooked.  Overproduction of oligodendrocytes occurs during development (Barres & Raff 1999), followed by refinement of cell numbers to match myelination requirements.  Intense physiological demand for remyelination following SCI may reactivate latent developmental programs predisposed to a developmental-like oligodendrocyte overproduction (to ensure remyelination completion).  Indeed, remyelination is thought to recapitulate developmental myelination, albeit with subtle deviations (Fancy et al. 2011).  Alternatively, the specific combination of factors at the lesion epicentre (e.g. growth factors, cytokines, chemokines, reactive species) might differentially effect maturing oligodendrocytes, as has been previously demonstrated for OPC-responsiveness in different environments (Bansal et al. 1996; Robinson & Miller 1999; Mason & Goldman 2002).    3.6 Oligodendrocyte apoptosis remote to the lesion site Oligodendrocyte apoptosis is a prominent feature of murine, non-human primate, and human SCI, contributing to the extensive oligodendrocyte loss at lesion epicentre (~50% by 1 dpi; ~93% by 7 dpi; murine models) (McTigue et al. 2001; Grossman et al. 2001; Lytle & Wrathall 2007).  Chronic oligodendrocyte apoptosis (2-3 wpi) along degenerating axonal tracts is attributed to loss of axon-derived trophic support (Casha et al. 2001; Crowe et al. 1997), however additional secondary 150  injury processes are thought to be required (specifically oxidative stress) (Sun et al. 2010; Almad et al. 2011).  In the current injury model, it is likely that both axonal damage/degeneration (confirmed by β-APP immunostaining) and oxidative stress (inferred from microgliosis and astrogliosis) are present remote to the lesion.  Indeed, we observed pre-existing apoptotic oligodendrocytes at 2 wpi (Casha et al. 2001; Crowe et al. 1997).  Various reactive microglia-derived factors induce oligodendrocyte death in vitro, including free radicals, pro-inflammatory cytokines, glutamate, and proteases (Buntinx et al. 2004; Jurewicz et al. 2005; Hövelmeyer et al. 2005).  Moreover, lower level exposure to microglia-derived factors (e.g. IFN-γ) increase oligodendrocyte vulnerability to reactive astrocyte-derived coupounds, oxidative stress, and/or growth factor depletion by proliferating cells (i.e. microglia or OPCs) (Buntinx et al. 2004; Lin et al. 2006).  What is the extent of cervical and lumbar oligodendrocyte loss following thoracic contusion injury?  Although OPC proliferation was rapidly increased (1 dpi for lumbar, 5 dpi for cervical) and maintained through 2 wpi, OPC density was not changed at 12 wpi, presumably due to the concomitant increase in oligodendrogenesis (as OPC apoptosis was not apparent).  If increased oligodendrogenesis compensates for lost oligodendrocytes (as might be expected from maintenance of committed oligodendrocyte density), cervical loss may be substantial as there is robust oligodendrogenesis (~2-fold increase in new oligodendrocytes at 12 wpi).  By contrast, lumbar oligodendrocyte loss may be less, as committed oligodendrocyte density was increased at 12 wpi despite a similar increase in oligodendrogenesis (i.e. ~2-fold), although lumbar tissue atrophy may complicate this interpretation.  Recurrent loss of newly-formed oligodendrocytes would also contribute to maintenance of cervical oligodendrocyte numbers despite continual oligodendrogenesis, as observed in models of ALS (Kang et al. 2013) and DWMI (Buser et al. 2012; Verney et al. 2012).  Indeed, apoptotic newly-formed oligodendrocytes were observed in 151  both the cervical and lumbar spinal cord at protracted times after injury (i.e. 2 wpi).  This may be attributed to maturation impairment, as observed in models of ALS (Kang et al. 2013).  Similarly, OPC-specific MyRF knockout (thus inhibiting maturation) leads to apoptosis of adult-born oligodendrocytes in murine models of contusion SCI and chemical-mediated demyelination (S. Manesh and G. Duncan, personal communication).  Unfortunately, quantification of apoptosis was unsuccessful as Casp3+ cells were scarce.  Utilization of a more rigous apoptotic stain (e.g. TUNEL) (Gorczyca et al. 1993) and/or increased sampling may address this in future.  As chronic autophagic oligodendrocyte death is also noted following SCI (>3 wpi) (revealed through beclin-1 immunostaining) (Kanno et al. 2009; Kanno et al. 2011), exclusive use of apoptotic markers may underestimate total oligodendrocyte loss.    3.7 OPC lineage plasticity: reliability of immunohistochemical markers It is widely accepted that OPCs generate mature myelinating oligodendrocytes (as revealed by genetic fate-mapping) in cortical stab injury (Dimou et al. 2008; Komitova et al. 2011), SCI (Barnabé-Heider et al. 2010), demyelination (Tripathi et al. 2010; Zawadzka et al. 2010), and ALS models (Kang et al. 2010).  Whether OPCs generate cells of other neural lineages is of great interest.  Conflicting reports exist for OPC astrogenesis, with low levels reported in SCI and TBI models (Barnabé-Heider et al. 2010; Komitova et al. 2011; Zawadzka et al. 2010), contrasted with complete absence in similar models of SCI, TBI, as well as other CNS pathologies (Dimou et al. 2008; Kang et al. 2010; Simon et al. 2011).  We did not observe any convincing evidence for OPC-derived astrocytes remote to the lesion site.  This analysis relied on Aldh1L1 and GFAP as astrocyte markers in immunohistochemical analyses.  Aldh1L1 (a folate metabolic enzyme) is also expressed by a NSPC population in the adult CNS that produces adult-born neuroblasts of the 152  rostral migratory stream (as revealed by genetic fate-mapping) (Foo & Dougherty 2013).  GFAP is considered the prototypical marker for immunohistochemical astrocyte identification, reliably labeling the majority (if not all) of reactive astrocytes in the injured CNS (Sofroniew & Vinters 2010).  Importantly, astrocytic GFAP expression is regionally heterogeneous and does not label all non-reactive astrocytes (e.g. in the healthy CNS or remote to a lesion) (Sofroniew 2009).  It is therefore feasible that GFAP- non-reactive OPC-derived astrocytes may exist remote to lesion, however subsequent Aldh1L1 immunostaining should have revealed their presence.  In addition to astrocytes, several CNS cell types that are defined as an extended ‘astroglial family’ express GFAP (Sofroniew & Vinters 2010), as do radial glial cells in adult neurogenic regions (i.e. hippocampal dentate gyrus and sub-ventricular zone) (Doetsch et al. 1999; Garcia et al. 2004; Imura et al. 2003; Seri et al. 2001).  As we did not observe any mGFP+/GFAP+ cells, this non-exclusivity does not complicate our interpretation.  In contrast to sparse evidence for astrogenesis, reports of OPC-derived Schwann cells are more substantive.  Genetic fate-mapping revealed OPC-derived Schwann cells in models of chemical demyelination (Zawadzka et al. 2010) and at the lesion epicentre in contusion SCI (Assinck et al. in revision.; Bartus et al. 2016).  Consistent with restriction to the lesion epicentre, we did not observe mGFP+/P0+ cells in either the cervical or lumbar spinal cord.  As P0 is only expressed by myelinating Schwann cells (Martini et al. 1995; D’Urso et al. 1990), it is feasible that OPC-derived pre-myelinating (i.e. P0-) Schwann cells may be present.  Immunostaining for p75-NTR would address this (Bentley & Lee 2000; Nickols et al. 2003), however this would also label motor neurons (Koliatsos et al. 1993).    153  3.8 Murine dorsal column sensory afferents: somatotopic vs. modality-based organization The murine cervical dorsal column is divided into a medial fasciculus gracilis and lateral fasciculus cuneatus (containing sensory afferents originating from DRG neurons below and above T6, respectively), located above the dorsal cortical spinal tract (dCST) running at the base (Watson 2009; Watson & Harrison 2012).  The prevailing view on dorsal column organization is the ‘somatotopic map’ model, which postulates that somatosensory fibres entering at successive rostral levels ascend laterally to those from lower spinal levels, supported by electrophysiological (Nord 1967; Johnson et al. 1968; Whitsel et al. 1969; Whitsel et al. 1970; Culberson & Brushart 1989), dye tracing (Maslany et al. 1991; Giuffrida & Rustioni 1992), and selective lesion studies (Smith & Deacon 1984).  Alternatively, a modality-based organization has been proposed (Uddenberg 1968; Dykes et al. 1982; Hummelsheim et al. 1985), with a somatotopic map existing within the same modality (i.e. proprioception vs. mechanosensory), with overlying ‘rough’ somatotopy (Niu et al. 2013).  The implications of this alternative view for the current study are limited as differential axonal damage between the fasciculus gracilis and fasciculus cuneatus (as assumed by a somatotopic organization) was confirmed by β-APP and SMI-312/NF-200 immunostaining.  As the purpose of sampling from these tracts was to assess the influence of axonal damage/degeneration on oligodendrocyte lineage cells, this does not substantially affect the presented interpretation.   3.9 Oligodendrogenesis in degenerating axonal tracts: maladaptive repair response? The revelation of robust oligodendrogenesis in the cervical fasciculus gracilis poses a conundrum: why produce new oligodendrocytes in a region replete with axonal injury/loss?  Indeed, enhancing 154  oligodendrogenesis without appropriate targets for myelination would appear to be maladaptive, and potentially detrimental (discussed below).      3.9.1 Activity-independent pathological stimulation of oligodendrogenesis Oligodendrogenesis in response to increased neural activity is thought to reflect a physiological demand for new myelination to optimize neural network function (Gibson et al. 2014; McKenzie et al. 2014; Chang et al. 2016), however this is presumably absent from damaged/degenerating axonal tracts.  Indeed, new myelination was substantially reduced in the fasciculus gracilis.  Interestingly, enhanced oligodendrogenesis localized to degenerating axonal tracts occurs in dorsal rhizotomy (Sun et al. 2010), axotomy (Nielson et al. 2006), and ALS models (Kang et al. 2013).  It is plausible that in contrast to activity-dependent signals reflecting a physiological demand for new myelination, aberrant oligodendrogenesis may also be triggered by activity-independent signals arising from pathological changes (irrespective of myelination demand), representing a pathologically-induced decoupling of oligodendrocyte developmental regulation (discussed below).  Indeed, robust oligodendrogenesis is observed in ischemic lesions (Zhang et al. 2013), neurodegeneration (Kang et al. 2013), and other pathologies in which axons and/or neurons are lost (Sun et al. 2010; Nielson et al. 2006).  Axonal degeneration-induced microglial activation, as well as astrogliosis, may be involved in this response, as reactive glia-derived factors are known to promote oligodendrogenesis (as discussed previously).   3.9.2 Detrimental effects of continual oligodendrogenesis? The production of new oligodendrocytes in an environment that does not need them is not only maladaptive and energetically wasteful, but also potentially damaging.  Indeed, it has been 155  proposed that continual cycling of OPC proliferation, differentiation, and subsequent death due to maturation failure among gray matter oligodendrocytes in a mouse model of ALS may accelerate damage to vulnerable axons by consuming resources (e.g. growth factors) and propagating reactive changes in other glial cells (e.g. microglia or astrocytes) (Kang et al. 2013).  Indeed, oligodendrocyte loss and/or damage activates microglia.  This would presumably require a form of death other than apoptosis (as parceling of cellular constituents into apoptotic bodies largely avoids immune activation) (Benedict et al. 2002).  Indeed, autophagic oligodendrocyte death is observed after SCI (Kanno et al. 2009; Kanno et al. 2011).  Furthermore, damaged oligodendrocytes release nucleotides that stimulate microglia purinergic receptors (e.g. P2X, P2Y), leading to microgliosis (Peferoen et al. 2014; Fields & Burnstock 2006).  Therefore, continual production and subsequent damage/loss of newly-formed oligodendrocytes may propagate detrimental inflammation through a positive feedback cycle (i.e. axonal damage/degeneration » microglial activation » oligodendrogenesis » newly-formed oligodendrocyte damage/loss » microglial activation » oligodendrogenesis, etc.).  Indeed, the pattern of microgliosis and OPC proliferation in the cervical fasciculus gracilis is consistent with a positive feedback cycle, as they both continually rise through 2 wpi.  The point at which oligodendrocyte maturation stalls (e.g. pre-myelinating stage in MyRF knockout) (Emery et al. 2009) prior to loss may be relevant.  For example, if nascent myelin sheaths are formed that contain myelin proteins (as seen in ALS models) (Kang et al. 2013), subsequent oligodendrocyte loss would generate myelin debris, which inhibits oligodendrocyte differentiation and axonal sprouting/regeneration (Kotter 2005; Kotter 2006; Plemel et al. 2013; Silver et al. 2015).  We did not observe any mGFP+ myelin debris (i.e. derived from newly-formed oligodendrocytes), but a more thorough assessment may be warranted. 156   3.9.3 Alternative non-myelin related function of the OPC response? It is feasible that the observed OPC proliferative response is elicited for another purpose than myelination, thus labeling it as ‘maladpative’ may be premature.  Indeed, several characteristics suggest that OPCs might have roles other than precursors of myelinating oligodendrocytes, including: (i) high cell density in the adult CNS (~5-8% of all neural cells), (ii) continual proliferation (i.e. not a dormant progenitor population), (iii) surveillance behaviour (i.e. continual movement through parenchyma, extension/retraction of processes), (iv) ability to closely monitor neural activity (via axo-glial synapses), and (iv) response to a variety of insults that do not involve demyelination as a primary feature (e.g. neurodegeneration) (Nishiyama et al. 2009; Hughes et al. 2013; Levine 2016; Dimou & Gallo 2015).  Indeed, targeted ablation of prefrontal cortical NG2+ glial cells (most likely OPCs) induces glutamatergic signaling deficits, associated with the emergence of depressive behaviours (Birey et al. 2015).  As OPC-derived bFGF is also implicated in astrocytic glutamate uptake (Birey et al. 2015) which is compromised in the injured CNS (Desilva et al. 2008; Loeliger et al. 2003; Raymond et al. 2011), increased OPC densities along degenerating axons may promote astrocytic glutamate uptake to mitigate secondary injury.  Therefore, the apparent maladaptive repair response seen in ALS (Kang et al. 2013), SCI (Sun et al. 2010), axotomy (Nielson et al. 2006), and dorsal rhizotomy (Sun et al. 2010) (as well as the current study) might be beneficial (at least in some regards), with oligodendrogenesis a side-effect of the OPC proliferative response.  Indeed, oligodendrocyte differentiation is density- and spatial-dependent (Rosenberg et al. 2008) and OPCs have tight spatial regulatory mechanisms that may promote differentiation when local densities are increased (Hughes et al. 2013).  This is purely speculative, but would be an interesting avenue for further research. 157   3.9.4 Pathological-induced decoupling of oligodendrocyte developmental regulation? Oligodendrocyte differentiation in vivo is considered terminal, requiring tight regulation of different developmental stages to ensure proper myelination and to maintain a sufficient post-natal OPC reservoir (Emery & Lu 2015).  It is plausible however that pathological effects may decouple this synchronous regulation.  Indeed, conduction block (via tetrodotoxin) is sufficient to induce OPC proliferation, but inhibits differentiation (Gautier et al. 2015).  Similarly, the current study suggests that axonal injury may stimulate OPC proliferation/differentiation despite the presumed absence of physiological demand for new myelination and potential injurious environment for maturing oligodendrocytes.  A largely similar response is observed in mouse models of ALS, as there is increased OPC proliferation and oligodendrogenesis, accompanied by maturation impairment, apoptosis, and myelin abnormalities (Kang et al. 2013).  This may be highly relevant for studies on the pathophysiology of demyelinating diseases (e.g. MS), as it is feasible that dysfunction at any stage of development could result in remyelination failure (e.g. impairment in OPC proliferation, differentiation, maturation, or myelination).  Indeed, impaired differentiation has been proposed as a cause of remyelination failure in MS (Franklin et al. 2002).  3.10 Remote effects of isolated SCI on oligodendrocyte lineage cells: potential relation to cognitive deficits observed in SCI patients? SCI patients display deficits in several cognitive functions (e.g. span memory, executive function, attention, processing speed, and learning), usually overlooked due to presumption of concomitant TBI (e.g. head trauma in motor vehicle accident or sports injury) (Davidoff et al. 1992; Roth et al. 1989; Dowler et al. 1995; Dowler et al 1997; Lazzaro et al. 2013; Jensen et al. 2007; Murray et al. 158  2007; Strubreither et al. 1997).  The demonstration of similar cognitive deficits in murine models subjected to an isolated thoracic injury directly implicates SCI-induced pathology (Wu et al. 2014a; Wu et al. 2014b).  Indeed, the observed cognitive deficits were associated with widespread inflammation and microglia activation across various brain regions (e.g. cortex, thalamus, hippocampus) (Wu et al. 2014a; Wu et al. 2014b).  The revelation of robust injury-induced oligodendrocyte lineage cell responses remote to the lesion raises the intriguing possibility that similar changes might occur in the brain.  As changes in myelination are linked to learning and cognitive functions (Bengtsson et al. 2005; Scholz et al. 2009; Zatorre et al. 2012; Chang et al. 2016), it is tempting to speculate that injury-induced changes in oligodendrocyte lineage cells might underlie neural circuit dysfunction manifesting as cognitive deficits.  It would be interesting to investigate injury-induced changes in the oligodendrocyte lineage cell populations of brain regions correlated with cognitive function (e.g. hippocampus, prefrontal cortex, etc.) following isolated SCI, specifically the extent of de novo myelination, OPC proliferation, oligodendrogenesis, and maturation of newly-formed oligodendrocytes.  Furthermore, as ablation of OPCs from the pre-frontal cortex is associated with glutamatergic signaling deficits (Birey et al. 2015) it would be interesting to correlate changes in the OPC population with neural circuit function (as revealed by electrophysiological and optogenetic approaches).   3.11 Proposed model of experimental observations To summarize, the following model is proposed to explain the observations presented here: (i) axonal damage/degeneration stimulates widespread microgliosis and focal astrogliosis remote to the injury site, (ii) microglia- and astrocyte-derived cytokines, altered neural activity, changes in biophysical parameters of the local microenvironment, and/or injury-induced changes in the 159  extracellular milieu stimulate OPC proliferation and oligodendrogenesis (even in the absence of demand for new myelination), (iii) newly-formed oligodendrocytes are born into a potential injurious environment (e.g. inflammation, astrogliosis, oxidative stress, axonal damage, etc.), which impairs their maturation (potentially due to intrinsic transcriptional dysregulation and/or lack of permissive substrate for myelination), (iv) the lack of axons prevents establishment of trophic interactions needed for maturation and survival, (v) culminating in apoptosis of a portion of newly-formed oligodendrocytes that do not mature to myelinating oligodendrocytes, potentially propagating tissue damage.  3.12 Conclusion Spinal cord injury (SCI) is a debilitating condition that imposes physical, psychological, financial, and emotional burdens on patients, families, caregivers, and the health system infrastructure at large (Dumont et al. 2001; Norenberg et al. 2004; Couris et al. 2009).  The development of therapeutic interventions that mitigate symptomatic severity is of significant interest.  To the best of our knowledge, the data presented here characterizes a previously unappreciated aspect of SCI pathophysiology (i.e. injury-induced changes in remote oligodendrocyte lineage cell populations).  In that regard, there are several notable findings to consider in the larger context of SCI research, including: (i) isolated injuries to the spinal cord have significant and rapid (~1 dpi) effects on multiple endogenous cell populations (e.g. microglia, OPCs, astrocytes) remote to the lesion site, (ii) maturation impairment among newly-formed oligodendrocytes may limit new myelination remote to the lesion, thus placing constraints on the amount of myelin plasticity that can occur in the post-injury environment,  (iii) maladaptive repair responses (i.e. OPC proliferation and oligodendrogenesis) may be stimulated by axonal injury and propagate tissue damage, and (iv) 160  pathological conditions may perturb regulation of oligodendrocyte development, decoupling synchrony of stage specific regulation.  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