PMCCPMCCPMCC

Search tips
Search criteria 

Advanced

 
Logo of nihpaAbout Author manuscriptsSubmit a manuscriptNIH Public Access; Author Manuscript; Accepted for publication in peer reviewed journal;
 
J Cell Mol Med. Author manuscript; available in PMC Nov 1, 2012.
Published in final edited form as:
PMCID: PMC3131486
NIHMSID: NIHMS256129
Differentiation and regeneration potential of mesenchymal progenitor cells derived from traumatized muscle tissue
WesleyM. Jackson,1,2 Thomas Lozito,1,3 Farida Djouad,1 Nastaran Z. Kuhn,1 Leon J. Nesti,1,2,4* and Rocky S. Tuan1,3*
1 Cartilage Biology and Orthopaedic Branch, National Institute of Arthritis, Musculoskeletal and Skin Disease, National Institutes of Health, Department of Health and Human Services, Bethesda, Maryland, USA
2 Clinical and Experimental Orthopaedics Laboratory, National Institute of Arthritis, Musculoskeletal and Skin Disease, National Institutes of Health, Department of Health and Human Services, Bethesda, Maryland, USA
3 Center for Cellular and Molecular Engineering, Department of Orthopaedic Surgery, University of Pittsburgh School of Medicine, Pittsburgh, PA
4 Department of Orthopaedics and Rehabilitation, Walter Reed Army Medical Center, Washington, DC, USA
Prior to Publication, address correspondence to: Wesley M. Jackson, PhD, 834 Evelyn Ave, Albany, CA 94706, Phone: (510) 813-8611, wjackson/at/gmail.com
Please use the following address in the manuscript for correspondence: Dr. Rocky Tuan, Department of Orthopaedic Surgery, Center for Cellular and Molecular Engineering, University of Pittsburgh, 450 Technology Drive, 2nd Floor, Room 206, Pittsburgh, PA 15219-3143, Phone: (412) 648-2222, Fax: (412) 624-5544, rst13/at/pitt.edu
*Dr. Nesti and Dr. Tuan were co-corresponding authors of this manuscript.
Mesenchymal stem cell (MSC) therapy is a promising approach to promote tissue regeneration by either differentiating the MSCs into the desired cell type or by using their trophic functions to promote endogenous tissue repair. These strategies of regenerative medicine are limited by the availability of MSCs at the point of clinical care. Our laboratory has recently identified multipotent mesenchymal progenitor cells (MPCs) in traumatically injured muscle tissue, and the objective of this study was to compare these cells to a typical population of bone marrow-derived MSCs. Our hypothesis was that the MPCs exhibit multilineage differentiation and expression of trophic properties that make functionally them equivalent to bone marrow-derived MSCs for tissue regeneration therapies. Quantitative evaluation of their proliferation, metabolic activity, expression of characteristic cell-surface markers and baseline gene expression profile demonstrate substantial similarity between the two cell types. The MPCs were capable of differentiation into osteoblasts, adipocytes and chondrocytes, but they appeared to demonstrate limited lineage commitment compared to the bone-marrow derived MSCs. The MPCs also exhibited trophic (i.e., immunoregulatory and pro-angiogenic) properties that were comparable to those of MSCs. These results suggest that the traumatized muscle-derived MPCs may not be a direct substitute for bone marrow-derived MSCs. However, because of their availability and abundance, particularly following orthopaedic injuries when traumatized muscle is available to harvest autologous cells, MPCs are a promising cell source for regenerative medicine therapies designed to take advantage of their trophic properties.
Keywords: Mesenchymal stem cells, progenitor cells, tissue engineering, regenerative medicine, immunoregulation, angiogenesis
Mesenchymal stem cells (MSCs) offer enormous promise in emerging strategies of cell-based therapeutics to promote the regeneration of damaged tissues and to modify the natural history of many diseases [14]. One attractive feature of MSCs is their ability to differentiate into multiple cell types, which makes them useful for tissue regeneration strategies to replace damaged, impaired or dead cells and tissues, either by cell replacement therapy [56] or in combination with a scaffold for tissue engineering [7]. MSCs also exert trophic functions in the environment of damaged tissues that promote endogenous wound healing mechanisms [8], such as promoting angiogenesis [910], reducing fibrosis [11] and modulating inflammation [12]. In particular, the immunoregulatory properties of MSCs have made them a useful therapeutic for the treatment of graft versus host disease (GvHD) [13]. Although the pro-regeneration functions of MSCs make them applicable for a variety of other clinical applications, there are significant challenges that limit their widespread use, including optimization of methods to harvest the cells, expand them ex vivo, and in the case of allogeneic MSCs, to transport and store the cells prior to implantation [14]. Many of these limitations could be overcome by using autogenic MSCs that can be harvested and used as a therapeutic agent at the point of clinical care [15].
Our laboratory has recently identified a novel population of cells in traumatically injured muscle tissue that resemble MSCs. Although multipotent stem cells have previously been isolated from untraumatized muscle using immuno-selective techniques [16], the plastic-adherent cells isolated from traumatized muscle can be harvested in substantially higher numbers [17]. We refer to this population as mesenchymal progenitor cells (MPCs) to indicate that these cells are likely be the descendants of stem cells (i.e., pericytes [18]) but are proliferating in the tissue at the time of harvest. The morphology and cell surface epitope profiles of MPCs are similar to those of bone marrow-derived MSCs, and they give rise to colony-forming-unit-fibroblasts (CFU-Fs), an indicator of a clonogenic, multipotent cell population [19]. Although the MPCs are capable of differentiation into osteoblasts, adipocytes and chondrocytes, they appeared to have a distinct osteogenic gene expression profile, which likely reflects their different tissue of origin and in vivo function [20]. It has been suggested that non-muscle progenitors participate in the process of muscle regeneration by facilitating the reparative function of myoblasts and myofibroblasts [21] and that the migration of MPCs into the traumatized muscle may be a part of the normal wound healing response [22].
The traumatized muscle-derived MPCs could be a useful cell source for cell-based therapies that follow musculoskeletal injury, particularly those related to orthopaedic reconstruction [23]. MPCs have been harvested from surgically debrided tissues from the wound margins of extremity injuries [19], which would provide an autogenic cell source that does not require a separate procedure for cell collection. Since the MPCs are harvested at relatively high concentrations, they may not require ex vivo expansion and could be ready for use almost immediately. However, there have been no quantitative comparisons of the MPCs to bone marrow-derived MSCs to evaluate whether the MPCs are a suitable substitute cell type. Furthermore, the ability of MPCs to exhibit trophic properties has not yet been investigated. The overall hypothesis of this study was that traumatized muscle-derived MPCs exhibit multilineage differentiation and expression of trophic properties that make them equivalent to bone marrow-derived MSCs to enhance tissue regeneration. Our specific aims were: (1) to compare the phenotype characteristics of MPCs to bone marrow derived MSCs, (2) to quantify the ability of the MPCs to differentiate into osteoblasts, adipocytes and osteoblasts, and (3) to determine whether MPCs exhibit trophic properties that are characteristic of MSCs.
MPC Harvest
Traumatically injured muscle was collected with informed consent and Institutional Review Board (IRB) approval from the Walter Reed Army Medical Center (WRAMC) using a previously described method [17,19]. Patients were considered for inclusion in this study if they had sustained traumatic injury with extensive soft tissue extremity wounds (n=24; age: 24.4 ± 4.7; sex: 100% male). Traumatized muscle was obtained from the healthy wound margin during serial debridement following orthopaedic trauma. Serial sections of the traumatized muscle were stained with hematoxylin and eosin to evaluate the tissue morphology and compared to control muscle sections (Biogenex, San Ramon, CA). Approximately 200 mg of the tissue was minced, incubated at 37°C for 2 hours with gentle agitation in digestion medium (DMEM with 0.5 mg/mL collagenase 2, Worthington Biosciences, Lakewood, NJ) and passed through a 40 μm cell strainer. The cells were pelleted by centrifugation, resuspended and plated in a T150 tissue culture flask in DMEM supplemented with 10% fetal bovine serum (FBS, Invitrogen) and 5 units/mL of penicillin, streptomycin and fungizone (PSF, Invitrogen). After 2 hours, the non-adherent cells were removed by washing with phosphate-buffered saline (PBS, Invitrogen). The adherent cells were cultured in growth medium (DMEM supplemented with 10% FBS and 1 unit/mL of PSF) and washed daily with PBS until subculture when tightly packed colony forming units (CFUs) were observed. Subsequent subcultures were performed when the cells were approximately 85% confluent. Bone-marrow derived MSCs were harvested from femoral heads obtained from total hip arthroplasties using an established technique [24] and with the consent of the patients (n=14; age: 54.9 ± 8.7; sex: 42.8% male) following an IRB approved protocol at the University of Washington. All cells were cryopreserved at the end of the first passage, and multiple biological replicates of each experiment were performed by thawing the cells simultaneously. Unless otherwise noted, all experiments were performed with third or fourth passage cultures.
Proliferation and Metabolic Activity Assays
Cells were plated in 24-well plates at an initial density of 1,000 cells/cm2. Cell number at the prescribed time-points was determined on the basis of dsDNA concentration using the PicoGreen assay (Invitrogen), and the doubling time was estimated using an exponential fit. Cellular metabolism based on the output of mitochondrial reductase was estimated using 3-(4,5-dimethylthiazol-2-yl)-2,5-diphenyltetrazolium bromide (MTT, Sigma). At the prescribed time points, MTT was added to the medium in 4 replicate wells to a final concentration of 1 mg/mL, and the cells were incubated at 37°C for two hours. The medium was aspirated, and the incorporated MTT reduction product was eluted with DMSO and quantified spectrophotometrically (A550) using a microtiter plate reader (Synergy HT, Bio-Tek, Winooski, VT).
Immunophenotyping
Flow cytometry was performed on cells at the end of the second passage as previously described [17]. Briefly, 100,000 cells were incubated with approximately 0.4 μg of phycoerythrin (PE) conjugated antibodies in 1.0 mL of FACS buffer (0.1% bovine serum albumin and 0.01% sodium azide in HBSS). All antibodies were mouse IgG1, κ isotype and reactive against human antigens (isotype control: clone MOPC-2I; CD14: clone M5A2; CD19: Clone HIB19; CD34: clone 563; CD45: clone HI30; CD49d: clone 9F10; CD73: clone AD2; CD79a: clone HM47; CD90: clone 5E10; CD146: clone P1H12; HLA-DR: clone TU36 -BD Biosciences; CD105: clone 6N6 - Serotec, Oxford, UK). The cells were incubated in the dark at 4°C for 40 minutes, washed once and resuspended in 100 μL of FACS buffer, and the fluorescence intensity profiles of the cells were analyzed using a FACSCalibur flow cytometer (BD Biosciences).
Differentiation Assays
The MPCs and MSCs were plated onto tissue culture plastic and cultured for up to 28 days in either (1) osteogenic induction medium (OM) containing GM supplemented with 10 mM β-glycerophosphate (Sigma, Saint Louis, MO), 50 μg/mL ascorbic acid (Sigma), 10 nM 1,25-dihydroxyvitamin D3 (Biomol, International, Plymouth Meeting, PA) and 0.01 μM dexamethasone (dex, Sigma) or (2) adipogenic induction medium (AM) containing GM supplemented with 0.5 mM IBMX (Arcos Organics, Geel, Belguim), 1 μM dex and 1 μg/ml insulin (Sigma). Chondrogenic differentiation was performed in pellet cultures containing 2.5×105 cells per pellet and by culturing the cells in chondrogenic medium (CM) containing DMEM supplemented with 1% ITS (BD Biosciences, San Jose, CA), 40 μg/mL L-proline, 10 ng/ml transforming growth factor-β3 (TGF-β3, R&D Systems, Minneapolis, MN), and 0.1 μM dex for 21 days. Osteogenesis was assessed using a bb Fast Blue alkaline phosphatase (ALP) kit (Sigma) and by staining mineralized matrix with alizarin red. Alkaline phosphatase activity was quantified with an ELISA-based assay [20]. Adipogenesis was assessed using oil red O to stain for intracellular lipid accumulation, which was eluted with isopropanol for spectrophotometric quantification (A510). Chondrogenesis was assayed histochemically by staining for sulfated glycosaminoglycans (GAGs) with alcian blue, and immunohistochemically by staining for collagen type II and aggrecan. GAG production was quantified using the Blyscan assay kit (Biocolor, Carrickfergus, UK).
qRT-PCR Array Analysis
Cells were lysed in TRIzol (Invitrogen, Carlsbad, CA), homogenized using QiaShredder columns (Qiagen, Hilden, Germany) and total RNA was extracted according to the manufacturer’s protocol, purified using RNeasy Mini columns (Qiagen), and RNA concentrations were estimated on the basis of A260. Real-time RT-PCR analysis was performed using SYBR Green (Bio-Rad) and a BioRad iCycler iQ real-time PCR detection system and gene expression was normalized to GAPDH using RT2 PCR arrays for MSC biology (SABiosciences, Frederick, MD).
Mixed Lymphocyte Reaction
Peripheral blood from healthy human donors was collected into heparinized containers (BD Biosciences), and mononuclear cells (PBMC) were isolated by Ficoll-Hypaque density-gradient centrifugation as described previously [25]. Responder human PBMC were resuspended in RPMI 1640 medium containing 10% FBS, 2 mM glutamine, 100 U/ml penicillin and 100 μg/ml streptomycin, 0.1 mM non-essential amino acids, 1 mM sodium pyruvate, 20 mM HEPES and 50 μM 2-mercaptoethanol (Invitrogen). PBMCs were seeded in triplicates at 1 × 105 cells/100 μl/well in 96-well round bottom plates (BD Biosciences). Phytohemagglutinin (PHA) was used at 5 μg/ml as a positive control mitogen to induce T-cell proliferation. MPCs were added at prescribed concentrations to obtain a 300 μl final volume. After 3 days of incubation, 1 μCi/well [3H]-thymidine (Amersham) was added overnight and radioactivity incorporation was determined by liquid scintillation counting.
Vascular Stability Assays
MPCs were cultured for 3 days in expansion medium and then with serum-free medium. After 1, 2, or 4 days, the media were then collected and concentrated by centrifugation in spin columns (Amicon Ultra-15 3 kDa NMWL; typical concentration yields: 20–40× by volume) to generate conditioned media (CM). Aliquots of the CM were analyzed by SDS-PAGE and western blotting for Vascular Endothelial Growth Factor (VEGF). HMEC-1 microvascular endothelial cells (ECs; Center for Disease Control [2627]) were cultured in CM that had been reconstituted to 1X in serum free medium supplemented with EGM-2-MV bullet kits (Cambrex, East Rutherford, NJ) [28]. After 48 or 72 hours, cell proliferation was quantified using the MTS Cell Proliferation Assay (Promega, Madison, WI).
Statistical Methods
Statistical significance for all tests was assigned to p<0.05 except for RT-PCR array gene expression levels, for which statistical significance was assigned to p<0.018 to limit the false discovery rate.
The phenotype of traumatized muscle-derived MPCs was similar to other populations of MSCs
The traumatized muscle exhibited substantial histological evidence of tissue damage (Figure 1A). Within two hours of plating the cells harvested from this tissue, adherent MPCs with an MSC-like morphology were observed for all of the 26 traumatized muscle tissue samples included in this study (Figure 1B). The MPCs were present at substantially greater numbers in the traumatized muscle than MSCs are present in the bone marrow, and the traumatized muscle also yielded a greater number of MPCs that formed ALP positive colonies (Table 1). There was no significant difference in the proliferation rate of the MPCs compared to bone marrow-derived MSC (Figure 1C), although the metabolic activity of the MPCs was significantly higher (Figure 1D). The cell surface epitope profile of the MPCs met the minimum criteria established for MSC populations [29] (Figure 2A), and there was no significant difference in the fluorescence intensities of the selected positive markers. Although the intensity of CD49d staining was low on the MPCs, greater than 97% of the MPCs expressed this surface marker, which is not uniformly expressed on bone-marrow derived MSCs [30]. Fewer than 5% of cells from either population expressed the pericyte marker CD146.
Figure 1
Figure 1
Morphology, proliferation and metabolism of traumatized muscle-derived MPC. (A) Representative sections of the traumatized muscle demonstrating the extent of tissue damage compared to control muscle. Bar=200 μm; hematoxylin and eosin. (B) MPC (more ...)
Table 1
Table 1
Properties of traumatized muscle derived MPCs and bone marrow-derived MSCs
Figure 2
Figure 2
Cell surface epitope profile of traumatized muscle-derived MPCs. (A) MPCs surface marker expression (black dots) compared to isotype control (grey dots) were compared using flow cytometry. All antibodies were PE conjugated (red), and the percentage of (more ...)
Multilineage differentiation activity of MPCs was lower than bone marrow MSCs
Under appropriate induction conditions, both cell types exhibited evidence of osteogenic, adipogenic and chondrogenic differentiation (Figure 3A). The MPCs up-regulated their surface expression of ALP and began to generate a mineralized matrix under osteogenic conditions (visualized with alizarin red), accumulated intracellular lipid stores under adipogenic conditions (visualized with oil red O), and they began to express and produce collagen type II and sulfated GAGs (visualized with alcian blue), including aggrecan, under chondrogenic conditions.
Figure 3
Figure 3
Trilineage differentiation of traumatized muscle-derived MPCs. (A) Histological evidence of differentiation in MPCs compared to bone marrow-derived MSCs grown in growth medium (GM), osteogenic induction medium (OM), adipogenic induction medium (AM), and (more ...)
Upon quantification, the MPCs appeared to exhibit a lower level of differentiation than bone marrow-derived MSCs. Although, the MPCs accumulated more intracellular lipids than MSCs under adipogenic conditions (Figure 3C), the up-regulation of surface ALP in MPCs was significantly lower than MSCs under osteogenic conditions (Figure 3B). Chondrogenic differentiation by MPCs was particularly limited, as they produced significantly smaller pellets, less sGAG per cell, and less overall sGAG than the bone marrow-derived MSCs (Figure 4D-F).
Figure 4
Figure 4
qPCR array for mesenchymal stem cell genes. (A) Comparison of the gene expression profiles for traumatized muscle-derived MPCs and bone marrow-derived MSCs. Genes with 4-fold differential expression (indicated by dashed lines) are labeled and select genes (more ...)
MPCs exhibit regenerative properties that are characteristic of MSCs
The gene expression profile of traumatized muscle-derived MPCs was nearly identical to that of bone marrow-derived MSCs (Figure 4A). No difference was detected between the two cell types in gene-expression levels of the MSC markers CD44 and ENG (CD105; Figure 4C), or baseline expression of PPARG2, SOX9 and RUNX2, the master regulators of adipogenesis, chondrogenesis and osteogenesis, respectively(Figure 4D). Notable differences in the gene expression profiles include THY1 (CD90) and NES (nestin), which are expressed at significantly higher levels in MPCs, and VCAM1 (vascular cellular adhesion molecule-1), which is expressed at significantly higher levels in bone marrow-derived MSCs (Figure 4B).
The MPCs also expressed similar levels of genes associated with the regenerative functions of MSCs. There was no significant difference in the gene expression of the immuno-regulatory genes IL6, IL10, HGF (hepatocyte growth factor) or TGFB3 (transforming growth factor-beta3) between the MPCs and MSCs, while IFNG (interferon-gamma) was expressed at slightly but significantly higher level in MPCs (Figure 5A). The immuno-regulatory function of the MPCs was verified using a mixed lymphocyte reaction assay (Figure 5B). The MPCs attenuated T-cell proliferation in response to antigen stimulation in a dose dependent manner, although MSCs appeared to generate greater attenuation of the immune response. The MPCs and MSCs expressed nearly identical levels of FGF2 (fibroblast growth factor-2), EGF (epithelial growth factor) and VEGFA (vascular endothelial growth factor-A), genes associated with vascular maintenance (Figure 6A), and the production of VEGFA, a potent angiogenic factor, by MPCs was verified in western blots (Figure 6B). The pro-angiogenic potential of MPCs was assayed by measuring the effect of their secreted factors on the proliferation of microvascular endothelial cells (Figure 6C). Similar to bone-marrow derived MSCs, the MPCs secreted factors that significantly increased microvascular endothelial cell proliferation within 48 hour of initial seeding, and there were no significant differences between the effects of the MPC or MSC secreted factors.
Figure 5
Figure 5
Immunosuppression by traumatized muscle-derived MPCs. (A) Comparison of immunoregulatory gene expression between MPCs and bone marrow-derived MSCs. (B) T-cell proliferation in a mixed lymphocyte reaction plotted as a percentage of total proliferation (more ...)
Figure 6
Figure 6
Vascular maintenance associated with traumatized muscle-derived MPCs. (A) Comparison of angiogenic gene expression between MPCs and bone marrow-derived MSCs. (B) Western blots to verify VEGF protein expression in the cell supernatants at days 1, 2 and (more ...)
MSCs may be a useful cell type for a variety of applications in regenerative medicine, although there is currently no clear consensus on the best method of sourcing and isolating MSCs for clinical use. In this study, we have evaluated a recently identified population of traumatized muscle-derived MPCs by quantitatively comparing them to a typical population of bone-marrow-derived MSCs. The traumatized muscle tissue is an attractive source of autolgous cells with MSC-like properties, as it is readily obtained in a clinical setting following orthopaedic injury without requiring additional tissue harvesting procedures [31]. We have demonstrated that MPCs can be harvested at high concentrations from traumatized muscle, and the proliferation rate, cell surface epitope profile and gene expression profile of the MPCs is substantially similar to that of bone marrow-derived MSCs. The MPCs also appear to maintain a higher level of metabolic activity relative to the MSCs. While they differentiate easily into adipocytes, the MPCs exhibit more limited differentiation into osteoblasts and chondrocytes. Finally, the MPCs express factors that modulate inflammatory responses and promote angiogenesis at functional levels that are comparable to the MSCs. Taken together, these results suggest that while traumatized muscle-derived MPCs may not be a direct substitute for bone marrow-derived MSCs, they perform specific trophic functions that will make them useful in regenerative medicine applications.
Several aspects of this study support our conclusions. First, both histological and quantitative analyses of the differentiation assays were included in our investigation. Compared to our previous qRT-PCR study showing the up-regulation of lineage specific genes by MPCs under the appropriate conditions [17,19], the protein-level assays reported in this studyprovide a more precise measure of lineage adoption. Using these higher sensitivity assays, we were able to observe the limitations of the MPCs to differentiate into osteoblasts and chondrocytes relative to the MSCs. Second, in addition to performing the in vitro differentiation assays on this novel population of MSC-like cells, we also investigated the trophic function of these cells, which are assumed to constitute at least part of the therapeutic benefit provided by MSCs. In addition to looking at the expression of genes associated with these functions, we performed functional assays to verify the activity of these trophic mechanisms by the traumatized muscle-derived MPCs. Finally, all of the experiments in this study were performed using human cells from a clinically relevant source. As a result, our findings are directly applicable to the development of therapies that can take advantage of this cell type.
Despite these strengths, a few caveats should be noted. First, this study was designed to investigate the spectrum of regenerative functions that MPCs are capable of performing, but it did not generate any insight into the mechanisms by which the MPCs were able perform these functions. Many of these mechanisms have been identified in MSCs, and it is tempting to assume that they are conserved between the two cell types. We are currently performing more comprehensive studies that are will investigate the mechanism of these functions in greater detail using additional experimental outputs. However, the present study was useful in that it provides a broad and quantitative comparison between the MPCs and bone marrow-derived MSCs. Second, all of the experiments in this study were performed in vitro, and further investigation will be necessary to verify that the traumatized muscle-derived MPCs will exhibit the same regenerative functions in vivo. Finally, the MPCs and MSCs were not age and sex matched due to limitations imposed by the demographics of the patients providing our tissue samples, in view of previous reports on the effects of age and sex on stem and progenitor cell function, the differences between the MPCs and MSCs reported here should be evaluated with caution.
Variations in differentiation potential [32] and trophic function [3334] of MSCs have previously been correlated with the age and sex of the donors from which the cells were harvested. The precise mechanism leading to these changes in MSC function are not completely understood, although age related changes have been attributed to telomere shortening [35], and the sex dimorphisms appear to be generated by preconditioning of the MSCs by sex hormones prior to harvest [36]. In the context of these donor effects, the differences in traumatized muscle-derived MPCs and bone-marrow-derived MSCs can be examined in greater detail. The substantially higher yield of harvested MPCs relative to MSCs is not consistent with the changes in cellularity that might be attributable to donor age or sex [3738], suggesting that the difference in cellularity is dominated by the tissue of origin and the cellular responses to injury. However, despite the donor mismatch, no significant differences in the proliferation rate or expression of lineage specific genes were observed between the two cell types. These results suggest that the difference in mean age between the two donor groups had a small effect on these outputs relative to the overall biological variability. We did find evidence of a potential sexual dimorphism in that the MSCs, which contained cells from female donors, appeared to more effectively suppress T-cell proliferation. However, the difference in immunosuppression between the cell types was slight, and only significantly different at two cell concentrations. The angiogenic function of the MPCs and MSCs also appeared to be equivalent. Taken together, the effects of donor sex also appear to be small relative to the biological variability. Finally, the impaired osteogenic and chondrogenic differentiation of the MPCs relative to MSCs might be expected due to the presence of females in the MSC group, but this result is inconsistent with the expected age-related effects [39]. As a result, these differences in the differentiation potential might be dominated by the difference in cell type.
Other progenitor cells with properties similar to MSCs have been harvested from human skeletal muscle tissue. Using immunoselection techniques, a population of cells can be isolated from digested muscle tissue that can differentiate into osteoblasts, adipocytes and chondrocytes, as well as into myoblasts [40]. Termed myoendothelial cells due to the myogenic (CD56) and endothelial cell (CD34 and CD144) markers that distinguish this cell type, they also promote regeneration of skeletal [16] and cardiac [41] muscle tissue, in part by secreting pro-angiogenic and pro-survival factors that promote the endogenous repair mechanisms. These cells demonstrate many characteristics of the muscle-derived stem cells (MDSCs), which can be isolated from murine skeletal muscle [42], although a direct equivalent of MDCSs has not been identified in human muscle tissue. It is noteworthy that the MDSC cell type is isolated on the basis of its slow adherence to tissue culture plastic (TCP) during the harvesting procedure [43], in contrast to the traumatized-muscle-derived MPCs, which rapidly adhere to the TCP in less than 2 hours.
In humans, it is assumed that the myoendothelial cells are related to pericytes, as both cell types share the perivascular niche. The pericytes also closely resemble MSCs in vitro, and there has been recent, compelling evidence that pericytes can be harvested from various tissues throughout the body and induced to exhibit MSC characteristics [18]. The MPCs may be an activated descendent of the myoendothelial/pericyte cell types that have (1) been activated in response to traumatic injury, (2) down-regulated the expression of surface proteins required for their vascular niche [44], and (3) began to proliferate in the tissue [22]. There is also recent evidence indicating that a multipotential progenitor cell population may be derived from the vasculature via epithelial to mesenchymal transition in response to injury [45]. Taken together, these studies suggest that the MPCs may arise from the vasculature in large numbers following trauma, and they may participate in the wound healing process by secreting trophic factors to promote tissue regeneration by mechanisms similar to those used by the myoendothelial cells and pericytes [46].
The MPCs may also be descendants of bone marrow-derived MSCs that entered the traumatized tissue via the bloodstream while homing to the site of injury [47]. Some differences were detected between the MPCs and bone marrow-derived MSCs, and these differences may reflect the tissues from which they were harvested and the extracellular environment immediately prior to harvest. The MPCs exhibited higher metabolic activity than the MSCs, which may indicate that they undergo mitochondrial biogenesis in response to injury and to prepare them for their contribution to the wound healing response [48]. The differences in the baseline gene expression profile may also be justified given that MPCs were in the regenerating muscle tissue at the time of harvest, whereas the MSCs were in their bone marrow niche [20]. For example, the MSCs expressed higher levels of VCAM1, which is characteristic of genes associated with bone physiology and maintenance by the marrow stroma [49], whereas the MPCs expressed higher levels of genes that indicate neuromuscular differentiation: THY1 [50] and NES [51]. It was not possible to definitively trace the origin of the MPCs to the muscle tissue or the bone marrow without using an in vivo injury model. However, it is likely that there are multiple sources of stem and progenitor cells that converge into the MPC phenotype once they are exposed to the biochemical milieu of the traumatized muscle tissue, and this heterogeneity could account for differences between MPCs and the more homogeneous population of bone marrow-derived MSCs.
Regardless of their origin, it appears that the muscle-derived MPCs may be a clinically useful population of autologous cells for regenerative medicine, particularly in orthopaedic applications [52]. The typical standard of care for musculoskeletal injuries is to debride to the wound margin until definitive closure is possible [31]. The results of this study indicate that the viable portions of debrided tissue, typically discarded as surgical waste, might instead be harvested to obtain MPCs, which can be used augment the wound healing process. The MPCs exhibit the trophic functions that are characteristic of bone marrow-derived MSCs, and they can be harvested without the need for additional procedure, such as bone marrow aspiration, which may be painful and exposes patient to additional surgical risks. Although the immunoregulatory functions of MPCs may be less effective than MSCs on a per cell basis, the MPCs can be harvested from traumatized muscle tissue at concentrations that are orders of magnitude higher than MSCs from bone marrow (Table 1). Therefore, the overall immunoregulatory function of MPCs harvested per gram of muscle tissue should be at least as effective as the MSCs harvested per gram of bone marrow (Figure 5B). Furthermore, the MPCs may be harvested at sufficient numbers for immediate clinical use without the need for ex vivo expansion. We are currently evaluating the use of autologous MPCs to promote regeneration of muscle, as well as other tissues that are typically damaged as a result of orthopaedic trauma, such as bone, nerve and blood vessels [5354]. MPCs may also be useful to manage GvHD following transplant of orthopaedic tissues [55].
In this study, we have evaluated the traumatized muscle-derived MPCs by quantitatively comparing their differentiation potential and trophic properties to bone marrow-derived MSCs. The two cell types share many similarities, although the extent to which the MPC population is able to undergo osteogenic and chondrogenic differentiation appears to be limited. Both cell types appear to exhibit immunoregulatory and pro-angiogenic trophic functions, which are an important component of the regenerative benefit of MSCs. Therefore, the traumatized muscle-derived MPCs appear to be an alternative source of autologous cells that are capable of performing the trophic functions that enhance tissue regeneration. The MPCs may have an advantage over bone marrow-derived cells in cellular therapy applications that follow orthopaedic trauma since they may be harvested without performing an additional surgical procedure, they do not require ex vivo expansion, and they appear activated by the trauma to participate in the wound healing response. We are continuing to investigate the mechanisms that mediate these trophic abilities in the MPCs and their function during wound healing. Simultaneously, we are developing regenerative medicine and tissue engineering strategies that take advantage of these functions to promote tissue regeneration.
Acknowledgments
Support was provided by the Military Amputee Research Program at WRAMC (PO5-A011), the Comprehensive Neurosciences Program (CNP-2008-CR01), the NIH Intramural Research Program (Z01 AR41131) and the University of Pennsylvania, Department of Health. Immunophenotyping was performed in the NIAMS Flow Cytometry Group with the assistance of James Simone. Technical assistance was provided by Jun Onodera, Yingjie Song and Ibardo Zambrano. We thank Dr. Paul Manner, Department of Orthopaedic Surgery at the University of Washington, for providing human skeletal tissues.
Footnotes
The authors confirm that there are no conflicts of interest.
1. Chen FH, Tuan RS. Adult stem cells for cartilage tissue engineering and regeneration. Curr Rhematol Rev. 2008;4:149–54.
2. Patterson TE, Kumagai K, Griffith L, et al. Cellular strategies for enhancement of fracture repair. J Bone Joint Surg Am. 2008;90 (Suppl 1):111–9. [PubMed]
3. Abdallah BM, Kassem M. Human mesenchymal stem cells: from basic biology to clinical applications. Gene Ther. 2007;15:109–16. [PubMed]
4. Nesselmann C, Ma N, Bieback K, et al. Mesenchymal stem cells and cardiac repair. J Cell Mol Med. 2008;12:1795–810. [PubMed]
5. Garcia-Castro J, Trigueros C, Madrenas J, et al. Mesenchymal stem cells and their use as cell replacement therapy and disease modelling tool. J Cell Mol Med. 2008;12:2552–65. [PubMed]
6. Newman RE, Yoo D, LeRoux MA, et al. Treatment of inflammatory diseases with mesenchymal stem cells. Inflamm Allergy Drug Targets. 2009;8:110–23. [PubMed]
7. Kuo CK, Tuan RS. Tissue engineering with mesenchymal stem cells. IEEE Eng Med Biol Mag. 2003;22:51–6. [PubMed]
8. Caplan AI. Why are MSCs therapeutic? New data: new insight. J Pathol. 2009;217:318–24. [PubMed]
9. Tang YL, Zhao Q, Zhang YC, et al. Autologous mesenchymal stem cell transplantation induce VEGF and neovascularization in ischemic myocardium. Regul Pept. 2004;117:3–10. [PubMed]
10. Martens TP, See F, Schuster MD, et al. Mesenchymal lineage precursor cells induce vascular network formation in ischemic myocardium. Nat Clin Pract Cardiovasc Med. 2006;3 (Suppl 1):S18–22. [PubMed]
11. Li L, Zhang Y, Li Y, et al. Mesenchymal stem cell transplantation attenuates cardiac fibrosis associated with isoproterenol-induced global heart failure. Transpl Int. 2008;21:1181–9. [PubMed]
12. Kode JA, Mukherjee S, Joglekar MV, et al. Mesenchymal stem cells: immunobiology and role in immunomodulation and tissue regeneration. Cytotherapy. 2009;11:377–91. [PubMed]
13. Le Blanc K, Rasmusson I, Sundberg B, et al. Treatment of severe acute graft-versus-host disease with third party haploidentical mesenchymal stem cells. Lancet. 2004;363:1439–41. [PubMed]
14. Habib NA, Gordon MY. Clinical applications of stem cell therapy--the pros and cons of stem cell sources. Regen Med. 2006;1:301–2. [PubMed]
15. Bongso A, Fong CY, Gauthaman K. Taking stem cells to the clinic: Major challenges. J Cell Biochem. 2008;105:1352–60. [PubMed]
16. Zheng B, Cao B, Crisan M, et al. Prospective identification of myogenic endothelial cells in human skeletal muscle. Nat Biotechnol. 2007;25:1025–34. [PubMed]
17. Jackson WM, Aragon AB, Djouad F, et al. Mesenchymal progenitor cells derived from traumatized human muscle. J Tissue Eng Regen Med. 2009;3:129–38. [PMC free article] [PubMed]
18. Crisan M, Yap S, Casteilla L, et al. A perivascular origin for mesenchymal stem cells in multiple human organs. Cell Stem Cell. 2008;3:301–13. [PubMed]
19. Nesti LJ, Jackson WM, Shanti RM, et al. Differentiation potential of multipotent progenitor cells derived from war-traumatized muscle tissue. J Bone Joint Surg Am. 2008;90:2390–8. [PubMed]
20. Jackson WM, Aragon AB, Bulken-Hoover JD, et al. Putative heterotopic ossification progenitor cells derived from traumatized muscle. J Orthop Res. 2009;27:1645–51. [PMC free article] [PubMed]
21. Grefte S, Kuijpers-Jagtman AM, Torensma R, et al. Skeletal muscle development and regeneration. Stem Cells Dev. 2007;16:857–68. [PubMed]
22. Bianco P, Robey P, Simmons P. Mesenchymal stem cells: revisiting history, concepts, and assays. Cell Stem Cell. 2008;2:313–9. [PMC free article] [PubMed]
23. Pountos I, Giannoudis PV. Biology of mesenchymal stem cells. Injury. 2005;36 (Suppl 3):S8–S12. [PubMed]
24. Caterson EJ, Nesti LJ, Danielson KG, et al. Human marrow-derived mesenchymal progenitor cells: isolation, culture expansion, and analysis of differentiation. Mol Biotechnol. 2002;20:245–56. [PubMed]
25. Djouad F, Jackson WM, Bobbick BE, et al. Activin A expression regulates multipotency of mesenchymal progenitor cells. Stem Cell Res Ther. 2010 Accepted. [PMC free article] [PubMed]
26. Ades EW, Candal FJ, Swerlick RA, et al. HMEC-1: establishment of an immortalized human microvascular endothelial cell line. J Invest Dermatol. 1992;99:683–90. [PubMed]
27. Xu Y, Swerlick RA, Sepp N, et al. Characterization of expression and modulation of cell adhesion molecules on an immortalized human dermal microvascular endothelial cell line (HMEC-1) J Invest Dermatol. 1994;102:833–7. [PubMed]
28. Lozito TP, Taboas JM, Kuo CK, et al. Mesenchymal stem cell modification of endothelial matrix regulates their vascular differentiation. J Cell Biochem. 2009;107:706–13. [PubMed]
29. Dominici M, Le Blanc K, Mueller I, et al. Minimal criteria for defining multipotent mesenchymal stromal cells. The International Society for Cellular Therapy position statement. Cytotherapy. 2006;8:315–7. [PubMed]
30. Conget PA, Minguell JJ. Phenotypical and functional properties of human bone marrow mesenchymal progenitor cells. J Cell Physiol. 1999;181:67–73. [PubMed]
31. Attinger CE, Bulan E, Blume PA. Surgical debridement. The key to successful wound healing and reconstruction. Clin Podiatr Med Surg. 2000;17:599–630. [PubMed]
32. Jiang Y, Mishima H, Sakai S, et al. Gene expression analysis of major lineage-defining factors in human bone marrow cells: effect of aging, gender, and age-related disorders. J Orthop Res. 2008;26:910–7. [PubMed]
33. Crisostomo PR, Wang M, Herring CM, et al. Sex dimorphisms in activated mesenchymal stem cell function. Shock. 2006;26:571–4. [PubMed]
34. Crisostomo PR, Markel TA, Wang M, et al. In the adult mesenchymal stem cell population, source gender is a biologically relevant aspect of protective power. Surgery. 2007;142:215–21. [PubMed]
35. Fehrer C, Lepperdinger G. Mesenchymal stem cell aging. Exp Gerontol. 2005;40:926–30. [PubMed]
36. Ray R, Novotny NM, Crisostomo PR, et al. Sex steroids and stem cell function. Mol Med. 2008;14:493–501. [PMC free article] [PubMed]
37. Muschler GF, Nitto H, Boehm CA, et al. Age- and gender-related changes in the cellularity of human bone marrow and the prevalence of osteoblastic progenitors. J Orthop Res. 2001;19:117–25. [PubMed]
38. Madonna R, Renna FV, Cellini C, et al. Age-dependent impairment of number and angiogenic potential of adipose tissue-derived progenitor cells. Eur J Clin Invest. 2010 [PubMed]
39. Zhou S, Greenberger JS, Epperly MW, et al. Age-related intrinsic changes in human bone-marrow-derived mesenchymal stem cells and their differentiation to osteoblasts. Aging Cell. 2008;7:335–43. [PMC free article] [PubMed]
40. Crisan M, Deasy B, Gavina M, et al. Purification and long-term culture of multipotent progenitor cells affiliated with the walls of human blood vessels: myoendothelial cells and pericytes. Methods Cell Biol. 2008;86:295–309. [PubMed]
41. Okamoto T, Aoyama T, Nakayama T, et al. Clonal heterogeneity in differentiation potential of immortalized human mesenchymal stem cells. Biochem Biophys Res Commun. 2002;295:354–61. [PubMed]
42. Qu-Petersen Z, Deasy B, Jankowski R, et al. Identification of a novel population of muscle stem cells in mice: potential for muscle regeneration. J Cell Biol. 2002;157:851–64. [PMC free article] [PubMed]
43. Gharaibeh B, Lu A, Tebbets J, et al. Isolation of a slowly adhering cell fraction containing stem cells from murine skeletal muscle by the preplate technique. Nat Protoc. 2008;3:1501–9. [PubMed]
44. Kolf CM, Cho E, Tuan RS. Mesenchymal stromal cells. Biology of adult mesenchymal stem cells: regulation of niche, self-renewal and differentiation. Arthritis Res Ther. 2007;9:204. [PMC free article] [PubMed]
45. Battula VL, Evans KW, Hollier BG, et al. Epithelial-mesenchymal transition-derived cells exhibit multilineage differentiation potential similar to mesenchymal stem cells. Stem Cells. 2010;28:1435–45. [PMC free article] [PubMed]
46. Corselli M, Chen CW, Crisan M, et al. Perivascular ancestors of adult multipotent stem cells. Arterioscler Thromb Vasc Biol. 2010;30:1104–9. [PubMed]
47. Tondreau T, Meuleman N, Delforge A, et al. Mesenchymal Stem Cells Derived from CD133-Positive Cells in Mobilized Peripheral Blood and Cord Blood: Proliferation, Oct4 Expression, and Plasticity. Stem Cells. 2005;23:1105–12. [PubMed]
48. Siggins RW, Zhang P, Welsh D, et al. Stem cells, phenotypic inversion, and differentiation. Int J Clin Exp Med. 2008;1:2–21. [PMC free article] [PubMed]
49. Levesque JP, Takamatsu Y, Nilsson SK, et al. Vascular cell adhesion molecule-1 (CD106) is cleaved by neutrophil proteases in the bone marrow following hematopoietic progenitor cell mobilization by granulocyte colony-stimulating factor. Blood. 2001;98:1289–97. [PubMed]
50. Fukada S, Yamamoto Y, Segawa M, et al. CD90-positive cells, an additional cell population, produce laminin alpha2 upon transplantation to dy(3k)/dy(3k) mice. Exp Cell Res. 2008;314:193–203. [PubMed]
51. Cizkova D, Soukup T, Mokry J. Nestin expression reflects formation, revascularization and reinnervation of new myofibers in regenerating rat hind limb skeletal muscles. Cells Tissues Organs. 2009;189:338–47. [PubMed]
52. Caplan AI. New era of cell-based orthopaedic therapies. Tissue Eng Part B Rev. 2009 [PMC free article] [PubMed]
53. Jackson WM, Nesti LJ, Tuan RS. Potential therapeutic applications of muscle-derived mesenchymal stem and progenitor cells. Expert Opin Biol Ther. 2010;10:1–13. [PMC free article] [PubMed]
54. Bulken-Hoover JD, Jackson WM, Zambrano I, et al. Optimization of neurotrophic factor production in mesenchymal stem and progenitor cells. Mol Biotechnol. 2010 Under Review.
55. Zhang X, Jiao C, Zhao S. Role of mesenchymal stem cells in immunological rejection of organ transplantation. Stem Cell Rev. 2009;5:402–9. [PubMed]
56. Haynesworth SE, Goshima J, Goldberg VM, et al. Characterization of cells with osteogenic potential from human marrow. Bone. 1992;13:81–8. [PubMed]