Published online before print February 15, 2007, doi: 10.1161/01.RES.0000260803.98329.1c
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© 2007 American Heart Association, Inc.
Cellular Biology |
Fusion of Human Hematopoietic Progenitor Cells and Murine Cardiomyocytes Is Mediated by 
4ß1 Integrin/Vascular Cell Adhesion Molecule-1 Interaction
From the Departments of Cardiology (S.Z., E.T.H.Y.) and Bone Marrow Transplantation (E.S.), The University of Texas M.D. Anderson Cancer Center; Brown Foundation (J.T.W., E.T.H.Y.), Institute of Molecular Medicine for the Prevention of Human Diseases, The University of Texas–Houston Health Science Center; and Texas Heart Institute (J.T.W., E.T.H.Y.), St. Luke’s Episcopal Hospital, Houston, Tex.
Correspondence to Edward T. H. Yeh, MD, Department of Cardiology, Box 449, University of Texas, M.D. Anderson Cancer Center, Houston, TX 77030. E-mail etyeh{at}mdanderson.org
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Abstract |
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Fusion of transplanted stem cells and host cells has been proposed as a major mechanism for the generation of hepatocytes, Purkinje neurons, and cardiomyocytes. However, the mechanism of cell fusion has not been precisely defined. Furthermore, the consequence of cell fusion remains unclear. We have previously shown that adult peripheral blood CD34-positive cells injected into severe combined immune deficiency (SCID) mice can transform into cardiomyocytes, endothelial cells, and smooth muscle cells following experimentally induced myocardial infarction and that most of the newly formed cardiomyocytes result from cell fusion. We therefore undertook this study to define the mechanism and consequences of cell fusion. Here we show that hypoxia and cytokines increase fusion of human peripheral blood CD34-positive cells and murine cardiomyocytes in vitro by up to 7-fold, and this is blocked by anti-
4ß1 or anti–vascular cell adhesion molecule (VCAM)-1. In vivo, fusion of progenitor cells and cardiomyocytes can also be blocked by anti-4ß1 or anti–VCAM-1, but not by anti–vascular endothelial growth factor. On the other hand, generation of human-derived endothelial cells is blocked by anti–vascular endothelial growth factor but not by anti-4ß1 antibodies. Two months following transplant, a high percentage of fused cells expressed cyclin B1 and incorporated bromodeoxyuridine. Thus, hematopoietic progenitor cell and cardiomyocyte fusion is mediated by 4ß1/VCAM-1 interaction, leading to cell cycle reentry and cellular proliferation.
Key Words: adult stem cell • myogenesis • angiogenesis
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Introduction |
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TopAbstractIntroductionMethods and MaterialsResultsDiscussionReferences |
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Fusion of transplanted stem cells and host cells has been proposed as a major mechanism for the generation of hepatocytes, Purkinje neurons, and cardiomyocytes.1–8 We have previously shown that adult peripheral blood CD34-positive cells injected into severe combined immune deficiency (SCID) mice can transform into cardiomyocytes, endothelial cells, and smooth muscle cells following experimentally induced myocardial infarction (MI)9 and that most of the newly formed cardiomyocytes result from cell fusion.10 However, the mechanism and the consequences of cell fusion remain unclear. This study is designed to define the factor(s) mediating cell fusion in the injured heart and to examine the consequence of fusion.
Adult progenitor cells derived from either the bone marrow or peripheral blood have been used to treat patients following acute MI or with chronic ischemic cardiomyopathy. Although most of the studies show benefit at clinical parameters, the precise mechanism whereby progenitor cell therapy benefit the heart is unclear.11–14 There is limited evidence for angiogenesis.15 However, there has not been any evidence for true myogenesis. It is difficult to show myogenesis in the transplant of autologous progenitor cells to the injured heart because there are no suitable markers. We used the human to mouse xenotransplant system to allow for tracking of human-derived cardiomyocytes or endothelial cells using human leukocyte antigen (HLA) as a convenient marker. Furthermore, we developed an in vitro model of cell fusion to identify the molecules that are involved in the fusion process. Vascular cell adhesion molecule (VCAM)-1 and 4ß1 were involved in the fusion of human progenitor cells with a murine cardiomyocyte cell line in vitro. The biological relevance of this finding was confirmed by blocking the generation of human-derived cardiomyocytes in vivo by pretreating the animals with anti–VCAM-1 or anti-4ß1 antibodies, but not by anti–vascular endothelial growth factor (VEGF) antibody. On the other hand, pretreatment with anti-VEGF antibody blocked the generation of human-derived endothelial cells, but not cardiomyocytes. Finally, we also studied the consequence of cell fusion and found that cell fusion could reprogram the newly fused cells to enter cell cycle and divide.
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Animals
Female SCID mice (C3H; The Jackson Laboratory, Bar Harbor, Me) were used as the recipients of human peripheral blood CD34-positive cells. Experimental MI was induced by ligation of the left anterior descending branch of the coronary artery. Control mice were treated with MI without cell transplantation. For the bromodeoxyuridine (BrdUrd) incorporation study, mice were injected with BrdUrd (50 mg/kg, IP; BD PharMingen, San Jose, Calif,) 4 hours before euthanasia. The Institutional Animal Care and Use Committees of The University of Texas Health Science Center and The University of Texas M.D. Anderson Cancer Center, Houston, approved all animal procedures, including human peripheral CD34-positive cell transplantation.
Cell Cultures
HL-1 cells16 were provided by Dr William Claycomb (Louisiana State University, New Orleans) and maintained in Claycomb medium (JRH Biosciences, Lenexa, Kan). The cells were cultured in 60-mm Petri dishes at 37°C in 95% air and 5% CO2. Cells from passages 67 to 75 were used in the experiments.
In Vitro Detection of Adhesion Molecules
HL-1 cells were harvested by trypsinization, washed twice in cold PBS containing 1% FBS, and then incubated with rat anti-mouse VCAM-1 conjugated with phycoerythrin (PE) or rat anti-mouse intercellular adhesion molecule (ICAM)-1 conjugated with fluorescein isothiocyanate (FITC) (1:100, Chemicon International, Temecula, Calif). Human CD34-positive cells were washed twice in cold PBS and then incubated with FITC- or PE-conjugated mouse anti-human 1ß4 (1:200) or lymphocyte function–associated antigen (LFA)-1 (1:100; both from Serotec, Raleigh, NC), or respected isotype control antibodies. After incubation cells were washed twice in cold fluorescence-activated cell-sorting (FACS) buffer (PBS containing 0.5% FBS and 0.1% sodium azide) and analyzed using an LSRII flow cytometer (BD Biosciences, San Jose, Calif).
Induction of MI
Experimental MI was induced using a previously described method.9 Briefly, mice were anesthetized. The animals were ventilated with 100% oxygen using a rodent ventilator (Inspira ASV; Harvard Apparatus Inc, South Natick, Mass). The chest was opened through a midline sternotomy, and a 7-0 silk suture (Ethicon Inc, Johnson & Johnson Co, Somerville, NJ) was passed with a tapered needle under the left anterior descending coronary artery 1 to 2 mm from the tip of the left auricle and tied. Human peripheral CD34-positive cells (2x106) suspended in 0.3 mL of saline were injected into the myocardium of the left ventricles 30 to 40 minutes after MI. Control mice underwent the same MI procedure but no cell transplantation.
Preparation of Single-Cell Suspension From the Heart
A single-cell suspension was obtained by enzyme digestion of the mouse heart, as described previously.17 The heart was perfused with ice-cold buffer containing (g/L) 6.8 NaCL, 0.4 KCl, 1.0 Glucose, 1.5 NaH2PO4, 0.1 MgSO4, and 4.76 Hepes, pH 7.38 before being removed from the animal, after which it was cut into six pieces and placed in a 10-mL beaker containing 2 mL of the enzymatic solution, the above buffer containing elastase (0.16 mg/mL; Worthington Biochemical, Freehold, NJ), type II collagenase (0.21 mg/mL; Worthington Biochemical), and pancreatin (0.6 mg/mL; Sigma). The tissues were then incubated for five 15-minute periods at 37°C with shaking (70 rpm). Cells were harvested after each incubation and suspended in FBS. The isolated cells were filtered through a cell strainer with a pore size of 70 µm.
FACS Analysis of Cells Isolated From the Heart
For the FACS analysis of cells isolated from the heart, cells were washed, fixed, and permeabilized.10 Cells were then incubated with monoclonal anti–cardiac troponin T (1:200, clone 1A11; Advanced Immunochemical) for 30 minutes at 4°C and washed twice with wash buffer. After incubation with the secondary antibody (goat anti-mouse IgG conjugated with Alexa Fluor 633 [Molecular Probes]) for 30 minutes, cells were washed 3 times, then incubated with PE-conjugated anti-human human leukocyte-associated antigen-ABC (HLA-ABC) (clone W6/32; Cedarlane Laboratories) before FACS analysis for cell fusion. For cell cycle marker detection, the cardiac troponin T–stained cells were incubated with FITC-conjugated anti–cyclin B1 (BD Biosciences, Mountain View, Calif) or with FITC-conjugated anti-BrdUrd (BD Biosciences, San Jose, Calif) for 30 minutes at 4°C. After 2 washes, cells were incubated with PE-conjugated anti-human HLA-ABC (clone W6/32; Cedarlane Laboratories). After 2 final washes, cells were analyzed using a FACSAria flow cytometer (BD Biosciences). Control cells were incubated with Alexa Fluor 633–conjugated anti-mouse IgG and with FITC- and PE- conjugated isotypic IgGs for anti–cyclin B1 and anti–HLA-ABC. To eliminate spillover of the different fluorochromes, individually stained cells were used to determine the compensation coefficient. In a triple-stained sample, a cyclin B1–positive or a BrdUrd-positive gate was established within the population of cells simultaneously expressing HLA-ABC and cardiac troponin T to distinguish the cells expressing all 3 markers. The triple-positive cells were then collected using a FACSAria flow cytometer. The collected cells were exposed to bright white light for 96 hours at 4°C until all fluorescence was bleached out.
Fluorescent In Situ Hybridization Analysis
The cells were spun onto a slide and fixed immediately with 3:1 methanol/acetic acid solution for 30 minutes. Complete diminishment of the residual fluorescence was confirmed by examination under an epifluorescence microscope (Nikon Eclipse TE-2000-U). Slides were then briefly fixed again in 3:1 methanol/acetic acid, then were predenatured, dehydrated, denatured, and hybridized with a FITC-conjugated DNA probe for mouse X chromosomes (ID Labs, London, Ontario, Canada) and a PE-conjugated probe for human X chromosomes (Qbiogene, Carlsbad, Calif) overnight at 37°C in a humidified chamber. After a posthybridization wash, slides were counterstained with 4',6-diamidino-2-phenylindole (DAPI) (0.02 µg/mL) and examined using an epifluorescence microscope (Nikon Eclipse TE 2000-U).
Statistics
Student t test was used in comparisons between means of 2 groups. When three or more means were compared, 1-way ANOVA followed by multiple comparisons among means was used.
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Results |
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Spontaneous Fusion Between Human CD34-Positive Cells and HL-1 Cells in Vitro
First, we established an in vitro model that could recreate the fusion of transplanted human peripheral blood CD34-positive cells and mouse cardiomyocytes observed in vivo.10 HL-1 mouse cardiomyocytes were cocultured with human peripheral blood CD34-positive cells at a 1:1 ratio for different times before cell fusion was analyzed using a combination of FACS and in situ fluorescence hybridization (FISH). We showed that HL-1 cells and human peripheral blood CD34-positive cells spontaneously fused at 24, 48, or 72 hours after the start of the coculture, as shown by HLA and troponin T positivity (Figure 1A and 1B). Under this simple coculture condition, the fused cells constituted less than 0.2% of the cells in culture. To confirm that HLA-positive, troponin T–positive cells were indeed resulted from fusion, and not transdifferentiation, the double-positive cells were sorted and analyzed using the FISH technique. Nuclei (227) were counted in 3 experiments, and 92±2.1% contained both human and mouse X chromosomes. A representative figure of nuclei analyzed by FISH is shown in Figure 1C. Human CD34+ cells (Figure 1D) and mouse cardiomyocytes (Figure 1E) were also hybridized with both human and mouse X chromosome probes to confirm the specificity of the FISH results. Thus, synkaryon formed as a result of coculturing of human CD34-positive cells with mouse cardiomyocytes.
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Hypoxia and Cytokines Enhance Cell Fusion
Heart damage can significantly increase in vivo transformation of transplanted stem cells into cardiomyocytes.7,9,18 To increase the fusion rate, we therefore simulated in vitro the hypoxia conditions that are present during an MI by culturing HL-1 cells in 5% CO2 and 95% N2 24 hours before coculturing them with human peripheral blood CD34-positive cells at a 1:1 ratio for another 24 hours. This significantly enhanced the fusion, compared with untreated cells (Figure 2A). The fused cells now constituted 0.36% of the cells in culture (Figure 2B). This showed that our in vitro model is able to recreate the in vivo conditions. Only HL-1 cells were exposed to hypoxia in the following experiments because human CD34-positive cells do not survive the hypoxic treatment.
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We then tested whether 4 different cytokines with concentrations that are elevated following acute MI19–21 might further augment cell fusion in the presence of hypoxia. When interleukin (IL)-4 or stromal cell–derived factor-1 was added to coculture of hypoxia-treated HL-1 cells and CD34-positive cells, there was no significant increase in the percentage of double-positive cells as compared with control (Figure 2C and 2D). However, with the addition of either IL-6 or tumor necrosis factor (TNF)-, the percentage of double-positive cells increased (Figure 2E and 2F). The best result was seen after combined treatment with IL-6 and TNF-, which caused the percentage of double-positive cells to increase 7.5-fold compared with control cells (Figure 2G).
To verify that these double-positive cells were a result of fusion, the cells were sorted and analyzed by FISH. The nuclei of more than 95% of the double-positive cells contained both human and mouse X chromosomes, whereas the cells positive only for cardiac troponin T contained only mouse X chromosomes. Representative nuclei are shown in Figure 2H and 2I, respectively. Thus, we successfully increased the rate of human CD34-positive cell and mouse cardiomyocyte fusion by up to 7.5-fold under conditions that exist during an MI.
4ß1/VCAM-1 Adhesion Molecule Pair Mediates Cell Fusion
We hypothesized that close contact of cells, which is mediated by surface adhesion molecules, is a prerequisite for cell fusion and that the increase in cell fusion is attributable to the induction/activation of adhesion molecules by cytokines and/or hypoxia.22–26 Thus, we examined surface expression of several key adhesion molecules that are induced by IL-6 and TNF-. We showed that 4ß1 was induced in human CD34-positive cells following combined treatment with IL-6 and TNF- (Figure 3A) but not with IL-4 and stromal cell–derived factor-1. Furthermore, VCAM-1 expression was increased in HL-1 cells following exposure to hypoxia (Figure 3B). However, there was no increase in ICAM-1 (Figure 3C) or LFA-1 (Figure 3D) expression on either human CD34-positive cells or HL-1 cells after the same treatment.
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To prove that the induction of the VCAM-1 or 4ß1 adhesion molecule pair is relevant to the fusion process, we tested whether fusion could be blocked in vitro when cells in coculture were incubated with anti-4ß1 antibody as opposed to an isotype-control antibody. As shown in Figure 3E, fusion was blocked by up to 70% in cells treated with the anti-4ß1 antibody, but not in cells treated with an isotype-matched control antibody. This is specific because anti–VCAM-1 also blocked up to 70% (Figure 3F), but anti-ICAM-1 could not block (data not shown).
We then sought to show whether a similar mechanism is also responsible for cell fusion in vivo. For this purpose, a SCID mouse was subjected to experimentally induced MI, as previously described.10 However, instead of injecting CD34-positive cells intraventricularly, human CD34-positive cells were injected directly into the myocardium adjacent to the infarct area to avoid the potential confounding effect of the antibody on blocking the homing of the injected cells. Isotype-matched control antibody or anti-4ß1 antibody was injected intraperitoneally 30 minutes before the injection of CD34-positive cells. A single-cell suspension was then prepared by enzymatic digestion of the heart, which was harvested 4 days after human CD34-positive cell injection and then analyzed by FACS. As shown in Figure 4A, anti-4ß1 antibody reduced the percentage of double-positive cells by 75% compared with those from animals treated with the isotype-matched control antibody. To verify that cell fusion, and not transdifferentiation, was blocked by the anti-4ß1, we collected the double-positive cells and subjected them to FISH analysis. We showed that more than 95% of the nuclei of the double-positive cells treated with isotype-matched control antibody contained both human and mouse X chromosomes, as did the double-positive cells treated with the anti-4ß1 antibody. Thus, anti-4ß1 blocked cell fusion in vivo.
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Because we have previously shown that injection of human CD34-positive cells into SCID mice following MI leads to the generation of endothelial cells through transdifferentiation, we reasoned that transdifferentiation is fundamentally different from cell fusion and should not be blocked by the anti-4ß1 antibody. We further reasoned that this would provide good control for our in vivo blocking experiments because it would enable us to demonstrate the blocking of cell fusion, but not transdifferentiation, in the same animal treated with anti-4ß1 antibody. To verify this, SCID mice were injected with isotype-matched or anti-4ß1 antibodies. Cell preparations used to analyze the fusion of CD34-positive cells, and cardiomyocytes were also used to analyze cells positive for HLA and vascular endothelial (VE)-cadherin, which are markers for transdifferentiation. Anti-4ß1 antibody significantly reduced the percentage of HLA-positive, troponin T–positive cells (Figure 4A) but not the percentage of HLA-positive, VE-cadherin–positive cells (Figure 4B). Similar findings were obtained when anti–VCAM-1 antibody was used (Figure 4C and 4D). As expected, the HLA-positive, VE-cadherin–positive cells contained only human X chromosomes, as detected by FISH (Figure 4G). This indicated that anti-4ß1 antibody blocks fusion, but not transdifferentiation, of these cells in vivo. To examine whether blocking the generation of donor cell–derived endothelial cells also inhibits myogenesis caused by cell fusion, mice were injected with anti-VEGF (50 µg/mouse, IP) or isotype control. We assumed that anti-VEGF treatment should reduce the generation of endothelial cells from human CD34+ cells, but not directly influence cell fusion. Results showed that anti-VEGF treatment significantly reduced the percentage of HLA+/VE-cadherin+ cells without affecting the percentage of HLA+/cardiac troponin T+ cells (Figure 4E and 4F).
Cardiomyocytes Reentered Cell Cycle After Fusion
It has been shown that cardiomyocytes can fuse with a variety of cells in vitro, which may cause them to reenter the cell cycle.27 Furthermore, a study involving an in vitro fusion method showed that human embryonic stem cells can fuse with somatic cells, which ultimately reprograms the fused cells, causing them to enter an embryonic state from which they can proliferate and differentiate.28 We therefore hypothesized that fusion of human peripheral blood CD34-positive cells with terminally differentiated mouse cardiomyocytes would cause the fused cells to reenter the cell cycle and proliferate. To test this hypothesis, mouse hearts were harvested either 1 to 4 days or 2 months after the infusion of human CD34-positive cells into a previously described SCID mouse MI model.9,10 Hearts were then digested with enzymes, and single-cell preparations were examined by FACS analysis to determine the dual expression of cardiac troponin T and HLA, an indicator of cell fusion. In the control experiment, in which MI was induced in the absence of the CD34-positive cell infusion, no HLA-positive cells were identified, as expected (Figure 5A). When CD34-positive cells were infused into the MI model, 
1% of the single-cell preparation was HLA positive and troponin T positive when examined at days 4 and 30 and 0.9% at day 60 (Table I in the online data supplement), suggesting that the cells were derived from infused human cells (Figure 5B). Furthermore, injection with CD34+ cells improved cardiac function examined by using MRI (supplemental Figure I). The dual-positive cells were then collected for FISH analysis, which was performed using specific probes for human and mouse X chromosomes, and at least 90% of the nuclei contained both the human and mouse X chromosomes, suggesting that cell and nuclear fusion had occurred (Figure 6A). Because the donor was male, only 1 human X chromosome was present, and 2 mouse X chromosomes were present in the female recipient cells. In contrast, all of the nuclei in the HLA-negative, troponin T–positive cells (mouse cardiomyocytes) expressed only 2 mouse X chromosomes (Figure 6B).
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Close examination of the nuclei of fused cells revealed that a number of the nuclei were in the process of dividing, which suggests that these terminally differentiated cardiomyocytes were actively in the cell cycle. (Figure 6C). In addition, 2 different sets of human and mouse X chromosomes were observed in the separating nuclei.
To confirm that the cells were actively dividing, we performed a 3-color cell-sorting analysis using HLA, troponin T, and cyclin B1, a specific marker for the G2-to-M phase of the cell cycle, as 3 distinct markers. Single-cell suspensions prepared from hearts harvested from SCID mice 4 days after MI were examined first. In nontransplanted hearts, the cyclin B1–positive cells were present in the cardiac troponin T–negative population (data not shown). However, in hearts from mice transplanted with human CD34-positive cells, 75.7% of the HLA-positive, troponin T–positive cells expressed cyclin B1 (Figure 5C). Furthermore, cyclin B1–positive cells made up less than 1% of the HLA-negative, troponin T–positive cell population. In addition, cells from hearts harvested 2 months after MI showed that 35.7% of the HLA-positive, troponin T–positive cells expressed cyclin B1. Again the cyclin B1–positive cells made up less than 1% of the HLA-negative, troponin T–positive cell population. Thus, cyclin B1–positive cells were highly enriched in the cell population that was derived mostly from fusion (HLA-positive, troponin T–positive cells) but not in the nonfused mouse cardiomyocytes (HLA-negative, troponin T–positive cells).
To confirm these results, we injected mice after transplantation with bromodeoxyuridine (BrdUrd, 50 mg/kg, IP) 4 hours before euthanasia to label the cells that had entered the S phase. BrdUrd-labeled cells were analyzed by 3-color fluorescent flow cytometry using HLA, troponin T, and BrdUrd as markers. In hearts from nontransplanted mice, BrdUrd-labeled cells were detected in only the noncardiomyocyte population (data not shown). However, as shown in Figure 5D, in hearts from the transplanted mice harvested on day 4 after MI, 89.0% of the cells in the HLA-positive, troponin T–positive population, as opposed to only 0.7% of the HLA-negative, troponin T–positive population, had incorporated BrdUrd into their DNA. In hearts harvested 2 months after MI, 76.4% of the cells in the HLA-positive, troponin T–positive population had incorporated BrdUrd into their DNA. Again, the HLA-negative, troponin T–positive population showed little evidence of BrdUrd incorporation. The results regarding the in vivo expression of cyclin B1 and the incorporation of BrdUrd in different cell populations as of days 4 and 60 after cell transplantation are summarized in Figure 5E and 5F. The BrdUrd results support our cyclin B1 data that most of the HLA-positive, troponin T–positive cell population is active in the cell cycle 4 days after human CD34-positive cell infusion, with evidence of cell cycling persisting for up to 2 months.
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Our results demonstrate that local environmental factors, such as hypoxia and cytokine release, which occur after MI promote the fusion of transplanted CD34-positive cells with the host cardiomyocytes, and that the cell adhesion molecules VCAM-1 and
1ß4 are the mediators involved in this fusion process. In adults, VCAM-1 is well known for its role in the recruitment of mononuclear cells during vascular inflammation. In embryos, on the other hand, VCAM-1 is required for chorioallantoic fusion and placentation29 and is essential for the development of the extraembryonic circulatory system and the embryonic heart.30 Thus, it is not surprising that the 4ß1/VCAM-1 adhesion molecule pair plays a critical role in the fusion of human progenitor cells with murine cardiomyocytes. However, we currently still do not fully understand the exact mechanism whereby the interaction of these 2 molecules increase the fusion rate. The observation that cell fusion can be blocked in vivo suggests to us that this interaction also play a critical role in cell fusion in vivo. Clinical studies on myocardial regeneration have shown a general beneficial effect in cardiac function after stem cell transplantation.14,31 However, it is difficult to explain the beneficial effect by simple fusion between the transplanted cells and the host cardiomyocytes, especially when the fate of the fused cells is unknown.4 The present study provides in vivo evidence that cell fusion triggers the reentry of fused cardiomyocytes into the cell cycle. The resultant proliferation of the fused cells, therefore, may contribute to the regeneration and, hence, healing, of damaged myocardial tissue.
Autologous adult progenitor cells have been used to treat patients following acute MI or with ischemic cardiomyopathy. Although many of these studies have shown clinical benefit, the mechanisms whereby adult progenitor cells repair the damaged myocardium remain unclear. It is not known whether angiogenesis, myogenesis, or both have taken place following transplanting progenitor cells into the diseased heart. To provide a rational basis for determining the true benefit of adult progenitor cell therapy for damaged heart, it is critically important to determine the relative contribution of angiogenesis versus myogenesis in the repair process. Results from our study suggest that it is possible to determine the beneficial effects on functional improvement in the heart that are observed after stem cell transplantation are attributable to myogenesis or angiogenesis or both by using selective antibody blockade. Our results show that myogenesis resulted mostly from fusion between transplanted stem cells and the host cardiomyocytes can be blocked in vivo by anti-4ß1. Angiogenesis, on the other hand, can be blocked by anti-VEGF. This could be very useful in determining the relative contribution of myogenesis and angiogenesis toward cardiac repair in the future.
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Acknowledgments |
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Sources of Funding
This work was supported by a Multidisciplinary Research Proposal from the M.D. Anderson Cancer Center (principal investigator, E.T.H.Y.).
Disclosures
None.
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Footnotes |
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Original received December 7, 2006; revision received January 29, 2007; accepted February 2, 2007.
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References |
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