doi: 10.1161/hc0102.101442
(Circulation. 2002;105:93.)
© 2002 American Heart Association, Inc.
Basic Science Reports |
Human Mesenchymal Stem Cells Differentiate to a Cardiomyocyte Phenotype in the Adult Murine Heart
From the Department of Medicine, Division of Cardiology, Johns Hopkins School of Medicine (C.T., P.D.K.), and Osiris Therapeutics, Inc (M.F.P.), Baltimore, Md; and the Powell Gene Therapy Center, Departments of Pediatrics, Molecular Genetics, and Microbiology, University of Florida School of Medicine, Gainesville, Fla (K.S.C., B.J.B.).
Correspondence to Mark Pittenger, PhD, Osiris Therapeutics Inc, 2001 Aliceanna St, Baltimore, MD 21231. E-mail mpittenger{at}osiristx.com
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Abstract |
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Background— Cellular cardiomyoplasty has been proposed as an alternative strategy for augmenting the function of diseased myocardium. We investigated the potential of human mesenchymal stem cells (hMSCs) from adult bone marrow to undergo myogenic differentiation once transplanted into the adult murine myocardium.
Methods and Results— A small bone marrow aspirate was taken from the iliac crest of healthy human volunteers, and hMSCs were isolated as previously described. The stem cells, labeled with lacZ, were injected into the left ventricle of CB17 SCID/beige adult mice. At 4 days after injection, none of the engrafted hMSCs expressed myogenic markers. A limited number of cells survived past 1 week and over time morphologically resembled the surrounding host cardiomyocytes. Immunohistochemistry revealed de novo expression of desmin, ß-myosin heavy chain, 
-actinin, cardiac troponin T, and phospholamban at levels comparable to those of the host cardiomyocytes; sarcomeric organization of the contractile proteins was observed. In comparison, neither cardiac troponin T nor phospholamban was detected in the myotubes formed in vitro by MyoD-transduced hMSCs.
Conclusions— The purified hMSCs from adult bone marrow engrafted in the myocardium appeared to differentiate into cardiomyocytes. The persistence of the engrafted hMSCs and their in situ differentiation in the heart may represent the basis for using these adult stem cells for cellular cardiomyoplasty.
Key Words: stem cells • genes • myocytes • heart failure
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Introduction |
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Adult cardiac muscle, unlike skeletal muscle, lacks the ability to regenerate after ischemic injury, and death of cardiomyocytes promotes a cascade that results in heart failure. This has prompted interest in identifying cell types capable of replenishing the injured myocardium with healthy cells and augmenting heart function. Over the past decade, cardiac cellular transplantation techniques have made significant gains, and a variety of cell types have been proposed as useful candidates (see review1). A particular interest has developed for the use of skeletal muscle satellite cells and myoblasts, because these cells have the potential for autologous transplantation (see review2). We have previously demonstrated survival and differentiation toward a slow-twitch muscle phenotype of C2C12 myoblasts arterially delivered into the murine heart.3 Subsequent work demonstrated, in addition to myoblast survival in the injured myocardium, positive effects on remodeling and improvement in left ventricle performance.4,5 The first human cellular cardiomyoplasty was recently reported using autologous myoblasts, with encouraging results.6
The ideal candidate for cellular cardiomyoplasty is likely to be a less committed cell that can undergo full cardiogenic differentiation. Such a cell population might be found in the adult bone marrow. It is now accepted that an adherent population of cells isolated from bone marrow and expanded in vitro represents a potential source of undifferentiated mesenchymal stem cells (MSCs) that can give rise to connective tissue cell types. Such human MSCs (hMSCs) were first isolated and shown to have multiple differentiation potential by Haynesworth et al.7 Recently, we clonally isolated hMSCs, culture-expanded them, and demonstrated their multilineage potential by in vitro methods.8
In vivo evidence supports the idea that MSCs undergo myogenic differentiation. For example, a murine MSC population isolated from mouse bone marrow and injected into the quadriceps of the mdx mouse demonstrated that the cells could locally contribute dystrophin to the muscle fiber sarcolemma.9 When bone marrow cells from a normal male mouse were injected intravenously into the tail vein of affected mdx female mice, skeletal myotubes in the recipients were found to contain Y-positive nuclei.10 Mouse MSC-like cells converted to a myogenic phenotype after infusion and homing to cardiotoxin injured skeletal muscle.11 Also, after treatment with the DNA demethylation agent 5-azacytidine, a murine MSC-like cell line was shown to express cardiac differentiation markers and exhibit spontaneous membrane depolarization in vitro.12 Recently, Bittner et al13 showed that bone marrow–derived cells could be recruited to both skeletal and cardiac muscle in the mdx mouse, in which these striated muscles undergo continual remodeling.
In the results reported here, hMSCs were injected into the hearts of mice to test whether hMSCs may undergo "milieu-dependent differentiation." Early passage, multipotential hMSCs were tagged with ß-galactosidase and injected into the left ventricle of immunodeficient mice. The animals were killed at several time points, and the hearts were analyzed for the differentiation of the engrafted hMSCs.
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Methods |
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Isolation of hMSCs
hMSCs were isolated as previously described.7,8 Briefly, after signed informed consent, a bone marrow aspirate was collected from the posterior iliac crest of healthy volunteers into a syringe containing 6000 U heparin. The marrow sample was washed with Dulbecco’s PBS, and cells were recovered after centrifugation at 900g. Nucleated cells were counted, and 1x108 nucleated cells were loaded onto 25 mL of 1.073 g/mL Percoll solution in a 50-mL conical tube. Cells were centrifuged at 1100g for 30 minutes at 20°C. The nucleated cells were collected from the upper layer and interface, diluted with 2 volumes of Dulbecco’s PBS, and collected by centrifugation at 900g. The cells were cultured in DMEM (low glucose) containing 10% FBS selected for hMSC outgrowth.14,15 The experiments described here used hMSCs grown in FBS lot AFE5185 from Hyclone Inc.
Adenovirus Vectors
Replication-deficient recombinant adenoviruses carrying the ß-galactosidase reporter gene lacZ under the control of either cytomegalovirus immediate-early promoter (CMV) or Rous sarcoma virus long-terminal repeat (RSV-LTR) promoters were purchased from the University of Iowa Gene Vector Core (Dr Richard Anderson). To test the vectors, hMSCs were plated at a density of 10 000 cells per well (subconfluent). Incremental concentrations of virus over the range of 100 to 2000 multiplicity of infection (MOI) were applied overnight in serum-free DMEM to determine an optimal transduction efficiency. The next day, the virus supernatant was removed, and the cells were washed with several changes of DMEM containing 10% FBS. Two days later, the cells were fixed with 0.2% glutaraldehyde for 10 minutes, and X-gal staining was performed according to procedure (Kirkegaard and Perry Laboratories). The ß-galactosidase–positive (ß-gal+) blue cells found in 10 microscopic fields (x20) were counted and expressed as a percentage of the total number of cells in those fields.
Cell Transplantation
Cells obtained from 4 different human donors were used for cell transplantation experiments. First passage hMSCs were plated at low density in DMEM with 10% FBS. Two to 5 days later, the cells were washed with serum-free DMEM and infected overnight with AdRSVlacZ at 1000 MOI. The supernatant was then removed, and cells were washed with DMEM with 10% FBS. The medium was repeatedly changed over 2 days (
5 changes) to ensure complete removal of viral particles and to allow for internalization of any virus particles remaining on the surface. On the day of the surgery, the cells were harvested with 0.02% trypsin-EDTA (Sigma), washed with PBS, and resuspended. The cells were kept on ice until they were implanted, usually within 20 minutes.
Immunodeficient CB17 SCID/beige male adult mice (4 to 10 weeks old) were used as recipients. These mice lack the ability to mount an adaptive immune response, either B- or T-cell–mediated, and also lack functional natural killer cell activity. All animal handling and surgical procedures were performed in accordance with the Johns Hopkins University Animal Care and Utilization Committee review and institutional guidelines. Inhalatory methoxyflurane was used for general anesthesia. A volume of 100 µL of cell suspension containing 500 000 or 1 million cells was injected into the left ventricle with a 32-gauge needle through a transdiaphragmatic approach. Once blood backflow was observed in the syringe, the needle was advanced 
1 mm farther, and the cell suspension was slowly injected. The abdominal wall incision was then closed with 5.0 silk suture. An aliquot of the cells for injection was replated and histochemically stained to confirm continued expression of ß-galactosidase at 1 day and 1 and 2 months.
Immunohistochemistry
The 16 animals analyzed for cell engraftment in the study were killed with an overdose of methoxyflurane at 30 minutes (n=1) and 4 (n=3), 14 (n=3), 21 (n=2), 30 (n=4), and 60 (n=3) days after surgery. Tissues were harvested, cryoprotected in 30% sucrose for 4 hours, and snap-frozen in melting isopentane/dry ice. The tissues were embedded in Tissue-Tek OCT (Sakura), and 10- to 16-µm cryostat sections were obtained. During cryosectioning of the hearts, 3 adjacent sections were cut and placed on separate Superfrost Plus slides (VWR). The middle section of 3 was used for X-gal staining, and the adjacent sections on each side were processed for immunohistochemistry. Tissue sections to be stained for ß-galactosidase were fixed with 0.2% glutaraldehyde and 2% formaldehyde for 15 minutes at room temperature and then incubated in X-gal solution overnight at room temperature. For immunostaining, the tissue sections were fixed for 2 minutes at room temperature with either acetone or acetone/methanol 50%/50% vol/vol and then incubated with PBS containing 5% goat serum. The sections were incubated for 60 minutes with a polyclonal rabbit anti–ß-galactosidase antibody (Corning Biochem) at a concentration of 10 µg/mL in 2% goat serum in PBS, followed by FITC-conjugated goat-anti rabbit IgG for 45 minutes. After thorough washing, the second primary antibody was applied for an additional 60 minutes. These included either a mouse monoclonal antibody directed against desmin (clone D33, used at 1:100 dilution, Dako Corp), striated muscle -actinin (EA-53, 1:100, Sigma), ß-myosin heavy chain (MHC) (A4.1025, 1:10, DSHB, University of Iowa, Iowa City), MyoD (5.8A, 1:50, Novacastra Biochemicals), or phospholamban (MA3-922, 10 µg/mL, Sigma/RBI). This was followed by TRITC-conjugated goat anti-mouse IgG (1:200, Jackson ImmunoResearch Laboratory) for 45 minutes. Cardiac troponin T staining was performed with a goat polyclonal antibody (C19, diluted 1:20); in this case, a TRITC-conjugated donkey anti-goat antibody was applied first, followed by the FITC-conjugated goat anti-rabbit antibody for ß-galactosidase identification. Fluorescence imaging was performed with a Zeiss Axiovert equipped for epifluorescence or a Nikon PCM 200 confocal microscope.
In Vitro Myogenic Differentiation
For comparison, first-passage hMSCs were used for in vitro skeletal myogenic differentiation experiments with MyoD. The cells were plated on 12-well plastic culture dishes at high density (50 000 cells/well) and infected the next day with an adenovirus containing the human MyoD gene under the control of the RSV promoter (kindly donated by Dr Charles Murry, University of Washington, Seattle) at 1000 MOI. The cells were maintained in low-serum medium (2% horse serum) for 1 month and processed for immunohistochemistry with the same antibodies and methods as described.
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Results |
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Stem Cell Transduction
The initial labeling experiments revealed that hMSCs could be efficiently transduced by use of recombinant adenoviruses at high MOI. The RSV promoter was more effective than the CMV promoter in expressing the lacZ gene in these cells (Figure 1). For the AdRSVlacZ, the maximum number of cells expressing ß-galactosidase was achieved at 1000 to 1500 MOI; this concentration was used for labeling cells for implantation experiments. Samples of transduced hMSCs maintained in culture revealed that the number of cells expressing ß-galactosidase was 90% at 1 month and 60% at 2 months.
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In Vivo Engraftment of Stem Cells
The goal of these experiments was to assess the ability of hMSCs to engraft and differentiate in the adult myocardium. For this, 5x105 to 10x105 AdRSVlacZ-labeled hMSCs were injected through the diaphragm into the left ventricular chamber of immunodeficient mice. At 4 days after injection, the majority of the ß-gal+ cells were identified in the spleen, liver, and lungs (Figure 2, B through D). In the heart, we did not identify hMSCs in 4 of 16 animals, probably as a result of the technical difficulties associated with injecting into the beating myocardium. In 12 animals, hMSCs were dispersed throughout the myocardium as individual cells at 4 to 60 days (Figure 2A and Figure 3). Quantification of the ß-gal+ hMSCs from serial sections estimated that 2200 hMSCs survived in the left ventricle 4 days after the injection of 500 000 cells (0.44%). Fewer hMSCs were identified in the myocardium at later time points, but 1 ß-gal+ cell per tissue section was found.
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With time, most of the ß-gal+ cells became morphologically indistinguishable from the surrounding cardiomyocytes. At 14 days and beyond, the persistent ß-gal+ hMSCs were rod-shaped and well aligned with the host cardiomyocytes (compare Figure 3A with 3B through 3D). Moreover, many of the engrafted cells became enlarged from a diameter of 20 to 30 µm at 4 days to a long axis of 50 to 70 µm and a diameter of 20 µm at 2 months. Immunofluorescence staining for ß-galactosidase also confirmed the presence of the engrafted hMSCs, although this method was less sensitive than X-gal staining. Double-label immunofluorescence staining with an anti–ß-galactosidase antibody was used to identify engrafted cells, and additional antibodies characterized the myogenic protein expression of the engrafted stem cells. Monoclonal antibodies directed against desmin, -actinin, phospholamban, ß-MHC, cardiac troponin T, and MyoD were used and visualized with a rhodamine-conjugated secondary antibody. None of the hMSCs from 4 different donors used in this study were reactive with these muscle protein antibodies when stained during in vitro culture.
Neither the hMSCs at the injection site nor those dispersed in the myocardium expressed desmin at 4 days (Figure 4, A and B, respectively). Desmin- and cardiac troponin T–expressing ß-gal+ cells were first identified at 14 days, whereas all identified hMSCs were positive for desmin expression at 60 days. Figure 5A clearly shows the double-positive cells in the same optical plane with anti–ß-galactosidase labeling (fluorescein green) and the cardiac troponin T labeling (rhodamine red) at 14 days. The ß-gal+ cells were also positive for -actinin, a component of the Z bands shown at 60 days in Figure 5B. Similarly, engrafted cells were positive for MHC (Figure 5C). Engrafted hMSCs were also found to express phospholamban, a phosphoprotein that plays a key role in modulating the cardiac sarcoplasmic reticulum Ca2+-ATPase (Figures 5D, 6E, and 6F). High-magnification imaging of these cells showed sarcomeric organization of the -actinin (Figure 6, C and D), cardiac troponin T, and desmin (not shown).
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None of the hMSCs identified on the cardiac sections reacted with the MyoD antibody. To further distinguish between cardiac and skeletal myogenic differentiation of the hMSCs implanted in the heart, we analyzed the expression of these markers to those expressed in myotubes formed by MyoD-transduced hMSCs in vitro. The MyoD-transduced hMSCs expressed high levels of nuclear MyoD shortly after infection with AdRSVMyoD. After MyoD transduction, adjacent hMSCs fused and formed multinucleated myotubes. At 1 month, the differentiated cells expressed desmin, -actinin, and ß-MHC, but no detectable levels of phospholamban or cardiac troponin T were observed (Figure 7). Again, sarcomeric organization of the contractile proteins was observed with high magnification, indicating differentiation to a mature myotube.
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Discussion |
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The isolation procedure for hMSCs from adult bone marrow used in this study yields a homogeneous cell population capable of differentiation to several mesenchymal lineages.8 At 4 days, it was noted that none of the ß-gal+ cells expressed desmin, which rules out the possibility that AdRSVlacZ particles were carried along with the hMSCs and infected native cardiomyocytes. hMSCs were found at the site of the needle injection very early, but at later times, the surviving hMSCs were mostly monodisperse. These persistent cells were found far from the needle site and most likely entered the myocardium by crossing the endothelium, although direct evidence of this is not available. This process may be similar to the tethering of leukocytes to the endothelium and involve integrins and other adhesion molecules present on the surface of hMSCs.8 In addition, capillary plugging and resultant microischemia may play a role in increasing the endothelial permeability. Transendothelial migration from the coronary capillaries and integration of cells into mouse cardiac muscle have been observed previously with both myoblasts3 and bone marrow–derived cells.11,13 Clearly, further understanding of these mechanisms of migration and cell localization may provide for more efficient myocardial delivery of hMSCs and less invasive delivery options.
Desmin is known to represent an early marker of myogenic differentiation synthesized by muscle progenitor cells. In these experiments, none of the engrafted hMSCs expressed desmin at 4 days, whereas at 60 days, all of the identifiable ß-gal+ cells expressed desmin. The hMSCs also expressed cardiac-specific troponin T, ß-MHC, -actinin, and phospholamban (Figure 5), and sarcomeric striations were evident at high magnification. Significantly, the intensity of immunostaining for these proteins in the engrafted hMSCs was indistinguishable from the surrounding cardiomyocytes.
The expression of myogenic markers in the engrafted cells was compared with the expression found in myotubes obtained by hMSCs expressing MyoD in vitro. It has been shown that forced expression of MyoD in cells of mesodermal origin results in myotube formation of both the fast- and slow-twitch types, indistinguishable from those formed by primary myoblasts.16,17 The MyoD-transduced hMSCs expressed desmin, -actinin, and ß-MHC but no detectable phospholamban or cardiac troponin T, two constitutive proteins restricted mainly to cardiac muscle. Together, the data are suggestive of conversion of engrafted hMSCs to a mature cardiac muscle rather than slow-twitch skeletal muscle. Although it is theoretically possible that the hMSCs have converted to a skeletal muscle phenotype and then adapted to the cardiac environment as suggested for C2C12 cells,3 we find this unlikely, given the data.
Bone marrow MSCs, as well as cardiomyocytes, are derived from early mesoderm, although they form from different embryonic regions. The cardiomyocyte progenitors form from a ventrally situated portion of the mesoderm, becoming committed to their fate under the influence of differentiation factors secreted by the closely adjacent endoderm (see review18). Adult cardiomyocytes actively produce insulin-like growth factor, transforming growth factor (TGF)-ß,19 and heparin-binding epidermal growth factor–like growth factor,20 which act as autocrine growth stimuli for cardiomyocytes. TGF was shown to induce cardiac differentiation in an avian embryonic mesodermal cell line,21 and the TGF-ß signaling molecule activin is involved in early cardiogenesis in amphibians and birds.18 Locally produced neuregulins promote survival and growth of cardiac myocytes via ErbB2 and B4 receptors, which also bind heparin-binding epidermal growth factor–like factor.22 In the present study, the differentiation process observed in vivo most likely involves a combination of paracrine growth signals and the electrical and mechanical stimulation present in the adult heart.
The recent reports of the ability of marrow cells to migrate from bone marrow and integrate and differentiate into the damaged skeletal and cardiac muscle suggest that such a process may normally contribute to tissue maintenance or regeneration,11,13,23 although this process may not be efficient.24 Other recent reports have demonstrated the potential for rat or mouse marrow cells to engraft in the infarcted myocardium14,15,23,25 and offer further support of this possibility. The present study demonstrates the ability of adult hMSCs to integrate and undergo striated muscle differentiation in the adult heart and opens the possibility for using these human adult stem cells for therapeutic cardiomyoplasty.
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Acknowledgments |
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This work was supported by the National Institute of Standards and Technologies Advanced Technology Program (NIST/ATP) CA#70NANB7H3061 and Osiris Therapeutics, Inc. We thank Dr Charles Murry for a gift of adenovirus encoding the human MyoD, and Miroslava Burysek, Jonathan Golob, Matthew Bernabei, and Stacey Porvasnik for excellent technical assistance. We also thank Drs Bradley Martin and Robert Deans for critical comments on the manuscript.
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Footnotes |
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Dr Toma is now at St Luke-Roosevelt Hospital, Department of Medicine, New York, NY; Dr Kessler is now at Gen-Vec Inc, Gaithersburg, Md.
Received June 28, 2001; revision received October 15, 2001; accepted October 25, 2001.
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