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Development of strategies to enhance the contribution of hematopoietic cells to skeletal muscle repair Long, Michael Anthony 2010

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DEVELOPMENT OF STRATEGIES TO ENHANCE THE CONTRIBUTION OF HEMATOPOIETIC CELLS TO SKELETAL MUSCLE REPAIR by MICHAEL ANTHONY LONG  B.Sc., McMaster University, 2001  A THESIS SUBMITTED IN PARTIAL FULFILLMENT OF THE REQUIREMENTS FOR THE DEGREE OF DOCTOR OF PHILOSOPHY in THE FACULTY OF GRADUATE STUDIES (Medical Genetics)  THE UNIVERSITY OF BRITISH COLUMBIA (Vancouver) August 2010 © Michael Anthony Long, 2010  ABSTRACT The ability of bone marrow derived cells to contribute to skeletal muscle repair may represent a novel means of cell therapy for myopathies. However, this phenomenon takes place at exceedingly low frequencies and has failed to yield any measurable functional improvement in disease models. In an effort to increase the efficiency of this process, we designed Cre/loxP-based tracing strategies to identify the lineages and mechanisms involved. However, these experiments were complicated by a previously unknown limitation of common Cre-reporter strains. We have studied the Z/AP, Z/EG and R26R-EYFP reporter strains and have demonstrated that although each reporter can be reliably activated by Cre during early development, exposure to Cre in adult hematopoietic cells results in a much lower frequency of reporter-positive cells in the Z/AP or Z/EG strains than in the R26R-EYFP strain. In reporter-negative cells derived from the Z/AP and Z/EG strains, the transgenic promoter is methylated and Cre-mediated recombination of the locus is inhibited. These findings suggest that the Z/AP and Z/EG strains may not be suitable for the investigation of developmental plasticity in adult models. As bone marrow derived cells are now believed to contribute to skeletal muscle repair primarily via fusion, we also constructed a chimeric measles hemagglutinin, Hα7, which efficiently mediates the fusion of diverse cell types with skeletal muscle. When compared directly to polyethylene glycol in vitro, Hα7 consistently generated a ten to fifteen fold increase in heterokaryon yield and induced insignificant levels of toxicity. More importantly, Hα7 was also capable of increasing the contribution of mouse and human bone marrow derived cells to skeletal muscle repair in vivo. ii  TABLE OF CONTENTS ABSTRACT................................................................................................................ii TABLE OF CONTENTS............................................................................................iii LIST OF FIGURES....................................................................................................vi CO-AUTHORSHIP STATEMENT ............................................................................vii CHAPTER 1. INTRODUCTION..................................................................................1 1.1 SKELETAL MUSCLE DEVELOPMENT AND REGENERATION...............1 1.2 MUSCULAR DYSTROPHY AND EXISTING THERAPIES ........................4 1.3 SKELETAL MUSCLE REPAIR BY BONE MARROW DERIVED CELLS...8 1.3.1 Hematopoiesis .............................................................................8 1.3.2 Discovery of a circulating myogenic progenitor..........................11 1.3.3 Phenotype of the circulating myogenic progenitor......................13 1.3.4 Recruitment of the circulating myogenic progenitor ...................17 1.3.5 Fusion versus transdifferentiation ..............................................20 1.3.6 Nuclear reprogramming .............................................................23 1.3.7 Prospects for therapy .................................................................26 1.4 SPECIFIC AIMS ......................................................................................28 1.4.1 Identification of hematopoietic lineages responsible for the generation of bone marrow derived muscle.....................................28 1.4.2 Identification of the mechanism responsible for the generation of bone marrow derived muscle...........................................................30 1.4.3 Increase the contribution of bone marrow derived cells to skeletal muscle repair. ..................................................................................33 1.5 REFERENCES ........................................................................................34 CHAPTER 2. SILENCING INHIBITS CRE-MEDIATED RECOMBINATION OF THE Z/AP AND Z/EG REPORTERS IN ADULT CELLS ..............................................44 2.1 INTRODUCTION .....................................................................................44 2.2 RESULTS ................................................................................................48 2.2.1 Inefficient activation of the Cre reporter in hematopoietic stem cells derived from adult Z/EG mice ..................................................48  iii  2.2.2 Inefficient activation of the Cre reporter in myeloid cells derived from adult Z/AP and Z/EG mice .......................................................51 2.2.3 The transgenic locus is resistant to Cre-mediated recombination in a subset of granulocytes derived from Z/EG mice.......................54 2.2.4 The transgenic locus is methylated in reporter-negative granulocytes derived from LysM-Cre/Z/EG and LysM-Cre/Z/AP mice .........................................................................................................56 2.2.5 Expression of the pre-excision reporter is also variegated in peripheral blood leukocytes derived from Z/AP and Z/EG mice.......59 2.3 DISCUSSION ..........................................................................................61 2.4 MATERIALS AND METHODS .................................................................64 2.4.1 Ethics statement.........................................................................64 2.4.2 Transgenic mice.........................................................................64 2.4.3 TIE2-tTA/Tet-O-Cre/Z/EG mice..................................................64 2.4.4 Flow cytometry...........................................................................65 2.4.5 Excision analysis........................................................................65 2.4.6 Bisulfite sequencing ...................................................................66 2.5 REFERENCES ........................................................................................67 CHAPTER 3. TARGETED CELL FUSION ENHANCES THE CONTRIBUTION OF HEMATOPOIETIC CELLS TO SKELETAL MUSCLE FIBERS............................70 3.1 INTRODUCTION .....................................................................................70 3.2 RESULTS ................................................................................................73 3.2.1 Design, construction and characterization of Hα7......................73 3.2.2 Verification of bona fide heterokaryons ......................................75 3.2.3 Comparison of Hα7 and PEG induced fusion.............................77 3.2.4 Nuclear reprogramming following Hα7 mediated fusion.............80 3.2.5 Hα7 mediated fusion in vivo .......................................................82 3.3 DISCUSSION ..........................................................................................85 3.4 METHODS...............................................................................................88 3.4.1 Construction of Hα7 ...................................................................88 3.4.2 In vitro fusion assays..................................................................89  iv  3.4.3 Immunofluorescence and FISH..................................................90 3.4.4 Quantitative real-time gene expression analysis........................91 3.4.5 Lentiviral vectors ........................................................................92 3.4.6 In vivo fusion assays ..................................................................93 3.5 REFERENCES ........................................................................................95 CHAPTER 4. CONCLUSION ...................................................................................98 4.1 CONCLUSION.........................................................................................98 4.2 REFERENCES ......................................................................................107 APPENDIX A. ANIMAL CARE CERTIFICATES ...................................................110  v  LIST OF FIGURES Figure 1.1 Tentative model of hematopoiesis…………………………………………...9 Figure 1.2 Phenotype of the circulating myogenic progenitor………………………..14 Figure 1.3 Signals affecting recruitment & differentiation of myogenic progenitors..18 Figure 1.4 Strategy for the identification of hematopoietic lineages capable of contributing to skeletal muscle repair…………………………………………………....29 Figure 1.5 Strategy to distinguish fusion from transdifferentiation in the generation of bone marrow derived muscle…………………………….……………………………....31 Figure 2.1 Genomic organization of Cre-reporter transgenes………………………..45 Figure 2.2 Activation of the EGFP reporter is inefficient in the blood of adult TIE2tTA/Tet-O-Cre/ZEG mice…………………………………………………………….…...49 Figure 2.3 Activation of the Cre-reporter gene is less efficient in Z/AP and Z/EG mice than in R26R-EYFP mice…………………………………………………………..52 Figure 2.4 The transgenic locus is resistant to Cre-mediated recombination in a subset of granulocytes derived from the Z/EG mice…………………………………...55 Figure 2.5 The transgenic locus is methylated in reporter-negative granulocytes derived from LysM-Cre/Z/EG and LysM-Cre/Z/AP mice………………………………57 Figure 2.6 Expression of the pre-excision reporter is variegated in peripheral blood leukocytes derived from Z/AP and Z/EG mice………………………………………….60 Figure 3.1 Design, construction and characterization of Hα7………………………..74 Figure 3.2 Formation of bona fide heterokaryons……………………………………..76 Figure 3.3 Comparison of Hα7 and PEG-mediated fusion efficiencies……………..78 Figure 3.4 Nuclear reprogramming following Hα7-mediated fusion…………………81 Figure 3.5 Hα7-mediated fusion in vivo………………………………………………...83 Figure 4.1 Model illustrating the proposed methylation status of the Z/EG locus during early embryogenesis………………………………………………………….…100  vi  CO-AUTHORSHIP STATEMENT Chapter 2 M. Long designed experiments, performed experiments, analyzed data and wrote the manuscript. F. Rossi designed experiments and edited the manuscript. Chapter 3 M. Long designed experiments, performed experiments, analyzed data and wrote the manuscript. J. Brind’Amour performed FISH analysis. F. Rossi designed experiments and edited the manuscript.  vii  CHAPTER 1. INTRODUCTION 1.1 SKELETAL MUSCLE DEVELOPMENT AND REGENERATION In vertebrate organisms, cells that are specialized for contraction may be broadly divided into four categories: skeletal muscle fibers, cardiomyocytes, smooth muscle cells and myoepithelial cells[1]. These cell types serve unique, non-overlapping functions and regulate a multitude of processes. Skeletal muscle, as the name implies, is connected by tendons to the bones of the skeleton and generates the forces required for virtually all voluntary movements. Conversely, cardiac muscle, smooth muscle and myoepithelial cells are responsible for involuntary movements such as the beating of the heart, peristalsis and dilation of the iris respectively. Although the ability to generate contractile forces is conserved among these cell types, they are otherwise dissimilar in many respects. Skeletal muscle fibers in particular, exhibit a number of characteristics that are the direct result of a unique mode of development and regeneration. During embryonic development, skeletal muscle formation is initiated by the migration of somite-derived progenitor cells into the developing limb buds as well as into the epaxial myotome, which eventually forms the muscles of the main body axis[2]. In these locations, a number of signals including Wnts stimulate the proliferation of skeletal muscle progenitor cells and induce expression of the myogenic determination factors Myf5 and MyoD[3]. These basic helix-loop-helix (bHLH) transcription factors are extremely powerful regulators of gene expression and initiate a hierarchical cascade that promotes myogenesis. In fact, expression of MyoD alone is sufficient to induce myogenesis in a number of differentiated adult  1  somatic cell types including fibroblasts and chondroblasts[4, 5]. Following the proliferative phase, the expression of Myf5 and MyoD is repressed, allowing other transcription factors including Myogenin and Mef2 family members to activate a differentiation program[6, 7]. However, unlike the vast majority of somatic cells, the differentiation of skeletal muscle progenitor cells ultimately culminates in large-scale cell fusion. As a result, differentiated skeletal muscle fibers, also known as myofibers or myotubes, are long cylindrical cells containing several hundred nuclei and can reach a length of 2-3cm in adult humans[1]. Following fusion, the nuclei contained within myofibers withdraw from the cell cycle and their transcriptional activity is altered to serve the function of adult skeletal muscle. This involves expression of all components of the contractile apparatus as well as secretion of a specialized extracellular matrix known as the basal lamina, which completely surrounds each myofiber, providing tensile strength and stability[8]. Importantly, during embryonic development not all skeletal muscle progenitor cells undergo the process of fusion. Following the initial wave of myogenesis described above, a second wave of progenitor cells, whose developmental origins remain uncertain, migrate into sites of developing muscle and take up residence between the plasma membrane and basal lamina of each myofiber[9]. These cells, known as satellite cells, persist during development and remain in the sublaminar position throughout adulthood[10]. Under homeostatic conditions in adult skeletal muscle, satellite cells are quiescent and are relatively rare, representing roughly 2-5% of the total nuclei present[11]. However, in response to injury, these cells proliferate and contribute to myofiber repair in a manner that recapitulates several aspects of  2  embryonic myogenesis. Although some of the factors involved in the repair of adult skeletal muscle tend to be dictated by the nature of the injury, the basic mechanisms underlying the regenerative process remain the same. In general, damaged muscle tissue is rapidly infiltrated by neutrophils and later by macrophages, which serve to phagocytose and eliminate fragments of necrotic myofibers[12]. These inflammatory cells, as well as damaged myofibers themselves also secrete a battery of growth factors including fibroblast growth factors (FGFs), which serve to activate satellite cells within 2-6hr of injury[13]. Reminiscent of embryonic myogenesis, this activation process is characterized by entry into the cell cycle and induced expression of MyoD[14]. At this stage, activated satellite cells are often referred to as myoblasts and undergo several rounds of division within 72hr of injury[15]. Unlike embryonic myogenesis however, the subsequent specification of cell fate results in two distinct outcomes. Most myoblasts differentiate in a myogenin dependent manner and fuse to form new myofibers within 7 days of damage. A subset of myoblasts however, prematurely downregulate MyoD, withdraw from the cell cycle and return to quiescence, thereby replenishing the satellite cell pool[16]. Although the signals governing these mutually exclusive outcomes are currently unclear, asymmetric divisions induced by Notch signaling are suspected to play a role[17]. Formation of nascent myotubes is finally followed by a maturation period during which innervation occurs, such that normal tissue architecture is restored within two weeks of injury[12].  3  1.2 MUSCULAR DYSTROPHY AND EXISTING THERAPIES By virtue of their central role in locomotion, skeletal muscle fibers are subjected to significant mechanical forces. However, these forces are not borne entirely by the myofibers themselves. As described above, each skeletal muscle fiber is surrounded by structure known as the basal lamina, which is composed largely of a lattice network of collagen IV and laminin[8]. Although the tensile strength of the lattice network itself provides a level of support to skeletal muscle fibers, the basal lamina must be linked across the plasma membrane to the cytoskeleton of each myofiber in order to provide mechanical strength and stability. In order to serve this purpose, differentiated skeletal muscle fibers also express the laminin binding receptors, alpha7 integrin and dystroglycan[18, 19]. These transmembrane receptors are bound to beta1 integrin and sarcoglycans respectively at the cell surface as well as to intracellular linker proteins including dystrophin, which in turn are bound to the actin cytoskeleton[20]. Although this system generally serves to maintain myofiber integrity during contraction, mutations in a number of the components have been described, which lead to various forms of a condition known collectively as muscular dystrophy. For example, the most common form of this condition, Duchenne muscular dystrophy, is known to be caused by mutations in the dystrophin gene[21]. Additionally, mutations in laminin and a number of sarcoglycans have been described, which result in congenital muscular dystrophy and limb-girdle muscular dystrophy respectively[22-26]. In each of these conditions, myofibers are not adequately stabilized by the basal lamina and as a result, the plasma membrane is damaged and eventually destroyed by the forces involved in simple voluntary  4  movements[27]. As described earlier, adult skeletal muscle retains a remarkable capacity for regeneration. However, this ability is not without limits. In all forms of muscular dystrophy, ongoing muscle damage eventually exceeds the capabilities of the satellite cell-mediated repair pathway. Although the clinical severity of this outcome can be relatively mild and involve generalized muscle weakness, patients suffering from Duchenne muscular dystrophy typically lose the ability to walk by the age of 10 and die by the age of 30[28]. At present, these patients are primarily treated with physical therapy and respiratory assistance in addition to the corticosteroid drugs Prednisone and Deflazacort, which are believed to exert their therapeutic benefit by a combination of anabolic and immunosuppressive effects[29]. However, long-term use of corticosteroids is well known to induce serious side effects including immunosuppression, osteoporosis and weight gain[30]. Moreover, these treatments merely delay the progression of symptoms and do not address the underlying cause of the disease. Given the efficiency of satellite cell-mediated skeletal muscle repair, myoblast transplantation has been an attractive candidate for treatment of muscular dystrophies[31]. In theory, satellite cells may be harvested via muscle biopsy from a healthy donor, expanded in culture and transplanted into patients where they will participate in muscle regeneration and provide nascent myofibers with functional copies of a defective gene. In reality however, clinical trials of this strategy over the past 20 years have yielded little success. The failure of these protocols is now generally attributed to two main factors. 5  I. Myoblasts survive poorly following transplantation. In fact, several groups have reported that at least 75% of donor myoblasts die within 72hr of transplantation[3234]. Once again, the precise mechanisms responsible for this phenomenon are unclear, although oxidative stress and free radicals derived from inflammatory cells are suspected to play a role[35]. Perhaps more importantly, all donor-derived myoblasts are rejected in less than two weeks if recipients are not placed on immunosuppressive therapy[36]. At present, strategies intended to overcome these limitations are relatively unsophisticated, involving transplantation of larger numbers of cells and sustained immunosuppression with FK506[31]. However, as with longterm use of corticosteroids, sustained treatment with FK506 is known to induce a number of serious side effects including nephrotoxicity and diabetes[30]. II. Myoblasts migrate poorly following transplantation. Due to the fact that myoblasts cannot be recruited from the circulation to sites of muscle damage, all clinical trials conducted to date have involved intramuscular transplantation of donor cells[37]. Unfortunately, following this mode of delivery, myoblasts do not migrate further than 200µm from the site of injection[38]. Therefore, systemic myoblastbased treatment of muscle tissue in Duchenne muscular dystrophy patients will require an enormous number of intramuscular injections. For example, in recent clinical trials, patients received 100 injections per cm2 to a total of 4000[39-41]. Although this protocol has yielded the most encouraging results of any clinical trial to date, in the best case, dystrophin expression was restored in only 35% of myofibers and no therapeutic effect was observed in any patient. Given the severity of the disease and the absence of alternative treatment options, patients suffering from  6  Duchenne muscular dystrophy may be willing to undergo this procedure on a larger scale. However, it is clear that cell therapy of muscular dystrophies would be greatly facilitated by the identification of an alternative myogenic progenitor that is capable of homing to sites of damage via the circulation.  7  1.3 SKELETAL MUSCLE REPAIR BY BONE MARROW DERIVED CELLS 1.3.1 Hematopoiesis In addition to the satellite cells of skeletal muscle, adult stem cells have been identified in numerous tissues including bone marrow[42], peripheral nervous system[43], central nervous system[44], myocardium[45], intestine[46], liver[47] and skin[48] where they appear to play a role in homeostatic maintenance and injury repair. Among these, hematopoietic stem cells residing in adult bone marrow are the most highly characterized. In adult vertebrates, these cells are responsible for the generation of all mature blood lineages via a process of stepwise commitment, which  progressively  restricts  the  differentiation  potential  of  intermediate  progenitors[49] (Figure 1.1).  8  Figure 1.1 Tentative model of hematopoiesis. Self-renewing, long-term reconstituting hematopoietic stem cells (LT-HSC) first give rise to transiently reconstituting, short-term hematopoietic stem cells (ST-HSC). ST-HSC in turn, produce common myeloid progenitors (CMP) and common lymphoid progenitors (CLP). CLP are the source of committed progenitors that eventually give rise to T and B lymphocytes. CMP further give rise to megakaryocyte-erythroid progenitors (MEP) and granulocyte-macrophage progenitors (GMP). MEP are the source of committed progenitors that eventually give rise to erythrocytes and megakaryocytes whereas GMP are the source of committed progenitors that eventually give rise to mast-cells, neutrophils, eosinophils and monocytes.  9  While the mechanisms involved in the differentiation of hematopoietic stem cells are extremely complex and remain the focus of intense study, it appears that both stochastic and instructive factors play a role in this process. According to the socalled, intrinsic theory, individual hematopoietic stem cells and multipotent progenitors co-express low levels of several lineage-specific transcription factors, with eventual differentiation being the result of stochastic reinforcement of a particular gene expression cascade and repression of all other alternatives[50]. Under homeostatic conditions, promiscuous gene expression in both hematopoietic stem cells and multipotent progenitors has been convincingly demonstrated[51, 52]. However, extrinsic signals also clearly play a role in hematopoiesis. For example, differentiation of T lymphocytes is absolutely dependent on IL-7 stimulation of progenitor cells within the thymus[53]. Importantly, signals derived from a number of pathological  conditions  may  also  instructively  alter  hematopoietic  lineage  differentiation. For example, in response to acute bacterial infection, bone marrow stromal cells secrete the cytokines, GM-CSF and GCSF, which stimulate multiple levels of granulopoiesis resulting in a selective increase in neutrophil levels[54]. Likewise, in response to anemia or hypoxia, the kidneys produce increased levels of erythropoietin, which stimulates increased production of erythrocytes from bone marrow derived progenitors[54]. Thus, adult hematopoietic stem cells and the process of hematopoiesis in general exhibit a remarkable plasticity and are able to respond appropriately to diverse physiological demands.  10  1.3.2 Discovery of a circulating myogenic progenitor Originally, it was believed that the repair of adult tissues was a semi-autonomous process as the multipotency of adult stem cells was restricted to regeneration of their tissue of origin. In 1998 however, a groundbreaking study of skeletal muscle repair called such dogma into question. This report by Ferrari et al. suggested that following a bone marrow transplant, donor derived cells were infrequently able to participate in the regeneration of skeletal muscle which had been injured with snake venom cardiotoxin[55]. Although bone marrow derived cells were found to contribute to muscle far less efficiently than satellite cells, the observation generated an enormous amount of interest in the plastic differentiation potential of adult stem cells and provided hope for a novel, cell-based therapeutic strategy aimed at the systemic treatment of muscle degenerative diseases. This study was quickly followed by further demonstrations of the ability of bone marrow derived cells to contribute to skeletal muscle repair[56, 57] as well as to other tissues including the heart[57], central nervous system[58, 59] and liver[60, 61]. Unfortunately, the rapid succession of such reports did not allow for a great deal of standardization of methodology. Therefore it may not be surprising that other groups, including at least two high profile reports, were unable to detect any contribution of bone marrow derived cells to the brain[62] or skeletal muscle[63], casting a degree of doubt on the existence of the phenomenon and creating a great deal of controversy. Although such discrepancies have subsequently been attributed to differences in experimental protocol[64], a great deal of controversy remained. At the heart of the debate was a simple hypothesis, which was proposed by the earliest reports describing the  11  formation of donor-derived tissues following bone marrow transplantation. That is, resembling their well-known role in the repopulation of all blood lineages, hematopoietic stem cells also retain the ability to differentiate into lineages of several other tissues. However, to date this has not been conclusively demonstrated. Thus, at the outset of the research described here, any effort to improve upon the exceedingly low frequency of this non-classical repair process for therapeutic purposes would first require a greater understanding of the cell types and mechanisms involved.  12  1.3.3 Phenotype of the circulating myogenic progenitor For at least five years following the initial report of Ferrari et al., the uncommitted nature of hematopoietic stem cells implicated them as the direct precursors of donor derived tissues. However, early studies utilizing whole bone marrow involved transplantation of a complex mixture of several lineages including mature hematopoietic cells, adipocytes, osteoblasts, and endothelial cells thereby rendering claims of hematopoietic stem cell plasticity somewhat premature. In subsequent years, few attempts have been made to identify the phenotype of the plastic bone marrow-derived lineage and most remain inconclusive due to technical difficulties. For example, studies involving intramuscular injection of fractionated bone marrow have been hampered by low viability of transplanted cells and have often yielded results differing form those obtained following a bone marrow transplant[65-67]. Alternatively, transplantation of purified hematopoietic stem cells has been equally ineffective in the confirmation of hematopoietic stem cell plasticity due to the potential heterogeneity of the ‘purified’ population. In an effort to circumvent these problems, a number of groups have demonstrated the occurrence of donor-derived muscle in mice whose hematopoietic system had been reconstituted with a single hematopoietic stem cell[63, 68-70]. While these experiments prove that the hematopoietic lineage contains a circulating myogenic progenitor, it remains a formal possibility that any daughter lineage of the hematopoietic stem cell is actually directly responsible for this phenomenon. In fact Camargo et al. have suggested that the hematopoietic cells participating in the repair of skeletal muscle are myelomonocytic in origin (Figure 1.2B)[69].  13  Figure 1.2 Phenotype of the circulating myogenic progenitor. The majority of skeletal muscle repair is performed by satellite cells, which are located between the sarcolemma and basal lamina of each muscle fiber (A). These cells are activated following muscle injury and fuse with multinucleated myofibers to repair damage. Recently it has been demonstrated that bone marrow derived cells also infrequently contribute to skeletal muscle repair and several models have emerged which attempt to define the lineage and mechanism involved. Camargo et al. have proposed that myelomonocytic cells are capable of fusing directly with damaged myofibers (B). Doyonnas et al. have demonstrated that c-kit+ myelomonocytic precursors are capable of contributing to skeletal muscle and evidence from the same group suggests that this may occur through a satellite cell intermediate (C). Sherwood et al. have found bone marrow derived cells in the satellite cell niche that appear to be non-hematopoietic in origin (D).  14  Unfortunately this study also remains inconclusive due to the fact that the selected marker of myeloid cells, Lysozyme-M, has also been shown to be expressed in a fraction of hematopoietic stem cells[52]. This caveat may eventually prove more academic than practical however, as other groups have also implicated myelomonocytic cells in the transfer of donor derived markers to the liver[71, 72] and skeletal muscle[73]. Utilizing fluorescence-activated cell sorting of bone marrow derived cells, Doyonnas et al. have demonstrated that only fractions containing c-kit+ immature myelomonocytic precursors are capable of contributing to myofibers following intramuscular injection (Figure 1.2C)[73]. However, in the absence of data describing the survival or expansion of each fraction following intramuscular injection, these results are difficult to interpret. Moreover, this technique has in the past proven to be less efficient than bone marrow transplantation in the generation of donor derived muscle[66, 74], suggesting that parameters such as sustained hematopoietic engraftment or recruitment from the circulation may be important in the process. Hence, lineages other than myelomonocytic cells may also possess the ability to contribute to skeletal muscle regeneration in other experimental models. As an alternative to the entire concept of developmental plasticity, Ratajczak et al. have proposed that circulating myogenic progenitors are simply satellite cells, which express the chemokine receptor CXCR4 and therefore accumulate in the bone marrow as a result of local expression of the CXCR4 ligand, stromal-derived factor 1 (SDF-1)[75]. Clearly, this theory does not account for the presence of donor-derived muscle in mice reconstituted with a single hematopoietic stem cell. However, it remains a formal possibility that bone marrow may indeed contain low numbers of  15  bona fide myoblasts or that other non-hematopoietic bone marrow derived lineages may also contribute to muscle repair. Along these lines, Sherwood et al. have recently demonstrated that the only subset of donor-derived cells found within the satellite cell niche which are capable of myogenesis upon co-culture with differentiating myoblasts do not express the pan-hematopoietic marker, CD45[76]. While it may be reasonable to concede that this marker could be down regulated in a myogenic environment, Sherwood et al. also demonstrate that this population is only generated following transplantation of whole bone marrow and not following transplantation  of  highly  purified  hematopoietic  stem  cells,  suggesting  a  mesenchymal rather than hematopoietic origin (Figure 1.2D). Thus short of once again invoking variations introduced by disparate methodologies, one must postulate the existence of multiple sources of a circulating myogenic progenitor in order to reconcile these various reports.  16  1.3.4 Recruitment of the circulating myogenic progenitor While many of the factors involved in the conversion of bone marrow derived cells to muscle remain unknown, it is now quite clear that muscle damage is an important requirement for this phenomenon. For example, intramuscular injection of snake venom toxins such as cardiotoxin or notexin has been demonstrated to reliably induce regeneration mediated via bone marrow derived cells, yet the effect is rarely observed in contralateral, uninjured muscles[68, 69]. In a more clinically relevant setting, the chronic myofiber degeneration observed in the murine mdx model of Duchenne muscular dystrophy appears to be sufficient to induce the myogenic conversion of bone marrow derived cells[69]. Notably, the recruitment of macrophages to damaged muscle is observed in both of these models and has been interpreted as further proof of the involvement of myelomonocytic cells in the contribution of bone marrow derived cells to skeletal myofibers[29, 69]. The process of inflammation however, involves extremely sophisticated means of recruiting cells from the circulation and the chemokines involved may also specifically summon other progenitor cells to damaged muscle. Recently, a role for SDF-1 in such trafficking has been suggested by a number of groups (Figure 1.3A)[77-79].  17  Figure 1.3 Signals affecting recruitment and differentiation of myogenic progenitors. A number of groups have demonstrated that the recruitment of bone marrow derived cells to damaged muscle is enhanced by local expression of stromal derived factor-1 (SDF-1) (A). However, delivery of this chemokine to uninjured muscle is insufficient to promote such recruitment, suggesting a role for unidentified, damage-induced factors in the homing process. Polesskaya et al. have shown that damaged muscle also upregulates various Wnt isoforms, which subsequently act upon resident CD45+/Sca1+ cells to induce expression of myogenic markers (B).  18  In addition to describing the SDF-1 based recruitment of satellite cells to bone marrow, Ratajczak et al. have also suggested that these cells are re-mobilized to peripheral blood and return to damaged muscle in response to a gradient of SDF-1 expressed therein following injury[77]. Likewise, myocardial infarction has been shown to transiently induce expression of SDF-1 in injured cardiac tissue, greatly enhancing the recruitment of bone marrow derived cells, including a c-kit+ fraction, to the heart[78, 79]. Conversely, expression of virally delivered or locally injected SDF-1 as well as G-CSF induced mobilization of bone marrow progenitor cells to the peripheral blood has proven insufficient to promote such homing in the absence of injury, suggesting a role for auxiliary damage-induced factors in the recruitment process[78, 79]. In agreement, Musaro et al. have demonstrated that recruitment of bone marrow derived cells to damaged skeletal muscle is also enhanced by local expression of insulin-like growth factor 1 (IGF-1)[80]. Interestingly, although this study as well as those of SDF-1 demonstrate significantly increased recruitment of bone marrow derived cells to damaged muscle tissue, none have reported a resultant increase in the frequency of donor-derived skeletal myofibers or cardiomyocytes. These observations imply that the mere recruitment of bone marrow derived cells to a site of injury may not be the rate limiting step in the formation of donor derived muscle and shift the focus to locally acting processes which follow homing.  19  1.3.5 Fusion versus transdifferentiation As mentioned earlier, the studies reported by Ferrari et al. as well as those that followed were immediately and widely attributed to so-called stem cell plasticity despite a paucity of evidence to support either the participation of stem cells or plasticity in the process. Therefore yet another caveat to temper this burgeoning field has been the possibility that the contribution of donor derived markers to nonhematopoietic recipient tissues is a consequence of fusion between donor and recipient cells as opposed to spontaneous, plastic differentiation of stem cells across lineage boundaries or transdifferentiation of mature lineages. This scenario was initially proposed following the observation of spontaneous fusion between cocultured embryonic stem cells and GFP positive bone marrow resulting in hyperdiploid cells expressing GFP yet retaining pluripotency[81]. Following up on this in vitro data, a number of subsequent reports have indeed confirmed the role of cell fusion in the transfer of donor derived markers to liver, heart and central nervous system tissues following a bone marrow transplant[82-86]. This is not to say that the process of environmentally induced reprogramming has been completely dismissed. In fact, fusion does not appear to play a role in the contribution of bone marrow-derived markers to insulin producing pancreatic islet cells[87] or epithelial cells of the lung, liver and skin[88]. In the case of skeletal muscle, the role of either fusion or transdifferentiation in the formation of myofibers expressing donor derived markers remains contentious for several reasons including the confounding effects of myoblast fusion in the physiological formation and repair of this tissue. This is certainly the case in studies that utilize the co-expression of  20  donor and recipient specific markers within a single myofiber to suggest fusion between donor-derived hematopoietic cells and mature host myofibers[69, 74]. Formally, these experiments do not exclude the possibility that such data may be due to the fusion of endogenous myoblasts with a nascent, autonomous, donorderived myofiber. Nor do they exclude the possibility that the donor-derived cell had converted to a myogenic phenotype prior to fusion with an existing myofiber. On the other hand, studies implicating transdifferentiation in the formation of myofibers expressing donor-derived markers are not without their own problems. For example, LaBarge and Blau have proposed an enticingly intuitive mechanism in which irradiation induced damage first recruits bone marrow-derived cells to the satellite cell niche beneath the basal lamina where local environmental cues induce these cells to convert to a resting myogenic phenotype. Subsequently, exerciseinduced damage activates these cells and elicits the contribution of donor-derived markers to regenerating myofibers via the physiological satellite cell mediated repair pathway (Figure 1.2C)[89]. Although subsequent studies have reported similar results[90, 91], others have contradicted both this model and each other, finding either no donor-derived satellite cells[69] or donor-derived cells occupying the satellite cell niche that, while expressing some myogenic markers, were not autonomously myogenic in vitro[76]. The latter discrepancy may be reconciled based on the fact that the myoblast isolation employed by LaBarge and Blau to test the myogenic potential of bone marrow-derived cells, included a nine-day expansion of mononuclear cells from the muscle of transplant recipients prior to re-plating in clonal conditions. This step therefore, may have recapitulated the co-culture of donor  21  derived and endogenous myofiber associated cells, which Sherwood et al. demonstrate to be effective in inducing a myogenic phenotype in a subset of myofiber-associated bone marrow derived cells[76]. Furthermore, both studies agree that the donor-derived cells occupying the satellite cell niche express a variety of myogenic markers and are capable of contributing to the formation of donor derived myofibers in vivo. As for the local environmental cues involved in the myogenic conversion of bone marrow derived cells, Polesskaya et al. have shown that the Wnt isoforms upregulated within damaged skeletal muscle, activate the canonical Wnt signaling pathway in resident CD45+/Sca1+ cells in vivo and induce expression of early myogenic markers including Pax 7 within the same cells in vitro (Figure 1.3B)[92]. Although expression of the time-tested hematopoietic marker, CD45, strongly suggests that these cells may be the progeny of hematopoietic stem cells, their origin in bone marrow has yet to formally proven. The biological progression from bone marrow through satellite cell to myofiber described above provides an attractive model for transdifferentiation. However, none of the relevant reports formally exclude the possibility that the intermediate myogenic precursors may be themselves the product of fusion. Karyotypic analysis suggesting that these cells are diploid was performed after in vitro expansion, during which reductive divisions may have led to the loss of the extra chromosomes and the reversion to a diploid genotype[89]. Therefore at present, the precise mechanism responsible for the generation of donor derived skeletal myofibers remains unclear and it is entirely possible that both fusion and transdifferentiation play a role. 22  1.3.6 Nuclear reprogramming Many reports have described the expression of donor-derived markers in nonhematopoietic recipient tissues following a bone marrow transplant as ‘plasticity’ or a ‘contribution’ of bone marrow derived cells. Yet very few studies have examined the degree to which gene expression is altered in donated nuclei or how significant this contribution is to the function of any given recipient tissue. In many cases, donorderived cells are marked with a transgene such as LacZ or GFP, which is ubiquitously  expressed  and  therefore  provides  no  evidence  of  nuclear  reprogramming. Furthermore, the expression of such genes would hardly be expected to contribute to the function of engrafted tissues. This however, is not to say that the nuclei of mammalian somatic cells are completely refractory to significant and meaningful alterations in gene expression. In fact, the ability of cytoplasmic factors to activate previously silenced genes was demonstrated in heterokaryons over two decades ago[93, 94]. More importantly, the generation of cloned animals via transfer of adult somatic cell nuclei to enucleated oocytes has demonstrated that terminally differentiated cells retain the potential to reestablish the gene expression profile required to produce functional cells of any tissue[95-97]. Although both of these examples are reminiscent of cell fusion as they involve the exposure of nuclei from one cell type to cytoplasmic factors in another, in vitro and in vivo examples of transdifferentiation are also known to occur. For example, the conversion of cultured myoblasts to adipocytes has been accomplished via ectopic expression of the transcription factors PPARγ and C/EBPα[98] or inhibition of Wnt signaling[99]. In vivo, the differentiation of pancreatic epithelial progenitor cells into  23  hepatocytes following engraftment of the adult rat liver has also been described[100]. More recently, the transfer of a small number of transcription factors including Oct4, Sox2, Klf4 and c-myc has been shown to revert several differentiated cell types to a state of pluripotency, thereby creating so-called ‘induced pluripotent stem cells’ or iPS cells[101-104]. These cells possess the capacity for differentiation into diverse lineages in vitro as well as the ability to contribute to chimeric animals in vivo[105-108]. As an extension of this phenomenon, the transfer of other small groups of transcription factors has been shown to directly convert pancreatic exocrine cells to insulin-producing beta cells as well as to stimulate the direct conversion  of  fibroblasts  to  functional  neurons[109,  110].  Thus,  nuclear  reprogramming of adult somatic cells is clearly possible. The issue now confronting the plasticity of bone marrow derived lineages regards the completeness of such reprogramming. In a limited number of cases, a degree of reprogramming has been demonstrated by activation of the muscle specific, myosin light chain 3F promoter[55, 69] or by expression of dystrophin[56, 57] following incorporation of bone marrow derived cells into skeletal myofibers. These experiments however, do not imply complete conversion from a hematopoietic to myogenic gene expression profile within donorderived nuclei. On the contrary, the generation of endogenous, revertant fibers via exon skipping in mdx mice is well documented, casting doubt on the reliability of dystrophin expression as a hallmark of reprogramming[111]. Furthermore, Lapidos et al. have recently reported that in a murine model of cardiomyopathy and muscular dystrophy caused by targeted disruption of the of δ-sarcoglycan gene, the majority of  24  donor derived cells engrafting either skeletal or cardiac muscle fail to activate δsarcoglycan expression despite exposure to syncytial myogenic transcription factors[112]. As stated by the authors however, the extent to which these donorderived nuclei may have been reprogrammed is unknown. Early markers of myogenesis such as MyoD or desmin may have been activated yet reprogramming may not have reached a threshold required to facilitate expression of late markers including δ-sarcoglycan[113]. Therefore, the infrequency with which bone marrow derived cells contribute to skeletal muscle also currently prevents systematic investigation of subsequent nuclear reprogramming processes.  25  1.3.7 Prospects for therapy To date, the most convincing demonstration of the ability of bone marrow derived cells to contribute to the repair of a non-hematopoietic tissue has been in a murine model of the genetic liver disease, fumarylacetoacetate hydrolase (Fah) deficiency. In this model, a functional copy of the Fah enzyme is provided to Fah-/- host hepatocytes via fusion with wild type myelomonocytic cells following a bone marrow transplant. As a result, donor derived hepatocytes are given a growth advantage allowing them to expand and regenerate the majority of the liver, curing mice of the disease. Unfortunately, an analogous myopathy model does not exist and with rare exceptions, the expansion of bone marrow derived myogenic cells has not been described[114]. More recently, a number of studies have demonstrated that transplanted bone marrow cells are also capable of contributing to functional improvement following stroke injury[115]. At present it appears that this effect is mediated by the secretion of cytokines and growth factors as well as by direct participation of bone marrow derived  cells  in  angiogenesis[116,  117].  Interestingly,  both  fusion  and  transdifferentiation have been shown to play a role in the formation of blood vessels following stroke and are responsible for generating bone marrow derived pericytes and endothelial cells respectively[118, 119]. Although this process, much like the generation of donor derived muscle is known to be enhanced by inflammation, very little is known about the particular mediators involved, thereby precluding significant enhancement of either phenomenon[118].  26  Aside from discordant reports of treatment success in animal models of muscular dystrophy, most groups agree that the frequency with which bone marrow-derived cells contribute to regenerating muscle is exceedingly low, thus precluding any clinical application[120-123]. Indeed, similarly low frequencies of donor-derived nuclei have been observed in myofibers of Duchenne’s patients who received a bone marrow transplant to treat an unrelated pathology[124]. Despite this lack of preclinical success, the desire to transfer findings from the bench to the bedside has led rapidly to clinical trials involving intracoronary transplantation of hematopoietic progenitor cells in an attempt to regenerate cardiac myocytes lost as a result of an infarct[125]. However, recent data demonstrates that under these conditions hematopoietic stem cells make no contribution to the ischemic myocardium, suggesting that this strategy may be ill founded[66, 67], or that the mechanism underlying potential beneficial effects is not based on myocyte replacement. These results underscore the fact that clinical advances must await more rigorous characterization of the mechanisms involved in the generation of bone marrow derived muscle as well as the development of strategies to increase its efficiency to therapeutic levels.  27  1.4 SPECIFIC AIMS 1.4.1 Identification of hematopoietic lineages responsible for the generation of bone marrow derived muscle. This aim was to be addressed utilizing a Cre-loxP based tracing strategy. In theory, specific hematopoietic lineages would be labeled by crossing Z/EG reporter mice[126] with transgenic strains expressing Cre-recombinase in T cells (LckCre)[127], B cells (CD19-Cre)[128] or myelomonocytic cells (LysM-Cre)[129]. Thus, following transplantation of labeled bone marrow into wildtype mice, the presence of GFP-positive myofibers in recipients would identify the capability of an individual lineage to contribute to skeletal muscle (Figure 1.4).  28  A pCAGGS  EGFP  β-geo  pCAGGS  EGFP  Cre  B Z/EG  x  Labeled-Lineage Mice  Bone Marrow Transplant into WT Mice  Lineage-Specific Cre Strain  C  Labeled Lineage Myofiber  GFP+ Myofiber  BM Transplant  Skeletal Muscle  Fusion or Transdifferentiation  Donor Derived Muscle  Figure 1.4 Strategy for the identification of hematopoietic lineages capable of contributing to skeletal muscle repair. (A) In Z/EG reporter mice, transcription of a β-galactosidase/neomycin phosphotransferase fusion gene (β-geo) is driven by a hybrid CMV/β-actin promoter (pCAGGS) and terminated by polyadenylation sites (white boxes). Cre recombines loxP sites (white triangles) to remove the β-geo gene, activating expression of enhanced green fluorescent protein (EGFP). (B) In the experimental set-up, the Z/EG strain is first crossed with mice expressing Crerecombinase in a specific hematopoietic lineage. Bone marrow from the resulting offspring is then transplanted into wild-type recipients. In this way, contribution of the labeled lineage to skeletal muscle will be easily detectable by expression of the donor-derived reporter in recipient muscle fibers (C).  29  1.4.2 Identification of the mechanism responsible for the generation of bone marrow derived muscle This aim was to be addressed utilizing a modification of the Cre-loxP based tracing strategy  described  above  in  order  to  distinguish  fusion  events  from  transdifferentiation events. PCX-NLS-Cre transgenic mice[130], which ubiquitously express Cre recombinase, were to be transplanted with bone marrow from the Z/EG reporter strain. Thus in transplant recipients, the expression of β-galactosidase would identify products of transdifferentiation while EGFP expression would distinguish products of fusion (Figure 1.5)  30  A  Z/EG Hematopoietic Cell Myofiber  GFP+ Myofiber  BM Transplant  PCX-NLS-Cre Skeletal Muscle  B  Recombination  Fusion  Donor Derived Muscle  Z/EG Hematopoietic Cell Myofiber  LacZ+ Myofiber  BM Transplant  PCX-NLS-Cre Skeletal Muscle  Transdifferentiation  Donor Derived Muscle  Figure 1.5 Strategy to distinguish fusion from transdifferentiation in the generation of bone marrow derived muscle. PCX-NLS-Cre mice are transplanted with bone marrow derived from the Z/EG reporter strain. Thus, if hematopoietic cells contribute to skeletal muscle by fusion, Cre expressed by recipient nuclei will activate EGFP expression in donor nuclei (A). Alternatively, if hematopoietic cells contribute to skeletal muscle by transdifferentiation, donor nuclei will not be exposed to Cre and continue to express β-galactosidase (B).  31  Clearly these experiments rely on efficient activation of the EGFP reporter in Z/EG mice. Therefore, we first analyzed the labeling efficiency of adult hematopoietic cells derived from this reporter strain. As described in Chapter 2, we discovered that epigenetic silencing inhibits Cre-mediated recombination of the Z/EG reporter in adult cells, thereby precluding our initial specific aims.  32  1.4.3 Increase the contribution of bone marrow derived cells to skeletal muscle repair. The contribution of bone marrow derived cells to skeletal muscle repair is now largely agreed to be due to the fusion of macrophages with damaged myofibers[69, 73]. However, the mechanisms involved in this process remain insufficiently understood to enhance its efficiency for therapeutic purposes. As stated earlier, recruitment of bone marrow derived cells to the site of injury does not appear to be the rate limiting step in the formation of donor derived muscle. In fact, in several human skeletal myopathies, muscle tissue is chronically inflamed as a result of ongoing fiber degeneration[131]. Therefore, we have designed a system to increase the efficiency with which these inflammatory cells fuse with skeletal muscle fibers. As described in Chapter 3, we have created a chimeric measles hemagglutinin, which specifically and efficiently mediates the fusion of diverse cell types with skeletal muscle both in vitro and in vivo. We anticipate that this reagent may facilitate the development of novel cell and gene therapies for skeletal myopathies.  33  1.5 REFERENCES 1.  Alberts, B., Molecular biology of the cell. 4th ed. 2002, New York: Garland Science. xxxiv, [1548] p.  2.  Shi, X. and D.J. Garry, Muscle stem cells in development, regeneration, and disease. Genes Dev, 2006. 20(13): p. 1692-708.  3.  Buckingham, M., et al., The formation of skeletal muscle: from somite to limb. J Anat, 2003. 202(1): p. 59-68.  4.  Davis, R.L., H. Weintraub, and A.B. Lassar, Expression of a single transfected cDNA converts fibroblasts to myoblasts. Cell, 1987. 51(6): p. 9871000.  5.  Choi, J., et al., MyoD converts primary dermal fibroblasts, chondroblasts, smooth muscle, and retinal pigmented epithelial cells into striated mononucleated myoblasts and multinucleated myotubes. Proc Natl Acad Sci U S A, 1990. 87(20): p. 7988-92.  6.  Hasty, P., et al., Muscle deficiency and neonatal death in mice with a targeted mutation in the myogenin gene. Nature, 1993. 364(6437): p. 501-6.  7.  Nabeshima, Y., et al., Myogenin gene disruption results in perinatal lethality because of severe muscle defect. Nature, 1993. 364(6437): p. 532-5.  8.  Sanes, J.R., The basement membrane/basal lamina of skeletal muscle. J Biol Chem, 2003. 278(15): p. 12601-4.  9.  Relaix, F., et al., A Pax3/Pax7-dependent population of skeletal muscle progenitor cells. Nature, 2005. 435(7044): p. 948-53.  10.  Mauro, A., Satellite cell of skeletal muscle fibers. J Biophys Biochem Cytol, 1961. 9: p. 493-5.  11.  Snow, M.H., The effects of aging on satellite cells in skeletal muscles of mice and rats. Cell Tissue Res, 1977. 185(3): p. 399-408.  12.  Goetsch, S.C., et al., Transcriptional profiling and regulation of the extracellular matrix during muscle regeneration. Physiol Genomics, 2003. 14(3): p. 261-71.  13.  Floss, T., H.H. Arnold, and T. Braun, A role for FGF-6 in skeletal muscle regeneration. Genes Dev, 1997. 11(16): p. 2040-51.  14.  Yablonka-Reuveni, Z. and A.J. Rivera, Temporal expression of regulatory and structural muscle proteins during myogenesis of satellite cells on isolated adult rat fibers. Dev Biol, 1994. 164(2): p. 588-603. 34  15.  Zammit, P.S., et al., Kinetics of myoblast proliferation show that resident satellite cells are competent to fully regenerate skeletal muscle fibers. Exp Cell Res, 2002. 281(1): p. 39-49.  16.  Zammit, P.S., et al., Muscle satellite cells adopt divergent fates: a mechanism for self-renewal? J Cell Biol, 2004. 166(3): p. 347-57.  17.  Conboy, I.M. and T.A. Rando, The regulation of Notch signaling controls satellite cell activation and cell fate determination in postnatal myogenesis. Dev Cell, 2002. 3(3): p. 397-409.  18.  von der Mark, H., et al., Skeletal myoblasts utilize a novel beta 1-series integrin and not alpha 6 beta 1 for binding to the E8 and T8 fragments of laminin. J Biol Chem, 1991. 266(35): p. 23593-601.  19.  Ibraghimov-Beskrovnaya, O., et al., Primary structure of dystrophinassociated glycoproteins linking dystrophin to the extracellular matrix. Nature, 1992. 355(6362): p. 696-702.  20.  Ervasti, J.M. and K.P. Campbell, A role for the dystrophin-glycoprotein complex as a transmembrane linker between laminin and actin. J Cell Biol, 1993. 122(4): p. 809-23.  21.  Hoffman, E.P., et al., Conservation of the Duchenne muscular dystrophy gene in mice and humans. Science, 1987. 238(4825): p. 347-50.  22.  Helbling-Leclerc, A., et al., Mutations in the laminin alpha 2-chain gene (LAMA2) cause merosin-deficient congenital muscular dystrophy. Nat Genet, 1995. 11(2): p. 216-8.  23.  Noguchi, S., et al., Mutations in the dystrophin-associated protein gammasarcoglycan in chromosome 13 muscular dystrophy. Science, 1995. 270(5237): p. 819-22.  24.  Piccolo, F., et al., Primary adhalinopathy: a common cause of autosomal recessive muscular dystrophy of variable severity. Nat Genet, 1995. 10(2): p. 243-5.  25.  Lim, L.E., et al., Beta-sarcoglycan: characterization and role in limb-girdle muscular dystrophy linked to 4q12. Nat Genet, 1995. 11(3): p. 257-65.  26.  Nigro, V., et al., Autosomal recessive limb-girdle muscular dystrophy, LGMD2F, is caused by a mutation in the delta-sarcoglycan gene. Nat Genet, 1996. 14(2): p. 195-8.  27.  Petrof, B.J., et al., Dystrophin protects the sarcolemma from stresses developed during muscle contraction. Proc Natl Acad Sci U S A, 1993. 90(8): p. 3710-4. 35  28.  Emery, A.E., The muscular dystrophies. Lancet, 2002. 359(9307): p. 687-95.  29.  Khurana, T.S. and K.E. Davies, Pharmacological strategies for muscular dystrophy. Nat Rev Drug Discov, 2003. 2(5): p. 379-90.  30.  Physicians' desk reference : PDR, Medical Economics Co.: Oradell, N.J. p. v.  31.  Peault, B., et al., Stem and progenitor cells in skeletal muscle development, maintenance, and therapy. Mol Ther, 2007. 15(5): p. 867-77.  32.  Huard, J., et al., Gene transfer into skeletal muscles by isogenic myoblasts. Hum Gene Ther, 1994. 5(8): p. 949-58.  33.  Fan, Y., et al., Rapid death of injected myoblasts in myoblast transfer therapy. Muscle Nerve, 1996. 19(7): p. 853-60.  34.  Guerette, B., et al., Control of inflammatory damage by anti-LFA-1: increase success of myoblast transplantation. Cell Transplant, 1997. 6(2): p. 101-7.  35.  Suzuki, K., et al., Dynamics and mediators of acute graft attrition after myoblast transplantation to the heart. FASEB J, 2004. 18(10): p. 1153-5.  36.  Guerette, B., et al., Lymphocyte infiltration following allo- and xenomyoblast transplantation in mice. Transplant Proc, 1994. 26(6): p. 3461-2.  37.  Neumeyer, A.M., D.M. DiGregorio, and R.H. Brown, Jr., Arterial delivery of myoblasts to skeletal muscle. Neurology, 1992. 42(12): p. 2258-62.  38.  Skuk, D., et al., Successful myoblast transplantation in primates depends on appropriate cell delivery and induction of regeneration in the host muscle. Exp Neurol, 1999. 155(1): p. 22-30.  39.  Skuk, D., et al., Dystrophin expression in myofibers of Duchenne muscular dystrophy patients following intramuscular injections of normal myogenic cells. Mol Ther, 2004. 9(3): p. 475-82.  40.  Skuk, D., et al., Dystrophin expression in muscles of duchenne muscular dystrophy patients after high-density injections of normal myogenic cells. J Neuropathol Exp Neurol, 2006. 65(4): p. 371-86.  41.  Skuk, D., et al., First test of a "high-density injection" protocol for myogenic cell transplantation throughout large volumes of muscles in a Duchenne muscular dystrophy patient: eighteen months follow-up. Neuromuscul Disord, 2007. 17(1): p. 38-46.  42.  Spangrude, G.J., S. Heimfeld, and I.L. Weissman, Purification and characterization of mouse hematopoietic stem cells. Science, 1988. 241(4861): p. 58-62.  36  43.  Kruger, G.M., et al., Neural crest stem cells persist in the adult gut but undergo changes in self-renewal, neuronal subtype potential, and factor responsiveness. Neuron, 2002. 35(4): p. 657-69.  44.  Rietze, R.L., et al., Purification of a pluripotent neural stem cell from the adult mouse brain. Nature, 2001. 412(6848): p. 736-9.  45.  Beltrami, A.P., et al., Adult cardiac stem cells are multipotent and support myocardial regeneration. Cell, 2003. 114(6): p. 763-76.  46.  Bjerknes, M. and H. Cheng, Clonal analysis of mouse intestinal epithelial progenitors. Gastroenterology, 1999. 116(1): p. 7-14.  47.  Wang, X., et al., The origin and liver repopulating capacity of murine oval cells. Proc Natl Acad Sci U S A, 2003. 100 Suppl 1: p. 11881-8.  48.  Alonso, L. and E. Fuchs, Stem cells of the skin epithelium. Proc Natl Acad Sci U S A, 2003. 100 Suppl 1: p. 11830-5.  49.  Orkin, S.H. and L.I. Zon, SnapShot: hematopoiesis. Cell, 2008. 132(4): p. 712.  50.  Enver, T. and M. Greaves, Loops, lineage, and leukemia. Cell, 1998. 94(1): p. 9-12.  51.  Papayannopoulou, T., et al., Hemopoietic lineage commitment decisions: in vivo evidence from a transgenic mouse model harboring micro LCR-betaproLacZ as a transgene. Blood, 2000. 95(4): p. 1274-82.  52.  Ye, M., et al., Hematopoietic stem cells expressing the myeloid lysozyme gene retain long-term, multilineage repopulation potential. Immunity, 2003. 19(5): p. 689-99.  53.  von Freeden-Jeffry, U., et al., Lymphopenia in interleukin (IL)-7 gene-deleted mice identifies IL-7 as a nonredundant cytokine. J Exp Med, 1995. 181(4): p. 1519-26.  54.  Zhu, J. and S.G. Emerson, Hematopoietic cytokines, transcription factors and lineage commitment. Oncogene, 2002. 21(21): p. 3295-313.  55.  Ferrari, G., et al., Muscle regeneration by bone marrow-derived myogenic progenitors. Science, 1998. 279(5356): p. 1528-30.  56.  Gussoni, E., et al., Dystrophin expression in the mdx mouse restored by stem cell transplantation. Nature, 1999. 401(6751): p. 390-4.  37  57.  Bittner, R.E., et al., Recruitment of bone-marrow-derived cells by skeletal and cardiac muscle in adult dystrophic mdx mice. Anat Embryol (Berl), 1999. 199(5): p. 391-6.  58.  Brazelton, T.R., et al., From marrow to brain: expression of neuronal phenotypes in adult mice. Science, 2000. 290(5497): p. 1775-9.  59.  Mezey, E., et al., Turning blood into brain: cells bearing neuronal antigens generated in vivo from bone marrow. Science, 2000. 290(5497): p. 1779-82.  60.  Petersen, B.E., et al., Bone marrow as a potential source of hepatic oval cells. Science, 1999. 284(5417): p. 1168-70.  61.  Alison, M.R., et al., Hepatocytes from non-hepatic adult stem cells. Nature, 2000. 406(6793): p. 257.  62.  Castro, R.F., et al., Failure of bone marrow cells to transdifferentiate into neural cells in vivo. Science, 2002. 297(5585): p. 1299.  63.  Wagers, A.J., et al., Little evidence for developmental plasticity of adult hematopoietic stem cells. Science, 2002. 297(5590): p. 2256-9.  64.  Blau, H., et al., Something in the eye of the beholder. Science, 2002. 298(5592): p. 361-2; author reply 362-3.  65.  Orlic, D., et al., Bone marrow cells regenerate infarcted myocardium. Nature, 2001. 410(6829): p. 701-5.  66.  Murry, C.E., et al., Haematopoietic stem cells do not transdifferentiate into cardiac myocytes in myocardial infarcts. Nature, 2004. 428(6983): p. 664-8.  67.  Balsam, L.B., et al., Haematopoietic stem cells adopt mature haematopoietic fates in ischaemic myocardium. Nature, 2004. 428(6983): p. 668-73.  68.  Corbel, S.Y., et al., Contribution of hematopoietic stem cells to skeletal muscle. Nat Med, 2003. 9(12): p. 1528-32.  69.  Camargo, F.D., et al., Single hematopoietic stem cells generate skeletal muscle through myeloid intermediates. Nat Med, 2003. 9(12): p. 1520-7.  70.  Krause, D.S., et al., Multi-organ, multi-lineage engraftment by a single bone marrow-derived stem cell. Cell, 2001. 105(3): p. 369-77.  71.  Willenbring, H., et al., Myelomonocytic cells are sufficient for therapeutic cell fusion in liver. Nat Med, 2004. 10(7): p. 744-8.  38  72.  Camargo, F.D., M. Finegold, and M.A. Goodell, Hematopoietic myelomonocytic cells are the major source of hepatocyte fusion partners. J Clin Invest, 2004. 113(9): p. 1266-70.  73.  Doyonnas, R., et al., Hematopoietic contribution to skeletal muscle regeneration by myelomonocytic precursors. Proc Natl Acad Sci U S A, 2004. 101(37): p. 13507-12.  74.  Sherwood, R.I., et al., Determinants of skeletal muscle contributions from circulating cells, bone marrow cells, and hematopoietic stem cells. Stem Cells, 2004. 22(7): p. 1292-304.  75.  Ratajczak, M.Z., et al., Stem cell plasticity revisited: CXCR4-positive cells expressing mRNA for early muscle, liver and neural cells 'hide out' in the bone marrow. Leukemia, 2004. 18(1): p. 29-40.  76.  Sherwood, R.I., et al., Isolation of adult mouse myogenic progenitors: functional heterogeneity of cells within and engrafting skeletal muscle. Cell, 2004. 119(4): p. 543-54.  77.  Ratajczak, M.Z., et al., Expression of functional CXCR4 by muscle satellite cells and secretion of SDF-1 by muscle-derived fibroblasts is associated with the presence of both muscle progenitors in bone marrow and hematopoietic stem/progenitor cells in muscles. Stem Cells, 2003. 21(3): p. 363-71.  78.  Abbott, J.D., et al., Stromal cell-derived factor-1alpha plays a critical role in stem cell recruitment to the heart after myocardial infarction but is not sufficient to induce homing in the absence of injury. Circulation, 2004. 110(21): p. 3300-5.  79.  Askari, A.T., et al., Effect of stromal-cell-derived factor 1 on stem-cell homing and tissue regeneration in ischaemic cardiomyopathy. Lancet, 2003. 362(9385): p. 697-703.  80.  Musaro, A., et al., Stem cell-mediated muscle regeneration is enhanced by local isoform of insulin-like growth factor 1. Proc Natl Acad Sci U S A, 2004. 101(5): p. 1206-10.  81.  Terada, N., et al., Bone marrow cells adopt the phenotype of other cells by spontaneous cell fusion. Nature, 2002. 416(6880): p. 542-5.  82.  Alvarez-Dolado, M., et al., Fusion of bone-marrow-derived cells with Purkinje neurons, cardiomyocytes and hepatocytes. Nature, 2003. 425(6961): p. 96873.  83.  Vassilopoulos, G., P.R. Wang, and D.W. Russell, Transplanted bone marrow regenerates liver by cell fusion. Nature, 2003. 422(6934): p. 901-4.  39  84.  Wang, X., et al., Cell fusion is the principal source of bone-marrow-derived hepatocytes. Nature, 2003. 422(6934): p. 897-901.  85.  Weimann, J.M., et al., Stable reprogrammed heterokaryons form spontaneously in Purkinje neurons after bone marrow transplant. Nat Cell Biol, 2003. 5(11): p. 959-66.  86.  Nygren, J.M., et al., Bone marrow-derived hematopoietic cells generate cardiomyocytes at a low frequency through cell fusion, but not transdifferentiation. Nat Med, 2004. 10(5): p. 494-501.  87.  Ianus, A., et al., In vivo derivation of glucose-competent pancreatic endocrine cells from bone marrow without evidence of cell fusion. J Clin Invest, 2003. 111(6): p. 843-50.  88.  Harris, R.G., et al., Lack of a fusion requirement for development of bone marrow-derived epithelia. Science, 2004. 305(5680): p. 90-3.  89.  LaBarge, M.A. and H.M. Blau, Biological progression from adult bone marrow to mononucleate muscle stem cell to multinucleate muscle fiber in response to injury. Cell, 2002. 111(4): p. 589-601.  90.  Dreyfus, P.A., et al., Adult bone marrow-derived stem cells in muscle connective tissue and satellite cell niches. Am J Pathol, 2004. 164(3): p. 7739.  91.  Fukada, S., et al., Muscle regeneration by reconstitution with bone marrow or fetal liver cells from green fluorescent protein-gene transgenic mice. J Cell Sci, 2002. 115(Pt 6): p. 1285-93.  92.  Polesskaya, A., P. Seale, and M.A. Rudnicki, Wnt signaling induces the myogenic specification of resident CD45+ adult stem cells during muscle regeneration. Cell, 2003. 113(7): p. 841-52.  93.  Blau, H.M., C.P. Chiu, and C. Webster, Cytoplasmic activation of human nuclear genes in stable heterocaryons. Cell, 1983. 32(4): p. 1171-80.  94.  Chiu, C.P. and H.M. Blau, Reprogramming cell differentiation in the absence of DNA synthesis. Cell, 1984. 37(3): p. 879-87.  95.  Wilmut, I., et al., Viable offspring derived from fetal and adult mammalian cells. Nature, 1997. 385(6619): p. 810-3.  96.  Hochedlinger, K. and R. Jaenisch, Monoclonal mice generated by nuclear transfer from mature B and T donor cells. Nature, 2002. 415(6875): p. 1035-8.  97.  Eggan, K., et al., Mice cloned from olfactory sensory neurons. Nature, 2004. 428(6978): p. 44-9. 40  98.  Hu, E., P. Tontonoz, and B.M. Spiegelman, Transdifferentiation of myoblasts by the adipogenic transcription factors PPAR gamma and C/EBP alpha. Proc Natl Acad Sci U S A, 1995. 92(21): p. 9856-60.  99.  Ross, S.E., et al., Inhibition of adipogenesis by Wnt signaling. Science, 2000. 289(5481): p. 950-3.  100.  Dabeva, M.D., et al., Differentiation of pancreatic epithelial progenitor cells into hepatocytes following transplantation into rat liver. Proc Natl Acad Sci U S A, 1997. 94(14): p. 7356-61.  101.  Takahashi, K. and S. Yamanaka, Induction of pluripotent stem cells from mouse embryonic and adult fibroblast cultures by defined factors. Cell, 2006. 126(4): p. 663-76.  102.  Aoi, T., et al., Generation of pluripotent stem cells from adult mouse liver and stomach cells. Science, 2008. 321(5889): p. 699-702.  103.  Hanna, J., et al., Direct reprogramming of terminally differentiated mature B lymphocytes to pluripotency. Cell, 2008. 133(2): p. 250-64.  104.  Kim, J.B., et al., Pluripotent stem cells induced from adult neural stem cells by reprogramming with two factors. Nature, 2008. 454(7204): p. 646-50.  105.  Dimos, J.T., et al., Induced pluripotent stem cells generated from patients with ALS can be differentiated into motor neurons. Science, 2008. 321(5893): p. 1218-21.  106.  Okita, K., T. Ichisaka, and S. Yamanaka, Generation of germline-competent induced pluripotent stem cells. Nature, 2007. 448(7151): p. 313-7.  107.  Wernig, M., et al., In vitro reprogramming of fibroblasts into a pluripotent EScell-like state. Nature, 2007. 448(7151): p. 318-24.  108.  Raya, A., et al., Disease-corrected haematopoietic progenitors from Fanconi anaemia induced pluripotent stem cells. Nature, 2009. 460(7251): p. 53-9.  109.  Zhou, Q., et al., In vivo reprogramming of adult pancreatic exocrine cells to beta-cells. Nature, 2008. 455(7213): p. 627-32.  110.  Vierbuchen, T., et al., Direct conversion of fibroblasts to functional neurons by defined factors. Nature. 463(7284): p. 1035-41.  111.  Lu, Q.L., et al., Massive idiosyncratic exon skipping corrects the nonsense mutation in dystrophic mouse muscle and produces functional revertant fibers by clonal expansion. J Cell Biol, 2000. 148(5): p. 985-96.  41  112.  Lapidos, K.A., et al., Transplanted hematopoietic stem cells demonstrate impaired sarcoglycan expression after engraftment into cardiac and skeletal muscle. J Clin Invest, 2004. 114(11): p. 1577-85.  113.  Noguchi, S., et al., Developmental expression of sarcoglycan gene products in cultured myocytes. Biochem Biophys Res Commun, 1999. 262(1): p. 8893.  114.  Abedi, M., et al., Robust conversion of marrow cells to skeletal muscle with formation of marrow-derived muscle cell colonies: a multifactorial process. Exp Hematol, 2004. 32(5): p. 426-34.  115.  Tang, Y., et al., Transplantation of bone marrow-derived stem cells: a promising therapy for stroke. Cell Transplant, 2007. 16(2): p. 159-69.  116.  Chen, J. and M. Chopp, Neurorestorative treatment of stroke: cell and pharmacological approaches. NeuroRx, 2006. 3(4): p. 466-73.  117.  Kokovay, E., L. Li, and L.A. Cunningham, Angiogenic recruitment of pericytes from bone marrow after stroke. J Cereb Blood Flow Metab, 2006. 26(4): p. 545-55.  118.  Piquer-Gil, M., et al., Cell fusion contributes to pericyte formation after stroke. J Cereb Blood Flow Metab, 2009. 29(3): p. 480-5.  119.  Bailey, A.S., et al., Myeloid lineage progenitors give rise to vascular endothelium. Proc Natl Acad Sci U S A, 2006. 103(35): p. 13156-61.  120.  Dell'Agnola, C., et al., Hematopoietic stem cell transplantation does not restore dystrophin expression in Duchenne muscular dystrophy dogs. Blood, 2004. 104(13): p. 4311-8.  121.  Torrente, Y., et al., Human circulating AC133(+) stem cells restore dystrophin expression and ameliorate function in dystrophic skeletal muscle. J Clin Invest, 2004. 114(2): p. 182-95.  122.  Ferrari, G., A. Stornaiuolo, and F. Mavilio, Failure to correct murine muscular dystrophy. Nature, 2001. 411(6841): p. 1014-5.  123.  Wagers, A.J. and I.L. Weissman, Plasticity of adult stem cells. Cell, 2004. 116(5): p. 639-48.  124.  Gussoni, E., et al., Long-term persistence of donor nuclei in a Duchenne muscular dystrophy patient receiving bone marrow transplantation. J Clin Invest, 2002. 110(6): p. 807-14.  42  125.  Assmus, B., et al., Transplantation of Progenitor Cells and Regeneration Enhancement in Acute Myocardial Infarction (TOPCARE-AMI). Circulation, 2002. 106(24): p. 3009-17.  126.  Novak, A., et al., Z/EG, a double reporter mouse line that expresses enhanced green fluorescent protein upon Cre-mediated excision. Genesis, 2000. 28(3-4): p. 147-55.  127.  Hennet, T., et al., T-cell-specific deletion of a polypeptide Nacetylgalactosaminyl-transferase gene by site-directed recombination. Proc Natl Acad Sci U S A, 1995. 92(26): p. 12070-4.  128.  Rickert, R.C., J. Roes, and K. Rajewsky, B lymphocyte-specific, Cre-mediated mutagenesis in mice. Nucleic Acids Res, 1997. 25(6): p. 1317-8.  129.  Clausen, B.E., et al., Conditional gene targeting in macrophages and granulocytes using LysMcre mice. Transgenic Res, 1999. 8(4): p. 265-77.  130.  Nagy, A., Cre recombinase: the universal reagent for genome tailoring. Genesis, 2000. 26(2): p. 99-109.  131.  Arahata, K. and A.G. Engel, Monoclonal antibody analysis of mononuclear cells in myopathies. I: Quantitation of subsets according to diagnosis and sites of accumulation and demonstration and counts of muscle fibers invaded by T cells. Ann Neurol, 1984. 16(2): p. 193-208.  43  CHAPTER 2. SILENCING INHIBITS CRE-MEDIATED RECOMBINATION OF THE Z/AP AND Z/EG REPORTERS IN ADULT CELLS1 2.1 INTRODUCTION The bacteriophage P1 enzyme, Cre, recognizes and recombines two copies of a specific 34 base-pair sequence known as a loxP sequence located at each end of the phages linear genome. This process converts the genome to a unit-copy circular plasmid ensuring proper replication and partitioning of the prophage [1,2,3]. This process does not require cofactors, therefore loxP sites inserted into the murine genome are also recognized and recombined by Cre, resulting in excision, inversion or translocation of chromosomal sequence depending on the orientation and location of the loxP sites [4]. To date, hundreds of transgenic mouse strains have been created which express Cre under the control of tissue specific or inducible promoters and in combination with transgenic mice containing loxP-flanked alleles, have revolutionized the study of the genetic factors involved in a multitude of biological processes. In order to facilitate such studies, several so-called Cre-reporter strains have also been created [4]. These mice generally carry the gene for an easily detectable marker, the expression of which is only activated following exposure to Cre. For example, in the Z/AP reporter strain (Figure 2.1A), transcription of a βgalactosidase/neomycin phosphotransferase fusion gene (β-geo) is driven by a hybrid CMV enhancer/chicken β-actin promoter (pCAGGS) and terminated by a trimer of SV40 polyadenylation sites [5].  1  A version of this chapter has been published. Long MA and Rossi FMV (2009) Silencing Inhibits Cre-Mediated Recombination of the Z/AP and Z/EG Reporters in Adult Cells. PLoS ONE. 4(5):e5435  44  Figure 2.1 Genomic organization of Cre-reporter transgenes. In the Z/AP strain (A) and Z/EG strain (B), transcription of a β-galactosidase/neomycin phosphotransferase fusion gene (β-geo) is driven by a hybrid CMV/β-actin promoter (pCAGGS) and terminated by polyadenylation sites (white boxes). Cre recombines loxP sites (white triangles) to remove the β-geo gene, activating expression of human placental alkaline phosphatase (hPLAP) or enhanced green fluorescent protein (EGFP) in the Z/AP and Z/EG strains respectively. In the R26R-EYFP strain (C), Cre-mediated recombination of loxP sites removes a phosphoglycerate kinase (PGK) promoter and a neomycin phosphotransferase gene (NeoR), allowing the endogenous ROSA26 genomic locus to drive expression of the downstream enhanced yellow fluorescent protein (EYFP) gene.  45  The β-geo gene and polyadenylation sites are flanked by loxP sites oriented in the same direction. Therefore, upon exposure to Cre, the β-geo gene is excised and transcription of the human placental alkaline phosphatase (hPLAP) gene is activated by proximity to the pCAGGS promoter. The Z/EG strain (Figure 2.1B) was derived from the Z/AP reporter and contains the cDNA for enhanced green fluorescent protein (EGFP) in place of hPLAP [6]. Thus, all cells of Z/AP and Z/EG mice are designed to exhibit a binary readout of Cre activity, expressing β-galactosidase by default and activating expression of hPLAP or EGFP respectively upon exposure to Cre. The R26R-EYFP strain (Figure 2.1C) was generated by targeted insertion of a Cre reporter cassette into the ROSA26 genomic locus, which has been shown to be ubiquitously expressed throughout development and in adult tissues [7,8]. The reporter cassette contains a PGK promoter driving expression of a neomycin phosphotransferase gene, both of which are removed by Cre mediated recombination of flanking loxP sites. Following excision, the endogenous ROSA26 promoter drives expression of the downstream enhanced yellow fluorescent protein (EYFP) gene. Cre-reporter strains have been utilized to validate the expression profile of Cre transgenes [9], to act as a surrogate marker for excision of a second allele [10], to irreversibly label cells for lineage tracing experiments [11] and to differentiate between fusion and transdifferentiation in studies of stem cell plasticity [12]. Although it is known that the chromosomal integration site of loxP sequences can affect the efficiency with which they are recombined [13], to date there has not been a systematic comparison of the labeling efficiencies of some of the most widely used  46  Cre-reporter strains. We have undertaken such a comparison and have demonstrated that the efficiency of reporter activation in adult cells derived from the Z/AP and Z/EG strain is much lower than in adult cells derived from the R26R-EYFP strain. Furthermore, our evidence suggests that the inefficient labeling efficiency observed in the Z/AP and Z/EG strains is due to methylation of the pCAGGS promoter which prevents both reporter expression and Cre-mediated recombination of the transgenic locus.  47  2.2 RESULTS 2.2.1 Inefficient activation of the Cre reporter in hematopoietic stem cells derived from adult Z/EG mice The Z/EG reporter strain has become a valuable tool for studying embryonic development [14,15]. However, we were interested in utilizing this strain to study adult hematopoiesis and therefore created the triple transgenic TIE2-tTA/Tet-OCre/Z/EG strain. In these mice, expression of the tetracycline-transactivator is driven by the TIE2 promoter, which has been shown to be active in hematopoietic stem cells [16,17]. Therefore following removal of doxycycline from the diet of mice, the tetracycline-transactivator is able to bind to the tet-operator and drive expression of Cre in hematopoietic stem cells, theoretically resulting in expression of the EGFP reporter in all hematopoietic lineages. In order to determine the kinetics of reporter activation in this system, we analyzed the blood of triple transgenic mice for expression of EGFP at several intervals following removal of doxycycline from the diet (Figure 2.2A-C).  48  Figure 2.2 Activation of the EGFP reporter is inefficient in the blood of adult TIE2-tTA/Tet-O-Cre/ZEG mice. EGFP expression in the blood of a representative triple transgenic mouse, 3, 6 and 9 months following removal of doxycycline from the diet is shown in (A-C) respectively. EGFP expression in the blood of a representative triple transgenic mouse, bred in the absence of doxycycline is shown in (D). In all triple transgenic mice analyzed, activation of the EGFP reporter was not observed in peripheral blood leukocytes (PBL) earlier than 6 months following removal of doxycycline and remained low at nine months post induction (E).  49  Although EGFP positive cells were eventually detected in the blood of all triple transgenic mice, in the best case less than 10 percent of peripheral blood leukocytes were labeled following nine months of induction (Figure 2.2E). Conversely, the blood of triple transgenic mice that developed in the absence of doxycycline was labeled quite efficiently, validating that the system does function properly during development (Figure 2.2D). While this suggests that the Z/EG reporter may not function as well in adult hematopoietic cells as it is during development, TIE2 expression has been proposed to maintain hematopoietic stem cells in a quiescent state, thus it remains possible that in triple transgenic mice, TIE2 positive hematopoietic stem cells are labeled quite efficiently and simply do not contribute significantly to the peripheral blood [18]. Furthermore, although unlikely, it is possible that following removal of doxycycline from the diet of triple transgenic mice, the levels of the drug remaining in vivo are sufficient to suppress expression of Cre.  50  2.2.2 Inefficient activation of the Cre reporter in myeloid cells derived from adult Z/AP and Z/EG mice In an effort to eliminate pharmacological complications and to test the hypothesis that the Z/EG reporter does not function as efficiently in adult hematopoietic cells as it does during development, we crossed Z/EG mice to the LysM-Cre strain, which expresses Cre in all myelomonocytic cells and to the general deleter strain pCXNLS-Cre [19,20]. As a basis for comparison, LysM-Cre and pCX-NLS-Cre mice were also bred to the Z/AP reporter strain as well as to the R26R-EYFP reporter strain and peripheral blood leukocytes were analyzed for expression of the appropriate post-excision reporter. As expected, the R26R-EYFP reporter was activated in all peripheral blood leukocytes of pCX-NLS-Cre/R26R-EYFP mice (Figure 2.3A) and in 85 percent of granulocytes of LysM-Cre/R26R-EYFP mice (Figure 2.3B).  51  Figure 2.3 Activation of the Cre-reporter gene is less efficient in Z/AP and Z/EG mice than in R26R-EYFP mice. (A) All reporter strains were crossed to the general deleter strain, pCX-NLS-Cre, and activation of each reporter gene was assessed by flow cytometry of peripheral blood leukocytes at 12 weeks of age. (B) All reporter strains were also crossed to the myeloid-specific LysM-Cre strain, and activation of each reporter gene was assessed by flow cytometry of peripheral blood granulocytes at 12 weeks of age. Representative histograms demonstrating reporter expression in peripheral blood granulocytes are shown in (C-E) for LysM-Cre/R26REYFP, LysM-Cre/ZAP and LysM-Cre/Z/EG mice respectively (solid lines). Reporter expression in BL/6 mice is shown in dotted lines.  52  A similar labeling efficiency of LysM-Cre/R26R-EYFP granulocytes has previously been reported and most likely approaches the maximum labeling efficiency of these short-lived cells utilizing a Cre transgene expressed from a promoter which is activated during their life-cycle [21]. In contrast to these results, the average labeling efficiency of granulocytes in the LysM-Cre/Z/AP and LysM-Cre/Z/EG strains was only 57 percent and 36 percent respectively (Figure 2.3B). A similar labeling efficiency has been reported for the Z/EG strain following a cross to a separate myeloid specific-Cre strain [9]. These data, taken together with the observation that it is possible to activate the hPLAP and EGFP reporter in virtually all blood cells of most pCX-NLS-Cre/Z/AP and pCX-NLS-Cre/Z/EG mice (Figure 2.3A), further supports the hypothesis that the Z/AP and Z/EG reporters do not function as efficiently in adult hematopoietic cells as they do during development. The presence of a significant percentage of hPLAP negative and EGFP negative cells in a subset of pCX-NLS-Cre/Z/AP and pCX-NLS-Cre/Z/EG mice (Figure 2.3A) has also been reported by others and demonstrates that even under conditions of embryonic exposure to high levels of Cre recombinase, the Z/AP and Z/EG reporters are not completely reliable [6,12,22].  53  2.2.3 The transgenic locus is resistant to Cre-mediated recombination in a subset of granulocytes derived from Z/EG mice The differential labeling efficiencies observed in LysM-Cre/R26R-EYFP, LysMCre/Z/AP and LysM-Cre/Z/EG mice are unlikely to be due to variable expression of functional Cre recombinase as all mice contain the same LysM-Cre transgene and were maintained on the same genetic background. Therefore, we reasoned that the inefficient labeling of Z/EG and Z/AP granulocytes may be due to impaired recombination and/or inefficient expression of the transgenic loci. In order to differentiate between these two scenarios, we designed a PCR-based strategy to examine the efficiency of Cre-mediated excision of the β-geo gene. Granulocytes from LysM-Cre/Z/EG mice were first sorted into reporter-negative and reporterpositive populations (Figure 2.4A). Genomic DNA from these groups was then subjected to PCR reactions containing primers designed to generate an amplicon only in the presence of a recombined transgenic locus. As seen in Figure 2.4B, reporter positive cells contain a recombined locus as expected. In the reporternegative population however, the recombined locus was not detected, suggesting that Cre may be unable to efficiently access the loxP sites in the genomic DNA of cells derived from the LysM-Cre/Z/EG strain.  54  Figure 2.4 The transgenic locus is resistant to Cre-mediated recombination in a subset of granulocytes derived from the Z/EG mice. (A) Gr-1 positive cells from a LysM-Cre/Z/EG mouse were sorted into reporter negative (-) and reporter positive (+) populations. (B) Genomic DNA from these populations was then analyzed by PCR utilizing primers binding within the pCAGGS promoter and EGFP cDNA. This combination of primers (Exc) generates a 240bp fragment in the presence of the recombined locus whereas the distance across the intact locus is too large to facilitate exponential amplification under the conditions used. Primers recognizing the IL-2 inducible T-cell kinase (ITK) gene were also utilized as a positive control in reporter negative (-) and reporter positive (+) reactions as well as in a no-template (NT) control.  55  2.2.4 The transgenic locus is methylated in reporter-negative granulocytes derived from LysM-Cre/Z/EG and LysM-Cre/Z/AP mice As DNA methylation is known to be one of the primary mechanisms by which eukaryotic cells silence foreign DNA, we hypothesized that the Z/EG transgene is methylated and incorporated into heterochromatin in adult hematopoietic cells, thereby reducing the accessibility of the loxP sites for Cre mediated recombination [23,24]. We therefore subjected a 200 base-pair segment of the pCAGGS promoter containing 36 CpG dinucleotides to bisulfite sequencing in order to examine its methylation status in both reporter-negative and reporter-positive granulocytes from the LysM-Cre/Z/EG strain. As seen in Figures 2.5A,C, this segment of the pCAGGS promoter is indeed methylated to a greater extent in reporter negative cells than it is in reporter positive cells.  56  Figure 2.5 The transgenic locus is methylated in reporter-negative granulocytes derived from LysM-Cre/Z/EG and LysM-Cre/Z/AP mice. Genomic DNA from reporter negative and reporter positive populations was subjected to bisulfite sequencing in order to determine the methylation status of a 200 bp region of the pCAGGS promoter. This region contains 36 CpG dinucleotides, significantly more of which were methylated (filled circles) in clones derived from reporter negative cells than in those derived from reporter positive cells (A,B). The degree of methylation at the pCAGGS and ROSA promoters in both reporter negative (-) and reporter positive (+) populations is shown in (C) and (D) for cells derived from LysMCre/ZEG and LysM-Cre/Z/AP mice respectively. The degree of methylation at the ROSA promoter in cells derived from the LysM-Cre/R26R-EYFP strain is shown in (E).  57  In order to demonstrate that hypermethylation is not a global phenomenon in reporter negative cells, we also determined the methylation status of a 253 base-pair segment of the endogenous ROSA26 promoter containing 28 CpG dinucleotides in both reporter-negative and reporter positive cells. As seen in Figure 2.5C, very little methylation was observed at this locus in all clones examined, regardless of reporter expression. A similar correlation between hypermethylation of the pCAGGS promoter and inefficient expression of the Cre-reporter gene was also observed in granulocytes obtained from LysM-Cre/Z/AP mice (Figures 2.5B,D). A corollary to our hypothesis that DNA methylation inhibits Cre-mediated activation of the reporter gene in the Z/AP and Z/EG strains is that the ROSA26 promoter should be unmethylated in granulocytes derived form the LysM-Cre/R26R-EYFP strain. As seen in Figure 2.5E, this is indeed the case.  58  2.2.5 Expression of the pre-excision reporter is also variegated in peripheral blood leukocytes derived from Z/AP and Z/EG mice As a final confirmation of the fact that the Z/AP and Z/EG loci are silenced in adult hematopoietic cells, we also quantified expression of the pre-excision reporter, βgalactosidase,  via  fluorescein-di-beta-D-galactopyranoside  (FDG)  staining  of  peripheral blood leukocytes taken from these mice. As seen in Figure 2.6, the preexcision reporter is also inefficiently expressed in adult hematopoietic cells of Z/AP and Z/EG mice presumably due to epigenetic silencing of the transgenic locus.  59  Figure 2.6 Expression of the pre-excision reporter is variegated in peripheral blood leukocytes derived from Z/AP and Z/EG mice. Silencing of the Z/AP and Z/EG transgenic loci was demonstrated via quantification of the percentage of peripheral blood leukocytes expressing β-galactosidase by FDG staining. Data are shown as mean ± s.d. (n=3).  60  2.3 DISCUSSION We have demonstrated a diminished sensitivity to Cre-mediated recombination in adult hematopoietic cells derived from Z/AP and Z/EG mice. These reporter cassettes were randomly inserted into the mouse genome and as such are more likely to be subjected to position effect variegation than the R26R-EYFP reporter, which was inserted into the ubiquitously expressed ROSA26 genomic locus. In accordance with this hypothesis we have also demonstrated that expression of the pre-excision reporter is variegated in adult hematopoietic cells derived from Z/AP and Z/EG mice. The difference in the degree of silencing observed between Z/AP and Z/EG cells is unlikely to be due to differences in the location or extent to which the pCAGGS promoter is methylated as these parameters appear to be quite similar between cells derived from the two strains (Figure 2.5A-D). Thus, differences in the chromatin conformation of the loci into which the transgenes have been inserted likely influence the frequency and not the extent to which the pCAGGS promoter is subjected to methylation, underlying the observed differences in silencing. Interestingly, the percentage of cells expressing the pre-excision reporter (Figure 2.6) in both Z/AP and Z/EG mice is remarkably similar to the percentage of cells expressing the post-excision reporter (Figure 2.3B) in LysM-Cre/Z/AP and LysMCre/Z/EG mice respectively. This observation suggests that the population of cells that express the pre-excision reporter may be the only cells capable of undergoing Cre-mediated activation of the post-excision reporter in adult granulocytes.  61  In order to explain these findings, we propose a model wherein the Z/AP and Z/EG loci are demethylated after fertilization as a result of the genome wide demethylation that is known to occur in pre-implantation embryos [25]. Therefore, if these transgenes are exposed to Cre recombinase shortly after fertilization, such as by crossing to the pCX-NLS-Cre strain, which ubiquitously expresses Cre, the locus is accessible, the β-geo gene is efficiently excised and the downstream reporter is expressed. As embryonic development progresses however, the Z/AP and Z/EG transgenes become methylated, resulting in the eventual incorporation of the transgenic loci into heterochromatin, which inhibits access of transcription factors and Cre recombinase. Therefore, if these transgenes are exposed to Cre recombinase in adult cells, such as by crossing to the LysM-Cre strain, the β-geo gene is inefficiently excised and the locus is inefficiently expressed. The presence of the β-galactosidase (LacZ) sequence within the β-geo gene may be a significant contributor to this effect as the CpG-rich LacZ cDNA is known to induce silencing of some genes to which it is fused [26]. In accordance with this notion, we have demonstrated that excision of the β-geo gene shortly after fertilization significantly reduces silencing of the reporter loci in adult cells (Figure 2.3A). Although we have restricted our analysis to the labeling efficiency of hematopoietic cells, other groups have also reported low labeling efficiency utilizing the Z/EG strain in the adult kidney, liver, testis, adrenal glands, fat tissue, lung, pituitary gland, spleen and retina [27,28]. Furthermore, Rotolo et al. have recently developed a method for the analysis of neuronal morphology, which is based in part on the inefficiency with which the Z/AP locus is recombined in the adult brain [29]. Our  62  findings highlight the potential shortcomings of utilizing these particular Cre-reporters as surrogate markers of excision or in lineage tracing experiments. Therefore, the R26R-EYFP reporter may be the strain of choice for researchers interested in tracing the expression of Cre beyond early development.  63  2.4 MATERIALS AND METHODS 2.4.1 Ethics statement All experiments were performed in accordance with the rules of the Animal Care Committee at the University of British Columbia. 2.4.2 Transgenic mice The Z/AP, Z/EG, pCX-NLS-Cre, TIE2-tTA and Tet-O-Cre strains were generously provided by Dr. Corrinne Lobe and the LysM-Cre and R26R-EYFP mice were generously provided by Dr. Thomas Graf. Mice were housed in a specific pathogen free facility and each strain was maintained by backcrossing to the C57BL/6 strain. 2.4.3 TIE2-tTA/Tet-O-Cre/Z/EG mice Double transgenic TIE2-tTA/Tet-O-Cre mice were crossed with Z/EG mice and breeders were fed Dox-Diet (Bio-Serv) containing 200mg/kg doxycycline. After weaning, triple transgenic pups were maintained on Dox-Diet until 8 weeks of age at which time doxycycline was removed from the diet. Peripheral blood then was taken daily for a week, weekly for a month and at 2, 3, 6 and 9 months post induction. Peripheral blood leukocytes were analyzed by flow cytometry for the expression of the Cre-reporter transgene, EGFP.  64  2.4.4 Flow cytometry Peripheral blood samples were taken from the tail vein of each mouse and erythrocytes were lysed in a hypotonic solution. For experiments requiring identification of granulocytes or hPLAP, cells were stained with a PE-conjugated anti-Gr-1 antibody (eBioscience) or an anti-human hPLAP antibody (Serotec) respectively. For FDG staining, blood cells were incubated in a hypotonic solution containing 1 mM fluorescein-di-beta-D-galactopyranoside for 1 minute at 37oC. The mixture was then diluted 10-fold in PBS and incubated for 1 hour on ice. All data was collected with a Becton-Dickinson FACSCalibur and analyzed with FlowJo software. 2.4.5 Excision analysis Peripheral blood samples were prepared and stained with a PE-conjugated antiGr-1 antibody as described above. Reporter negative and reporter positive granulocytes were sorted utilizing a Becton-Dickinson FACSVantage and genomic DNA was prepared from roughly 3x104 sorted cells from each population (DNeasy Blood and Tissue Kit, QIAGEN). PCR primers designed to amplify a segment of the ITK gene were as follows: ITK-F: 5’-GCCGTAAATGAACAGGTGGTG-3’ and ITK-R: 5’-TGCTCCAGACTGTGAGAGTCG-3’. Primers designed to identify a recombined Z/EG locus were pCAGGS-F: 5’-GGGCAACGTGCTGGTTGT-3’ and EGFP-R: 5’CCAGCTCGACCAGGATGG-3’.  65  2.4.6 Bisulfite sequencing Genomic DNA from reporter negative and reporter positive populations was converted utilizing the EpiTect Bisulfite Kit (QIAGEN). For analysis of the pCAGGS promoter, DNA was subjected to a semi-nested PCR reaction utilizing primers BABF6:  5’-GGAGAGGTGYGGYGGTAGTTAATTAGAG-3’  and  BABR5d:  5’-  AAACCCCTCAAAACTTTCACRCAACCACAA-3’ for the first round followed by BABF6 and BABR4c: 5’-TCATTAAACCAAACRCTAATTACAACCC-3’ for the second round. Analysis of the ROSA26 promoter utilized the primers ROSAF2: 5’GGAAAYGTTATTGATYGTAYGGGGATT-3’  and  ROSAR3:  5′-  ACTATCTCACAAAACRACTCCACCAC-3′. PCR products were cloned into the pCR2.1 vector (Invitrogen) and sequenced from the T7 priming site utilizing Applied Biosystems BigDye v3.1 Terminator Chemistry at the NAPS Unit, UBC.  66  2.5 REFERENCES 1.  Sternberg, N. and D. Hamilton, Bacteriophage P1 site-specific recombination. I. Recombination between loxP sites. J Mol Biol, 1981. 150(4): p. 467-86.  2.  Austin, S., M. Ziese, and N. Sternberg, A novel role for site-specific recombination in maintenance of bacterial replicons. Cell, 1981. 25(3): p. 72936.  3.  Sternberg, N., et al., Bacteriophage P1 cre gene and its regulatory region. Evidence for multiple promoters and for regulation by DNA methylation. J Mol Biol, 1986. 187(2): p. 197-212.  4.  Branda, C.S. and S.M. Dymecki, Talking about a revolution: The impact of site-specific recombinases on genetic analyses in mice. Dev Cell, 2004. 6(1): p. 7-28.  5.  Lobe, C.G., et al., Z/AP, a double reporter for cre-mediated recombination. Dev Biol, 1999. 208(2): p. 281-92.  6.  Novak, A., et al., Z/EG, a double reporter mouse line that expresses enhanced green fluorescent protein upon Cre-mediated excision. Genesis, 2000. 28(3-4): p. 147-55.  7.  Srinivas, S., et al., Cre reporter strains produced by targeted insertion of EYFP and ECFP into the ROSA26 locus. BMC Dev Biol, 2001. 1: p. 4.  8.  Zambrowicz, B.P., et al., Disruption of overlapping transcripts in the ROSA beta geo 26 gene trap strain leads to widespread expression of betagalactosidase in mouse embryos and hematopoietic cells. Proc Natl Acad Sci U S A, 1997. 94(8): p. 3789-94.  9.  Ferron, M. and J. Vacher, Targeted expression of Cre recombinase in macrophages and osteoclasts in transgenic mice. Genesis, 2005. 41(3): p. 138-45.  10.  Muncan, V., et al., Rapid loss of intestinal crypts upon conditional deletion of the Wnt/Tcf-4 target gene c-Myc. Mol Cell Biol, 2006. 26(22): p. 8418-26.  11.  Jiang, X., et al., Fate of the mammalian cardiac neural crest. Development, 2000. 127(8): p. 1607-16.  12.  Harris, R.G., et al., Lack of a fusion requirement for development of bone marrow-derived epithelia. Science, 2004. 305(5680): p. 90-3.  13.  Vooijs, M., J. Jonkers, and A. Berns, A highly efficient ligand-regulated Cre recombinase mouse line shows that LoxP recombination is position dependent. EMBO Rep, 2001. 2(4): p. 292-7. 67  14.  Zhu, X., D.E. Bergles, and A. Nishiyama, NG2 cells generate both oligodendrocytes and gray matter astrocytes. Development, 2008. 135(1): p. 145-57.  15.  Kidder, B.L., et al., Embryonic stem cells contribute to mouse chimeras in the absence of detectable cell fusion. Cloning Stem Cells, 2008. 10(2): p. 231-48.  16.  Iwama, A., et al., Molecular cloning and characterization of mouse TIE and TEK receptor tyrosine kinase genes and their expression in hematopoietic stem cells. Biochem Biophys Res Commun, 1993. 195(1): p. 301-9.  17.  Yano, M., et al., Expression and function of murine receptor tyrosine kinases, TIE and TEK, in hematopoietic stem cells. Blood, 1997. 89(12): p. 4317-26.  18.  Arai, F., et al., Tie2/angiopoietin-1 signaling regulates hematopoietic stem cell quiescence in the bone marrow niche. Cell, 2004. 118(2): p. 149-61.  19.  Clausen, B.E., et al., Conditional gene targeting in macrophages and granulocytes using LysMcre mice. Transgenic Res, 1999. 8(4): p. 265-77.  20.  Nagy, A., Cre recombinase: the universal reagent for genome tailoring. Genesis, 2000. 26(2): p. 99-109.  21.  Ye, M., et al., Hematopoietic stem cells expressing the myeloid lysozyme gene retain long-term, multilineage repopulation potential. Immunity, 2003. 19(5): p. 689-99.  22.  Guo, C., W. Yang, and C.G. Lobe, A Cre recombinase transgene with mosaic, widespread tamoxifen-inducible action. Genesis, 2002. 32(1): p. 818.  23.  Walsh, C.P., J.R. Chaillet, and T.H. Bestor, Transcription of IAP endogenous retroviruses is constrained by cytosine methylation. Nat Genet, 1998. 20(2): p. 116-7.  24.  Jahner, D., et al., De novo methylation and expression of retroviral genomes during mouse embryogenesis. Nature, 1982. 298(5875): p. 623-8.  25.  Rougier, N., et al., Chromosome methylation patterns during mammalian preimplantation development. Genes Dev, 1998. 12(14): p. 2108-13.  26.  Cohen-Tannoudji, M., C. Babinet, and D. Morello, lacZ and ubiquitously expressed genes: should divorce be pronounced? Transgenic Res, 2000. 9(3): p. 233-5.  27.  Jullien, N., et al., Conditional transgenesis using Dimerizable Cre (DiCre). PLoS ONE, 2007. 2(12): p. e1355.  68  28.  Zhang, X.M., et al., Transgenic mice expressing Cre-recombinase specifically in retinal rod bipolar neurons. Invest Ophthalmol Vis Sci, 2005. 46(10): p. 3515-20.  29.  Rotolo, T., et al., Genetically-directed, cell type-specific sparse labeling for the analysis of neuronal morphology. PLoS ONE, 2008. 3(12): p. e4099.  69  CHAPTER 3. TARGETED CELL FUSION ENHANCES THE CONTRIBUTION OF HEMATOPOIETIC CELLS TO SKELETAL MUSCLE FIBERS1 3.1 INTRODUCTION The induced fusion of cells in vitro has been an essential technique for research in a number of fields including the study of nuclear reprogramming[1], the production of monoclonal antibodies[2] and the generation of dendritic-cell hybrids for cancer immunotherapy[3]. However, advances in these and other areas are currently inhibited by the limitations of traditional fusogenic agents. The most commonly utilized techniques for inducing cell fusion in vitro, namely polyethylene glycol (PEG)[4] and electrofusion[5] were first described roughly thirty years ago and although incremental refinements have gradually increased their efficacy, each of these methods remain notoriously inefficient. As a result of mechanisms that rely on random aggregation and membrane damage in order to achieve cell fusion, PEG and electrofusion protocols generally produce heterokaryons with low efficiency and high toxicity. Methods that employ micromanipulation[6], affinity crosslinking[7] or microfluidic devices[8] to properly pair two cell types are capable of increasing the efficiency of fusion. However, these systems continue to rely on PEG or electric pulses to initiate membrane fusion.  1  A version of this chapter will be submitted for publication. Long MA and Rossi FMV (2010) Targeted Cell Fusion Enhances the Contributiuon of Hematopoietic Cells to Skeletal Muscle Fibers.  70  In vivo, the ability of bone marrow derived cells to contribute to the repair of several organs is largely thought to be due to the fusion of circulating cells with damaged tissues[9]. This discovery has raised the prospect that cell fusion may represent a viable therapeutic strategy for several genetic and degenerative diseases. However, the inefficiency with which this phenomenon occurs has also precluded its therapeutic utility. Attempts to increase the efficiency of this process, including injection of snake venom toxins, appear to function by simply damaging tissue which in turn recruits inflammatory cells to the site of injury where they infrequently fuse to regenerating tissue[10]. Clearly these reagents, like PEG and electrofusion are unlikely to be suitable for clinical application. Members of the Paramyxoviridae family of viruses, including measles and Sendai virus have long been known to induce cell fusion in vivo and in vitro[11, 12]. In the case of measles virus, infection is initiated via recognition of human CD46 or CD150 on the surface of cells by the viral hemagglutinin (H) protein[13, 14]. This interaction is believed to induce a conformational change in the associated viral fusion (F) protein, exposing a hydrophobic peptide, which inserts into the target plasma membrane and mediates fusion of the virus with the cell[15]. Subsequent display of measles H and F on the surface of infected cells then initiates fusion between neighbouring cells, ultimately resulting in large multinucleated syncitia, which die primarily as a result apoptosis[16]. This cytopathic effect has motivated the development of oncolytic measles viruses, capable of specifically recognizing and infecting tumour cells. A number of groups have accomplished this by the addition of peptides[17], growth factors[18], single chain antibodies (scFv)[19] or cytokines[20]  71  to the carboxyl-terminus of the hemagglutinin protein, effectively retargeting the tropism of the measles virus, resulting in fusion and death of cells expressing the cognate receptor or antigen. To date however, chimeric hemagglutinin glycoproteins have not been developed for the generation of stable heterokaryons in vitro or for cell therapy in vivo. Here we present a system based on a chimeric measles virus hemagglutinin glycoprotein, which is capable of generating stable heterokaryons with high efficiency both in vitro and in vivo. This modified measles virus hemagglutinin, Hα7, was produced by addition of a scFv that recognizes the muscle specific integrin, α7, to the carboxyl-terminus of a mutant hemagglutinin. Co-transfection of plasmids encoding Hα7 and measles F, induced fusion of all cell types tested with cultured skeletal muscle fibers. Moreover, the efficiency of Hα7-mediated fusion was clearly superior to PEG mediated fusion and demonstrated insignificant levels of toxicity. Interspecific heterokaryons generated via Hα7 mediated fusion retained morphology characteristic of differentiated myotubes and activated transcription of human myogenic genes. Expression of Hα7 on both mouse and human inflammatory cells also increased the contribution of these cells to muscle fiber repair in vivo.  72  3.2 RESULTS 3.2.1 Design, construction and characterization of Hα7 We generated a muscle-specific fusion reagent, Hα7, by adding an anti-alpha7integrin scFv to the carboxyl terminus of a mutated measles hemagglutinin, H481A,533A, which lacks the ability to bind either measles receptor[21] (Figure 3.1A). The scFv was constructed from the well-characterized CA5.5 monoclonal antibody, which has been employed extensively in the purification and characterization of myoblasts[22]. As seen in Figures 3.1B-E, the scFv retains the specificity and affinity of the parental monoclonal antibody, as demonstrated by its ability to stain C2C12 myoblasts but not NIH/3T3 fibroblasts. We then tested the ability of cells expressing our chimeric hemagglutinin to fuse with differentiated skeletal myotubes. In order to accomplish this, 293T cells were co-transfected with plasmids encoding Hα7, F and GFP. The following day, these cells were mixed with cultures of differentiated C2C12 myotubes and twenty-four hours after mixing, the percentage of GFP-positive myotubes was determined. As seen in Figure 3.1F, transfected 293T cells can be made to fuse with nearly every myotube in the culture, with an efficiency proportional to the amount of input plasmid. Importantly, insignificant numbers of GFP-positive myotubes were observed in the same assay when H481A,533A was used in place of Hα7 or when Hα7 or F were omitted from the transfection (Figure 3.1F). Moreover, myotubes remained viable following fusion and exhibited a morphology characteristic of differentiated C2C12 cells (Figure 3.1G).  73  Figure 3.1 Design, construction and characterization of Hα7. (A) Schematic representation of Hα7, approximating the locations each blade (β1-β6) in the βpropellor fold [23, 24] as well as the location of mutations that abrogate CD46 binding (Y481A) and CD150 binding (R533A). The anti-α7 integrin scFv is displayed as a carboxy-terminal extension of the type II transmembrane glycoprotein. Standard one-letter abbreviations are used to denote amino acid residues. N: Amino-terminal cytoplasmic tail. TM: Transmembrane domain. (B-E) Evaluation of the anti-α7 integrin scFv by flow cytometry. The scFv (B, solid line) retains the ability of the parental monoclonal antibody (D, solid line) to stain C2C12 myoblasts, whereas neither antibody stains NIH/3T3 fibroblasts (C and F, solid line). In all plots, the staining level of cells incubated with secondary antibody alone is shown in dotted lines. (F) Hα7 mediates fusion of transfected 293T cells and differentiated C2C12 myotubes with an efficiency that is proportional to the amount of transfected plasmid and is absolutely dependent on the presence of the anti-α7 integrin scFv and the measles F protein. Data are shown as mean ± s.d. (n=3). (G) Morphology of myotubes following fusion. 74  3.2.2 Verification of bona fide heterokaryons In order to eliminate the possibility that the multinucleated, GFP-positive cells observed in co-cultures were exclusively derived from the homotypic fusion of transfected cells, we first cultured 293T cells in myogenic differentiation medium following co-transfection with Hα7, F and GFP. This treatment did not result in the formation of syncitia (Figure 3.2A), suggesting that transfected 293T cells are unable to autonomously initiate the fusion process and demonstrating the inability of Hα7 to facilitate fusion between cells that do not express alpha7 integrin. In co-cultures however, multinucleated, GFP-positive cells were found to express murine myosin heavy chain (Figure 3.2B-D), confirming the presence of proteins derived from both 293T and C2C12 cells within these syncitia. Furthermore, in order to confirm that these cells were heterokaryons by definition, we identified the presence of both human and murine nuclei within syncitia by differential DAPI staining (Figure 3.2E, F) as well as by fluorescent-in situ-hybridization (FISH) staining of human and murine satellite repeat DNA (Figure 3.2G). In the FISH assay, double positive nuclei were never observed, confirming that the Hα7-mediated fusion of 293T cells with differentiated C2C12 myotubes results in the generation of true heterokaryons.  75  Figure 3.2 Formation of bona fide heterokaryons. (A) 293T cells co-transfected with Hα7, F and GFP do not fuse with one another. (B-D) Following co-culture of transfected human 293T cells with differentiated mouse C2C12 myotubes, elongated GFP positive cells (B) express mouse myosin heavy-chain (C) and contain multiple nuclei (D, merged). (E,F) Differential DAPI staining demonstrates the presence of both human and mouse nuclei within heterokaryons. Mouse nuclei contain dense chromocenters while human nuclei (arrows) stain diffusely and exhibit dark nucleoli. (G) Fluorescent in situ hybridization of human α-satellite DNA (red) and mouse γsatellite DNA (green) further confirms the presence of both human and mouse nuclei within heterokaryons.  76  3.2.3 Comparison of Hα7 and PEG induced fusion PEG remains the most widely used fusogenic agent for the production of heterokaryons. Therefore, we sought to compare the efficiency Hα7-mediated fusion with that of a standard PEG-mediated fusion protocol. In this case, 293T cells were either co-transfected with plasmids encoding Hα7, F and GFP or transfected with a plasmid encoding GFP alone. The following day, equal numbers of 293THα7,F,GFP or 293TGFP cells were mixed with cultures of differentiating C2C12 cells and wells containing 293TGFP were treated with PEG to induce fusion. The number of GFPpositive myotubes as well as the total number of myotubes per low-power field was determined daily thereafter for each condition. As seen in Figure 3.3, the number of GFP-positive myotubes generated by Hα7-mediated fusion was roughly ten to fifteen-fold greater than the number generated by standard PEG-mediated fusion. This finding is unlikely to be due to improper use of PEG, as previous studies employing this method have reported similar fusion efficiencies[25]. A slight increase in the percentage of GFP positive myotubes was observed in wells subjected to Hα7 mediated fusion between day-one and day-two post fusion (Figure 3.3A versus Figure 3.3B). This observation is most likely due to that fact that differentiation of C2C12 myoblasts continues and terminates over this time frame. Thus, in the first 24 hours of co-culture, nascent myotubes are forming at a rate that simply outpaces the ability of cells expressing Hα7 to encounter and fuse with them.  77  Figure 3.3 Comparison of Hα7 and PEG-mediated fusion efficiencies. (A-C) The total number of myotubes (white bars) as well as the number of GFP-positive myotubes (green bars) was determined by visual inspection of randomly selected, low power (5x) fields at (A) 1 day (B) 2 days and (C) 3 days post fusion. ***: P<0.001 (unpaired t-test). Data are shown as mean ± s.d. (n=6). Values above each bar represent the average fusion efficiency.  78  At all timepoints, the total number of myotubes surviving in the Hα7 treatment group was nearly twice as great as the number surviving PEG treatment. In fact, the total number of myotubes present following Hα7-mediated fusion was not significantly different from controls lacking any fusogen, demonstrating the greatly reduced toxicity of this method with respect to PEG. A decrease in the total number of myotubes was observed on day three post-fusion as differentiated muscle cells began to contract and detach from the dish. However, this phenomenon uniformly affected the total number of myotubes across all treatment groups and did not preferentially affect GFP-positive myotubes within any group.  79  3.2.4 Nuclear reprogramming following Hα7 mediated fusion To investigate the potential utility of Hα7 mediated fusion for nuclear reprogramming studies, we analyzed the expression of human muscle transcripts in heterokaryons comprised of MRC-5 human lung fibroblasts and differentiating C2C12 myotubes. Similar experiments performed with PEG have demonstrated that following the fusion of several human cell types with C2C12 myotubes, muscle genes are activated in human nuclei with kinetics that resemble myogenic differentiation[26, 27]. Therefore, we chose to examine the expression of MyoD and myogenin, as markers of early and late myogenic differentiation respectively. As seen in Figure 3.4, isolated MRC-5 cells do not express either of these transcription factors. However, MyoD expression was upregulated within 72 hours of Hα7 mediated fusion and reached a peak roughly 48 hours later (Figure 3.4A). On the other hand, the levels of myogenin transcript remained relatively low at early time points while steadily increasing over time (Figure 3.4B). These data are in agreement with previous reports[26, 27], confirming that heterokaryons generated via Hα7 mediated fusion retain the capacity to undergo nuclear reprogramming.  80  Figure 3.4 Nuclear reprogramming following Hα7-mediated fusion. (A,B) Quantitative RT-PCR analysis of (A) human MyoD and (B) human myogenin transcript levels at daily intervals following Hα7-mediated fusion of MRC-5 cells and differentiating C2C12 myotubes. All values were normalized to β-actin transcript levels and subsequently to the mean expression level on Day 1. Data are shown as mean ± s.d. (n=3). (C) Endpoint RT-PCR reactions demonstrating the specificity of human MyoD and myogenin primers.  81  3.2.5 Hα7 mediated fusion in vivo Finally, to evaluate the potential utility of targeted cell fusion for regenerative medicine, we investigated the ability of Hα7 to increase the efficiency of fusion between inflammatory cells and muscle fibers in vivo. Bone marrow derived from the Z/AP transgenic strain[28], which expresses β-galactosidase in the majority of hematopoietic cells was infected with the lentiviral vectors LV-HIG and LV-FIY, which encode Hα7-IRES-GFP and F-IRES-YFP respectively. Infected bone marrow was then transplanted into congenic C57BL/6 recipients and four weeks later, muscle damage was induced in the tibialis anterior muscle by notexin injection. All recipients remained healthy throughout the treatment and at eight weeks posttransplant, mice were sacrificed and hind limbs were examined for the presence of β-galactosidase-positive muscle fibers. Unfortunately, the efficiency of lentiviral transduction in all recipients was low (Figure 3.5A,B). However, significantly more donor-derived muscle fibers were observed in a subset of recipients that were transplanted with infected bone marrow as compared to control recipients that received uninfected bone marrow transplants (Figure 3.5C).  82  Figure 3.5 Hα7-mediated fusion in vivo. (A,B) Expression of lentiviral transgenes in the peripheral blood of mice four weeks after transplantation with (A) infected or (B) uninfected bone marrow. (C) The maximum number of donor derived muscle fibers per section at eight weeks post transplant in recipients of infected and uninfected bone marrow. (D,E) Expression of lentiviral transgenes in U937 cells following infection with (D) LV-HIG and LV-FIY or (E) LV-FIY alone. (F) Donor derived, GFP positive muscle fibers in NOD/SCID recipients one week following transplantation of U937Hα7,F. (G) Same image displayed in (F) with the green channel removed to emphasize intact basal lamina surrounding donor derived muscle fibers.  83  As a model of human inflammatory cells, the monocyte cell line U937, was also infected with LV-HIG and LV-FIY (Figure 3.5D). Doubly infected U937Hα7,F cells or single infected U937F control cells (Figure 3.5E) were then injected into the tibialis anterior muscle of NOD/SCID recipients. One week after transplantation, mice were sacrificed and hind limbs were examined for the presence of GFP positive muscle fibers. As seen in Figures 3.5F and G, donor derived fibers exhibiting normal morphology and surrounded by basal lamina were regularly observed in recipients of U937Hα7,F cells. In all tissues obtained from control animals however, only a single GFP-positive muscle fiber was ever observed.  84  3.3 DISCUSSION We have created a cell fusion reagent, Hα7, which overcomes the low efficiency, high toxicity and lack of specificity exhibited by existing chemical and physical fusogens. As opposed to PEG and electrofusion, our system is based on a specific ligand-receptor interaction, which simultaneously promotes the proper pairing and efficient fusion of cells. This feature maximizes the generation of heterokaryons and virtually eliminates the non-productive formation of homokaryons. In vitro, Hα7 mediated fusion efficiencies routinely exceeded ninety percent and consistently generated ten to fifteen fold more heterokaryons than a standard PEG mediated protocol. A similar increase in fusion efficiency has recently been described utilizing a microfluidic device to control cell pairing[8]. While this represents a significant improvement over existing techniques, the microfluidic device is limited to the manipulation of a maximum of six thousand cell pairs per run. Our method on the other hand, has no such limitations and is therefore capable of producing far more of heterokaryons per experiment. The ability of Hα7 to increase fusion efficiency was not gained at the expense of cell viability. At all time points analyzed, the total number of myotubes present following Hα7-mediated fusion was not significantly different from controls lacking any fusogen. Moreover, heterokaryons generated via Hα7 treatment exhibited normal healthy morphology, characteristic of differentiated C2C12 myotubes. In contrast, PEG mediated fusion resulted in the death of roughly fifty percent of myotubes. This difference is likely due to the fact that unlike PEG, the measles fusion glycoprotein complex is capable of initiating and stabilizing the fusion process  85  without relying on the induction of membrane damage. Following Hα7 mediated fusion of MRC-5 human lung fibroblasts and differentiating C2C12 myotubes, expression of the myogenic transcription factors MyoD and myogenin was induced in human nuclei. The kinetics of this reprogramming process was nearly identical to previous reports based on PEG mediated fusion[26, 27]. While it remains possible that the expression of Hα7 and F may alter gene expression in heterokaryons, it is unlikely that this effect will be greater than the perturbations caused by the toxic effects of PEG. Therefore, we anticipate that the increased yield and quality of heterokaryons generated via Hα7 mediated fusion will facilitate the systematic identification of the factors and mechanisms involved in the process of nuclear reprogramming. Skeletal muscle is naturally repaired by satellite cells, which proliferate and fuse to multinucleated myofibers[29]. However, in several human skeletal myopathies, ongoing cycles of fiber degeneration overwhelm this regenerative process and muscle tissue becomes chronically inflamed as a result[30]. Therefore, we examined the ability of Hα7 to increase the efficiency of fusion between inflammatory cells and multinucleated myofibers in vivo. In two distinct models, Hα7 expression increased the contribution of mouse and human inflammatory cells to skeletal muscle repair. Unfortunately, in the mouse bone-marrow transplantation model, the magnitude of this effect was limited by the low efficiency of lentiviral transduction. Despite this fact, these results demonstrate that Hα7 is capable of enhancing fusion of inflammatory cells and skeletal muscle fibers in an immunocompetent host. In agreement, Iankov et al, have demonstrated that cells infected with measles virus are capable of 86  undergoing fusion in vivo even in the presence of pre-existing humoral immunity[31]. Therefore we further anticipate that the creation of circulating myogenic progenitors via expression of Hα7 on inflammatory cells will facilitate the development of novel cell and gene therapies for skeletal myopathies.  87  3.4 METHODS 3.4.1 Construction of Hα7 RNA was prepared from the CA5.5 hybridoma (RNeasy, Qiagen) and cDNA was produced utilizing Superscript II (Invitrogen) and an oligo-dT primer. The variable region of the immunoglobulin heavy chain was then amplified utilizing Platinum Pfx DNA Polymerase (Invitrogen) and the degenerate primers VH: 5’-TGA GGT GCA GCT GGA GGA GTC-3’ and Cγ: 5’-AGA CCG ATG GGG CTG TTG TTT TGG C-3’. The variable region of the immunoglobulin light chain was amplified utilizing the degenerate primers Vκ: 5’-GAC ATT CTG ATG ACC CAG TCT-3’ and Cκ: 5’-TGG ATA CAG TTG GTG CAG CAT CAG C-3’. PCR products were cloned into pCRBluntII-TOPO (Invitrogen) and sequenced from the T7 priming site. (Applied Biosystems BigDye v3.1 Terminator Chemistry, NAPS Unit, UBC). These sequences were utilized to identify the particular variable region genes expressed in CA5.5 cells and gene specific primers were designed. The variable region of the immunoglobulin heavy chain was then reamplified from CA5.5 cDNA utilizing the primers CA5.5H-F: 5’-AAA AGA TCT GGC CCA GCC GGC CCA GGT GCA GCT GAA GGA GTC-3’and CA5.5H-R: 5’-GAC GGT GAC CAT GAC TCC TTG G-3’. The variable region of the immunoglobulin light chain was reamplified utilizing the primers CA5.5L-F: 5’-AAA GAG CTC GCT GAC CCA GTC TCC TGC TTT G-3’ and CA5.5L-R: 5’-AAA CTC GAG CGG CCG CCC GTT TCA ATT CCA GCT TGG TGC-3’. A complete scFv was then assembled by cloning the heavy chain fragment upstream and the light chain fragment downstream of a glycine-serine (G4S1)3 linker contained in pASK85-9E10 utilizing BglII, BstEII and SacI, XhoI sites respectively, thereby creating pCA5.5scFv.  88  The CA5.5scFv was subsequently fused to a human light chain constant region by subcloning into pLC-huCκ with BglII and NotI. This plasmid, pCA5.5scFv- huCκ, was transiently transfected into 293T cells via standard calcium phosphate precipitation and 48 hours later, neat supernatant containing the CA5.5scFv-huCκ fusion protein was utilized to stain C2C12 myoblasts and NIH/3T3 fibroblasts in parallel with a 0.5µg/mL dilution of the CA5.5 monoclonal antibody. The goat anti-human-kappa-PE (Southern Biotech) and the goat anti-rat-PE secondary (Southern Biotech) antibodies were used to detect the CA5.5scFv-huCκ and CA5.5 staining respectively. All flow cytometry data was collected with a Becton-Dickinson FACSCalibur and analyzed with FlowJo software. Following confirmation of specificity, the CA5.5scFv was fused to the carboxylterminus of a mutant measles hemagglutinin contained in pTNH6-Haa using SfiI and NotI, thereby creating pHα7. 3.4.2 In vitro fusion assays 293T and C2C12 cells were maintained in DMEM (Gibco) supplemented with 10% and 20% fetal bovine serum (Gibco) respectively. To induce differentiation, C2C12 cells were plated in DMEM supplemented with 2% horse serum (Invitrogen) on collagen-coated dishes (Sigma, Becton Dickinson) at a density of 4x104 cells/cm2. Twenty-four hours later, cytosine β-D-arabinofuranoside (Ara-C) (Sigma) was added to a concentration of 1x10-5M in order to eliminate proliferating myoblasts. 293T cells were transfected with calcium phosphate twenty-four hours prior to co-culture and were plated onto C2C12 cells at a density of 4x104 cells/cm2. Co-cultures were initiated following two or five days of C2C12 differentiation and are referred to as 89  differentiating or differentiated cultures respectively. PEG mediated fusion of cells was carried out as described previously. Briefly, 293T cells were mixed with differentiating C2C12 myoblasts and allowed to settle and adhere for four to six hours. Medium was then completely aspirated and replaced with prewarmed 50% PEG 1500 (Roche) for sixty seconds. PEG was then removed and cells were washed three times in prewarmed DMEM. Cultures were subsequently maintained in DMEM supplemented with 2% horse serum, 1x10-5M Ara-C and 1x10-5M ouabbain (Sigma) to eliminate unfused human cells. Fusion efficiency was quantified at selected intervals by enumerating the total number of myotubes as well as the number of GFP-positive myotubes present in at least six randomly selected low power (5x) fields. 3.4.3 Immunofluorescence and FISH To detect mouse myosin-heavy chain expression, heterokaryons were first fixed in 4% paraformaldehyde (PFA) for 5 minutes at room temperature, washed in PBS and permeabilized in 0.5% Triton X-100 for 5 minutes at room temperature. Cells were then stained with mouse anti-mouse myosin-heavy chain (Developmental Studies Hybridoma Bank) overnight at 4oC, followed by a 1 hour incubation with goat antimouse Alexa 568 (Molecular Probes) at room temperature. Nuclei were counterstained with 4´,6-diamidino-2-phenylindole (DAPI) (1µg/mL). For FISH, cells were post-fixed with 4% formaldehyde, treated with 1 mg/ml pepsin, dehydrated in increasing series of ethanol and air-dried. Cells were denatured for 3 minutes at 80°C in hybridization mixture (70% formamide, 0.5 µg/ml 90  of  Cy-3–conjugated  PNA probe  specific  to  human  α-satellite  sequences  (CTCCAAATATCCACTTGC), 0.5 µg/ml of Cy-5–conjugated PNA probe specific to mouse major satellite (GAAGGACCTGGAATATGG) and 0.25% (w/v) blocking reagent (DuPont) in 10 mM Tris (pH 7)). Hybridization was performed at room temperature for 1 hour and slides were then washed with 70% formamide/10 mM Tris (pH 7.2; twice for 15 min each) and with 0.05 M Tris/0.15 M NaCl (pH 7.2) containing 0.05% Tween-20 (three times for 5 min each). Slides were dehydrated, air dried and counterstained with 0.2 µg of DAPI/ml, and mounted in antifading solution (DABCO). The images were acquired with the DeltaVision RT imaging system (Applied Precision) on an inverted microscope (IX70 Olympus) equipped with a Coolsnap HQ digital camera. Images stacks were acquired for each wavelength in 12 bit grey scale through a 60/1.4 oil immersion lens. Deconvolution was performed on the Deltavision RT imaging system and single plane projection of individual images was done using SoftWoRx software (Applied Precision). 3.4.4 Quantitative real-time gene expression analysis Heterokaryons were generated in 24-well plates as described above. Following fusion, RNA was harvested daily (RNeasy, Qiagen) from a single well for each treatment condition for a total of eight days. Purified RNA was treated with DNAse (Fermentas) and cDNA was then produced utilizing Superscript II (Invitrogen) and random hexamer primers (Invitrogen). qPCR reactions were set up with Maxima SYBR Green/ROX qPCR Master Mix (Fermentas) and the following primers pairs hMyoDF: 5’-CAC TCC GGT CCC AAA TGT AG-3’ and hMyoDR: 5’-GGT ATA AAC GTA CAA ATT CCC TGT A-3’. hMyogeninF: 5’-CAG CGA ATG CAG CTC TCA C-3’  91  and hMyogeninR: 5’-CAG AAG TAG TGG CAT CTG TGG-3’. β-actinF: 5’-TTT GAG ACC TTC AAC ACC CCA GCC-3’ and β-actinR: 5’-AAT GTC ACG CAC GAT TTC CCG C-3’. Gene expression was quantified using a 7900HT Fast Real-Time PCR System and the 7000 SDS relative quantification software (Applied Biosystems). 3.4.5 Lentiviral vectors The transfer vector, pLV-HIG, was constructed by inserting the Hα7 cDNA contained in pHα7, downstream of the EF1α promoter in the third generation lentiviral vector, pCCL.sin.cPPT.EF1α.SET7.IRES.GFP.WPRE utilizing BamH1. The transfer vector, pLV-FIY, was constructed by first inserting the measles fusion protein cDNA contained in pCGF, downstream of the EF1α promoter in same the third generation lentiviral vector described above, utilizing Xma1 and Xba1. The EYFP cDNA contained in pEYFP (Clontech) was then cloned downstream of the IRES utilizing Nco1 and BsrG1. Lentiviruses were produced by cotransfecting 293T cells with the appropriate transfer vector as well as with the packaging plasmids, pMDL, pRev and pVSVG utilizing calcium phosphate. Supernatant was harvested 36 to 60 hours later, filtered through a 0.45µm filter (Pall) and concentrated by ultracentrifugation at 19,400rpm for 2hr at 20oC in a Beckman SW28 rotor. Viral pellets were resuspended in 100µL of HBSS and stored in 20µL aliquots at -80oC. In order to determine the biological titer of each lentivirus preparation, 293T cells were infected with serial dilutions of stock solutions and infection efficiency was quantified 48hr later via flow cytometry (BD FACScan and FlowJo software).  92  3.4.6 In vivo fusion assays In the mouse bone marrow transplantation model, 1.5x105 Sca-postitive bone marrow cells derived from a Z/AP transgenic mouse were plated in StemSpan SFEM (StemCell Technologies) supplemented with rhFlt-3 ligand, rhSCF, rhTPO (CC110, StemCell Technologies), all at a concentration of 100ng/mL. Following a 2 hour prestimulation, cells were incubated for 4 hours with 1.5x106 infectious units of each lentivirus, vHα7-IRES-GFP and vF-IRES-YFP, in the presence of 5µg/mL polybrene. 7.5x104 cells were then transplanted via tail vein injection into C57BL/6 recipients that had been lethally irradiated (11Gy) in two doses, nine hours and five hours prior to transplantation. Four weeks after transplantation, the tibialis anterior muscle of each recipient was injured via intramuscular injection of 100ng of notexin. Muscle tissue was then allowed to heal for four weeks prior to analysis. In the human monocyte transplantation model, U937 cells maintained in RPMI medium supplemented with 10% fetal bovine serum were infected with vHα7-IRESGFP  and  vF-IRES-YFP.  Infected  cells  were  sorted  (Becton-Dickinson  FACSVantage) and 1x105 cells were injected into the tibialis anterior muscle of NOD/SCID recipients. Muscle tissue was then allowed to heal for one week prior to analysis. For analysis, mice were first terminally anesthetized with avertin, then perfused with PBS containing 10mM EDTA and finally perfused with 4% PFA in PBS. All lower leg muscles were then removed from recipients and post-fixed in 4% PFA overnight prior to overnight cryoprotection in 20% sucrose. Muscle tissue was then embedded (OCT, Sakura) and cut into 20µm sections (Leica CM3050 S). In the 93  mouse bone marrow transplantation model, sections were stained overnight with 5bromo-4-chloro-3-indolyl β-D-galactopyranoside (X-gal) (Sigma). In the human monocyte transplantation model, sections were stained with rabbit anti-mouse laminin (Abcam) overnight at 4oC, followed by a 1 hour incubation with goat antirabbit Alexa 568 (Molecular Probes) at room temperature. In both cases the presence of donor-derived fibers was detected by examination using a Zeiss Axioplan2 microscope.  94  3.5 REFERENCES 1.  Blau, H.M., C.P. Chiu, and C. Webster, Cytoplasmic activation of human nuclear genes in stable heterocaryons. Cell, 1983. 32(4): p. 1171-80.  2.  Kohler, G. and C. Milstein, Continuous cultures of fused cells secreting antibody of predefined specificity. Nature, 1975. 256(5517): p. 495-7.  3.  Gong, J., et al., Induction of antitumor activity by immunization with fusions of dendritic and carcinoma cells. Nat Med, 1997. 3(5): p. 558-61.  4.  Pontecorvo, G., Production of mammalian somatic cell hybrids by means of polyethylene glycol treatment. Somatic Cell Genet, 1975. 1(4): p. 397-400.  5.  Zimmermann, U. and J. Vienken, Electric field-induced cell-to-cell fusion. J Membr Biol, 1982. 67(3): p. 165-82.  6.  Stromberg, A., et al., Manipulating the genetic identity and biochemical surface properties of individual cells with electric-field-induced fusion. Proc Natl Acad Sci U S A, 2000. 97(1): p. 7-11.  7.  Bakker Schut, T.C., et al., Selective electrofusion of conjugated cells in flow. Biophys J, 1993. 65(2): p. 568-72.  8.  Skelley, A.M., et al., Microfluidic control of cell pairing and fusion. Nat Methods, 2009. 6(2): p. 147-52.  9.  Alvarez-Dolado, M., et al., Fusion of bone-marrow-derived cells with Purkinje neurons, cardiomyocytes and hepatocytes. Nature, 2003. 425(6961): p. 96873.  10.  Camargo, F.D., et al., Single hematopoietic stem cells generate skeletal muscle through myeloid intermediates. Nat Med, 2003. 9(12): p. 1520-7.  11.  Warthin, A.S., Occurrence of numerous large giant cells in the tonsils and pharyngeal mucosa in the prodromal stage of measles - Report of four cases. Archives of Pathology, 1931. 11(6): p. 864-874.  12.  Okada, Y., Analysis of giant polynuclear cell formation caused by HVJ virus from Ehrlich's ascites tumor cells. I. Microscopic observation of giant polynuclear cell formation. Exp Cell Res, 1962. 26: p. 98-107.  13.  Dorig, R.E., et al., The human CD46 molecule is a receptor for measles virus (Edmonston strain). Cell, 1993. 75(2): p. 295-305.  14.  Tatsuo, H., et al., SLAM (CDw150) is a cellular receptor for measles virus. Nature, 2000. 406(6798): p. 893-7.  95  15.  Yin, H.S., et al., Structure of the parainfluenza virus 5 F protein in its metastable, prefusion conformation. Nature, 2006. 439(7072): p. 38-44.  16.  Esolen, L.M., et al., Apoptosis as a cause of death in measles virus-infected cells. J Virol, 1995. 69(6): p. 3955-8.  17.  Hallak, L.K., et al., Targeted measles virus vector displaying echistatin infects endothelial cells via alpha(v)beta3 and leads to tumor regression. Cancer Res, 2005. 65(12): p. 5292-300.  18.  Schneider, U., et al., Recombinant measles viruses efficiently entering cells through targeted receptors. J Virol, 2000. 74(21): p. 9928-36.  19.  Peng, K.W., et al., Oncolytic measles viruses displaying a single-chain antibody against CD38, a myeloma cell marker. Blood, 2003. 101(7): p. 255762.  20.  Allen, C., et al., Interleukin-13 displaying retargeted oncolytic measles virus strains have significant activity against gliomas with improved specificity. Mol Ther, 2008. 16(9): p. 1556-64.  21.  Nakamura, T., et al., Antibody-targeted cell fusion. Nat Biotechnol, 2004. 22(3): p. 331-6.  22.  Blanco-Bose, W.E., et al., Purification of mouse primary myoblasts based on alpha 7 integrin expression. Exp Cell Res, 2001. 265(2): p. 212-20.  23.  Hashiguchi, T., et al., Crystal structure of measles virus hemagglutinin provides insight into effective vaccines. Proc Natl Acad Sci U S A, 2007. 104(49): p. 19535-40.  24.  Colf, L.A., Z.S. Juo, and K.C. Garcia, Structure of the measles virus hemagglutinin. Nat Struct Mol Biol, 2007. 14(12): p. 1227-8.  25.  Palermo, A., et al., Nuclear reprogramming in heterokaryons is rapid, extensive, and bidirectional. FASEB J, 2009. 23(5): p. 1431-40.  26.  Terranova, R., et al., Acquisition and extinction of gene expression programs are separable events in heterokaryon reprogramming. J Cell Sci, 2006. 119(Pt 10): p. 2065-72.  27.  Pomerantz, J.H., et al., Reprogramming to a muscle fate by fusion recapitulates differentiation. J Cell Sci, 2009. 122(Pt 7): p. 1045-53.  28.  Long, M.A. and F.M. Rossi, Silencing inhibits Cre-mediated recombination of the Z/AP and Z/EG reporters in adult cells. PLoS One, 2009. 4(5): p. e5435.  96  29.  Lipton, B.H. and E. Schultz, Developmental fate of skeletal muscle satellite cells. Science, 1979. 205(4412): p. 1292-4.  30.  Arahata, K. and A.G. Engel, Monoclonal antibody analysis of mononuclear cells in myopathies. I: Quantitation of subsets according to diagnosis and sites of accumulation and demonstration and counts of muscle fibers invaded by T cells. Ann Neurol, 1984. 16(2): p. 193-208.  31.  Iankov, I.D., et al., Infected cell carriers: a new strategy for systemic delivery of oncolytic measles viruses in cancer virotherapy. Mol Ther, 2007. 15(1): p. 114-22.  97  CHAPTER 4. CONCLUSION 4.1 CONCLUSION Existing models of cell therapy for the treatment of skeletal myopathies are mainly based on the transplantation of satellite cells[1]. Unfortunately, these cells exhibit several properties which limit their clinical utility, namely immunogenicity[2], relative rarity[3], and inability to migrate from the circulation to muscle tissue[4]. Several groups have attempted to overcome these limitations via immunosupression[5], expansion in culture[6] and local injection[7], respectively. However, recent evidence demonstrates that expansion in culture significantly reduces the regenerative capacity of myoblasts[8]. Moreover, the large number of local injections required to systemically treat heritable myopathies remains a major limitation of these protocols and certainly hampers regeneration of poorly accessible sites, such as the diaphragm. Clearly, the treatment of skeletal myopathies would be greatly facilitated by an alternative, abundant source of cells capable of delivery via the circulation. The ability of bone marrow derived cells to participate in the repair of skeletal muscle initially generated hope that this phenomenon may represent an alternative means of cell therapy[9]. Unfortunately, this process has been shown to occur at exceedingly low frequencies, thereby precluding clinical application. At the outset of the work described here, the factors involved in the generation of bone marrow derived muscle were almost completely uncharacterized. As a result, early attempts to increase the efficiency of the process were ill founded and met with failure due to a lack of knowledge regarding the cell types and mechanisms involved[10, 11]. Therefore we designed a Cre/loxP based tracing strategy to identify the  98  hematopoietic lineages responsible for the generation of bone marrow derived muscle as well as a strategy to identify the role of fusion or transdifferentiation in the process (Figures 1.4 and 1.5). However, these strategies were not pursued upon discovery of inefficient labeling in our chosen Cre-reporter strain. We analyzed the labeling efficiency of hematopoietic cells in three Cre-reporter strains, Z/AP, Z/EG and R26R-EYFP[12-14]. Each of these reporter strains was capable of efficient hematopoietic labeling when exposed to Cre during early embryonic development. However, when Cre was expressed in adult hematopoietic cells, the labeling efficiency of the Z/AP and Z/EG reporter was much lower than the R26R-EYFP reporter. We subsequently demonstrated that in unlabeled adult hematopoietic cells derived from Z/AP and Z/EG mice, the transgenic promoter was methylated and Cre-mediated recombination of the locus was inhibited. In order to explain these data, we propose the following model (Figure 4.1).  99  Cre Expression Global DNA Methylation  A  PCX-NLS Cre  LysM Cre  0  B 3.5  DPC  C  D 7.5  Figure 4.1 Model illustrating the proposed methylation status of the Z/EG locus during early embryogenesis. (A) The Z/EG locus is methylated (white circles) and refractory to Cre-mediated recombination in a subset of adult cells. (B) Following fertilization, this epigenetic modification is removed during a genome wide demethylation process, which is known to peak at 3.5 days post-coitum (DPC). The demethylated locus may be recombined by Cre-recombinase present during this time. (C) Although we do not know the precise kinetics of the process, we hypothesize that if recombination does not occur within this temporal window, the βgeo transgene nucleates methylation of the locus during a genome wide methylation process, which is known to peak at 7.5 DPC. (D) This epigenetic modification then spreads to the promoter, silencing expression of β-geo and inhibiting Cre-mediated recombination. Thus, Cre transgenes expressed following this window, such as LysM-Cre, are unable to efficiently activate expression of the EGFP reporter.  100  After fertilization, the Z/AP and Z/EG loci are demethylated as a result of the genome wide demethylation that is known to occur in pre-implantation embryos[15]. This process provides a window wherein Cre recombinase may efficiently access the reporter locus and excise the β-geo transgene. However, if Cre-recombinase is not present during this developmental time window, the β-geo transgene remains intact and nucleates methylation of the locus as development proceeds. This prediction is based in part on the fact that removal of the β-geo transgene during early embryogenesis prevents silencing of the locus in adult cells (Figure 2.3A). We propose that methylation then spreads outward and inhibits both expression of β-geo transgene and Cre-mediated recombination of loxP sites. These effects may be mediated by direct inhibitory methylation of transcription factor binding sites[16] or loxP sites[17]. However DNA methylation is also known to be involved in the establishment and maintenance of a repressive chromatin state[18]. Therefore these effects may also be due to incorporation of the transgenic loci into heterochromatin. In support of this model, it should be noted that the percentage of cells expressing βgeo in both Z/AP and Z/EG mice (Figure 2.6) is remarkably similar to the percentage of cells expressing the post-excision reporter (Figure 2.3B) following exposure to Cre in adult cells. This observation suggests that the population of cells that express the pre-excision reporter may be the only cells capable of undergoing Cre-mediated activation of the post-excision reporter in adult cells. Admittedly, this model is largely based on correlative data. In order to establish a causative relationship, the methylation status of the Z/AP and Z/EG loci would need to be manipulated dynamically. However, at present it is not possible to alter the 101  methylation status of a single locus. Thus, the treatment required to manipulate DNA methylation would certainly result in an alteration of the expression level of many genes and therefore this experiment and would not directly link the methylation status of reporter loci to the efficiency with which they are recombined. Ultimately, this research provides a caveat for the future use of the Z/AP and Z/EG reporters as surrogate markers of excision or in lineage tracing experiments. However, a number of groups have employed the Z/EG reporter to investigate the role of fusion in the formation of donor-derived tissue in a manner similar to our initial specific aim[19, 20]. These reports suggest that fusion does not play a role in the formation of donor-derived epithelia or in the contribution of embryonic stem cells to chimeras and are based on the lack of EGFP expression following transplantation. While these findings may be valid, they should be reconsidered in the light of our data. Our final specific aim was to increase the contribution of bone marrow derived cells to skeletal muscle repair. Although our own efforts to identify the factors involved in the generation of donor derived muscle were precluded by the technical limitations described above, contemporary studies revealed that this phenomenon is primarily due to fusion of inflammatory cells with damaged myofibers[21, 22]. Unfortunately, despite its involvement in the formation and maintenance of a number of tissues including the placenta, osteoclasts and skeletal muscle itself, the process of cell fusion remains insufficiently understood to facilitate its enhancement for therapeutic purposes. Therefore, in order to specifically target and enhance the fusion of bone marrow derived cells with skeletal muscle fibers, we adapted a 102  fusogenic membrane glycoprotein complex derived from measles virus. Measles virus has recently emerged as a promising vector for the treatment of malignancies[23]. This oncolytic effect is based in part on the ability of the Edmonston B strain of measles virus to preferentially infect various human tumours based on elevated expression of CD46[24]. However, in an effort to further increase the specificity of this vector, Nakamura et al. have created a mutant measles hemagglutinin, H481A,533A, which is unable to bind its natural receptors and may be retargeted to virtually any cell surface antigen by the addition of a suitable polypeptide to its carboxyl-terminus[25]. Measles viruses expressing chimeric hemagglutinin proteins based on H481A,533A, have been shown to successfully infect malignant cells expressing the target antigen, leading to regression in a number of tumour models[26]. The measles virus fusogenic membrane glycoprotein complex is capable of mediating cell fusion in the absence of viral infection[27]. Therefore, in order to utilize this system for our purposes, we generated a single chain antibody, which recognizes the muscle specific integrin, alpha7, and fused it to the carboxyl-terminus of H481A,533A. This reagent, Hα7, specifically and efficiently mediated fusion of all cell types tested with skeletal muscle fibers in vitro and in vivo. When compared directly to polyethylene glycol, Hα7 consistently generated a ten to fifteen fold increase in heterokaryon yield and induced insignificant levels of toxicity. We also demonstrated that Hα7-mediated fusion results in the generation of true heterokaryons, which retain the capacity for nuclear reprogramming. Although our third specific aim was  103  originally focused entirely on increasing the frequency of fusion for therapeutic purposes, the striking efficiency of Hα7-mediated fusion in vitro was worth pursuing. A number of significant discoveries in the field of nuclear reprogramming have been made via fusion of various cell types with differentiated muscle in vitro[28, 29]. However, the low yield of existing fusogenic agents has generally prohibited advances in our understanding of this phenomenon. We anticipate that the increased yield and quality of heterokaryons generated via Hα7 mediated fusion will facilitate the systematic identification of the factors and mechanisms involved in the process of nuclear reprogramming. Finally, we performed proof of principle experiments demonstrating that Hα7 is able to increase the fusion of both mouse and human bone marrow derived cells with  skeletal muscle  in  vivo.  Unfortunately,  in  the  mouse  bone-marrow  transplantation model, the magnitude of this effect was limited by the low efficiency of lentiviral transduction. We suspect that this result may be due to inadequacy of the lentiviral vector itself and not due to expression of the measles virus proteins. Barrette et al. have demonstrated that ecotropic murine retroviruses may transduce mouse hematopoietic stem cells more efficiently than VSV-G pseudotyped lentiviral vectors[30]. Therefore, we are currently constructing MSCV-based retroviral vectors encoding Hα7 and F in an effort to increase the transduction efficiency of mouse bone marrow. However, transduction of long-term repopulating hematopoietic stem cells may not be necessary to meet the clinical goal. It is conceivable that a transient wave of circulating fusogenic cells derived from the transduction of short term hematopoietic progenitors may prove sufficient to deliver healthy or ex vivo  104  corrected nuclei to ailing myofibers. In fact, this strategy may even be preferable as it may avoid the risk of generating leukemias via insertional mutagenesis of hematopoietic stem cells[31]. Cell fusion itself has also been proposed to be a potential cause of malignancy[32]. According to this theory, the proliferation of hybrid cells results in aberrant chromosome segregation, genomic instability and stochastic loss of tumor suppressor genes, eventually leading to neoplastic transformation. However, empirical evidence for this model is lacking. While it is known that many tumor cells cell types are more fusogenic than their untransformed counterparts[33], it remains to be proven that cell fusion is an initiating event in tumorigenesis as opposed to a by product of the process. Furthermore, the Hα7-mediated fusion of bone marrow derived cells to skeletal muscle is unlikely to result in tumorigenesis as myofibers are post-mitotic and the simultaneous loss of a tumor suppressor from all syncytial myonuclei is extremely improbable. On the other hand, hybrid cells generated via Hα7 mediated fusion of bone marrow derived cells and satellite cells may be prone to aneuploidy following proliferation. However, we have not investigated the extent to which this type of fusion occurs or the viability of resultant hybrids. Clearly, these and other experiments should be performed in order to resolve whether malignancy is a legitimate concern for therapies involving cell fusion. At present, a more pragmatic limitation of Hα7-mediated fusion therapy may be the inability to treat a subset of human myopathies. While diseases caused by the simple absence of a single gene such as Duchenne muscular dystrophy would likely  105  be treatable by the integration of healthy nuclei into preexisting myofibers, treatment of other diseases with a dominant negative etiology including myotonic dystrophy[34] and oculopharyngeal muscular dystrophy[35] may require the generation of new myofibers exclusively composed of healthy nuclei. However, the autonomous generation of donor-derived myofibers in vivo has not been observed with any myogenic progenitor including myoblasts and may represent an intrinsic limitation to cell based therapies of skeletal myopathies. Fortunately, skeletal myopathies exhibiting a dominant mode of inheritance are relatively rare and are associated with comparatively mild symptoms[36]. Therefore we remain confident that the creation of circulating myogenic progenitors via expression of Hα7 on inflammatory cells will facilitate the development of novel cell and gene therapies for the majority of skeletal myopathies.  106  4.2 REFERENCES 1.  Peault, B., et al., Stem and progenitor cells in skeletal muscle development, maintenance, and therapy. Mol Ther, 2007. 15(5): p. 867-77.  2.  Guerette, B., et al., Lymphocyte infiltration following allo- and xenomyoblast transplantation in mice. Transplant Proc, 1994. 26(6): p. 3461-2.  3.  Snow, M.H., The effects of aging on satellite cells in skeletal muscles of mice and rats. Cell Tissue Res, 1977. 185(3): p. 399-408.  4.  Neumeyer, A.M., D.M. DiGregorio, and R.H. Brown, Jr., Arterial delivery of myoblasts to skeletal muscle. Neurology, 1992. 42(12): p. 2258-62.  5.  Karpati, G., et al., Myoblast transfer in Duchenne muscular dystrophy. Ann Neurol, 1993. 34(1): p. 8-17.  6.  Gussoni, E., et al., Normal dystrophin transcripts detected in Duchenne muscular dystrophy patients after myoblast transplantation. Nature, 1992. 356(6368): p. 435-8.  7.  Skuk, D., et al., Successful myoblast transplantation in primates depends on appropriate cell delivery and induction of regeneration in the host muscle. Exp Neurol, 1999. 155(1): p. 22-30.  8.  Montarras, D., et al., Direct isolation of satellite cells for skeletal muscle regeneration. Science, 2005. 309(5743): p. 2064-7.  9.  Sullivan, S. and K. Eggan, The potential of cell fusion for human therapy. Stem Cell Rev, 2006. 2(4): p. 341-9.  10.  Abbott, J.D., et al., Stromal cell-derived factor-1alpha plays a critical role in stem cell recruitment to the heart after myocardial infarction but is not sufficient to induce homing in the absence of injury. Circulation, 2004. 110(21): p. 3300-5.  11.  Musaro, A., et al., Stem cell-mediated muscle regeneration is enhanced by local isoform of insulin-like growth factor 1. Proc Natl Acad Sci U S A, 2004. 101(5): p. 1206-10.  12.  Lobe, C.G., et al., Z/AP, a double reporter for cre-mediated recombination. Dev Biol, 1999. 208(2): p. 281-92.  13.  Novak, A., et al., Z/EG, a double reporter mouse line that expresses enhanced green fluorescent protein upon Cre-mediated excision. Genesis, 2000. 28(3-4): p. 147-55.  107  14.  Srinivas, S., et al., Cre reporter strains produced by targeted insertion of EYFP and ECFP into the ROSA26 locus. BMC Dev Biol, 2001. 1: p. 4.  15.  Rougier, N., et al., Chromosome methylation patterns during mammalian preimplantation development. Genes Dev, 1998. 12(14): p. 2108-13.  16.  Watt, F. and P.L. Molloy, Cytosine methylation prevents binding to DNA of a HeLa cell transcription factor required for optimal expression of the adenovirus major late promoter. Genes Dev, 1988. 2(9): p. 1136-43.  17.  Rassoulzadegan, M., M. Magliano, and F. Cuzin, Transvection effects involving DNA methylation during meiosis in the mouse. EMBO J, 2002. 21(3): p. 440-50.  18.  Esteve, P.O., et al., Direct interaction between DNMT1 and G9a coordinates DNA and histone methylation during replication. Genes Dev, 2006. 20(22): p. 3089-103.  19.  Harris, R.G., et al., Lack of a fusion requirement for development of bone marrow-derived epithelia. Science, 2004. 305(5680): p. 90-3.  20.  Kidder, B.L., et al., Embryonic stem cells contribute to mouse chimeras in the absence of detectable cell fusion. Cloning Stem Cells, 2008. 10(2): p. 231-48.  21.  Camargo, F.D., et al., Single hematopoietic stem cells generate skeletal muscle through myeloid intermediates. Nat Med, 2003. 9(12): p. 1520-7.  22.  Doyonnas, R., et al., Hematopoietic contribution to skeletal muscle regeneration by myelomonocytic precursors. Proc Natl Acad Sci U S A, 2004. 101(37): p. 13507-12.  23.  Peng, K.W., et al., Systemic therapy of myeloma xenografts by an attenuated measles virus. Blood, 2001. 98(7): p. 2002-7.  24.  Anderson, B.D., et al., High CD46 receptor density determines preferential killing of tumor cells by oncolytic measles virus. Cancer Res, 2004. 64(14): p. 4919-26.  25.  Nakamura, T., et al., Antibody-targeted cell fusion. Nat Biotechnol, 2004. 22(3): p. 331-6.  26.  Nakamura, T., et al., Rescue and propagation of fully retargeted oncolytic measles viruses. Nat Biotechnol, 2005. 23(2): p. 209-14.  27.  Wild, T.F., E. Malvoisin, and R. Buckland, Measles virus: both the haemagglutinin and fusion glycoproteins are required for fusion. J Gen Virol, 1991. 72 ( Pt 2): p. 439-42.  108  28.  Blau, H.M., C.P. Chiu, and C. Webster, Cytoplasmic activation of human nuclear genes in stable heterocaryons. Cell, 1983. 32(4): p. 1171-80.  29.  Chiu, C.P. and H.M. Blau, 5-Azacytidine permits gene activation in a previously noninducible cell type. Cell, 1985. 40(2): p. 417-24.  30.  Barrette, S., et al., Lentivirus-based vectors transduce mouse hematopoietic stem cells with similar efficiency to moloney murine leukemia virus-based vectors. Blood, 2000. 96(10): p. 3385-91.  31.  Hacein-Bey-Abina, S., et al., LMO2-associated clonal T cell proliferation in two patients after gene therapy for SCID-X1. Science, 2003. 302(5644): p. 415-9.  32.  Duelli, D. and Y. Lazebnik, Cell fusion: a hidden enemy? Cancer Cell, 2003. 3(5): p. 445-8.  33.  Wakeling, W.F., J. Greetham, and D.C. Bennett, Efficient spontaneous fusion between some co-cultured cells, especially murine melanoma cells. Cell Biol Int, 1994. 18(3): p. 207-10.  34.  Philips, A.V., L.T. Timchenko, and T.A. Cooper, Disruption of splicing regulated by a CUG-binding protein in myotonic dystrophy. Science, 1998. 280(5364): p. 737-41.  35.  Brais, B., et al., Short GCG expansions in the PABP2 gene cause oculopharyngeal muscular dystrophy. Nat Genet, 1998. 18(2): p. 164-7.  36.  Emery, A.E., The muscular dystrophies. Lancet, 2002. 359(9307): p. 687-95.  109  APPENDIX A. ANIMAL CARE CERTIFICATES  110  111  112  113  

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