LITHUANIAN UNIVERSITY OF HEALTH SCIENCES MEDICAL ACADEMY
Arvydas Ūsas
REGENERATION OF INJURED BONE
AND ARTICULAR CARTILAGE USING
MOUSE MUSCLE DERIVED
STEM CELLS
Doctoral Dissertation
Biomedical Sciences, Medicine (07 B)
The investigations were performed at the Children’s Hospital of Pittsburgh and the University of Pittsburgh, Pittsburgh, Pennsylvania, USA during the period of 2000–2010, and the dissertation has been prepared at Kaunas University of Medicine during the period of 2009–2010.
Dissertation is defended extramurally.
Scientific Consultant:
Prof. Dr. Romaldas Mačiulaitis (Lithuanian University of Health Sciences, Medical Academy, Biomedical Sciences, Medicine – 07 B)
LIETUVOS SVEIKATOS MOKSLŲ UNIVERSITETAS MEDICINOS AKADEMIJA
Arvydas Ūsas
PAŽEISTO KAULINIO IR KREMZLINIO
AUDINIO REGENERACIJOS TYRIMAI
PANAUDOJANT PELĖS RAUMENINĖS
KILMĖS KAMIENINES LĄSTELES
Daktaro disertacija
Biomedicinos mokslai, medicina (07 B)
Disertacija ginama eksternu.
Mokslinis konsultantas:
Prof. dr. Romaldas Mačiulaitis (Lietuvos sveikatos mokslų universitetas, Medicinos akademija, biomedicinos mokslai, medicina – 07 B)
CONTENT
CONTENT ... 5
ABBREVIATIONS ... 8
1. INTRODUCTION ... 9
1.1. Study objective and specific aims ... 9
1.2. Research significance and novelty ... 9
1.3. Review of the literature ... 10
2. MATERIALS AND METHODS ... 19
2.1. Cell isolation and characterization ... 19
2.1.1. Isolation of muscle-derived cell populations and MDSC clones ... 19
2.1.2. Immunohistochemistry ... 20
2.1.3. RT-PCR analysis ... 20
2.1.4. Flow cytometry ... 21
2.1.5. Cytogenetic and tumorigenicity assay ... 21
2.1.6. In vitro cell stimulation with rhBMP-2 or rhBMP-4 ... 22
2.1.7. In vivo myogenic differentiation of the mc13 cells ... 23
2.2. Retroviral vectors and transduction of MDSCs ... 23
2.2.1. Construction of retroviral vectors ... 23
2.2.2. Transduction of MDSCs ... 24
2.2.3. BMP4 bioassay and detection of protein secretion ... 24
2.3. Evaluation of osteogenic and chondrogenic differentiation of transduced MDSCs in vitro ... 25
2.3.1. Detection of alkaline phosphatase activity after transduction ... 25
2.3.2. Chondrogenic stimulation in monolayer and type II collagen immunohistochemistry ... 25
2.3.3. Cell pellet culture and staining for glycosaminoglycans ... 26
2.4. Evaluation of osteogenic and chondrogenic differentiation of MDSCs in vivo ... 26
2.4.1. Intramuscular transplantation of MDSCs ... 26
2.4.2. Evaluation of β-galactosidase and osteocalcin expression ... 27
2.4.3. Immune response to allogeneic cell implantation ... 27
2.4.4. Alcian blue staining for cartilage formation ... 27
2.4.5. Skull defect model ... 28
2.4.6. Von-Kossa staining ... 28
2.4.7. Repair of osteochondral defects ... 28
2.4.8. Macroscopic evaluation and detection of LacZ transgene expression ... 29
2.4.9. Evaluation of β-galactosidase and type II collagen expression ... 29
2.5. Evaluation of osteogenic capacity of BMP4- and
VEGF-expressing MDSCs in vivo... 30
2.5.1. Ectopic bone formation ... 30
2.5.2. Skull defect healing... 30
2.5.3. Immunostaining for vascular structures ... 30
2.5.4. Apoptosis assay ... 31
2.6. Evaluation of chondrogenic capacity of BMP4- and VEGF-expressing MDSCs... 31
2.6.1. Pellet culture for in vitro chondrogenesis ... 31
2.6.2. Alcian blue staining ... 32
2.6.3. RT-PCR analysis ... 32
2.6.4. Repair of osteochondral defects ... 32
2.6.5. Histological evaluation and grading of cartilage repair ... 33
3. RESULTS... 34
3.1. Characterization of muscle-derived cell populations ... 34
3.1.1. Marker analysis of the pp6 cells ... 34
3.1.2. Marker analysis of the mc13 cell clone ... 35
3.1.3. Osteogenic differentiation after in vitro stimulation with rhBMP-2 .. 36
3.1.4. Osteogenic differentiation after stimulation with rhBMP4 ... 40
3.1.5. Standard cytogenetic and tumorigenicity assay ... 40
3.1.6. In vivo myogenic differentiation of the mc13 cells ... 41
3.2. Osteogenic potential of MDSCs ... 43
3.2.1. In vitro osteogenic differentiation of transduced MDSCs ... 43
3.2.2. Protein expression from transduced MDSCs ... 44
3.2.3. Ectopic bone formation induced by BMP4-expressing MDSCs ... 45
3.2.4. Immune reaction at the site of cell implantation ... 46
3.2.5. In vivo osteogenic differentiation of transduced MDSCs ... 47
3.2.6. Enhancement of bone healing in skull defect model ... 48
3.3. Chondrogenic potential of MDSCs ... 49
3.3.1. In vitro chondrogenic differentiation of MDSCs ... 49
3.3.2. Cell viability detected by LacZ transgene expression ... 51
3.3.3. In vivo chondrogenic differentiation of MDSCs ... 52
3.3.4. Macroscopic findings at the cartilage repair site ... 53
3.3.5. Histological evaluation of the cartilage repair ... 54
3.3.6. Histological grading of the cartilage repair ... 56
3.4. The effect of VEGF supply on BMP4 induced bone formation ... 56
3.4.1. Enhancement of endochondral bone formation ... 56
3.4.2. Enhancement of angiogenesis ... 59
3.4.3. Enhancement of skull defect healing ... 60
3.4.5. Importance of proper ratio of VEGF to BMP4 in bone regeneration . 64
3.4.5. Inhibition of bone formation by VEGF-specific antagonist sFlt1 ... 66
3.5. The effect of BMP4, VEGF and sFlt1 on chondrogenic potential of MDSCs ... 68
3.5.1. Expression of BMP4, VEGF, and sFlt1 by transduced MDSCs ... 68
3.5.2. Chondrogenesis in pellet culture ... 68
3.5.3. Macroscopic and histological evaluation of cartilage repair ... 71
3.5.4. Histological grading of the cartilage repair ... 74
4. DISCUSSION ... 76
CONCLUSIONS ... 84
CLINICAL RELEVANCE OF THE STUDY ... 85
PUBLISHED ARTICLES OF DISSERTATION ... 86
ABBREVIATIONS
ALP – alkaline phosphatase Bcl-2 – B-cell lymphoma 2 BMP2 – bone morphogenetic protein 2 BMP4 – bone morphogenetic protein 4 CD – cluster of differentiation CFU – colony forming units
DAPI – 4’, 6-diamidino-2-phenylindole DMEM – Dulbecco’s modified Eagle’s medium EDTA – ethylenediaminetetraacetic acid FACS – fluorescence-activated cell sorting FBS – phosphate buffered saline
FITC – fluorescein isothiocyanate MDSCs – muscle-derived stem cells MOI – multiplicity of infection PCR – polymerase chain reaction
pVSVG – plasmid vesicular stomatitis virus glycoprotein rhBMP2 – recombinant human bone morphogenetic protein 2 rhBMP4 – recombinant human bone morphogenetic protein 4 Sca-1 – stem cell antigen-1
SCID – severe combined immunodeficiency sFlt1 – soluble fms-like tyrosine kinase 1 TGFβ1 – transforming growth factor β1 TGFβ3 – transforming growth factor β3 VEGF – vascular endothelial growth factor
1. INTRODUCTION
1.1. Study objective and specific aims
Study objective:
To characterize muscle-derived stem cells (MDSCs) isolated from mouse skeletal muscle and evaluate their regenerative potential to repair injured bone and articular cartilage.
Specific aims:
1. Isolate MDSCs from mouse skeletal muscle, to determine their phenotypic characteristics and differentiation potential in vitro. 2. Evaluate osteogenic and chondrogenic regeneration capacity of
MDSCs genetically engineered to express bone morphogenetic protein 4 (BMP4).
3. Evaluate osteogenic and chondrogenic differentiation potential of MDSCs-expressing BMP4 in vivo.
4. Investigate the possibility for MDSCs-expressing BMP4 and vascular endothelial growth factor (VEGF) to enhance bone repair. 5. Investigate the possibility for MDSCs-expressing BMP4 and
anti-angiogenic factor sFlt1 to enhance articular cartilage repair.
1.2. Research significance and novelty
Skeletal muscle represents a convenient source of stem cells for cell mediated therapies based on cell transplantation, gene therapy and tissue engineering. Satellite cells, known as muscle stem cells, are myogenic precursors that can regenerate muscle and undergo self-renewal; however, these cells are committed to the myogenic lineage. Muscle-derived stem cells (MDSCs), a possible predecessor of the satellite cells, are considered to be a distinct population. MDSCs are early progenitors not limited to the myogenic lineage and are capable of differentiating into multiple lineages (osteogenic, chondrogenic, adipogenic, neural, endothelial, and hematopoietic). These cells display long-term proliferation, high self-renewal, and immune-privileged behaviors and are able to resist oxidative stress.
This investigation addresses important issues regarding: 1) stem cell isolation from skeletal muscle and their specific characteristics that enable us to identify these cells from other populations of stem cells in skeletal
muscle; 2) the efficiency of muscle-derived stem cell-mediated bone and cartilage formation in vivo using retroviral vectors to express bone morphogenetic protein 4 (BMP4); 3) the enhancement of osteogenic and chondrogenic potential of BMP4-secreting muscle-derived stem cells by addition of MDSCs engineered to express vascular endothelial growth factor (VEGF) or its antagonist, sFlt1.
The novelty of this investigation is that for the first time we show the existence of a clonal cell population (mc13) contained by highly purified muscle-derived cells (pp6) isolated from a mouse skeletal muscle based on cell adhesion to collagen coated flasks and phenotypic characteristics. We demonstrate that these clonal cells express both early myogenic progenitor markers (desmin, c-met, MNF, and Bcl-2) and stem cell markers (Sca-1, Flk-1). We show that these cells can survive better after transplantation, can avoid host immune responses, and exhibit a multipotent differentiation potential with the capacity to undergo myogenic, osteogenic, and chondrogenic differentiation in vitro and in vivo. We also demonstrate that by using gene therapy techniques and retroviral vectors carrying different growth factors, these clonal MDSCs can heal a critical-size bone defect created in the skull of a mouse, and an osteochondral defect created in a rat femur. Overall, we provide clear evidence for the existence of MDSCs in skeletal muscle and present a new strategy for the repair of various musculoskeletal tissues by using gene therapy and tissue-engineering based on MDSCs.
1.3. Review of the literature
Stem cells play a crucial role in the development and regeneration of various tissues. They are defined as unspecialized cells that are capable continuously renewing themselves, while maintaining the ability to differentiate into various cell types [1]. Stem cells are found at all developmental stages, from embryonic stem cells that can differentiate into all cell types found in the body, to adult stem cells that have a more limited differentiation capacity, and are responsible for post-natal tissue regeneration and growth [2-3]. The controversies regarding the use of embryonic stem cells for research are complicated by the ethical issue of using human embryos, also because of the evidence that embryonic stem cells can cause the formation of teratomas, and share genetic programs with cancer stem cells [4]. Postnatal adult stem cells are immunocompatible, have no ethical issues related to their use, and are more easily acquired. In addition, multiple studies have demonstrated that under certain
microenvironmental conditions or specific stimuli, adult stem cells can give rise to other cell types [5]. Multiple tissue types and organs in mammalian organisms have been identified as harboring adult stem cells. Among them include: bone marrow, skeletal and cardiac muscle, periosteum, synovial membrane, blood, brain, lung, liver, pancreas, digestive tract, skin, retina, breast, ovaries, prostate and testis. In general, the adult stem/progenitor cells are localized within a specialized microenvironment, designated as a niche, which consists of neighboring cells such as fibroblasts, endothelial cells and/or stromal components that tightly regulate their functions through the direct interaction and by the release of specific soluble factors [6]. Currently, there is no clear consent on whether each pool of stem cells is a separate entity or whether they are all descendents of one common stem cell that circulates and homes to tissues as needed [7]. In fact, growing evidence seems to suggest that the source of these post-natal adult stem cells is the blood vessel wall and that the number of stem cells found in a particular tissue correlates with the degree of vascularization within that tissue [8].
Several characteristics of skeletal muscle make it an attractive source of adult stem cells. Skeletal muscle comprises the largest proportion of total body mass of any adult tissue, making it readily available from almost any part of the body and can be obtained relatively simply through a minimally invasive biopsy. Also, skeletal muscle is a highly dynamic tissue that possesses an inherent ability to regenerate following damage caused by injury, exercise, immobilization, or by the administration of certain chemical agents. This regenerative capacity is due to the presence of a tissue-specific population of myogenic stem cells termed satellite cells, first discovered by Mauro in 1961. The satellite cell is so-called because of its peripheral location to the mature skeletal muscle cell, also called a multi-nucleated myofibre. Satellite cells are specifically located between the plasma membrane and surrounding basal lamina of the myofibres [9]. The identification of satellite cell markers such as the transcription factors Pax7 and Myf5 along with the cell surface markers M-cadherin and CD34 in murine muscle enabled researchers to demonstrate that satellite cells are a major source of skeletal myoblasts that are responsible for repopulating damaged and growing post-natal skeletal muscle as well as forming new myofibres [10]. Satellite cells remain quiescent in the muscle until an external stimuli such as injury occurs, at which point they re-enter the cell cycle, proliferate, and differentiate into myoblasts, which subsequently fuse to form new myofibres [11-12]. In addition, these cells harbor the ability to repopulate the stem cell niche through a self-renewal mechanism [13-16]. When stimulated with the appropriate growth factors in vitro, satellite cells are able to undergo osteogenic, chondrogenic, and adipogenic differentiation
which indicates their multipotentiality [17]; however, these cells express proteins, such as MyoD and Pax7, which also indicate their commitment to the myogenic lineage.
Over the last decade research has identified several other myogenic cell populations, including muscle-derived stem cells (MDSCs), side-population (SP) cells, pericytes, and mesangioblasts, that can participate in muscle regeneration under experimental conditions. The variations in isolation methods make it difficult to determine and/or directly compare the origin and localization of these stem cell populations. In addition, their relationship to satellite cells remains unknown, as does their relevance to muscle regeneration under physiopathological conditions [18]. Recent findings indicate that the MDSCs may originate from vascular and perivascular regions in the muscle [19-20], and suggest that their ultimate origin is the walls of blood vessel [21].
Different methods have been used to separate distinct muscle-derived cell populations including by their adhesiveness, proliferation behaviors, and stem cell marker expression profiles. These methods include density-gra-dient fractionation, pre-plating culture series, repeated culture using freeze-thaw technique, magnetic cell sorting (MCS), and fluorescence-activated cell sorting (FACS). MCS and FACS have been employed to sort cells on the basis of their surface marker profiles [22-23] or their ability to exclude Hoechst dye [24-25]. Muscle-derived stem and progenitor cell populations are defined by cell markers, some of which are restricted to these populations, whereas other markers are shared with other populations. However, marker-profile-based isolation methods are limited because they rely exclusively on the expression of cell surface markers that are variable and may change under cell culture conditions [26]. Presently, the most prevalent method to isolate MDSCs is a marker-profile-independent modified preplate technique which is based on variable adherence of freshly dissociated muscle cells to collagen-coated flasks [27-30]. A skeletal muscle biopsy is digested using dispase, collagenase, and trypsin, and the resulting single cell suspension is plated onto a series of collagen-coated flasks. The cells that adhere early, within 12-24 hours after plating, are known as rapidly adhering cells (RACs) and are comprised of fibroblastic-like cells. Cells that adhere between 48-96 hours are known as slowly adhering cells (SACs) that contain myoblasts already committed to the myogenic lineage. The SACs that are slowest to adhere contain a population of early myogenic progenitor cells, and after continuing to further passage the SACs, we obtained population of long term proliferating cells, known as MDSCs [29-31]. The MDSC population has been characterized using immunostaining for certain stem cell markers in vivo and in vitro, and also by FACS. Several
modifications of this preplate technique have been performed to isolate muscle-derived cells from various species [32-33]. The variations include the use of different types of enzymes to digest the tissue, different types of material used to coat the tissue culture flasks, and different periods of time between each preplate.
Although MDSCs are similar to satellite cells in their ability to regenerate skeletal muscle, MDSCs are a separate population of cells that expresses distinct markers and phenotypes [26, 31]. Satellite cells, whether active or quiescent, typically express the paired box-7 (Pax-7) gene [34-36], whereas MDSCs are more heterogenous but express stem cell antigen-1 (Sca-1) consistently, and often express cluster of differentiation 34 (CD34). Pax-7 and Sca-1 positive cells have not been found to co-localize in skeletal muscle, providing further evidence that satellite cells and MDSCs are distinct populations [37]. MDSCs also appear to be more primitive than satellite cells, as they have the ability to differentiate into a broader number of cell types, thus displaying greater plasticity [31, 38]. MDSCs are a unique cell population whose characteristics, including marker profile, proliferation and differentiation kinetics, and regenerative capacity, are distinct from myoblasts [39-40]. It is likely that MDSCs are situated hierarchically upstream of Pax7+ cells and constitute the initial stage during myogenic differentiation [21, 31, 41-42].
In vivo studies provide additional evidence for the stem cell nature of
MDSCs. It has been demonstrated that MDSCs can self-renew, differentiate into multiple lineages, and have the potential to regenerate various adult tissues [43-46]. In fact, a recent study revealed that MDSCs have an extended replicative lifetime and a substantial self-renewal capacity comparable with that exhibited by embryonic stem cells [40]. They retained their phenotype and ability to promote muscle regeneration in vivo when cultured up to 200 population doublings. MDSCs also display a superior regenerative capacity relative to satellite cells following their transplantation into dystrophic muscle in a murine model of muscular dystrophy (mdx) [26, 31, 40]. MDSCs appear to also be at least partially immune-privileged, as transplantation with MDSCs results in dystrophin expression in mdx mice (dystrophin-deficient) for over 3 months after injection [31]. It has been reported that stem cells exhibit enhanced resistance to oxidative stress and may avoid the oxidative damage associated with transplantation and the host immune response [47-50]. Indeed, MDSCs showed lower rates of oxidative and inflammatory stress-induced cell death which could, at least partially, explain their increased regenerative capacity and to some extent justify the sex related differences observed between male and female MDSCs in myogenic differentiation and skeletal muscle regeneration [51-52]. When
compared to myoblasts, MDSCs implanted into infarcted hearts exhibited greater and more persistent engraftment, induced more neoangiogenesis, prevented cardiac remodeling, and elicited significant improvements in cardiac function [46, 53]. MDSCs were also found to be more resistant to oxidative stress induced apoptosis and ischemia in comparison to myoblasts and cardiomyocytes which may explain their increased survival and engraftment capacity when transplanted into the injured area of the heart [46, 53]. In a lethally irradiated mouse model, implantation of a subpopulation of MDSCs expressing cluster of differentiation 117 (c-kit) and Sca-1 was able to rescue the animal indicating that these cells have the ability to differentiate into hematopoietic cells [54]. In addition, it has been shown that donor-derived bone-marrow cells isolated from recipient mice by FACS, could repopulate the bone marrow of secondary, lethally irradiated animals and retained their myogenic differentiation capacity both
in vitro and in vivo [43].
MDSCs could be used in a variety of applications utilizing cellular therapy. MDSCs are particularly attractive for cell therapy purposes because they are easy to isolate, are adult-derived, are able to self-renew, can differentiate into multiple lineages, express greater levels of important paracrine factors (i.e. VEGF) than their more differentiated counterparts such as myoblasts, and are, at least to some degree, protected from host immune responses after implantation [40, 50]. A human counterpart to the murine MDSCs has recently been isolated based on expression of both myogenic and endothelial cell surface markers. These cells have been shown to be better than cells that express only myogenic cell markers or endothelial cell markers for both skeletal muscle repair [19] and cardiac repair following myocardial infarction [55].
There are many potential uses for MDSCs in cell replacement-based therapies, and promising results have been obtained in treating muscle atrophy associated with aging, muscle wasting and various musculoskeletal and neuromuscular degenerative disorders such as Duchene and Becker muscular dystrophies and amyotrophic lateral sclerosis as well as with cardiovascular and urological degenerative disorders [21, 56-57]. Currently, autologous human muscle-derived cells being tested in a clinical trial for the treatment of urinary incontinence [58] and phase I trials for cardiac repair after a myocardial infarction will begin in the near future.
The formation of extra-skeletal bone in muscle is a widely observed phenomenon suggesting that skeletal muscle contains osteoprogenitor cells [59]. Several studies demonstrated the feasibility of using primary muscle derived cells in ex vivo gene therapy to produce bone [60-62]. It has been shown that BMP2 prevents in vivo myogenic differentiation, promotes
osteogenic differentiation and has no adverse effect on cell survival of muscle-derived cells virally transduced with the BMP2 gene [63]. Retrovirally transduced MDSCs expressing BMP4 were found superior in terms of their capacity to induce more endochondral bone formation compared to other, less purified, muscle derived cell populations transduced similarly. The improved bone formation capacity of the transduced MDSCs has been linked to prolonged cell survival due to enhanced proliferation and lower immunogenicity [44].
There are a variety of orthopedic conditions that would be treatable with cellular therapy using muscle-derived cells, including delayed fracture unions or non-unions, segmental bone loss resulting from injury or tumor removal, and articular cartilage degeneration due to traumatic injury or osteoarthritis. Since the formation of new bone with the appropriate mechanical properties is critical for bone repair, research has focused on scaffolds and cells that release paracrine factors to induce osteogenesis. While efforts are still being made to engineer biomaterials capable of releasing osteoinductive factors, cell-based growth factor delivery appears to be the most controllable system. Various growth factors have been investigated for their capacity to improve bone healing. The bone morphogenetic proteins (BMPs) are cytokines that belong to the transforming growth factor beta (TGF-β) family well known for their capacity to induce bone formation. The activity of BMPs was discovered by Urist in the 1960s, who described a substance in bone matrix that had inductive properties for the development of bone and cartilage [64]. To date, at least 20 BMPs have been identified that possess varying degrees of inductive activities, some of which have been shown capable of stimulating stem cell differentiation into osteoblasts [65]. BMP signaling is required for endochondral ossification and maintenance of the adult skeleton [66]. Bone morphogenetic protein 4 (BMP4) has important roles in embryonic develop-ment and bone formation. It appears to be an early stimulator of endochond-ral ossification and is highly expressed during the early phases of fracture healing [67]. BMP4 mRNA was detected in primitive mesenchymal and chondrocytic cells, the cambium layer of the periosteum, the bone marrow cavity, and skeletal muscle adjacent to the fracture site [68].The genetic sequences of the BMPs have been identified, and recombinant gene technology has been used to produce these proteins for clinical application. Osteogenic BMPs, including BMP2, BMP4, and BMP7, have been used to induce bone formation and repair bone defects in various animal models [69-70]. BMP2 and BMP7 already have clinical approval for use in cases of non-unions, open tibial fractures, and spinal fusions [71-74].
Although direct application of exogenous BMP can stimulate bone growth in vivo, large quantities of the protein are required and it results in bone growth that is structurally less hardy in comparison to bone induced by gene therapy technique which enables the delivery of sustained quantities of the desired protein [75]. The ex vivo approach to deliver different BMPs to enhance bone healing has been employed by many investigators [70, 76-77]. Different cell lines and primary cells isolated from the bone marrow, skeletal muscle, periosteum, articular cartilage and skin have all been utilized as vectors [61, 78-79].
Bone regeneration involves several cellular pathways. One such pathway, angiogenesis mediated by vascular endothelial growth factor (VEGF), is important for bone formation during endochondral ossification in bone development [80-82]. Recent findings imply that VEGF is not only involved in endochondral ossification, but also implicated in intramembraneous ossification [83-85]. VEGF promotes angiogenesis, stimulates and affects chemotaxis, proliferation, survival, and the activity of osteoblasts, osteoclasts, and chondrocytes [86-89]. VEGF blockade leads to the suppression of blood vessel invasion, impairs cartilage resorption, and reduces trabecular bone formation at the growth plate [80]. Inhibition of VEGF activity decreases blood flow, delays bone repair and may lead to nonunion. Recent studies demonstrated that combined delivery of angiogenic and osteogenic growth factors improved blood vessel formation and bone growth, suggesting interplay between these growth factors for early bone regeneration [90-93]. Although the interactions between the BMPs and VEGF in vitro have been previously established [94], the effects and mechanisms of these interactions during in vivo bone formation remained poorly understood. In order to investigate whether VEGF function is essential for bone formation induced by osteogenic BMPs, and which cellular mechanisms are involved in this interaction, an ex vivo approach using cells as gene delivery vehicles to deliver osteogenic and angiogenic genes was employed. MDSC-mediated delivery of exogenous VEGF enhanced BMP2-induced endochondral bone formation and the healing of a skull defect in mice by improving angiogenesis, which consecutively led to accelerated cartilage resorption and augmented mineralized matrix deposition [95]. This study and others also showed that the synergistic interaction between VEGF and BMP2 depends on their expression ratio, with higher proportions of VEGF leading to reduced synergism [95-96]. Most recent reports demonstrated that mesenchymal stem cells expressing BMP2 and VEGF were able to enhance the healing of a segmental tibiae defect in mice after systemic delivery [97].
Articular cartilage is a naturally avascular tissue and has a very limited capacity to heal itself. None of the currently available treatment options, including abrasion arthroplasty, microfracture, meniscal or osteochondral allografts, and autologous chondrocyte implantation (ACI), can fully restore damaged cartilage. The use of autologous chondrocytes, a natural choice for cartilage repair, is limited due to donor site morbidity, the inability to harvest a sufficient number of cells, and the loss of the cell’s chondrocytic phenotype during long-term in vitro culturing which results in the formation of a fibrocartilage [98-99]. Adult mesenchymal stem cells (MSCs) derived from bone marrow, periosteum, synovium or fat are increasingly being considered as promising alternative cell types to differentiated chondrocytes for use in cell-based cartilage repair strategies because of their chondrogenic potential [98, 100]. MSCs are available in larger quantities and are easier to isolate culture and manipulate ex vivo compared to autologous chondrocytes. Skeletal muscle-derived cells also constitute an attractive alternative for articular cartilage repair. It has been demonstrated that purified muscle cells can repair cartilage defects to a degree equivalent to chondrocyte transplantation [101]. Although that pilot study did not provide any evidence regarding muscle cell differentiation into chondrocytes, an assumption was made that a population of stem cells existing in the skeletal muscle probably contributed to the repair process. It has been shown that purification for Sca-1 may further enhance chondrogenic potential of stem cells, including MDSCs [102-104]. In addition, male MDSCs have been found to have a superior chondrogenic differentiation potential and cartilage regeneration potential compared to female MDSCs [104].
Several growth factors, including transforming growth factor β (TGFβ), BMP2, insulin-like growth factor 1 (IGF1), and basic fibroblast growth factor (bFGF), have been able to improve chondrocyte proliferation and extracellular matrix synthesis [105-107]. BMP4 also was identified as a promising candidate for the enhancement of chondrogenesis [98, 108-110]. Cells overexpressing growth factor genes represent a novel treatment option for cartilage tissue engineering [111].
The control of angiogenesis during chondrogenic differentiation is one of the most important issues affecting the application of stem cells for cartilage repair. In the growth plate, VEGF has been reported to play an essential role in cartilage vascularization and absorption of hypertrophic chondrocytes, which together lead to endochondral ossification [80, 82]. Previous research has shown that VEGF expression by chondrocytes in osteoarthritic joints may be related to articular cartilage destruction and osteophyte formation during osteoarthritis development also involves VEGF signaling [112-114]. Recent findings reveal the expression of VEGF and its receptors (Flt1 and
Flk1) in osteoarthritic cartilage and reflect the ability of VEGF to enhance catabolic pathways by stimulating matrix metalloproteinase (MMP) activity and reducing tissue inhibitors of MMPs [114-116]. These findings suggest that, apart from the effect of VEGF on cartilage vascularization and proliferation of cells in the synovial membrane, chondrocyte-derived VEGF promotes catabolic pathways in the cartilage itself, thereby leading to a progressive breakdown of the extracellular matrix of articular cartilage. Blocking VEGF signaling and inhibiting the bioactivity of VEGF after osteochondral damage has been shown to suppress chondrocyte apoptosis and improve articular cartilage repair [117-118].
2.
MATERIALS AND METHODS
2.1. Cell isolation and characterization
2.1.1. Isolation of muscle-derived cell populations and MDSC clones
Primary muscle cell cultures were obtained from 3-wk-old mdx (mouse model for Duchene Muscular Dystrophy) mice (C57BL/10ScSn mdx/mdx, The Jackson Laboratories) using a modified version of a previously described preplate technique [27-28, 119]. Briefly, the hindlimb muscles of mice were removed and minced into coarse slurry using fine scissors. The muscle tissue was enzimatically dissociated at 37 °C in 0.2% collagenase-type XI (Sigma-Aldrich) for 1h, in dispase (GIBCO BRL), prepared as 2.4 units/1ml HBSS, for 45 min, and then incubated in 0.1% trypsin-EDTA (GIBCO BRL) diluted in HBSS for 30 min. Each step was followed by centrifugation at 3500 rpm for 5 min. The isolated cells were resuspended in proliferation medium (DME containing 10% horse serum, 10% FBS, 0.5% chick embryo extract, and 1% penicillin-streptomycin) and plated on collagen type 1-coated flask. After 1–2 hours, the supernatant containing nonadherent cells was withdrawn from the flask (pp1) and transferred to a new flask (pp2). After 24 h, the floating cells in pp2 were collected, centrifuged, and plated on new flask (pp3). The serial replating procedure was repeated at 24 h intervals until preplates (pp4-pp6) were obtained. It took an additional 24–72 h for the pp6 cell population to adhere to collagen-coated flask after transfer from pp5. The obtained pp6 culture was enriched with small, round cells (Fig. 2.1.1.1).
Fig. 2.1.1.1 Schematic presentation of muscle-derived stem cell
Most of the pp6 cells die within 1–2 wk of culturing, with very few of the adherent surviving cells proliferating and forming clonal colonies. To isolate clones from highly purified pp6 cell population, the cells were transfected with 10 µg of the linear plasmid containing β-galactosidase (lacZ), minidystrophin and neomycin resistance gene using the lipofectamine reagent (GIBCO BRL) according to the manufacturer’s instructions. After 72 h cells were selected with 3000 µg/ml of G418 (GIBCO BRL) for 10 days until colonies appeared. Colonies were carefully collected, expanded to obtain lar-ge quantity of the transfected cells, and tested for the expression of lacZ and dystrophin genes. One of these clones, mc13, was selected for further studies.
2.1.2. Immunohistochemistry
Cells were seeded in a 6-well plate and fixed with cold methanol for 1 min. After rinsing with PBS, cells were blocked with 5% horse serum at room temperature for 1 h. The primary antibodies were mouse anti-desmin (1:100; Sigma-Aldrich), biotin anti-mouse CD34 (1:200; BD PharMingen), biotin anti-Sca-1 (1:200; BD PharMingen), rabbit anti-mouse Bcl-2 (1:1000; BD PharMingen), rabbit anti-mouse Flk-1 (1:100; Research Diagnostics), rabbit anti-mouse m-cadherin (1:50), mouse anti-MyoD (1:100; BD PharMingen), and mouse anti-rat myogenin (1:100; BD PharMingen). The primary antibodies were applied overnight and appropriate secondary antibodies were applied for 1h at room temperature. The cells were rinsed with PBS and then incubated with 1:300 streptavidine conjugated with Cy3 fluorochrome for 1h at room temperature. The percentage of positive cells was calculated by counting cells expressing specific marker in 10 randomly chosen 20× fields.
2.1.3. RT-PCR analysis
Total RNA was isolated using TRIzol reagent (Life Technologies). Reverse transcription (RT) was performed using SuperScript™ preamplifi-cation system for first strand cDNA synthesis (Life Technologies) according to the manufacturer’s instructions. PCR amplification of the targets was performed in 50 μl reaction mixture containing 2 μl of RT-material, 5 U / 100 μl Taq DNA polymerase (Life Technologies), and 1.5 mM MgCl2. CD34
primers were designed by using Oligo software. The sequences of the other primers are from references as follows: myogenin [120], c-met [121] and MNF [122]. Following parameters were used: 94 °C 45 s, 50 °C (for CD34) 60 s; 60 °C (for myogenin and c-met) 60 s; and 58 °C (for MNF) 60 s, 72 °C 90 s for 40 cycles. PCR products were checked by 1% agarose-TBE-ethidium
bromide gels. The expected products sizes are: CD34, 147 bp; myogenin, 86 bp; c-met, 370 bp and MNF, 305 bp. To exclude genomic DNA contamina-tion, two controls were used: parallel RT without reverse transcriptase, and amplification of β-actin using a primer set that spans an intron (Clonetech).
2.1.4. Flow cytometry
Cultured cells were trypsinized, spun, washed, and counted. The cells were then divided into two aliquots (experimental and control) and spun to a pellet. A 1:10 mouse serum (Sigma-Aldrich) in PBS solution and Fc Block (rat anti-mouse CD16/CD32; BD PharMingen) was used to re-suspend each pellet, and the suspensions were incubated for 10 min on ice. Optimal amounts of rat anti-mouse monoclonal antibodies were predetermined and added directly to each tube for 30 min. Each experimental tube received FITC-conjugated CD45, R-PE-conjugated CD117(c-kit) and biotin-conjugated Sca-1 (all from BD PharMingen). Control tubes for each cell type received equivalent amounts of FITC-conjugated, biotin-conjugated, and R-PE-conjugated isoty-pe standards (BD PharMingen). After several rinses, streptavidin-allophyco-cyanin conjugate (APC; BD PharMingen) was added to each tube, including controls, and incubated on ice for 20 min. Just before analysis, 7-amino-actinomycin D (7-AAD, Via-Probe, BD PharMingen) was added to each tube for dead cell exclusion. A minimum of 10,000 live cell events were analyzed on a FACSCalibur® (Becton Dickinson) flow cytometer using Cell Quest software.
2.1.5. Cytogenetic and tumorigenicity assay
The standard cytogenetic method for metaphase preparation of mc13 was performed using a previously described protocol [123]. Briefly, the cells were grown to near confluency in a T75 collagen coated flask in DMEM supplemented with 10% horse serum, 10% FBS, 0.5% chick embryo extract and 1% Pen-Strep solution (all from GIBCO BRL). Cells had 15 ml of fresh medium added along with 0.2 ug/ml Colcemid Solution (GIBCO BRL) and were allowed to incubate an additional 2 hr. at 37 °C. Cells were then harvested by adding 0.1% Trypsin-EDTA to the cells until they lifted. Cells were pelleted and resuspended in 5 ml 0.75 M KCl and incubated 8 min at 37 °C. 1 ml Carnoy’s fixative (3:1 methanol to glacial acetic acid) was next added to solution and the cells pelleted. Fresh fixative was added and cells were again pelleted. This rinsing and pelleting was performed 3 times. Cells were then dropped on glass slides, heated at 60 °C for 30 min and GTG banded. 32 metaphase cells were then counted for modal number using a
magnification of 630X. Two images were capture on a Cytoscan cytogenetics analyzer.
The technique of cell growth on soft agar was performed as previously described [124]. A 1.275% bacto-Agar (Difco) solution was first prepared in distilled water and sterilized in an autoclave. The agar medium containing 20 ml of culture medium at twice the concentration used was mixed with 20 ml of 1.275% bacto-Agar at 45 OC and 10 ml of Fetal Bovine Serum (FBS). Petri dishes (60 mm) were filled with 5 ml of agar medium and the agar was allowed to solidify during 1 hour at room temperature. The cells to be tested were trypsinized and resuspended in the culture medium with 15 % FBS to obtain a concentration of 500,000 cells per ml. This suspension was passed twice through a 20G needle to dissociate any aggregate. One million cells (2 ml) were then mixed with 5 ml of agar medium. A sample of 1.5 ml of this agar cell suspension was finally plated over the solidified agar in the Petri dishes. When the new agar layer containing the cells has solidified, 1 ml of culture medium was added to the Petri dishes. The culture medium overlaying the agar was changed once a week for two weeks. The cell behavior was monitored every two days and picture was taken. Different cell populations were tested: mc13 late passages (>20), mc13 early passage (<5), HEK293 cells (adenovirus permissive cells), and a freshly isolated primary myoblast cell culture.
2.1.6. In vitro cell stimulation with rhBMP-2 or rhBMP-4
The mc13 and non purified muscle derived cells were plated in triplicate at a density of 1–2 × 104 per well in 12 well collagen-coated flasks. The cells were stimulated by addition of 200 ng/ml recombinant human BMP-2 (rhBMP-2) to the media. The media was changed on days 1, 3, and 5 after initial plating. The control group also had media changed on these days, without rhBMP-2. After 6 days of stimulation, cells were counted using a hemacytometer, and cell lysates were harvested by repeated freeze-thaw cycles. The alkaline phosphatase (ALP) activity in the cell lysate was measured using a commercially available kit (Sigma-Aldrich) which utilizes color change in the reagent due to the hydrolysis of inorganic phosphate from p-nitrophenyl phosphate. The color change was analyzed on a spectrophotometer, and the data was expressed in internatio-nal units ALP activity per liter normalized to 106 cells (U/L/mil cells). Statistical difference among the different groups was analyzed using student’s t-test (*p<0.05). Stimulation with rhBMP-4 was performed similarly.
2.1.7. In vivo myogenic differentiation of the mc13 cells
The mc13 cells were injected (5 × 105 cells) intramuscularly in the hind limb muscle of mdx mice. The mdx mice were sacrificed at 7 days post-injection, and the injected muscles were frozen, cryostat sectioned, and assayed for dystrophin and LacZ expression [28]. For dystrophin staining, sheep-anti-human D-10 antibody (1:250 dilution in PBS) was used as the primary antibody. Following several rinses in PBS, a biotin-conjugated anti-sheep was subsequently used (1:250 dilution in PBS). Streptavidin-FITC (bone) and streptavidin-Cy3 (muscle) at a dilution of 1:250 in PBS were used; immunoreaction was obser-ved by fluorescence microscopy (Nikon, Eclipse E-800). For LacZ staining, the sections were fixed in 1% glutaraldehide, incubated overnight with X-gal substrate at 37 °C, and counterstained with eosin. The mc13 cells (5 × 105 cells) were also injected intravenously in the tail vein of mdx mice. The mice were sacrificed at 7 days post-injection and various tissues, including the hind limb muscle, lung, liver, spleen, kidney and brain, were isolated and assayed for dystrophin and LacZ using the same protocol described above.
2.2. Retroviral vectors and transduction of MDSCs
2.2.1. Construction of retroviral vectors
Retroviral vectors containing BMP2/4 constructs were generated by subclo-ning the BMP2/4 constructs from pBluescript II KS(-) into CLX. Vector CLX was constructed by removing the SV40 promoter and the Neo resistant gene from LXSN (provided by A.D. Miller) and replacing the U3 region in the 5’ long-terminal repeat with a cytomegalovirus (CMV) promoter to increase the titer of virus produced by transient transfection. The resultant vectors were termed CLBMP2/4.
Human BMP4 and BMP2 cDNAs were amplified by PCR from the respective phage clones (ATCC). A Kozak sequence was introduced to the 5’ end of each cDNA to enhance protein expression in mammalian cells. BMP4-1 and BMP4-2 primers were used to generate the BMP4 expression construct. Similarly, BMP2-1 and BMP2-2 primers were used to amplify the BMP2 construct. Following cloning into pBleuscript II KS(-), sequences of selected clones were verified by sequencing both strands of cDNA.
The BMP2/4 hybrid construct, in which the original sequence coding for the pro-peptide of BMP4 was replaced by the corresponding pro-peptide sequence of the BMP2 cDNA, was generated as described previously [125]. This modification led to a markedly higher level of BMP4 secretion in transfected cells. The modified BMP4 cDNA was subcloned into retroviral
vector pCLX, derived from pLXSN (from A. Dusty Miller of Fred Hutchinson Cancer Research Center) by removing the SV40 promoter and the Neomycin resistant gene, and replacing the U3 in the 5' LTR with the human CMV promoter. The resultant vector was termed pCLBMP4. Retro-viral vectors expressing human VEGF, sFlt1, or the bacterial LacZ gene were constructed by cloning the VEGF165 cDNA (from InvivoGen), cDNA
encoding human sFlt1 (from InvivoGen), or the LacZ gene into pCLX, resulting in pCLVEGF, pCLsFlt1, and pCLlacZ, respectively.
Vector DNA was converted into replication-defective retrovirus by co-transfection, with calcium-phosphate precipitation, into packaging cell line GP-293 (Clontech) with a plasmid, pVSVG, which expressed vascular stomatitis virus glycoprotein as the viral envelope. The titer of the viral vectors was estimated to be 5×105 to 2×106 CFU/ml by limiting dilution. The retroviruses expressing BMP4, VEGF, sFlt1, or LacZ were termed retroBMP4, retroVEGF, retrosFlt1, and retroLacZ, respectively.
2.2.2. Transduction of MDSCs
The MDSCs (mc13 clone) cells were transduced separately with retrovi-ral vectors at a multiplicity of infection of 5 in the presence of 8 μg/ml polybrene. The transduced cells were expanded for 2 weeks before being used in animal experiments. At the end of expansion, cells and the condi-tioned media were sampled to determine the level of transgene expression. For the cartilage repair study some of the LacZ-transduced cells then were transduced with the retroBMP4; this process resulted in co-expression of LacZ and BMP4 by the MDSCs.
2.2.3. BMP4 bioassay and detection of protein secretion
The level of functional BMP4 secreted from the transduced cells was estimated with a BMP4 bioassay [125]. On day 1, C2C12 myoblast clones were seeded at 8×103 cells/100 μL/well in a 96-well plate and allowed to grow overnight. The following day, growth medium from the transduced cells was collected and stored temporarily at 4 °C. Overnight growth me-dium was removed from the C2C12 plates, and 50 μL of fresh meme-dium was added to each well. Eight serial 2-fold dilutions of the growth medium from transduced cells were prepared in a 96-well plate, and 50 μL of the dilutions were added to corresponding wells of the C2C12 plate. Each medium sample was tested in triplicate. A similar 2-fold serial dilution of rhBMP4 (RDI), starting from 200 ng/well, was used as a positive control. Plates were
incubated overnight. On day 3, the C2C12 plates were prepared for ALP lysate assay. The levels of VEGF or sFlt secreted from the transduced cells were measured using enzyme-linked immunosorbent assay (ELISA) kits (R&D).
2.3. Evaluation of osteogenic and chondrogenic
differentiation of transduced MDSCs in vitro
2.3.1. Detection of alkaline phosphatase activity after transduction
Transduced and control cells were counted weekly using a hemacytome-ter, and cell lysates were harvested by 2 freeze/thaw cycles. Alkaline phosphatase (a pre-osteogenic marker) activity of the cell lysate was measu-red using a commercially available Alkaline Phosphatase (AP) Leukocyte kit (Sigma-Aldrich) that uses color change in the reagent secondary to hydrolysis of inorganic phosphate from p-nitrophenyl phosphate. The degree of color change was analyzed spectrophotometrically, and the data were expressed in international units of ALP activity per liter normalized to 1 million cells (U/L/×106 cells).
2.3.2. Chondrogenic stimulation in monolayer and type II collagen immunohistochemistry
MDSC and MDSC-B4 were plated at a density of 2500 cells/chamber on Falcon culture slides (Becton Dickinson Labware) and cultured in 1 of 3 types of media: 1) normal differentiation medium (Dulbecco’s modified Eagle medium [DMEM] supplemented with 1% fetal bovine serum, 1% horse serum, 0.5% chicken embryo extract, and 2% penicillin/streptomycin), 2) CM (high-glucose DMEM supplemented with dexamethasone, sodium pyruvate, ascorbate-2-phosphate, proline, insulin-transferrin-selenium+ Premix, L-glutamine, and 1% penicillin/streptomycin) (Cambrex Bioscience), or 3) CM (see description above) supplemented with TGF-β1 (10 ng/ml). After 4 days in culture, cells were fixed with cold acetone for 20 minutes, blocked for 60 minutes with 10% sheep serum containing 0.5% Triton X-100, and incubated with primary antibody (rabbit anti-rat collagen type II, 1:300; Chemicon) in blocking solution for 3 hours at room temperature (RT) in a humid chamber. The secondary antibody was sheep anti-rabbit Cy3 conjugated (1:300; Sigma Chemical), which was applied for 1 hour at RT. Nuclei were revealed with DAPI. Four chambers of the culture slide were used for each experimental group, and a Nikon E-800 microscope (Nikon Instruments) was used with
Northern Eclipse software (Empix Imaging) to calculate the number of collagen type II-positive colonies in 5 fields per chamber.
2.3.3. Cell pellet culture and staining for glycosaminoglycans
Cell pellets were made with MDSC and MDSC-B4 as described previously [126]. The cells first were trypsinized and counted. Next, 2×105 cells in 0.5 ml of CM were centrifuged at 500 g in 15 ml polypropylene coni-cal tubes. The pellets were incubated at 37 °C in the presence of 5% CO2
using chondrogenic medium with or without TGF-β1 (10 ng/ml). The medium was changed every 2 to 3 days. Pellets were harvested after 14 days, fixed overnight in 10% buffered formalin, dehydrated, and embedded in paraffin. After sectioning, the 5 µm-thick pellet sections were deparaffinized, placed in 3% acetic acid for 3 minutes, and transferred into Alcian blue solution for 30 minutes. The slides were rinsed with running tap water for 1 minute and counterstained with nuclear fast red.
2.4. Evaluation of osteogenic and chondrogenic
differentiation of MDSCs in vivo
2.4.1. Intramuscular transplantation of MDSCs
Two weeks after transduction with retrovirus encoding BMP4, CLBMP2/4, the transduced cells were trypsinized, centrifuged, and washed with Hanks’ balanced salt solution (HBSS), and 3×105 cells were injected percutaneously into the gastrocnemius muscle of SCID and immunocom-petent C57BL/6J mice (Jackson Laboratory) using a 30-gauge needle on a gas-tight syringe. Non-transduced cells were injected into the contra-lateral limb as a control. Fourteen days after transplantation, animals were sacrificed by cervical dislocation. The hind limbs were analyzed grossly and radiographically. The gastrocnemius muscles were isolated and flash frozen in 2-methylbutane buffered in phosphate buffered saline (PBS) pre-cooled in liquid nitrogen. The frozen samples were cut into 5 to 10 micron sections using a cryostat (Micron, HM 505 E, Fisher Scientific) and stored at –80 °C.
2.4.2. Evaluation of β-galactosidase and osteocalcin expression
The muscle sections were first fixed with cold acetone (100%), followed by a 1-hour blocking step with horse serum (10%) and incubated with a biotinylated anti-β-galactosidase antibody (1:100 in PBS; Sigma) followed by streptavidin conjugated to fluorescein (1:1000 in PBS, Sigma), to stain the β-galactosidase-expressing nuclei in green fluorescence. Then they were treated with a goat anti-mouse osteocalcin (1:50 in PBS, Biomedical Technologies). The muscle sections were subsequently incubated with a Cy3 conjugated anti-goat antibody (1:100 in PBS, Sigma) to label the osteocalcin-expressing cells in red fluorescence. The colocalization of the β-galactosidase and osteocalcin-expressing cells (yellow fluorescence) was visualized by fluorescence microscopy using an E-800 Nikon microscope.
2.4.3. Immune response to allogeneic cell implantation
Additional muscle sections were stained for CD4- and CD8-activated lymphocytes surface markers using a protocol previously described [127]. Briefly, muscle sections were fixed with cols acetone for 10 min and the non-specific binding site was blocked with a 3% mixture of goat and horse serum for 10 min followed by rinsing with PBS. The primary antibodies used were a rat monoclonal antibody against CD4 and CD8 (both from Pharmingen) for 2 h at room temperature. Subsequently, the endogenous peroxidase activity was blocked with 1% hydrogen peroxide for 20 min. Following several rinses in PBS, the sections were incubated with a biotinylated goat anti-rat antibody (1:500, Dako) for 1 h. This was followed by several rinses in PBS and incubation with Vectastain Elite ABC (5 ml PBS + two drop A and B, Vector) for 30 min. The peroxidase activity was revealed using 3’, 3’-diaminobenzidine (1 mg/ml; Sigma) and 0.03% hydro-gen peroxide. Muscle sections were then mounted with Gel Mount (Biomeda) and visualized by light microscopy.
2.4.4. Alcian blue staining for cartilage formation
Cartilage formation was determined by alcian blue staining using the following technique. Muscle sections were fixed with 4% formaldehyde and rinsed in PBS. The sections were subsequently incubated in the Alcian blue solution (1% Alcian blue solution; Alcian blue 8GX, 5 g and acetic acid 3%, pH 2.5) for 30 minutes, washed in distilled water, counterstained with eosin, and analyzed by light microscopy.
2.4.5. Skull defect model
Three 8-week-old male normal mice (Jackson Laboratories) were used in both the control and experimental groups. The mice were anesthetized with ketamine/xylazine mixture via intraperitoneal injection and placed prone on the operating table. Using a number 15 blade, the scalp was dissected to the skull, and the periosteum was stripped. A 5 mm full-thickness circular skull defect was created using a 1.5 mm dental burr, with minimal penetration of the dura. A 6 mm diameter collagen sponge matrix (Gelfoam, Pharmacia & Upjohn) seeded with 5.0×105 MDSCs, either with or without retrovirus-BMP4 transduction, was placed to fill the skull defect. The scalp was closed using a 4-0 nylon suture, and the animals were allowed food and activity. After 3 weeks, the mice were sacrificed, and harvested skull specimens were analyzed both grossly and microscopically.
2.4.6. Von-Kossa staining
For von Kossa staining, slides were fixed in 4% formaldehyde, and then soaked in 0.1 M AgNO3 solution for 15 minutes in the dark. After exposure
to light for at least 15 minutes, the slides were washed with PBS and stained with hematoxylin and eosin for histological evaluation.
2.4.7. Repair of osteochondral defects
Thirty-six 12-week-old athymic rats were used for the cartilage repair study. The animals were anesthetized via exposure to 3% isoflurane and O2
gas (1.5 liter/minute) delivered through an inhalation mask. A medial parapatellar skin incision was made, and the knee joint was exposed via lateral dislocation of the patella. A 1.5 mm biopsy punch (Robbins Instru-ments) was used to create a full-thickness articular cartilage defect (1.5×1.5×2.0 mm) in the trochlear groove of each femur. Cells were mixed with fibrin glue (Tisseel VH; Baxter Healthcare) before transplantation. The animals were divided into 3 treatment groups. The defects in the rats in group 1 were treated with 5.0×105 MDSC embedded in fibrin glue, defects in the rats in group 2 were treated with 5.0×105 MDSC-B4 embedded in fibrin glue, and defects in rats in group 3 (controls) were treated with acellular fibrin glue. After surgery, animals were allowed to move freely within their cages. Rats were sacrificed at 4, 8, 12, and 24 weeks after surgery.
2.4.8. Macroscopic evaluation and detection of LacZ transgene expression
At each time point, 6 knees from each treatment group were examined for gross appearance of the defects, smoothness and integration of the repaired tissue with the surrounding normal cartilage. The repaired tissues from 2 samples were sharply excised, flash frozen in liquid nitrogen, and cryostat-sectioned. The sections were stained with β-galactosidase (β-gal) and analyzed to determine the average percentage of LacZ-positive cells.
2.4.9. Evaluation of β-galactosidase and type II collagen expression
Sections obtained from the defect areas at the 4-week time point were stained with β-gal to assess LacZ expression and with Alcian blue to assess proteoglycan expression. Also, immunohistochemical staining was perfor-med to evaluate the differentiation of transplanted cells into chondrocytes by colocalizing β-gal and type II collagen. Staining for type II collagen was performed via the same protocol described for the in vitro monolayer culture above. The dilution of the primary antibody (rabbit anti-rat collagen type II) and secondary antibody (sheep anti-rabbit Cy3 conjugated) was 1:100. To stain for β-gal, tissue sections were blocked with 2% horse serum for 1 hour and incubated overnight at RT in a humid chamber with the primary antibody (monoclonal mouse anti-β-gal, 1:200; Sigma) in 1% horse serum. The sections were then incubated with streptavidin 488 for 1 hour (1:1000; Sigma) and mounted on slides.
2.4.10. Histological evaluation and grading of cartilage repair
After macroscopic examination, 4 distal femora per group per time point were dissected and fixed with 10% buffered formalin for 48–72 hours. They then were decalcified with decalcifying solution (Decalcifier II; Surgipath) for 24 hours and embedded in paraffin. Sagittal sections (5 µm thick) were obtained from the center of each defect and were stained with Safranin O-fast green. The histologic grading scale described by O’Driscoll et al. [128] was used to evaluate the quality of the repaired tissue. The total score on the grading scale ranges from 0 points (no cartilage repair) to 24 points (normal cartilage). Histological grading is based on the predominant nature of the repair tissue, matrix staining, regularity of the surface, structural integrity, thickness of the repair, apposition between the repaired cartilage and surrounding normal cartilage, freedom from degenerative signs in repair tissue, and freedom from degenerative changes of the surrounding normal cartilage.
2.5. Evaluation of osteogenic capacity of BMP4- and
VEGF-expressing MDSCs in vivo
2.5.1. Ectopic bone formation
Eight C57BL/6J mice (male, 12 weeks old) were randomly divided into two groups. Under general anesthesia, an 8-mm incision was made over the lateral aspect of each femur, a 7 mm Gelfoam disk (Pharmacia & Upjohn) impregnated with 6×105 of transduced cells was implanted into an intramuscular pocket created by blunt dissection, and the wound was closed with suture. Left hind limbs received BMP4-expressing MDSCs, while right hind limbs received mixture of BMP4- and VEGF-expressing cells at the ratio of 5:1 (BMP4-/VEGF-expressing cells). Ectopic bone formation was monitored radiographically and histologically at 7, 10, and 14 days post-implantation. The relative bone area and bone density detected by radiography were quantified with Northern Eclipse Version 6.0 software (Empix Imaging, Inc., North Tonarwanda, NY). Statistical differences between groups were analyzed using a Student’s t test with Microsoft Excel (Microsoft, Seattle, WA).
2.5.2. Skull defect healing
Twelve C57BL/6J mice (male, 12 weeks old) were randomly divided into three groups. Under general anesthesia, a 6 mm diameter defect was created in the parietal bone without breaching the dura and a 7-mm Gelfoam disk impregnated with BMP4-expressing cells (6×105), both BMP4-expressing and VEGF-BMP4-expressingcells (5×105 : 1×105), or Lacz-expressing and VEGF-expressing cells (5×105 : 1×105) was implanted into the defect.. Bone healing was monitored radiographically and histologically at designa-ted time points after surgery. Quantitative analyses of bone area and bone density as well as histological evaluation of cartilage formation and mineralization were performed as described above.
2.5.3. Immunostaining for vascular structures
Cryosections of bone samples were stained with a rat anti-mouse CD31 monoclonal antibody (clone MEC13.3, PharMingen) diluted 1:100 with reagents in the Vectastain Elite ABC kit (Vector Laboratories) following the protocol recommended by the manufacturer. The sections were counter-stained with hematoxylin and eosin. The relative capillary density was
estimated by counting the capillary number and measuring the length of the capillaries in the defined area after image digitization.
2.5.4. Apoptosis assay
Bone samples were fixed with 4% paraformaldehyde and the apoptotic cells were detected using the In Situ Cell Death Detection, POD kit (Roche Molecular Biochemicals) according to the instructions provided by the manufacturer. More than 500 cells in each quadruplicated sample were counted to determine the number of apoptotic cells in each group. The statistical differences were analyzed using a two-tailed Student’s t test.
2.6. Evaluation of chondrogenic capacity of BMP4- and
VEGF-expressing MDSCs
2.6.1. Pellet culture for in vitro chondrogenesis
Pellet culture was done as described previously [126]. Cell pellets were made with the following: 1) 1.4×105 of nontransduced MDSCs and 1.4×105 of BMP4-expressing MDSCs (BMP4 group); 2) 1.4×105 VEGF-expressing MDSCs and 1.4×105 BMP4-expressing MDSCs (BMP4+VEGF group); 3) 1.4×105 sFlt1-expressing MDSCs and 1.4×105 BMP4-expressing MDSCs (BMP4+sFlt1 group); 4) 2.8×105 primary chondrocytes derived from the mouse knee (Chond group); and three more groups (5, 6, and 7) were crea-ted using 2.8×105 nontransduced MDSCs. Pellets from groups 1–4 were cultured in 0.5 ml of chondrogenic medium (CM) that contained DMEM supplemented with 1% penicillin/streptomycin, 10-7M dexamethasone, 50 μg/ml ascorbate-2-phosphate, 40 μg/ml proline, 100 μg/ml pyruvate, and 1% ITS+Premix (Becton Dickinson) with 10 ng/ml of transforming growth factor β3 (TGFβ3; R&D Systems). Pellets in Group 5 (made with nontransduced cells) were cultured in chondrogenic medium without the TGFβ3 supplement (C group). Group 6 pellets were fed with chondrogenic medium without TGFβ3 but with 50 ng/ml BMP4 added (C+B4 group). Finally, group 7 pellets were fed with chondrogenic medium, supplemented with 10 ng/ml of TGFβ3 (C+T group). All pellets were incubated at 37 °C in 5% CO2, and the medium was changed every 2 to 3 days. Pellets were
2.6.2. Alcian blue staining
Pellets were fixed overnight in 10% neutral buffered formalin, dehydra-ted, embedded in paraffin, and sectioned in 5 µm-thick slices. Pellet sections were deparaffinized, placed in 3% acetic acid for 3 minutes, and transferred into Alcian blue solution for 30 minutes. The slides were then rinsed with running tap water for 10 minutes and counterstained with nuclear fast red.
2.6.3. RT-PCR analysis
mRNA was isolated using the RNeasy Plus Kit (Qiagen), according to the manufacturer’s instructions. After RNA extraction, quantitative real-time PCR (qPCR) analysis was carried out as described previously [129-130]. Gene expression levels were calculated based on the ∆CT method
(separate tubes). All target genes were normalized to the reference housekeeping gene, 18S. 18S primers and probes were designed by and purchased from Applied Biosystems. Primers and probes were designed for type II collagen, sox9, and type X collagen according to GenBank sequence. All target gene primers and probes were purchased from Integrated DNA Technologies Inc. Each experimental value is reported as the mean ± SEM results of triplicate treatments. For quantitative PCR assays, the coefficient of variation (CV) was calculated from three assay replicates. For all treatment groups and target genes analyzed, the CV did not exceed 3%. One-way ANOVA followed by Tukey-Kramer’s post hoc test using Stat View software was performed to determine significance among treatment groups. P values less than 0.05 were regarded as statistically significant.
2.6.4. Repair of osteochondral defects
Twenty-eight, 10-week-old nude rats (NIH-Whn NIHRNU-M, Taconic) were used in this study. The animals were anesthetized via exposure to 3% isofluorane and O2 gas (1.5 liter/minute) delivered through an inhalation
mask. The knee joint was exposed by medial parapatellar approach, and the trochlear groove was exposed by lateral dislocation of the patella. A 1.8 mm outer diameter trephine drill was used to create an osteochondral defect (1.8×2.0 mm) in the trochlear groove of each femur. Mouse skeletal muscle-derived stem cells were mixed with fibrin glue (Tisseel VH; Baxter Healthcare) before transplantation. The animals were divided into 7 treatment groups, and the defects in each group were treated as follows: 1) Group 1 (no-cell control) rats were treated with a(no-cellular fibrin glue; 2) Group 2 (MDSC group) rats were treated with 500,000 MDSCs embedded in fibrin glue;
3) Group 3 (VEGF group) rats were treated with 250,000 MDSC-VEGF cells plus 250,000 MDSCs embedded in fibrin glue; 4) Group 4 (sFlt1 group) rats were treated with 250,000 MDSC-sFlt1 cells plus 250,000 MDSCs embedded in fibrin glue; 5) Group 5 (B4 group) rats were treated with 250,000 MDSC-B4 cells plus 250,000 MDSCs embedded in fibrin glue; 6) Group 6 (B4+VEGF group) rats were treated with 250,000 MDSC-B4 cells plus 250,000 MDSC-VEGF cells embedded in fibrin glue; and, 7) Group 7 (B4+sFlt1 group) rats were treated with 250,000 MDSC-B4 cells plus 250,000 MDSC-sFlt1 cells embedded in fibrin glue. Four defects (two rats) were made for each group (n=4). The rats were allowed to move freely within their cages after surgery. Rats were euthanized 8 and 16 weeks after surgery. Groups including those receiving VEGF treatment were killed 8 weeks after surgery because the deterioration of the knee joint impaired the animal’s ability to move freely.
2.6.5. Histological evaluation and grading of cartilage repair
After macroscopic examination, 4 distal femora per group per time point were dissected and fixed with 10% neutral buffered formalin for 48 hours, followed by decalcification with 10% EDTA for 2 weeks and paraffin embedding. Sagittal sections, 5 µm in thickness, were obtained from the center of each defect and stained with Safranin O-fast green. The histolo-gical grading scale described by Sellers et al. [105] was used to evaluate the quality of the repaired tissue. The total score on the grading scale ranges from 0 points (normal cartilage) to 31 points (no repair). The modified scale allows evaluating all relevant aspects of repair of a full-thickness defect of articular cartilage. The categories included are: 1) filling of the defect rela-tive to surface of normal adjacent cartilage; 2) integration of repair tissue with surrounding articular cartilage; 3) matrix staining with Safranin O-fast green; 4) cellular morphology; 5) architecture within entire defect; 6) archi-tecture of surface; 7) percentage of new subchondral bone; and 8) formation of tidemark. All data were expressed as the mean ± SD. Differences of each category and total score were analyzed by Kruskal-Wallis and Mann-Whitney U tests using SPSS software (SPSS v12.0.1). P values less than 0.05 were regarded as significant.
3. RESULTS
3.1. Characterization of muscle-derived cell populations
3.1.1. Marker analysis of the pp6 cells
We isolated population of slowly adhering cells (pp6) from mdx mice and analyzed these cells for the expression of various markers using immuno-histochemistry, RT-PCR, and flow cytometry (Table 3.1.1.1).
Table 3.1.1.1. Immunohistochemical, RT-PCR, and flow cytometry analysis
of pp6 cells
Biochemical markers Immunohistochemistry RT-PCR Flow cytometry
Desmin + nd nd CD34 + + nd Bcl-2 + nd nd Flk-1 + nd nd Sca-1 + nd + M-cadherin –/+ nd nd Myogenin –/+ + nd C-met nd + nd MNF nd + nd MyoD +/– + nd C-kit nd nd – CD45 nd nd –
+, >95%; –, <2%; +/–, 40–80% of cells expressed the antigen; –/+, 5–30% of cells expressed the antigen; nd, not determined.
As shown in Table 3.1.1.1, pp6 cells expressed myogenic markers, inclu-ding desmin+, MyoD+/–, and Myogenin–/+. These cells were also c-met and MNF positive (RT-PCR), two genes which are expressed at an early stage of myogenesis [131]. The pp6 showed a lower percentage of cells expressing m-cadherin (–/+), a satellite cell-specific marker [132], but a higher percentage of cells expressing Bcl-2, a marker limited to cells in the early stages of myogenesis [133], and CD34, a marker identified in human hematopoietic progenitor cells as well as stromal cell precursors in bone marrow [134-137]. The pp6 cells were also highly positive for the
expression of Flk-1, a mouse homologue of human KDR gene which was recently identified as a marker of hematopoietic cells with stem cell-like characteristics [138]. Similarly, the pp6 cells were also found positive for Sca-1, a marker present in sub-populations of both skeletal muscle and hematopoietic cells with stem cell-like characteristics[25]. Finally, the pp6 cells were also found CD45 and c-kit negative.
3.1.2. Marker analysis of the mc13 cell clone
The biochemical markers expressed by mc13 cells were analyzed using RT-PCR, immunohistochemistry and flow cytometry. As summarized in Table 3.1.2.1, mc13 cells were positive for the expression of desmin, c-met, MNF, myogenin (+/–) and MyoD (RT-PCR). These results suggest that this clonal population contained cells at different stages of differentiation. The mc13 cells were positive for m-cadherin (+/–) and Bcl-2 (+/–) but negative for CD34 expression. Similar to that observed with the pp6 cells, the mc13 cells were highly positive for the expression of Flk-1 and Sca-1, but were negative for CD45 and c-kit (Table 3.1.2.1).
Table 3.1.2.1. Immunohistochemical, RT-PCR, and flow cytometry analysis
of mc13 cells
Biochemical markers Immunohistochemistry RT-PCR Flow cytometry
Desmin + nd nd CD34 – – nd Bcl-2 +/– nd nd Flk-1 + nd nd Sca-1 + nd + M-cadherin +/– nd nd Myogenin +/– + nd C-met nd + nd MNF nd + nd MyoD nd + nd C-kit nd nd – CD45 nd nd –
+, >95%; –, <2%; +/–, 40–80% of cells expressed the antigen; –/+, 5-30% of cells expressed the antigen; nd, not determined.
