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The role of protein tyrosine phosphatase alpha (PTPalpha) in oligodendrocyte development and CNS myelination Wang, Pei-Shan 2011

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The Role of Protein Tyrosine Phosphatase Alpha (PTPalpha) in Oligodendrocyte Development and CNS myelination  by  Pei-Shan Wang M.Sc., National Yang-Ming University, 2000  A THESIS SUBMITTED IN PARTIAL FULFILLMENT OF THE REQUIREMENTS FOR THE DEGREE OF  DOCTOR OF PHILOSOPHY  in  The Faculty of Graduate Studies (Pathology and Laboratory Medicine)  THE UNIVERSITY OF BRITISH COLUMBIA (Vancouver)  April 2011  © Pei-Shan Wang, 2011  ABSTRACT  Oligodendrocytes are specialized cells of the central nervous system (CNS) that are responsible for axonal myelination. They are derived from precursors termed oligodendrocyte progenitor cells (OPCs). In OPCs, the signal transduction pathways that regulate self-renewal versus differentiation remain poorly defined. The tyrosine kinase Fyn plays a critical role in oligodendrocyte differentiation and myelination in the CNS. However, the upstream molecules that regulate Fyn activity in these cells are not well characterized. In these studies, I utilized two model systems, the rat CG4 oligodendroglial cell line and a genetically modified mouse with germline PTP! knock-out, to investigate the role of protein tyrosine phosphatase alpha (PTP!) in OPC differentiation. My results demonstrate that PTP! is required for OPC differentiation and functions in at least two distinct phases during the lifespan and maturation of OPCs. Firstly, upon induction of OPC differentiation, Fyn activation and signaling are significantly reduced in the absence of PTP!, as measured by enhanced negative regulatory phosphorylation of Fyn and reduced activation or inhibition of Fyn downstream effectors such as focal adhesion kinase, Rac1, Cdc42, and Rho. Furthermore, myelination is defective in the brains of PTP!-/- mice, suggesting that OPC maturation requires PTP!-mediated signal transduction cascades. Secondly, OPCs must cease proliferation in order for differentiation to occur, and I found that PTP!-/- cells exhibit enhanced proliferation, as well as decreased cell cycle exit and increased survival. Interestingly, the activities of Ras and the Rho GTPases Rac1, Cdc42, and Rho were significantly increased, and p27 protein levels were significantly decreased, in the absence of PTP!. Moreover, Fyn is responsible for Rho inactivation and p27 accumulation. In conclusion, I propose that PTP! negatively regulates OPC proliferation and promotes cell cycle exit, causing  !  ""!  cell fate commitment. Subsequently, PTP! positively regulates and promotes differentiation and maturation. These functions of PTP! in OPC proliferation and differentiation processes are mainly exerted through PTP!-mediated dephosphorylation and activation of Fyn to induce the process-specific regulation of common and distinct Fyn effector molecules.  !  """!  PREFACE  The research project presented in this dissertation was originally developed in Dr. Catherine J. Pallen's laboratory in collaboration with Dr. Xiao. I have participated in experimental design under Dr. Pallen's supervision and conducted all listed experiments. I was also responsible for data collection and analysis, as well as preparation of manuscripts for peerreviewed journal publication. Mouse colony was maintained by Dr. Jing Wang. List of Publication: 1. Wang PS, Wang J, Xiao ZC, Pallen CJ. (2009) Protein-tyrosine phosphatase ! acts as an upstream regulator of Fyn signaling to promote oligodendrocyte differentiation and myelination. Journal of Biological Chemistry. 284:33692-702. The data included in this manuscript is located in Chapter 2, 3 and 4. Animal care and use followed the guidelines of the University of British Columbia and the Canadian Council on Animal Care, and were reviewed and approved by the University of British Columbia Animal Care Committee with Certificate Number A10-0194 (Neurobiology of PTP Alpha) and with Certificate Number A10-0122 (PTP Alpha Mouse Breeding).  !  "#!  TABLE OF CONTENTS  ABSTRACT................................................................................................................................ ii PREFACE.................................................................................................................................. iv TABLE OF CONTENTS ...........................................................................................................v LIST OF FIGURES ....................................................................................................................x LIST OF ABBREVIATIONS ................................................................................................. xii ACKNOWLEDGEMENTS .....................................................................................................xv 1. INTRODUCTION.................................................................................................................1 1.1. Oligodendrocyte development and CNS myelination...............................................1 1.1.1. Structure and function of the myelinated axon................................................1 1.1.2. Origin of oligodendrocyte progenitor cells .......................................................2 1.1.3. Migration of oligodendrocyte progenitor cells .................................................3 1.1.4. Proliferation of oligodendrocyte progenitor cells ............................................4 1.1.5. Differentiation stages of oligodendrocyte progenitor cells..............................4 1.1.6. Control of the numbers of oligodendrocyte progenitor cells and mature oligodendrocytes..................................................................................................6 1.1.7. Oligodendrocyte-related diseases ......................................................................7 1.1.8. Models of demyelination.....................................................................................8 1.2. Protein tyrosine phosphatases (PTPs) in the development and myelination of myelinating glia ..........................................................................................................10 1.2.1. Protein tyrosine phosphatase (PTP) superfamily ..........................................10 1.2.2. Protein tyrosine phosphatase alpha (PTP!) ...................................................12 1.2.2.1. Structure of PTP! ..............................................................................12 1.2.2.2. Regulation of PTP!............................................................................13 1.2.2.3. Substrates of PTP! ............................................................................17 1.2.3. PTPs are involved in oligodendrocyte development and CNS myelination ........................................................................................................18 1.2.3.1. PTP" ....................................................................................................19 1.2.3.2. CD45....................................................................................................20  !  #!  1.2.3.3. SHP-1 (Src homology region 2 domain-containing phosphatase-1)....................................................................................21 1.2.4. PTPs are involved in PNS myelination ...........................................................22 1.3. Src Family Kinases (SFKs) in oligodendrocyte development and CNS myelination .................................................................................................................23 1.3.1. Structure and activation of the SFKs..............................................................24 1.3.2. SFKs are involved in receptor and adhesion molecule-mediated oligodendrocyte proliferation, migration and survival .................................27 1.3.2.1. PDGF receptor (PDGFR)..................................................................27 1.3.2.2. IGF-1 receptor (IGF-1R)...................................................................28 1.3.2.3. Thromboxane A2 receptor (TXA2 receptor)....................................29 1.3.2.4. Muscarinic acetylcholine receptor (mAChR)..................................29 1.3.2.5. Integrins ..............................................................................................30 1.3.3. Fyn is involved in receptor and adhesion molecule-mediated changes in oligodendrocyte differentiation and myelination...........................................32 1.3.3.1. Insulin-like growth factor-1 receptor (IGF-1R)..............................33 1.3.3.2. Transferrin receptor..........................................................................33 1.3.3.3. The common # chain of immunoglobulin Fc receptors (FcR#).....34 1.3.3.4. Large myelin associated glycoprotein (L-MAG).............................35 1.3.3.5. GPI-anchored proteins (F3/Contactin and NCAM) .......................36 1.3.3.6. Integrins ..............................................................................................37 1.3.3.7. Deleted in colorectal carcinoma (Dcc)..............................................38 1.3.3.8. Leucine-rich repeats and Ig domain-containing, Nogo receptorinteracting protein-1 (LINGO-1)......................................................39 1.3.3.9. Opioid receptor ..................................................................................40 1.3.4. Fyn signaling in oligodendrocyte morphological differentiation and myelination ........................................................................................................40 1.3.4.1. Tau.......................................................................................................41 1.3.4.2. Focal adhesion kinase (FAK) ............................................................41 1.3.4.3. Rho Family GTPases (RhoA, Rac1 and Cdc42)..............................42 1.3.4.4. Rho GTPase activating proteins (p190RhoGAP and  !  #"!  p250RhoGAP) ....................................................................................44 1.3.5. Fyn controls MBP expression via multiple mechanisms ...............................45 1.4. Rationale and hypothesis ............................................................................................46 2. MATERIALS AND METHODS .......................................................................................55 2.1. Materials .....................................................................................................................55 2.1.1. Mice ....................................................................................................................55 2.1.2. Reagents .............................................................................................................55 2.1.3. Antibodies ..........................................................................................................55 2.1.4. Growth factors ..................................................................................................56 2.2. Cell lines and primary cell cultures..........................................................................56 2.2.1. OLN-93 cells ......................................................................................................56 2.2.2. CG4 cells ............................................................................................................57 2.2.3. Primary mouse neural progenitor/stem cells and OPCs ...............................57 2.2.3.1. Isolation and culture of primary OPCs from postnatal mice..........57 2.2.3.2. Isolation and culture of primar neural progenitor/stem cells and OPCs from mouse embryos ..................................................................58 2.3. Immunofluorescence labeling of cells and tissues ...................................................60 2.4. Immunoblotting .........................................................................................................61 2.5. Immunoprecipitation.................................................................................................61 2.6. siRNA transfection.....................................................................................................62 2.7. Ras and Rho family GTPase activities .....................................................................62 2.8. Cell cycle analysis and BrdU incorporation assay ..................................................63 2.9. Data analysis...............................................................................................................63 3. ESTABLISHMENT AND CHARACTERIZATION OF OLIGODENDROGLIAL CELL MODEL SYSTEMS................................................................................................65 3.1. Oligodendroglia cell lines ..........................................................................................65 3.1.1. Characterization of OLN-93 cell differentiation............................................66 3.1.2. Characterization of CG4 cell differentiation..................................................66 3.1.3. Ablation of PTP! by RNA interference (RNAi) in OLN-93 and CG4 cell lines ..............................................................................................................67 3.2. Characterization of primary mouse oligodendroglia cells isolated and  !  #""!  cultured from postnatal day 1 (P1) mice..................................................................68 3.3. Characterization of primary mouse neural progenitor/stem cells and OPCs isolated and cultured from mouse embryos.............................................................70 3.3.1. Forebrain ...........................................................................................................71 3.3.2. Spinal cord.........................................................................................................73 3.4. Discussion ...................................................................................................................74 3.5. Summary.....................................................................................................................75 4. PROTEIN TYROSINE PHOSPHATASE ! ACTS AS AN UPSTREAM REGULATOR OF FYN SIGNALING TO PROMOTE OLIGODENDROCYTE PROGENITOR CELL DIFFERENTIATION ................................................................84 4.1. Introdcution and rationale ........................................................................................84 4.2. Results .........................................................................................................................85 4.2.1. PTP! is required for CG4 differentiation......................................................85 4.2.2. PTP! is required for the activation of Fyn and the Fyn effectors FAK, Rac1, and Cdc42 during CG4 differentiation ................................................87 4.2.3. PTP! is not required for Fyn-mediated signaling to p190RhoGAP, but is required for Rho inactivation during CG4 differentiation .......................89 4.2.4. PTP! is required for primary mouse OPC differentiation ..........................90 4.2.5. PTP! selectively regulates Fyn activation and signaling in primary mouse OPCs.......................................................................................................91 4.2.6. Ablation of PTP! results in decreased MBP protein expression in primary mouse OLs and leads to defective myelination ...............................93 4.3. Discussion ...................................................................................................................94 4.4. Summary.....................................................................................................................98 5. PROTEIN TYROSINE PHOSPHATASE ! IS A NEGATIVE REGULATOR OF OLIGODENDROCYTE PROGENITOR CELL SELF-RENEWAL .........................112 5.1. Introduction and rationale ......................................................................................112 5.2. Results .......................................................................................................................113 5.2.1. Abnormal population of OPCs and oligodendrocytes in the corpus callosum of PTP!-/- mice.................................................................................113 5.2.2. PTP! negatively regulates OPC growth and growth factor !  #"""!  dependency ......................................................................................................114 5.2.3. PTP! negatively regulates OPC proliferation, cell cycle entry and survival in response to PDGF/bFGF .............................................................116 5.2.4. Lack of PTP! does not affect proliferation of neural stem/progenitor cells ...................................................................................................................117 5.2.5. PTP! negatively regulates the activities of Ras and Rho Family small GTPases, Cdc42, Rac1 and Rho, in OPCs ....................................................118 5.2.6. PTP! negatively regulates protein expression of p27Kip1 (p27)................119 5.2.7. Hyperproliferation of PTP!-/- OPCs is not due to upregulation of PDGFR! ..........................................................................................................120 5.2.8. Higher Ras activity in PTP!-/- OPCs is not due to sequesteration of the negative regulator p120 RasGAP by higher levels of FAK.........................121 5.2.9. PTP! negatively regulates multiple signaling pathways and is required for p27 accumulation in CG4 cells ......................................................................122 5.2.10. Fyn is required for Rho inactivation and p27 accumulation in CG4 cells ...................................................................................................................123 5.3. Discussion .................................................................................................................124 5.4. Summary...................................................................................................................130 6. GENERAL DISCUSSION ...............................................................................................145 6.1. The role of PTP! in OPC self-renewal and cell fate commitment......................145 6.2. The role of PTP! in OPC differentiation ..............................................................149 6.3. Future direction .......................................................................................................150 REFERENCES........................................................................................................................155  !  "$!  LIST OF FIGURES Figure 1.1 Schematic representation of a myelinated axon................................................49 Figure 1.2 Schematic representation of the major stages in the development of oligodendrocytes...................................................................................................50 Figure 1.3 Schematic representation of the PTP superfamily ...........................................51 Figure 1.4 Structure and phosphorylation sites in PTP!...................................................52 Figure 1.5 Structure and activation of SFKs .......................................................................53 Figure 1.6 SFKs signalings in oligodendrocyte proliferation and migration ...................54 Figure 2.1 Flow chart of isolation and culture of primary mouse neural progenitor/ stem cells and OPCs ...........................................................................................64 Figure 3.1 Characterization of OLN-93 cell differentiation...............................................76 Figure 3.2 Characterization of CG4 cell differentiation.....................................................77 Figure 3.3 Ablation of PTP! by siRNA in two oligodendroglial cell clines......................78 Figure 3.4 Characterization of primary mouse oligodendroglia cells differentiation .....79 Figure 3.5 Characterization of primary mouse NSC differentiation (forebrain) ............80 Figure 3.6 Characterization of primary mouse OPC differentiation (forebrain) ............81 Figure 3.7 Characterization of primary mouse NSC differentiation (spinal cord)..........82 Figure 3.8 Characterization of primary mouse OPC differentiation (spinal cord) .........83 Figure 4.1 Protein expression of PTP! and SFKs and activity of Fyn in CG4 cells during differentiation ..........................................................................................99 Figure 4.2 PTP! is required for CG4 morphological differentiation .............................100 Figure 4.3 PTP! is required for CNPase expression in CG4 cells upon differentiation.....................................................................................................101 Figure 4.4 PTP! is required for activation of Fyn and its downstream effectors FAK in CG4 cells.........................................................................................................102 Figure 4.5 PTP! is required for activation of Rac1 and Cdc42 in CG4 cells.................103 Figure 4.6 PTP! is not required for tyrosine phosphorylation of p190RhoGAP (p190) and Fyn-p190 interaction in CG4 cells ............................................................104 Figure 4.7 PTP! is required for Rho inactivation in CG4 cells ......................................105  !  $!  Figure 4.8 Protein expression of PTP!, Fyn and CNPase in primary mouse OPCs upon differentiation.....................................................................................................106 Figure 4.9 CNPase expression decreased in PTP!-null OPCs upon differentiation .....107 Figure 4.10 Differentiation of PTP!-null OPCs is impaired .............................................108 Figure 4.11 PTP! is a regulator of Fyn-FAK signaling in OPCs and OLs ......................109 Figure 4.12 PTP! is not a regulator of Fyn-p190 signaling in OPCs and OLs................110 Figure 4.13 Decreased MBP expression in PTP! -/- OLs and PTP! -/- mouse brain.........111 Figure 5.1 Abnormal population of OPCs and oligodendrocytes in PTP! -/- mouse brain ....................................................................................................................132 Figure 5.2 Increased growth of OPCs in the absence of PTP! ........................................133 Figure 5.3 Decreased growth factor dependency of OPCs in the absence of PTP!.......134 Figure 5.4 Increased proliferation of OPCs in the absence of PTP! ..............................135 Figure 5.5 Decreased cell cycle exit and apoptosis of OPCs in the absence of PTP! ....136 Figure 5.6 Lack of PTP! does not affect proliferation and cell cycle exit in neural stem cells ......................................................................................................................137 Figure 5.7 Increased activities of Ras and Rho family small GTPases, Cdc42, Rac1 and Rho, in PTP! -/- OPCs ........................................................................................138 Figure 5.8 Phosphorylation of ERK1/2 and Akt were not affected in the absence of PTP! ...................................................................................................................139 Figure 5.9 Decreased expression of p27 in PTP! -/- OPCs and differentiating OLs.......140 Figure 5.10 PDGFR! expression is not affected by the absence of PTP! ........................141 Figure 5.11 Increased protein level of FAK in PTP! -/- cells does not promote FAK-p120 RasGAP association to sustain Ras activity ....................................................142 Figure 5.12 PTP! negatively regulates Ras, Rho GTPases, MAPK and Akt and is required for p27 accumulation .........................................................................143 Figure 5.13 PTP!-Fyn signaling is required for Rho inactivation and p27 required for p27 accumulation...............................................................................................144 Figure 6.1 Proposed mechanisms by which PTP! regulates OPC proliferation and differentiation.....................................................................................................153 Figure 6.2 Proposed mechanisms by which PTP! regulates p27 accumulation............154  !  $"!  LIST OF ABBREVIATIONS A2RE  A2 response element  bFGF  basic fibroblast growth factor  C/EBP  CCAAT-enhancer-binding protein  Cbp/PAG  Csk-binding protein or Phosphoprotein Associated with glycosphingolipid-enriched microdomains  Cdc42  cell division control protein 42 homolog  CDK  cyclin-dependent kinase  CNPase  2', 3'-cyclic nucleotide 3'-phosphodiesterase  CNS  central nervous system  CNTF  ciliary neurotrophic factor  Csk  c-src tyrosine kinase  CXCL1  chemokine (C-X-C motif) ligand 1  Dcc  deleted in colorectal carcinoma  DMEM  Dulbecco's Modified Eagle Medium  EAE  experimental autoimmune encephalomyelitis  ECM  extracellular matrix  EDTA  ethylenediaminetetraacetic acid  EGF  epidermal growth factor  ES  embryonic stem  Eya  eyes absent  FAK  focal adhesion kinase  FcR#  the commom # chain of immunoglobulin Fc receptor  FRET  cluorescence resonance energy transfer  FTS  S-trans, transfarnesylthiosalicylic acid  GFAP  glial fibrillary acidic protein  GFP  green fluorescent protein  GPI  glycosylphosphatidylinositol  !  $""!  HBSS  Hank's Buffered Salt Solution  HGF  hepatocyte growth factor  hnRNP  heterogeneous nuclear ribonucleoprotein  Ig  immunoglobulin  IGF-1  insulin-like growth factor 1  IL-6  interleukin-6  JNK/SAPK  stress-activated protein kinase or c-Jun NH2-terminal kinase  LINGO-1  leucine-rich repeats and Ig domain-containing, Nogo receptorinteracting protein-1  LMW  low molecular weight  mAChR  muscarinic acetylcholine receptors  MAG  myelin Associated Glycoprotein  MAM  meprin-A5-mu  MAP2  microtubule-associated protein 2  MAPK  mitogen-activated protein kinases  MBP  myelin basic protein  MOG  myelinoligodendrocyte glycoprotein  MS  multiple slerosis  NCAM  neural Cell Adhesion Molecule  NCM  neural culture medium  NMDA  N-Methyl-D-aspartic acid  NRG  neuregulin  NT-3  neurotrophin-3  OL  oligodendrocyte  OPC  oligodendrocyte progenitor cell  p27  p27Kip1  PBS  phosphate buffered saline  PCNA  proliferating Cell Nuclear Antigen  PDGF  platelet-derived growth factor  PDGFR  platelet-derived growth factor receptor  PDL  poly-D-lysine  !  $"""!  PDLO  poly-DL-ornithine  PI3K  Phosphatidylinositol 3-kinases  PKC  protein kinase C  PLP  proteolipid protein  pNPP  p-nitrophenyl phosphate  PP2A  Protein phosphatase 2A  PSA  polysialic acid  PTK  protein tyrosine kinase  PTP  protein tyrosine phosphatase  PVDF  polyvinylidene Fluoride  QKI  quaking homolog, KH domain RNA binding  Rac1  Ras-related C3 botulinum toxin substrate 1  RhoGAP  Rho GTPase activating protein  SDS-PAGE  sodium dodecyl sulfate polyacrylamide gel electrophoresis  SFK  src family kinase  SH1  Src Homology 1  SH2  Src Homology 2  SH3  Src Homology 3  SHP-1  Src homology region 2 domain-containing phosphatase-1  STAT3  signal transducer and activator of transcription 3  T3  triiodothyronine  TXA2  thromboxane A2  VEGF  vascular endothelial growth factor  WAVE2  Wiskott-Aldrich syndrome protein family member 2  WT  wild-type  !  $"#!  ACKNOWLEDGEMENTS Foremost, I would like to express my sincere gratitude to my supervisor, Dr. Catherine J. Pallen for her continuous support of my PhD study and research, as well as for her patience, enthusiasm, and immense knowledge. Her guidance has helped me tremendously in throughout research and thesis writing. I could not have imagined having a better mentor for my PhD study. In addition, I am deeply grateful to my supervisory committee: Dr. Janet Chantler, Dr. Peter van den Elzen, Dr. Cheryl Wellington and Dr. Vanessa Auld, for their efforts in reading my thesis as well as their insightful comments. All the lab members in Pallen lab made it an enjoyable place to work. In particular, I would like to thank Dr. Jing Wang for managing the lab, teaching me techniques and generating and maintaining mice for my studies in this thesis. Noteworthy, I really enjoyed all the delicious food made by Dr. Wang over the past four years. Last but not the least, I would like to thank my family: my husband Fu-Sheng Chou for his encouragement, support and help at every stage of my PhD study, and my parents Te-Lung Wang and Chin Chen and my brother Pei-Chang Wang for supporting me spiritually throughout my life.  !  $#!  Chapter 1. INTRODUCTION  1.1  Oligodendrocyte development and CNS myelination The vertebrate central nervous system (CNS) contains three major classes of  neural cells: neurons, astrocytes, and oligodendrocytes. Oligodendrocytes are the myelin-forming cells of the CNS. They are derived from oligodendrocyte progenitors that undergo a series of developmental stages before acquiring the mature myelinating function (Pfeiffer et al., 1993). Oligodendrocyte progenitors arise and proliferate in certain locations and subsequently migrate through the CNS and stop at their destination. The majority of oligodendrocyte progenitor proliferation occurs in developing white matter. After a sufficient number of oligodendrocyte progenitors have been generated, these cells differentiate into immature oligodendrocytes and mature into myelinating oligodendrocytes (Miller, 2002) Upon maturation, oligodendrocytes increase their expression of myelin associated molecules and assemble myelin sheaths around the appropriate axons. The myelin sheath is a fatty insulation composed of modified membrane that promotes the rapid conduction of electrical impulses along myelinated axons and prevents axonal degeneration (Bunge, 1968; Nguyen et al., 2009).  1.1.1 Structure and function of the myelinated axon Myelination of the axons is carried out by oligodendrocytes in the CNS, leading to the formation of myelinated regions intervened by the unmyelinated nodes of Ranvier. Clustering of Na+ channels in the nodes of Ranvier allows action potentials to occur only in the nodes, therefore increasing the conducting velocity. In addition,  !  1!  myelination greatly reduces ATP consumption in axons due to the restricted distribution of ATP-dependent ion channels in the nodes (Nave, 2010). Moreover, it has recently been shown that myelination effectively prevents axonal degeneration, which is possibly related to the ability of the myelinated axons to maintain sufficient ATP as an energy source for membrane repolarization to occur. A multiple sclerosis model demonstrated that, without proper repolarization, abnomal calcium entry may follow, leading to proteolysis and axonal degeneration (Edgar et al., 2009; Griffiths et al., 1998; Lappe-Siefke et al., 2003). On the edges of the myelinated axonal segments are paranodes and juxtaparanodes. The paranode is a specialized axon-glia contact zone that seals the internodal periaxonal space from the outside milieu. The abundant axonal adhesion proteins in this zone are linked via axonal scaffolding proteins to Na+ channels in the nodes of Ranvier and to K+ channels in the juxtaparanodes and interact with a variety of oligodendroglial surface molecules. (Fig. 1.1).  1.1.2 Origin of oligodendrocyte progenitor cells In the spinal cord, most oligodendrocytes arise from a pool of progenitors derived from the ventral ventricular zone, which proliferate and migrate dorsally and finally differentiate into myelin-forming oligodendrocytes. A dorsal source of progenitors also arises and contributes to spinal gliogenesis (Cai et al., 2005; Vallstedt et al., 2005). In the forebrain, three sequential waves of oligodendrocyte progenitors are generated (Kessaris et al., 2006). The first wave of oligodendrocyte progenitors arises in the medial ganglionic eminence and anterior entopeduncular area in the ventral forebrain.  !  2!  These progenitors populate the entire embryonic telecephalon including the cerebral cortex, and then are joined by a second wave of progenitors derived from the lateral and/or caudal ganglionic eminences. The third wave of progenitors arises within the postnatal cortex. These functionally redundant populations of progenitors compete for space in the developing brain. When one population is destroyed (i.e. by the targeted expression of diphtheria toxin), the remaining cells take over and restore the normal distribution of oligodendrocyte progenitors (Kessaris et al., 2006).  1.1.3  Migration of oligodendrocyte progenitor cells The oligodendrocyte progenitors have to travel long distances from the site of  origin to developing white matter. Migration of these progenitors is guided by regulatory signals including secreted molecules and contacts with extracellular matrices, axonal tracts and astrocytic surfaces. Growth factors including PDGF, bFGF, EGF, VEGF and HGF; extracellular matrix such as fibronectin; chemotropic molecules including netrins and semaphorins, and the chemokine CXCL1, have been shown to be involved in the migration of oligodendrocyte progenitors (de Castro and Bribián, 2005). For example, either fibronectin and merosin alone as well as in combination with PDGF promote migration, and tenascin-C selectively inhibits the migration of oligodendrocytes that respond to fibronectin but not to merosin (Frost et al., 1996). PSA-NCAM has been shown to be crucial for the PDGF-induced OPC migration (Zhang et al., 2004).  !  3!  1.1.4 Proliferation of oligodendrocyte progenitor cells The expansion of oligodendrocyte numbers occurs in the ventricular and subventricular zone, but the majority of oligodendrocyte proliferation occurs after migration to the developing white matter (Miller et al., 1997). Many factors have been identified as regulators of oligodendrocyte proliferation, such as growth factors including PDGF, bFGF, CNTF, IGF-1, NT-3, HGF and neuregulins; axonal contact molecules including Notch/Jagged; and extracellular matrix components including vitronectin and fibronectin (Baron et al., 2005; McTigue and Tripathi, 2008; Ohya et al., 2007). The best characterized mitogens for OPCs are PDGF, which induces OPCs to proliferate for a number of divisions, thereby preventing premature differentiation (Noble et al., 1988; Raff et al., 1988; Richardson et al., 1988) and bFGF, which maintains a high level of PDGFR! in OPCs and prevents differentiation into oligodendrocytes (Bogler et al., 1990; McKinnon et al., 1990; McKinnon et al., 1993). The control of oligodendrocyte progenitor number is discussed in section 1.1.6. After a sufficient number of oligodendrocyte progenitors have been generated, these progenitors differentiate and mature into myelin-forming cells.  1.1.5 Differentiation stages of oligodendrocyte progenitor cells The progression of oligodendrocyte progenitors to mature oligodendrocytes can be divided into four stages: early oligodendrocyte progenitors, pro-oligodendrocytes, immature oligodendrocytes, and mature oligodendrocytes (Deng and Poretz, 2003). Oligodendrocyte progenitors are bipolar, migratory, proliferative cells (Fig. 1.2). They elaborate more processes as they mature into pro-oligodendrocytes. The PDGF receptor  !  4!  ! (PDGFR!) is expressed by progenitors at very early stages of oligodendrocyte development (McMorris and McKinnon, 1996). Along with the PDGFR!, NG2 proteoglycan is detected on progenitors and pro-oligodendrocytes (Dawson et al., 2000). In addition, a number of glycolipids are detected on oligodendroglial lineage cells. The monoclonal antibody A2B5 (that detects several gangliosides) is used to identify the bipolar progenitors (Pfeiffer et al., 1993). The pro-oligodendrocytes can be detected by the monoclonal antibody O4, which recognizes sulfatides and an unidentified sulfated glycoconjugate (Bansal et al., 1992). The immunoreactivity toward O4 is retained throughout oligodendrocyte development. The monoclonal O1 antibody, which recognizes galactocerebroside, monogalactosyldiglycerides and an unidentified antigen, can be used to identify immature oligodendrocytes, which are post-mitotic cells with more complex morphology. One of the earliest oligodendrocyte specific proteins is 2’,3’-cyclic nucleotide 3’-phosphodihydrolase (CNPase), which can be detected on immature oligodendrocytes. Finally, the oligodendrocytes enter terminal differentiation and extend membrane-sheets to enwrap axons. The mature oligodendrocytes express many oligodendrocyte-specific proteins, including myelin basic proteins (MBP), proteolipid protein (PLP) and myelin-associated glycoproteins (MAG) (Fig. 1.2) (Pfeiffer et al., 1993).  1.1.6  Control of the numbers of oligodendrocyte progenitor cells and mature  oligodendrocytes In order to match the number of oligodendrocytes and axons, the proliferation, differentiation and survival of the oligodendrocyte lineage cells are tightly controlled.  !  5!  Several molecules have been demonstrated to have opposite effects on oligodendrocyte progenitor proliferation and differentiation (Deng and Poretz, 2003; McTigue and Tripathi, 2008), such as PDGF, bFGF, CNTF, HGF, neuregulins and Notch/Jagged. These molecules promote proliferation and inhibit differentiation of oligodendrocyte lineage cells to generate sufficient numbers of oligodendrocyte progenitors. In the absence of mitogens and survival factors, oligodendrocytes will differentiate or die, respectively (Temple and Raff, 1985). The survival of oligodendrocyte lineage cells is regulated by growth factors and axonal contacts. Several growth factors that promote oligodendrocyte proliferation also promote survival of these cells, such as PDGF, bFGF, IGF-1, NT-3, CNTF and neregulins (Baron et al., 2005; Deng and Poretz, 2003; Yasuda et al., 1995). Newly formed oligodendrocytes compete for the limiting amount of survival signals from target cells that prevent apoptosis and then mature into myelinforming oligodendrocytes. About half of the oligodendrocytes will die due to failure to contact axons (Barres et al., 1992a). Laminin-2 is one of the contact-mediated survival signals from axons, and it amplifies the survival signals from PDGF and neuregulins upon binding to "6"1 integrins of oligodendrocytes (Baron et al., 2005). Laminin-2 is also an important axonal signal that promotes oligodendrocyte maturation and myelination, since the binding of laminin-2 to "6"1 integrins switches the neuregulin signaling from PI3K-mediated proliferation to MAPK-mediated differentiation. In addition, laminin-2/integrin interactions enhance process extension and myelin membrane formation (Buttery and ffrench-Constant, 1999; Chun et al., 2003; Colognato et al., 2004). Neuregulin-1 is another axonal signal that regulates myelination, but the role of neuregulin-1 in CNS myelination is still not clear  !  6!  (Brinkmann et al., 2008; Taveggia et al., 2008). Moreover, CNS myelination may also be determined by the degree of neuronal differentiation, since blockade of sodiumdependent action potentials inhibits myelination in developing optic nerves and increased neuronal firing enhances myelination (Demerens et al., 1996). Firing of action potentials leads to the release of adenosine and ATP, which promote oligodendrocyte differentiation and myelination, respectively (Ishibashi et al., 2006; Stevens et al., 2002).  1.1.7 Oligodendrocyte-related diseases In addition to enwrapping axons to enhance conduction of electrical impulses, oligodendrocytes have been demonstrated to have other functions, such as maintaining a healthy axon segment, regulating sodium channel clustering along axon segments, and providing trophic support for neuronal somas (McTigue and Tripathi, 2008). Recently, researchers found that there are heterogeneous populations of NG2-positive cells throughout the CNS. These cells not only serve as the primary source of remyelinating cells, but also have unique functions in the adult CNS including forming synapses with neurons and being able to generate action potentials (Karadottir et al., 2008; Nishiyama et al., 2009). In accord with the variety of functions of oligodendrocyte lineage cells, these cells have been indicated to be involved in numerous neurological disorders. The most well known human disease involving loss of oligodendrocytes is multiple sclerosis (MS), an inflammatory disease in which the myelin sheath of the brain and spinal cord are damaged. Imperfect development of the myelin sheath, mostly caused by gene mutations, such as in SURF1 and ARSA, results  !  7!  in leukodystrophies (Harvey et al., 1993; Rahman et al., 2001). In addition, virus infection, stroke and traumatic injury of the CNS also cause loss of oligodendrocytes and demyelination (Crowe et al., 1997; Emery et al., 1998; Fazakerley and Walker, 2003; Kovari et al., 2004). Oligodendrocyte damage and loss also occur in Alzheimer’s disease and Huntington’s disease (Bartzokis et al., 2007; Roth et al., 2005; Sjobeck et al., 2005). Altered expression of oligodendroglia-related genes (MAG, ERBB, TF, PLP1, MOG, MOBP), ultrastructural changes and altered oligodendroglia have been found in brain regions from patients with schizophrenia, bipolar disorders, major depressive disorders or substance abuse (Sokolov, 2007). However, little is known about the etiology and the relationship between oligodendrocytes and these diseases. Since oligodendrocytes and myelin have so many important functions, understanding the biology of oligodendrocytes and myelin and discovering new strategies for treatment are imperative.  1.1.8 Models of demyelination There are several experimental models for inducing demyelination in rodents, such as experimental autoimmune encephalomyelitis (EAE), and cuprizone and lysolecithin treatments. The EAE model has been described for over 70 years and publications using the EAE model has number more than 7000 articles. There are several rodent EAE models with different procedures and they exhibit some similarities and differences to MS, therefore, there is no general agreement on the right animal model (Gold et al., 2006; Krishnamoorthy et al., 2006). Generally, EAE is induced by injection of antigens, such as CNS myelin, MBP, MOG or PLP, into the brain or spinal  !  8!  cord, leading to autoimmune-mediated inflammation of the CNS. The first clinical signs of disease often present within 9-12 days of sensitization. Similarities to MS include T-cell inflammation and demyelination, but these models only cause transient or mild demyelination, or do not cause spontaneous disease, thus differening from MS. Neurotoxicants, such as cuprizone, can induce noninflammatory demyelination. Cuprizone is a copper chelator that can be administered in the diet, resulting in decreased oxidative phosphorylation of oligodendroglial mitochondria due to depletion of cytochrome oxidase activity (Carey and Freeman, 1983). Demyelination is observed after 3-6 weeks exposure to cuprizone (Carey and Freeman, 1983; Matsushima and Morell, 2001). Replacement with normal diet allows for complete remyelination within 4 weeks (Matsushima and Morell, 2001; Merrill, 2009). Lysolecithin is a detergent-like membrane solubilizing agent that induces myelin breakdown and apoptosis of oligodendrocytes (Blakemore et al., 2002; Shields et al., 1999; Wallace et al., 2003) and has been used to induce local demyelination by injection (Jeffery and Blakemore, 1995). Demyelination is observed one week post-injection followed by rapid remyelination (begining by 7 days after lesioning and complete by one month) (Jeffery and Blakemore, 1995; Merrill, 2009; Triarhou and Herndon, 1986).  1.2  Protein tyrosine phosphatases (PTPs) in the development and myelination of  myelinating glia Regulation of protein tyrosine phosphorylation is an key signalling mechanism to control different functions of eukaryotic cells. The relative activities of protein tyrosine  !  9!  kinases (PTKs) and PTPs regulate important signalling pathways that are involved in the control of cell proliferation, adhesion, migration, survival, differentiation, metabolism, cell-cell communication, ion channel activity and the immune response (Hunter, 1998). Disruption of this balance is associated with human diseases, including cancers (Ostman et al., 2006).  1.2.1  Protein tyrosine phosphatase (PTP) superfamily The PTP superfamily contains over 100 members, and a total of 107 PTPs are  encoded by the human genome (Alonso et al., 2004). They are defined by a unique signature motif, HC(X)5R, and can be divided into four classes based on their structure and substrate specificity: Class I, classical PTPs, dual-specificity PTPs; Class II, low molecular weight (LMW) PTPs; Class III, the cdc25 phosphatases and Class IV, the eye absent family of PTPs (EYA) (Julien et al., 2011; Wang et al., 2003) (Fig. 1.3). Among the PTP superfamily, 38 belong to the ‘classical’ PTPs that show specificity for phosphotyrosine (Alonso et al., 2004). Classical PTPs are divided into receptor-like forms and non-receptor forms (Paul and Lombroso, 2003). The receptorlike PTPs have a single transmembrane domain, variable extracellular domains and one or two cytoplasmic PTP domains. Most of the receptor-like PTPs contain two PTP domains (D1 and D2; see Fig. 1.3) with most, if not all, of the catalytic activity residing in the membrane proximal PTP domain D1. The function of the membrane distal domain is not well characterized, but has been suggested to play a role in substrate targeting (Andersen et al., 2001). The extracellular regions typically contain domains implicated in cell adhesion, including immunoglobulin-like repeats, fibronectin type III  !  10!  repeats, carbonic anhydrase-like domains, meprin-A5-mu (MAM) domains or cysteinerich regions (Beltran and Bixby, 2003; Brady-Kalnay et al., 1995). Non-receptor PTPs have a single catalytic domain and striking structural diversity and often contain sequences that target them to specific subcellular locations or enable their binding to specific proteins (Mauro and Dixon, 1994) (Fig. 1.3). Dual-specificity PTPs are a heterogeneous group of PTPs that dephosphorylate protein substrates on tyrosine, serine and threonine residues, as well as lipid substrates (Patterson et al., 2009). There are 61 dual-specificity PTPs of heterogeneous form and function, which can be grouped on the basis of the presence of specific domains and sequence similarity. LMW-PTPs are a group of 18-kDa enzymes that are widely expressed (Raugei et al., 2002), and these enzymes have been shown to interact with several receptor tyrosine kinases and docking proteins, including platelet-derived growth factor receptor (PDGFR) (Chiarugi et al., 1995), ephrin A2 receptor (Eph2A) (Kikawa et al., 2002), and "-catenin (Taddei et al., 2002). The cdc25 phosphatases are highly conserved dual specificity phosphatases, but are classified separately due to their distant structural relation to other PTPs (Wang et al., 2003). They activate cyclin-dependent kinase (CDK) complexes, which in turn regulate cell cycle progression, and are also key components of the checkpoint pathways that become activated in the event of DNA damage (Boutros et al., 2007). The transcription factor Eyes absent (Eya) possesses protein phosphatase activity and may regulate the phosphorylation state of either itself or its transcriptional cofactors (Jemc and Rebay, 2007). Eya is the prototype for a metal-dependent PTP family that differs from traditional thiol-based PTPs by using an aspartic acid rather than a cysteine as the nucleophile during catalysis (Alonso et al.,  !  11!  2004; Collet et al., 1998; Li et al., 2003; Rayapureddi et al., 2003; Selengut, 2001; Selengut and Levine, 2000; Tootle et al., 2003). Eya possess intrinsic phosphatase activity toward the low molecular weight synthetic substrate p-nitrophenyl phosphate (pNPP) and has specificity toward phosphotyrosine, although activity of Eya family proteins toward tyrosine phosphorylated peptides is lower than that of the classical PTP, PTP1B (Li et al., 2003; Rayapureddi et al., 2003; Tootle et al., 2003).  1.2.2  Protein tyrosine phosphatase alpha (PTP!)  1.2.2.1  Structure of PTP!  PTP! is a ubiquitously expressed transmembrane PTP that is highly enriched in brain and is the most abundant PTP in OLs (Ranjan and Hudson, 1996; Shock et al., 1995).! It is often designated as a receptor protein-tyrosine phosphatase (RPTP), however no extracellular ligand is known. Unlike other PTPs, PTP! has a short, heavily glycosylated extracellular domain (~123 residues) that lacks cell adhesion motifs (Fig. 1.4). Unglycosylated PTP! is a ~85 kD protein, and it becomes ~130 kD due to extensive N- and O- glycosylation. Like other PTPs, PTP! has two tandem cytoplasmic PTP domains (D1 and D2) of which the D1 domain (residues 241–500) is active and the D2 domain (residues 501–802) exhibits minimal (0.1%) activity (Kaplan et al., 1990; Wang and Pallen, 1991). The D1 and D2 domains not only differ in catalytic activity, but also in substrate recognition and specificity. For example, PTP!D2 possesses in vitro catalytic activity towards pNPP but towards phosphotyrosyl peptide substrates (Lim et al., 1997; Wu et al., 1997). Since PTP!-D2 exhibits minimal activity towards phosphotyrosine and no physiological substrate for PTP!-D2 has been  !  12!  found so far, the D2 domain may have a regulatory role rather than a catalytic one. Indeed, PTP!-D2 may act in a non-enzymatic capacity to form protein-protein interactions that could regulate direct substrate recognition or position PTP! in a multicomponent complex for D1-mediated hydrolysis. For example, PTP!-D2 interacts with the PDZ2 domain of PSD95 to enable PSD95-linked Src activation and regulation of NMDA receptor activity (Lei et al., 2002). In addition, PTP!-D1 can trans-associate with PTP!-D2, which could affect PTP! dimer formation and activity (Jiang et al., 1999). PTP! has three phosphorylation sites that exhibit regulatory roles, Ser 180/204 in the juxtamembrane region and Tyr 789 in the C-terminal tail (den Hertog et al., 1995; Stetak et al., 2001; Zheng et al., 2002; Zheng and Shalloway, 2001). This is described further in section 1.2.2.2.  1.2.2.2 Regulation of PTP! Dimerization PTP! activity may be negatively regulated by dimerization or intermolecular associations with other PTPs (Jiang et al., 2000). Crystallization of PTP!-D1 revealed a dimer formed by interaction of a helix-turn-helix at the N-terminal region of one PTP!-D1 monomer with the catalytic cleft of the other monomer (Bilwes et al., 1996). This interaction would interfere with substrate binding, suggesting that PTP! dimerization may inhibit its catalytic activity. Overexpression of PTP! allows detection of constitutively formed homodimers by FRET (fluorescence resonance energy transfer) and on gels after chemical cross-linking (Jiang et al., 2000; Tertoolen et al., 2001). However, the dimers are not detectable by co-immunoprecipitation assay  !  13!  (Blanchetot and den Hertog, 2000), suggesting that PTP! dimers may not be stable. More stable PTP! dimers can be induced by oxidation upon H2O2, UV or heat shock treatment due to inactivation of PTP!-D1 and conformational changes of PTP!-D2 (Blanchetot et al., 2002; van der Wijk et al., 2004). However, whether PTP! forms dimers at endogenous cellular levels remains unknown, because all the studies described above were conducted with cells overexpressing PTP!.  Phosphorylation PTP! activity can be stimulated by protein kinase C (PKC)-mediated serine phosphorylation on Ser180 and Ser204 (den Hertog et al., 1995; Tracy et al., 1995). The PKC isoform that is responsible for PTP! phosphorylation remains unknown, although PKC# is the only PKC isoform that can physically associate with PTP! (Stetak et al., 2001). Controversial results were reported describing the effects of phosphorylation of PTP! at Ser180 and/or Ser204 in regulation of mitosis, probably due to the different model systems that were used. Zheng et al. (Zheng et al., 2002; Zheng and Shalloway, 2001) used overexpression of PTP! and Src to demonstrate that serine phosphorylation of PTP! facilitates the activation of Src during mitosis by two mechanisms: one is to increase the specificity of PTP!, the other is to reduce Grb2 binding to PTP! and thus relieve Grb2-mediated inhibition of PTP!-catalyzed Src activation. These effects were abrogated by mutation of Ser180 and/or Ser204, indicating that phosphorylation of PTP! at Ser180 and/or Ser204 is required for Src binding and the mitotic reduction in PTP!-Grb2 association. However, in contrast, Vacaru et al. (Vacaru and den Hertog, 2010) recently reported that phosphorylation of  !  14!  PTP! at Ser204 was almost completely undetectable in mitotic NIH 3T3 and HeLa cells. They also found that the phosphorylation of PTP! at Ser180 and Ser204 did not significantly affect the intrinsic catalytic activity of endogenous PTP!. They proposed a new model for mitotic activation of Src in which PP2A-mediated dephosphorylation of  PTP! at  Ser204  facilitates  Src  binding,  leading  to  PTP!-mediated  dephosphorylation of Src at Tyr527 and Tyr416, and resulting in modest activation of Src. PTP! can also be phosphorylated at Tyr789. However, whether tyrosine phosphorylation at this site affects the catalytic activity of PTP! remains unclear, since conflicting results were reported by different groups. Some groups showed that Tyr789 phosphorylation has no effect on the catalytic activity of PTP! (Su et al., 1996; Zheng et al., 2000), while another group reported that PTP! activity decreased upon phosphorylation at this site (den Hertog et al., 1994). Src may be the kinase phosphorylating PTP! at Tyr789 since co-expression of Src and PTP! increased Tyr789 phosphorylation of PTP! (den Hertog et al., 1994). However, Hao et al. (Hao et al., 2006) demonstrated Src/Fyn/Yes-independent tyrosine phosphorylation of PTP! at Tyr789. Phosphorylated Tyr789 of PTP! has been shown to serve as binding site for the SH2 domain of other proteins, such as Grb2 and Src (den Hertog et al., 1994; Su et al., 1994; Zheng et al., 2000). Almost all tyrosine phosphorylated PTP! is found to be associated with Grb2 (den Hertog et al., 1994; Su et al., 1994). However, the Grb2 binding partner, Sos, has not been found in the complex of PTP!-Grb2, suggesting that PTP! may compete with other Grb2 binding proteins to negatively regulate Grb2mediated signaling (den Hertog and Hunter, 1996; den Hertog et al., 1994; Su et al.,  !  15!  1996). On the other hand, upon Grb2 binding, phospho-Tyr789 can be protected from dephosphorylation and can be prevented from interacting with other proteins. Although it has been shown that Src activation is dependent on phospho-Tyr789 of PTP! in some situations, phospho-PTP!-independent Src activation has been demonstrated in two different cell types, PC12 neuronal cells and embryonic fibroblasts, as demonstrated by a mutant form of PTP! lacking the Tyr789 residue (Y789F PTP!) (Chen et al., 2006; Kapp et al., 2003; Vacaru and den Hertog, 2010; Yang et al., 2002). Recently, it has been shown that integrin and IGF-1 stimulate PTP! Tyr789 phosphorylation, and phospho-Tyr789 of PTP! is required for cell migration (Chen et al., 2006; Chen et al., 2009). Therefore, PTP! tyrosine phosphorylation may have multiple roles and be involved in different signaling pathways.  Proteolysis The function of PTP! can also be regulated by proteolysis, since truncated PTP! (p66 PTP!) generated by calpain-mediated cleavage in vivo is unable to dephosphorylate Src and the Kv2.1 potassium channel (Gil-Henn et al., 2001). Treatment of living cells with a variety of calpain inhibitors resulted in significant reduction of cleavage (Gil-Henn et al., 2001). However, only a small proportion of PTP! is cleaved by calpain in primary cortical neurons. Moreover, the PTP! C433S mutant was cleaved to generate a 75 kD form of PTP! upon treatment with pervanadate, a non-specific inhibitor of PTPs (Lammers et al., 2000). This truncated form was no longer associated with focal adhesions. These findings indicate that PTP! can be regulated by proteolysis to alter its cellular localization.  !  16!  1.2.2.3  Substrates of PTP!  Src family kinases (SFKs) The SFKs comprise of nine members in vertebrates: Src, Yes, Fgr, Yrk, Fyn, Lyn, Hck, Lck and Blk (Thomas and Brugge, 1997b). The structure and activation of SFKs are discussed in section 1.3.1. PTP! dephosphorylates the C-terminal negative regulatory site (e.g. mouse Src Tyr527) of SFKs and thus activates SFKs (den Hertog et al., 1993; Zheng et al., 1992). PTP! has been shown to dephosphorylate and associate with Src, Fyn and Yes in cells expressing these SFKs and PTP! (Bhandari et al., 1998; Harder et al., 1998). PTP!-null mouse brain and embryonic fibroblasts exhibit only 30-50% Src and Fyn activities compared with wild-type (WT) tissue/cells, correlating with enhanced phosphorylation of Src and Fyn at Tyr527 (Ponniah et al., 1999; Su et al., 1999), indicating that PTP! is involved in SFK activation and downstream signaling.  Voltage-gated potassium channels (Kv channels) Kv channels are transmembrane channels specific for potassium and are sensitive to voltage changes. Four !-subunits form a functional, tetrameric Kv channel. Each !subunit contains six transmembrane helices (S1-S6), flanking the P region, which is responsible for ion conduction (Robertson, 1997). At least 11 main gene families of potassium channels have been characterized (Hugnot et al., 1996; Salinas et al., 1997). The largest group is formed by the Kv1 channels (Pongs, 1992). The members of this family are expressed in the brain at different levels (Veh et al., 1995). PTP! has been  !  17!  shown to participate in the m1 muscarinic acetylcholine receptor (mAChR)-mediated regulation of Kv1.2 activity (Tsai et al., 1999). The m1 mAChR induces the tyrosine phosphorylation of PTP! and PTP!-Kv1.2 association in 293 cells stably expressing the m1 AchR, the Kv1.2 ! subunit and the Kv"2 subunit. PTP! also binds the N-and C-terminal of Kv1.2 in vitro. Increased resting Kv1.2 current was observed in Xenopus oocytes overexpressing PTP!. In addition to Kv1.2, PTP! has been shown to regulate Kv2.1 activity (Tiran et al., 2006). PTP! dephosphorylates Kv2.1 in vitro and constitutively associates with Kv2.1 in HEK293 cells co-expressing Kv2.1 and PTP!. Moreover, increased Kv channel current in Schwann cells and increased phosphorylation of Kv2.1 in sciatic nerves of PTP!-null mice were demonstrated (Tiran et al., 2006), indicating that PTP! play a role in regulating Kv channel activity.  1.2.3 PTPs are involved in oligodendrocyte development and CNS myelination In oligodendrocyte lineage cells, 11 distinct PTPs have been detected using a differential display strategy (Ranjan and Hudson, 1996). Four major PTPs; PTP!, PTP", PTP# and PTP$, are expressed in both oligodendrocyte progenitors and oligodendrocytes, while PTP% is expressed in more differentiated cells (Krueger and Saito, 1992; Matthews et al., 1990; Ranjan and Hudson, 1996; Sahin and Hockfield, 1993; Walton et al., 1993; Yan et al., 1993). Inhibition of PTP activities by a tyrosine phosphatase inhibitor, orthovanadate, results in decreased CG4 differentiation, indicating the crucial roles of PTPs in oligodendrocyte differentiation (Ranjan and Hudson, 1996). Roles for several of these receptor PTPs in in vivo myelination have been described. CNS myelination appears normal in PTP$-null mice, although myelin  !  18!  stability may be reduced (Harroch et al., 2000).!However, remyelination is impaired in PTP$-null mice after experimental autoimmune encephalomyelitis (EAE)-induced demyelination (Harroch et al., 2002). Mice null for another receptor PTP, CD45, have reduced numbers of MAG-positive myelinating OLs and exhibit general, mild dysmyelination (Nakahara et al., 2005).!Myelination defects are also detected in mice lacking the cytosolic PTP SHP-1, and MBP expression is reduced in OLs from the mice (Massa et al., 2004).  1.2.3.1 PTP$ PTP$ (encoded by the PTPRB gene) is one member of a subfamily of PTP that contains a CAH (carbonic anhydrase homology) domain followed by fibronectin type III repeats in the extracellular domain. Three different isoforms of PTP$ are expressed as a result of alternative mRNA splicing: two transmembrane forms and a secreted form composed of only the extracellular domain of PTP$, also known as phosphacan (Krueger and Saito, 1992; Levy et al., 1993). PTP$ is expressed predominantly by astrocytes, oligodendrocytes, and Schwann cells but also by neurons throughout the developing and adult nervous system (Canoll et al., 1993; Shintani et al., 1998). Both transmembrane forms of PTP$ are predominantly expressed in glial progenitor cells, while phosphacan is expressed at high levels by more mature glial cells (Canoll et al., 1996b). These results suggest that the PTP$ expression is regulated during glial cell differentiation and different isoforms may have different functions. Moreover, glial PTP$ binds to a cell recognition complex on neurons, which consist of contactin, Caspr (Peles et al., 1997), and NCAM (Sakurai et al., 1997). Therefore, it was suggested that  !  19!  PTP$ is involved in myelination and formation of the node (Einheber et al., 1997). Harroch et al. (Harroch et al., 2000) generated mice deficient in the three isoforms of PTP$ and showed that these mice are viable, fertile and have no gross abnormalities. Although the ultrastructure of the paranodes remained normal in these mice, a fragility of the myelin was observed. Moreover, PTP$-deficient mice are more susceptible to experimental autoimmune encephalomyelitis (EAE), a model of multiple sclerosis (Harroch et al., 2002). These mice showed impaired recovery from EAE induced by myelin oligodendrocyte glycoprotein (MOG) peptide, and this may due to increased apoptosis of mature oligodendrocytes of mutant mice at the peak of inflammation. In addition, increased expression of the human homolog of PTP$, PTPRZ1, was observed in remyelinating oligodendrocytes in multiple sclerosis lesions. These results suggest that PTP$ may have a role in oligodendrocyte survival and in recovery from multiple sclerosis.  1.2.3.2 CD45 CD45 (encoded by the PTPRC gene) is a receptor-type PTP that contains three fibronectin type III repeats in the extracellular region (Okumura et al., 1996), and is expressed in all differentiated hematopoietic cells except erythrocytes and platelets (Thomas, 1989). The different CD45 isoforms generated from alternative splicing of a single gene contain different extracellular domains with an identical transmembrane and cytoplasmic region, and are expressed in a specific manner according to cell type and stage of differentiation (Koretzky, 1993). Nakahara et al. (Nakahara et al., 2005) demonstrated that CD45 expression increased in OPCs upon differentiation induced by  !  20!  immunoglobulin G (IgG), and FcR%-stimulated CD45-/- OPCs show impaired morphological differentiation and a lack of MBP upregulation. In normal OPCs, CD45 is complexed with Fyn and the Fyn negative regulatory kinase Csk, suggesting that CD45 is a candidate activator of Fyn and Fyn-MBP signaling in response to FcR% engagement. In P10 mouse forebrain, few CD45-positive cells were detected in the subventricular zone, where OPCs are generated from neural stem cells. Most of the CD45-positive cells are myelin-associated glycoprotein (MAG) positive and were located in the corpus callosum, the fimbria and the internal capsule, indicating that CD45  is  expressed  in  differentiating  oligodendrocytes  in vivo.  Moreover,  dysmyelination and reduced MAG-positive cells were observed in P10 CD45-deficient mice and persistent dysmyelination was observed in adult CD45-deficient mice. These results suggest that CD45 is crucial in oligodendrocyte differentiation and myelin maintenance.  1.2.3.3 SHP-1 (Src homology region 2 domain-containing phosphatase-1) SHP-1 (encoded by the PTPN6 gene) is a non-receptor type PTP containing two tandem SH2 domains, which mediate the interaction of this PTP with its substrates. Two different tissue-specific promoters regulate expression of the two forms of SHP-1 protein. One is active in all cells of non-hematopoietic origin, whereas the other is active exclusively in cells of hematopoietic lineage. SHP-1 is predominantly expressed in hematopoietic cells. It downregulates transmembrane receptor signaling, such as by growth factor receptors, cytokine receptors and receptors involved in the immune response (Wu et al., 2003). Massa et al. (Massa et al., 2000) first reported that SHP-1 is  !  21!  also expressed in oligodendrocytes and that SHP-1 negatively regulates IL-6 signaling as demonstrated by higher STAT3 phosphorylation and STAT3-responsive c-fos gene expression in response to IL-6 in oligodendrocytes lacking SHP-1. In addition, SHP-1 deficient mice (motheaten mice) exhibit dysmyelination as demonstrated by reduced MBP expression in the brain and spinal cord, decreased MBP immunoreactivity in the corpus callosum, optic nerve and trigeminal nerve and decreased CNPase immunoreactivity in the corpus callosum (Massa et al., 2000; Massa et al., 2004; Wishcamper et al., 2001). Moreover, the decreased percentage of myelinated axons and myelin sheath thickness in the spinal cord of SHP-1 deficient mice and decreased MBP expression in purified O1-positive oligodendrocytes lacking SHP-1 further support a role for SHP-1 as a critical regulator of terminal differentiation and myelin sheath formation by oligodendrocytes (Massa et al., 2004).  1.2.4 PTPs are involved in PNS myelination Although PTP! and PTP& have not yet been described in CNS myelination, they have been shown to be involved in PNS myelination (Peretz et al., 2000; Tiran et al., 2006). Two major isoforms of PTP& (encoded by the PTPRE gene) generated by two different promoters have been reported: one is a transmembranal, receptor-type PTP (RPTP&), which is closely related to PTP!; the other is a cytoplasmic PTP (cyt-PTP&) (Elson and Leder, 1995a; Elson and Leder, 1995b; Krueger et al., 1990; Nakamura et al., 1996; Tanuma et al., 1999). The differences in the N-terminal sequences determine their subcellular localization and physiological roles (Elson and Leder, 1995a). Both forms of PTP& bind Grb2 (Toledano-Katchalski and Elson, 1999). RPTP& has been  !  22!  linked to promotion of mammary tumorigenesis by dephosphorylating Src, and cytPTP& may regulate osteoclast function (Chiusaroli et al., 2004; Gil-Henn and Elson, 2003). Schwann cells express cyt-PTP& and the early post-natal (P5) PTP&-deficient mice exhibit hypomyelination of sciatic nerve as demonstrated by decreased myelin sheath thickness (Peretz et al., 2000). Similar results were observed in PTP!-deficient mice, and the myelination defects were more severe in PTP! and PTP& double knockout mice (Tiran et al., 2006). Lack of either PTP increases voltage-gated potassium channel (Kv) activity and phosphorylation in Schwann cells. PTP! inhibits Kv channels more strongly than PTP&, probably due to constitutive association of PTP! with Kv2.1. PTP!, but not PTP&, activates Src in sciatic nerve extracts. Taken together, both PTPs support PNS myelination but are not fully redundant.  1.3  Src Family Kinases (SFKs) in oligodendrocyte development and CNS  myelination Since SFKs are important substrates of PTP! (Pallen, 2003) and have been implicated in oligodendrocyte development and CNS myelination (as described below), this section describes the roles of SFKs in these events. The nonreceptor cytoplasmic tyrosine kinase Src was the first discovered protooncogene in the vertebrate genome and is an important regulator of many signaling pathways and cellular functions (Martin, 2001). Five members of the SFK family, Src, Fyn, Yes, Lyn and Lck, are expressed in brain (Thomas and Brugge, 1997b). Among these, Fyn plays a unique role in CNS myelination, since myelin deficits are only found in Fyn-/- mice, but not in Src-/-, Lyn-/- or Yes-/- mice (Sperber et al., 2001; Umemori et  !  23!  al., 1999). In addition, loss of Fyn on the C57BL/6 genetic background results in premature death, severe hydrocephalus and defects in oligodendrocyte development. These mice showed enlarged lateral ventricles with thinner cerebral cortices and degenerating axons in the corpus callosum. The number of oligodendrocytes was reduced and the morphogenesis of oligodendrocytes was impaired in the cerebral cortex (Goto et al., 2008). Recently, Buckley et al. (Buckley et al., 2010) showed that inhibition of SFK activities in zebrafish larvae by a SFK inhibitor PP2 resulted in a significant decrease in mbp transcripts that encode myelin basic protein (MBP), a major constituent of the myelin sheath in the nervous system. Src, Fyn, Lyn and Yes are expressed in oligodendrocyte lineage cells (Krämer et al., 1999; Miyamoto et al., 2008; Osterhout et al., 1999; Sperber et al., 2001; Wang et al., 2009). However, the expression patterns and activities of these SFKs in oligodendrocyte lineage cells are different. Fyn is the only SFK with significant kinase activity and with increased expression during differentiation (Osterhout et al., 1999; Wang et al., 2009), suggesting these SFKs may have different roles in oligodendrocyte development.  1.3.1  Structure and activation of the SFKs Members of the Src family are 52-62 kDa proteins and they share a common  structure composed of following functional domains (Guarino, 2010; Kim et al., 2009; Thomas and Brugge, 1997a) (Fig. 1.5A): (1) The N-terminal region contains a myristoylation sequence, which is essential for SFK localization at the inner surface of the cell membrane. (2) The unique sequence provides unique functions and specificity to each member of the Src family. (3) The SH3 domain and (4) the SH2 domain bind  !  24!  proline-rich sequences and phosphorylated tyrosine residues, respectively, to mediate protein-protein interactions. (5) The SH1 catalytic domain contains a positiveregulatory autophosphorylation site at Tyr416 (chicken Src), which is important for maximizing its kinase activity. (6) The C-terminal tail contains a negative-regulatory site at Tyr527 (chicken Src), which binds to the SH2 domain of the same protein when phosphorylated, resulting in a closed conformation that prevents the interaction with substrates. In general, the activity of SFKs can be regulated by phosphorylation and interaction with their binding proteins (Bjorge et al., 2000b; Thomas and Brugge, 1997b). The SH2 and SH3 domains play an essential role in the regulation of SFK catalytic activity (Pawson, 1997; Sicheri and Kuriyan, 1997; Xu et al., 1997). The first important phosphorylation site of Src is its C-terminal negative-regulatory site Tyr527 (Cooper et al., 1986; Kmiecik and Shalloway, 1987). When Src is in the inactive state, the phosphorylated Tyr527 at its C-terminal tail interacts with its SH2 domain (Liu et al., 1993; Roussel et al., 1991), and the SH2-SH1 linker interacts with the SH3 domain, resulting in a closed conformation of the kinase (Fig. 1.5B). Several upstream regulators can regulate the activity of SFKs through phosphorylating or dephosphorylating the negative-regulatory site of SFKs, such as tyrosine kinases and protein tyrosine phosphatases (PTPs) (Roskoski, 2005). The major tyrosine kinase responsible for phosphorylating this site is a cytoplasmic tyrosine kinase called c-Src kinase (Csk). A transmembrane phosphoprotein, Csk binding protein (Cbp) or phosphoprotein associated with glycosphingolipid-enriched microdomains (PAG), binds to and recruits Csk to the plasma membrane, where it can phosphorylate and  !  25!  inactivate Src (Brdicka et al., 2000; Kawabuchi et al., 2000). PTP!, PTP', SHP-1 (or PTP-1C), SHP-2, PTP-1B and CD45 are phosphatases that have been shown to dephosphorylate the negative-regulatory site of Src (Bjorge et al., 2000a; Fang et al., 1994b; Peng and Cartwright, 1995; Somani et al., 1997; Thomas and Brown, 1999; Wang et al., 2002; Zheng et al., 1992). The tyrosine kinase Csk and these phosphatases control the balance between Src phosphorylation/dephosphorylation, leading to upregulation or downregulation of Src activity (Nada et al., 1991; Okada et al., 1991; Zheng et al., 1992). The second important phosphorylation site of Src is Tyr416 in its SH1 kinase domain. Tyr416 can be phosphorylated by another Src or by other tyrosine kinases (Broome and Hunter, 1997; Chiang and Sefton, 2000; Gould and Hunter, 1988; Ralston and Bishop, 1985; Stover et al., 1996). Phosphorylation of this site allows Src to interact with its substrates (Xu et al., 1999; Xu et al., 1997). Src can also be dephosphorylated at this site by phosphatases, such as PTP!, PTP' and SHP-1 (Bjorge et al., 2000a; Fang et al., 1994a; Fang et al., 1994b; Somani et al., 1997; Zheng et al., 1992). Another mechanism of regulation of Src activity is through interaction with binding proteins, leading to activation of the kinase activity and relocalization of Src into proximity with its substrates (Bjorge et al., 2000a). Several Src-binding proteins have been shown to perturb the intramolecular interactions of Src, including PDGFR (Alonso et al., 1995; Kypta et al., 1990) and FAK (Cobb et al., 1994; Eide et al., 1995; Schaller et al., 1994), leading to Src activation. SFKs have been implicated in various receptor signaling pathways, such as those initiated by immune recognition receptors and major histocompatibility receptors, integrin and other adhesion receptors, receptor protein tyrosine kinases, G-protein-  !  26!  coupled receptors, cytokine receptors, GPI-linked receptors, voltage-gated and ligandgated channels (Thomas and Brugge, 1997a). Therefore, SFKs are important in a variety of cellular events, such as regulation of the cell cycle, apoptosis, differentiation, adhesion and migration (Thomas and Brugge, 1997a).  1.3.2  SFKs are involved in receptor and adhesion molecule-mediated  oligodendrocyte proliferation, migration and survival 1.3.2.1 PDGF receptor (PDGFR) One of the best-characterized mitogens for oligodendrocyte progenitors is PDGF (Baron et al., 2000), provided by neurons and astrocytes in the CNS (Richardson et al., 1988; Yeh et al., 1991). PDGF is also a chemoattractant and survival factor for oligodendrocyte progenitors (Armstrong et al., 1990; Barres et al., 1992b). The only PDGF receptor expressed in oligodendrocyte progenitors is PDGFR!, and the PDGFR" is absent in these cells (Hart et al., 1989). Complete loss of PDGFR! results in embryonic lethality, and loss of PDGFA, a PDGFR!-specific ligand, results in a severely reduced number of oligodendrocyte progenitors in the mouse brain (Betsholtz et al., 2001). SFKs have been shown to bind and be activated by PDGFR! in response to PDGF-AA (Thomas and Brugge, 1997b). Mice harboring mutations in the PDGFR! that selectively eliminate its capacity to activate SFKs exhibit neurological defects including shaking, seizure and decreased limb mobility (Klinghoffer et al., 2002). These mice also have a reduced number of oligodendrocytes progenitors in the spinal cord and hypomyelination in the cerebellum, corpus callosum and spinal cord. In vitro studies showed that inhibition of the activities of SFKs by PP2 inhibits PDGF-induced  !  27!  proliferation of oligodendrocyte progenitors, and SFKs promote PDGF-induced proliferation through upregulating the expression of Kv1.5, a potassium channel for delayed outward-rectifying K+ currents found predominantly in oligodendrocytes (Fig. 1.6A) (Attali et al., 1997; Soliven et al., 2003). Inhibition of the activities of SFKs by PP1 or knockdown of Fyn inhibit PDGF-mediated migration of oligodendrocyte progenitors, and Fyn promotes migration of oligodendrocyte progenitors through phosphorylating Cdk5 at Tyr15 in response to PDGF (Fig. 1.6B) (Miyamoto et al., 2008). These results indicate that SFKs are involved in PDGFR!-signaling that is required for normal oligodendrocyte development including proliferation and migration.  1.3.2.2 IGF-1 receptor (IGF-1R) In addition to PDGF and bFGF, there are many other molecules that promote oligodendrocyte proliferation and survival, such as neurotrophin-3 (NT-3), neuregulins and insulin-like growth factor 1 (IGF-1) (McTigue and Tripathi, 2008). IGF-1 and IGF1R are expressed in oligodendrocytes (Masters et al., 1991; McMorris et al., 1986; Shinar and McMorris, 1995). IGF-1-overexpressing mice showed a significant increase in brain weight and oligodendrocyte number (Carson et al., 1993; Zumkeller, 1997), while IGF-1 knockout mice exhibit a decreased number of oligodendrocytes (Beck et al., 1995; Ye et al., 2002). Moreover, disruption of the gene encoding IGF-1R reduced the proliferation of oligodendrocyte progenitors in mouse brain (Mason et al., 2003). In vitro studies demonstrated that IGF-1 promotes proliferation and survival of oligodendrocyte progenitors (Barres et al., 1992a; Barres et al., 1993). IGF-1 activates Fyn and Lyn in oligodendrocyte progenitors (Cui et al., 2005), and inhibition of the  !  28!  activities of SFKs by PP2 decreased IGF-1-stimulated proliferation of oligodendrocyte progenitors (Cui and Almazan, 2007), but had no significant effect on IGF-1-promoted survival (Cui et al., 2005). In addition, IGF-1 stimulation activates Akt and ERK1/2 to promote proliferation of oligodendrocyte progenitors, and IGF-1-mediated activation of Akt and ERK1/2 was blocked by PP2 (Fig. 1.6C) (Cui and Almazan, 2007).  1.3.2.3 Thromboxane A2 receptor (TXA2 receptor) TXA2 receptor is a member of the seven transmembrane domain, G-proteincoupled receptor superfamily (Hirata et al., 1991; Kim et al., 1992). It has been shown to be highly concentrated in myelinated regions of the CNS and to be expressed in oligodendrocytes (Blackman et al., 1998; Borg et al., 1994). A TXA2 mimetic, U46619, stimulates the proliferation of oligodendrocyte progenitors and promotes the survival of oligodendrocytes (Lin et al., 2005). Moreover, U46619 caused activation of ERK1/2 in oligodendrocyte progenitors and this activation was blocked by inhibition of SFKs by PP2 (Lin et al., 2005), suggesting that SFKs may be involved in TXA2-mediated proliferation and survival of oligodendrocyte progenitors (Fig. 1.6C).  1.3.2.4 Muscarinic acetylcholine receptor (mAChR) High-affinity mAChRs were detected in brain myelin and in oligodendrocytes (Cohen and Almazan, 1994; Larocca and Almazan, 1997; Larocca et al., 1987). The predominant mAChR subtype expressed in oligodendrocytes is M3, followed by M4, M1, M2 and M5 (Ragheb et al., 2001). A stable analog of acetylcholine, carbachol, promotes M3 receptor-mediated proliferation of oligodendrocyte progenitors (Cohen et al., 1996). It has been shown that carbachol also protects oligodendrocyte progenitors !  29!  from apoptosis induced by growth factor withdrawal, and SFKs are involved in this protective effect (Cui et al., 2006).  1.3.2.5 Integrins Integrins are the major cell surface receptors for extracellular matrix (ECM) ligands, and the intracellular signaling triggered by integrins regulates cell growth, survival, migration and differentiation (Cabodi et al.; Janik et al., 2010). In oligodendrocytes, the collagen/laminin receptor !1"1, the fibronectin receptor !5"1, the laminin receptor !6"1, and the vitronectin/fibronectin receptors !v"1, !v"3, !v"5, !6"1 have been detected, and the integrated actions of integrins with growth factors and their receptors has been demonstrated (Baron et al., 2005). PDGFR! associates with !v"3 integrin in oligodendrocyte progenitors and !6"1 integrin in newly-formed oligodendrocytes (Baron et al., 2003; Baron et al., 2002). Proliferation of oligodendrocyte progenitors in response to physiological PDGF concentrations (0.1-1 ng/ml) can be enhanced by engagement of !v"3 integrin (Baron et al., 2002), and this enhancement can be blocked by depletion of Lyn, but not Src or Fyn (Colognato et al., 2004). Newly formed oligodendrocytes have an increased dependency on survival factors such as PDGF and neuregulins (NRG) (Barres et al., 1993; Calver et al., 1998; Canoll et al., 1996a; Fernandez et al., 2000), and myelinating axon tracts express laminins to potentiate the effects of these soluble factors (Colognato and Yurchenco, 1999; Frost et al., 1999). Engagement of !6"1 integrin by laminin-2 amplifies PDGFmediated survival in newly formed oligodendrocytes, and Fyn is required for this amplification of the survival-promoting effects of PDGF (Colognato et al., 2004). Lyn  !  30!  depletion has a smaller effect on laminin-2-mediated amplification of PDGF survival, and Src depletion does not affect this amplification (Colognato et al., 2004). In addition to the integration of integrin and PDGF receptor actions, it has been shown that the Ig superfamily member F3/contactin is associated with integrins in oligodendrocytes (Laursen et al., 2009). The association of F3/contactin and integrins further amplifies PDGF survival signaling. Integrin stimulation induces dephosphorylation of the inhibitory Tyr531 residue of Fyn, whereas F3/contactin upregulates phosphorylation of both Tyr531 and the activating Tyr420, leading to enhanced activity of Fyn (Laursen et al., 2009). These results suggest that Fyn may be required for the synergistic effects of integrin and F3/contactin, which amplify PDGF survival signaling. Recently, Watzlawik et al. (Watzlawik et al., 2010) demonstrated that human remyelination promoting IgM mAb (rHIgM22) co-localized with "3 integrin and upregulates the expression and activity of Lyn in oligodendrocytes. The rHIgM22 antibody strongly inhibits apoptotic signaling in oligodendrocytes, and inhibition of Lyn activity by PP2 blocks rHIgM22-mediated inhibition of apoptosis (Watzlawik et al., 2010). These results suggest that Lyn may also play a role in "3 integrin-mediated survival of oligodendrocytes. 1.3.3  Fyn is involved in receptor and adhesion molecule-mediated changes in  oligodendrocyte differentiation and myelination Fyn is the only SFK that has been reported to be crucial for oligodendrocyte differentiation and myelination. Mice null for Src, Lyn and Yes do not exhibit defects in CNS myelination, but mice with mutant Fyn or lacking Fyn exhibit reduced myelin content and hypomyelination in the brain (Sperber et al., 2001; Umemori et al., 1999;  !  31!  Umemori et al., 1994). However, myelination in the spinal cord is normal in fyndeficient mice, indicating that CNS myelination exhibits regional differences. Reduced oligodendrocyte numbers are found in the corpus callosum, but not in the spinal cord of fyn-deficient mice, correlating well with the myelination phenotypes (Sperber et al., 2001). Reduced expression of myelin-associated genes, such as MBP and MOG, is also found in fyn-deficient mice (Goto et al., 2004). These findings indicate that Fyn is essential for CNS myelination, probably due to abnormal development of oligodendrocytes and expression of myelin-associated genes. In vitro studies further confirm that activation of Fyn is required for oligodendrocyte differentiation (Osterhout et al., 1999; Sperber and McMorris, 2001). Although Src, Lyn and Yes are expressed in oligodendrocytes, Fyn is the only SFK with significant kinase activity and with increased expression during differentiation (Câmara et al., 2009; Osterhout et al., 1999). Inhibition of Fyn activity by PP1 or by expression of a dominant-negative mutant Fyn reduces process extension and myelin membrane formation (Osterhout et al., 1999). However, it has been shown that in the absence of Fyn, some oligodendrocytes can still differentiate and express myelin specific genes, such as CNP and MBP, but the number of differentiated oligodendrocytes was decreased in fyndeficient mouse glial cell cultures (Sperber and McMorris, 2001). This controversy may be due to non-specific effects of the SFK inhibitor or compensatory effects in fyndeficient glial cultures. It is also possible that in addition to regulating morphological differentiation, Fyn has other roles in oligodendrocyte development. Indeed, as described above, Fyn is also involved in survival and migration of oligodendrocyte  !  32!  lineage cells, and there may be other functions that need to be identified, such as cell cycle regulation and self-renewal.  1.3.3.1 Insulin-like growth factor-1 receptor (IGF-1R) As described in 1.3.2.2, IGF-1 promotes survival of oligodendrocytes. In addition, IGF-1 also promotes oligodendrocyte differentiation and myelination (McMorris and McKinnon, 1996). IGF-1 treatment increased the morphological complexity of WT oligodendrocytes, but fyn-deficient oligodendrocytes are unable to respond to IGF-1 stimulation (Sperber and McMorris, 2001). These results suggest that Fyn is required for IGF-1-induced morphological changes of oligodendrocytes.  1.3.3.2 Transferrin receptor Transferrin receptors play a role in the uptake of transferrin-bound iron (Jandl and Katz, 1963). Apotransferrin has been shown to induce oligodendrocyte differentiation and myelin deposition (Escobar Cabrera et al., 1994; Escobar Cabrera et al., 1997; Marta et al., 2000; Paez et al., 2002) and to inhibit the response of oligodendrocyte progenitors to PDGF-induced proliferation (Paez et al., 2006). Mice overexpressing the human transferrin gene showed a significant increase in the expression of myelin basic protein (MBP), tubulin, Tau, and stable tubulin only peptide (STOP) (Marta et al., 2002). Similar results were found using apotransferrin-injected rats (Cabrera et al., 2000). Apotransferrin increases Fyn expression and autophosphorylation, and decreases Fyn colocalization with lipid rafts. Blocking Fyn-Tau-microtubule (MTs) interactions by using a dominant-negative form of Fyn inhibited apotransferrin-induced  !  33!  oligodendrocyte morphological differentiation. Apotransferrin also inhibits RhoA activity in an immortalized oligodendroglial cell line (N19) (Perez et al., 2009). These results suggest that Fyn plays an important role in the morphological differentiation of oligodendrocytes promoted by apotransferrin.  1.3.3.3 The common % chain of immunoglobulin Fc receptors (FcR%) FcR$ (receptors for IgG) belong to the immunoglobulin superfamily and are the most  important  Fc  receptors  for  inducing phagocytosis of opsonized microbes  (Hunkapiller and Hood, 1989; Springer, 1990). FcR% is expressed in oligodendrocytes. Cross-linking of FcRg induces Fyn signaling, myelin basic protein expression and morphological differentiation of oligodendrocytes. FcR%-deficient mice exhibit hypomyelination and reduced MBP expression (Nakahara et al., 2003). Cross-linking of FcR% also induces protein tyrosine phosphatase CD45 expression, and the interaction of Fyn with CD45 and Csk, suggesting that CD45 is involved in Fyn signaling during oligodendrocyte differentiation in response to immunoglobulin stimulation. Indeed, CD45-deficient oligodendrocyte progenitor cells failed to morphologically differentiate in response to immunoglobulin stimulation (Nakahara et al., 2005). Moreover, remyelination stimulated by the herbal medicine Ninjin’yoeito requires FcR%/Fyn signaling (Seiwa et al., 2007). In addition, FcR% and Fyn double knockout mice fail to activate Rac1 and p38MAPK as well as to remyelinate in response to Ninjin’yoeito. These results indicate that FcR%/Fyn signaling is crucial for oligodendrocyte differentiation and myelination during development as well as remyelination.  !  34!  1.3.3.4 Large myelin associated glycoprotein (L-MAG) MAG is a glycoprotein of the Ig-superfamily of adhesion molecules and is a minor constituent of myelin, comprising ~1% of all myelin proteins in the CNS. Two MAG isoforms, small (~67 kD, S-MAG) and large (~72 kD, L-MAG) MAG, resulting from alternative splicing of the MAG transcript, have been discovered. L-MAG is the abundant isoform in the developing CNS, while S-MAG is the predominant isoform in the adult CNS. MAG mediates interactions between axons and myelin-forming cells and initiates myelination (Schachner and Bartsch, 2000). MAG and Fyn are both expressed in oligodendrocytes, and can be coimmunoprecipitated. The association between L-MAG and Fyn requires the N-terminal region of Fyn including the SH2 and SH3 domains. Cross-linking of MAG with antibodies enhanced Fyn activity in COS cells cotransfected with Fyn and L-MAG. These results indicate that L-MAG interacts with and activates Fyn during the initial stage of myelination (Umemori et al., 1994). In addition, hypomyelination of the optic nerve of Fyn-deficient mice was more severe when compared with MAG-deficient mice, and the MAG/Fyn double-deficient mice exhibit the most severe phenotype of hypomyelination among these mutants (Biffiger et al., 2000). These results suggest that in addition to MAG, there may be other molecules that initiate myelination through activating Fyn.  1.3.3.5 GPI-anchored proteins (F3/Contactin and NCAM) F3/contactin and NCAM are both glycosyl-phosphatidylinositol (GPI)-linked neural adhesion molecules belonging to the Ig-superfamily. F3/contactin is expressed by neurons and oligodendrocytes (Einheber et al., 1997; Gennarini et al., 1989; Koch et  !  35!  al., 1997; Ranscht, 1988). F3/contactin has been shown to cooperate with other molecules in myelination, such as Tenascin-R, PTP!, Caspr, NogoA and Notch 1 (Hu et al., 2004). The spliced variants of neural cell adhesion molecule (NCAM) encode three isoforms with molecular weights of 120, 140 and 180 kD, of which NCAM-120 is the predominant isoform expressed in oligodendrocytes (Bhat and Silberberg, 1986; Bhat and Silberberg, 1988; Trotter et al., 1989). During early development, the majority of NCAM is posttranslationally modified to carry polysialic acid (PSA), which acts as a negative regulator of cell-cell interactions, and this modification is downregulated during late development (Bartsch et al., 1990; Edelman and Chuong, 1982; Rougon et al., 1982). Since both neurons and oligodendrocytes express NCAM, the modication of NCAM during early development may prevent premature myelination. Indeed, removal of PSA from NCAM expressed on axons is required for myelination (Charles et al., 2000; Fewou et al., 2007). Moreover, it has been shown that NCAM increases survival of premyelinating oligodendrocytes and promotes oligodendrocyte progenitor process outgrowth (Palser et al., 2009). Therefore, following removal of PSA, NCAM may be a signaling molecule promoting survival and process outgrowth of oligodendrocytes to initiate myelination. F3/contactin, NCAM and Fyn are expressed in oligodendrocyte progenitors, but they are not stably associated. In mature oligodendrocytes, Fyn is colocalized with F3/contactin and NCAM in rafts. Cross-linking F3/contactin with antibodies stimulates Fyn activity in rafts (Krämer et al., 1999). In addition, binding of the neuronal adhesion molecule L1 to oligodendroglial F3/contactin results in Fyn activation, which leads to enhanced phosphorylation of heterogeneous nuclear ribonucleoprotein A2 (hnRNP A2),  !  36!  facilitating translation of MBP mRNA (White et al., 2008). These results suggest that Fyn is responsible for F3/contactin and NCAM-mediated initiation of myelination.  1.3.3.6 Integrins As described in section 1.3.2.5, a variety of integrin isoforms are expressed in oligodendrocyte lineage cells. Laminin binding to !6"1 integrin not only promotes survival of oligodendrocytes, but also promotes differentiation into MBP-positive mature oligodendrocytes and morphological complexity (Colognato et al., 2004). Moreover, Fyn, but not Lyn and Src, is required for these processes. The SFK activity is upregulated by laminin in oligodendrocytes, not progenitors, and FAK is required for this upregulation (Colognato et al., 2004; Hoshina et al., 2007). Protein expression and phosphorylation of Fyn in oligodendrocytes also increase when these cells are cultured on fibronectin-coated dishes and "1 integrin is required for these processes (Liang et al., 2004). An in vivo study using laminin-deficient mice showed increased Fyn repression, and elevated levels of the SFK negative regulator Csk and its adaptor Cbp in these mice (Relucio et al., 2009). Relucio et al. (Relucio et al., 2009) also found delayed oligodendrocyte  maturation,  impaired  myelination  and  accumulation  of  oligodendrocyte progenitor cells in the brain of laminin-deficient mice. These results further confirm that integrin signaling, especially that triggered by laminin, is crucial for oligodendrocyte differentiation and myelination.  !  37!  1.3.3.7 Deleted in colorectal carcinoma (Dcc) The mammalian Dcc protein functions as a netrin receptor, based on a study demonstrating that Dcc-blocking antibodies inhibit netrin-stimulated outgrowth of neuron axons in vitro (Keino-Masu et al., 1996). The C. elegans unc-5 was identified to play a role in dorsalventral axon guidance (Hedgecock et al., 1990; Hedgecock et al., 1987). Three vertebrate homologues of this gene, such as UNC5, have been cloned and shown to bind netrin-1 in vitro (Ackerman et al., 1997; Leonardo et al., 1997). Oligodendrocyte progenitor cells express netrin-1 receptors Dcc and UNC5, but not netrin-1 itself (Jarjour et al., 2003). Netrin-1 acts as a chemorepellent and antagonist of PDGF-induced chemoattraction for oligodendrocyte progenitor cells to initiate oligodendrocyte precursor dispersal from the ventral to the dorsal embryonic spinal cord (Jarjour et al., 2003; Tsai and Miller, 2002; Tsai et al., 2003). In the adult CNS, netrin-1 is expressed by oligodendrocytes (Manitt et al., 2001). Netrin-1 promotes outgrowth by immature oligodendrocytes in vivo and in vitro, and induces Dccdependent process branching and myelin-like membrane sheet extension by mature oligodendrocytes in vitro (Rajasekharan et al., 2009). Rajasekharan et al. (Rajasekharan et al., 2009) also found that netrin-1 binding to Dcc recruits Fyn to a complex containg FAK and inhibits RhoA activity, and that Fyn is required for oligodendrocyte process extension and branching. These findings suggest a role for netrin 1 in promoting oligodendrocyte morphological changes.  !  38!  1.3.3.8 Leucine-rich repeats and Ig domain-containing, Nogo receptor-interacting protein-1 (LINGO-1) On neurons, LINGO-1 simultaneously interacts with the Nogo-66 receptor and p75NTR (neurotrophin receptor) or TROY (TNF receptor) to form a receptor complex that interacts with NogoA, MAG and oligodendrocyte myelin glycoprotein (OMgp), leading to the restriction of axonal elongation via activation of RhoA (Mi et al., 2004; Park et al., 2005; Shao et al., 2005). LINGO-1 is expressed in oligodendrocytes and negatively regulates oligodendrocyte differentiation, myelination and remyelination, probably due to its activation of RhoA and inhibition of Fyn activity (Lee et al., 2007; Mi et al., 2007; Mi et al., 2005; Mi et al., 2009). In addition, LINGO-1 knockout mice show early-onset CNS myelination, but no obvious changes in peripheral nervous system myelination (Mi et al., 2005). Treatment with anti-LINGO-1 antibody augments remyelination in lysophosphatidylcholine and cuprizone-induced demyelination (Mi et al., 2009). Ablation of LINGO-1 or treatment with anti-LINGO-1 antibody results in enhanced remyelination and mitigated diseases symptoms in rodent experimental autoimmune encephalomyelitis (EAE), a mouse model of multiple sclerosis. Bourikas et al. (Bourikas et al., 2010) demonstrated that LINGO-1 inhibits process extension and MBP expression in the oligodendroglial cell lines MO3.13 and CG4, respectively. They also found that LINGO-1-mediated inhibition of oligodendrocyte differentiation requires p75NTR signaling. These results suggest that LINGO-1 is a negative regulator of oligodendrocyte differentiation and myelination, and that Fyn is involved in these events.  !  39!  1.3.3.9 Opioid receptor The µ- and (-opioid receptors are expressed in oligodendrocytes in a developmentally regulated manner (Knapp et al., 1998). Oligodendrocytes from cultured adult rat hippocampal progenitors decreased in number upon treatments with µ- and #-opioid receptor antagonists (Persson et al., 2003). Buprenorphine, a µ-opioid receptor partial agonist and (-opioid receptor antagonist, affects myelination in the developing brain (Sanchez et al., 2008). Buprenorphine alters the levels and developmental expression of MBP isoforms and the number of myelinated axons, and reduces relative thickness of the myelin sheath. Buprenorphine also affects the expression of MAG and its glycosylation as well as altering MAG-Fyn complex formation. These results suggest that Fyn may play a role in opioid-mediated oligodendrocyte development and myelination.  1.3.4  Fyn signaling in oligodendrocyte morphological differentiation and  myelination Several downstream targets of Fyn that are involved in oligodendrocyte differentiation and myelination have been identified, such as Tau, FAK, p190RhoGAP, p250RhoGAP, Rac1 and Cdc42. These molecules regulate cytoskeletal rearrangement to promote process outgrowth and/or wrapping of axons, which are critical processes in oligodendrocyte morphological differentiation and CNS myelination.  !  40!  1.3.4.1 Tau Tau is a microtubule-associated protein that induces microtubule assembly and bundle formation (Brandt, 1996). It also interacts with components of the plasma membrane in neurons (Brandt et al., 1995; Maas et al., 2000). Tau has been shown to interact with the Fyn SH3 domain in a neuroblastoma cell line (Lee et al., 1998). In addition to neurons, Tau is also expressed in oligodendrocytes (LoPresti et al., 2001; LoPresti et al., 1995; Muller et al., 1997; Richter-Landsberg, 2000; Richter-Landsberg and Gorath, 1999; Song et al., 2001). Klein et al. (Klein et al., 2002) demonstrated that Fyn is associated with Tau and tubulin in primary oligodendrocytes. Tau binds to the SH2 domain of Fyn, while !-tubulin interacts with the SH2 and SH3 domains of Fyn. Inhibition of the interaction between Fyn and Tau by overexpressing a mutant Tau that competes with endogenous Fyn-Tau binding downregulates process numbers and process length in an oligodendroglial cell line, Oli-neu. These results indicate that FynTau signaling is essential in oligodendrocyte morphological differentiation.  1.3.4.2 Focal adhesion kinase (FAK) FAK is a ubiquitously expressed cytoplasmic tyrosine kinase that is a downstream effector of integrins and various extracellular signals and is well recognized as a central regulator of cell adhesion and motility (Mitra et al., 2005). The loss of FAK expression disrupts microtubule polarization and results in defect in focal contact turnover, which has been linked to the FAK-mediated regulation of Rho family GTPases in cells (Palazzo et al., 2004; Ren et al., 2000). Rho family GTPases control the formation and disassembly of actin cytoskeletal structures as described in section  !  41!  1.3.4.3. FAK is autophosphorylated at Tyr397 upon integrin stimulation. This phosphotyrosine residue is a binding site for the SH2 domain of SFKs (Cary et al., 1996). FAK is then phosphorylated at several tyrosine residues by the recruited SFKs, leading to full activation (Calalb et al., 1995). FAK is expressed in oligodendrocyte lineage cells and is present in myelin (Bacon et al., 2007; Kilpatrick et al., 2000). Since a ubiquitous FAK knockout is early embryonic lethal (Furuta et al., 1995; Ilic et al., 1995a; Ilic et al., 1995b), Forrest et al. (Forrest et al., 2009) generated oligodendrocytespecific and inducible FAK knockout mice to investigate its role in myelination. They observed a significant reduction in the number of myelinated fibers on postnatal day 14 when inducing FAK ablation prior to and during active myelination. Hoshina et al. (Hoshina et al., 2007) found that tyrosine phosphorylation of FAK increased during differentiation of an oligodendroglial cell line, CG4, in a Fyn-dependent manner. FAK phosphorylation was induced by laminin stimulation and laminin-stimulated process outgrowth was impaired in FAK-knockdown CG4 cells. Moreover, FAK is required for the activation of SFKs, Rac1 and Cdc42 in laminin-stimulated CG4 cells. These results indicate that Fyn-FAK signaling activates the downstream effectors Rac1 and Cdc42 and is required for laminin-induced oligodendrocyte process outgrowth.  1.3.4.3 Rho Family GTPases (RhoA, Rac1 and Cdc42) Rho family GTPases play important roles in regulating cytoskeletal rearrangement (Visvikis et al., 2010). Rho GTPases can be either in an active form (bound to GTP) or in an inactive form (bound to GDP) (Jaffe and Hall, 2005). The activities of Rho GTPases can be regulated by (1) guanine nucleotide exchange factors  !  42!  (GEFs) that catalyze the exchange of GDP for GTP (Schmidt and Hall, 2002); (2) GTPase-activating proteins (GAPs) that stimulate the intrinsic GTPase activity (Bernards, 2003); and (3) guanine nucleotide dissociation inhibitors (GDIs) that block spontaneous activation (Olofsson, 1999). In fibroblasts, Rho activation promotes stress fiber formation, while Rac and Cdc42 activation induce lamellipodia and filopodia, respectively (Kozma et al., 1995; Nobes and Hall, 1995; Ridley et al., 1992). Rho and Cdc42-Rac1 exert opposing effects on oligodendrocyte morphological differentiation. RhoA activity decreases and Cdc42 and Rac1 activities increase in oligodendrocyte progenitor cells during differentiation (Liang et al., 2004). By using dominant-negative and constitutively active Rho, Rac1 and Cdc42, it was demonstrated that Rho activition inhibits oligodendrocyte process outgrowth, while the activation of Rac1 and Cdc42 induces this process (Wolf et al., 2001). However, Thurnherr et al. (Thurnherr et al., 2006b) reported that Cdc42 is not required for oligodendrocyte morphological differentiation by using Cdc42-null oligodendrocyte progenitor cells. An in vivo study showed that ablation of both Rac1 and Cdc42 resulted in an abnormal myelination phenotype (Thurnherr et al., 2006b). Fyn is involved in the activation of Rac1 and Cdc42 in oligodendrocytes, since inhibition of integrin engagement and Fyn activation blocked the activation of Rac1 and Cdc42 (Liang et al., 2004). These results suggest that signaling from integrin to Fyn to RhoGTPases is crucial in oligodendrocyte differentiation and myelination.  !  43!  1.3.4.4 Rho GTPase activating proteins (p190RhoGAP and p250RhoGAP) RhoGAPs, potent regulators of Rho GTPases, accelerate the intrinsic GTPase activity of Rho family GTPases to inactivate the GTP-bound (on) state of the Rho GTPases (Settleman, 2003). The p190RhoGAP was identified as a Fyn interacting protein in a yeast two hybrid screen using Fyn as bait (Wolf et al., 2001), and is a major substrate of SFKs in the adult brain (Brouns et al., 2001). Phosphorylated p190RhoGAP binds to the Fyn SH2 domain. In addition, phosphorylation of p190RhoGAP by Src and Fyn increases the GAP activity (Arthur et al., 2000; Fincham et al., 1999; Wolf et al., 2001). Phosphorylation of p190RhoGAP increases during oligodendrocyte differentiation and the phosphorylation of p190RhoGAP requires Fyn activation. By overexpressing WT and mutant p190RhoGAP in primary oligdendrocyte progenitor cells, Liang et al. (Liang et al., 2004) showed that p190RhoGAP promotes oligodendrocyte morphological differentiation and inhibits RhoA activation. Another RhoGAP, p250RhoGAP (also known as p200RhoGAP, Grit and RICS), is a brainenriched RhoGAP (Moon and Zheng, 2003; Nakamura et al., 2002; Okabe et al., 2003). The p250RhoGAP is recruited to activated receptor tyrosine kinases and is tyrosine phosphorylated upon ligand stimulation (Nakamura et al., 2002). It has been shown that p250RhoGAP is tyrosine phosphorylated by Src and Fyn (Moon and Zheng, 2003; Taniguchi et al., 2003). Taniguchi et al. (Taniguchi et al., 2003) also found that tyrosine phosphorylation of p250RhoGAP increased in CG4 cells upon differentiation. These results indicate that RhoGAPs are important regulators of oligodendrocyte morphological differentiation.  !  44!  1.3.5 Fyn controls MBP expression via multiple mechanisms It has been shown that Fyn controls MBP expression by transcriptional, posttranscriptional and translational regulation (White et al., 2008). Transcriptional activation of the MBP gene by Fyn was demonstrated by CAT reporter assay, and the Fyn response sequence has been identified (bp -675 to -647 of the MBP promoter). Binding of transcription factors (probably CCAAT/enhancer binding protein (C/EBP) family of transcription factors) to the Fyn response element is developmentally regulated, indicating that Fyn stimulates transcription factor binding to the promoter of the MBP gene during myelination (Umemori et al., 1999). Since a preferential reduction of exon2-containing MBP mRNA isoforms derived from alternative splicing was found in Fyn-deficient mouse forebrain, Fyn may be required for posttranscriptional regulation of MBP. A RNA-binding protein maintaining the stability of MBP mRNA, QKI, can be phosphorylated by Fyn (Lu et al., 2005a). SFKs act as negative regulators of QKI functions, since the opposite expression patterns of MBP isoforms are detected in Fyn-deficient compared to QKIdeficient mouse forebrains (Lu et al., 2005a). Also, phosphorylation of QKI at a Cterminal tyrosine by SFKs inhibits the ability of QKI to bind MBP mRNA (Zhang et al., 2003). These results indicate that Fyn phosphorylates QKI to regulate the homeostasis of MBP mRNA. MBP mRNA is transported in RNA granules to the processes of oligodendrocytes, and translated locally to synthesize of large amounts of MBP at the axon-glia contacts. Therefore, the translation of MBP mRNA has to be repressed until the mRNA reaches its destination. MBP mRNA contains an A2 response element (A2RE), an 11-  !  45!  nucleotide sequence that hnRNP A2 binds to (Ainger et al., 1997; Ainger et al., 1993). A2RE-containing mRNA (such as MBP mRNA) and hnRNP A2 assemble in RNA granules, which recruit hnRNP E1 to inhibit translation of A2RE-containing mRNA (Kosturko et al., 2006). By using immunoaffinity chromatography to purify tyrosine phosphorylated proteins following cross-linking of oligodendroglial F3/contactin to activate Fyn, hnRNP A2 was identified to be a candidate substrate of Fyn. The phosphorylation of hnRNP A2 by Fyn was confirmed by in vitro kinase assay and coimmunoprecipitation. Fyn phosphorylates hnRNP A2 in response to neuronal L1, leading to release of hnRNP E1 and hnRNP A2 from the RNA granules (termination of translation repression), and enhancing translation of MBP mRNA (White et al., 2008). These results suggest that Fyn is a crucial regulator of the local translation of MBP mRNA at axon-glia contacts.  1.4  Rationale and Hypothesis Central nervous system (CNS) myelination is crucial in maintaining normal  functions of the brain and spinal cord. Defects in CNS myelination result in several demyelinating diseases that fall into two main groups: acquired diseases (i.e. multiple sclerosis) and hereditary neurodegenerative disorders (i.e. the leukodystrophies) (Keyoung and Goldman, 2007; Nguyen et al., 2006). Events such as virus infection, stroke, traumatic brain injury and spinal cord injury can also result in demyelination (Medana and Esiri, 2003; Mueller et al., 2005). Without myelin, nerve impulses are slowed or stopped, leading to a constellation of neurological symptoms. Myelin, a multilamellar insulating membrane that ensheathes the axons, is made by membranes of  !  46!  oligodendrocytes in the brain and spinal cord, and of Schwann cells in peripheral nerves. Oligodendrocytes arise during development from oligodendrocyte precursor cells (OPCs) and express myelin-specific genes. The development of therapies to promote remyelination and treat demyelinating diseases such as multiple sclerosis requires understanding of the molecular mechanisms that promote oligodendrocyte differentiation and myelination. Myelin production by oligodendrocytes in the CNS is regulated by signaling between adhesion molecules on both axon and oligodendrocyte cell surfaces (Baron et al., 2005; Hu et al., 2006). The role of protein tyrosine kinases in oligodendrocyte development and myelination has been extensively studied (Roy et al., 2007; Seiwa et al., 2007). However, the importance of protein tyrosine phosphatases (PTPs) in the development and myelination of oligodendrocytes remains poorly defined. It is therefore necessary to define the role of PTPs in oligodendrocyte development and CNS myelination. A focus of research in our lab is the widely expressed receptor PTP, PTP!. Several findings indicate that PTP! might play an important role in oligodendrocyte development and CNS myelination. First, PTP! is highly expressed in brain and is the most abundant PTP in oligodendrocytes (Ranjan and Hudson, 1996). Second, our lab demonstrated that PTP! forms a complex with the cell surface adhesion molecule F3/contactin (Zeng et al., 1999), and that F3/contactin is a Notch ligand that upregulates oligodendrocyte maturation (Hu et al., 2003). Third, PTP! associates with and activates Fyn in cultured cells, in brain and in integrin signaling (Bhandari et al., 1998; Ponniah et al., 1999; Zeng et al., 1999); and Fyn and integrin signaling have been implicated in oligodendrocyte differentiation and CNS myelination (Sperber et al., 2001). Therefore,  !  47!  I  hypothesize  that  PTP!  promotes  oligodendrocyte  differentiation  and  myelination by dephosphorylating and activating Fyn. This will be investigated through the following specific objectives: Objective 1: Establish in vitro oligodendroglial cell model systems with and without PTP!, that can be manipulated to differentiate and are amenable to the characterization of associated morphological and molecular changes. Objective 2: Determine if PTP!-Fyn signaling is required for oligodendrocyte differentiation and myelination and if so, investigate the molecular mechanisms of this signaling. Objective 3: Determine if PTP!-Fyn signaling regulates oligodendrocyte selfrenewal, and if so, investigate the molecular mechanisms of this signaling. The results of these investigations will lead to a better understanding of oligodendrocyte development and CNS myelination, and may shed light on the pathogenesis of oligodendrocyte-related diseases. Furthermore, the demonstration of roles of PTP! and the elucidation of PTP!-regulated events in oligodendrocyte development that determine myelination would indicate that drugs targeting PTP! or PTP! signaling events could be an effective treatment for enhancing remyelination in demyelinating diseases such as multiple sclerosis and leukodystrophie#$!  !  48!  !  !  49!  !  50!  !  51!  !  52!  !  53!  !  54!  Chapter 2. MATERIALS AND METHODS  2.1  Materials  2.1.1 Mice The 129SvEv PTP!&/& mice (Ponniah et al., 1999) were backcrossed with C57BL/6 mice for 10 generations. PTP!&/& and wild type (WT) C57BL/6 mice were housed under specific pathogen-free conditions. Animal care and use followed the guidelines of the University of British Columbia and the Canadian Council on Animal Care, and were reviewed and approved by the University of British Columbia.  2.1.2 Reagents Reagents were obtained from Sigma-Aldrich Canada (Oakville, ON, Canada) unless otherwise indicated. DNase I was purchased from Invitrogen Canada (Burlington, ON, Canada).  2.1.3 Antibodies Anti-PTP! antiserum has been described previously (Chen et al., 2006).! Antibodies to A2B5, O4, NG2, MBP, Ras, PDGFR! and phosphotyrosine (4G10) were purchased from Millipore (Billerica, MA, USA). Antibodies to phosphoTyr527-Src and phosphoTyr576-FAK were purchased from Biosource (Camarillo, CA, USA). Antibodies to Fyn, FAK, Rac1, Cdc42, p190 RhoGAP and p27Kip1 were purchased from BD Transduction Laboratories (San Jose, CA, USA). Antibodies to cleaved caspase-3, phosphoSer473-Akt, Akt, phosphoThr202/Tyr204-ERK1/2, and ERK were  !  55!  purchased from Cell signaling (Dever, MA, USA). Antibody to Lyn and p120RasGAP and antibody for the immunoprecipitation of Fyn were purchased from Santa Cruz Biotechnology (Santa Cruz, CA, USA). Antibodies to actin and CNPase were purchased from Sigma-Aldrich Canada (Oakville, ON, Canada). Antibody to Src was purchased from Oncogene Science (Cambridge, MA, USA). Antibody to Rho was purchased from Stressgen Biotechnologies (Victoria, BC, Canada). Antibody to Ki-67 was purchased from Dako Canada (Burlington, ON, Canada). Secondary antibodies conjugated with Alexa Fluor 488 or 594 (Molecular Probes) were purchased from Invitrogen Canada (Burlington, Ontario, Canada).  2.1.4 Growth factors Human recombinant PDGF-AA, bFGF and EGF were purchased from PeproTech (Rocky Hill, NJ, USA). Human recombinant IGF-1 was purchased from BioVision (Mountain View, CA, USA).  2.2  Cell line and primary cell cultures  2.2.1 OLN-93 cells OLN-93 cells were kindly provided by Dr. Zhi-Cheng Xiao (Institute of Molecular and Cell Biology, Singapore) and were maintained in DMEM (high glucose, Cat. # SH30243.01, Thermo Scientific) with 10% FBS. To promote differentiation, cells were seeded on poly-D-lysine (PDL, 10 µg/ml)-coated dishes at a density of 1x104/cm2 overnight. Cells were then gently washed with PBS and subsequently cultured in DMEM with 0.5% FBS for various times.  !  56!  2.2.2 CG4 cells CG4 cells were kindly provided by Dr. Y. Feng (Emory University School of Medicine, USA) and were maintained in CG4 proliferation medium (DMEM [high glucose, Cat. # SH30243.01, Thermo Scientific], 1% FBS, 5 µg/ml insulin, 50 µg/ml transferrin, 30 nM sodium selenite, 100 µM putrescine, 20 nM progesterone, 10 ng/ml biotin, 10 ng/ml PDGF, 10 ng/ml bFGF). To promote differentiation, cells were seeded on PDL (10 µg/ml)-coated dishes at a density of 1.5x104/cm2. After attachment (~3h), cells were gently washed with PBS and subsequently cultured in CG4 differentiation medium (DMEM, 0.5% FBS, 5 µg/ml insulin, 50 µg/ml transferrin, 30 nM sodium selenite, 50 nM tri-iodothyronine [T3]) for various times.  2.2.3 Primary mouse neural progenitor/stem cells and OPCs 2.2.3.1 Isolation and culture of primary OPCs from postnatal mice This procedure was established according to the protocol described previously (McCarthy and de Vellis, 1980). After removal of meninges and cerebellum in ice-cold Hank’s buffered saline solution (HBSS, Cat. #24020, Gibco), cerebral cortex tissue from postnatal day 1 (P1) mice was placed in a sterile dish containing HBSS, cut into small pieces, transferred into a 15 ml tube and centrifuged at 200 x g for 1 min to remove HBSS. The pellet was resuspended in 3ml of 0.25% trypsin-EDTA (Gibco) for 10 min at 37°C followed by the addion of 7 ml DMEM with 10% FBS to stop digestion. The digested tissue was collected and resuspended in 3 ml of DMEM with 1 µl of DNase I. The tissues were triturated until the cell suspension had no or very few  !  57!  small clumps, then filtered through a 70 µM cell strainer, and plated at 5-8x106 cells per 75 cm2 flask in Basal Medium Eagle’s with Earle’s balanced salts (Cat. # 21010, Gibco) containing 15% FBS, 0.1% glutamine, 0.6% glucose (BME medium). Cells were cultured overnight and then the medium was replaced. After culturing for 9 days with the medium replaced every other day, the medium was replaced again. At day 10, the cells were rinsed 3 times with BME medium to remove floating cells. Fresh medium was added and the flasks were allowed to equilibrate in the CO2-incubator for 2 hours followed by shaking at 37°C at 250 rpm for 15-18 hours. The medium was then collected, filtered through 70µm and then 40 µm cell strainers (BD Falcon, Mississauga, ON, Canada) followed by centrifugation at 1000rpm for 5 min. Cells were seeded on PDL-coated dishes/coverslips at 3x104 cells/cm2 for the indicated times.  2.2.3.2 Isolation and culture of primary neural progenitor/stem cells and OPCs from mouse embryos Primary mouse neural progenitor/stem cells and OPCs were generated from neurospheres as described previously (Chen et al., 2007; Pedraza et al., 2008b) with some modifications. A schematic diagram of the procedure is shown in Fig. 2.1. In brief, after removal of meninges and cerebellum in ice-cold Hank’s buffered saline solution (HBSS), cerebral cortex tissue from E14.5-E17.5 mouse embryos was placed in a sterile dish containing HBSS, cut into small pieces and transferred into ice-cold neural culture medium (NCM) supplemented with 20 ng/ml bFGF and 20 ng/ml EGF (0.5 ml per brain). The tissues were then mechanically triturated with a 1-ml Gilson pipette until the cell suspension had no or very few small clumps, filtered through a 70 µM cell strainer,  !  58!  and plated at 5x104 cells/ml in a six-well plate (4 ml per well of NCM supplemented with 20 ng/ml bFGF and 20 ng/ml EGF). NCM contains DMEM/F12 (Cat. # 11330, Gibco), 25 µg/ml insulin, 100 µg/ml apo-transferrin, 20 nM progesterone, 60 µM putrescine and 30 nM sodium selenite. After 3-4 days, floating neurospheres were passaged at a 1:3 ratio in the same medium every 3-4 days. Passage 2-6 neurospheres were used for experiments, cryopreserved in full media containing 10% DMSO, or mechanically dissociated into a single cell suspension and resuspended in NCM supplemented with 20 ng/ml PDGF-AA and 20 ng/ml bFGF (oligosphere medium) to induce oligosphere formation. After 72 h, cell aggregates were passaged at a 1:2 ratio every 4-6 days. Oligospheres (passage 2-6) were dissociated using the NeuroCult Chemical Dissociation Kit (mouse) (StemCell Technologies, Alberta, Canada) and plated on poly-D,L-ornithine (PDLO, 50 µg/ml)-coated chamber slides or dishes at a density of 3x104/cm2 in oligosphere medium for 2 days. To induce differentiation, the medium was changed to NCM supplemented with 5 µg/ml N-acetyl-L-cysteine and 50 nM T3 for 2-4 days. For spinal cord mouse neural progenitor/stem cells and OPCs, mouse E14.517.5 spinal cord was dissected in ice-cold Hank’s buffered saline solution (HBSS). The tissues were then placed in a sterile dish containing HBSS, cut into small pieces and transferred into ice-cold NCM supplemented with 20 ng/ml bFGF and 20 ng/ml EGF (0.5 ml per brain) followed by the procedures described above. Embryonic spinal cord cells initially attached to the plate, and the floating neurospheres formed within 5– 7 days in culture. The primary spheres were plated at 5x104 cells/ml in a six-well plate in the same medium for 3-4 days. Passage 3-6 neurospheres were used for experiments  !  59!  or mechanically dissociated into a single cell suspension and induced to form oligospheres. Culture and differentiation of the oligospheres were performed as described above.  2.3  Immunofluoresence labeling of cells and tissues Cells grown on PDL or PDLO-coated coverslips or chamber slides (Nalgene  Nunc International, Rochester, NY, USA) were fixed with 4% paraformaldehyde for 15 min at room temperature and then washed three times with PBS. For other experiments, animals were anesthetized and perfused intracardially with 4% paraformaldehyde in 0.1 M phosphate buffer, pH 7.2. The brain was removed, post-fixed in the same solution overnight at 4°C, and then cryopreserved in 30% sucrose in 0.1M phosphate buffer and embedded in OCT media (Sakura Finetek, Torrance CA, USA). Cells and tissue sections (10 µm) were incubated with blocking buffer (0.1M phosphate buffer, 0.1% gelatin, 1% BSA, 0.02% sodium azide, 10% goat serum) for 30 min (0.5% Triton-X100 was added to the blocking buffer if permeablization was required), followed by incubation with primary antibodies overnight at 4°C. After washing three times with PBS, cells and tissues were incubated with secondary antibodies for 2 h at room temperature. The slides were washed three times with PBS followed by mounting in Prolong Gold Antifade Reagent (Invitrogen Canada) with DAPI and viewed using an Axioplan2 fluorescence microscope (Carl Zeiss MicroImaging, Thornwood, NY).  !  60!  2.4  Immunoblotting Cells were harvested by washing twice with ice-cold PBS on ice. For preparation  of lysates, cells were lysed on ice by adding RIPA lysis buffer (50mM Tris-HCl pH 7.4, 150mM NaCl, 0.5% sodium deoxycholate, 1% NP-40, 0.1% SDS, 1mM EDTA, 2 mM sodium orthovanadate, 50 mM sodium fluoride, 10 µg/ml aprotinin, 10 µg/ml leupeptin, 1 mM PMSF) or NP-40 lysis buffer (RIPA lysis buffer without sodium deoxycholate and SDS) directly onto the cells. Cell lysates were then transferred to microtubes and incubated for 30 min on ice, centrifuged at 12000 rpm for 10 min at 4°C, and the supernatants collected to obtain protein extracts. Protein concentration was determined with the BCA Protein Assay Kit (Pierce, Rockford, IL, USA). Protein extracts were resolved by SDS-PAGE and transferred to a PVDF membrane, which was then blocked with 3% bovine serum albumin in PBS with 0.1% Tween 20 for 1 h at room temperature. The membranes were probed overnight at 4°C with the appropriate antibodies, washed, and probed again with species-specific secondary antibodies coupled to horseradish peroxidase. Chemiluminescent reagents were then added for signal detection.  2.5  Immunoprecipitation Cell  lysates  (50-100  µg  protein)  prepared  with  RIPA  buffer  were  immunoprecipitated with the indicated antibody at 4ºC overnight, followed by incubation with 40 µl protein A/G-agarose (Santa Cruz Biotechnology, Santa Cruz, CA, USA) at 4ºC for 3h. After washing three times with lysis buffer, the immunoprecipitates were analyzed by immunoblotting.  !  61!  2.6  siRNA transfection The following siRNAs (Dharmacon, Chicago, IL, USA) were used: Control  (siCONTROL Non-Targeting siRNA Pool #2 D-001206-14-20), PTP! (ONTARGETplus SMARTpool L-080089-01-0050, Rat PTPRA, NM_012763) and Fyn (ON-TARGETplus SMARTpool L-089444-00-0010, Rat Fyn, NM_012755). CG4 cells were seeded in CG4 proliferation medium (3x104/cm2). After overnight attachment, cells were incubated with 20 nM siRNA and Lipofectamine RNAiMax (Invitrogen Canada) in OPTI-MEM I (Cat. # 31985, Gibco) for 24 hours. Cells were trypsinized and seeded on PDL-coated chamber slides or dishes in CG4 proliferation medium. After 3 hours, cells were gently washed and incubated in CG4 differentiation medium for various times.  2.7  Ras and Rho family GTPase activities Ras activity was measured by GST-Raf1 RBD (Ras-binding domain) pulldown  assays performed using the Ras Activation Assay Kit (Upstate, Temecula, CA, USA). Rho activity was measured by GST-RBD (Rhotekin-binding domain) pulldown assays performed using the Rho Activation Assay Kit (Upstate, Temecula, CA, USA). Rac1 and Cdc42 activities were measured by GST-PBD (PAK-binding domain) pull-down assays (Manser et al., 1994). Cells were lysed on ice by adding RIPA lysis buffer directly onto the cells. Cell lysates (15-100 µg) were incubated with 10 µg GST-PBD bound to glutathione-Sepharose beads. Samples were washed with lysis buffer and then immunoblotted with anti-Rac1 and Cdc42 antibodies. Lysates were directly  !  62!  immunoblotted to determine the total amount of Rho, Rac1 or Cdc42 proteins. Levels of active Rho, Rac1 and Cdc42 were normalized to those of total Rho, Rac1 and Cdc42.  2.8  Cell cycle analysis and BrdU incorporation assay Cell cycle analysis was conducted using flow cytometry. Cells were collected and  washed in PBS. Cells were then fixed with ice-cold 70% ethanol at 4°C for periods of 30 mins up to a week. Cell pellets were collected by centrifugation at approximately 2,000 rpm for 5 mins, and then washed twice in PBS. Cells were stained by adding 20 µl of 7-AAD (BD Pharmingen, Mississauga, ON, Canada) per 106 cells and then analyzed by flow cytometry. About 20000 events per sample were collected. BrdU incorporation assay was performed using the In Situ Cell Proliferation Kit, FLUOS (Roche, Mannheim, Germany).  2.9  Data analysis Densitometric quantification of immunoblots and cell differentiation data were  statistically analyzed using ANOVA (single factor).  !  63!  !  64!  Chapter  3.  ESTABLISHMENT  AND  CHARACTERIZATION  OF  OLIGODENDROGLIAL CELL MODEL SYSTEMS  3.1  Oligodendroglia cell lines To study the role of PTP! in oligodendrocyte development and myelination, I  choose two types of cultured oligodendrocyte cell lines, OLN93 and CG4, as models. PTP!-deficient OLN93 and CG4 cells were generated by siRNA-mediated silencing. There are some advantages of using these cell lines as models. For example, they are easy to culture, expand and manipulate, and they generate highly pure populations of oligodendrocytes. Therefore, they are suitable for biochemical and molecular studies. These two cell lines were both derived from primary glial cultures of neonatal rat brain. OLN-93 cells are A2B5-negative cells and CG4 cells are A2B5-positive OPCs (Louis et al., 1992; Richter-Landsberg and Heinrich, 1996). These cell lines undergo a maturation/differentiation process upon serum or growth factor withdrawal that resembles myelination, during which the cells lose the bipolar phenotype typical of immature oligodendrocytes and extend multiple cell processes. In addition, the expression of surface antigens and other proteins characteristic of mature oligodendrocytes (O1, CNPase, MBP, MAG) are upregulated (de Castro and Bribián, 2005). The expression of these markers can be detected by immunoblotting or immunofluoresence labeling.  !  65!  3.1.1 Characterization of OLN-93 cell differentiation OLN-93 cells are A2B5-negative, spontaneously transformed cells, and they represent an oligodendroglia cell line at a later stage of differentiation. Their morphological features and antigenic properties resemble 5-10-day-old (postnatal day) cultured rat brain oligodendrocytes (Richter-Landsberg and Heinrich, 1996). When OLN-93 cells were plated on PDL-coated dishes and incubated in DMEM with 0.5% FBS for up to 6 days, they underwent differentiation with increased expression of the oligodendrocyte markers CNPase and MAG and extended multiple branched processes (Fig. 3.1A). To confirm that specific the oligodendrocyte markers were upregulated in these cells after differentiation was induced, cell lysates were subjected to immunoblotting with anti-CNPase antibody. CNPase protein levels were upregulated in OLN-93 cells during differentiation (Fig. 3.1B).  3.1.2 Characterization of CG4 cell differentiation CG4 cells are A2B5-positive OPCs, which display the karyotype of normal rat cells. They can be induced to differentiate into oligodendrocytes and type-2 astrocytes by mitogen withdrawal or high serum concentrations. They do not proliferate in the absence of mitogens secreted by the neuronal B104 cell line, suggesting that they are not transformed (Louis et al., 1992). When CG4 cells were plated on PDL-coated dishes and incubated in CG4 differentiation medium for up to 6 days, they underwent differentiation  from  A2B5-positive  bipolar  OPCs  into  CNPase-positive  oligodendrocytes that extended multiple branched processes (Fig. 3.2A). To confirm that specific oligodendrocyte markers were upregulated in these cells after  !  66!  differentiation was induced, cell lysates were subjected to immunoblotting with antiCNPase antibody. CNPase protein levels were upregulated in CG4 cells during differentiation (Fig. 3.2B).  3.1.3 Ablation of PTP! by RNA interference (RNAi) in OLN-93 and CG4 cell lines To optimize the conditions for siRNA transfection, two different transfection reagents (DharmaFECT3 and Lipofectamine 2000) were tested. When DharmaFECT3 was used as the transfection reagent, ~70% PTP! knockdown efficiency was achieved by transfecting 50 nM PTP! siRNA in OLN-93 cells, while only ~30% PTP! knockdown was observed in CG4 cells. When Lipofectamine 2000 was used as the transfection reagent, ~70% PTP! knockdown was observed in OLN-93 cells, and ~90% PTP! knockdown was achieved in CG4 cells (data not shown). To improve the poor efficiency of PTP! knockdown in OLN-93 cells, another transfection reagent, Lipofectamine RNAiMAX was tested. This effected a ~99% knockdown of PTP! expression in OLN-93 cells. To avoid off-target effects and to determine if efficient knockdown could be achieved during differentiation, OLN-93 and CG4 cells were transfected using Lipofectamine RNAiMAX and 10 nM siRNA and the cells were then induced to differentiate. This reproducibly resulted in a greater than 90% knockdown of PTP! in both OLN-93 and CG4 cells after differentiation for 2 days (Fig. 3.3A and B). Therefore, these conditions were used for further experiments. To generate stable PTP!-deficient cell lines, SMARTvector shRNA lentiviral particles were purchased from Dharmacon. The SMARTvector includes a TurboGFP reporter gene to facilitate assessment of transduction efficiencies and to be used for  !  67!  fluorescence activated cell sorting, and a puromycin resistance gene to allow selection and isolation of clonal populations when generating stable cell lines. After transducing OLN-93 and CG4 cells with three PTP! shRNA with different target sequences or empty vector, cells were selected by 0.8 µg/ml puromycin. After several passages, PTP! expression was determined by immunoblot. No significant PTP! knockdown was observed in any OLN-93 and CG4 clones transduced with PTP! shRNA-1, 2 or 3 (data not shown). To exclude the possibility that puromycin selection was not efficient or that puromycin-resistant cells were present in the clones, CG4 clones transduced with vector, PTP! shRNA-1 or PTP! shRNA-3 were analysed by flow cytometry using GFP as a positive control. GFP-positive cell populations of 81.57%, 79.95% and 74.45% were observed in CG4 clones transduced with vector, PTP! shRNA-1 or PTP! shRNA-3, respectively. To exclude GFP-negative or low GFP-expressing cells, cells with high GFP expression were selected by sorting. However, there was still no significant PTP! knockdown observed in these high GFP-positive CG4 clones transduced with PTP! shRNA-3 (data not shown), suggesting that the target sequence of this PTP! shRNA (and likely the others) was not efficient. Therefore, I focused on the isolation and culture of primary mouse OPCs from wild-type (WT) and PTP!-/- mice.  3.2  Characterization of primary mouse oligodendroglia cells isolated and  cultured from postnatal day 1 (P1) mice In addition to knocking down PTP! expression by siRNA in oligodendroglia cell lines, I wished to use primary OPCs isolated from WT and PTP!-/- mice as another model system. The traditional method for establishing oligodendroglial cell cultures  !  68!  from rat brain was developed in 1980 (McCarthy and de Vellis, 1980). This method is based on (1) the absence of viable neurons in cultures prepared from postnatal rat cerebrum, (2) the stratification of astrocytes and oligodendrocytes in cultures, and (3) the selective detachment of the overlaying oligodendrocytes from the underlying astrocytes. Although this procedure generates highly pure rat oligodendroglia cells, it is not suitable for isolation of oligodendroglia cells from the mouse. I attempted several times to culture primary mouse OPCs according to this method (see Materials and Methods, section 2.2.3.1), but found that the quantity and purity of the cells were insufficient. Mouse OPCs are reportedly more difficult to isolate as cultured cells than rat OPCs, because they do not share all of the cell surface antigens with rat, such as A2B5, and they tend to differentiate in in vitro mixed glial cultures (Chen et al., 2007; Fanarraga et al., 1995; Vitry et al., 1999). WT oligodendroglia cells prepared according to McCarthy et al. (McCarthy and de Vellis, 1980) were seeded on PDL-coated chamber slides for 3 days in BME medium followed by incubation for 3-6 days in CG4 differentiation medium. Cells were immunostained with antibodies against A2B5, GFAP and MAP2. As shown in Fig. 3.4A, most of the cells were A2B5-positive OPCs before induction of differentiation. However, some cells were GFAP- and MAP2-positive, suggesting that the cultures contained some astrocytes and neurons. After differentiation for 3 days, A2B5 signal decreased, some cells differentiated into astrocytes, and the MAP2-positive cells extended multiple processes (Fig. 3.4A). Significant cell death was observed after differentiation for 6 days, suggesting that some of the differentiated cells need trophic factors for survival that were not present in the differentiation medium. BME medium  !  69!  contains a high concentration of FBS (15%), which may promote survival of other types of neural cells and differentiation of OPCs into astrocytes. I next tried to seed the cells in CG4 proliferation medium instead of BME medium, and cultured them in the medium for only one day rather than three days. The cells were then induced to differentiate for only 1-3 days, to circumvent the extensive cell deathobserved after longer periods. As shown in Fig. 3.4B, most cells were A2B5-positive with some O4positive pre-oligodendrocytes present before differentiation (day 0), indicating that the oligodendroglia cultures contain mixed populations of OPCs and pre-oligodendrocytes, as reported by others (Holmes et al., 1988). After differentiation for 1 day, some A2B5positive cells extended multiple processes and some O4-positive cells were observed (Fig. 3.4B). After differentiation for 3 days, A2B5 signal decreased and more O4positive cells were observed (Fig. 3.4B). The lack of purity and the insufficient number of cells that could be isolated in one preparation indicated that cells generated by this procedure are not ideal for biochemical and molecular studies. I thus sought another method for isolation and culture of mouse OPCs.  3.3  Characterization of primary mouse neural progenitor/stem cells and OPCs  isolated and cultured from mouse embryos Several studies have determined that self-renewing OPCs can be generated from neural progenitor/stem cells of different species (Avellana-Adalid et al., 1996; Chen et al., 2007; Pedraza et al., 2008a; Vitry et al., 1999; Zhang et al., 1998a; Zhang et al., 1998b), and I utilized these procedures according to the protocols reported by Chen et al. (Chen et al., 2007) and Pedraza et al. (Pedraza et al., 2008a) (see Materials and  !  70!  Methods, section 2.2.3.2) to isolate and culture OPCs from WT and PTP!-/- mouse embryos. In addition to isolating and culturing primary mouse OPCs from forebrain, I also successfully adapted and used this method for isolation and culture of primary OPCs from spinal cord.  3.3.1 Forebrain WT mouse neural progenitor/stem cells from passage 2 neurospheres were seeded on PDLO/gelatin-coated dishes in neural culture medium with EGF/bFGF and maintained for two days. Cells were fixed and immunostained with antibodies against the neural progenitor/stem cell marker nestin. As shown in Fig. 3.5, greater than 90% of WT cells were positive for nestin. The cells were then induced to differentiate along three different lineages by culture in neural culture medium with 1% FBS for 4 days. Cells were fixed and immunostained with antibodies against the neuron marker MAP2, the pre-oligodendrocyte marker O4, and the astrocyte marker GFAP. As shown in Fig. 3.5, most cells differentiated into astrocytes when cultured in the presence of 1% FBS, and some of them differentiated into neurons and pre-oligodendrocytes, indicating that the cells derived from neurospheres are multipotential neural progenitor/stem cells. Neuronal markers are not detected in OPC cultures derived from oligospheres (Pedraza et al., 2008a), suggesting that these OPCs are mainly oligodendrocyte-type-2 astrocyte (O-2A) progenitors that can differentiate into oligodendrocytes or type-2 astrocytes. WT neurospheres were dissociated and cultured in NCM with 20 ng/ml PDGF and bFGF to induce oligosphere formation. WT mouse OPCs from passage 2 oligospheres were seeded on PDLO-coated dishes in OPC proliferation medium for two  !  71!  days. Cells were fixed and immunostained with antibodies against the OPC marker NG2, the pre-oligodendrocyte marker O4, and the astrocyte marker GFAP. As shown in Fig. 3.6, more than 95% of WT mouse OPCs were positive for NG2 and negative for either O4 or GFAP, confirming them to be progenitors. The cells were then induced to differentiate by mitogen withdrawal and thyroid hormone (T3) exposure (Barres et al., 1994). In addition, N-acetyl-L-cysteine was added to support cell survival (Mayer and Noble, 1994). Two growth factors, insulin-like growth factor 1 (IGF-1) (Cui et al., 2005; McMorris and Dubois-Dalcq, 1988) and ciliary neurotrophic factor (CNTF) (Mayer et al., 1994) can promote OPC differentiation and survival. Therefore, I also examined the effects of IGF-1 and CNTF on the differentiation of mouse OPCs. I found that after differentiation for 2 days, the numbers of NG2-positive cells decreased and O4-positive cells increased in all conditions tested (Fig. 3.6). In addition, O4 immunostaining revealed that more cells extended highly branched processes in the presence of IGF-1, and that there were less O4-positive cells but more GFAP-positive cells in the presence of CNTF. Moreover, very few GFAP-positive cells were detected in OPC differentiation medium with or without IGF-1. These results are consistent with previous studies showing that IGF-1 induces OPCs to differentiate into OLs (Hsieh et al., 2004) while CNTF also induces OPCs to differentiate into type-2 astrocytes (Hughes et al., 1988). Therefore, to enrich the oligodendrocyte population after OPC differentiation, I used OPC differentiation medium with or without IGF-1 for further studies.  !  72!  3.3.2 Spinal cord WT mouse neural progenitor/stem cells from P2 neurospheres were seeded on PDLO/gelatin-coated dishes in neural culture medium with EGF/bFGF for two days. The cells were then induced to differentiate along three different lineages by culture in neural culture medium with 1% FBS for 4 days. Cells were fixed and immunostained with antibodies against the neuron marker MAP2, the pre-oligodendrocyte marker O4, and the astrocyte marker GFAP. As shown in Fig. 3.7, most cells differentiated into astrocytes when cultured in the presence of 1% FBS, and some of them differentiated into neurons and pre-oligodendrocytes, indicating that the cells derived from neurospheres are multipotential neural progenitor/stem cells. Spinal cord oligospheres were induced to form as described in section 3.3.1. WT mouse OPCs from passage 2 oligospheres were seeded on PDLO-coated dishes in OPC proliferation medium for two days. Cells were fixed and immunostained with antibodies against the OPC marker NG2, the pre-oligodendrocyte marker O4, and the astrocyte marker GFAP. As shown in Fig. 3.8, more than 90% of WT mouse OPCs were positive for NG2 and negative for either O4 or GFAP, confirming them to be progenitors. The cells were then induced to differentiate by mitogen withdrawal and thyroid hormone (T3) exposure as described in 3.3.1. I found that the numbers of NG2-positive cells decreased and O4-positive cells increased after differentiation for 2 days with or without IGF-1 (Fig. 3.8). In addition, there were no O4-positive cells but more GFAPpositive cells in the presence of CNTF. Moreover, very few GFAP-positive cells were detected in OPC differentiation medium with or without IGF-1.  !  73!  3.4  Discussion OLN-93 cells are transformed cells and do not express OPC markers. Therefore,  they are easy to culture and differentiate without extensive cell death. In addition, they extend highly complex processes in response to serum-withdrawal, so they are suitable for studying oligodendrocyte terminal differentiation and myelination. However, their physiology of proliferation and cell cycle regulation are different from primary OPCs, so they are not ideal models for the study of OPC proliferation and differentiation. CG4 cells are non-transformed OPCs and express OPC markers. They also require PDGF and bFGF for self-renewing proliferation and will differentiate in the absence of these two growth factors, similar to primary OPCs. Therefore, they are suitable for studying proliferation in the presence of PDGF/bFGF and differentiation in response to mitogenwithdrawal. Therefore, I chose the CG4 cell line as a model to study the role of PTP! in OPC proliferation and differentiation. Due to the different nature of primary OPCs derived from rat and mouse, I tested the established protocol for culturing primary rat OPCs, but found it unsuitable for culturing OPCs from mouse origin. Instead, based on published protocols for isolation and culture of primary mouse OPCs derived from neural stem/progenitor cells, I successfully established a protocol for primary mouse (forebrain) OPC culture. This permits the generation of a large quantity of cells that can be used for biochemical/molecular studies. In addition, I also established a protocol for inducing OPC differentiation into oligodendrocytes without extensive astrocytic differentiation. Therefore, my studies will be performed using OPCs obtained using this protocol.  !  74!  Moreover, I have demonstrated that using this protocol, primary mouse OPCs isolated and cultured from spinal cord can also be generated and induced to differentiate. Each system has unique advantages. The CG4 cell line provides a ready, easy to maintain, and abundant source of cells that are amenable to manipulation such as targeted, albeit transient, knockdown of PTP! and other molecules. The primary OPCs are more difficult to obtain both procedurally and in respect to quantity, but do not require manipulation to ablate PTP! as they are derived from genetically modified PTP!-null and counterpart WT mice. The in vitro generation of these OPCs from neurospheres allows more precise knowledge of their position in the oligodendrocyte maturation and differentiation pathways.  3.5  Summary I successfully established two in vitro oligodendroglial cell model systems for  studying  the  role  of  PTP!  in  oligodendrocyte  development:  rat-derived  oligodendroglial cell lines in which PTP! is silenced by RNAi; and oligosphere-derived primary mouse OPCs isolated and cultured from WT and PTP!-/- mouse embryos. The complementary features of the two model systems will optimize my investigations and be useful in cross-validating my findings.  !  75!  !  76!  !  !  !  77!  !  !  78!  !  !  79!  !  !  80!  !!  !  81!  !  !!  !  82!  !  !!!  !  83!  Chapter 4. PROTEIN TYROSINE PHOSPHATASE ! ACTS AS AN UPSTREAM REGULATOR OF FYN SIGNALING TO PROMOTE OLIGODENDROCYTE PROGENITOR CELL DIFFERENTIATION  4.1  Introduction and rationale The Src family tyrosine kinase (SFK) Fyn is an essential participant and central  co-ordinator of oligodendrocyte differentiation, maturation, and myelination. In vitro studies have linked Fyn activation or inhibition to several stimuli that respectively induce or inhibit oligodendrocyte differentiation and maturation. Fyn signals to several molecules that are important for oligodendrocyte morphological changes that require cytoskeletal rearrangement and process extension and elaboration, such as focal adhesion kinase (FAK), the Rho GTPases Rho, Rac1, and Cdc42, the Rho regulators p190 and p250 RhoGAP, tau protein, and possibly via the kinase Cdk5 to paxillin (Hoshina et al., 2007; Klein et al., 2002; Liang et al., 2004; Miyamoto et al., 2007; Taniguchi et al., 2003; Wolf et al., 2001). It also controls myelin production transcriptionally and postranscriptionally (Lu et al., 2005a; Umemori et al., 1999). The potential roles of PTPs in coupling upstream signals, many of which involve engagement of catalytically inactive receptors, to Fyn activation in oligodendrocyte differentiation have not been extensively investigated. Although PTP! mRNA is upregulated in oligodendrocyte differentiation, a role for PTP! in this process and in CNS myelination has not been described. I therefore investigated whether PTP! is required for oligodendrocyte differentiation and for Fyn activation and signaling in this process using two model systems; the cultured CG4 oligodendroglial cell line in which  !  84!  PTP! expression was ablated by siRNA-mediated silencing, and primary OPCs derived from wild-type (WT) and PTP!-/- mouse embryos.  4.2  Results  4.2.1 PTP! is required for CG4 differentiation It has been reported that PTP! mRNA expression increases during oligodendrocyte differentiation (Ranjan and Hudson, 1996). As shown in Fig. 4.1A, the PTP! protein level was upregulated ~1.5 fold in CG4 cells after differentiation for 2 days and was gradually upregulated to ~2.5 fold during differentiation day 3-6, indicating that PTP! may play a role in this process. Since PTP! can dephosphorylate and activate SFKs (Pallen, 2003), the protein expression of the three SFKs that are present in oligodendrocytes; Fyn, Lyn, and Src (Umemori et al., 1999), was characterized in differentiating CG4 cells. Among these SFKs, Fyn plays a unique role in myelination, since myelin deficits are only found in Fyn-/- mice and not in Lyn-/- or Src-/- mice (Sperber et al., 2001). As shown in Fig. 4.1B, Fyn protein level rapidly increased (2.8-fold) over the first 2 days of CG4 differentiation, and then increased slightly further and was maintained for the remainder of the 6 day differentiation period. Consistent with previous studies (Colognato et al., 2004; Lu et al., 2005a; Osterhout et al., 1999), we found that Lyn protein level increased by 1.7-fold over the first 2 days of differentiation, and then decreased over subsequent days to return to the starting level or somewhat lower by day 5-6. Src protein continually decreased during differentiation, and by day 6 was reduced to 35% of the starting level. In conjunction with these findings, and as Fyn is reported to be the  !  85!  only SFK with significant kinase activity in either cultured OPCs or oligodendrocytes (Osterhout et al., 1999), Fyn activity was investigated in differentiating CG4 cells by determining its phosphorylation status. Phosphorylation of Fyn at the negative regulatory C-terminal tail residue Tyr531 was significantly reduced during CG4 differentiation, especially over the first 2 days (Fig. 4.1C), indicative of Fyn activation. Overall, Fyn Tyr531 phosphorylation per unit Fyn protein decreased to 34% of the starting level by day 6. To investigate the role of PTP! in oligodendrocyte differentiation, PTP!deficient CG4 cells were generated using siRNA (as described in section 3.1.3). Cells were cultured in proliferation medium and transfected, then 24h later they were seeded on PDL-coated plates. After 3 hours of attachment in proliferation medium, the medium was changed to CG4 differentiation medium (differentiation day 0). Lysates of control siRNA- and PTP! siRNA-treated CG4 cells were prepared from cells maintained for 03 days in differentiation medium, and examined by immunoblotting to determine the effectiveness of siRNA-mediated knockdown (Fig. 4.2A). PTP! expression was reduced by more than 90% during the 24-72 h following siRNA transfection (differentiation day 0-2). To evaluate the differentiation of control siRNA- and PTP! siRNA-treated CG4 cells after 2 days in differentiation medium, they were immunostained with anti-A2B5 and anti-CNPase antibodies for microscopic visualization and quantitative measurements. The numbers of A2B5-positive cells were counted, and about 4 times more cells in the PTP!-knockdown CG4 population were found to remain A2B5-positive (progenitor-like) compared to the control siRNA-treated cell population (Fig. 4.2B and 4.2C). CNPase immunofluorescence revealed multiple  !  86!  branched processes that were formed by the control siRNA-treated cells, but that were lacking in the PTP!-directed siRNA-treated cells (Fig. 4.2B, bottom panels). As cell process extension is an indicator of differentiation, I counted the number of processes per cell with a length greater than that of the cell body. After 2 days in differentiation medium, significantly more PTP!-knockdown CG4 cells had a low number (2 or less) of extended processes per cell, and fewer PTP!-knockdown CG4 cells had a high number (4 or 5) of extended processes per cell, compared to control siRNA treated cells (Fig. 4.2B and 4.2D). Moreover, CNPase protein expression was lower in PTP!knockdown CG4 cells than in control siRNA-treated cells after differentiation (Fig. 4.3). These results indicate that PTP! is required for the differentiation of CG4 cells into OLs.  4.2.2 PTP! is required for the activation of Fyn and the Fyn effectors FAK, Rac1, and Cdc42 during CG4 differentiation To determine if PTP! is required to dephosphorylate and activate Fyn in differentiating CG4 cells, the phosphorylation of Fyn at Tyr531 was determined in control siRNA- and PTP! siRNA-treated CG4 cells that were induced to differentiate for 2 days. Fyn immunoprecipitates from PTP!-directed siRNA-treated cells contained higher levels of phosphoTyr531-Fyn than those from control siRNA-treated cells (Fig. 4.4A), indicating that PTP! is required to dephosphorylate this tyrosine residue and activate Fyn in differentiating CG4 cells. The effect of silencing PTP! on the activation of several downstream effectors of Fyn was examined. FAK activation involves the phosphorylation of FAK Tyr576  !  87!  (Calalb et al., 1995), and this is reported to be upregulated in a Fyn-dependent manner during differentiation of CG4 cells (Hoshina et al., 2007). I confirmed that the phosphorylation of FAK Tyr576 increases during CG4 differentiation (Fig. 4.4B, upper panels). This required PTP!, since compared to control siRNA-treated CG4 cells, the PTP! siRNA-treated cells displayed significantly reduced (by ~40%) phosphoTyr576FAK after induction of differentiation (Fig. 4.4B, lower panels). The Rho family GTPases Rac1 and Cdc42 play important roles in cytoskeleton rearrangement and are crucial for the morphological differentiation of oligodendrocytes and myelination (Liang et al., 2004; Thurnherr et al., 2006a). It has also been reported that activation of Rac1 and Cdc42 is dependent on the activity of Fyn and FAK in differentiating oligodendrocytes (Hoshina et al., 2007; Liang et al., 2004). I therefore investigated whether the activities of Rac1 and Cdc42 were affected by PTP! silencing in CG4 cells following the induction of differentiation. Using GST-PBD (PAK-binding domain) pull-down assays to measure the levels of active GTP-bound Rac1 and Cdc42, I found that both Rac1 and Cdc42 were activated in CG4 cells during differentiation (Fig. 4.5, upper panels). However, the differentiation- induced activity of both Rac1 and Cdc42 was significantly reduced by more than 50% in PTP!-knockdown CG4 cells placed in differentiation medium for 2 days (Fig. 4.5, lower panels). These results indicate that PTP! is required for Fyn-mediated signaling to FAK, Rac1 and Cdc42 in differentiating CG4 cells.  !  88!  4.2.3 PTP! is not required for Fyn-mediated signaling to p190RhoGAP, but is required for Rho inactivation during CG4 differentiation During oligodendrocyte differentiation, the Fyn interacting protein and substrate p190RhoGAP is tyrosine phosphorylated, resulting in increased p190RhoGAP activity that promotes Rho inhibition and oligodendrocyte differentiation (Liang et al., 2004; Wolf et al., 2001). In accord with these findings, p190RhoGAP co-immunoprecipitated with Fyn in both progenitor and differentiating CG4 cells and the tyrosine phosphorylation of p190RhoGAP increased after differentiation for 2 days (Fig. 4.6A). To investigate if PTP! is an upstream regulator of differentiation-induced Fyn signaling to p190RhoGAP, the Fyn and p190RhoGAP association and p190RhoGAP tyrosine phosphorylation were determined in CG4 cells that were treated with control siRNA or PTP!-directed siRNA and induced to differentiate for 2 days. Surprisingly, both the Fyn-p190RhoGAP interaction and tyrosine phosphorylation of p190RhoGAP were not affected by PTP! siRNA treatment (Fig. 4.6B). These results suggest that PTP! does not act upstream of Fyn-mediated regulation of p190RhoGAP, and that PTP! thus regulates specific aspects of Fyn signaling in differentiating CG4 cells. The Rho family GTPase Rho plays important roles in controlling cellular morphology. Overexpression of constitutively active Rho inhibits process extension in oligodendrocytes, whereas overexpression of dominant-negative Rho results in a hyperextension of oligodendrocyte processes (Wolf et al., 2001). Since the primary function of p190RhoGAP is to inactivate Rho, I determined Rho activities to determine if, like p190RhoGAP, this was unaffected upon PTP! silencing. GST-RBD (Rhotekinbinding domain) pull-down assays were utilized to measure the levels of active GTP-  !  89!  bound Rho, and demonstrated that Rho was inactivated in CG4 cells during differentiation (Fig. 4.7). Compared to control CG4 cells, Rho activity was significantly increased by ~3 fold in PTP!-knockdown CG4 cells placed in differentiation medium for 2 days (Fig. 4.7). Taken together, these results indicate that PTP! is required for Rho inactivation in differentiating CG4 cells, but in a manner independent of Fynp190RhoGAP signaling.  4.2.4 PTP! is required for primary mouse OPC differentiation To extend the findings described above from CG4 cells where PTP! expression was transiently silenced, I investigated the role of PTP! in OPC differentiation in a different model system where PTP! expression was permanently ablated, using primary OPCs isolated and cultured from PTP!-null mouse embryos. To characterize the role of PTP! in primary mouse OLs, PTP! protein expression during WT OPC differentiation was first examined. Consistent with observations in CG4 cells, PTP! protein expression, as well as that of Fyn and CNPase, increased during the differentiation of primary mouse OPCs (Fig. 4.8). To investigate the role of PTP! in the differentiation of primary mouse OPCs, WT and PTP!-/- OPCs were induced to differentiate for 2 days in OPC differentiation medium with or without IGF-1. After differentiation in both conditions, PTP!-/oligodendrocytes expressed less of the oligodendrocyte marker CNPase than did WT OLs (Fig. 4.9), suggesting that their differentiation was impaired. This was confirmed by immunostaining with antibodies against the progenitor marker NG2 and the preoligodendrocyte marker O4. As shown in Fig. 4.10A, after differentiation for 2 days in  !  90!  media with or without IGF-1, fewer WT cells than PTP!-/- cells were NG2-positive (top panels), and more WT cells than PTP!-/- cells were O4-positive (lower panels). The NG2-positive and O4-positive cells of each genotype were quantified (Fig. 4.10B), demonstrating that about 3-4-fold higher populations of O4-positive WT cells than PTP!-/- cells, and about 3-3.5-fold lower populations of NG2-positive WT cells than PTP!-/- cells, were present after 2 days in differentiation medium either lacking or containing IGF-1. These results further confirm that PTP! is required for OPC differentiation.  4.2.5 PTP! selectively regulates Fyn activation and signaling in primary mouse OPCs To confirm that PTP! is required for Fyn dephosphorylation at its negative regulatory site, lysates from WT and PTP!-/- progenitors or differentiating oligodendrocytes were immunoprecipitated with anti-Fyn antibody followed by immunoblotting. Fyn phospho-Tyr528 (the equivalent of rat Fyn Tyr531) was enhanced 2-fold in PTP!-/- OPCs and differentiating oligodendrocytes compared to the WT group, suggesting that PTP! is required for Fyn dephosphorylation at Tyr528 in both progenitors and oligodendrocytes (Fig. 4.11A). Fyn phospho-Tyr528 decreased by ~40% in both WT and PTP!-/- cells after differentiation in the presence or absence of IGF-1 for 2 days (Fig. 4.11A), suggesting that there are also other regulators of Fyn activation during OPC differentiation. Despite the decreased phosphorylation of Fyn Tyr528 that occurs in both WT and PTP!-/- cells during differentiation, the level of Fyn Tyr528 phosphorylation in the PTP!-/- cells after 2 days differentiation was not  !  91!  significantly lower than that in undifferentiated WT cells (Fig. 4.11A). In conjunction with the impaired differentiation of PTP!-/- cells, this suggests that non-PTP!-mediated Fyn Tyr528 activation is insufficient to promote mouse OPC differentiation. To investigate whether PTP! is required for FAK activation in primary mouse OPCs, FAK Tyr576 phosphorylation was determined in WT and PTP!-/- progenitors and differentiating oligodendrocytes. As shown in Fig. 4.11B, FAK phospho-Tyr576 was significantly reduced by 60% and 50%, respectively, in PTP!-/- progenitors and differentiating oligodendrocytes compared to WT cells. Although FAK phosphorylation at Tyr576 increased in both WT and PTP!-/- cells after differentiation was induced, the FAK phosphorylation level in the PTP!-/- cells only increased to a level equivalent to that in undifferentiated WT cells (Fig. 4.11B). Thus the differentiation-induced modulation of FAK in both WT and PTP!-/- cells correlated closely with that of Fyn, further indicating that PTP! is required for Fyn-mediated FAK activity in primary mouse OPCs and differentiating oligodendrocytes. To determine whether PTP! is required for Fyn-p190RhoGAP signaling in the primary mouse cell system, p190RhoGAP immunoprecipitates were prepared from lysates of WT and PTP!-/- progenitors and differentiating oligodendrocytes and analyzed. Increased amounts of p190RhoGAP immunoprecipitated from both WT and PTP!-/- cells that had been induced to differentiate for 2 days compared to the undifferentiated OPCs (Fig. 4.12A, middle panel), but determination of the phosphotyrosine incorporated into p190RhoGAP protein revealed that there were no differentiation-induced changes in the tyrosine phosphorylation of p190RhoGAP in either cell type nor between the cell types (Fig. 4.12A, top panel and graph). It has been  !  92!  reported that p190RhoGAP associates with p120RasGAP in oligodendrocytes (Wolf et al., 2001), and that the SH2 domain of p120RasGAP binds tyrosine phosphorylated p190RhoGAP (Bryant et al., 1995). I investigated whether PTP! might affect p190RhoGAP:p120RasGAP  complex  formation  upon  differentiation.  While  p120RasGAP was detected in p190RhoGAP immunoprecipitates, no difference in the extent of association was apparent between WT and PTP!-/- cells or between progenitors and oligodendrocytes of each genotype (Fig. 4.12A, bottom panel), indicating that PTP! is not required for Fyn-mediated p190RhoGAP-p120RasGAP signaling in progenitors and oligodendrocytes. To determine if PTP! is required for Rac1 and Cdc42 activation and Rho inactivation in the primary mouse cell system, I examined Rac1, Cdc42 and Rho activities in WT and PTP!-/- oligodendrocytes. As shown in Fig. 4.12B and 4.12C, Rac1 and Cdc42 activities were reduced, and Rho activity was increased in PTP!-/oligodendrocytes compared to WT cells.  4.2.6 Ablation of PTP! results in decreased MBP protein expression in primary mouse oligodendrocytes and leads to defective myelination Fyn directly stimulates the promoter activity of the MBP gene and is involved in posttranscriptional regulation of MBP mRNA (Lu et al., 2005a; Umemori et al., 1999). Therefore, I investigated if PTP! is required for Fyn-MBP signaling. The WT and PTP!-/- OPCs were induced to differentiate for 2 days in OPC differentiation medium with  or  without  IGF-1.  After  differentiation  in  both  conditions,  PTP!-/-  oligodendrocytes expressed less MBP than WT oligodendrocytes (Fig. 4.13A),  !  93!  suggesting that PTP! is also required for Fyn-mediated upregulation of MBP expression. In support of the above in vitro results, myelinated fibers in WT and PTP!-/mouse brains were examined for MBP immunoreactivity. Fewer myelinated fibers could be observed in the corpus callosum of P18 PTP!-/- mouse brains (Fig. 4.13B) and in the cortex and striatum of P10 and P18 PTP!-/- mouse brains (Fig. 4.13C). Taken together, these results demonstrate that PTP! is involved in regulating MBP expression during OPC differentiation and thus is required for proper myelination in the brain.  4.3  Discussion In this study, I have demonstrated that PTP! is required for OPC differentiation  using two distinct model cell systems. The siRNA-mediated silencing of PTP! in the rat CG4 OPC cell line results in impaired differentiation to oligodendrocytes as evidenced by the prolonged maintenance of a high population of A2B5-positive population of progenitor cells, the inhibition of process extension, and the reduced expression of the maturation marker CNPase that is localized to cell bodies and processes. Oligosphere-derived OPCs isolated from PTP!-/- mouse embryos likewise exhibit an oligodendrocyte differentiation defect as determined by elevated NG2positive and reduced O4-positive populations, the appearance of few oligodendrocytes with a mature morphology of multiple/branched processes, and reduced CNPase and MBP expression as compared to the cells isolated from WT mouse embryos. Furthermore, defective differentiation of PTP!-deficient OPCs correlates with a physiological defect in CNS myelination, since relative to WT mice, PTP!-/- mice have  !  94!  a readily apparent overall reduction in myelin in forebrain sections detected by immunostaining of MBP. Fyn is activated during OPC differentiation, and this is critical for morphological differentiation, maturation and CNS myelination (Biffiger et al., 2000; Osterhout et al., 1999; Sperber et al., 2001; Umemori et al., 1999). Several upstream molecules stimulate Fyn activity in this process, including the ligand-receptor interactions of ECM components like vitronectin and fibronectin with "1 integrins, laminin 2 binding to !6"1 integrin, and the laminin family member netrin 1 and its receptor Dcc (Colognato et al., 2004; Liang et al., 2004; Rajasekharan et al., 2009). Other receptors such as FcR% (upon crosslinking of bound immunoglobulin G) (Nakahara et al., 2003) and the PTP CD45 (Nakahara et al., 2005) can also promote Fyn activation during oligodendrocyte differentiation/myelination. My findings identify PTP! as an additional upstream activator of Fyn in oligodendrocyte differentiation. In CG4 cells, the differentiationassociated activation of Fyn, as measured by reduced phosphorylation of its inhibitory tyrosine residue, is reduced by siRNA-mediated silencing of PTP! expression. Likewise, differentiating primary mouse OPCs lacking PTP! contain less activated Fyn than do WT mouse OPCs, irrespective of whether differentiation was induced by PDGF/bFGF withdrawal in the presence or absence of IGF-1. Interestingly, despite the reduced level of activated Fyn in differentiating PTP!-null OPCs, differentiation cues still stimulated some Fyn activation in PTP!-/- cells. Thus, Fyn activation during this process is not exclusively regulated by PTP! but is also controlled by PTP!independent mechanisms. These could involve inhibition of the C-terminal Src kinase (Csk) that phosphorylates the negative regulatory site of SFKs, or dephosphorylation by  !  95!  other PTPs (Brauninger et al., 1992; Granot-Attas and Elson, 2004; Nakahara et al., 2005). Nevertheless, in the absence of PTP!-activated Fyn, the Fyn activation that is mediated by these other mechanisms is insufficient to promote oligodendrocyte differentiation. This may be because other upstream activators cannot stimulate Fyn activity to a level required for differentiation, as supported by the finding that the level of active Fyn detected in PTP!-/- cells after 2 days of differentiation was similar to that in undifferentiated WT OPCS, and/or because PTP! has other unique actions that are required for differentiation. In accord with the notion of there being insufficient Fyn activation in differentiation-induced PTP!-depleted or -null OPCs to effectively promote differentiation, I observed significantly impaired regulation of the Fyn downstream effectors FAK and the RhoGTPases Rac1, Cdc42, and Rho. FAK has been implicated in CNS myelination, and is proposed to regulate oligodendrocyte process outgrowth and/or remodeling (Forrest et al., 2009; Hoshina et al., 2007). Fyn-mediated signaling through activation of Rac1 and Cdc42 and inhibition of Rho is important for cytoskeletal alterations involved in process extension and branching that occur during the morphological differentiation of oligodendrocytes (Liang et al., 2004; Thurnherr et al., 2006a; Wolf et al., 2001). Diverse Fyn-FAK signaling mechanisms that regulate these RhoGTPases during oligodendrocyte differentiation have been described. For example,  laminin  stimulation  induces  Fyn-FAK-Rac1/Cdc42  signaling  in  oligodendrocyte differentiation (Hoshina et al., 2007), while netrin-1 stimulates recruitment of Fyn to the netrin-1 receptor Dcc that is complexed with FAK and thereby promotes the inhibition of Rho without affecting Rac1/Cdc42 (Rajasekharan et al.,  !  96!  2009). The ablated Fyn-FAK to RhoGTPase signaling in PTP!-deficient cells, comprising impaired activation of Rac1 and Cdc42 and defective inhibition of Rho, is likely a major defect contributing to their reduced process extension and maturation. Fyn-mediated inhibition of Rho in differentiating oligodendrocytes is regulated by Fyn phosphorylating and activating p190RhoGAP, and is essential for process extension and differentiation (Liang et al., 2004). The differentiation-induced phosphorylation of p190RhoGAP, as well as its interaction with Fyn and p120RasGAP are not altered by the lack of PTP! in differentiation-induced CG4 cells or primary mouse oligodendrocytes. This indicates that although essential, p190RhoGAP activation is not sufficient to inhibit Rho or promote process extension. Furthermore, my results suggest that oligodendrocyte differentiation involves distinct pathways that regulate Rho; one that appears to involve Fyn-mediated p190RhoGAP activation that is PTP!-independent, and another that requires PTP! and may be Fyn-dependent but is p190RhoGAP-independent. The latter may represent a distinct, possibly specific action of PTP!, and could utilize other Fyn regulated RhoGAPs, such as the p250RhoGAP implicated in oligodendrocyte differentiation (Taniguchi et al., 2003). Pending the identification of the specific Rho regulator(s) involved it is nonetheless clear that it is critical  for  optimal  PTP!-mediated  Rho  inhibition  during  oligodendrocyte  differentiation. The development of OPCs into mature oligodendrocytes is a complex process that requires exit from the cell cycle, expression of oligodendrocyte-specific genes, and extension of processes and myelin sheets. Another possible role for PTP! in regulating oligodendrocyte differentiation is that PTP! functions in progenitor cells to regulate  !  97!  survival, proliferation or cell cycle exit. Indeed, I observe an increased number of PTP!-/- primary mouse cells compared to WT cells when they are grown as oligospheres, but not with cells grown as neurospheres (data not shown), suggesting that PTP! functions to control proliferation or survival of oligodendrocyte lineage cells but not cells that are at an earlier stage of development. Whether PTP! is necessary for these or other OPC processes that position the progenitor cells to respond appropriately to differentiation stimuli requires further investigation.  4.4  Summary In summary, I have identified PTP! as a novel regulator of oligodendrocyte  differentiation and in vivo CNS myelination. I propose that the major function of PTP! in promoting these processes is through activation of Fyn, in accord with the well characterized role of PTP! as an activator of SFKs (Pallen, 2003) and with the overlapping phenotypes of defective forebrain myelination in PTP!-/- and Fyn-/- mice. This study reveals PTP! to be an essential, but not the sole, regulator of Fyn in differentiating OPCs. Furthermore, PTP! is required for activation of the Fyn effectors FAK, Rac1, and Cdc42 and for Rho inhibition during oligodendrocyte differentiation, and it mediates the latter through a p190RhoGAP-independent mechanism. This suggests that upstream regulators such as PTP! are differentially coupled to various Fyn-effector signaling modules to provide stimulus-specific responses that determine aspects of the profound changes in gene expression and morphology that occur during oligodendrocyte differentiation.  !  98!  !  99!  !  100!  !  101!  !  102!  !  103!  !  104!  !  105!  !  106!  !  107!  !  108!  !  109!  !  110!  !  111!  Chapter 5. PROTEIN TYROSINE PHOSPHATASE ! (PTP!) IS A NEGATIVE REGULATOR OF OLIGODENDROCYTE PROGENITOR CELL SELF-RENEWAL  5.1  Introduction and rationale Oligodendrocytes are the myelin-forming cells of the CNS. They are derived from  oligodendrocyte progenitor cells (OPCs) that undergo a series of developmental stages before acquiring their mature myelinating function (Pfeiffer et al., 1993). OPCs arise and proliferate in certain locations and subsequently migrate through the CNS and stop at the correct location. The majority of OPC proliferation occurs in developing white matter. After a sufficient number of OPCs have been generated, these cells differentiate and mature into myelinating oligodendrocyte. The development of OPCs into mature oligodendrocytes is a complex process that requires exit from the cell cycle, expression of oligodendrocyte-specific genes, and extension of processes and myelin sheets. This is regulated by the presence or absence of growth factors, extracellular matrix and axonal signals (Miller, 2002; Rogister et al., 1999). A variety of growth factors have been shown to regulate oligodendrocyte development. The best characterized mitogens for OPCs are platelet-derived growth factor (PDGF) and basic fibroblast growth factor (bFGF) (Baron et al., 2000). PDGF induces OPCs to proliferate for a number of divisions, thereby preventing premature differentiation (Noble et al., 1988; Raff et al., 1988). bFGF blocks terminal differentiation and myelin gene expression at the late progenitor stage (Bansal and Pfeiffer, 1994; McKinnon et al., 1990). Moreover, bFGF can maintain a high level of  !  112!  expression of the PDGFR!, the only PDGF receptor expressed in oligodendrocytes (Hart et al., 1989; McKinnon et al., 1990). PDGF and bFGF cooperatively promote selfrenewal and survival and inhibit differentiation of OPCs (Barres et al., 1992a; Bogler et al., 1990; Yasuda et al., 1995). However, the downstream regulators of these growth factors remain poorly defined. My studies described in Chapter 4 demonstrate that PTP! is expressed and regulates Fyn signaling in OPCs to promote their differentiation upon cessation of proliferation. These results raise the question of whether PTP! regulates other processes that appropriately position OPCs to differentiate, such as proliferation and cell cycle exit. Indeed, PTP! has been shown to have positive and negative roles in growth control, which depend on the lineage of cells and the signals received. For example, PTP! activates Src and induces tumorigenesis in fibroblasts (Ardini et al., 2000), but inhibits proliferation of HER2/neu-transfected MCF-7 breast cancer cells and reduces their tumorigenicity (Ardini et al., 2000). I therefore investigated the role of PTP! in the proliferation of neural stem/progenitor cells and their derivative oligospheres and OPCs, prepared from wild-type (WT) and PTP!-/- mouse embryos, and in the CG4 oligodendroglial cell line.  5.2  Results  5.2.1 Abnormal populations of OPCs and oligodendrocytes in the corpus callosum of PTP! -/- mice I have shown that PTP! promotes oligodendrocyte differentiation by activating Fyn signaling and that fewer myelinated fibers are detected in PTP!-/- mouse brain as  !  113!  determined by immunostaining of myelin basic protein (Wang et al., 2009). To determine whether the maturation of OPCs into myelin-forming cells was impaired in vivo, immunophenotypic analysis of oligodendrocyte lineage cells in brain sections of WT and PTP!-/- mice at postnatal days 10 and 18 was conducted. Double immunofluorescence with antibodies against NG2 and CC1 was used to identify progenitors and mature oligodendrocytes, respectively. Images were acquired and the relative proportion of NG2-positive and CC1-positive cells was calculated as a percentage of the total oligodendrocyte lineage population. At P10, the oligodendrocyte lineage population in the corpus callosum of WT mice was composed of 55.3% ± 3.2% CC1-positive cells while in PTP!-/- mice only 37% ± 6.8% of the cells were CC1positive (Fig. 5.1A). However, the difference in CC1-positive populations between WT and PTP!-/- mice decreased by P18 (76.1% ± 3.1% vs. 66.7% ± 3.4%, respectively) compared to P10, suggesting that the lack of PTP! delayed but did not completely block the maturation of oligodendrocyte lineage cells. This result was confirmed by immunofluoresence using antibodies specific for NG2 and for the lipid sulfatide recognized by O4 (Fig. 5.1B).  5.2.2 PTP! negatively regulates OPC growth and growth factor dependency Cell proliferation and differentiation are highly coordinated processes during development. If cells persist in the cell cycle, they do not differentiate properly. Therefore, it is possible that PTP! functions in progenitor cells as regulators of proliferation, cell cycle exit and survival. To test this hypothesis, I used primary OPCs isolated and cultured from WT and PTP!-null (PTP!-/-) mouse embryos (see Materials  !  114!  and Methods, section 3.3). I first examined the growth of WT and PTP!-/- OPCs cultured as oligospheres. As shown in Fig. 5.2A, after culture in OPC proliferation medium for 5 days, PTP!-/- oligospheres were larger than WT oligospheres. The size of the oligospheres was measured and the percentage of oligospheres larger than 100µm was calculated, revealing that a higher percentage of PTP!-/- oligospheres than WT oligospheres had a size bigger than 100µm. It has been shown that neurospheres are motile and they aggregate under culture conditions (Singec et al., 2006). To rule out the possibility that PTP!-/- oligospheres tend to merge to form larger spheres and confirm that the sizes of WT and PTP!-/- cells are different, WT and PTP!-/- oligospheres were collected and dissociated and total cell numbers were counted. This demonstrated that indeed the PTP!-/- oligospheres were composed of more cells than the WT oligospheres. To further confirm that PTP!-/- OPCs grow faster than WT OPCs in adherent culture, oligospheres were dissociated and seeded on PDLO-coated dishes in OPC proliferation medium for 2 weeks. PTP!-/- OPCs grew to a higher density and formed larger colonies compared to WT OPCs (Fig. 5.2B). These results indicate that the proliferation of PTP!-/- OPCs is faster than WT OPCs. The best-characterized mitogens for OPCs are PDGF and bFGF (Baron et al., 2000), and both PDGF and bFGF are survival factors for OPCs (Barres et al., 1992a; Yasuda et al., 1995). In the oligosphere and OPC culture system, cells undergo selfrenewing cell division in OPC proliferation medium, which contains PDGF and bFGF. I next determined if PTP!-/- OPCs are hypersensitive to PDGF/bFGF stimulation. As shown in Fig. 5.3, both WT and PTP!-/- OPCs cannot form new spheres in the absence of these growth factors, but PTP!-/- OPCs can form more and larger new spheres at  !  115!  lower concentrations of growth factors than WT OPCs. This indicates that PTP!-/OPCs are hypersensitive to PDGF/bFGF, but that PTP! loss did not render oligosphere formation growth factor-independent.  5.2.3 PTP! negatively regulates OPC proliferation, cell cycle entry and survival in response to PDGF/bFGF The accumulation of large numbers of PTP!-/- OPCs may result from an increase in cell proliferation, a decrease in cell cycle exit and/or a decrease in cell death. To address which mechanism PTP! affects to regulate OPC growth, I assessed these parameters in WT and PTP!-/- OPCs. WT and PTP!-/- oligospheres were dissociated and analyzed by flow cytometry. As shown in Fig. 5.4A, more PTP!-/- cells than WT cells were in S phase. This was confirmed by BrdU incorporation and immunostaining with BrdU antibody. WT and PTP!-/- oligospheres were dissociated and seeded on PDLO-coated chamber slides in proliferation medium for 2 days followed by culture in fresh medium with or without PDGF/bFGF for another 24 hours. BrdU incorporation was significantly increased in PTP!-/- OPCs compared with WT OPCs in the presence or absence of PDGF/bFGF (Fig. 5.4B). Ki-67 is a protein associated with proliferation that is present in cells that are within the cell cycle but not in cells that have exited the cell cycle. To address whether PTP! regulates cell cycle exit, immunofluoresence with antibody against Ki-67 was performed. This demonstrated that more PTP!-/- cells than WT cells are in the cell cycle in the presence of PDGF/bFGF (Fig. 5.5A). In addition, the percentage of Ki-67-positive WT cells cultured in the absence of PDGF/bFGF decreased by 19.12% compared to that cultured in the presence of PDGF/bFGF, while  !  116!  the percentage of Ki-67-positive PTP!-/- cells cultured in the absence of PDGF/bFGF only decreased by 4.71% compared to that cultured in the presence of PDGF/bFGF. This indicates that PTP!-/- cells are more resistant to PDGF/bFGF withdrawal-induced cell cycle exit. Since PDGF and bFGF are both survival factors for OPCs, I next determined if PDGF/bFGF-mediated survival is increased in PTP!-/- OPCs. To induce apoptosis, supplement (including insulin) was removed from the culture medium for 24 hours, and the cells were labeled with the apoptosis marker, cleaved caspase-3. Significantly fewer PTP!-/- OPCs compared than WT OPCs exhibited caspase-3 cleavage when cultured in medium without supplement, while the percentages of apoptotic cells in WT and PTP!-/- OPCs cultured under normal conditions (with supplement) were similar (Fig. 5.5B). Therefore, PDGF/bFGF-mediated survival is increased in PTP!-/- OPCs, since PTP!-/- OPCs are more resistant to supplement withdrawal-induced apoptosis.  5.2.4 Lack of PTP! does not affect proliferation of neural stem/progenitor cells The effect of PTP! ablation of enhancing OPC proliferation raised the question of how early PTP! is expressed during development and whether it regulates the proliferation and cell cycle exit of cells at an earlier stage of differentiation (neural stem/progenitor cells). It has been shown that PTP! mRNA is not detectable in undifferentiated mouse embryonic pluripotent stem cells (P19 embryonic carcinoma cells), but is expressed in neuroectoderm-like cells (retinoic acid-treated P19 derivatives) (den Hertog et al., 1993). Therefore, the expression of PTP! protein in neurospheres composed of neural stem/progenitor cells was determined and compared with that in  !  117!  oligospheres composed of OPCs. PTP! protein and its substrate Fyn were detectable in neurospheres, but at lower levels than in oligospheres (Fig. 5.6A). I next examined the expression of the proliferation marker PCNA in WT and PTP!-/- neurospheres and oligospheres. As shown in Fig. 5.6B, in contrast to the different expression levels of PCNA in WT and PTP!-/- oligospheres, PCNA expression levels are similar in WT and PTP!-/- neurospheres, suggesting that PTP! does not have a significant role in regulating proliferation of neural stem/progenitor cells in response to EGF/bFGF. This finding was confirmed by determining BrdU incorporation and Ki-67 positivity, revealing that PTP! ablation does not significantly affect proliferation and cell cycle exit of neural stem/progenitor cells (Fig. 5.6C).  5.2.5 PTP! negatively regulates the activities of the Ras and Rho family small GTPases, Cdc42, Rac1 and Rho, in OPCs Ras proteins are small guanine nucleotide-binding proteins that control a variety of cellular processes including proliferation, survival and differentiation, and are important downstream effectors of growth factor receptor tyrosine kinase signaling pathways (Karnoub and Weinberg, 2008). It has also been reported that Ras plays an important role in oligodendrocyte development (Barnett and Crouch, 1995; Barnett et al., 1998; Bennett et al., 2003). I therefore investigated whether the activity of Ras was affected by PTP! ablation in OPCs. GST-Raf1 RBD pulldown assays were utilized to measure the levels of active GTP-bound Ras, and this demonstrated that in both PTP!-/oligospheres and OPCs, Ras activities were increased compared to those in WT oligospheres and OPCs (Fig. 5.7A). These results suggest that PTP! negatively  !  118!  regulates Ras activity in OPCs. Ras can activate at least three downstream pathways: Rho family small GTPases, mitogen-activated protein kinase (MAPK) and Akt pathways (Zhu and Parada, 2002). I next determined which downstream effector was affected by PTP! ablation. Using GST-PBD pulldown assays to measure the levels of active GTP-bound Cdc42 and Rac1 demonstrated that PTP!-/- OPCs exhibit increased activities of Cdc42 and Rac1 (Fig. 5.7B). Also, Rho activity was significantly increased in PTP!-/- OPCs, as detected using GST-RBD pulldown assays (Fig. 5.7B). However, the phosphorylation of ERK1/2 (extracellular signal-regulated kinase 1/2) at Thr202/Tyr204 and that of Akt at Ser473 were not affected in PTP!-/- OPCs after culturing in proliferation medium for 2 days (Fig. 5.8). These results indicate that PTP! negatively regulates the activities of Rho, Rac1 and Cdc42, but does not affect the phosphorylation status of ERK1/2 and Akt in this cell culture condition and at the time point of investigation.  5.2.6 PTP! promotes the expression of p27Kip1 protein The cell cycle inhibitory protein p27Kip1 (p27) plays a critical role in cell cycle regulation in response to the extracellular environment, such as growth factors and the extracellular matrix. The levels of p27 must remain low to permit cell cycle progression, and the signaling pathways active in proliferating cells are responsible for the suppression of p27. For example, Ras activation is required for the suppression of p27 levels throughout the cell cycle (Sa and Stacey, 2004). On the other hand, p27 can also regulate Ras activation by preventing Grb2-SOS formation (Moeller et al., 2003). Overexpression of p27 reduces cell proliferation and self-renewal and promotes cell  !  119!  death in neural progenitor cells (Li et al., 2009). In oligodendrocyte lineage cells, p27 is a crucial regulator of the decision to withdraw from the cell cycle (Casaccia-Bonnefil et al., 1997). Therefore, I determined if p27 expression is altered in PTP!-/- OPCs. As shown in Fig. 5.9A, decreased (by about 30%) p27 expression was detected in both PTP!-/- oligospheres and OPCs, indicating that PTP! may upregulate p27 expression to promote cell cycle withdrawal. Since accumulation of p27 is required for OPC differentiation (Casaccia-Bonnefil et al., 1997; Durand et al., 1997; Dyer and Cepko, 2001; Friessen et al., 1997) and since PTP!-/- OPCs fail to differentiate with proper timing (Chapter 4, and Wang et al., 2009), p27 expression was monitored before and after differentiation. As shown in Fig. 5.9B, differentiation induction by mitogenwithdrawal and T3 exposure induced p27 accumulation in both WT and PTP!-/- cells, but a lower expression of p27 was observed in both PTP!-/- OPCs and differentiating OLs compared to WT cells. These results suggest that PTP! promotes OPC differentiation, at least partially, by facilitating p27 accumulation and cell cycle exit, leading to decreased self-renewing proliferation and increased cell fate commitment.  5.2.7 Hyperproliferation of PTP! -/- OPCs is not due to upregulation of PDGFR! Activation of PDGFR! promotes proliferation and inhibits premature differentiation of OPCs (Noble et al., 1988; Raff et al., 1988). It has been shown that SFKs contribute to degradation of the PDGFR! through c-Cbl (Rosenkranz et al., 2000) and that activation of Fyn/c-Cbl pathways results in the reduced level of the PDGFR! in OPCs (Li et al., 2007). Therefore, it is possible that PTP! acts through a Fyn/c-Cbl  !  120!  pathway to downregulate PDGFR! expression. However, no difference in PDGFR! expression was detected in between PTP!-/- and WT oligospheres or OPCs (Fig. 5.10).  5.2.8 Higher Ras activity in PTP! -/- OPCs is not due to sequestration of the negative regulator p120 RasGAP by higher levels of FAK p120 RasGAP is a negative regulator of Ras, and it interacts with Ras to trigger Ras intrinsic GTPase activity (Vogel et al., 1988). It has been reported that elevated expression of FAK can promote Ras activity through the competitive recruitment of p120 RasGAP, thereby diminishing the association of p120 RasGAP with active Ras (Hecker et al., 2004). Since FAK protein expression is increased in PTP!-/- OPCs (Fig. 5.11A), I investigated whether the higher Ras activity in PTP!-/- OPCs is due to the elevated expression of FAK and consequently increased binding to FAK of p120RasGAP. According to Hecker et al. (Hecker et al., 2004), the phosphorylation of tyrosine residue 397 (Tyr397) of FAK is required for its association with p120 RasGAP. I assessed the phosphorylation status of FAK at Tyr397 in WT and PTP!-/- cells, and intriguingly, found that this was significantly decreased in PTP!-/- oligospheres and OPCs, even though the protein levels of FAK increased (Fig. 5.11B). In addition, I observed a decrease rather than an increase in FAK-p120 RasGAP complex formation in PTP!-/- oligospheres (Fig. 5.11C). Togather, these results suggest that the elevated expression of FAK in PTP!-/- cells does not result in increased sequestration of p120 RasGAP to promote Ras activity.  !  121!  5.2.9 PTP! negatively regulates multiple signaling pathways and is required for p27 accumulation in CG4 cells CG4 cells were used for subsequent investigation of PTP!-dependent signaling in oligodendrocyte precursor/progenitor cells due to the following reasons. Firstly, some of the primary OPCs differentiate spontaneously, leading to a heterogeneous population of cells. Secondly, a large number of the limited and difficult to prepare primary OPCs is required for biochemical studies to elucidate molecular signaling mechanisms. Thirdly, it is difficult to synchronize the primary OPCs without inducing differentiation, therefore, some signaling events are difficult to manipulate and monitor. To study PTP!-dependent signaling in response to growth factor stimulation, CG4 cells were transfected with control siRNA and PTP! siRNA and starved overnight, followed by stimulation with CG4 proliferation medium for 3h. The increased activities of Ras and the Rho GTPases, Rac1, Cdc42 and Rho were confirmed in PTP!-knockdown CG4 cells (Fig. 5.12A, 5.12B). Phosphorylation of ERK1/2 and Akt was also determined by immunoblotting. In contrast to the results obtained with primary OPCs, the phosphorylation of ERK1/2 at Thr202/Tyr204 and that of Akt at Ser473 were upregulated 3h after stimulation in PTP!-knockdown CG4 cells (Fig. 5.12C). However, after stimulation for 24h, the phosphorylation of ERK1/2 at Thr202/Tyr204 and that of Akt at Ser473 decreased compared to 3h, and exhibited no significant difference between control and PTP!-knockdown CG4 cells. These results suggest that PTP! negatively regulates the phosphorylation of ERK1/2 and Akt in CG4 cells. Moreover, the expression of p27 was monitored in CG4 cells stimulated for 3h and 24h. As shown in Fig. 5.12D, the p27 expression level is similar in control siRNA and PTP! siRNA  !  122!  treated CG4 cells after stimulation for 3h. However, after 24h stimulation, the p27 protein level increased in control CG4 cells, but not in PTP!-knockdown CG4 cells (Fig. 5.12D). Therefore, in accord with findings in OPCs, PTP! negatively regulates the activities of Ras and Rho GTPases, Rac1, Cdc42 and Rho, and is required for p27 accumulation in OPCs.  5.2.10 Fyn is required for Rho inactivation and p27 accumulation in CG4 cells Since SFKs are substrates of PTP!, I investigated if SFKs are responsible for the inhibition of OPC growth. Treatment of CG4 cells with the SFK inhibitors SU6656 (2µm and 10µM) or PP2 (2µm and 10µM) resulted in a decrease in the number of cells, suggesting that SFKs are required for CG4 growth (data not shown). This result is similar to previous findings that SFKs are required for PDGF-induced OPC proliferation (Attali et al., 1997; Soliven et al., 2003). Fyn has been reported to promote growth arrest and differentiation of keratinocytes (Cabodi et al., 2000) and neuroblastoma cells (Berwanger et al., 2002), suggesting a role of Fyn in cell cycle regulation. It has been shown that fewer oligodendroglial cells (O4-positive pro-OLs) developed in Fyn-/- mixed glia cell cultures (Sperber and McMorris, 2001), suggesting that Fyn may be required for oligodendrocyte lineage commitment. Since Fyn is an important substrate of PTP! and I have shown that Fyn activity decreased in PTP!-/- OPCs (Wang et al., 2009), I next determined if Fyn is involved in PTP!-mediated signaling that inhibits OPC proliferation and promotes cell cycle exit. I first confirmed that Fyn activity decreased in PTP!-knockdown CG4 cells stimulated for 3h and 24h (Fig. 5.13A). Since Fyn  !  123!  phosphorylates p190RhoGAP (Wolf et al., 2001) and p250RhoGAP (Taniguchi et al., 2003), leading to Rho inactivation, I determined if ablation of Fyn results in hyperactivation of Rho. As shown in Fig. 5.13B, Rho activity increased in both PTP!knockdown and Fyn-knockdown CG4 cells, suggesting that PTP! may act through Fyn to inactivate Rho. Fyn and Rho have been implicated in regulating p27 expression. Overexpression of Fyn results in increased expression of p27 (Cabodi et al., 2000). Inhibition of Rho, by either lovastatin or C3 exoenzyme, increases the translational efficiency of p27 mRNA (Vidal et al., 2002). Therefore, it is possible that PTP! activates Fyn to inactivate Rho, leading to increased p27 protein expression. In keeping with this scenario, p27 protein levels were similar in control cells and cells in which PTP! and Fyn expression was silenced at the earlier time of 3h stimulation, but at 24h stimulation the p27 level more than doubled in control cells, but did not significantly increase in PTP!-knockdown or Fyn-knockdown CG4 cells. Moreover, the p27 level was significantly lower in PTP!-knockdown and in Fyn-knockdown CG4 cells compared to control cells at 24h stimulation (Fig. 5.13C). These results suggest that PTP! may act through Fyn to promote Rho inactivation and p27 accumulation.  5.3  Discussion Uncovering the molecular control of progenitor self-renewal and differentiation is  an important step in understanding tissue homeostasis and repair as well as tumor development. Adult OPCs can self-renew, differentiate and remyelinate damaged regions in the CNS (Chong and Chan). On the other hand, they may be the source of  !  124!  gliomas, since the marker phenotypes, morphologies, and migratory properties of cells in gliomas strongly resemble glial progenitors in many ways (Canoll and Goldman, 2008). In this study, I have investigated the effects of PTP! ablation in mouse OPC proliferation, cell cycle exit and survival. PTP!-null mice display a delayed maturation of OPCs into myelin-forming cells. In vitro studies using neurosphere-derived oligospheres showed that the growth rates of PTP!-null OPCs in both suspension (grown as oligospheres) and adherent (grown on poly-DL-ornithine-coated dishes) cultures are increased compared to WT OPCs. In addition, dissociated PTP!-null OPCs form new spheres in response to lower concentrations of PDGF and bFGF compared to WT OPCs, suggesting that PTP!-null OPCs are hypersensitive to PDGF/bFGF. However, it is difficult to dissect which growth factor signaling pathways are affected by PTP!, since culturing OPCs in medium without either growth factor results in cell death or spontaneous differentiation, increasing the complexity of the experimental conditions. Increased proliferation of PTP!-null OPCs was confirmed by cell cycle analysis, BrdU incorporation and Ki-67 positivity, indicating that more cells enter S phase (higher % of BrdU-positive cells) and fewer cells exit from the cell cycle (lower % of Ki-67-negative cells) in the PTP!-null population. The difference in Ki-67 expression between WT and PTP!-null OPCs is more pronounced after culture in the absence of PDGF and bFGF for 24 hours, suggesting that PTP!-null OPCs are more resistant to PDGF/bFGF withdrawal-induced cell cycle exit. I also found that PTP!null OPCs are more resistant to supplement withdrawal-induced apoptosis in response to PDGF/bFGF, as determined by the presence of cleaved caspase-3.  !  125!  Protein expression of PTP! and Fyn is much lower in neural stem cells than in OPCs and PTP! ablation does not affect proliferation of neural stem/progenitor cells. These results suggest that PTP! does not have a significant role in regulating the proliferation of neural stem/progenitor cells. This may be due to several reasons. First, the protein expression of PTP! in neural stem/progenitor cells may not reach a level sufficient to affect proliferation. Second, some downstream effectors of PTP! expressed in OPCs may not be expressed in neural stem/progenitor cells. Third, in contrast to the culture medium of OPCs which contains PDGF and bFGF, the growth factors bFGF and EGF are provided in the culture medium of neural stem/progenitor cells. PTP! may not have a significant role in bFGF and EGF-induced proliferation of neural stem/progenitor cells. Therefore, a more detailed examination is required to clarify the basis of the differential involvement of PTP! in the proliferation of OPCs and not of neural stem/progenitor cells. According to my findings and those of others, PTP! expression is absent in pluripotent stem cells (den Hertog et al., 1993), low in neural stem/progenitor cells (Section 5.2.4, and den Hertog et al., 1993), and increases during differentiation; and PTP! is required for the differentiation of neurons (den Hertog et al., 1993) and oligodendrocytes (section 4.2.1 and 4.2.4). Ablation of PTP! results in enhanced proliferation and cell cycle progression of OPCs (section 5.2.2 and 5.2.3). Moreover, the PTP! substrate Fyn promotes growth arrest and differentiation of keratinocytes (Cabodi et al., 2000) and neuroblastoma cells (Berwanger et al., 2002). These results suggest that PTP!-Fyn signaling may serve as a negative regulator of cell cycle progression to facilitate cell fate commitment, in addition to its role in cytoskeletal rearrangement.  !  126!  Ras proteins are small guanine nucleotide-binding proteins that control a variety of cellular processes including proliferation, survival, differentiation and migration (Karnoub and Weinberg, 2008). Ras has been shown to play an important role in oligodendrocyte development. Ras activation in OPCs in response to bFGF has been reported. Moreover, lack of neurofibromin, a negative regulator of Ras, resultes in the abnormal accumulation of OPCs (Bennett et al., 2003). Receptor-mediated activation of Ras signaling is required for the maintenance of glioblastomas, and Ras inhibition by farnesylthiosalicylic acid (FTS) or dominant-negative Ras downregulates ERK and Akt signaling and promotes apoptosis of these cancer cells (Blum et al., 2006). My findings identify PTP! as a negative regulator of Ras in OPCs in response to PDGF/bFGF, indicating that the lack of PTP! facilitates growth factor-mediated Ras activation and downstream events. However, the precise molecular mechanism of this PTP!-mediated negative regulation needs to be further investigated. Rho family small GTPases regulate many important processes, including the organization of the actin cytoskeleton, gene transcription, cell cycle progression, and membrane trafficking. Their activity is regulated by signals originating from surface receptors (Kjoller and Hall, 1999). Rho, Rac1 and Cdc42 promote entry into G1 and progression to S phase when expressed in quiescent fibroblasts. Unlike Ras, none of the Rho GTPases activate the MAPK cascade; instead, Rac1 and Cdc42, but not Rho, can stimulate a distinct MAP kinase, the c-Jun kinase JNK/SAPK (Jun NH2-terminal kinase or stress-activated protein kinase) (Olson et al., 1995). Rho is required for the expression of p21 and p27, inhibitors of G1 cyclin/Cdk, whereas Rac1 and Rho promote transcription and translation of cyclin D (Etienne-Manneville and Hall, 2002;  !  127!  Hu et al., 1999; Vidal et al., 2002). In addition, all three of these Rho GTPases have been implicated in Ras-mediated transformation, suggesting that they act downstream of Ras in the control of cell proliferation (Khosravi-Far et al., 1995; Prendergast et al., 1995; Qiu et al., 1997; Qiu et al., 1995). Rho negatively regulates process formation in oligodendrocytes (Liang et al., 2004; Wolf et al., 2001) and has been implicated in PDGF-induced proliferation of glioma cells (Wolf et al., 2003; Zohrabian et al., 2009). Rac1 and Cdc42 play important roles in cytoskeleton rearrangement and are required for process outgrowth of oligodendrocytes and myelination (Liang et al., 2004; Thurnherr et al., 2006a). However, the roles of Rho, Rac1 and Cdc42 in the proliferation of OPCs remain unknown. Here I showed that PTP! is a negative regulator of Rho, Rac1 and Cdc42 in OPCs in response to PDGF/bFGF. The increased activities of these GTPases correlate with the increased proliferation and decreased cell cycle exit in PTP!-null OPCs, suggesting that these GTPases may be downstream effectors of PDGF or bFGF in OPCs to promote proliferation and cell cycle progression. I previously showed that PTP! activates Rac1 and Cdc42 in oligodendrocytes during differentiation (Chapter 4, and Wang et al., 2009). Therefore, PTP! may have distinct roles in regulating Rho family small GTPases in OPCs and oligodendrocytes, depending on the extrinsic signals received and the intrinsic developmental stages of the cells. MAPK and Akt are important downstream effectors of Ras. Ras activates Akt through PI3K (phosphatidylinositol 3-kinase) to promote survival and activates ERK1/2 through Raf and MEK1/2 (mitogen-activated protein kinase kinase1/2) to promote gene expression (Karnoub and Weinberg, 2008). Increased or altered phosphorylation of  !  128!  ERK1/2 and Akt was not evident in proliferating PTP!-null primary mouse OPCs, suggesting that PTP! is not involved in ERK1/2 and Akt signaling in response to PDGF/bFGF. However, the phosphorylation of ERK1/2 and Akt was enhanced in proliferating PTP!-siRNA-treated CG4 cells compared to control CG4 cells. Since the culture conditions of mouse OPCs (serum-free medium) and CG4 cells (medium containing 1% FBS) are different, it is likely that distinct components of the culture medium (such as other growth factors present in FBS), induce signaling to downstream effectors, such as ERK1/2 and Akt in a manner that is regulated by PTP!. Therefore, the role of PTP! in other growth factor-mediated signaling pathways needs to be further investigated. Since ERK1/2 and Akt are also downstream effectors of the insulin receptor and the concentrations of insulin are different in the culture medium for primary OPCs (25 ng/ml) and CG4 cells (5 ng/ml), it is possible that the phosphorylation of ERK1/2 and Akt that was detected in primary OPCs largely results from insulin receptor signaling, such that the phosphorylation of ERK1/2 and Akt induced by PDGF and bFGF receptor signaling is masked. Another possibility is that the primary OPCs were not starved prior to stimulation, and were cultured in proliferation medium for 2 days, resulting in non-obvious differences in the phosphorylation of ERK1/2 and Akt or the return of this phosphorylation to basal levels after this extended time. This is supported by the observation that the phosphorylation of ERK1/2 and Akt decreased in both control and PTP!-siRNA-treated CG4 cells after stimulation for 24h and the absence of a difference between control and PTP!-siRNAtreated CG4 cells. Therefore, the role of PTP! in regulating MAPK signaling in primary OPCs needs to be further investigated. It is also possible that there are  !  129!  compensatory effects in cells with a stable, long-term lack of PTP! (PTP!-null OPCs) that do not occur in transient, short-term knockdown (PTP!-siRNA-treated CG4 cells) conditions. Finally, intrinsic differences between primary mouse OPCs and rat-derived CG4 cells may underlie the distinct activation responses of ERK1/2 and Akt. The CDK inhibitor p27 has been shown to be crucial in OPC differentiation due to its function in promoting growth arrest (Casaccia-Bonnefil et al., 1999; Tang et al., 1999; Tikoo et al., 1998). Ablation of p27 enhances proliferation and impairs differentiation of OPCs in vitro (Casaccia-Bonnefil et al., 1997). I found that ablation of PTP! or Fyn results in a decreased p27 protein level in OPCs, suggesting a role of PTP!-Fyn signaling in the regulation of p27 protein expression and growth arrest. The mechanisms through which Fyn regulates p27 expression remain unclear. I found that ablation of PTP! or Fyn results in increased Rho activity in CG4 cells, indicating that PTP!-Fyn signaling may downregulate Rho activity. Since RhoA has been shown to downregulate p27 protein levels (Hu et al., 1999; Vidal et al., 2002), PTP! may activate Fyn to inhibit Rho-mediated suppression of p27 expression, leading to accumulation of p27 in oligodendrocyte lineage cells and facilitating cell cycle arrest and differentiation of these cells.  5.4  Summary  !  In summary, my findings demonstrate an important role of PTP! in negatively  regulating growth factor-mediated proliferation, cell cycle entry and survival, and thereby self-renewal, in OPCs. PTP! is required for Fyn activation in proliferating OPCs (Fig. 4.11) and CG4 cells, and I have identified Ras, Rho GTPases (Rac1, Cdc42  !  130!  and Rho), MAPK, Akt, and p27 as targets of PTP! signaling in proliferating OPCs and/or CG4 cells. Furthermore, I propose that PTP! inhibits these processes at least partially by acting through Fyn to negatively regulate growth factor-mediated activation of Rho and suppression of p27 expression. Therefore, loss of PTP! may contribute to the progression of oligodendrocyte-related diseases, such as multiple sclerosis and glioma.  !  131!  !  132!  !  133!  !  134!  !  135!  !  136!  !  137!  !  138!  !  139!  !  140!  !  141!  !  !  142!  !  143!  !  !  144!  Chapter 6. GENERAL DISCUSSION  I have demonstrated that PTP! has dual roles in regulating oligodendrocyte differentiation: one is to inhibit proliferation and promote growth arrest by inhibiting Ras, Rac1, Cdc42 and Rho, and facilitating p27 accumulation; the other is, upon cessation of proliferation and induction of differentiation, to promote morphological changes by activating FAK, Rac1 and Cdc42 and inactivating Rho (Fig. 6.1). I also showed that Fyn is an important substrate and downstream effector of PTP! that mediates many of these effects, although there may be other Fyn-independent actions of PTP! that have not yet been discovered.  6.1  The role of PTP! in OPC self-renewal and cell fate commitment At the early stage of development, embryonic stem (ES) cells do not express  detectable PTP! (den Hertog et al., 1993). According to den Hertog et al. (den Hertog et al., 1993) and my study, as the ES cells differentiate into neural stem cells, they start to express PTP!. I also found that PTP! expression level increases upon differentiation into OPCs and is upregulated during OL differentiation. These results suggest the possibility that PTP! serves as a negative regulator of self-renewal to promote cell fate commitment and differentiation and therefore, the expression of PTP! is suppressed in cells having a requirement for self-renewal. Unlike some of the molecules that promote OL differentiation that are only expressed in differentiating OLs (such as PTP&), I and others found that PTP! and its substrate Fyn are expressed in both OPCs and differentiating OLs (Lu et al., 2005a;  !  145!  Ranjan and Hudson, 1996), suggesting that PTP! and Fyn may have other roles in oligodendrocyte development (such as proliferation, survival and migration). Indeed, I found that PTP! negatively regulates the PDGF/bFGF-mediated activation of Ras and downstream signaling pathways such as those involving ERK1/2 and Akt, and also the activation of the Rho family GTPases Rac1, Cdc42 and Rho (Fig. 6.1), to prevent hyperproliferation and ensure the proper timing of differentiation once the levels of growth factors decrease. Furthermore, my results indicate that Fyn may be involved in regulating these events. Fyn has been implicated in the regulation of growth arrest and differentiation in two other cell types, keratinocytes and neuroblastoma cells (Berwanger et al., 2002; Cabodi et al., 2000). Taken together, these findings indicate that PTP!-Fyn signaling may be crucial in promoting cell cycle arrest and cell fate commitment, as well as differentiation, in some specific cell types. One of the most important molecules that controls the decision to self-renew or differentiate in oligodendrocyte lineage cells is p27. During the G1 phase, cells receive extracellular signals that determine whether to proliferate or differentiate (Frame and Balmain, 2000). The coordinated activation of two CDKs, CDK4/6 and CDK2, is critical to facilitate the progression of cells from G1 to S phase (Resnitzky and Reed, 1995; Sherr and Roberts, 1995). The activities of these two CDKs are tightly controlled by both positive and negative regulators (Morgan, 1997). Kip proteins (p21, p27 and p57) are inhibitors of both CDKs (Sherr and Roberts, 1995). Differentiation is associated with a decrease in the G1 CDK activity (Kato and Sherr, 1993; Skapek et al., 1995), and the accumulation of CDK inhibitors (Matsuoka et al., 1995; Parker et al., 1995). Mice deficient for the CDK inhibitory function of p27 showed enhanced growth  !  146!  (Fero et al., 1996; Kiyokawa et al., 1996; Nakayama et al., 1996), and this growth correlates with an increase in proliferating cells in tissues undergoing postnatal maturation, suggesting that p27 affects the decision to proliferate or withdraw from the cell cycle in response to extracellular signals (Kiyokawa et al., 1996). Accumulation of p27 is observed in OPCs within 6 h of PDGF deprivation (Durand et al., 1997), and is observed in CG4 cells upon astrocytic differentiation (Tikoo et al., 1997). Ablation of p27 enhances proliferation and impairs differentiation of OPCs in vitro (CasacciaBonnefil et al., 1997). Overexpression of p27 in oligodendocyte lineage cells results in cell cycle arrest, even in the presence of mitogens, however, this is not sufficient to induce differentiation (Casaccia-Bonnefil et al., 1999; Tang et al., 1999; Tikoo et al., 1998; Tokumoto et al., 2002). In addition, activation of "-adrenergic and glutaminergic receptors by kainite in OPCs results in accumulation of p27 and G1 arrest (Ghiani et al., 1999). The accumulation of p27 in OPCs is controlled posttranscriptionally and is temperature-sensitive (Tokumoto et al., 2002). I found that ablation of PTP! results in a decreased p27 protein level in OPCs, suggesting a role of PTP! in the regulation of p27 protein expression and growth arrest. Indeed, I found that PTP! negatively regulates the activation of Ras, Rac1 and Rho and the phosphorylation of ERK1/2 and Akt, molecules that have all been reported to play roles in suppressing the p27 protein level (Bond et al., 2008; Coleman et al., 2004; Sa and Stacey, 2004). Therefore, it is possible that the reduced level of p27 protein in PTP!-null OPCs is due to hyperactivation of the signaling molecules described above. On the other hand, the PTP! substrate Fyn promotes growth arrest and causes upregulation of p27 in keratinocytes, but not in dermal fibroblasts (Cabodi et al., 2000).  !  147!  Moreover, active Fyn induces G1 arrest and differentiation of neuroblastoma cells (Berwanger et al., 2002), suggesting a role of Fyn in suppressing or terminating proliferation prior to differentiation. I found that ablation of Fyn in CG4 cells results in decreased p27 protein level. Therefore, it is possible that PTP! downregulates p27 protein expression through dephosphorylating and activating Fyn. The mechanisms by which Fyn regulates p27 expression remain unclear, but could involve the following two possibilities (Fig. 6.2). First, Fyn could downregulate Rho activity by phosphorylating RhoGAPs, thus stimulating the intrinsic GTPase activity of Rho, leading to Rho inactivation. RhoA has been shown to downregulate p27 protein levels posttranscriptionally (Vidal et al., 2002) and posttranslationally (Hu et al., 1999). I found that ablation of Fyn indeed results in increased Rho activity in CG4 cells. Therefore, Fyn may upregulate p27 protein level by phosphorylating RhoGAP to inhibit Rho-mediated suppression of p27 expression. Second, the RNA-binding protein QKI can be phosphorylated by Fyn, and this phosphorylation alters QKI binding with mRNA (Lu et al., 2005b). QKI-6 and -7 induce G0/G1 cell cycle arrest in OPCs and upregulate p27 expression by stabilizing its mRNA (Larocque et al., 2005). Therefore, Fyn may phosphorylate QKI to promote QKI-mediated protection of p27 mRNA. In both scenarios, PTP! is envisaged to function as an upstream activator of Fyn. Although p27 is required for cell cycle arrest, it is not sufficient to induce differentiation. Therefore, PTP!, acting in Fyn-dependent or -independent manners, likely regulates additional targets to negatively regulate self-renewal and promote cell fate commitment. For example, downregulation of Ras and Rho GTPase signaling pathways by PTP! may inhibit the expression of genes that are required for OPC self-  !  148!  renewal. In addition, PTP! may upregulate the expression of genes that are crucial for differentiation through activating Fyn. This is further discussed below in section 6.2.  6.2  The role of PTP! in OPC differentiation Differentiation of cells requires growth arrest, induction of specific gene  expression  and  morphological  changes.  Fyn  upregulates  MBP  expression  transcriptionally and posttranscriptionally, and I found that MBP expression decreased in differentiating PTP!-/- OLs and in the brains of PTP!-/- mice, suggesting that PTP! is involved in Fyn-mediated regulation of MBP expression. As described in section 1.3.5, Fyn regulates MBP gene expression via multiple mechanisms: stimulation of transcription factor binding to target genes; phosphorylation of QKI to regulate the stability of target mRNA; and phosphorylation of hnRNP A2 to enhance the translation of target mRNA. Although MBP is the only Fyn target gene in oligodendrocyte lineage cells that has been studied, there must be other Fyn target genes in these cells that have not been discovered that are important in cell fate commitment and differentiation. Therefore, PTP!-Fyn signaling in the context of regulating gene expression in differentiating oligodendrocytes warrants further investigation. In addition to gene expression, OPC differentiation requires extensive reorganization of the cytoskeleton. PTP! and Fyn are well known regulators of cytoskeletal rearrangement and are downstream effectors of integrin and IGF-1 signaling, both of which have been implicated in promoting OPC morphological differentiation (as described in sections 1.3.3.1 and 1.3.3.6). Therefore, PTP!-Fyn signaling may be crucial in mediating the cytoskeletal rearrangement of  !  149!  oligodendrocyte lineage cells to promote their differentiation. Indeed, I found that PTP! is required for Fyn dephosphorylation at its negative regulatory site, an event that is critical in Fyn activation, in both OPCs and differentiating OLs. Moreover, I found that an important substrate of Fyn in OLs, FAK, is also regulated by PTP!, as demonstrated by decreased phosphorylation at FAK Tyr576 in the absence of PTP!. Rac1 and Cdc42 are downstream effectors of FAK and are important in promoting process outgrowth of OLs. I also showed that the differentiation-induced activation of Rac1 and Cdc42 was significantly reduced in the absence of PTP!. This function of PTP! as a Rac1/Cdc42 activator is opposite to its function in proliferation, where PTP! acts to inhibit Rac1 and Cdc42. Since Rac1 and Cdc42 play important roles in a variety of cellular events, PTP! may regulate these two Rho GTPases differently in response to different extracellular signals and in different cell types. In interesting contrast to the differential roles of PTP! as a Rac/Cdc42 regulator, PTP! is required for Rho inactivation in both OPCs and differentiating OLs. The phosphorylation of p190RhoGAP, another Fyn substrate and negative regulator of Rho, was not affected in the absence of PTP!. This suggests that PTP! may be an upstream regulator of other molecules that regulate Rho activity, such as RhoGEFs, RhoGDIs and other RhoGAPs.  6.3  Future directions As described in sections 1.3.2, 1.3.3 and 1.3.5, SFKs (especially Fyn) are  involved in various receptor- and adhesion molecule-mediated oligodendrocyte processes such as proliferation, survival, migration, morphological differentiation and gene expression. However, which of these receptor/adhesion molecule-initiated  !  150!  signaling pathways require or utilize PTP! remains largely unknown. It is necessary to further investigate the role of PTP! in these specific signaling pathways and cellular functions to fully understand the biology of oligodendrocyte development and CNS myelination. Since CNS myelination is regulated by signaling between adhesion molecules on both axon and oligodendrocyte cell surfaces (Barres and Raff, 1999; Hu et al., 2004; Nave, 2010), and PTP! is also expressed in neurons and has been shown to interact with F3/contactin, axonal PTP! may also play roles in regulating oligodendrocyte development and CNS myelination. Co-culture of WT OPCs with WT and PTP!-/neurons can be performed to address this hypothesis. Oligodendrocytes have been implicated in many neurological disorders (as described in section 1.1.7). Therefore, it may be informative to investigate whether PTP! is involved in the development and/or progression of these diseases, for example, if PTP! expression or functions correlate with disease progression in patients. In addition, the role of PTP! in disease progression and its potential aberrant or loss of function in such disease states can be investigated using in vitro cell culture and in vivo mouse models of disease. Based on my studies described above, the loss or inhibition of PTP! may contribute to the progression of demyelinating diseases, such as leukodystrophies and multiple sclerosis, since PTP! is required for oligodendrocyte differentiation and myelination. Mouse models for studying remyelination (as described in section 1.1.8) can be used to address this hypothesis. Moreover, loss of PTP! may also contribute to the progression of glioma, especially pediatric high-grade glioma (HGG), since PTP! negatively regulates the self-renewing proliferation of OPCs driven  !  151!  by PDGF/bFGF and PDGFRA is the predominant target of amplification in pediatric HGG (Paugh et al.). A xenograft model or a PDGF-driven glioma model can be used to address this hypothesis.  !  152!  !  153!  !  154!  REFERENCES Ackerman, S. L., Kozak, L. P., Przyborski, S. A., Rund, L. A., Boyer, B. B., and Knowles, B. B. (1997). The mouse rostral cerebellar malformation gene encodes an UNC-5-like protein. 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