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Characterization of the role of CD34 in adult skeletal muscle regeneration Alfaro, Leslie Ann So 2011

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CHARACTERIZATION OF THE ROLE OF CD34 IN ADULT SKELETAL MUSCLE REGENERATION by Leslie Ann So Alfaro B.Sc., The University of British Columbia, 2005 A THESIS SUBMITTED IN PARTIAL FULFILLMENT OF THE REQUIREMENTS FOR THE DEGREE OF DOCTOR OF PHILOSOPHY in THE FACULTY OF GRADUATE STUDIES (Experimental Medicine) THE UNIVERSITY OF BRITISH COLUMBIA (Vancouver)  November 2011 © Leslie Ann So Alfaro, 2011  Abstract  Expression of the cell surface sialomucin CD34 is common to many adult stem cell types, including muscle satellite cells. However, no clear stem cell or regeneration-related phenotype has ever been reported in mice lacking CD34, and its function on these cells remains poorly understood. Here, we assess the functional role of CD34 on satellite cellmediated muscle regeneration. Using an optimized flow cytometry-based method to analyze myogenic progenitors, we show that CD34’s expression is tightly regulated early during the muscle regeneration process. Following this, we show that Cd34-/- mice, which have no obvious developmental phenotype, display a defect in muscle regeneration when challenged with either acute or chronic muscle injury, resulting in impaired myofibre hypertrophy. In vivo engraftment efficiency and BrdU proliferation assays comparing WT and Cd34-/myogenic progenitors attribute this defect to impaired myogenic progenitor cell function in Cd34-/- animals. Lastly, the culture of isolated single myofibres demonstrate that this overall muscle regenerative defect is caused by a delay in the activation of satellite cells lacking CD34 as well as impaired proliferation following activation. Consistent with the reported anti-adhesive function of CD34, Cd34-/- satellite cells also show decreased motility along their host myofibre. Altogether, our results identify a role for CD34 in the poorly understood early steps of satellite cell activation, and provide the first evidence that beyond being a stem cell marker, CD34 may play an important function in modulating satellite cell activity.  ii  Preface  Chapter 2 includes work published in Joe, A.W.B.J., Yi, L., Natarajan, A., Le Grand, F., So, L., Wang, J., Rudnicki, M.A., Rossi, F.M.V. 2010. Muscle injury activates resident fibro/adipogenic progenitors that facilitate myogenesis. Nature Cell Biology 12(2): 153-163. All authors contributed to designing and performing experiments, and also data interpretation. L.S. (Alfaro) contributed to the development, optimization, and verification of flow cytometry based analysis and isolation of MPC and FAP cells.  Chapters 2, 3, and 4 include data published in Alfaro, L.A., Dick, S.A., Siegel, A.L., Anonuevo, A.S., McNagny, K.M., Megeney, L.A., Cornelison, D.D., Rossi, F.M. 2011. CD34 promotes satellite cell motility and entry into proliferation to facilitate efficient skeletal muscle regeneration. Stem Cells Dec; 29(12): 2030-2041. L.A.S.A. designed and performed experiments, analyzed data, interpreted results, and wrote the manuscript. S.A.D. performed single fibre isolations over a timecourse following culture and immunostained with Pax7 and MyoD. A.L.S. and D.D.W.C. performed single fibre isolations, live videoimaging of satellite cell movement on fibres during culture, and pp38 expression on satellite cells by immunofluorescence. A.A. assisted in cross-sectional area measurements of regenerating fibres in WT and Cd34-/- animals. L.A.M. and K.M.M. designed experiments and interpreted results. F.M.V.R. designed experiments, interpreted results, and wrote the manuscript.  iii  Figures 4.10 and 4.13 were generated by our collaborator, Ashley L. Siegel from the University of Missouri.  Figure 5.3 was generated by our collaborators Jamie Haddon and Kelly M. McNagny, Biomedical Research Centre, University of British Columbia.  Data obtained to generate Figures 4.5, 4.7-4.9, 4.11-4.12 were collected by our collaborators, Ashley L. Siegel and Dr. D.D.W. Cornelison, from the University of Missouri.  Data obtained to generate Figures 4.1-4.4 and Table 4.1 were collected by our collaborator, Sarah A. Dick from the Ottawa Health Research Institute.  This work was approved by the University of British Columbia Animal Care Committee under Certificate Numbers A09-0216 and A09-0364.  iv  Table of Contents  Abstract .............................................................................................................................. ii Preface ............................................................................................................................... iii Table of Contents ................................................................................................................v List of Tables .......................................................................................................................x List of Figures.....................................................................................................................xi List of Abbreviations........................................................................................................xiv Acknowledgements..........................................................................................................xvii Dedication.........................................................................................................................xix Chapter 1: Introduction...................................................................................................1 1.1  Skeletal muscle .............................................................................................................................. 1  1.1.1  Structure and function............................................................................................................ 1  1.1.2  Adult skeletal muscle regeneration ....................................................................................... 2  1.1.2.1  Historical perspectives of muscle regeneration ............................................................ 3  1.1.2.2  Muscle regeneration: a complex process ...................................................................... 5  1.1.2.3  Major steps in muscle regeneration............................................................................... 5  1.1.3  Molecular regulation of muscle regeneration....................................................................... 8  1.1.3.1  Myogenic regulatory factors (MRFs)............................................................................ 8  1.1.3.2  Pax3/7............................................................................................................................ 10  1.1.3.2.1  Embryonic muscle development.......................................................................... 10  1.1.3.2.2  Adult skeletal muscle and regeneration............................................................... 11  1.1.3.3  Secreted factors ............................................................................................................ 13  1.1.3.3.1  Hepatocyte growth factor ..................................................................................... 14  1.1.3.3.2  Fibroblast growth factors ..................................................................................... 16  1.1.3.3.3  Insulin-like growth factors ................................................................................... 17  1.1.3.3.4  Transforming growth factor beta ......................................................................... 19  1.1.3.3.5  Interleukins 6 and 4 .............................................................................................. 21  1.1.3.4  Signaling pathways ...................................................................................................... 23  v  1.1.4  1.1.3.4.1  p38 MAPK ............................................................................................................ 23  1.1.3.4.2  Notch pathway ...................................................................................................... 26  1.1.3.4.3  Wnt pathway ......................................................................................................... 28  Major cell types involved in the myoregenerative process ............................................... 30  1.1.4.1  Satellite cells ................................................................................................................. 31  1.1.4.1.1  Origin of satellite cells ......................................................................................... 31  1.1.4.1.2  Identification of satellite cells.............................................................................. 31  1.1.4.1.3  Satellite cells as myogenic progenitors ............................................................... 33  1.1.4.1.4  Isolation of satellite cells...................................................................................... 34  1.1.4.2  Inflammatory cells........................................................................................................ 36  1.1.4.3  Fibro-adipogenic progenitors ...................................................................................... 39  1.1.5  Summary of muscle regeneration ....................................................................................... 42  1.1.5.1  Muscle homeostasis...................................................................................................... 42  1.1.5.2  Myotrauma.................................................................................................................... 42  1.1.5.3  Myoblast proliferation and migration ......................................................................... 43  1.1.5.4  Fusion and differentiation............................................................................................ 44  1.1.6  Models of skeletal muscle regeneration ............................................................................. 46  1.1.6.1  In vivo models............................................................................................................... 46  1.1.6.1.1  Acute damage........................................................................................................ 46  1.1.6.1.2  Chronic damage .................................................................................................... 48  1.1.6.2  In vitro models.............................................................................................................. 49  1.1.6.2.1 1.2  Single fibre cultures.............................................................................................. 50  CD34............................................................................................................................................. 50  1.2.1  CD34: a stem cell marker .................................................................................................... 51  1.2.2  CD34 and CD34-related family members: structure and expression ............................... 51  1.2.3  Proposed functions of CD34 ............................................................................................... 55  1.2.3.1  CD34 marks cells at an activated state........................................................................ 55  1.2.3.2  CD34 promotes proliferation and inhibits differentiation ......................................... 57  1.2.3.3  CD34 as a pro- or anti- adhesion molecule to facilitate migration ........................... 58  1.2.3.4  CD34 and signaling...................................................................................................... 60  1.3  CD34 and muscle regeneration................................................................................................... 61  1.4  Hypothesis and specific objectives............................................................................................. 63  1.4.1  CD34 maintains satellite cell quiescence ........................................................................... 63  vi  1.4.2  CD34 enhances myogenic progenitor cell progression through activation and  proliferation ....................................................................................................................................... 64 1.4.3  CD34 promotes satellite cell motility ................................................................................. 64  1.4.4  CD34 prevents premature myoblast differentiation........................................................... 65  Chapter 2: Characterization of CD34 expression on myogenic progenitor cells during adult muscle regeneration.................................................................................................66 2.1  Introduction and rationale ........................................................................................................... 66  2.2  CD34 is an excellent marker for identification and isolation of murine MPCs from adult  skeletal muscle....................................................................................................................................... 68 2.3  Alpha7 integrin as an alternative marker to CD34 for MPC isolation during regeneration ... 73  2.4  CD34 isoform and surface protein expression is regulated on MPCs during in vivo muscle  regeneration ........................................................................................................................................... 77 2.5  CD34 surface expression is regulated on satellite cells during in vitro myogenesis............... 81  2.6  Summary ...................................................................................................................................... 83  2.7  Materials and methods................................................................................................................. 83  2.7.1  Mice ..................................................................................................................................... 83  2.7.2  Single fibre isolation and culture ........................................................................................ 84  2.7.3  Preparation of skeletal muscle tissue for flow cytometry/FACS..................................... 84  2.7.4  Isolation of MPCs and analysis using flow cytometry ..................................................... 85  2.7.5  Acute damage with NTX.................................................................................................... 85  2.7.6  Limiting-dilution assays ..................................................................................................... 86  2.7.7  Intramuscular transplant of sorted MPCs .......................................................................... 86  2.7.8  Immunofluorescent staining and analysis by microscopy................................................ 87  2.7.9  Quantitative real-time and RT-PCR primers..................................................................... 87  2.7.10  Statistical analysis............................................................................................................. 87  Chapter 3: CD34 is necessary for proper myogenic progenitor cell function during adult myogenesis. ..............................................................................................................88 3.1  Introduction and rationale ........................................................................................................... 88  3.2  CD34 is necessary for efficient muscle regeneration in response to both acute and chronic  damage ................................................................................................................................................... 89 3.3  Impaired muscle regeneration observed in Cd34-/- can be attributed to a specific defect in  their MPCs ............................................................................................................................................. 96 3.4  CD34 promotes in vivo MPC proliferation .............................................................................. 102  vii  3.5  CD34 is dispensable for in vitro myogenic differentiation..................................................... 106  3.6  Summary .................................................................................................................................... 108  3.7  Materials and methods............................................................................................................... 108  3.7.1  Mice ................................................................................................................................... 108  3.7.2  Acute damage with NTX................................................................................................... 109  3.7.3  Cross-sectional area measurements ................................................................................. 110  3.7.4  Preparation of skeletal muscle tissue for flow cytometry/FACS................................... 110  3.7.5  Isolation of MPCs and analysis using flow cytometry .................................................... 111  3.7.6  Cytospin............................................................................................................................. 111  3.7.7  Transplantation and engraftment ..................................................................................... 111  3.7.8  BM transplantation and NTX injection ........................................................................... 112  3.7.9  BrdU analysis .................................................................................................................... 112  3.7.10  MPC culture: growth and differentiation ...................................................................... 113  3.7.11  Fusion index calculations ............................................................................................... 113  3.7.12  Statistical analysis........................................................................................................... 114  Chapter 4: CD34 is necessary for progression of satellite cells through the myogenic program, specifically in activation, migration, and proliferation. ................................115 4.1  Introduction and rationale ......................................................................................................... 115  4.2  CD34 is dispensable for maintenance of satellite cell numbers during homeostasis ............ 116  4.3  CD34 is necessary for satellite cells to progress through the myogenic program on cultured  single fibres.......................................................................................................................................... 118 4.4  Satellite cells lacking CD34 are delayed in cell cycle entry and overall proliferation ......... 122  4.5  CD34 is necessary for efficient satellite cell migration .......................................................... 125  4.6  Summary .................................................................................................................................... 132  4.7  Materials and methods............................................................................................................... 132  4.7.1  Mice ................................................................................................................................... 132  4.7.2  Single fibre isolation and culture ..................................................................................... 133  4.7.3  3D video time-lapse imaging ........................................................................................... 133  4.7.4  p38 MAPK detection by Western blot and immunofluorescence .................................. 133  4.7.5  Immunofluorescence......................................................................................................... 134  4.7.6  Freeze damage injury........................................................................................................ 134  4.7.7  Statistical analysis............................................................................................................. 134  Chapter 5: Conclusion .................................................................................................135 viii  5.1  Data summary ............................................................................................................................ 135  5.2  General discussion ..................................................................................................................... 137  5.2.1  CD34 mediates satellite cell activation and subsequent myoblast proliferation during  myogenic progression..................................................................................................................... 138 5.2.2  CD34 facilitates migration of myogenic progenitors ..................................................... 140  5.2.3  CD34 and signal transduction .......................................................................................... 142  5.3  Future directions ........................................................................................................................ 144  5.3.1  Contribution of other cell types ....................................................................................... 144  5.3.1.1  Hematopoietic subsets............................................................................................... 145  5.3.1.2  Fibro-adipogenic progenitors .................................................................................... 147  5.3.1.3  Endothelial cells ......................................................................................................... 148  5.3.2  Identification of CD34 binding partners........................................................................... 148  5.3.3  CD34 in human satellite cells and skeletal muscle......................................................... 149  5.4  Significance of this work........................................................................................................... 151  References........................................................................................................................152  ix  List of Tables  Table 2.1  Only CD34+ cells are capable of forming myogenic colonies. ...........................77  Table 4.1  Statistical analysis of the 2 myogenic progenitor populations on WT and Cd34-/-  myofibres during culture. ..................................................................................................120  x  List of Figures  Figure 1.1 Skeletal muscle structure. ..................................................................................2 Figure 1.2 Adult muscle regeneration and involvement of myogenic progenitors...................7 Figure 1.3 Skeletal muscle satellite cell. ...........................................................................32 Figure 1.4  Myogenic progression of cells from cultured single fibres. ..............................35  Figure 1.5  Distinct roles for FAPs during muscle regeneration and repair.........................41  Figure 1.6 Schematic representation of key events during adult muscle regeneration. .......45 Figure 1.7 Schematic representation of the CD34 family. .................................................53 Figure 2.1  CD34 is expressed on satellite cells found on isolated single myofibres. ..........69  Figure 2.2 Schematic diagram of skeletal muscle processing to isolate MPCs...................70 Figure 2.3  Gating strategy for prospective isolation MPCs by FACS. ...............................71  Figure 2.4 Sorted Sca1- CD34+ MPCs from adult skeletal muscle are myogenic. ..............72 Figure 2.5 Four distinct populations exist within skeletal muscle following Hoechst+, PI-, CD31-, CD45-, Sca1- gating when assessing CD34 and alpha7 integrin expression. .............74 Figure 2.6  Alpha7 integrin+ cells within adult skeletal muscle are myogenic. ...................75  Figure 2.7 Satellite cells are contained only within the CD34+ fraction. ............................76 Figure 2.8  CD34 surface expression on MPCs is regulated during in vivo muscle  regeneration. .......................................................................................................................78 Figure 2.9 Total CD34 mRNA expression is down-regulated during the first week of muscle regeneration following acute damage.......................................................................79 Figure 2.10 CD34 surface expression remains constant on FAPs during in vivo muscle regeneration. .......................................................................................................................80 Figure 2.11 CD34 isoform expression varies during in vivo muscle regeneration. .............81 Figure 2.12 CD34 expression is regulated on satellite cells during in vitro culture of single fibres...................................................................................................................................82 Figure 3.1  Comparable areas of damage at day 5 post-NTX injection between WT and  Cd34-/- animals. ...................................................................................................................90 Figure 3.2 Increased levels of necrosis in Cd34-/- animals at day 5 post-NTX damage. .....91 Figure 3.3  H&E analysis of WT vs Cd34-/- skeletal muscle at various timepoints post-NTX  damage................................................................................................................................92 xi  Figure 3.4  Cd34-/- regenerating myofibres fail to undergo hypertrophy following acute  NTX damage.......................................................................................................................93 Figure 3.5  H&E analysis of mdx vs. mdx/Cd34-/- skeletal muscle at vaious ages. ..............95  Figure 3.6 Schematic of experimental outline for cell transplantation. ..............................96 Figure 3.7  Comparison of WT LacZ+ donor cell engraftment with WT/GFP+ or Cd34-/-  /GFP+ MPCs. ......................................................................................................................97 Figure 3.8  Defective engraftment of Cd34-/- MPCs. ..........................................................98  Figure 3.9  Recipient genotype does not affect engraftment efficiency of transplanted  MPCs. .................................................................................................................................99 Figure 3.10 Comparable satellite cell purity from freshly sorted WT and Cd34-/- MPCs. 100 Figure 3.11 The lack of CD34 on BM-derived cells does not significantly affect muscle regeneration. .....................................................................................................................101 Figure 3.12 Representative FACS plots showing distinct BrdU+ MPC population post-NTX damage..............................................................................................................................103 Figure 3.13 Inefficient proliferation of Cd34-/- MPCs during skeletal muscle..................104 Figure 3.14 CD34 does not affect FAP proliferation during skeletal muscle regeneration. .105 Figure 3.15 Little to no cell death observed in WT and Cd34-/- muscle 3 days following NTX damage..............................................................................................................................106 Figure 3.16 Cd34-/- MPCs can generate multinucleated myotubes and is dispensible for myogenic differentiation in vitro. ......................................................................................107 Figure 4.1 Pax7 readily identifies satellite cells in WT and Cd34-/- isolated myofibres.......117 Figure 4.2 Comparable satellite cell numbers between WT and Cd34-/- single fibres immediately after isolation. ...............................................................................................117 Figure 4.3 Immunofluorescent imaging of Pax7 and MyoD staining on single fibres identifies activated satellite cells and differentiation-committed myoblasts. ......................................118 Figure 4.4 Cd34-/- satellite cells have delayed activation and progression through the myogenic program. ...........................................................................................................119 Figure 4.5. WT and Cd34-/- satellite cells have delayed p38 MAPK activation during single fibre culture.......................................................................................................................120 Figure 4.6. Western blot analysis using sorted WT and Cd34-/- MPCs show decreased p38 MAPK activation in Cd34-/- MPCs ....................................................................................121 xii  Figure 4.7 Cd34-/- satellite cells do not expand efficiently on cultured single fibres. ..........122 Figure 4.8 Cd34-/- satellite cells divide significantly less than WT controls........................124 Figure 4.9 Cd34-/- satellite cells undergo cell division significantly later than WT controls. ..........................................................................................................................................124 Figure 4.10 Representative images showing tracking of WT and Cd34-/- satellite cell movement on cultured single fibres. ..................................................................................126 Figure 4.11 Cd34-/- satellite cells move significantly slower than WT controls. .................127 Figure 4.12 Cd34-/- satellite cells travel less overall distance compared to WT controls. ....127 Figure 4.13 Instantaneous velocity measurements of WT and Cd34-/- satellite cells on cultured fibres. ..................................................................................................................128 Figure 4.14 Increased necrotic area in Cd34-/- skeletal muscle following freeze damage. ...129 Figure 4.15 Cd34-/- satellite cells efficiently exit their niche. .............................................131 Figure 5.1 Comparable transwell migration of WT and Cd34-/- MPCs. ..............................143 Figure 5.2 WT BM transplantation to WT and Cd34-/- recipients results in no significant difference in regenerating myofibre sizes. .........................................................................145 Figure 5.3 Mast cells do not significantly contribute to muscle regeneration following acute damage..............................................................................................................................147 Figure 5.4 Human CD34 is expressed in MPCs. ................................................................150  xiii  List of Abbreviations  !-cat  Beta-catenin  bFGF  Basic fibroblast growth factor  bHLH  Basic helix-loop-helix  BM  Bone marrow  Cd34-/-  Cd34-null  CD34CT  CD34 full-length isoform  CD34FL  CD34 t runcated isoform  CDKI  Cyclin dependent kinase inhibitor  CME  Crushed muscle extract  CNFs  Centrally-nucleated fibres  CSA  Cross-sectional area  CTX  Cardiotoxin  DGC  Dystroglycan complex  DMD  Duchenne muscular dystrophy  ECM  Extra-cellular matrix  EDL  Extensor digitorum longus  FACS  Fluoresecence-activated cell sorting  FAP  Fibro-adipogenic progenitor  FGF  Fibroblast growth factor  FGFR  Fibroblast growth factor receptor  G-CSF  Granulocyte-colony stimulating factor  xiv  GDF-8  Growth and differentiation factor 8  GFP  Green fluorescent protein  hCD34  Human CD34  hCD34Tg  Human CD34 transgenic  H&E  Hematoxylin and eosin  HEV  High endothelial venule  HGF  Hepatocyte growth factor  HSC  Hematopoietic stem cell  HSPG  Heperain sulfate proteoglycan  IGF  Insulin-like growth factor  IGF-R  Insulin growth factor receptor  IL-4  Interleukin-4  IL-6  Interleukin-6  i.m.  Intramuscular  i.p.  Intraperitoneal  i.v.  Intravenous  kDa  kilo daltons  L-sel  L-selectin  MEF2  Myocyte enhancer factor-2  MPC  Myogenic progenitor cell  MRF  Myogenic regulator factor  MyHC  Myosin heavy chain  NTX  Notexin  xv  PDGF  Platelet-derived growth factor  PI  Propidium iodide  PKC  Protein kinase C  pp38  Phospho-p38 MAPK  qRT-PCR  quantitative real-time polymerase chain reaction  SDF-1  Stromal derived factor 1  SF  Scatter factor  TA  Tibialis anterior  TGF-!  Transforming growth factor-!  TGF-!R  Transforming growth factor-! receptor  WT  Wild-type  xvi  Acknowledgements  I would like to thank all of the past and present Rossi Lab members who have made my time in the lab very memorable and from whom I’ve learned a tremendous amount from. I would like to particularly thank Dr. Stephane Corbel, Jeffrey Duenas, and Lin Yi who very patiently trained me when I first started in the lab. Thank you to Dr. Bernhard Lehnertz and Dr. Michael Long for the project critiques and guidance along the way – I couldn’t have asked for better fellow graduate students to look up to. I would also like to thank my supervisory committee members, Dr. Kelly McNagny and Dr. Michael Underhill, for all the support, project insights, and discussions. Lastly, a special thank you to my supervisor, Dr. Fabio Rossi, who was gave me support when I needed it, but most importantly, showed me how to grow into an independent person. It has truly been a very memorable experience working with you and your lab.  Thank you to everyone at The Biomedical Research Centre, and in particular Core and Administrative Staff Members Andy Johnson, Justin Wong, Taka Murakami, Krista Ranta, Helen Merkens, Michael Williams, Les Rollins, Nicole Voglmaier, George Gill, Eunice Yao, Joseph Liau, and Patricia Snider. The BRC would fall apart without you guys.  To the many friends, colleagues, and collaborators that I have met throughout the years and had the privilege and pleasure to work with – thank you for all the advice, encouragement, and being there every time I needed help.  xvii  On a more personal and special note, I would like to say thank you to Dr. Bahareh Ajami for being a wonderful friend and keeping me sane throughout the years. Thanks for always lending a listening ear, sharing your advice (“that’s why you’re the older graduate student”), and for just being there. I would never have been able to do this without you. Thank you also to Dr. Poh Tan, Jan Khoo, Katy Zhang, and Rehana Ishmail for listening to all my “stories” and for being wonderful friends that never let me forget that there is life outside the lab. Most importantly, a very big and heartfelt thank you to my family for being my strongest support network and cheering team – all of you have always been there for me and I can’t say thank you enough for that. Lastly, to my incredible husband, Dr. Gabriel Alfaro, thank you for never leaving my side and never letting me fall. It has been an amazing journey and I look forward to our next.  This research was made possible by grants from CIHR and the Stem Cell Network to the Dr. Fabio Rossi, as well as personal fellowships received from NSERC and MSFHR.  xviii  Dedication  This work is dedicated to all my family – never-ending gratitude for the support and inspiration.  ...but most especially to my husband, my best friend, and my favourite person, Gabriel Alfaro – thank you for everything…  xix  Chapter 1: Introduction  1.1  Skeletal muscle  Skeletal muscle works with nerves, connective tissue, and vasculature to provide the body with structure and the capability to perform voluntary, precise movements. Due to this indispensable role, many intricate histological, biochemical, and biological studies have been done to characterize skeletal muscle. These studies have culminated in the large body of knowledge we currently have regarding skeletal muscle.  1.1.1  Structure and function  Muscle cells, more often referred to as a muscle fibres, are unlike other cell types, since they are multinucleated, composed of hundreds of post-mitotic nuclei that lie within a sarcolemma, the plasma membrane that ensheathes each fibre. Specialized mononucleated cells called satellite cells are found outside the sarcolemma but beneath a basal lamina, hence the derivation of the name “satellite cell”. Satellite cells are also known as muscle stem cells, as they are the source of myogenic progenitors needed for regeneration. In addition, other cell types such as capillaries and fibro-adipogenic progenitors (FAPs) can be found just outside the basal lamina of myofibres. Overall, myofibres are connected by an endomysial lining and are grouped into muscle bundles called fascicles, which are surrounded by a perimysium and attached to the bone through tendons (Figure 1.1).  1  Figure 1.1  Skeletal muscle structure.  Schematic diagram of skeletal muscle and associated structures and cell types. Reprinted from Current Topics in Developmental Biology, volume 96, Paylor et al., “nonmyogenic cells in skeletal muscle regeneration”, pp. 139-165, ! 2011, with permission from Elsevier.  1.1.2  Adult skeletal muscle regeneration  Like many tissues in the body, damage to skeletal muscle can occur. Fortunately, under normal circumstances, skeletal muscle has the remarkable capacity to regenerate. However, under some circumstances, such as genetic disease, the normal regenerative mechanism is insufficient to fully restore muscle structure and function. Thus, significant work to better understand the cell types, factors, and molecular and cellular processes involved in muscle  2  regeneration are continually being done, making the muscle regeneration field an evergrowing area of research.  1.1.2.1  Historical perspectives of muscle regeneration  During embryonic and post-natal development into adulthood, skeletal muscle undergoes substantial growth (Buckingham & Relaix, 2007; White, Bierinx, Gnocchi, & Zammit, 2010). In contrast, adult skeletal muscle has long been thought to be a stable tissue that does not undergo much turnover. Despite evidence to the contrary, even the idea of regeneration following injury has been largely discounted for many years.  Two distinct hypotheses were initially proposed for how skeletal muscle regenerates itself. The first one postulates that mononucleated cells bud off from injured muscle fibres, generating new nuclei without mitosis. The second hypothesis, and now accepted dogma, proposes that mononuclear myogenic precursors are present in muscle fibres and survive the injury. These cells become myoblasts and proliferate mitotically upon damage and fuse to form myotubes that mature to new myofibres (Sloper & Partridge, 1980).  The initial discovery of the satellite cell by Mauro in 1961 (Mauro, 1961) provided support to the second hypothesis of myogenic precursors as he proposed these precursors to be derived, at least in part, from satellite cells. Moreover, work by Stockdale and Holtzer (Stockdale & Holtzer, 1961) conclusively showed that nuclei in myofibres do not undergo DNA synthesis, confirming that myonuclei do not replicate and are post-mitotic. Breakthrough publications 3  by Konigsberg (Konigsberg, 1963) and Yaffe (Yaffe, 1969) demonstrated that multinucleated myotubes are derived from the progeny of a single myogenic precursor in both the chick and mouse systems, respectively. These results, along with extensive morphological studies of growing or regenerating muscle (Sloper & Partridge, 1980), added further evidence that regeneration occurs through myogenic precursors. However, it is through autoradiography experiments that put forward the strongest evidence to support the myogenic precursor hypothesis, specifically the fact that satellite cells are the source of these progenitors. Work by Moss and Leblond first demonstrated that satellite cells proliferate and are responsible for the new addition of myonuclei during post-natal growth (Moss & Leblond, 1970). Consistent with this, Snow beautifully demonstrated that myonuclei addition during regeneration are derived from satellite cells (Snow, 1978).  These seminal works proved that the myogenic progenitor hypothesis was indeed the correct one and, overall, provided a solid foundation for the field of adult myogenesis. Over the last few decades, extensive research into the factors that regulate muscle regeneration (Charge & Rudnicki, 2004; Hawke & Garry, 2001) contributed greatly to our current understanding of this complex process. However, much still remains to be discovered, leaving this field as exciting as it was decades ago.  4  1.1.2.2  Muscle regeneration: a complex process  In normal adults, skeletal muscle has the remarkable capacity to completely restore its structure and function following the onset of damage. For example, athletes who frequently undergo training exercises constantly damage their muscle, yet fully recover and build strength and endurance. In order to do so, skeletal muscle contains resident satellite cells that ensure proper regeneration for the lifetime of an individual. However, upon severe physical trauma to muscle (Jarvinen, Jarvinen, Kaariainen, Kalimo, & Jarvinen, 2005) or in severe myodegenerative diseases, such as Duchenne muscular dystrophy (DMD) (Serrano, et al., 2011), the normal regenerative capacity of skeletal muscle is insufficient. Thus, it is clear that mechanisms that ensure efficient muscle regeneration need to be further elucidated and understood in order to develop effective therapeutic strategies for a wide range of muscle injuries.  1.1.2.3  Major steps in muscle regeneration  At rest, adult skeletal muscle is stable with very little myonuclei turnover (1-2% per week in rats) (Decary, et al., 1997; Schmalbruch & Lewis, 2000). Upon damage, muscle regeneration initiates. This cyclic process can be classified into several major steps (Figure 1.2): (1) Satellite cell quiescence. In unperturbed muscle, satellite cells lie within their niche, between the myofibre sarcolemma and basal lamina. At this point, satellite cells are quiescent and possess morphological features such as a high nuclei to cytoplasm ratio. (2) Satellite cell activation. During damage, satellite cells become activated. The nucleus and cytoplasm 5  enlarge and protein synthesis is upregulated. (3) Satellite cell migration. Following activation, satellite cells exit their sublaminar niche and migrate towards the site of damage to participate in the repair process. (4) Myoblast proliferation. Extensive proliferation of satellite cell-derived myoblasts occurs. Whether migration occurs before or after proliferation is still not well understood. Likely, both do occur as in vitro inhibitory studies and live videoimaging of satellite cells show proliferation can happen despite impaired migration. (5) Myoblast fusion and differentiation. Upon cessation of proliferation, myoblasts fuse to each other or to damaged fibres, generating terminally differentiated myofibres.  6  Figure 1.2 Adult muscle regeneration and involvement of myogenic progenitors. (A) At resting state, quiescent satellite cells are present at their niche, outside the myofibre sarcolemma. Upon myotrauma, satellite cells activate to generate myoblasts that fuse to the damaged fibre or to each other, generating centrally-nucleated fibres, a characteristic feature of regeneration. Under normal circumstances, muscle regeneration completes and fibres return to its resting state. (B) H/E staining to show histological features of muscle regeneration (cytoplasm, pink; nuclei, blue). At resting state, nuclei are peripherally located, due to the dense amount of protein structures needed for muscle function. Upon damage, inflammatory cells (not depicted here) invade to remove necrotic and damaged tissue. While new proteins are synthesized, myonuclei are centrally-located, a distinct feature of regeneration. Once regeneration is complete, muscle returns to its original resting state.  7  1.1.3  Molecular regulation of muscle regeneration  Muscle regeneration is a beautifully orchestrated process commencing with satellite cell activation and concluding with differentiation and myofibre formation. This process requires organization within each stage but also the smooth interplay and transition between steps. Despite the seemingly simple process described above, intricately regulated cell types and regulatory factors are involved that ensures muscle regeneration proceeds efficiently. Below, we summarize a few such cell types and key regulators. Although each group below is discussed separately, these regulators do not act in isolation.  1.1.3.1  Myogenic regulatory factors (MRFs)  A family of transcription factors known as myogenic regulatory factors (MRFs) regulate skeletal muscle development and differentiation. Four transcription factors belong in the family of MRFs: Myf5, MyoD, MRF4/Myf6, and Myogenin. These proteins share a homologous basic helix-loop-helix (bHLH) domain required for DNA binding and also for interaction with the E-protein family of transcription factors. MRF-E protein heterodimers then bind a specific CANNTG sequence, otherwise known as the E-box, on the promoters of many muscle-specific genes (Parker, Seale, & Rudnicki, 2003). Although E-proteins are not exclusively expressed in skeletal muscle, MRFs are muscle specific. The specificity of MRFs in combination with their timed expression allows MRFs to specifically direct myogenesis. Additionally, the myocyte enhancer factor-2 (MEF2) MADS box family of transcription  8  factors are also known activators of myogenic genes and act in concert with MRFs to regulate myogenic gene transcription (Black & Olson, 1998). However, MEF2 factors cannot activate muscle genes on their own. All four MRFs are considered master regulators of muscle in that they can each activate the complete myogenic program when introduced to non-myogenic cells.  The MRFs act during different stages of myogenic progression (Zammit, et al., 2004). Much of what is known about the roles of MRFs comes from studies performed using knock-out animals. The presence of multipotent, putative myogenic progenitors but complete lack of muscle in Myf5:MyoD double knock-out mice suggests an early role for Myf5 and MyoD in myogenic precursor specification (Rudnicki, et al., 1993). The severe lack of differentiated myofibres in Myogenin knock-out mice, despite the presence of myogenic progenitors, along with increased Myogenin expression and deficient myogenesis in MRF4 null mice suggests that the latter two MRFs, MRF4 and Myogenin, play a role in myogenic differentiation (Hasty, et al., 1993; Nabeshima, et al., 1993; Patapoutian, et al., 1995; Rawls, et al., 1995; Venuti, Morris, Vivian, Olson, & Klein, 1995). In general, Myf5 and MyoD are termed “early MRFs” regulating myogenic specification and MRF4 and Myogenin are the “late MRFs” regulating myogenic differentiation (Francetic & Li, 2011).  9  1.1.3.2  Pax3/7  Pax genes (Pax1 through -9) are a group of tissue specific transcription factors characterized by the presence of a paired domain that provides sequence specific binding to DNA. An octapeptide motif and homeobox DNA-binding domain may also be present. Based on the presence or absence of these domains, Pax genes are arranged into 4 subfamilies. Paralogues Pax3 and Pax7 belong in one such subfamily and are the only Pax genes expressed in skeletal muscle (Buckingham & Relaix, 2007).  1.1.3.2.1  Embryonic muscle development  During embryonic muscle development, Pax3 and Pax7 are expressed in somites, segments of paraxial mesoderm that form on either side of the neural tube that eventually give rise to skeletal muscle in the trunk and limbs. Pax3 is required for the formation of trunk muscles, regulating the lateral migration of myogenic precursors to the limb buds (Bober, Franz, Arnold, Gruss, & Tremblay, 1994; Daston, Lamar, Olivier, & Goulding, 1996; Tremblay, et al., 1998), and has been proposed to have a role in muscle cell survival (Borycki, Li, Jin, Emerson, & Epstein, 1999), a function that can be compensated for by Pax7 (Relaix, Rocancourt, Mansouri, & Buckingham, 2004). Moreover, Pax3 activates Myf5 (Bajard, et al., 2006) and MyoD (Relaix, et al., 2003; Tajbakhsh, Rocancourt, Cossu, & Buckingham, 1997), with Pax7 also being able to substitute for this function in the absence of Pax3. Conversely, Pax7 seems to be dispensable during embryonic myogenesis as Pax7 null mice appear normal at birth and survive until about 2-3 weeks of age (Mansouri, Stoykova, Torres, 10  & Gruss, 1996; Seale, et al., 2000). Notably, head myogenesis appear to be independent of Pax3 and Pax7.  1.1.3.2.2  Adult skeletal muscle and regeneration  A Pax3/7-expressing population of cells has been shown to constitute myogenic progenitors that eventually form adult skeletal muscle (Relaix, Rocancourt, Mansouri, & Buckingham, 2005). In adult skeletal muscle, Pax3 expression can be abundantly found in the diaphragm, forelimbs, and gracilis muscle in the hindlimbs. The remainder of the hindlimbs has notably lower amounts of Pax3. In the remainder of the body, ventral trunk and intercostal muscles lack Pax3 expression, whereas body wall muscles such as serratus caudalis dorsalis are positive. Head muscles do not express Pax3 (Relaix, et al., 2003). The isolation and characterization of Pax3-expressing primary cells demonstrate that Pax3 can be used to efficiently mark and isolate satellite cells from adults (Montarras, et al., 2005). Moreover, coexpression of Pax3 with Pax7 in the diaphragm confirms that Pax3 is present in satellite cells (Relaix, et al., 2006). A comprehensive study by Relaix and colleagues outlines a role for Pax3 and Pax7 in adult myogenic progenitors. Experiments with satellite cells infected with dominant negative Pax3 or Pax7 show a striking reduction in MyoD expression, while leaving Myf5 expression relatively unchanged, showing that both Pax3 and Pax7 specifically regulate MyoD expression in satellite cells (Relaix, et al., 2006). However, this overlapping role of Pax3 and Pax7 diverges in later stages of myogenic differentiation (Relaix, et al., 2006).  11  Like its Pax3 paralogue, Pax7 is expressed in skeletal muscle, more specifically in quiescent and activated satellite cells (Seale, et al., 2000), although Pax7 can be detected in murine adult brains (Day, Shefer, Richardson, Enikolopov, & Yablonka-Reuveni, 2007). Relaix et al. observed reduced proliferation and increased detection of apoptotic cells in Pax7 null cells compared to wild-type (WT) controls in vitro, hinting towards a pro-survival role for Pax7 in satellite cells (Relaix, et al., 2006). The introduction of a dominant-negative form of Pax7 in WT animals resulting in increased satellite cell death further confirms these results. Interestingly, a similar analysis with Pax3 indicates that Pax3 does not share this pro-survival role. Therefore, Pax7 appears to have a positive role in cell survival (Relaix, et al., 2006). Altogether, Pax3 and Pax7 seem to have distinct but also overlapping roles in adult muscle regeneration and satellite cells. Both Pax3 and Pax7 appear to regulate myogenesis through MyoD. However, only Pax7 appears to have protective, anti-apoptotic functions in satellite cells (Buckingham, 2007).  Because Pax7 null mice severely lack satellite cells in postnatal muscle (Mansouri, et al., 1996; Seale, et al., 2000), a functional role for Pax7 for satellite cell specification was initially suggested (Seale, et al., 2000). However, a later study showing the presence of satellite cells, although reduced, in the body wall and limb muscles of P11 Pax7 null mice contradicts this notion (Oustanina, Hause, & Braun, 2004). In their study, Oustanina and colleagues compared Pax7+/- with Pax7 null animals and saw no significant difference in satellite cell numbers or myofibre size. However, upon culture and differentiation of primary myogenic cells, Pax7 null cells show significantly decreased production of myogenic colonies, a difference not seen early in culture. Taken together, an alternative hypothesis is  12  proposed in which Pax7 is involved in continued proliferation and maintenance of satellite cells as opposed to their specification (Oustanina, et al., 2004). Interestingly, recent work by Lepper et al. has shed some light on whether Pax7 is indeed necessary for satellite cell selfrenewal and muscle regeneration (Lepper, Conway, & Fan, 2009). Unlike previous works by others, Lepper and colleagues examined Pax7 conditional knock-out animals. Surprisingly, their studies demonstrate that Pax7 is dispensable in adults and is only required for satellite cell function and muscle regeneration during the early stages of post-natal muscle growth (up to P21), when progenitor cells transition into quiescent satellite cells (Lepper, et al., 2009). Despite this, it still remains that Pax7+ satellite cells are the most significant contributors to, and are indispensable for, adult skeletal muscle regeneration (Lepper, Partridge, & Fan, 2011; Murphy, Lawson, Mathew, Hutcheson, & Kardon, 2011; Sambasivan, et al., 2011).  1.1.3.3  Secreted factors  Secreted factors play a significant role in regulating regeneration. The initial discovery that crushed muscle extract (CME), and not extract from intact muscle, can increase satellite cell proliferation (Bischoff, 1986a, 1986b; Chen & Quinn, 1992) fuelled the search for such factors that can potentially regulate the various stages of muscle regeneration. Below, we summarize a few key factors that have been proposed to have functional roles.  13  1.1.3.3.1  Hepatocyte growth factor  The exact identification of the mitogen within the CME was largely unknown for quite some time, although attempts to do so resulted in some characterizations. In his work, Bischoff described that this mitogen is heat and trypsin sensitive, carried a molecular weight of greater than 30 kDa (kilo daltons), and specifically increased proliferation of satellite cells, and not that of non-muscle cells such as fibroblasts (Bischoff, 1986b). Chen and Quinn confirmed these results and also performed combinatorial studies with other known mitogens, such as basic fibroblast growth factor (bFGF), insulin-like growth factor (IGF)-1, platelet-derived growth factor (PDGF), epidermal growth factor, and macrophage-colony stimulating factor, demonstrating that the active compound in CME is not one of these factors (Chen & Quinn, 1992). Ultimately, the active mitogen was identified as scatter factor (SF), otherwise known as hepatocyte growth factor (HGF). The scattering effect of CME on cultured MDCK cells, characteristic of SF/HGF, and the abolished satellite cell mitogen activity of CME upon the addition of anti-HGF antibodies confirms this (Tatsumi, Anderson, Nevoret, Halevy, & Allen, 1998).  HGF is a cytokine that can be synthesized and secreted by satellite cells to act in an autocrine fashion (Sheehan, Tatsumi, Temm-Grove, & Allen, 2000). HGF binds to its only receptor, cmet, a receptor tyrosine kinase. Upon ligand binding, c-met autophosphorylates to activate downstream signaling pathways, such as the MAPK pathway (Wozniak, Kong, Bock, Pilipowicz, & Anderson, 2005). C-met is a known satellite cell marker (Cornelison & Wold,  14  1997) and as satellite cells are the only cell types within muscle to express c-met, HGF can act specifically on satellite cells.  In resting muscle, HGF is not expressed; however, upon damage, HGF expression is upregulated early during regeneration and eventually down-regulated at later timepoints (Jennische, Ekberg, & Matejka, 1993). Injection of exogenous HGF to animals early during repair results in increased myogenic progenitor numbers but blocked differentiation, whereas later injections of HGF into damaged muscle have no affect (Miller, Thaloor, Matteson, & Pavlath, 2000; Tatsumi, et al., 1998). Moreover, Gal-Levi et al. show that ectopic HGF expression on primary satellite cells prevents the expression of MyoD, MRF4, Myogenin, MEF2, and MyHC (Gal-Levi, Leshem, Aoki, Nakamura, & Halevy, 1998). Notably, HGF has also been proposed to have chemotactic abilities mediated by the Ras/Ral pathway and possibly regulating satellite cell migration during adult regeneration (Bischoff, 1997; J. Suzuki, Yamazaki, Li, Kaziro, & Koide, 2000). Overall, these studies indicate that HGF primarily acts during the early stages of muscle regeneration (Miller, et al., 2000), promoting satellite cell activation and proliferation, while inhibiting differentiation.  A large body of knowledge regarding the role of HGF/c-met in embryonic myogenesis exists that describes the necessary role for HGF in the migration of myogenic progenitors from the somites to the limb bud during development (Bladt, Riethmacher, Isenmann, Aguzzi, & Birchmeier, 1995; Dietrich, et al., 1999; Maina, et al., 1996). Unfortunately, the detailed mechanism by which HGF regulates satellite cells during adult muscle regeneration is yet to  15  be defined. Currently, it is understood that HGF can promote activation, migration, and proliferation of satellite cells.  1.1.3.3.2  Fibroblast growth factors  The fibroblast growth factor (FGF) family is composed of nine members (FGF-1 through -9) and four FGF receptors (FGFR-1 through -4). FGFs also have a high affinity for heparin or heparin sulfate. FGF binds to both FGFR and heparin sulfate proteoglycans (HSPG) to form a signaling complex that enhances the mitogenic effects of FGF. Without HSPG binding, FGF-FGFR binding is decreased along with cell proliferation and myogenic differentiation is induced (Rapraeger, Krufka, & Olwin, 1991). Interestingly, transmembrane HSPGs such as syndecan-3 and syndecan-4 can be found on satellite cells and have been shown to be important in satellite cell activation, proliferation, and differentiation (Cornelison, Filla, Stanley, Rapraeger, & Olwin, 2001; Cornelison, et al., 2004).  All FGFs and FGFRs can be found in adult myofibres (Kastner, Elias, Rivera, & YablonkaReuveni, 2000). Interestingly, unlike other family members, FGF-6 was uniquely expressed by myofibres only and not connective tissue cells, suggesting a specific role in muscle (Kastner, et al., 2000). Unfortunately, contradictory results from Floss et al. (Floss, Arnold, & Braun, 1997) and Fiore et al. (Fiore, et al., 1997) in the characterization of FGF-6-null mice leaves us uncertain as to the in vivo functional role of FGF-6. However, direct injection of FGF-6 to damaged muscle accelerates regeneration, suggesting it has beneficial effects on the process (A. S. Armand, et al., 2003). FGF-2 neutralization in skeletal muscle at the time 16  of damage decreases the number and size of regenerating fibres (Lefaucheur & Sebille, 1995b) and injection of FGF-2 to dystrophic mdx animals improves regeneration (Lefaucheur & Sebille, 1995a), indicating a positive role for FGF-2 in regeneration. Similarly, increasing the expression of FGFR-1 results in increased satellite cell proliferation and delayed differentiation (Scata, Bernard, Fox, & Swain, 1999).  During in vitro studies of myogenic progression, Hannon et al. demonstrate that FGF-1, -2, and -6 are expressed in proliferating cells, while FGF-5 and -7 are expressed in differentiating myofibres; FGF-3, -4, and -8 were not detectable in either (Hannon, Kudla, McAvoy, Clase, & Olwin, 1996). This indicates that specific FGFs act during different stages of regeneration. In support of this, a detailed analysis of each FGF shows that only FGF-1, 2, and -6 could enhance satellite cell proliferation (Kastner, et al., 2000).  Overall, during muscle regeneration, FGFs and FGFRs stimulate satellite cell proliferation (Allen & Boxhorn, 1989; Lefaucheur & Sebille, 1995a; Olwin & Hauschka, 1986) and inhibit myogenic differentiation (Allen & Boxhorn, 1989; Linkhart, Clegg, & Hauschika, 1981).  1.1.3.3.3  Insulin-like growth factors  Insulin-like growth factors consist of two ligands (IGF-I and IGF-II) and two receptors (IGFR-1 and IGFR-2). In mice, significantly decreased body weights of IGF-1 null, IGF-II null, and of IGF-1:IGF-II knock-outs have been reported (Baker, Liu, Robertson, & 17  Efstratiadis, 1993; Liu, Baker, Perkins, Robertson, & Efstratiadis, 1993), which can partially be attributed to underdeveloped muscle tissue (Powell-Braxton, et al., 1993). Conversely, IGF-1 over-expression in mice results in increased body weight, increased muscle strength, and enlarged muscle fibres due to hypertrophy (Mathews, et al., 1988). Additionally, IGF-1 over-expression prevents age-related morphological changes in old mice, such atrophy and decreased muscle strength, (Barton-Davis, Shoturma, Musaro, Rosenthal, & Sweeney, 1998; Musaro, et al., 2001) and has even been shown to ameliorate dystrophic phenotypes in mdx mice (Barton, Morris, Musaro, Rosenthal, & Sweeney, 2002). Overall, these data suggest that IGFs are important skeletal muscle growth and regeneration.  During adult muscle regeneration, IGF-I and -II expression transiently increases, suggesting that IGFs play a role in this process (Bamman, et al., 2001; Hambrecht, et al., 2005; Levinovitz, Jennische, Oldfors, Edwall, & Norstedt, 1992). In vitro, IGF-II over-expression or addition of IGF-I or -II to myoblasts increases the rate of differentiation (Barton, et al., 2002; Barton-Davis, et al., 1998; Coleman, et al., 1995; Stewart, James, Fant, & Rotwein, 1996). Conversely, IGF-II inhibition disrupts myogenic differentiation (Florini, Ewton, & Roof, 1991; Florini, Magri, et al., 1991). In combination with results from IGF-I or -II null animals, these data suggest that IGFs are important in myogenic cell proliferation and differentiation. In addition, IGFs have also been implicated in promoting cell survival (Lawlor, et al., 2000; Lawlor & Rotwein, 2000a) and satellite cell migration (Schabort, van der Merwe, & Niesler, 2011).  18  Upon ligand binding to IGF receptors, autophosphorylation ensues and downstream signaling cascades activate. To mediate myoblast proliferation, cell survival, and differentiation, IGF/IGF-R binding activates downstream pathways such as PI3K-Akt and Raf-Mek/Erk1/2 (El-Shewy, Lee, Obeid, Jaffa, & Luttrell, 2007; Lawlor, et al., 2000; Lawlor & Rotwein, 2000a, 2000b). In summary, IGFs function in regeneration by promoting myoblast proliferation, surivival, and differentiation.  1.1.3.3.4  Transforming growth factor beta  Traditionally, the transforming growth factor beta (TGF!) superfamily is composed of three ligand isoforms, TGF!1, TGF!2, and TGF!3, and two receptor types, TGF!R-I and TGF!RII. These receptors form a heterodimer complex (Kingsley, 1994). Ligands first bind the TGF!R-II receptor, which, in turn, recruits and phosphorylates the TGF!R-I receptor. The now activated TGF!R-I phosphorylates and activates Smad proteins, which translocates to the nucleus to activate target genes such as p21, a cyclin-dependent kinase inhibitor (CDKI). In muscle, TGF!1-3 inhibits both myoblast proliferation and differentiation (Allen & Boxhorn, 1987, 1989; Olson, Sternberg, Hu, Spizz, & Wilcox, 1986; Schabort, et al., 2011). Moreover, recent work has shown that TGF!1 can attenuate satellite cell activation, despite the presence of HGF (Rathbone, et al., 2011).  In 1997, a new member of the TGF! superfamily was discovered and named growth and differentiation factor-8 (GDF-8), now more commonly known as myostatin (McPherron, Lawler, & Lee, 1997). Initial characterization of GDF-8/myostatin null animals showed 19  significantly increased overall body size and, more importantly, significantly increased muscle mass. Both hypertrophy (increased fibre size) and hyperplasia (increased fibre number) account for this increased muscle mass (McPherron, et al., 1997), suggesting that myostatin functions as a potent negative regulator of myogenesis. Cell cycle analysis demonstrates that myostatin inhibits myoblast proliferation, specifically by preventing G1 to S phase cell cycle progression through upregulation of p21 (McCroskery, Thomas, Maxwell, Sharma, & Kambadur, 2003; Thomas, et al., 2000).  In vivo, satellite cells express myostatin indicating a possible role for myostatin in regeneration. In support of this, myostatin null animals have accelerated regeneration and reduced fibrosis following notexin damage (McCroskery, et al., 2005). Decreased chemotaxis of myoblasts upon the addition of myostatin led the authors to propose that this accelerated regeneration can be at least partially attributed to increased myoblast migration (McCroskery, et al., 2005). Moreover, significantly higher satellite cell numbers and BrdUexpressing satellite cells can be found in myostatin null myofibres, suggesting that myostatin negatively regulates satellite cell activation and self-renewal (McCroskery, et al., 2003). Overall, the generally accepted function for myostatin in regeneration is that it is a negative regulator of myogenesis, inhibiting satellite cell activation, proliferation, and migration. Recently, Amthor and colleagues have challenged this dogma (Amthor, et al., 2009). Using myostatin null mice in combination with biological concentrations of exogenous myostatin on WT cells, these authors claim that the muscle hypertrophy observed with myostatin blockade is not actually dependant on myostatin regulation of myogenic progenitors, but instead relies on modulating protein synthesis within the fibres themselves (Amthor, et al.,  20  2009). Clearly, further investigations of myostatin’s functional role in adult regeneration are warranted.  1.1.3.3.5  Interleukins 6 and 4  Interleukin-6 (IL-6) is a cytokine secreted by T-cells and macrophages to modulate immune response. In addition, muscle has also been shown to be a source of IL-6 (Bartoccioni, Michaelis, & Hohlfeld, 1994; Hiscock, Chan, Bisucci, Darby, & Febbraio, 2004; Keller, et al., 2001), which is upregulated following muscle injury (Kami & Senba, 1998; Kurek, Nouri, Kannourakis, Murphy, & Austin, 1996). For many years, no reports have been made on a functional role for IL-6 in muscle regeneration. However, a report by Beaza-Raja and Munoz-Canoves describes that IL-6 expression increases during C2C12 myoblast differentiation (Baeza-Raja & Munoz-Canoves, 2004). Importantly, inhibition of IL-6 reduces C2C12 myogenic differentiation while its over-expression increases differentiation, suggesting a promyogenic role for IL-6 (Baeza-Raja & Munoz-Canoves, 2004). Serrano et al. then asked whether IL-6 is necessary for in vivo regeneration. Their study also shows increased IL-6 expression during muscle regeneration in WT mice, confirming previous in vitro results. Furthermore, the authors observed impaired muscle regeneration in IL-6 null mice that is attributed to defective satellite cell proliferation and myonuclear accretion (Serrano, Baeza-Raja, Perdiguero, Jardi, & Munoz-Canoves, 2008). An examination of possible signaling mechanisms involved in IL-6 regulation of proliferation indicates that activation of the JAK/STAT3 pathway is required for myoblast proliferation and migration. In support of this hypothesis, retroviral expression of STAT3 in WT myoblasts enhanced 21  proliferation and migration and STAT3 expression in IL-6 deficient myoblasts rescued their defective proliferation and migration (Serrano, et al., 2008). In humans, IL-6 has also been shown to induce satellite cell proliferation following damage (Toth, et al., 2011). Thus, for the first time, a defined promyogenic role for IL-6 in promoting satellite cell proliferation amd myogenic differentiation has been established.  Interleukin-4 (IL-4), like IL-6, has also been proposed to have a positive role in regulating muscle growth (Horsley, Jansen, Mills, & Pavlath, 2003). During myoblast growth and differentiation in vitro, IL-4 is present only on small, nascent myotubes after the induction of differentiation conditions and absent on mature myotubes (Horsley, et al., 2003). This regulated expression of IL-4 indicates a potential role during regeneration. Indeed, IL-4 deficient mice have defective muscle regeneration, with significantly smaller regenerating myofibres and decreased myonuclei. Interestingly, when myofibre sizes were assessed during various stages of regeneration, Horsley et al. observed that the size differences between WT and IL-4 deficient muscle only manifested during later stages (Horsley, et al., 2003). Furthermore, in vitro experiments with WT and IL-4 deficient myoblasts confirm a defect in myoblast differentiation in the absence of IL-4. More specifically, co-culture experiments between myotubes (WT or IL-4 deficient) and mononuclear cells (WT or IL-4 deficient) demonstrate that IL-4 acts on mononuclear cells to promote their recruitment and fusion to myotubes (Horsley, et al., 2003). However, the exact mechanism by which IL-4 induces myoblast recruitment and fusion are still under investigation, although activation of downstream signaling pathways such as PI3K and MAPK have been suggested, in addition to increasing expression of adhesion molecules (Horsley, et al., 2003).  22  1.1.3.4  Signaling pathways  In adult skeletal muscle regeneration, signaling pathways also have significant effects on satellite cells and their progression along the myogenic lineage. However, these pathways by no means act alone or solely on one population. Below, we summarize a few such regulators specifically of satellite cells and adult skeletal muscle regeneration.  1.1.3.4.1  p38 MAPK  Three major MAPK pathways exist in mammals: MAPK/ERK, SAPK/JNK, and p38 MAPK. MAPK signaling has been generally associated with regulating proliferation and differentiation of various tissues. A specific look at the role of p38 MAPK in adult myogenesis shows that it plays a functional role. Using the C2C12 myogenic cell line, Cuenda and Cohen show that p38 MAPK activity specifically increases during differentiation, while others, such as p42 MAPK/ERK, do not (Cuenda & Cohen, 1999). Inhibition of p38 MAPK activity with SB203580 prevented differentiation in a dosedependant manner, showing that p38 MAPK is necessary for this process. Moreover, induction of differentiation markers such as Myogenin, MyHC, and p21 was also significantly decreased with p38 MAPK inhibition, but not with p42 MAPK/ERK inhibition (Cuenda & Cohen, 1999). Complementary to this, exogenous activation of p38 MAPK results in precocious differentiation (Wu, et al., 2000).  23  Wu and colleagues also provide evidence that p38 MAPK is involved in the regulation of myogenic genes such as MyoD, MEF2A, and MEF2C (Wu, et al., 2000). Other works have further examined the mechanism by which p38 MAPK controls myogenic gene expression to modulate myoblast proliferation and differentiation. In muscle, p38 MAPK phosphorylates MEF2A and MEF2C (Wu, et al., 2000; Zetser, Gredinger, & Bengal, 1999), suggesting that p38 MAPK may contribute to the synergistic interaction of MRF (ie: MyoD) and MEF2 proteins. p38 MAPK also phosphoryles the E-protein, E47, significantly improving MyoD/E47 association to activate muscle specific gene transcription (Lluis, Ballestar, Suelves, Esteller, & Munoz-Canoves, 2005; Lluis, Perdiguero, Nebreda, & Munoz-Canoves, 2006). Altogether, these studies demonstrate that p38 MAPK is indeed essential for myogenic differentiation and can regulate this process at the level of gene transcription (Lluis, et al., 2006).  p38 MAPK has four family members, p38", p38!, p38#, and p38$. Phosphorylation of these members activates their kinase activity. p38" and p38# are the most abundant isoforms in skeletal muscle and are activated during myogenic differentiation (Perdiguero, Ruiz-Bonilla, Gresh, et al., 2007). Since SB203580 specifically inhibits p38" and p38! isoforms, the work by Cuenda and Cohen (Cuenda & Cohen, 1999) demonstrates the functional role of these isoforms in myogenic differentiation. Unfortunately, no such inhibitor exists for p38# and p38$ MAPKs. Thus, mouse knock-out models were used to examined the contribution of p38", p38!, p38#, and p38$ MAPKs to muscle regeneration (Perdiguero, Ruiz-Bonilla, Gresh, et al., 2007). Embryonic primary myoblasts deficient in p38", p38#, p38$, or p38% MAPKs were examined for their ability to differentiate in comparison to WT controls  24  (Perdiguero, Ruiz-Bonilla, Gresh, et al., 2007). Only p38" MAPK deficient myoblasts showed clear defects in myogenic progression, failing to form differentiated myotubes. Further analyses show that this is due to failed cell-cycle exit, thus resulting in continued cell proliferation despite differentiation conditions (Perdiguero, Ruiz-Bonilla, Serrano, & MunozCanoves, 2007). Increased activation of JNK was observed in p38" MAPK deficient myoblasts and inhibition of JNK rescued the proliferation defect in p38" MAPK cultures, suggesting that p38" MAPK normally regulates myogenesis by antagonizing JNK activation (Perdiguero, Ruiz-Bonilla, Gresh, et al., 2007; Perdiguero, Ruiz-Bonilla, Serrano, et al., 2007). Unfortunately, p38" MAPK null mice are embryonic/neonatal lethal. However, analysis of p38#, p38$, and p38% MAPK null mice and adult primary myoblasts show unaffected regeneration and differentiation, suggesting that p38" MAPK is the functionally significant isoform in skeletal muscle (Ruiz-Bonilla, et al., 2008). In addition to promoting cell cycle exit and differentiation, p38"/! MAPK has been proposed to be a molecular switch that regulates satellite cell activation from its quiescent state (Jones, et al., 2005). By examining satellite cells on cultured single fibres, Jones et al. showed that p38"/! MAPK activation coincides with satellite cell activation, defined as the upregulation of MyoD, and that inhibition of p38"/! MAPK by SB203580 substantially decreases satellite cell activation (Jones, et al., 2005). Collectively, these works demonstrate the important role of p38"/! MAPK in promoting efficient satellite cell activation and myogenic differentiation.  The work performed on embryos and adult mice deficient in p38", p38#, p38$, and p38% MAPK largely discounted the contribution of p38# and p38$ in muscle regeneration. However, the impaired muscle regeneration in p38$ MAPK null mice associated with  25  decreased satellite cell numbers suggests that p38# MAPK indeed plays a functional role (Gillespie, et al., 2009). In vitro experiments further demonstrate that, contrary to phenotypes seen with p38" MAPK deficient cells, cultured p38$ MAPK deficient myoblasts prematurely differentiate. Gillespie et al. further show that p38# MAPK can directly phosphorylate MyoD, thereby enhancing its occupancy of the Myogenin promoter, which intriguingly coincides with increased repressive H3K9 lysine methylation on the Myogenin promoter (Gillespie, et al., 2009). In essence, Gillespie et al. show, for the first time, that p38# MAPK is essential for preventing precocious differentiation through MyoD-mediated repression of Myogenin expression. Thus, the role of p38 MAPKs in muscle regeneration seem to be divided between the inverse, but complementary, roles of p38"/! MAPK in promoting cell cycle exit and differentiation with that of p38# MAPK in promoting cell proliferation and inhibiting premature differentiation (Lassar, 2009). However, the means by which the balance between p38" and p38# is delicately regulated is still unknown.  1.1.3.4.2  Notch pathway  Notch is a cell surface receptor that binds to its ligands, Delta-like ligand and Jagged. Upon binding, the intracellular portion of Notch is cleaved and translocates to the nucleus to mediate gene transcription. Numb, a Notch signaling pathway inhibitor, can prevent this process by interacting with the cleaved Notch intracellular domain to inhibit its nuclear translocation. In skeletal muscle, Notch activity can regulate myogenic progression. In C2C12 myogenic cells lines, Nofziger et al. showed that constitutively active Notch inhibits myogenic differentiation (Nofziger, Miyamoto, Lyons, & Weinmaster, 1999). In vivo, 26  Conboy et al. showed that upon muscle injury, Notch is expressed in activated satellite cells. Additionally, Notch activation increased satellite cell proliferation and inhibited differentiation (Conboy & Rando, 2002). Overall, this work provided initial insights into the role of Notch in satellite cells during adult regeneration.  A comprehensive compilation of work has been done to better understand the role of Notch signaling, with special regards to its contribution to age-related decline of satellite cell activity. In 2003, Conboy et al. showed that although satellite cell numbers are similar between young (2-3 months) and old (23-24 months) mice, satellite cell proliferation is significantly decreased, indicating impaired activation (Conboy, Conboy, Smythe, & Rando, 2003). During damage, satellite cell expression of Notch ligand, Delta, increases, whereas expression of Notch inhibitor, Numb, decreases (Conboy, et al., 2003; Conboy & Rando, 2002). This does not occur in aged animals, suggesting decreased overall Notch activity with age. Moreover, Notch inhibition in young mice results in decreased proliferation and increased myogenic differentiation, strikingly similar to phenotypes seen in aged groups; conversely, forced Notch activation in aged animals reversed impaired muscle regeneration. Therefore, the authors concluded that Notch is necessary for regeneration in young mice and sufficient to promote efficient repair in old mice (Conboy, et al., 2003).  In search for a mechanistic regulation of Notch activity, Conboy et al. later used heterochronic, parabiotic pairings between young and old mice (Conboy, et al., 2005), where partners establish shared circulation leading to blood exchange and chimerism. These experiments show that exposure of aged animals to a young environment rejuvenates aged  27  myogenic progenitors and results in efficient regeneration. Culture of satellite cells from young and old mice show that exposure of aged satellite cells to young serum increases Delta expression and Notch activity, rendering them functionally equivalent to young satellite cells. This serum-derived factor has yet to be identified (Conboy, et al., 2005). Interestingly, excessive TGF! expression has also been reported in old mice (Carlson, Hsu, & Conboy, 2008). Indeed, exposure of young satellite cells to old satellite cells results in increased TGF! production and “premature aging”. Notch and Smad3 antagonize each other, with Notch inhibiting TGF!-dependent activation of CDKIs by affecting the binding of Smads to CDKI promoters. Altogether, the authors propose that in aged animals, an imbalance between Notch and Smad exists, triggering expression of CDKIs old animals and preventing satellite cell proliferation (Carlson, et al., 2008).  Overall, this extensive research into the role of Notch in muscle regeneration, especially in terms of satellite cell decline in aged animals, demonstrates that Notch activation is important in satellite cell activation and proliferation in order to maintain proper regenerative potential (Luo, Renault, & Rando, 2005).  1.1.3.4.3  Wnt pathway  The canonical Wnt signaling pathway consists of a network of proteins involved in the regulation of many processes, including myogenic differentiation. In the absence of Wnt ligands, beta-catenin (!-cat) is phosphorylated by phospho-GSK3! and maintained in its APC/Axin destruction complex and targeted for proteolytic degradation. Upon Wnt ligand 28  binding to their receptor, Frizzled, !-cat is dephosphorylated and released from its destruction complex and translocates to the nucleus to activate gene transcription.  Wnt signaling has been shown to have an essential role in myogenic differentiation. In vitro, stable expression of Wnt or !-cat results in increased differentiation of P19 myogenic cells (Petropoulos & Skerjanc, 2002). In vivo, Wnt signaling is inactive in undamaged muscle, but is activated in myogenic progenitors upon injury (Brack, Conboy, Conboy, Shen, & Rando, 2008). Correspondingly, phospho-GSK3! expression, normally inhibiting Wnt activity, declines at the same time as Wnt upregulation. The timing of Wnt activation coincides with the switch of myoblasts from a proliferative state to that of differentiation, suggesting Wnt signaling to be important in this process. Indeed, Wnt activators accelerate myogenic progression in vitro and in vivo, resulting in premature differentiation. However, this is not associated with enhanced regeneration, as premature differentiation depletes myogenic progenitors.  Since Notch has been implicated as a regulator of satellite cell function, the authors asked whether there is possible cross-talk between Wnt and Notch signaling and examined Wnt signaling readouts in response to Notch inactivation. When Notch is inhibited, both Wnt and the inactive form of GSK3! are increased, suggesting that a balance between Notch and Wnt activation through GSK3! is necessary to mediate proper myogenic progression, specifically in the switch from proliferation to differentiation (Brack, et al., 2008). Overall, this study highlights a role for Wnt signaling in promoting myogenic progression.  29  Wnt signaling has also been implicated in the increased fibrosis observed in aged mice (Brack, et al., 2007). By parabiosing young and aged mice, Brack et al. show that serum components from aged mice are responsible for increased collagen deposition and decreased myogenic cell proliferation in young partners. These serum components from aged animals bind Frizzled, strongly implicating Wnt ligands as the active component. Conversely, Wnt inhibitors can suppress the fibrotic phenotype in aged animals. Thus, the authors propose a model in which myogenic cells convert to a fibrogenic phenotype through a Wnt signal activated pathway and contributes to the impaired regeneration and increased fibrosis in aged skeletal muscle (Brack, et al., 2007).  Therefore, Wnt signaling activity is not only important in promoting the switch of myogenic progenitors from proliferation to differentiation, but also plays a pivotal role in preventing age-related fibrosis.  1.1.4  Major cell types involved in the myoregenerative process  Upon muscle injury, numerous local and peripheral cells are involved that clean up the damage and repair the muscle, resulting in a state that is indistinguishable from that of the original state. Much effort has been put towards characterizing the cell types that facilitate this process. Despite this, new cell types and factors are still likely to be identified. Below, we discuss some major players involved in muscle repair.  30  1.1.4.1  1.1.4.1.1  Satellite cells  Origin of satellite cells  Historically, the developmental origin of satellite cells was largely debated between two distinct, but not necessarily mutually exclusive, hypotheses. The first hypothesis, supported by quail-chick chimeric experiments, proposes that satellite cells are derived from somites, which are the segmental derivatives of the paraxial mesoderm (O. Armand, Boutineau, Mauger, Pautou, & Kieny, 1983). The second hypothesis, arising from the discovery of skeletal muscle progenitors from the embryonic dorsal aorta, suggests that satellite cells are not exclusively derived from somites (De Angelis, et al., 1999). Settling this dispute, Gros et al., used GFP electroporation in chick somites in combination with video-microscopy to show that the dermomyotome (dorsal compartment of the somite) is the origin of myogenic progenitors and satellite cells (Gros, Manceau, Thome, & Marcelle, 2005), confirming the first hypothesis that satellite cells are derived from the somite.  1.1.4.1.2  Identification of satellite cells  Satellite cells, as their namesake, are characterized and identified based on their anatomical location between the myofibre’s sacrolemma and basal lamina. Although electron microscopy was traditionally used to identify these muscle progenitors, other methods that rely on satellite cell markers are now more commonly used. Examples of such markers  31  include Pax7, Myf5, c-met, m-cadherin, syndecan3/4, CXCR4, alpha7 integrin, and CD34 (Beauchamp, et al., 2000; Blanco-Bose, Yao, Kramer, & Blau, 2001; Cornelison, et al., 2001; Seale, et al., 2000; Sherwood, et al., 2004) (Figure 1.3).  Figure 1.3  Skeletal muscle satellite cell.  (A) Electron microcopic image of a satellite cell from rat sartorius muscle (22,000x magnification). The sarcolemma is marked as the intersection between the muscle (mp) and satellite cell plasma membrane (sp). ! Mauro, 1961. Originally published in the Journal of biophysical and biochemical cytology (Journal of Cell Biology). 9:493-495. (B) Immunofluorescent image of a satellite cell from a freshly isolated mouse EDL muscle fibre (scalebar, 10µm). The satellite cell is located under the basal lamina (laminin, blue) and identified with markers Pax7 (red) and CD34 (green).  In our studies, we define satellite cells as cells found in this distinct anatomical niche and marked by satellite cell specific markers; on the other hand, we define myogenic progenitor cells (MPCs) as a heterogenous population of activated satellite cells and committed myogenic progenitors. MPCs are located outside of the satellite cell niche, but still express satellite cell specific markers. Both satellite cells and MPCs are capable of undergoing the complete process of myogenic differentiation.  32  1.1.4.1.3  Satellite cells as myogenic progenitors  Mauro’s initial discovery of the satellite cell (Mauro, 1961) led to its extensive characterization. Early studies using autoradiography confirmed that satellite cells indeed contribute to adult skeletal muscle growth and, more importantly, regeneration. However, the exclusivity and significance of this contribution to repair was put in question upon the discovery that bone marrow (BM)-derived, circulating cells can also contribute to establishing regenerated myofibres and even restore dystrophin expression in mdx dystrophic mice (Ferrari, et al., 1998; Gussoni, et al., 1999; LaBarge & Blau, 2002). Later works further show BM-derived cells can contribute to muscle repair even at the single cell level (Corbel, et al., 2003) and that it is specifically the myelomonocytic fraction that contains myogenic precursors (Doyonnas, LaBarge, Sacco, Charlton, & Blau, 2004). Together, these discoveries made circulating cells a very attractive source for therapy that could repair muscle throughout the body.  However, it was not until the work of Sherwood et al. was published that this debate of endogenous myogenic precursors versus circulating BM cell contribution was put to rest (Sherwood, et al., 2004). This group isolated and intricately tested various proposed cells that contribute to muscle regeneration. Circulating BM hematopoietic stem cell (HSC)-derived cells and non-circulating endogenous muscle-derived cells were isolated by flow cytometry and compared for their in vitro and in vivo myogenic activity. Their results showed that although circulating cells do contribute to the generation of new muscle fibres upon injury, this contribution needs to be induced with damage or co-culture and is not significant when  33  compared to that of local myogenic progenitors (Sherwood, et al., 2004). In essence, satellite cells, the local myogenic progenitors, were “re-enthroned” as the most significant and primary cell type directly contributing to muscle regeneration (Partridge, 2004).  1.1.4.1.4  Isolation of satellite cells  Because of the tremendous capacity of satellite cells to generate skeletal muscle, there has always been a need to efficiently isolate these cells, both for experimental studies and also for possible cell-based treatments. More traditional methods have relied on culturing dissociated muscle from young animals and selecting for myogenic populations based on adherence to the culture plates under specific culture conditions (Rando & Blau, 1994; Richler & Yaffe, 1970). Unfortunately, this method may not always result in consistent populations among different groups, and, more importantly, do not always yield pure populations of myogenic cells.  Single fibres isolations are commonly used as it allows for the isolation of specific groups of muscles, such as the extensor digitorum longus (EDL) muscle, composed mainly of fasttwitch fibres, or the soleus muscle, composed mainly of slow-twitch fibres. Moreover, this is the only method thus far that allows for the isolation of satellite cells within their niche. The subsequent culture of these myofibres activates satellite cells, which then exit their niche and continue to progress through the myogenic program, from proliferation to differentiation and myotube formation (Collins & Zammit, 2009; Rosenblatt, Lunt, Parry, & Partridge, 1995) (Figure 1.4). A caveat in this process is that it does not fully exclude the presence of non34  satellite, fibre-associated cell types. Nevertheless, satellite cells are heavily enriched for in this system. Albeit an in vitro model, single fibre culture is extensively used as a model of myogenesis.  Figure 1.4  Myogenic progression of cells from cultured single fibres.  Brightfield imaging of cultured single fibre. (A) Upon culture of isolated single fibres, mononuclear satellite cells migrate out of their niche and adhere to the culture plate. (B) Subsequent removal of the isolated fibre leaves the surrounding cells undisturbed. (C) Myogenic cells proliferate and begin to fuse. (D) Differentiated myotubes are formed by two weeks. ! With kind permission from Springer Science + Business media: In Vitro Cellular & Developmental Biology – Animal, Culturing satellite cells from living single muscle fiber explants, 199, 1995, 773-779, Rosenblatt, et al., Figure 1.  35  Because of the availability of many satellite cell markers (some mentioned above) and reporter mice (ie: Pax7-YFP, Myf5/LacZ, etc), another commonly used system to purify and isolate satellite cells from adult tissues is through fluorescence-activated cell sorting (FACS). Here, muscle tissue is enzymatically dissociated and then labeled with antibodies to allow for negative exclusion or positive inclusion of specific cell types. A combination of markers eventually allow for the isolation of a specific myogenic population that is relatively pure and consistent among labs (Joe, et al., 2010; Kuang, Kuroda, Le Grand, & Rudnicki, 2007; Montarras, et al., 2005; Sacco, Doyonnas, Kraft, Vitorovic, & Blau, 2008; Uezumi, Fukada, Yamamoto, Takeda, & Tsuchida, 2010). Altogether, several methods of satellite cell purification are now well established, allowing researchers to study and characterize various stages of myogenesis.  1.1.4.2  Inflammatory cells  Upon muscle injury, damaged myofibres release factors that recruit local and circulating leukocytes to the site of damage. The first types of cells to arrive are neutrophils, accumulating as rapid as within the first 2 hours post-damage. Neutrophil numbers peak between 6-24 hours post-damage and then subsequently decline. The second types of cells to invade muscle are macrophages. Two subtypes of macrophages play a significant role in regeneration: pro- and anti-inflammatory macrophages. Initially, phagocytic, proinflammatory macrophages, otherwise known as M1 macrophages, arrive at damaged areas at around 24 hours post-injury and have continued presence that peak at about 2 days postinjury. Following this initial wave of M1 macrophages, non-phagocytic, anti-inflammatory, 36  M2 macrophages take over and peak at about 4 days post-injury (Arnold, et al., 2007; Chazaud, et al., 2009; Chazaud, et al., 2003; Tidball & Villalta, 2010). At 10 days postinjury, macrophage numbers begin to decline.  Despite the large body of work surrounding the role of inflammatory cells in adult muscle regeneration, the exact mechanisms by which these cells modulate muscle repair has only begun to be understood. Early studies characterizing the role of neutrophils in muscle regeneration initially yielded contradictory results as to whether or not neutrophils played any role in this process, with one group showing neutrophil blockage decreases membrane damage (Brickson, et al., 2003), while another group showed unaffected muscle regeneration in neutropenic mice (Lowe, Warren, Ingalls, Boorstein, & Armstrong, 1995). The current understanding of neutrophil function in muscle regeneration is that they do indeed play an important role in initiating the regenerative process (Butterfield, Best, & Merrick, 2006). Neutrophils can release myeloperoxidase, which oxidizes low-density lipoprotein thereby promoting binding to its receptor and macrophage marker CD68. This receptor-ligand binding activates phagocytosis and pro-inflammatory cyokine release by M1 macrophages (Ottnad, et al., 1995; Ramprasad, et al., 1995; Van Velzen, Da Silva, Gordon, & Van Berkel, 1997). Importantly, this serves as an example of a regulatory interface between neutrophils and macrophages during regeneration.  Macrophages also play an essential role in skeletal muscle regeneration. In vitro studies show that macrophages can release a factor that selectively increases the proliferation and differentiation of myogenic cells (Cantini & Carraro, 1995). In vivo studies with models of  37  reduced macrophage entry to muscle show delayed regeneration and delayed appearances of regenerating fibres (Shireman, et al., 2007; Summan, et al., 2006). Subsequent studies that completely ablated macrophage recruitment showed impaired muscle repair, demonstrating the necessary role macrophages have in this process (Arnold, et al., 2007). As mentioned above, two types of macrophages are proposed to participate in regeneration, pro- and antiinflammatory macrophages. Currently, it is thought that M1 pro-inflammatory macrophages first enter the site of damage during regeneration to perform phagocytic functions. M1 macrophages can be identified as CD45+, F4/80lo, Ly6C+ and CX3CR1lo cells and through the release of factors such as TNF", IL-6, and reactive oxygen species. M1 macrophages promote muscle regeneration by phagocytic removal cellular debris. After this initial wave, M2 anti-inflammatory macrophages appear. These cells can be identified as CD45+, F4/80hi, Ly6C-, and CX3CR1hi cells and through the release of anti-inflammatory cytokines such as IL-10, TGF-!, and PDGF. M2 macrophages suppress inflammation by deactivating M1 macrophages and mediate healing and repair (Villalta, Nguyen, Deng, Gotoh, & Tidball, 2009).  In vitro studies using conditioned media during the culture of myogenic cells show that medium from pro-inflammatory macrophages decrease myogenic fusion and differentiation, while increasing motility. Conversely, conditioned media from anti-inflammatory macrophages enhance myogenic fusion and differentiation, while decreasing motility (Chazaud, et al., 2009). These observations coincide well with the timing of M1 versus M2 appearance during certain stages of in vivo regeneration. M1 macrophages appear early during the repair process (days 1-2) and likely aids in preventing premature differentiation by  38  increasing myoblast motility. On the contrary, M2 macrophages appear later (days 4-10) and likely promote fusion and differentiation.  In summary, inflammatory cells play an important and functional role to mediate efficient (Butterfield, et al., 2006; Chazaud, et al., 2009; Tidball & Villalta, 2010)muscle regeneration in adults, with neutrophils initiating the inflammatory response, M1 macrophages promoting myoblast motility and proliferation, and M2 macrophages promoting myogenic differentiation.  1.1.4.3  Fibro-adipogenic progenitors  Like inflammatory cells, cell types exist that can modulate efficient muscle regeneration without directly contributing to muscle formation themselves. Recently, a population of mesenchymal progenitors was found in skeletal muscle. These cells, termed FAPs, can be isolated by FACS as CD45- CD31- alpha7- Sca1+ CD34+ and PDFGR"+ cells and were identified simultaneously by two independent groups (Joe, et al., 2010; Uezumi, et al., 2010). Together, their works describe a novel cell type that has both the ability to enhance muscle regeneration and the bipotentiality to differentiate into adipocytes and fibroblasts.  The authors propose that upon muscle injury, both FAPs and satellite cells activate and proliferate. The satellite cells directly contribute to regenerating muscle, while FAPs indirectly contribute to regeneration by promoting myogenic differentiation. During normal muscle repair and full recovery from the initial injury, satellite cells and FAPs are thought to 39  positively interact with each other. Through direct contact, satellite cell-derived myofibres inhibit the adipocyte or fibroblast differentiation of FAPs (Uezumi, et al., 2010). In turn, FAPs enhance differentiation of myogenic cells through an unknown secreted factor (Joe, et al., 2010). These interactions between muscle and FAPs are proposed to help ensure successful regeneration and restoration of normal muscle.  In aged, diseased, or chronic damage models, muscle regeneration becomes inefficient and eventually fails to restore normal muscle structure and fibrosis ensues. The origin of the cells responsible for fibrosis remains debated, with some works suggesting that satellite cells themselves undergo an alternative path of differentiation to generate fibrotic tissue instead of normal muscle (Brack, et al., 2007; Shefer, Wleklinski-Lee, & Yablonka-Reuveni, 2004). The discovery of FAPs provides an alternative hypothesis as to the origin of fibrosis and fatty degeneration. Since FAPs are capable of producing both adipocytes and fibroblasts, this new cell type has been proposed to directly contribute to fibrosis and fatty infiltration when muscle regeneration is impaired. Indeed, both Uezumi et al. and Joe et al. demonstrate that transplant of FAPs into damaged muscle results in donor-derived fat, providing first-hand evidence that FAPs can generate fat in vivo (Joe, et al., 2010; Uezumi, et al., 2010). A balance exists between the contribution of FAPs to efficient muscle regeneration and to scartissue forming fibrosis (Natarajan, Lemos, & Rossi, 2010) (Figure 1.5). However, the exact regulatory factors involved in modulating communication between FAPs, satellite cells, the microenvironment, and other cell types involved remain unknown. Further elucidation of this intricate balance of FAP contribution to regeneration or fibrosis would be of great interest, especially when considering therapeutic potential.  40  Figure 1.5  Distinct roles for FAPs during muscle regeneration and repair.  (A) During homeostatsis, both satellite cells and FAPS are quiescent. (B) Upon muscle damage, satellite cells and FAPs activate and proliferate. (C) FAPs secrete a factor that positively influences myogenic differentiation. (D) Regeneration successfully progresses and myofibres inhibit FAP differentiation through direct contact. The muscle then returns to its normal state. (E) Under some circumstances, regeneration is insufficient and FAPs differentiate into adipocytes and fibroblasts. ! Cell cycle, 2010, by permission from Fabio M.V. Rossi.  41  1.1.5  Summary of muscle regeneration  Clearly, the process of muscle regeneration is a diverse system involving many cell types and extra-cellular factors that must cohesively function to ensure proper regulation of the various steps of the repair process (Charge & Rudnicki, 2004; Hawke & Garry, 2001). A misregulation of this intricate network results in delayed or defective regeneration. Here, we give a brief summary of the major cell types and factors discussed in this thesis (Figure 1.6).  1.1.5.1  Muscle homeostasis  In resting muscle, satellite cells are quiescent and found in their niche, between the sarcolemma and basal lamina of the myofibre (Mauro, 1961). A number of markers such as Pax7, Pax3, Myf5, c-met, and CD34 can be used to detect satellite cells (Beauchamp, et al., 2000; Cornelison & Wold, 1997; Seale, et al., 2000). FAPs are also located on the myofibre (Joe, et al., 2010; Uezumi, et al., 2010).  1.1.5.2  Myotrauma  Upon muscle damage, satellite cells activate, thus beginning the myogenic program. MRFs MyoD and Myf5 are upregulated and Pax7 expression remains (Zammit, JCS, 2006). HGF, some of which is directly synthesized and secreted by satellite cells, promotes satellite cell activation (Miller, et al., 2000; Sheehan, et al., 2000). HGF binds to its receptor, c-met, and  42  activates a variety of downstream signaling cascades such as Ras-Ral and p38 MAPK (Bischoff, 1986a; J. Suzuki, et al., 2000). These pathways have been associated with promoting later stages of regeneration such as migration and differentiation of myoblasts, respectively. In addition, p38"/! MAPK activation has been proposed to be a molecular switch that activates quiescent satellite cells (Jones, et al., 2005). Conversely, other factors such as TGF!/myostatin can inhibit satellite cell activation (McCroskery, et al., 2003; Rathbone, et al., 2011). At this time, inflammatory cells begin to accumulate in the damaged tissue. Neutrophils arrive first, followed by macrophages. Neutrophils can promote M1 macrophage-mediated phagocytosis and release of pro-inflammatory cytokines to further recruit macrophages (Chazaud, et al., 2009; Tidball & Villalta, 2010). Moreover, FAPs begin to proliferate (Joe, et al., 2010).  1.1.5.3  Myoblast proliferation and migration  Activated satellite cells exit their niche, a process mediated by lamellipodia formation (Otto, Collins-Hooper, Patel, Dash, & Patel, 2011), and satellite-cell derived myoblasts proliferate. Pax7 expression is down-regulated and MyoD expression is upregulated (Zammit, et al., 2006). Factors such as HGF, FGF, IGF, IL-6, p38# MAPK, and Notch signaling promote myoblast proliferation (Allen & Boxhorn, 1987, 1989; Conboy & Rando, 2002; Serrano, et al., 2008), while TGF!/myostatin and p38"/! MAPK signaling inhibit this process (McCroskery, et al., 2003; Perdiguero, Ruiz-Bonilla, Gresh, et al., 2007; Perdiguero, RuizBonilla, Serrano, et al., 2007). Motile myoblasts, mediated mostly by blebbing (Otto, et al., 2011), migrate to the site of damage. Factors such as HGF, IGF, and IL-4 promote this 43  motility (Allen & Boxhorn, 1989; Horsley, et al., 2003). M1 pro-inflammatory macrophages are now abundantly found at this stage and secrete a factor that also promotes myoblast motility (Chazaud, et al., 2009).  1.1.5.4  Fusion and differentiation  Myoblasts exit the cell cycle, commit to differentiation, and can fuse with the damaged fibres or to each other. MRF4 and Myogenin expression is upregulated and Pax7 and MyoD expression is down-regulated. However, some cells down-regulate MyoD while maintaining Pax7 expression and return to a quiescent state (Zammit, et al., 2006). p38"/! MAPK signaling promotes myoblast cell cycle exit and commitment to differentiation (Perdiguero, Ruiz-Bonilla, Serrano, et al., 2007), while IL-4 promotes myoblast fusion (Horsley, et al., 2003). M2 anti-inflammatory macrophages are abundantly found at this stage and secrete a factor that inhibits myoblast motility to promote differentiation (Chazaud, et al., 2009). Additional factors such as IGF, IL-6, Wnt, and p38"/! MAPK signaling also promotes differentiatiation (Brack, et al., 2008; Cuenda & Cohen, 1999; Petropoulos & Skerjanc, 2002; Serrano, et al., 2008), while FGF, TGF!, and p38# MAPK signaling inhibits it (Allen & Boxhorn, 1989; Gillespie, et al., 2009; Linkhart, et al., 1981; McCroskery, et al., 2003). Upon the conclusion of differentiation, the muscle returns to its normal resting state.  44  Figure 1.6  Schematic representation of key events during adult muscle regeneration.  (A) Adult myofibre at homeostasis, with quiescent satellite cells and FAPs located beneath the basal lamina. (B) Upon damage, satellite cells become activated and inflammatory cells invade the tissue. FAPs proliferate. (C) Satellite cells exit their niche to derive myoblasts, which then proliferate. At this point, M1 pro-inflammatory macrophages are present. (D) Myoblasts are motile and migrate to the site of damage. Myoblasts fuse with the fibre or to each other, beginning myogenic differentiation. At this point, M2 anti-inflammatory macrophages are present. (E) Terminal differentiation occurs and the regenerating myofibre acquires its characteristic centralnucleation. (F) The myofibre returns to its resting state. MRFs MyoD and Myf5 are involved in the early stages of regeneration while Myogenin and MRF4 are involved in the late stages. Also, a variety of secreted factors and signaling pathways help regulate the different stages of regeneration. Positive regulators are shown in green while negative regulators are shown in red.  45  1.1.6  Models of skeletal muscle regeneration  As with the study of any given system or tissue, models need to be available that adequately and consistently mimic the system being studied, but also have the flexibility to allow for experimental manipulation. For the study of adult myogenesis or muscle regeneration, in vivo and in vitro models exist. Each model has its own benefits and drawbacks. Significant differences between in vivo and in vitro models exist (Cornelison, 2008) and no one given model perfectly recapitulates the entire process of regeneration. We summarize below some models that have been well established and commonly used.  1.1.6.1  In vivo models  In vivo models provide the advantage of physiological and environmental conditions. However, because in vivo models involve the use of animals, the disadvantage lies in the ethical issues, cost, and availability of animal models. Nevertheless, the use of in vivo models is commonplace and almost mandatory in the field of regenerative medicine.  1.1.6.1.1  Acute damage  Acute muscle damage occurs as a result of one traumatic event. Generally, because normal animals that undergo acute injury models fully recover in a given timeframe, acute damage models are used to study specific events that occur during one cycle of normal muscle regeneration or to assess the effects of various factors or genes on regeneration. 46  Myotoxins such as bupivacaine, cardiotoxin (CTX), and notexin (NTX), are commonly used to induce muscle injury and subsequent regeneration in vivo. Bupivacaine is a known local anesthetic that can also cause skeletal muscle injury (Hall-Craggs, 1974; Rosenblatt & Woods, 1992), although the exact mechanism behind its myotoxicity is unknown. CTX and NTX are both snake venoms isolated from spitting cobra and tiger snake, respectively. CTX is a peptide that acts as a protein kinase C-specific inhibitor that is thought to disrupt muscle membrane organization and cause cell lysis by inducing depolarization and muscle contraction. NTX is a phospholipase A2 neurotoxin peptide that also causes membrane depolarization and disruption, resulting in muscle damage (Harris & MacDonell, 1981). A tremendous benefit of myotoxin use is that only differentiated myofibres are specifically damaged, whereas nerves, blood vessels, and satellite cells are undisturbed. Because of technical ease of direct intramuscular (i.m.) injection of the agent, availability, cost, and overall reproducibility of the damage generated, myotoxins are one of the most popular methods used to produce a regeneration timecourse in vivo. However, the degree of damage and time required for complete recovery depends on the type and amount of myotoxic agent used. In our experiments, we use NTX to induce acute muscle damage in mice.  Another method employed to induce regeneration is by physically damaging the muscle of interest. Several methods are popularly used for this. Crush injury involves the application of a weight directly on to the muscle, thereby causing injury. Although relatively crude, this model allows for variations in the weight used and the resulting incurred damage. In the freeze damage model, a liquid-nitrogen cooled rod, or alternatively, a piece of dry ice, is applied directly to an area of muscle (Gayraud-Morel, et al., 2007). The duration and number  47  of repetitions can vary, depending on the amount of damage one is required to induce. Like CTX or NTX damage, the muscle membrane is left intact; however, unlike myotoxins, which result in a relatively even distribution of cellular debris, freeze injured muscle has a distinguishable damaged (proximal to direct site of freezing) and regenerating (distal to the site of freezing) zone.  1.1.6.1.2  Chronic damage  Chronic muscle damage occurs as a result of repeated trauma to the muscle over time. Contrary to acute damage models, continuous chronic damage over a long period of time results in dysregulated muscle regeneration, usually associated with scar tissue formation and fatty infiltration (fibrosis). Generally, chronic injury models are used to mimic human injuries or disease, or when fibrosis is being examined.  Genetic disease modeling is commonly used when the gene mutation or disruption causing the disease is known. This type of model is especially popular when trying to better understand human disease. Among myodegenerative diseases, DMD is the most common and devastating, affecting 1 in 3500 boys. The myopathy seen in DMD patients is caused by a mutation in the dystrophin gene resulting in a functional loss of the dystrophin protein (Hoffman, Brown, & Kunkel, 1987). The lack of a functional dystrophin, a key component of the dystroglycan complex (DGC) that links the cytoskeleton of muscle to extracellular matrix (ECM) (Henry & Campbell, 1999), results in constant muscle damage and repair and eventual fibrosis. Like DMD patients, mdx mice carry a point mutation in the dystrophin 48  gene that results in the functional loss dystrophin (Sicinski, et al., 1989). Thus, mdx animals are used to model DMD or chronic injury. However, despite the genetic match of mdx mice to DMD patients, mdx mice have a comparatively mild phenotype and live relatively normal lives (Tanabe, Esaki, & Nomura, 1986). So, in order to better mimic human disease, mdx mice can be crossed to other strains such as utrophin knock-out (Grady, et al., 1997) or telomerase knock-out animals (Sacco, et al., 2010). Moreover, like dystrophin, mutations in any structurally important membrane protein or component of the DGC can also result in muscle disruption and damage (Durbeej & Campbell, 2002).  1.1.6.2  In vitro models  Cell culture techniques are commonly used to study a wide-range of events. These in vitro models are relatively inexpensive compared to in vivo models, have less variability, and are easier to scale-up when needed. Also, in vitro models allow for a more controlled environment with endless possibilities to manipulate the system. However, in vitro events never perfectly recapitulate those that occur in vivo (Cornelison, 2008) and are thus generally complemented with in vivo experiments.  49  1.1.6.2.1  Single fibre cultures  One of the most commonly used systems to isolate satellite cells and also to study myogenic progression is with single fibre cultures. In this model, groups of muscle such as the EDL and soleus can be delicately dissected out from mice and digested with collagenase. The digestion results in the release of single fibres from the original muscle group, which can then be individually picked and separately cultured. Satellite cells, still intact on their host fibres immediately after isolation, activate shortly after culture and the rest of myogenic progression (migration, proliferation, and differentiation) proceeds (Collins & Zammit, 2009; Rosenblatt, et al., 1995). Because satellite cells are still under the basal lamina upon immediate isolation, single fibre cultures are currently the only way to harvest niche-bound satellite cells. The beauty of this technique is that it incorporates the benefits of cell culture, while still maintaining the ability to have an overview of the entire process of muscle regeneration, from quiescence to differentiation. Notably, i.m. injections of isolated single fibres have been used to study satellite cell engraftment and self-renewal (Collins, et al., 2005). In our experiments, we also use single fibre cultures to examine myogenic progression as a whole.  1.2  CD34  Since its initial discovery and continued use to clinically isolate stem cells, the quest to define a functional role for CD34 began. Surprisingly, despite the enormous effort put into this task, not much is know about the function of this elusive protein, even with respect to the  50  well-characterized hematopoietic system. Below, we provide a general overview of the current knowledge of CD34.  1.2.1  CD34: a stem cell marker  The discovery of the existence and therapeutic potential of hematopoietic stem or progenitor cells fuelled the search for stem cell markers that could be used to isolate these powerful cells. In a screen of antibodies generated against human hematopoietic progenitors, CD34 was first identified (Andrews, Singer, & Bernstein, 1986; Civin, et al., 1984; Katz, Tindle, Sutherland, & Greaves, 1985). In 1984, Civin et al. first showed that a monoclonal antibody reacts to a 115 kDa antigen present on normal human hematopoietic progenitor cells and leukemic marrow cells, but absent from mature cell types (Civin, et al., 1984). Subsequent in vivo transplants demonstrating that the CD34+ fraction of BM can engraft and replenish the hematopoietic system of lethally irradiated baboons (Berenson, et al., 1988) and human cancer patients (Berenson, et al., 1991) further added to its heralded role as a “stem cell marker”. Currently, CD34 is still the most widely used marker to clinically isolate hematopoietic progenitors (Sutherland, Stewart, & Keating, 1993).  1.2.2  CD34 and CD34-related family members: structure and expression  CD34 belongs to a family of CD34-family of proteins that also include Podocalyxin and Endoglycan as members (Figure 1.7). All three family members have a similar genomic organization of eight exons. At the mRNA level, alternative splicing of CD34 (Suda, et al.,  51  1992) and Podocalyxin (Li, Li, Brophy, & Kershawt, 2001) can occur, resulting in two protein isoforms. The full-length isoform (CD34FL) has a shorter transcript but longer protein; conversely, the cytoplasmic truncated isoform (CD34CT) has a longer transcript but shorter protein that lacks most of the cytoplasmic domain. The inverse relationship between transcript length and protein size is attributed to the presence of exonX in the CD34CT isoform. ExonX is located between exons 7 and 8 and that contains a premature stop codon (Krause, Fackler, Civin, & May, 1996; Suda, et al., 1992).  CD34 and CD34-family members are type-I, transmembrane, sialomucins and classified in the same family based on domain organization. Although there are slight variations among the three members, CD34 and CD34-family members are generally structured as follows (Furness & McNagny, 2006; Nielsen & McNagny, 2008): (1) An extracellular region consisting of an N-terminal mucin domain followed by a cysteine-bonded globular domain and stalk. The mucin domain contains a few putative N-linked and many O-linked glycosylations, some of which have sialic acid modifications. This mucin structure contributes to the highly net negative charge of CD34. (2) A transmembrane region passing through the bilipid layer of the cell membrane. (3) An intacellular cytoplasmic tail containing putative phosphorylation sites and a PDZ-binding motif. The cytoplasmic tail is proposed to have signaling-related functions (Fackler, Civin, & May, 1992; Fackler, Civin, Sutherland, Baker, & May, 1990; Felschow, McVeigh, Hoehn, Civin, & Fackler, 2001).  52  Figure 1.7  Schematic representation of the CD34 family.  (A) Genomic organization of CD34 family. Each colour corresponds to a specific protein domain. (B) Alternative splicing showing the derivation two isoforms for CD34 and Podocalyxin. (C) Protein structures of CD34, Podocalyxin, and Endoglycan. Horizontal lines and horizontal lines with arrowheads represent Oglycosylations and sialylated residues, respectively. Lines with circles represent putative N-glycosylations. The extracellular component is composed of a mucin domain (green), cysteine rich globular domain (blue), and a stalk domain (yellow). This is followed by a transmembrane region (light blue) and intracellular cytoplasmic tail (red) containing putative phosphorylation sites. ! Adapted with permission. Nielsen and McNagny, Journal of Cell Science, 2008;121(Pt 22):3683-3692.  53  In adult tissues, the three proteins have overlapping expression on hematopoietic precursors (Doyonnas, et al., 2005; Fieger, Sassetti, & Rosen, 2003; Krause, et al., 1994; McNagny, et al., 1997) and vascular endothelia (Baumheter, et al., 1993; Doyonnas, et al., 2005; Sassetti, Tangemann, Singer, Kershaw, & Rosen, 1998; Sassetti, Van Zante, & Rosen, 2000). In addition, each family member is also expressed on its own in a number of cell types. CD34 is expressed on mast cells (Drew, Merkens, Chelliah, Doyonnas, & McNagny, 2002), eosinophils (Blanchet, et al., 2007; Radinger, Johansson, Sitkauskiene, Sjostrand, & Lotvall, 2004), muscle satellite cells (Beauchamp, et al., 2000), and hair follicle stem cells (Trempus, et al., 2003). Podocalyxin, otherwise known as MEP21, podocalyxin-like protein 1, thrombomucin, and gp135, is expressed on kidney glomerular podocytes (hence its name “Podocalyxin”) (Kerjaschki, Sharkey, & Farquhar, 1984), platelets, megakaryocytes (Miettinen, et al., 1999), and nucleated erythrocytes (Doyonnas, et al., 2005). Endoglycan is expressed on smooth muscle (Sassetti, et al., 2000), T- and B-cells (Kerr, Fieger, Snapp, & Rosen, 2008).  Although several studies assessing the function of Podocalyxin and Endoglycan have been done (Furness & McNagny, 2006; Nielsen & McNagny, 2008), the work presented here specifically focuses on CD34’s role in adult muscle regeneration and myogenic progenitors, which only express CD34 and not Podocalyxin or Endoglycan. Below, we summarize the current proposed functions of CD34.  54  1.2.3  Proposed functions of CD34  With CD34 being a stem cell marker, many studies investigating the role of CD34 have been done. The generation and availability of Cd34 null mice greatly helped in such studies (Cheng, et al., 1996; A. Suzuki, et al., 1996) and several proposed functions have now been put forth for this sialomucin. However, since these proposed roles are still debatable, work is still ongoing to provide further insight and elucidate possible mechanisms.  1.2.3.1  CD34 marks cells at an activated state  The demonstration of CD34+ BM-derived cells being able to reconstitute the complete hematopoietic system of irradiated hosts (Berenson, et al., 1988; Berenson, et al., 1991) indicated that a stem or multipotent progenitor cell type exists within this fraction. However, when tested, the CD34- fraction is also able to reconstitute irradiated hosts (Bhatia, Bonnet, Murdoch, Gan, & Dick, 1998; Goodell, Brose, Paradis, Conner, & Mulligan, 1996; Goodell, et al., 1997; Osawa, Hanada, Hamada, & Nakauchi, 1996). These contradictory sets of data were controversial as CD34 was already being used to clinically isolate stem cells. The beautifully simple, yet straightforward, work by Sato et al. provided some insight to the expression of CD34 on hematopoietic stem cells (Sato et al, 1999, blood). These authors show that the stem cell fraction of BM, with long-term reconstituting activity, lies mostly within the CD34- fraction. However, upon 5-FU depletion of proliferating, circulating cells, stem cell activity is present in both CD34- and CD34+ fractions, suggesting that CD34 marks an activated state of progenitors. Serial transplants of CD34- and CD34+ BM show that 55  activated CD34+ cells revert back to a CD34- state (Sato, Laver, & Ogawa, 1999). Follow up experiments later published by the same group confirm that, upon granulocyte-colony stimulating factor (G-CSF) mobilization of BM cells, the long-term engrafting HSCs are CD34+ and become CD34- later on (Tajima, Sato, Laver, & Ogawa, 2000). Analysis of the cell cycle state of CD34- and CD34+ HSCs further show that quiescent, G0 HSCs are CD34and G-CSF mobilized CD34+ HSCs are in a G0/G1 activated state (Roberts & Metcalf, 1995; Uchida, et al., 1997). Together, these data clarifies the conundrum in finding stem cell activity in both CD34- and CD34+ fractions of HSCs. Interestingly, when looking at CD34+ HSC frequency as a function of age, Ito et al. show that all HSCs in young, 5-week old, mice are CD34+. This frequency declines starting at 7-weeks of age and in adults, only 20% of HSCs are CD34+ (Ito, Tajima, & Ogawa, 2000; Ogawa, et al., 2001). Together, these data demonstrate that although HSCs express CD34 in young animals, CD34 marks only an activated state of HSCs in adults and that this expression is reversible (Ogawa, 2002). Importantly, these works suggest that CD34 may play an important role in mediating the process of activation or have a direct function on activated cells.  A study by Trempus et al. looking at the role of CD34 in tumor development shows that in comparison to WT controls, Cd34 null animals have decreased tumorigenesis. Analyzing the state of the Cd34 deficient hair follicles revealed that they remained in a non-cycling state, rather than transitioning to a growth state (Trempus, et al., 2007). Since hair follicle stem cells are CD34+ and proposed to be a target of carcinogens in certain models of skin carcinogenesis (Trempus, et al., 2007), these results suggest that CD34 is involved in hair  56  follicle stem cell activation, thus providing support to the notion that CD34 may be important in mediating activation.  1.2.3.2  CD34 promotes proliferation and inhibits differentiation  A role for CD34 in promoting proliferation and inhibiting differentiation has long been suggested, mostly due to its expression on activated HSCs and lack thereof on most mature hematopoietic cell types. However, few lines of work provide data that directly support this hypothesis. Analysis of hematopoietic progenitors from the yolk sac or fetal livers from Cd34 null embryos by Cheng et al. show decreased colony-forming activity and decreased overall numbers in these cells (Cheng, et al., 1996). In adult Cd34 null mice, BM- and spleenderived cells also have decreased colony-forming activity, suggesting that CD34 is important in HSC progenitor cell formation in developing and adult mice (Cheng, et al., 1996). Lastly, CD34+ leukemic subsets have increased proliferation and decreased apoptosis compared to CD34- subsets (Shman, Savitski, Fedasenka, & Aleinikova, 2007). Together, these results suggest that CD34 is necessary for efficient proliferation.  The role for CD34 in preventing differentiation is generally supported by the presence of this marker on immature, proliferative, hematopoietic progenitors. CD34’s subsequent downregulation as these progenitors differentiate hints towards a role for CD34 in preventing differentiation. Using M1 myeloblastic leukemia cell lines, Fackler et al. provides more direct evidence for CD34’s inhibitory role in differentiation (Fackler, Krause, Smith, Civin, & May, 1995). M1 cells normally express CD34 in its undifferentiated state and lose this 57  expression upon terminal differentiation. When CD34 is overexpressed in M1 cells, terminal differentiation is inhibited. Surprisingly, only the CD34FL isoform has this inhibitory function (Fackler, et al., 1995). This work directly demonstrates that CD34 plays a role in preventing differentiation. Moreover, the differential inhibitory activity of the two isoforms, mainly differing in the presence (CD34FL) or absence (CD34CT) of the cytoplasmic domain containing putative phosphorylation sites (Blom, Gammeltoft, & Brunak, 1999; Blom, Sicheritz-Ponten, Gupta, Gammeltoft, & Brunak, 2004), further suggests that the intracellular region could be involved in preventing differentiation possibly through signaling pathways (discussed below).  1.2.3.3  CD34 as a pro- or anti- adhesion molecule to facilitate migration  The identification of extracellular and intracellular binding partners for CD34 could potentially elucidate more roles and even provide insight into the mechanism that enables CD34 to perform such roles. L-selectin (L-sel) has been reported to be an extracellular binding partner for CD34 (Baumheter, et al., 1993). As leukocyte rolling depends on the lowaffinity binding of L-sel+ leukocytes to specialized endothelia called high endothelial venules (HEVs), CD34 has been suggested to be a pro-adhesive molecule. Indeed, in vitro laminar flow assays show that up to half of the L-sel mediated lymphocyte tethering and rolling can be attributed to CD34; conversely, CD34- cells tether less and roll faster (Puri, Finger, Gaudernack, & Springer, 1995). Although this data provides evidence for a pro-adhesive role for CD34 in leukocyte tethering and rolling, the Sialyl-Lewis-X CD34 modification required for L-sel binding is uniquely found on lymph node HEVs (Paavonen & Renkonen, 1992) and 58  in no other CD34+ cell types. This suggests that this pro-adhesive function of CD34 may be more of an exception rather than a rule.  Conversely, an anti-adhesion, or “molecular teflon”, role for CD34 has also been proposed (Drew, Merzaban, Seo, Ziltener, & McNagny, 2005). Using WT and Cd34 null mast cells, Drew et al., observed significantly increased homotypic aggregation in Cd34 null groups (Drew, Merzaban, et al., 2005). This can be reversed upon re-expression of CD34, indicating that CD34 has anti-adhesive properties. Moreover, although there is no significant difference in the frequency of mast cell progenitors between WT and Cd34 null animals, CD34 is required for mast cell repopulation and hematopoietic progenitor reconstitution in vivo, a function attributed to its anti-adhesive properties (Drew, Merzaban, et al., 2005). Overall, this suggests that CD34 plays an important role in mediating proper adhesion required for migration.  Later studies by the same group further show that CD34 is required for the migration of inflammatory cells to mediate disease progression. Decreased susceptibility and delayed pathology of mice to allergic asthma and Salmonella-induced gastroenteritis, respectively, is observed in Cd34 null mice (Blanchet, et al., 2007; Grassl, et al., 2010). Cytokine profiling performed on Cd34 null cells in both models showed comparable levels to WT controls. However, analysis of eosinophil and neutrophil kinetics, both important inflammatory mediators, show that Cd34 null cells have impaired migration to the lung and gut (Blanchet, et al., 2007; Grassl, et al., 2010), providing an explanation to the decreased disease susceptibility and pathology in Cd34 null animals. The mechanism by which CD34 promotes  59  efficient migration remains unknown, although it has been speculated that CD34 inhibits adhesion by steric hindrance due to its bulky, negatively charged mucin domain, with its cytoplasmic tail being involved in the re-localization or downregulation of surface CD34 when this hindrance is no longer needed (Furness & McNagny, 2006; Nielsen & McNagny, 2008). Despite all this, CD34 null adult mice have a relatively normal phenotype and hematopoietic lineage distribution. Therefore, CD34 is likely not necessary for cell maintenance at a homeostatic state, but rather, for proper cell functions upon tissue injury.  1.2.3.4  CD34 and signaling  To date, no direct evidence has been described that directly links CD34 to known signaling pathways. Nonetheless, several publications do provide some support that CD34 is involved in signal transduction. Studies show that CD34 can be directly phosphorylated by protein kinase C (PKC) on hematopoietic (Fackler, et al., 1990) and acute lymphoblastic leukemia cells (Sutherland, Fackler, May, Matthews, & Baker, 1992). PKC phosphorylation upregulates CD34 expression as fast as 1 minute post-phosphorylation (Fackler, et al., 1992). To identify specific phosphorylation sites on the cytplasmic tail of CD34, Deterding et al. used mass spectroscopy (Deterding, et al., 2011). Surprisingly, the in vivo sites determined were not consensus PKC phosphorylation sites, but, instead, were potential sites for AKT2 phosphorylation (Deterding, et al., 2011). These data suggests that PKC, AKT2, and possibly other kinases, are likely involved in CD34 signaling functions.  60  The only known intracellular binding partner of CD34 is CrkL (Felschow, et al., 2001), a member of the Crk family of adaptor proteins. Crk proteins contain one SH2 domain and two SH3 domains and generally link non-kinase proteins to intracellular signaling cascades that allow such proteins to indirectly tranduce signals. The exact binding site of CrkL on CD34 is still unidentified. But, Felschow et al. have shown that the SH3 domain of CrkL binds CD34 and that this binding is dependent on a 10 amino-acid sequence on the intracellular domain of CD34 present in both isoforms (Felschow, et al., 2001). Lastly, crosslinking studies with antibodies to CD34 show that antibody-induced capping of CD34 results in actin polymerization and phosphorylation of Lyn and Syk (Tada, Omine, Suda, & Yamaguchi, 1999), possibly providing a link between CD34’s role in adhesion and signaling.  In summary, despite the array of proposed roles for CD34, the exact mechanisms for these have yet to be defined. Moreover, specific functions for CD34 may depend on the cell type it is expressed on.  1.3  CD34 and muscle regeneration  Currently, there is no efficient way to treat severe myodegenerative diseases. Thus, regulators of muscle regeneration are continually being investigated in hopes of developing novel therapeutic strategies. CD34 is a commonly used satellite cell marker whose expression is regulated during muscle regeneration. However, its function in this process is unknown. In this thesis, we investigate the potential role of CD34 in satellite cell function and overall muscle regeneration.  61  Beauchamp and colleagues were the first to show that various myogenic cell lines and primary myogenic progenitors express CD34 (Beauchamp, et al., 2000). Moreover, in situ hybridization analysis of whole mount embryos at different stages show that CD34 transcripts are present in skeletal muscle later during development, starting at E16.5, specifically in putative satellite cell locations (Beauchamp, et al., 2000). This pattern of expression is identical to that of m-cadherin, a marker of myogenic cells (Irintchev, Zeschnigk, Starzinski-Powitz, & Wernig, 1994). Single fibre isolations from adult mice confirm that CD34 is indeed present on m-cadherin+ satellite cells. Upon culture of the single fibres, this group reported that CD34 transcripts undergo alternative splicing resulting in differential isoform expression during the progression of satellite cells through the myogenic program (Beauchamp, et al., 2000). The isoform switch from CD34CT to CD34FL occurs during the early stages, such as activation and proliferation, but expression is completely down-regulated at later stages, such as differentiation (Beauchamp, et al., 2000).  However, despite these intriguing results that provided initial insight into CD34 and satellite cells, not much work has been done to define a function for this protein during skeletal muscle regeneration. Thus, upon the initiation of the work presented in this thesis, the only link between CD34 and muscle regeneration known was its use as a marker for quiescent satellite cells and that upon in vitro culture of single fibres, alternative splice variants of CD34 are expressed, followed by its complete transcriptional downregulation during myogenic differentiation.  62  1.4  Hypothesis and specific objectives  The objective of this thesis is to investigate the role of CD34 in satellite cells and muscle regeneration, focusing on elucidating what function CD34 serves on satellite cells during the different stages of adult regeneration. Because of the precise timing of specific CD34 transcript expression (Beauchamp, et al., 2000), we hypothesize that CD34 is important during the early stages of regeneration, such as satellite cell quiescence, activation, and proliferation. Additionally, we hypothesize that CD34 may play similar functions in satellite cells as those proposed in hematopoietic cells.  1.4.1  CD34 maintains satellite cell quiescence  The work by Beauchamp and colleagues show that CD34 is expressed on quiescent satellite cells and is shut down later during myogenic progression (Beauchamp, et al., 2000). This regulated expression is reminiscent of that described in the hematopoietic system, where CD34 is expressed on immature progenitor subsets and absent from differentiated cells. Thus, as previously suggested for CD34 in hematopoietic progenitors, we hypothesize that CD34 plays a role in maintaining the quiescent state of satellite cells.  63  1.4.2  CD34 enhances myogenic progenitor cell progression through activation and  proliferation  In addition to CD34’s presence on quiescent satellite cells, the regulated expression of CD34 isoforms coinciding with satellite cell activation and proliferation leads us to hypothesize that CD34 has a functional role in these processes, possibly through downstream signaling or intracellular cytoskeletal interactions via its cytoplasmic tail. Lastly, similar to the transient expression of CD34 on mobilized HSCs (Ogawa, et al., 2001; Sato, et al., 1999), CD34 may also mark activated and cycling satellite cells.  1.4.3  CD34 promotes satellite cell motility  The most characterized role for CD34 has been its function as an anti-adhesive molecule on hematopoietic cells to promote efficient migration during inflammation (Furness & McNagny, 2006; Nielsen & McNagny, 2008). Similarly, during in vivo (Phillips, Hoffman, & Knighton, 1990; Schultz, Jaryszak, & Valliere, 1985) and in vitro (Otto, et al., 2011; Siegel, Atchison, Fisher, Davis, & Cornelison, 2009; Siegel, Kuhlmann, & Cornelison, 2011) models of regeneration, satellite cells have been proposed to be extremely motile. Based on this, we hypothesize that CD34 plays an important role in promoting efficient motility of satellite cells during regeneration through its anti-adhesive properties. As such, CD34’s downregulation on differentiating myogenic cells may aid in promoting adhesion necessary for myoblast fusion.  64  1.4.4  CD34 prevents premature myoblast differentiation  CD34FL has been shown to prevent differentiation of M1 cell lines (Fackler, et al., 1995), suggesting that it is specifically the cytoplasmic tail that provides this inhibitory function. Like M1 cell lines, myogenic cells down-regulate CD34 upon differentiation, indicating CD34 may be necessary in stages leading up to differentiation. Taken together, we propose that CD34 may also play a role in preventing premature myoblast differentiation in adult regeneration.  Overall, we hypothesize that CD34 is not only a marker for satellite cells, but that it also plays a fundamental role in the different stages of adult muscle regeneration. Thus, the specific objectives of this work are: •  To characterize CD34 expression on WT satellite cells in vivo  •  To determine if CD34 is necessary for efficient muscle regeneration in adult mice  •  To determine in which stage(s) of muscle regeneration CD34 plays a functional role  65  Chapter 2: Characterization of CD34 expression on myogenic progenitor cells during adult muscle regeneration.  2.1 Introduction and rationale  Skeletal muscle damage, caused by exercise or genetic disease, triggers the activation of quiescent satellite cells, myogenic precursors that are the primary and most significant contributors to muscle regeneration. These cells expand and eventually fuse to each other or into ailing myofibers to repair muscle. Ineffective treatments for chronic myodegenerative diseases such as DMD (Partridge, 2002) have prompted the interest and necessity to better understand old or identify new molecular mechanisms that promote efficient muscle regeneration. These include, but are not limited to, identifying novel cell types that could be a viable resource for cell therapy (Meregalli, Farini, Parolini, Maciotta, & Torrente, 2010; Peault, et al., 2007), developing tools to enhance current outcomes of myoblast/satellite cell transplant, having a better understanding of the steps of myogenic progression and identifying regulatory factors that could enhance each of these steps.  In this work, we propose that CD34 may be one such regulator. Previous work by Beauchamp et al. showing tightly-regulated expression of CD34 on satellite cells in vitro early during myogenic progression suggests a role for this sialomucin in muscle regeneration (Beauchamp, et al., 2000). CD34 was initially discovered as a hematopoietic stem cell marker is is widely used to clinically isolate these powerful cells. Investigations using the hematopoietic system have yieled several possible roles for CD34 (summarized in the  66  introduction) (Furness & McNagny, 2006; Nielsen & McNagny, 2008). To date, no consensus has been made on its exact function although there is strong evidence for its importance in promoting hematopoietic cell migration during inflammation (Blanchet, et al., 2007; Grassl, et al., 2010). Similarly, CD34 is also commonly used to isolate murine satellite cells, the myogenic precursors of adult skeletal muscle. Little is known CD34’s function during adult muscle regeneration. But, based on the tightly regulated expression during myogenic progression (Beauchamp, et al., 2000), a functional role for CD34 in muscle regeneration has been proposed. Yet, to date, no concrete evidence supporting this notion has been provided.  Here, we investigated the possible role of CD34 in muscle regeneration by first assessing its expression on adult MPCs during myogenesis, defined here as the process of MPCs progressing through the different stages of the myogenic program, in both in vitro and in vivo settings. These specific stages include activation, migration, proliferation, and differentiation. We began by developing a flow cytometry based method to analyze and prospectively isolate MPCs. With this, we show that CD34 is indeed an excellent marker to isolate MPCs from undamaged adult skeletal muscle. Importantly, our results also demonstrate that during regeneration, CD34 expression specifically on MPCs is regulated in vivo. In vitro analysis of satellite cells on cultured single fibres confirm these findings.  Overall, to our knowledge, the observations and results described in this chapter are the first to provide evidence showing in vivo regulation of CD34 specifically in MPCs during skeletal  67  muscle regeneration. Based on this early regulation of CD34 in the myogenic program, we propose that such role is likely to be in satellite cell activation, migration, or proliferation.  2.2  CD34 is an excellent marker for identification and isolation of murine MPCs from  adult skeletal muscle  To begin our investigation of CD34 in adult muscle regeneration, we first confirmed the validity of this surface protein as a reliable satellite cell and MPC marker. Thus, individual, single myofibres were isolated from WT EDL muscles, immediately fixed, and stained for CD34 and Pax7, a satellite-cell specific transcription factor. Immunofluorescent analyses of these fibres confirm that CD34 is indeed present on the surface satellite cells (Figure 2.1).  68  Figure 2.1  CD34 is expressed on satellite cells found on isolated single myofibres.  Single myofibres were isolated from EDL muscles of WT adult mice and immediately fixed and stained. Confocal microscopy was used to visualize expression of Pax7, CD34, and laminin (CD34, green; Pax7, red; laminin, blue). Four separate fibres are shown above as an example.  Although isolating single fibres is a commonly used technique to study satellite cells, we also required a method that will allow us to reliably and consistently isolate large numbers of satellite cells or MPCs. For this, we chose FACS. Traditionally, MPCs have been isolated with Percoll-gradient isolations or pre-plating followed by culture of digested muscle tissue (Rando & Blau, 1994). However, MPC isolation by FACS has been gaining popularity  69  recently since it is relatively easy, cost effective, and the technique can be performed reliably among different groups.  Based on some published works (Montarras, et al., 2005; Sherwood, et al., 2004), we optimized a flow-cytometry based method for MPC isolation. Briefly, hind-limb muscles are dissected from adult animals and undergo a 2-step digestion with collagenaseII followed by collagenaseD/Dispase. After filtering, the resulting single cell suspension is stained with fluorescently-labeled antibodies (Figure 2.2).  Figure 2.2  Schematic diagram of skeletal muscle processing to isolate MPCs.  Hoechst+ and propidium iodide+ (PI) gates were used to for nucleated, live cells, respectively. CD31- and CD45- gates were used to exclude endothelial and hematopoietic cells, respectively. Lastly, in a Sca1 vs. CD34 plot, we find that the MPCs exclusively fall within the Sca1- CD34+ population (Joe, et al., 2010), verifying our method and that CD34 is expressed on adult myogenic progenitors. Samples containing isotype control antibodies were used to determine specificity of the gating (Figure 2.3).  70  Figure 2.3  Gating strategy for prospective isolation MPCs by FACS.  Representative plots showing gating used for MPC isolation by flow cytometry. MPCs were isolated by gating for the nucleated, live cell fraction (Hoechst+ and PI-, respectively), followed by exclusion of hematopoietic and endothelial cells (identified as CD45- and CD31-, respectively), leaving MPCs exclusively within the Sca1CD34+ population. An isotype control was used to verify antibody specificity and gating accuracy.  The myogenicity of the Sca1- CD34+ population was assessed both in vitro and in vivo. Limiting-dilution colony assays show that the frequency of myogenic cells within this population is approximately 1 in 16 (Joe, et al., 2010). Multinucleated, terminally differentiated myotubes, marked by Myosin heavy chain (MyHC), were observed during in vitro culture; complementary to this, when MPCs were isolated from mice ubiquitously expressing green fluorescent protein (GFP) mice, donor-derived, GFP+ myofibres were observed following in vivo transplant (Figure 2.4).  71  Figure 2.4  Sorted Sca1- CD34+ MPCs from adult skeletal muscle are myogenic.  (A) Immunofluorescent imaging showing GFP+, donor-derived, myofibres in WT recipient muscle 3 weeks following i.m. Sca1- CD34+ MPC transplantation. (GFP, green; Laminin, red; Hoechst, blue). (B) Sca1- CD34+ MPCs from WT animals differentiate to form multinucleated myotubes. (MyHC, green; Hoechst, blue). Representative images are shown.  Interestingly, another CD34+ population distinct from MPCs exists within skeletal muscle. Termed FAPs, this Sca1+ CD34+ population is not myogenic, but instead has the bi-potential capability to form adipocytes or fibroblasts (Joe, et al., 2010; Uezumi, et al., 2010). These cells have been proposed to be regulated by myofibres (Uezumi, et al., 2010) and, in turn,  72  FAPs have also been proposed to positively influence myogenic differentiation (Joe, et al., 2010). The potential importance of CD34 on these cells and contribution of FAPs was also taken into consideration during our studies.  Altogether, these results clearly demonstrate that CD34 is an excellent marker to identify and isolate satellite cells and MPCs. Moreover, we have shown here an optimized method to isolate MPCs by FACS, providing us with an efficient tool to obtain sufficient numbers of MPCs needed for further experiments.  2.3  Alpha7 integrin as an alternative marker to CD34 for MPC isolation during  regeneration  Despite CD34 being a satellite cell and MPC marker, it has been reported to be downregulated on satellite cells following in vitro culture (Beauchamp, et al., 2000). Thus, in order to determine if this also occurs during in vivo muscle regeneration, we first had to find an alternate marker to CD34 that is not down-regulated during regeneration. Several reported markers, such as CXCR4 (Sherwood, et al., 2004), syndecan3/4 (Cornelison, et al., 2001), SM/C-2.6 (Fukada, et al., 2004), and alpha7 integrin (Blanco-Bose, et al., 2001), were taken into consideration. However, due to the availability, quality, and consistency of the antibody, alpha7 integrin was chosen and tested using the original FACS strategy described above. To assess the expression of alpha7 integrin on primary skeletal muscle, we gated for Hoechst+, PI-, CD31-, CD45-, Sca1- cells and looked at CD34 and alpha7 integrin. Four  73  resulting populations were clearly observed (Figure 2.5): (1) CD34- alpha7 integrin-, (2) CD34+ alpha7 integrin-, (3) CD34+ alpha7 integrin+, and (4) CD34- alpha7 integrin+.  Figure 2.5  Four distinct populations exist within skeletal muscle following Hoechst+, PI-, CD31-, CD45-,  Sca1- gating when assessing CD34 and alpha7 integrin expression. CD34- alpha7- (lower left), CD34+ alpha7- (upper left), CD34+ alpha7+ (upper right), CD34- alpha7+ (lower right) populations exist within skeletal muscle following Hoechst+, PI-, CD31-, CD45-, Sca1- gating. Isotype controls were used to verify antibody specificity and gating accuracy. Representative FACS plots are shown.  Culture these four populations under myogenic conditions showed that only alpha7 integrin+ populations had myogenic potential, confirmed by the presence of MyHC+ myotubes (Figure 2.6). Thus, from hereon, “MPCs” will always refer to the Sca1- alpha7 integrin+ population.  74  Figure 2.6  Alpha7 integrin + cells within adult skeletal muscle are myogenic.  Brightfield (A) and immunofluorescent (B) images of all 4 populations (from Figure 2.5) following 8 days of culture under myogenic conditions (MyHC, green; Hoechst, blue). Representative images are shown.  75  We also assessed the myogenic potential of the CD34- and CD34+ fractions within Sca1alpha7 integrin+ MPCs by isolating these populations from Myf5LacZ mice. In these animals, !-galactosidase expression is under the control of the Myf5 promoter (Tajbakhsh, Rocancourt, & Buckingham, 1996). As Myf5 is one of the four MRFs, satellite cells in Myf5LacZ animals can be easily identified by LacZ staining. Following sorting of CD34- and CD34+ MPCs, cells were cytospun and assessed for !-galactosidase activity. Our results showed that !-galactosidase activity was only found in the CD34+ fraction (Figure 2.7).  Figure 2.7  Satellite cells are contained only within the CD34+ fraction.  (A) CD34- and CD34= fractions were isolated from Myf5LacZ animals and stained for LacZ to assess !– galactosidase activity. Quantification of LacZ+ cells shows 72.3 ± 3.4% of the total cells isolated are myogenic. Error bars represent ± SEM for n=3 animals. (B) Representative image of LacZ-stained CD34- and CD34+ MPCs is shown (LacZ+, blue). Results are from 2 independent experiments.  Moreover, limiting dilution assays showed that the frequency of cells capable of initiating colonies containing multinucleated, MyHC+, myotubes is negligible within the CD34fraction (1 in 2921 cells in CD34- fraction vs. 1 in 31 cells in CD34+ fraction) (Table 2.1). Therefore, essentially all myogenic activity in sorted MPCs is contained within the CD34+ subset.  76  Table 2.1  Only CD34+ cells are capable of forming myogenic colonies.  (A) Analysis for the presence of mygenic colonies in wells plated with CD34- or CD34+ MPCs at different densities. (B) 95% confidence intervals of limiting dilution assay results to assess myogenicity of CD34- or CD34+ MPCs.  2.4  CD34 isoform and surface protein expression is regulated on MPCs during in vivo  muscle regeneration  With alpha7 integrin as a confirmed marker for MPCs that can substitute for CD34, we then characterized the regulation of CD34 expression during in vivo muscle regeneration following acute NTX damage. Muscles were harvested at specific days throughout the regeneration timecourse (day 0 to day 21) and analyzed by flow cytometry. FACS analysis revealed that, similar to in vitro observations (Beauchamp, et al., 2000), CD34 is downregulated from the surface of MPCs starting at day 3 post-damage, and is essentially absent 77  by day 5. At day 10, a time that follows the cessation of their proliferation (Joe, et al., 2010), MPCs begin to re-express CD34 on their surface and by day 21, CD34 surface expression is fully restored (Figure 2.8).  Figure 2.8  CD34 surface expression on MPCs is regulated during in vivo muscle regeneration.  Representative FACS plots showing CD34 expression on MPCs following NTX damage (n=3–5 mice per timepoint). An isotype antibody control was used to verify specificity. Results are from 4 independent experiment.  This result was confirmed with quantitative real-time PCR (qRT-PCR) analysis of sorted MPCs showing the down-regulation of total CD34 mRNA at the early timepoints following acute damage (figure 2.9).  78  Figure 2.9  Total CD34 mRNA expression is down-regulated during the first week of muscle  regeneration following acute damage. qRT-PCR analysis of total CD34 expression in MPCs after NTX damage. Error bars represent ± SEM for n=2. Results are from 3 independent experiments.  In contrast, the expression of CD34 on Sca1+ alpha7 integrin- FAPs remained constant throughout the same regeneration timecourse (Figure 2.10), despite the fact that FAPs are recruited during proliferation to the same extent as myogenic cells (Joe, et al., 2010), indicating that CD34 regulation is specific to MPCs.  79  Figure 2.10 CD34 surface expression remains constant on FAPs during in vivo muscle regeneration. Representative histograms showing consistent CD34 expression on FAPs during a regeneration timecourse following NTX damage (n=3–5 mice per timepoint). An isotype control was used to verify specificity. Results are from 3 independent experiments.  In addition, we assessed whether CD34 isoform expression is also regulated during muscle regeneration using end-point PCR with primer sets that allowed us to distinguish between CD34FL and CD34CT transcripts. Our results show that in non-damaged muscle, MPCs exclusively express CD34FL. Upon satellite cell activation and proliferation (days 1-5 postNTX damage), both CD34FL and CD34CT are co-expressed. By day 7, when regeneration is well underway, the CD34FL again becomes the predominant isoform (Figure 2.11).  80  Figure 2.11  CD34 isoform expression varies during in vivo muscle regeneration.  End-point RT-PCR analysis of CD34FL and CD34CT isoform expression in MPCs after NTX damage. Results are from 3 independent experiments.  2.5  CD34 surface expression is regulated on satellite cells during in vitro myogenesis  Because in vivo and in vitro conditions for myogenesis do not always recapitulate each other (Cornelison, 2008), we also assessed the expression of CD34 on satellite cells from isolated myofibres during culture. Single fibres were obtained from the EDL muscle and cultured for 0, 12, 24, and 42 hours and then stained for CD34. Co-staining with Pax7 identified satellite cells and the localization of CD34 was examined by confocal microscopy. Our results show that immediately after isolation, CD34 is expressed in a uniform distribution on the surface of satellite cells. Following 12 hours in culture, CD34 remains expressed on the surface; however, its expression appears patchy, possibly indicating the initial down-regulation of CD34 from the surface of satellite cells. After 24 hours in culture, down-regulation of CD34 is evident. Finally, at 42 hours, CD34 is absent from the surface of satellite cells (Figure 2.12).  81  Figure 2.12  CD34 expression is regulated on satellite cells during in vitro culture of single fibres.  Analysis of CD34 localization (B, E, H, K) in Pax7+ cells (A, D, G, J) on WT fibers at 0, 12, 24, and 42 hours post-culture (Pax7, red; CD34, green; Hoechst, blue). Representative images are shown.  Altogether, these results demonstrate that CD34 mRNA isoform and surface protein expression is dynamically regulated during the early stages of in vitro and in vivo myogenesis.  82  2.6  Summary  Our analysis of primary muscle satellite cells in both in vitro and in vivo systems confirms that CD34 is an excellent marker for satellite cells, but only in undamaged muscle. Since CD34 expression is rapidly down-regulated on cultured satellite cells early in myogenic progression, we have identified the use of alpha7 integrin as an alternative marker to CD34. With our optimized FACS-based method to prospectively analyze and isolate MPCs, we show that CD34 is down-regulated on MPCs during muscle regeneration. Thus, our in vivo results are consistent with in vitro published work (Beauchamp, et al., 2000).  This regulation of CD34 during the early stages of adult myogenesis provides evidence that CD34 may be important in activation, migration, and proliferation, as opposed to differentiation where it is no longer expressed on MPCs. With optimized tools and initial results supporting our hypothesis that CD34 plays in muscle regeneration, we continued our investigation of CD34’s role in regeneration by analyzing CD34-null (Cd34-/-) animals, presented in the next chapter.  2.7  2.7.1  Materials and methods  Mice  Animals were housed in the main animal facility of the Biomedical Research Centre in the University of British Columbia. Mice were kept under sterile conditions and handled following guidelines approved by the University of British Columbia’s Animal Care 83  Committee. EGFP expression in the GFP+CD45.2 C56BL/6 mice is under the control of cytomegalovirus enhancer-chicken beta–actin hybrid promoter and were generously provided Dr. Irving I. Weissman (Stanford University). Myf5/LacZ animals express the betagalactosidase gene under the control of the Myf5 promoter and were kindly provided by Dr. Michael Rudnicki (Ottawa Health Research Institute). Mice genotypes for Cd34-/-, GFP+, and LacZ+ were determined by PCR, fluorescence microscopy, or beta-galactosidase activity using X-gal. C56Bl/6 mice were used as WT, Ly5.2, controls, unless otherwise indicated. All mice were bred in-house.  2.7.2  Single fibre isolation and culture  Single fibre isolations were performed as per standard protocol. Briefly, the EDL muscle was gently harvested following sacrifice of the mouse and care was taken to handle only the tendons. Collagenase I (Worthington# LS004197, 400 U/mL) digestion for approximately 1 hour in 37OC was performed to obtain live, single fibres. Fibres were then cultured in Ham’s F-12 complete media + 15% horse serum + 0.5 nM bFGF + 1% P/S, harvested, and fixed with 4% PFA at specific timepoints.  2.7.3  Preparation of skeletal muscle tissue for flow cytometry/FACS  FACS isolation of myogenic progenitors was performed as previously published in Joe, et al., 2010 (Joe, et al., 2010). Briefly, primary adult murine myogenic progenitors were obtained as follows: whole hind-limb or tibialis anterior (TA) muscles were carefully  84  harvested from adult mice (6-12 weeks of age) and finely minced into small pieces using forceps. Muscles then underwent a 2-step enzymatic digestion with 0.2% collagenase type II (Roche# C6885) for 30 minutes followed by collagenase D (Roche# 1088882, 1.5 U/mL) and dispase type II (Roche# 295825, 2.4 U/mL) at 37OC for 1 hour. The homogenized muscle samples were then filtered through a 40 µm cell strainer using PBS and the cell suspension is then stained with antibodies against CD45, CD31, Sca1, and alpha7 integrin. Hoechst and propidium iodide were also added. Gating for the sorted MPC population was performed as follows: Hoechst+, PI-, CD45-, CD31-, Sca1-, and alpha7 integrin+. All cell surface staining was done on ice and isotype controls were used to determine gating. Antibodies to CD34 were used when needed.  2.7.4  Isolation of MPCs and analysis using flow cytometry  Cells were sorted on a BD FACSVantage SETM machine using BD FACSDivaTM version 4.0.1.2 software. Purity checks were done following the sort to ensure sorting efficiency and accuracy. Analysis of samples was performed using FlowJo (version 8.7)  2.7.5  Acute damage with NTX  To induce acute damage, 10 µL of notexin (Latoxan# L8104, 10 µg/mL) was injected into WT TA muscle using a 3/10 CC insulin syringe. Mice were sacrificed and muscles were harvested at days 5, 7, 10, 14, 21 post-notexin damage. TA muscles were then dissected and processed for flow cytometry analysis or MPC isolation.  85  2.7.6  Limiting-dilution assays  Freshly sorted MPCs were obtained from WT animals (8-12 weeks of age) and immediately placed into 96-well tissue culture treated plates coated with BD MatrigelTM (BD# 356235). Cells were placed at densities of 1, 5, 10, 30, 50, and 100 cells per well. Cells were maintained under growth conditions (DMEM + 20% FBS + 10% horse serum + 1% P/S + 0.1 mM bFGF) for a total of 3 weeks and then fixed with 4% PFA. Immunofluorescent staining with MyHC and Hoechst were performed to determine if myotubes were present in the wells. Myogenicity was then determined by http://bioinf.wehi.edu.au/software/elda/index.html, which is described in further detail in Hu, Y and Smyth, GK (Hu & Smyth, 2009).  2.7.7  Intramuscular transplant of sorted MPCs  Freshly sorted MPCs were obtained from GFP+CD45.2 adult mice (8-12 weeks of age). 20,000 GFP+ cells were then injected into the TA muscles of adult WT mice in a 20 µL volume of PBS using a 3/10 CC insulin syringe. Recipient mice were sacrificed 3 weeks post-transplant and perfused with PBS + 10mM EDTA followed by 4% PFA. Lower hindlimb muscles were harvested and left overnight in 20% sucrose at 4OC. Muscle tissues were then embedded in cryomolds with OCT and stored at -80OC. Serial sections of 20 µm were analyzed for engraftment of GFP+ cells and production of GFP+ donor-derived myofibres.  86  2.7.8  Immunofluorescent staining and analysis by microscopy  Following blocking using blocking buffer (25% normal goat serum, 3% BSA, 0.3% TritonX), immunofluoresecent staining on single fibres was done using antibodies to Pax7 (DSHB), CD34 (clone RAM34, eBioscience# 13-0341), and laminin (rabbit polyclonal, abcam# ab11575) diluted in 0.3% TritonX. Analysis was done by confocal microscopy (Nikon C1 laser scanning confocal microscope). For differentiated myotubes from sorted MPCs cultured in vitro, an antibody to MyHC (clone A.41025, homemade) diluted in 0.3% TritonX was used. For myofibres generated by injection of sorted MPCs into WT animals, an antibody to laminin was used. Hoechst 33342 dye (Sigma# B2261) was used to mark nuclei. Analysis was done using a Zeiss Axioplan2 microscope.  2.7.9  Quantitative real-time and RT-PCR primers  Probes for qRT-PCR analysis of CD34 were purchased from Applied Biosystems (ABI assay ID Mm00519283_m1*). The sequence for RT-PCR primers used to distinguish transcripts for CD34FL and CD34CT isoforms were 5’-AGCACAGAACTTCCCAGCAA-3’ in exons 5/6 and 5’-CCTCCACCATTCTCCGTGTA-3’ in exon 8 (Beauchamp, et al., 2000).  2.7.10 Statistical analysis  Student’s two-tailed t-test was used on all statistical analyses performed between groups. Statistical significance was set at p % 0.05.  87  Chapter 3: CD34 is necessary for proper myogenic progenitor cell function during adult myogenesis.  3.1  Introduction and rationale  Our results presented in Chapter 2 demonstrates the regulation of CD34 expression during in vitro and in vivo myogenesis, thus providing evidence that CD34 could play a functional role early during muscle regeneration. This, in combination with optimized tools to specifically look at myogenic progenitors, prompted us to further investigate CD34 in skeletal muscle regeneration. In particular, we hypothesize that CD34’s role is significant during the time frame in which it is being regulated, in particular during the early stages (such as activation, migration, and proliferation) as opposed a later stage (such as differentiation).  To test our hypothesis, we compared the myoregenerative response of WT and Cd34-/animals. Our results show that CD34 is necessary for myofibre hypertrophy in both acute and chronic damage models, a necessary component of muscle regeneration (Adams, 2006; Snow, 1990). In support of a cell autonomous role of CD34 in MPCs, our results reveal that Cd34-/- MPCs engraft with significantly decreased efficiency compared to WT controls. Further comparisons between WT and Cd34-/- MPCs isolated during various timepoints following damage show that MPCs lacking CD34 have defective proliferation following damage in vivo. However, the process of differentiation appears unaffected. Altogether, these results obtained support our hypothesis that CD34 is necessary for efficient muscle regeneration and plays an active role during the early stages of muscle regeneration.  88  3.2  CD34 is necessary for efficient muscle regeneration in response to both acute and  chronic damage  To investigate the role of CD34 in adult muscle regeneration, we began by comparing the regenerative potential of Cd34-/- mice with that of WT controls. Cd34-/- mice, generated independently by two different labs in 1996 (Cheng, et al., 1996; A. Suzuki, et al., 1996), are viable and show no obvious phenotypes under homeostatic conditions. However, phenotypes can be observed when specific systems are perturbed (Blanchet, et al., 2010; Blanchet, et al., 2007; Drew, Huettner, Tenen, & McNagny, 2005; Drew, Merzaban, et al., 2005; Grassl, et al., 2010). So, to assess whether CD34 is required for skeletal muscle regeneration, we performed histological analysis of TA muscles following acute damage with NTX. TA muscles from adult WT and Cd34-/- mice were harvested at days 0, 5, 10, 14, and 21 postdamage and analyzed by hematoxylin and eosin (H&E) staining.  The extent of total areas of damage at day 5, the earliest timepoint analyzed post-NTX damage, in the two groups were comparable (Figure 3.1), suggesting that Cd34-/- mice are not more sensitive to NTX than WT.  89  Figure 3.1  Comparable areas of damage at day 5 post-NTX injection between WT and Cd34-/- animals.  (A) H&E staining of WT and Cd34-/- muscle at 5 days after NTX. Representative images are shown. (B) Bar graph showing comparable total areas of damage between WT and CD34-/-. Error bars represent ± SEM for n=4-6 animals. Results are from 2 independent experiments.  90  However, a significant increase in the amount of necrotic myofibres was clearly visible in Cd34-/- animals as early as 5 days post-damage (Figure 3.2).  Figure 3.2  Increased levels of necrosis in Cd34-/- animals at day 5 post-NTX damage.  (A) Representative H&E staining of WT and Cd34-/- muscles 5 days post-NTX damage (B) Quantification of necrotic and regenerating areas of damage 5 days after NTX. Error bars represent ± SEM for n=4-6 mice. Results are from 2 independent experiments.  In addition, elevated levels of necrosis can be observed in Cd34-/- animals at all post-damage time points analyzed, indicating a consistent difference in regeneration efficiency between WT and Cd34-/- animals (Figure 3.3).  91  Figure 3.3  H&E analysis of WT vs Cd34-/- skeletal muscle at various timepoints post-NTX damage.  (A-J) H&E staining of WT and Cd34-/- muscles following acute NTX damage. TA muscles were analyzed at days 0, 5, 10, 14, and 21 post-NTX damage (n=5-6 mice). Representative image are shown. Results are from 2 independent experiments.  To quantify this difference, we performed standard cross-sectional area (CSA) measurements on non-damaged fibres, peripherally-nucleated fibres, and regenerating, centrally-nucleated,  92  fibres (CNFs) from both WT and Cd34-/- animals 0 and 21 days after injury, respectively. Morphometric analyses confirmed that WT CNFs are significantly larger than those from Cd34-/- animals. However, this difference in size is not seen in non-damaged myofibres, (Figure 3.4). This suggests that even though Cd34-/- animals are capable of initiating the regenerative process, Cd34-/- regenerating fibres fail to undergo hypertrophy during repair, a necessary component of muscle regeneration (Adams, 2006; Snow, 1990).  Figure 3.4  Cd34-/- regenerating myofibres fail to undergo hypertrophy following acute NTX damage.  (A) H&E staining of muscle sections showing centrally nucleated myofibers 21 days after NTX damage. (B) Myofiber CSA measurements were performed on undamaged (day 0) and damaged (day 21) myofibers. Error bars represent ± SEM for n=5-6 animals with >200 fibers per animal. Results are from 2 independent experiments.  93  Since the kinetics of satellite cell activation and proliferation may differ between acute and chronic muscle damage, we asked whether the regeneration defect observed in acutely damaged Cd34-/- mice is also present during chronic damage, as for example seen in mdx mice, the murine model for DMD. Thus, we crossed mdx mice with Cd34-/- mice to obtain mdx/Cd34-/- offspring. Groups of mdx and mdx/Cd34-/- muscles from animals of different ages underwent the same histological and morphometric analyses described above.  H&E staining shows histological differences reminiscent of those observed after NTX damage (Figure 3.5). CSA measurements of CNFs show a marked decrease in regenerating myofibre sizes of mdx/Cd34-/- animals at 4 weeks of age, a time when the first wave of myodegeneration takes place in this model (DiMario, Uzman, & Strohman, 1991), and at 6 months of age, relative to mdx controls. Interestingly, when aged, 18-month old mice were analyzed, no significant differences are observed between mdx/Cd34-/- and mdx groups (Figure 3.5). Thus, data from both acute and chronic damage models support a key role for CD34 in ensuring efficient muscle repair.  94  Figure 3.5  H&E analysis of mdx vs. mdx/Cd34-/- skeletal muscle at vaious ages.  (A-F) H&E staining of mdx and mdx/Cd34-/- muscle sections at 1 (A, D), 6 (B, E), and 18 (C, F) months of age. (G) CSA measurements performed on regenerating myofibers of mdx and mdx/Cd34-/- muscles at 1, 6, and 18 months of age. Error bars represent ± SEM for n=3-5 mice with > 200 fibers per animal. Resuls are from one experiment.  95  3.3  Impaired muscle regeneration observed in Cd34-/- can be attributed to a specific  defect in their MPCs  Many CD34+ cell types, including vascular endothelial cells, hematopoietic stem cells, mast cells, and FAPs (Baumheter, et al., 1993; Drew, et al., 2002; Joe, et al., 2010; Krause, et al., 1994; Uezumi, et al., 2010), have been proposed to participate in muscle regeneration. To test whether the observed muscle regeneration defect in Cd34-/- animals reflects a direct effect of CD34 loss on MPCs, we functionally compared WT and Cd34-/- MPCs through in vivo using cell transplantation. MPCs were freshly sorted from three groups of mice: (1) Z/AP mice ubiquitously expressing LacZ, (2) GFP+CD45.2 mice ubiquitously expressing GFP, and (3) GFP+CD45.2 lacking CD34. These 3 groups will, from hereafter, be referred to as LacZ+, WT/GFP+, and Cd34-/-/GFP+, respectively. LacZ+ MPCs, initially used as internal standards, and were mixed with either WT/GFP+ or Cd34-/-/GFP+ MPCs. These cells were then injected into non-damaged TA muscles of WT recipients and donor-cell engraftment was assessed three weeks later (Figure 3.6).  Figure 3.6  Schematic of experimental outline for cell transplantation.  96  Comparable numbers of LacZ+ fibres were observed in all samples (Figure 3.7), confirming that no bias was introduced during the manipulation of cells prior to transplantation.  Figure 3.7  Comparison of WT LacZ+ donor cell engraftment with WT/GFP+ or Cd34-/-/GFP+ MPCs.  Direct enumeration and comparison of WT LacZ+ donor-derived myofibres used as internal standards in transplantation experiments. Bar graphs displaying the relative amount of LacZ+ fibers injected with WT/GFP+ or Cd34-/-/GFP+ MPCs. Error bars represent ± SEM for n=3 mice. Results are from 5 independent experiments.  In support of a cell autonomous role of CD34 in MPCs, our results reveal that Cd34-/-/GFP+ MPCs engraft with significantly decreased efficiency compared to WT/GFP+ MPCs (Figure 3.8). In addition, experimental controls of WT or Cd34-/ MPCs transplants into WT and Cd34-/ -recipients showing no difference of engraftment based on recipient genotype (Figure 3.9) provides further evidence of a cell autonomous role for CD34 on MPCs.  97  Figure 3.8  Defective engraftment of Cd34-/- MPCs.  (A) Representative image showing engraftment of WT/GFP+ and Cd34-/-/GFP+ MPCs 3 weeks following injection into non-damaged WT recipients (GFP, green; Laminin, red). (B) Quantification of engraftment. GFP+ donor-derived myofibers were counted and normalized to the number of LacZ+ donor fibers. Ratios were then normalized to WT controls. Error bars represent ± SEM for n=3-5. Results are from 5 independent experiments.  98  Figure 3.9  Recipient genotype does not affect engraftment efficiency of transplanted MPCs.  Quantification of GFP+ MPC engraftment from WT or Cd34-/- donors to generate donor-derived myofibers in WT or Cd34-/- undamaged recipients. Error bars represent ± SEM for n=3. Results are from 2 independent experiments.  To ensure that the difference in engraftment efficiency is not due to a difference in the frequency of myogenic cells contained within WT and Cd34-/- MPCs, we sorted MPCs from WT and Cd34-/- mice carrying a Myf5LacZ transgene. No significant difference in the number of LacZ+ cells between the two groups indicates equal MPC frequency in both groups (Figure 3.10).  99  Figure 3.10  Comparable satellite cell purity from freshly sorted WT and Cd34-/- MPCs.  (A) MPCs from WT and Cd34-/- mice on a Myf5LacZ background were sorted, cytospun, and stained for LacZ to assess !-galactosidase activity. Representative images displaying LacZ+ cells in both groups are shown. (B) Frequency of LacZ+ cells in WT and Cd34-/- sorted MPCs. Error bars represent ± SEM for n=3 mice. Results are from 2 independent experiments.  Lastly, because hematopoietic cells are important in muscle regeneration and CD34 has been proposed to have a role in hematopoietic cell migration, we compared the regenerative capacity of WT mice transplanted with total WT or Cd34-/- BM following NTX damage. We hypothesized that if an impairment of Cd34-/- BM-derived cells significantly contributed to 100  the muscle regeneration defect seen in Cd34-/- animals, it would be evident in this assay. Morphometric analysis of CSA of regenerating fibres 21 days following NTX damage showed no significant difference between the 2 groups (Figure 3.11), indicating that the lack of CD34 on BM-derived cells does not significantly affect muscle regeneration.  Figure 3.11  The lack of CD34 on BM-derived cells does not significantly affect muscle regeneration.  CSA measurements performed on regenerating myofibers of WT animals transplanted with WT or Cd34-/- BM 21 days following NTX damage. Error bars represent ± SEM for n=3 mice with > 200 fibers per animal. Results are from one experiment.  In summary, these data demonstrate that CD34 expression on MPCs is required for the efficient generation of myofibres following in vivo transplant and suggest that the muscle regeneration defect seen in Cd34-/- mice can be attributed to a cell autonomous defect specific to MPCs.  101  3.4  CD34 promotes in vivo MPC proliferation  Recent publications using bioluminescence and fluorescence imaging-based tracking of transplanted MPCs suggest that a burst of proliferation occurs following transplant of MPCs to recipient muscle (Sacco, et al., 2008; Xu, Yang, Liu, & Wang, 2010). Thus, we hypothesized that inefficient proliferation of Cd34-/- MPCs may account for their relatively poor engraftment and failure to repair damaged muscle in Cd34-/- mice. To directly assess WT and Cd34-/- MPC proliferation in vivo, TA muscles of BrdU-injected mice were damaged with NTX and harvested at 0, 1, 2, 3, 5, and 7 days later. The frequency of MPCs that incorporated BrdU was assessed at all timepoints by flow cytometry (Figure 3.12).  102  Figure 3.12  Representative FACS plots showing distinct BrdU+ MPC population post-NTX damage.  Representative FACS plots showing detection of a distinct BrdU+ MPC population at 0, 1, 2, 3, 5, and 7 days following NTX damage in WT and Cd34-/- animals.  103  Our analysis showed significantly decreased BrdU incorporation in Cd34-/- MPCs at day 3 post-damage, the timepoint in which maximum BrdU incorporation occurs in WT MPCs (Joe, et al., 2010) (Figure 3.13). A similar trend of inefficient Cd34-/- MPC proliferation is observed at day 5.  Figure 3.13  Inefficient proliferation of Cd34-/- MPCs during skeletal muscle.  Graph showing the frequency of BrdU+ WT and Cd34-/- MPCs at 0, 1, 2, 3, 5, and 7 days following NTX damage. Error bars represent ± SEM for n=3-5 mice per timepoint. From the point of damage until the day of harvest, BrdU was administered i.p. twice daily and also put in the drinking water. Results are from 4 independent experiments.  Notably, this proliferation defect is not observed in the FAP population, again indicating that CD34 plays a selective role in MPC function (Figure 3.14).  104  Figure 3.14 CD34 does not affect FAP proliferation during skeletal muscle regeneration. Frequency of BrdU+ WT and Cd34-/- FAPs at 0, 1, 2, 3, 5, 7 days post-NTX damage. Error bars represent ± SEM for n=3-5 mice per timepoint. From the point of damage until the day of harvest, BrdU was administered i.p. twice daily and also put in the drinking water. Results are from 4 independent experiments.  The lower frequency of BrdU+ MPCs could reflect either a reduction in their proliferative response or the selective loss of cells lacking CD34. To distinguish between these possibilities, we used TUNEL staining of WT and Cd34-/- muscle sections harvested 3 days post-damage to detect cells actively undergoing apoptosis. We observed little to no apoptosis in both groups (Figure 3.15), indicating that the decreased amount of proliferating Cd34-/MPCs is not due to increased apoptosis.  105  Figure 3.15 Little to no cell death observed in WT and Cd34-/- muscle 3 days following NTX damage. Immunofluorescent detection apoptotic cells at day 3 post-NTX damage by TUNEL on WT and Cd34-/- muscle sections. A DNase treated WT section at the same timepoint was used as a positive control. Representative images are shown (TUNEL, green; laminin, red). Results are from 2 independent experiments.  Overall, our results suggest that CD34 is required for the efficient expansion of MPCs after muscle injury in vivo.  3.5  CD34 is dispensable for in vitro myogenic differentiation  Differentiation, whereby MPCs fuse to each other or to damaged fibres, is the last step of regeneration. Although CD34 is no longer expressed on WT MPCs at this late stage of the myogenic program, a defect in differentiation of Cd34-/- MPCs could also account for the defective regeneration phenotype observed in Cd34-/ mice. Accordingly, we tested the ability of WT and Cd34-/- MPCs to differentiate and form syncytial myotubes in vitro. Sorted cells were expanded under high serum, growth conditions for 5 days and then exposed to low serum, differentiation conditions for another 5 days. Multinucleated, MyHC+ cells were  106  readily observed in both WT and Cd34-/- samples. Fusion index calculations (percent of myonuclei over total nuclei counted), a standard measure of differentiation efficiency, confirmed that Cd34-/- MPCs differentiate in vitro as efficiently as WT (Figure 3.16). These results are not unexpected, given that CD34 disappears from the cell surface prior to the appearance of mature myofibres, suggesting it would have little impact on differentiation.  Figure 3.16 Cd34-/- MPCs can generate multinucleated myotubes and is dispensible for myogenic differentiation in vitro. (A) Representative images showing WT and Cd34-/- differentiated MPCs forming multinucleated myotubes (MyHC, green; Hoechst, blue). (B) Fusion index (percent of total nuclei found in myotubes) for WT and Cd34-/myotubes. Error bars represent ± SEM for n=3 mice with 15 random fields of view per animal. Results are from 2 independent experiments.  107  3.6  Summary  Our results presented in this chapter demonstrate that, for the first time, CD34 is necessary for efficient muscle regeneration in both acute and chronic damage models. The inefficient myofibre hypertrophy observed in Cd34-/- animals during regeneration can be attributed specifically to a cell autonomous defect of myogenic progenitors lacking CD34, as shown by the defective engraftment of transplanted Cd34-/- MPCs. Moreover, experiments comparing WT and Cd34-/- MPCs further demonstrate that CD34 is necessary for efficient proliferation of MPCs, but not for differentiation. Altogether, these results provide strong support to our hypothesis that CD34 is important on MPCs during the early stages of adult muscle regeneration in vivo, specifically in proliferation.  Although bulk isolations of MPCs allow us to assess myogenic progenitors in vivo, we cannot fully assess the entire myogenic progression of satellite cells since the FACS isolation process most likely activates the satellite cells already. Thus, in the following chapter, we use the single fibre isolation model to complement our studies with MPCs.  3.7  3.7.1  Materials and methods  Mice  Animals were housed in the main animal facility of the Biomedical Research Centre in the University of British Columbia. Mice were kept under sterile conditions and handled following guidelines approved by the University of British Columbia’s Animal Care 108  Committee. Cd34-/- mice were kindly provided by Dr. Kelly M. McNagny (University of British Columbia). Cd34-/- mice were then crossed onto the GFP+CD45.2 background to obtain CD34-/-GFP+CD45.2 mice. LacZ in the Z/AP mice and EGFP expression in the GFP+CD45.2 C56BL/6 mice are both under the control of cytomegalovirus enhancer-chicken beta–actin hybrid promoter. These strains were used as WT controls. The Z/AP and GFP+CD45.2 mice were generously provided by Dr. Corrinne Lobe (MaRS Centre) and Dr. Irving I. Weissman (Stanford University), respectively. Mdx mice contain a point mutation in the dystrophin gene yielding complete absence of the protein. Myf5LacZ animals express the beta-galactosidase gene under the control of the Myf5 promoter. Both these mice were kindly provided by Dr. Michael Rudnicki (Ottawa Health Research Institute). Mice genotypes for Cd34-/-, mdx, GFP+, and LacZ+ were determined by PCR, fluorescence microscopy, or betagalactosidase activity using X-gal. C56Bl/6 mice were used as WT, Ly5.2, controls, unless otherwise indicated. All mice were bred in-house.  3.7.2  Acute damage with NTX  To induce acute damage, 10 µL of notexin (Latoxan# L8104, 10 µg/mL) was injected in TA muscle using a 3/10 CC insulin syringe. WT and Cd34-/- mice were age and sex-matched accordingly. Mice were sacrificed and muscles were harvested at days 5, 7, 10, 14, 21 postnotexin damage. Muscles were paraffin embedded and serially sectioned at 5 µm. Slides were H&E stained following standard procedures. Paraffin embedded tissues were sectioned and stained by Wax-it Histology Services, Inc. (University of British Columbia).  109  3.7.3  Cross-sectional area measurements  Area measurements were performed using images taken from H&E stained slides of crosssectioned muscle. Images were taken with a light microscope (Zeiss Axioplan2 Imaging) and measurements done on density slices calculated with OpenLabTM software (version 4.0.4). All measured fibres were verified to ensure measurements were done on individual fibres. Regenerating fibres were defined as CNFs, having centrally located nuclei. Non-regenerating fibres were defined as fibres with peripherally located nuclei.  3.7.4  Preparation of skeletal muscle tissue for flow cytometry/FACS  FACS isolation of myogenic progenitors was performed as previously published in Joe, et al., 2010. Briefly, primary adult murine myogenic progenitors were obtained as follows: whole hind-limb or TA muscles were carefully harvested from adult mice (6-12 weeks of age) and finely minced into small pieces using forceps. Muscles then underwent a 2-step enzymatic digestion with 0.2% collagenase type II (Roche# C6885) for 30 minutes followed by collagenase D (Roche# 1088882, 1.5 U/mL) and dispase type II (Roche# 295825, 2.4 U/mL) at 37OC for 1 hour. The homogenized muscle samples were then filtered through a 40 µm cell strainer using PBS and the cell suspension is then stained with antibodies against CD45, CD31, Sca1, and alpha7 integrin. Hoechst and propidium iodide were also added. Gating for the sorted MPC population was performed as follows: Hoechst+, PI-, CD45-, CD31-, Sca1-, and alpha7 integrin+. All cell surface staining was done on ice and isotype controls were used to determine gating. Antibodies to CD34 were used when needed.  110  3.7.5  Isolation of MPCs and analysis using flow cytometry  Cells were sorted on a BD FACSVantage SETM machine using BD FACSDivaTM version 4.0.1.2 software. Purity checks were done following the sort to ensure sorting efficiency and accuracy. Analysis of samples was performed using FlowJo (version 8.7)  3.7.6  Cytospin  Freshly sorted MPCs were cytospun onto non-coated, sterile slides at 750 rpm for 5 minutes. Slides were then allowed to dry for 10 minutes and then fixed with 2% PFA for 2 minutes. Standard LacZ detection using X-gal was then performed.  3.7.7  Transplantation and engraftment  Freshly sorted myogenic progenitors were obtained from Z/AP+, Cd34-/-GFP+CD45.2, and GFP+CD45.2 adult mice (8-12 weeks of age). 20,000 GFP+ cells from either WT or Cd34-/animals were mixed with 40,000 LacZ+ cells in PBS. Mixed cells were then injected into the TA muscles of adult WT mice in a 20 µL volume of PBS using a 3/10 CC insulin syringe. Recipient mice were sacrificed 3 weeks post-transplant and perfused with PBS + 10mM EDTA followed by 4% paraformaldehyde. Lower limb muscles were harvested and left overnight in 20% sucrose at 4OC. Muscle tissues were then embedded in cryomolds with OCT and stored at -80OC. Serial sections of 20 µm were analyzed for engraftment of GFP+  111  and LacZ+ cells. A ratio was then obtained by taking the maximum number of GFP+ fibres and dividing it by the maximum number of LacZ+ fibres on an adjacent section.  3.7.8  BM transplantation and NTX injection  Donor BM cells were harvested from WT or Cd34-/- animals and injected intravenously (i.v.) into lethally irradiated WT recipients. The hematopoietic contribution of donor cells was assessed by flow cytometry and only those mice with >85% chimerism were used for analysis. NTX damage was induced 16 weeks post-BM transplant and muscles were harvested and processed for H&E staining 3 weeks post-NTX damage.  3.7.9  BrdU analysis  Following NTX damage, mice were intraperitoneally (i.p.) injected twice daily with 200 µL BrdU (10 mg/mL) for the first 10 days following muscle damage and once daily after day 10. BrdU (0.8 mgl/mL) was also added to the drinking water and changed daily. All mice were sacrificed and harvested on the same day to ensure consistency among samples. Muscle tissues were processed and stained as per our normal MPC isolation protocol. Subsequent permeabilization and denaturation with 0.1% saponin and DNAse (300 µg/mL) treatment were performed followed by addition of anti-BrdU-FITC antibody. All data samples were collected by flow cytometry using a BD FACSLSRIITM machine and BD FACSDivaTM version 4.0.1.2 software. Analysis of samples was performed using FlowJo (version 8.7).  112  NTX damaged TAs from mice that did not receive BrdU were used as controls. Individual TAs were maintained as separate samples throughout the experiment.  3.7.10 MPC culture: growth and differentiation  Freshly sorted MPCs were obtained from WT animals (8-12 weeks of age) and immediately placed into tissue culture treated wells, coated with MatrigelTM (BD# 356235) or collagen (Sigma# C2249). Cells were maintained under growth conditions (DMEM + 20% FBS + 10% horse serum + 1% P/S + 0.1 mM bFGF) for 5 days following initial isolation and another 5 days under differentiation conditions (DMEM + 2% horse serum + 1% P/S).  3.7.11 Fusion index calculations  WT and Cd34-/- MPCs were obtained from individual animals and kept separate throughout the duration of the experiment. Each mouse was split into 3 separate wells and cultured as described above. After 10 total days in culture, all wells were fixed with 4% PFA and stained with MyHC and Hoechst. The fusion index was calculated by counting the number of nuclei detected within myotubes, which are defined as MyHC+ structures containing 2 or more nuclei, and dividing by the total number of nuclei counted within a given field. 15 random fields of view were counted.  113  3.7.12 Statistical analysis  Student’s two-tailed t-test was used on all statistical analyses performed between groups. Statistical significance was set at p % 0.05.  114  Chapter 4: CD34 is necessary for progression of satellite cells through the myogenic program, specifically in activation, migration, and proliferation.  4.1  Introduction and rationale  The results described in Chapter 3 show CD34’s necessity in muscle regeneration, specifically, and at least, in MPC proliferation following damage. In this chapter, we focus on having a more detailed look at the discrete stages that occur early in the myogenic program. Contrary to bulk sorted MPCs, we wanted to have a model that allows us to look at individual satellite cells. For this, we performed single fibre isolations. In this model, satellite cells are isolated within their niche and the culture process allows them to activate and progress through the myogenic program (Rosenblatt, et al., 1995). Single myofibres cultures can be used to model myogenesis as satellite cells at specific stages can be identified using molecular markers such as Pax7 and MyoD (Zammit, et al., 2006).  By comparing WT and Cd34-/- satellite cells in these single fibres, we observed that Cd34-/satellite cells are delayed in progression through the myogenic program. More specifically, this delay initiates at the stage of activation, where Pax7 is down-regulated. Notably, p38 MAPK pathway activation, a proposed molecular switch governing satellite cell entry from quiescence to activation (Jones, et al., 2005), is significantly decreased and delayed in Cd34-/MPCs and satellite cells. Moreover, satellite cells lacking CD34 are also delayed in entry into cell cycle and have decreased overall expansion, consistent with results obtained using sorted MPCs. However, CD34 does not appear to be important for maintenance of satellite cell  115  numbers at homeostatic conditions. Interestingly, video time-lapse imaging of satellite cell movement on host myofibres show that Cd34-/- satellite cell motilty is significantly impaired, which is further confirmed in vivo, using a freeze damage model.  Overall, the results presented here are consistent with those obtained from MPCs, in that CD34 is necessary for efficient muscle regeneration and satellite cell function. More importantly, these data suggest that CD34 is important specifically in satellite cell activation, migration, and proliferation – roles that are well in-line with our hypothesis and previously proposed functions for CD34 in other cell types.  4.2  CD34 is dispensable for maintenance of satellite cell numbers during homeostasis  Differential amounts of quiescent satellite cells between WT and Cd34-/- muscle during homeostatic conditions could account for the decreased MPC proliferation and overall defect in myofibre hypertrophy during regeneration of Cd34-/- animals. To determine if Cd34-/muscle have decreased satellite cell numbers, single myofibres were isolated from EDL muscles of WT and Cd34-/- animals. As shown in Figure 4.1, Pax7+ satellite cells are readily detectable in both groups (Figure 4.1).  116  Figure 4.1 Pax7 readily identifies satellite cells in WT and Cd34-/- isolated myofibres. Immunofluorescent detection of satellite cells on WT and Cd34-/- single fibers (Pax7, red; Hoechst, blue).  Quantification of Pax7+ satellite cells immediately after fibre isolation showed no significant difference in numbers between WT and Cd34-/- fibres (Figure 4.2), indicating that CD34 is dispensable for the maintenance of quiescent satellite cell numbers during homeostasis, and also likely during embryonic muscle development.  Figure 4.2 Comparable satellite cell numbers between WT and Cd34-/- single fibres immediately after isolation. Direct enumeration of satellite cells on WT and Cd34-/- single fibers immediately following isolation. Error bars represent ± standard deviation for n=5-7 animals. Results are from 2 independent experiments.  117  4.3  CD34 is necessary for satellite cells to progress through the myogenic program on  cultured single fibres  Another potential explanation for the observed reduction in proliferating MPCs could be a delay in satellite cell activation. We thus examined the progression of satellite cells through the myogenic program by assessing Pax7 and MyoD expression. Pax7 is highly expressed in quiescent satellite cells; but, upon entry into cycle, these cells initiate expression of MyoD and eventually down-regulate Pax7 as they commit to differentiation (Cornelison & Wold, 1997; Zammit, et al., 2006). With this, 2 myogenic progenitor populations can then be identified: activated satellite cells (Pax7+ MyoD+) and differentiation-committed myoblasts (Pax7- MyoD+) (Figure 4.3).  Figure 4.3 Immunofluorescent imaging of Pax7 and MyoD staining on single fibres identifies activated satellite cells and differentiation-committed myoblasts. Combined Pax7 and MyoD staining on single fibers identify two subpopulations of myogenic progenitors: Pax7+ MyoD+ as activated satellite cells and Pax7- MyoD+ as differentiation-committed myoblasts (Pax7, red; MyoD, green; Hoechst, blue).  118  Analysis of these 2 populations over time in WT and Cd34-/- single fibre cultures revealed no significant difference in the frequency of Pax7+ cells that activate MyoD expression. After 48 hours of culture, the differentiation-committed myoblast population becomes evident in WT cultures; however, at this timepoint, this population is absent in Cd34-/- fibres. At the last timepoint analyzed, 72 hours in culture, although differentiation-committed myoblasts are now readily detectable on Cd34-/- fibres, their frequency is still significantly lower than on WT fibres (Figure 4.4 and summarized in Table 4.1). Thus, the lack of CD34 on satellite cells results in a significant delay in their progression along the myogenic program.  Figure 4.4 Cd34-/- satellite cells have delayed activation and progression through the myogenic program. Quantification of activated satellite cells and differentiation committed myoblasts on WT and Cd34-/- cultured fibers. Error bars represent ± standard deviation for n=3-7 animals per timepoint. Results are from 2 independent experiments.  119  Table 4.1  Statistical analysis of the 2 myogenic progenitor populations on WT and Cd34-/- myofibres  during culture.  Work by Jones and colleagues supports a requirement for p38 MAPK phosphorylation in satellite cell activation (Jones, et al., 2005). Thus, we investigated phospho-p38 MAPK (pp38) expression in satellite cells on single fibres. Analysis of freshly isolated fibres showed comparable pp38+ satellite cells on WT and Cd34-/-. However, further culture revealed that within 12 hours, over 80% of WT satellite cells had activated this pathway, while Cd34-/samples required 48 hours to reach comparable levels (Figure 4.5).  Figure 4.5. WT and Cd34-/- satellite cells have delayed p38 MAPK activation during single fibre culture Analysis of pp38 expression on Pax7+ satellite cells on cultured WT and Cd34-/- single fibers over time. Bar graphs represent the frequency of pp38+ satellite cells (n=11-87 satellite cells). Results are from one experiment.  120  A similar analysis and quantification by Western blot for pp38 using freshly sorted WT and Cd34-/- MPCs revealed significantly lower levels of pp38 in Cd34-/- MPCs sorted from undamaged muscle, but not in whole muscle extracts (Figure 4.6), providing further confirmation that p38 MAPK activation is impaired in Cd34-/- myogenic cells. Together, these results indicate that the presence of CD34 is required for the efficient activation of adult myogenic cells.  Figure 4.6. Western blot analysis using sorted WT and Cd34-/- MPCs show decreased p38 MAPK activation in Cd34-/- MPCs (A) Western blot analysis of pan- and pp38 MAPK in WT and Cd34-/- whole muscle extracts and sorted MPCs. (B) Quantification of pan-p38 and pp38 from WT and Cd34-/- whole muscle extracts and sorted MPCs. Ratios were normalized to WT controls. Error bars represent ± SEM for n=2. Results are from 2 independent experiments.  121  Collectively, these data suggest that the lack of CD34 on satellite cells results in their significant delay in the initiation of the myogenic program, likely beginning at the activation stage. Next, we examined whether their proliferation is also similarly delayed.  4.4  Satellite cells lacking CD34 are delayed in cell cycle entry and overall  proliferation  Proliferation can be assessed in single fibre cultures by enumerating all myogenic cells, defined here as Pax7+ and/or MyoD+ expressing cells, on the fibres at each timepoint. On WT fibres, an increase in the total number of myogenic cells is clearly observed at 48 hours following culture initiation. In contrast, no increase in is detected on Cd34-/- fibres until 72 hours following culture. At both timepoints, WT fibres harbor significantly more myogenic cells than Cd34-/- fibres (Figure 4.7).  Figure 4.7 Cd34-/- satellite cells do not expand efficiently on cultured single fibres. Total myogenic cells counted per fiber. Error bars represent ± SEM for n=3-7 animals per timepoint. Results are from 2 independent experiments.  122  Because these differences were not present initially, these results indicate a striking reduction in the ability of Cd34-/- satellite cells to expand. It is noteworthy that Cd34-/- myogenic progenitors can and do proliferate, as can be seen by the sharp increase in average cells per fibre between 48–72 hours in culture. However, this proliferation is delayed when compared to WT fibres, whose associated myogenic progenitors numbers begin to increase between 24–48 hours in culture (Figure 4.7).  Our results from single fibre cultures suggest that Cd34-/- satellite cells may be delayed in initiating proliferation. As a further confirmation of these results, we performed 3D video time-lapse imaging of cultured single fibres (Siegel, et al., 2009), allowing us to directly observe satellite cell divisions in real time. In agreement with our previous results, we observed fewer satellite cell divisions on Cd34-/- fibres from 24 to 48 hours after myofibre harvest compared to WT controls (Figure 4.8), reflecting a smaller percentage of Cd34-/- cells entering the first cell cycle (44.2 vs. 20.6%). Moreover, this analysis revealed a significant delay in the first division of Cd34-/- satellite cells (Figure 4.9).  123  Figure 4.8 Cd34-/- satellite cells divide significantly less than WT controls. Number of satellite cell divisions detected using time-lapse microscopy between 24 and 48 hours after fiber culture initiation. Error bars represent ± SEM for n=42-107 satellite cells. Results are from 5 independent experiments.  Figure 4.9 Cd34-/- satellite cells undergo cell division significantly later than WT controls. Timing of the first division of individual satellite cells. Line represents the mean value for n=19-22 satellite cells. Results are from 5 independent experiments.  124  The lack of CD34 resulted in the overall delay of progression through myogenesis. The data presented here not only correspond well with the decreased proliferation seen through BrdU analysis in Cd34-/- MPCs during in vivo regeneration, but also show that the overall delay in Cd34-/- animals begins at the stage of satellite cell activation and follows through to the first cell division and subsequent expansion of myogenic cells.  4.5  CD34 is necessary for efficient satellite cell migration  It has been previously proposed that satellite cells migrate during muscle regeneration (Otto, et al., 2011; Phillips, et al., 1990; Schultz, et al., 1985). In general, efficient migration requires proper adhesive interactions with the extracellular milieu mediated through a number of specialized molecules present on the cell surface (Palecek, Loftus, Ginsberg, Lauffenburger, & Horwitz, 1997). Because CD34 has been proposed to promote the efficient migration of hematopoetic cells through its anti-adhesive functions (Blanchet, et al., 2007; Drew, Huettner, et al., 2005; Nielsen & McNagny, 2008) and since satellite cells have been shown to undergo extensive movement and migratory behavior on single fibres during culture (Otto, et al., 2011; Siegel, et al., 2009), we extended our characterization of Cd34-/satellite cells by evaluating their total motility on a native substrate, the myofibre.  Live imaging of WT and Cd34-/- isolated single fibres was initiated 24 hours after fibre isolation and continued for an additional 24 hours (Siegel, et al., 2009). During this period, both WT and Cd34-/- cells were found to be actively motile, but WT cells exhibited  125  substantially more movement than Cd34-/- cells. An example of direct satellite cell tracking is shown in Figure 4.10.  Figure 4.10 Representative images showing tracking of WT and Cd34-/- satellite cell movement on cultured single fibres. Representative images of WT and Cd34-/- satellite cell tracking on single fibers based on time-lapse microscopy imaging. Each color represents a different cell tracked.  The quantification of individual satellite cell movements show dramatic and significant reductions in average speed (Figure 4.11) and overall distance traveled (Figure 4.12) by Cd34-/- satellite cells in comparison to WT controls.  126  Figure 4.11 Cd34-/- satellite cells move significantly slower than WT controls. WT and Cd34-/- satellite cell velocities as determined using time-lapse microscopy. Error bars represent ± SEM for n=42-107 satellite cells. Results are from 5 independent experiments.  Figure 4.12 Cd34-/- satellite cells travel less overall distance compared to WT controls. Total distances traveled for WT and Cd34-/- satellite cell were determined. Error bars represent ± SEM for n=42-107 satellite cells. Results are from 5 independent experiments.  Furthermore, evaluation of the instantaneous velocities of WT and Cd34-/- individual cells provides direct evidence that WT cells move faster throughout this period, suggesting that the decreased motility of Cd34-/- cells is not merely due to a delay in initiating movement, but  127  rather to a defect in migration (Figure 4.13). Overall, these data support a key role for CD34 in promoting efficient satellite cell movement.  Figure 4.13 Instantaneous velocity measurements of WT and Cd34-/- satellite cells on cultured fibres. Measurement of frame-by-frame instantaneous velocities for individual WT and Cd34-/- satellite cells. Results are from 5 independent experiments.  To test if impaired migration of Cd34-/- myogenic progenitors also occurs in vivo, we performed freeze damage injury to WT and Cd34-/- TA muscles of adult mice. In this model, a liquid-nitrogen cooled rod is placed directly on the TA muscle. Extensive damage and necrosis occurs at site of lesion, whereas regenerating and non-regenerating fibres can be found distal to this region (Gayraud-Morel, et al., 2007). In vivo, satellite cells migrate from the viable area to the site of regeneration (Phillips, et al., 1990; Schultz, et al., 1985). So, given this, if Cd34-/- satellite cells do have a defect in migration during muscle regeneration, it should be evident by relatively increased amount of damaged tissue remaining in Cd34-/muscles compared to WT controls. At 7 days following freeze injury, WT and Cd34-/muscles were harvested and H&E stained. Measurements of the distinct area closest to the 128  site of lesion, where the most damage was done, showed that Cd34-/- animals have a significantly larger damaged area remaining when compared to WT controls (Figure 4.14). This demonstrates that in vivo, Cd34-/- satellite cells are also impaired in migration.  Figure 4.14 Increased necrotic area in Cd34-/- skeletal muscle following freeze damage. (A) H&E staining of WT and Cd34-/- muscle sections at day 7 following freeze damage. (B) The total necrotic area resulting from freeze damage is plotted individually for WT and Cd34-/- groups. Line represents the mean value of total for n=5-6 mice. Results are from 2 independent experiments.  Lastly, because one of the earliest motility-associated phenomena that take place during satellite cell activation is their exit from the niche beneath the basement membrane, we tested  129  whether this process is also delayed in Cd34-/-mice. Single fibres from WT and Cd34-/- were harvested and fixed at 0, 24, 48, and 72 hours following culture and stained for Pax7 and laminin. The position of individual Pax7+ satellite cells on cultured fibres relative to the laminin+ basement membrane was then assessed by confocal microscopy. We observed no difference in the proportion of WT and Cd34-/- cells located above or below the basement membrane, indicating that the inefficient motility of Cd34-/- satellite cells does not affect their ability to exit from the niche (Figure 4.15).  130  Figure 4.15 Cd34-/- satellite cells efficiently exit their niche. WT and Cd34-/- single fibers were cultured and harvested at 0, 24, 48, and 72 hours post-culture. Immunofluorescent staining for satellite cells (Pax7) and the basal lamina (laminin) was performed and the location of satellite cells relative to the basal lamina was determined (Pax7, red; laminin, green). Representative images are shown. Bar graphs on the right show the relative proportion of satellite cells below or above the basal lamina in WT and Cd34-/- groups, n=3 animals. Results are from one experiment.  131  4.6  Summary  Although an in vitro model system, cultured single fibres allow us to directly assess satellite cells starting from a quiescent state, when they are still within their niche, and follow their progression through the myogenic program. Using this model, our results demonstrate that CD34, despite being dispensable for maintenance of satellite cell numbers during homeostatic conditions, is required for efficient satellite cell activation, migration, and proliferation. Defects in these stages of the myogenic program correspond well with the results from sorted MPCs presented in Chapter 3 and the overall defective regeneration observed in Cd34-/- animals upon damage.  4.7  4.7.1  Materials and methods  Mice  Animals were housed in the main animal facility of the Biomedical Research Centre in the University of British Columbia. Mice were kept under sterile conditions and handled following guidelines approved by the University of British Columbia’s Animal Care Committee. Cd34-/- mice were kindly provided by Dr. Kelly M. McNagny (University of British Columbia) and genotypes determined by PCR. C56Bl/6 mice were used as WT, Ly5.2, controls, unless otherwise indicated. All mice were bred in-house.  132  4.7.2  Single fibre isolation and culture  Single fibre isolations were performed as per standard protocol. Briefly, the EDL muscle was gently harvested following sacrifice of the mouse and care was taken to handle only the tendons. Collagenase I (Worthington# LS004197, 400 U/mL) digestion for approximately 1 hour in 37OC was performed to obtain live, single fibres. Fibres were then cultured in Ham’s F-12 complete media + 15% horse serum + 0.5nM bFGF + 1% P/S, harvested, and fixed with 4% PFA at specific timepoints.  4.7.3  3D video time-lapse imaging  Real-time video imaging and analysis was performed on WT and Cd34-/- single fibre cultures as initially described in Siegel, et al., 2010 (Siegel, et al., 2009).  4.7.4  p38 MAPK detection by Western blot and immunofluorescence  Western blot procedures were performed as described in Perdiguero, et al., 2007. Pan- and phospho-p38 MAPK (Cell signaling# 9212 and 9216, respectively) were probed on the same blot with fluorescently-labeled secondary antibodies. The Li-COR Odyssey was used to determine relative quantification of the bands. The same primary antibodies were used to detect pan- and phospho-p38 MAPK on isolated single fibres.  133  4.7.5  Immunofluorescence  Following blocking using blocking buffer (25% normal goat serum, 3% BSA, 0.3% TritonX), immunofluoresecent staining on single fibres was done using antibodies to Pax7 (DSHB), MyoD (clone C20, Santa Cruz# sc-304), and laminin (abcam# ab11575) diluted in 0.3% TritonX. Analysis of Pax7 and/or MyoD myogenic progenitors was done with a (Zeiss Axioplan2 Imaging microscope. Analysis was of satellite cell location relative to the niche was done by confocal microscopy (Nikon C1 laser scanning confocal microscope).  4.7.6  Freeze damage injury  All experimental mice were anaesthetized with isofluorane and a small incision of the skin was made above the TA muscle. Freeze injury was caused by a liquid-nitrogen cooled copper rod placed lightly on top of the exposed TA muscle for 10 seconds. This was repeated for a total of 3 rounds of freezing on both legs. The incision closed using staples. Mice were carefully monitored for health status and harvested at the timepoints indicated.  4.7.7  Statistical analysis  Student’s two-tailed t-test was used on all statistical analyses performed between groups. The only exception to this was the statistical analysis used to test the significance between phospho-p38 MAPK presence on WT vs. Cd34-/- satellite cells on fibres. In this case, the Fisher exact test (F-test) was used. Statistical significance was set at p ! 0.05.  134  Chapter 5: Conclusion  5.1  Data summary  The goal of this thesis was to define a role for CD34 in adult muscle regeneration. In the work presented here, we have done this in 3 steps. In Chapter 2, we present an optimized flow cytometry-based method to prospectively isolate MPCs from adult skeletal muscle. Although we demonstrate that CD34 is an excellent marker for MPCs from undamaged muscle, we have also verified alpha7 integrin as an alternative marker since CD34 expression has been previously reported to be down-regulated following satellite cell activation in vitro (Beauchamp, et al., 2000). Indeed, analysis of MPCs isolated from various timepoints following acute NTX damage demonstrates that CD34 expression is dynamically regulated early during regeneration. Both qRT-PCR and flow cytometry results show that CD34 expression is significantly decreased shortly after damage, demonstrating that this regulation occurs both at the transcript and protein level. Interestingly, CD34 isoform expression also varies in the same timeframe. Under non-damaged conditions, MPCs exclusively express the CD34FL. Upon injury, these cells express both CD34FL and CD34CT isoforms. By day 7 postdamage, when regeneration is well underway, MPCs once again exclusively express CD34FL. Overall, this chapter demonstrates that CD34 expression is regulated in vivo during the early stages muscle regeneration, suggesting that CD34 likely plays a role during satellite cell quiescence, activation, migration, and proliferation.  135  In Chapter 3, we assessed the regenerative capacity of WT and Cd34-/- animals in both acute and chronic damage models. Elevated levels of necrosis and impaired hypertrophy in Cd34-/mice show that CD34 is necessary for efficient muscle regeneration. Defective engraftment of Cd34-/- MPCs indicate that the overall muscle regeneration defect in Cd34-/- animals can be specifically attributed to impaired myogenic precursors. Although in vitro fusion index assays show that CD34 is dispensable for myogenic differentiation, in vivo BrdU incorporation assays demonstrate that CD34 is necessary for MPC proliferation following damage. A similar analysis of FAPs, another CD34+ cell population found in muscle, did not show any regulated CD34 expression on WT FAPs nor any difference in proliferation between WT and Cd34-/- FAPs, providing further evidence that CD34 is specifically important for MPC function. Together, these data are consistent with our hypothesis and show that CD34 is specifically required for MPC function, particularly during the proliferative stage of muscle regeneration.  Lastly, in Chapter 4, we use single fibre cultures to model adult myogenesis as a whole, beginning with quiescence and following through the rest of the myogenic program (activation, migration, proliferation). Here, we demonstrate that CD34 is not necessary for maintenance of quiescent satellite cells under homeostatic conditions, but its lack on satellite cells results in delayed myogenic progression beginning with activation and subsequently results in impaired migration and myoblast expansion. In summary, these experiments enabled us to identify which stages of regeneration CD34 is important in with regards to satellite cell function.  136  Together, our work demonstrates that: (1) CD34 is expressed on satellite cells and MPCs. This expression is regulated during the early stages of adult myogenesis, both in vitro and in vivo. (2) CD34 is necessary for MPC function and thus to promote efficient muscle regeneration following acute and chronic damage. (3) CD34 is specifically necessary for the early stages of satellite cell activation, migration, and proliferation.  5.2  General discussion  CD34 is a well-known surface marker used to isolate various progenitor cells, yet is also notorious for leaving researchers questioning its exact functional role. Previous reports speculate that CD34 may function as a homing receptor, a blocker of differentiation, a proadhesive receptor, or, conversely, an anti-adhesion molecule (Furness & McNagny, 2006; Nielsen & McNagny, 2008). Efforts to directly reveal a function for CD34 have been hampered by the fact that CD34 is one member of a functionally redundant family of three sialomucins (CD34, podocalyxin, endoglycan) with an overlapping tissue distribution (Doyonnas, et al., 2001; Nielsen & McNagny, 2008; Sassetti, et al., 2000). This may, in part, explain why Cd34-/- mice show no obvious defects in tissues where its homologues are expressed. Recent studies, however, showed that phenotypes in these mice can be revealed when a specific system is challenged with damage or infection, thus leading to more precise hypotheses of CD34’s function (Blanchet, et al., 2010; Blanchet, et al., 2007; Grassl, et al., 2010; Strilic, et al., 2009). In our work, we evaluated muscle regeneration in adult Cd34-/-  137  animals and observed a clear defect in both acute and chronic damage models caused by functional impairment of Cd34-/- satellite cells in activation, migration, and proliferation. Below, we discuss our findings in relation to our current understanding of CD34 in muscle and other systems.  5.2.1  CD34 mediates satellite cell activation and subsequent myoblast proliferation  during myogenic progression  Our data demonstrates that CD34 is required for MPCs and satellite cells to progress through activation, defined here as the period spanning from the onset of damage to the first satellite cell division. Our work provides several lines of evidence to support this notion. Satellite cells on cultured single fibres have significantly delayed progression through the myogenic program. Upon investigation of the different stages of myogenesis, we find that quiescent satellite cell numbers are not significantly different between WT and Cd34-/- animals. However, significantly decreased myogenic progenitor cell expansion occurs in Cd34-/- fibre cultures, indicating that the activation step that occurs between quiescence and cell proliferation is delayed or impaired. This is further supported by analysis of satellite cell expression of Pax7 and/or MyoD, showing slower appearance of differentiation-committed myoblast population in Cd34-/- cells associated with the down-regulation of Pax7. Moreover, p38 MAPK activation, previously proposed to act as a molecular switch from satellite cell quiescence to activation (Jones, et al., 2005), is also decreased in Cd34-/- MPCs and is delayed in satellite cells during single fibre culture. Overall, this impaired activation due to the lack of CD34 results in delayed cell cycle entry and significantly decreased myoblast  138  expansion, which correlates well with significantly smaller myofibre sizes (impaired hypertrophy) observed in Cd34-/-animals during acute or chronic damage models of muscle regeneration. Notably, in the chronic damage model, this defect was no longer apparent in aged, 18-month old animals. However, since impaired regeneration in mdx mice has been attributed to satellite cell exhaustion (Luz, Marques, & Santo Neto, 2002; Sacco, et al., 2010), the defects in satellite cell function caused by the lack of CD34 could be masked in aged animals.  The proposition that CD34 may be involved in activation is not unique to satellite cells. Sato et al. originally characterized that CD34 expression is associated with a reversible, activated state of hematopoietic stem cells (Sato, et al., 1999). In support of a role for CD34 in facilitating the entry of quiescent cells into proliferation, Trempus et al. reported that Cd34-/animals failed to develop papillomas upon DMBA/TPA induction (Trempus, et al., 2007). A detailed analysis of the cell cycle state of hair follicles shows that Cd34-/- follicles retain BrdU and remain in a resting phase, as opposed to an actively growing phase, demonstrating that Cd34-/- cells do not efficiently undergo activation. Our work presented here adds further evidence that supports a role for CD34 as a mediator of activation.  Studies comparing CD34+ and CD34- HSC subsets show a higher percentage of proliferating cells in the CD34+ population (Shman, et al., 2007) providing a link between CD34 and proliferation. It is possible that CD34 is present on activated cells to allow for interaction with binding partners to mediate downstream extra- or intra-cellular signal transduction that can regulate entry to cell proliferation, the stage following activation. In terms of muscle  139  regeneration, once satellite cells pass activation and continue through proliferation, myogenic cells down-regulate CD34 expression, as seen with our MPCs during regeneration, and proceed with differentiation, which requires the prior cessation of proliferation.  5.2.2  CD34 facilitates migration of myogenic progenitors  Another proposed role for CD34 is to mediate cell motility and migration. The relationship between satellite cell movement and proliferation remains debatable, with some suggesting movement follows division (Kuang, et al., 2007), while our data further supports the notion that proliferation can proceed despite blocked movement (Siegel, et al., 2009). Evidence that both proliferation and motility are affected in Cd34-/- satellite cells suggests that a functional link between the two may exist. But, it is also possible that CD34 plays independent roles in each of these processes. Interestingly, links between CD34 and both proliferation and motility have been independently established in other systems. For example, Cd34-/- mast cells and eosinophils display defective inflammatory migration, an effect that has been ascribed to increased adhesion caused by the loss of CD34 (Blanchet, et al., 2007; Drew, Huettner, et al., 2005).  Unfortunately, the mechanism by which CD34 regulates cell motility and proliferation is unknown. Two key pieces of data presented in this work could provide some initial insights into possible mechanistic pathways: (1) Regulated CD34 expression during muscle regeneration. (2) Decreased p38 MAPK activation in Cd34-/- myogenic progenitors (discussed later). The changes in isoform expression on MPCs during various stages of  140  regeneration (from exclusively CD34FL to both CD34FL and CD34CT and then back to just CD34FL) may indicate that the cytoplasmic tail is not as important during activation, migration, and proliferation. Alternatively, their concurrent expression during activation, migration, and proliferation could also indicate that dual and independent roles for each isoform could exist during these stages. The functional difference, if any, between CD34FL and CD34CT is still debatable. Drew et al. show that the homotypic aggregation of Cd34-/mast cells can be abolished by re-expression of either CD34 isoform; curiously, CD34CT appears to do so more efficiently (Drew, Merzaban, et al., 2005). The authors propose that the cytoplasmic domain present in CD34FL allows it to be more easily cleared from the surface by intracellular ligands. Since CD34CT lacks this intracellular interaction, its surface expression is better retained, thus allowing it to perform its anti-adhesive function more efficiently. Moreover, since over-expression of CD34FL, but not CD34CT, has been shown to inhibit M1 cell line differentiation, the cytoplasmic domain may be linked to signals that prevent differentiation (Fackler, et al., 1995).  During muscle regeneration, CD34 may behave in a similar fashion: (1) During homeostatic conditions, CD34FL interacts with intracellular binding partners in satellite cells, allowing them to maintain a quiescent state and prevent premature differentiation. (2) Upon activation and proliferation, MPCs express both CD34FL and CD34CT, with CD34FL providing signals to still inhibit differentiation, while anti-adhesive properties of CD34CT allow for MPC migration to facilitate activation and proliferation. (3) During differentiation, when MPCs have reached the site of damage, CD34 is significantly down-regulated, allowing for efficient myoblast fusion and myofibre repair. The proposal above does not exclude a possible active  141  role for CD34FL in mediating MPC motility. Podocalyxin, a CD34-family member, has been shown to modulate cell morphology, polarization, and adhesion of cancer cell lines (Somasiri, et al., 2004), all critical aspects of overall cell movement. Podocalyxin’s intracellular binding partners, NHERF-1 and NHERF-2 (Tan, et al., 2006), contain PDZ and ERM binding domains that link Podocalyxin to the actin-cytoskeleton and likely contributes to a mechanistic role (Furness & McNagny, 2006; Nielsen & McNagny, 2008). By analogy, CD34FL could also similarly mediate MPC motility during muscle regeneration through intracellular interactions.  5.2.3  CD34 and signal transduction  Although we presented the decreased and delayed p38 MAPK activation in Cd34-/- myogenic cells as a read-out of inefficient activation, it is also one of the few times that CD34 has been implicated in a signaling pathway. Whether the delayed p38 MAPK activation is causally linked with the delayed activation and initiation of proliferation in Cd34-/- myogenic progenitors or merely a consequence of it is currently unclear, as is the exact relationship between this pathway and CD34. Interestingly, another sialomucin, CD164, has been described to regulate myoblast motility and fusion through modulation of satellite cell responses to the chemokine stromal derived factor-1 (SDF-1) (Bae, et al., 2008). Moreover, it has been shown that CD34+ myogenic cells express higher levels of the SDF-1 receptor, CXCR4, compared to the CD34- population (Ieronimakis, et al., 2010), suggesting a role for CD34 in CXCR4 signaling. However, our preliminary results failed to detect any change in  142  the ability of Cd34-/- MPCs to migrate in response to SDF-1 (Figure 5.1), suggesting that CD34 acts through other mechanisms.  Figure 5.1 Comparable transwell migration of WT and Cd34-/- MPCs. % MPCs that have migrated towards an SDF + HGF or SDF only gradient using a transwell migration assay. Error bars represent ± SEM for n=2 mice. Results are from 2 independent experiments.  The presence of putative phosphorylation sites in the CD34FL cytoplasmic domain suggests that interaction with an intracellular ligand could also link CD34 to signal transduction. CrkL is the only identified intracellular binding partner to CD34 thus far. Although the specific interaction of CD34/CrkL has not been directly linked to any known signaling pathways or downstream interacting partners, Crk signaling has been implicated in regulating cell migration and preventing apoptosis (Cho & Klemke, 2000; Feller, et al., 1998; Stupack, Cho, & Klemke, 2000). Moreover, Crk and CrkL proteins have also been shown to bind Gab1, an adaptor protein known to play a central role in growth and apoptosis, in response to HGF/cmet activation (Furge, Zhang, & Vande Woude, 2000). Together, these provide a potential 143  link between HGF/c-met signaling, which is known to be important in satellite cell activation, proliferation, and motility, and CD34 signaling through CrkL.  5.3  Future directions  Despite much effort by many groups, the exact mechanism by which CD34 performs its functions in various cell types is unknown. Likewise, how CD34 positively regulates satellite cell activation, proliferation, and motility still eludes us. Overall, futher studies are warranted and would be of great interest to many groups.  5.3.1  Contribution of other cell types  Currently, the only knock-out mice available to study CD34 is the complete Cd34-/- mouse (Cheng, et al., 1996; A. Suzuki, et al., 1996). The generation of other CD34 transgenic mice would allow for temporal, cell specific, or conditional deletion of CD34 and further dissection of observed phenotypes in specific models. Moreover, it would allow for more sophisticated in vivo studies that could automatically exclude the contribution of other cell types. Other CD34+ cell types besides satellite cells are also involved in muscle regeneration. For example, hematopoietic cells, FAPs, and endothelial cells express CD34 and participate in muscle regeneration. Although our data shows a cell autonomous defect in Cd34-/- MPCs that contributes to the muscle regeneration defect in Cd34-/- animals, we cannot formally exclude the involvement of defective non-MPC cell types that may also contribute to this phenotype.  144  5.3.1.1  Hematopoietic subsets  Our previous results showed no significant difference in the CSAs of regenerating fibres in WT mice reconstituted with WT or Cd34-/- BM following NTX damage (Figure 3.11), indicating that the lack of CD34 on hematopoietic cells does not significantly affect regeneration. However, a similar analysis of WT and Cd34-/- mice transplanted with WT BM experiment (Figure 5.2) showed no significant difference in regenerating fibres sizes, suggesting that the introduction of WT BM can rescue the regeneration defect normally present in Cd34-/- animals (Figure 3.4).  Figure 5.2 WT BM transplantation to WT and Cd34-/- recipients results in no significant difference in regenerating myofibre sizes. CSA measurements performed on regenerating myofibers of WT and Cd34-/- animals transplanted with WT BM 21 days following NTX damage. Error bars represent ± SEM for n=3 mice with > 200 fibers per animal. Results are from one experiment.  Given that our earlier results (Figure 3.11) show no difference in regenerating myofibre CSAs between WT mice transplanted with WT or Cd34-/- BM, we hypothesize that the full 145  phenotype of defective muscle regeneration in Cd34-/- animals can only be seen in the absence of CD34 from both myogenic and hematopoietic progenitors, suggesting that CD34 plays a functional role in both cell types during regeneration. In addition, the process of lethally irradiating the animals to perform the BM transplants may somehow dampen their regenerative capacity, a caveat that has to also be taken into consideration when interpreting these preliminary results. Clearly, further investigations with more specific BM fractions may provide further insight and would be of great interest.  Mast cells have been hypothesized to be involved in the progression of DMD (Gorospe, Tharp, Demitsu, & Hoffman, 1994; Gorospe, Tharp, Hinckley, Kornegay, & Hoffman, 1994). As CD34 is a marker for mast cells and has been shown to affect mast cell function (Drew, et al., 2002; Drew, Merzaban, et al., 2005), we sought to determine if these cells indeed affected muscle regeneration following acute damage. A comparison of necrotic tissue areas among WT, W/Wv (mast-cell deficient), and W/Wv mice reconstituted with WT mast cells following NTX damage showed no difference in regenerative capacity among the three groups, indicating that mast cells do not play a significant role in muscle repair following acute damage (Figure 5.3).  146  Figure 5.3 Mast cells do not significantly contribute to muscle regeneration following acute damage. Quantification of necrotic areas of damage 5 days after NTX in WT, W/Wv, and W/Wv mice reconstituted with WT mast cells. Error bars represent ± SEM for n=3 mice. This figure is from Jamie Haddon and Kelly M. McNagny (unpublished results). Results are from one experiment.  Since Cd34-/- animals have been reported to have impaired inflammatory cell migration (Blanchet, et al., 2007; Grassl, et al., 2010), a similar analysis of other inflammatory cells would be provide further elucidation as to which hematopioetic cell type CD34 plays a significant role in during muscle regeneration.  5.3.1.2  Fibro-adipogenic progenitors  FAPs have been shown to positively regulate myogenic differentiation (Joe, et al., 2010). Since FAPs express CD34, the muscle regeneration defect observed in Cd34-/- mice could be partly attributed to impaired FAP function. We performed a similar analysis of CD34 expression on FAPs and, unlike with MPCs, found CD34 to be consistently expressed on the 147  surface these cells during regeneration. Moreover, a comparison between WT and Cd34-/FAP proliferation following NTX damage shows no difference between the two groups. Overall, this preliminary data suggests that FAP function is not affected in Cd34-/- animals. However, the exact mechanism by which FAPs influences myogenesis still unknown (Joe, et al., 2010). Thus, a better understanding of the exact role of FAPs in vivo will allow us to design more precise experiments to better evaluate CD34’s role in FAPs.  5.3.1.3  Endothelial cells  Interestingly, work by Blanchet et al. describe an exacerbated phenotype in Cd34-/- animals during autoimmune arthritis, which authors attribute to increased vascular leakage, providing evidence that CD34 is required to maintain vascular integrity (Blanchet, et al., 2010). Given that proper vascularization is necessary for efficient muscle repair (Faulkner, Weiss, & McGeachie, 1983; Grounds, 1987), further evaluation of CD34’s contribution to vascular development and regeneration would be of great interest.  5.3.2 Identification of CD34 binding partners  The identification of both intra- and extra-cellular partners for CD34 would be extremely beneficial in moving current studies from a phase that investigates cell functions that require CD34 to the development of mechanistic pathways that link CD34 to a given cell function. Moreover, it would be of particular interest to compare the similarities and differences in CD34 ligands based on cell types to determine which CD34 functions are general and which 148  are cell-type specific. The identification of binding partners in would be optimal in beginning to dissect a mechanism for CD34.  5.3.3  CD34 in human satellite cells and skeletal muscle  All of the data presented here were generated with murine models. The question of whether CD34 is present on human satellite cells or MPCs has, for a long time, been asked but difficult to directly answer. Interestingly, although literature suggests that CD34 is not present on human satellite cells (Peault, et al., 2007), concrete evidence is still lacking. In fact, a recent publication by Pisani and colleagues demonstrate that a subset of CD34+ muscle-derived human cells have myogenic potential, alluding to the notion of CD34 expression on human satellite cells, although their data also shows a CD34- population that is myogenic (Pisani, et al., 2010). We addressed this question by using a human CD34 (hCD34) transgenic mouse model (hCD34Tg), containing a 160 kb genomic DNA fragment that includes the hCD34 coding exons, promoter elements, and upstream and downstream flanking regions (Okuno, Huettner, et al., 2002; Radomska, et al., 2002). This mouse faithfully recapitulates hCD34 transcript expression and has been used to assess hCD34 expression on a variety of cell types (Drew, Huettner, et al., 2005; Okuno, Huettner, et al., 2002; Okuno, Iwasaki, et al., 2002). Flow cytometric analysis of hCD34Tg digested muscle shows hCD34 expression within the MPC population. The absence of hCD34 staining on WT C57Bl/6 mice confirms the specificity of our staining. In addition, RT-PCR for hCD34 on freshly sorted MPCs confirms the flow cytometry data (Figure 5.4).  149  Figure 5.4 Human CD34 is expressed in MPCs. (A) Flow cytometric analysis of skeletal muscle from hCD34 transgenic mice show expression of hCD34 within the CD34+ MPCs. (B) WT C57Bl/6 mice were used as controls. (C) RT-PCR analysis for hCD34 gene expression using sorted MPCs from hCD34Tg animals. WT C57Bl/6 mice were used as controls. Results are from 2 independent experiments.  Altogether, these results show that CD34 is expressed on all murine MPCs and, for the first time, suggest that CD34 may be similarly regulated in mice and humans. Confirming the expression of CD34 on satellite cells and an analysis of its regulation during regeneration in humans could provide novel insights to regulatory mechanisms of human muscle regeneration.  150  5.4  Significance of this work  Considering that the discovery of CD34 as a stem cell marker was over 25 years ago, relatively little is known about the exact function of this sialomucin. Recently, more work comparing WT and Cd34-/- mice or specific cell types have added to our understanding of CD34’s importance in various cell functions. For the first time, the work presented here shows that CD34 is actively involved in satellite cell function during muscle regeneration, specifically in activation, migration, and proliferation. Our data contributes to two general fields. Firstly, the role of CD34 has remained an enigma to the hematopoietic field for so long. A functional role for CD34 in a specific cell type is likely translatable to other cell types, so identification of new roles or further confirmation of proposed functions will greatly contribute to promoting CD34 from being a mere marker to a functional protein. Our data presented here are well in line with proposed functions of CD34 in activation, migration, and proliferation. Secondly, our data provides more insight to the largely enigmatic state of satellite cell activation. To date, very few regulators are identified for this transient state during regeneration. 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