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. 2009 Feb;57(2):167–176. doi: 10.1369/jhc.2008.952507

Histopathological Study of Healing After Allogenic Mesenchymal Stem Cell Delivery in Myocardial Infarction in Dogs

Deborah C Vela 1, Guilherme V Silva 1, Joao AR Assad 1, Andre LS Sousa 1, Stephanie Coulter 1, Marlos R Fernandes 1, Emerson C Perin 1, James T Willerson 1, L Maximilian Buja 1
PMCID: PMC2628321  PMID: 19001635

Abstract

In this histological study, we assessed the role of mesenchymal stem cells (MSCs) in the healing process that takes place during the subacute phase of myocardial infarction in dogs. Seven days after occlusion of the left anterior descending coronary artery, adult mongrel dogs received 100 × 106 4′-6-diamidino-2-phenylindole (DAPI)-labeled allogenic bone marrow–derived MSCs by the transendocardial (TE, n=6) and intracoronary (IC, n=4) routes; control dogs (n=6) received no infusion. The dogs were euthanized at 21 days after occlusion. Hearts were excised and sliced from apex to base into four transverse sections, which were divided into nine segments. Paraffin sections from each segment were stained with hematoxylin and eosin, trichrome, picrosirius red, and antibodies against several extracellular matrix components. Frozen sections were immunostained for host cardiac phenotypical markers and analyzed by epifluorescence and deconvolution fluorescence microscopy (DFM). We found less unresolved necrotic myocardium and more extracellular matrix deposition in MSC-treated dogs than in controls 2 weeks after cell delivery. By DFM, no DAPI+ MSC nuclei were observed within native cardiac cells. MSCs delivered during the subacute phase of acute myocardial infarction positively affect healing, apparently by mechanisms other than differentiation into mature native cardiac cells. (J Histochem Cytochem 57:167–176, 2009)

Keywords: mesenchymal stem cells, myocardial scar, infarct healing, extracellular matrix

The wound healing process that occurs after acute myocardial infarction (AMI) is important because of its implications in left ventricular remodeling (Sun and Weber 2000). During the healing process, the extracellular collagen matrix (ECM) undergoes significant changes and plays an important role not only by providing a structural support, but also by modulating cellular and signaling processes in the proliferative phase of infarct healing (Lindsey et al. 2003). The sequence of events that take place shortly after an acute ischemic injury in dogs has been described in detail (Richard et al. 1995; Jugdutt 2002; Dewald et al. 2004; Dobaczewski et al. 2006).

Preliminary studies of transplantation of bone marrow mesenchymal stem cells (MSCs) have shown promising results for cardiac repair and have reported improved ventricular function (Barbash et al. 2003; Amado et al. 2005; Silva et al. 2005; Tang et al. 2006; Imanishi et al. 2007; Schuleri et al. 2008). Both intracoronary (IC) and transendocardial (TE) methods have been used to deliver stem cells in ischemic heart disease. The IC route of delivery has been most frequently used in clinical trials (Abdel-Latif et al. 2007). In a previous safety and feasibility study of both IC and TE delivery, we reported functional improvement and increased vascularization in dogs treated with MSCs after AMI (Perin et al. 2008).

However, the focus of preliminary studies has been on angiogenic responses, enhanced coronary blood flow, and the controversial topic of differentiation of MSCs into cardiomyoblasts. The role of MSCs in the postinfarct wound healing process and the development of myocardial scar have received less attention.

Thus, in this histological study, we examine the role of MSCs in the healing process when delivered by the IC and TE routes during the subacute phase of myocardial infarction in dogs.

Materials and Methods

To study the histological characteristics and extracellular matrix deposition within the infarcts of dogs treated with allogenic MSCs, we used histological tissue samples from a previous set of experiments performed by our group (Perin et al. 2008). The original study was reviewed and approved by The University of Texas Health Science Center at Houston's Animal Welfare Committee.

Model of Acute Ischemia in Dogs

Healthy male and female adult mongrel dogs (n=18) underwent left thoracotomy; anesthesia was induced with pentothal (17 mg/kg, intravenously) and maintained with isoflurane (1.5%–2.0%). Acute myocardial ischemia was produced by a transient 3-hr occlusion of the proximal left anterior descending coronary artery followed by reperfusion; the diagonal branch was ligated to decrease collateral flow to the infarct area. Seven days after the infarction, the dogs received 100 × 106 4′-6-diamidino-2-phenylindole (DAPI)- and chloromethylated 1,1′-dioctadecyl-3,3,3′,3′-tetramethylindocarbocyanine perchlorate analog–labeled allogenic MSCs by either IC delivery or 25 electromechanical mapping–guided TE injections targeted to the viable border zone of the infarction. Control dogs underwent occlusion and reperfusion but did not receive any IC infusions or TE injections. Sixteen dogs completed the study (TE = 6, IC = 4, control = 6) and were euthanized 2 weeks after MSC delivery. None of the animals received immunosuppression.

MSC Isolation, Expansion, and Labeling

The allogenic canine MSCs used in this study were isolated and prepared at Osiris Therapeutics (Baltimore, MD), as previously described (Pittenger et al. 1999). Briefly, a purified bone marrow MSC population was expanded in culture. Cells were harvested, labeled with the nuclear stain DAPI and the cross-linkable membrane dye CM-DiI (Molecular Probes; Eugene, OR), and frozen in cryocyte bags. Frozen cells were stored in the vapor phase of liquid nitrogen until implantation. Before injection, the cells were thoroughly washed and resuspended in a 5-ml volume of saline (20 × 106 MSC/ml). Cell viability at delivery was confirmed to be >90%.

Tissue Acquisition and Histology

After euthanasia, hearts were excised, weighed, and perfused; infarct size and bed-at-risk were measured. Hearts were sliced from apex to base into four transverse sections, which were cut into nine segments (labeled anterior, anterolateral, lateral, posterolateral, and posterior left ventricular free wall; anterior, mid, and posterior interventricular septum; and right ventricular free wall) and formalin fixed. A thin slice from each transverse section was cut into the same nine segments, each of which was placed in embedding medium for frozen tissue specimens and kept frozen at −80C. Histological sections from formalin-fixed and paraffin-embedded tissues were cut at 5 μm thickness and stained with hematoxylin and eosin (H&E), Masson's trichrome, picrosirius red, and various IHC stains. Fresh frozen tissues were immunostained (see below) for analysis by deconvolution fluorescence microscopy.

IHC

Paraffin-embedded sections were deparaffinized and hydrated with distilled water and stained for the following antibodies at the respective dilutions: laminin, 1:50 (#L9393; Sigma-Aldrich, St. Louis, MO); α-smooth muscle actin, 1:40,000 (#A2547; Sigma-Aldrich); fibronectin clone IST-9, 1:200 (#ab6328; Abcam, Cambridge, MA); anti-macrophage clone mac387, 1:200 (#MS-148-P; Lab Vision, Fremont, CA); CD3, 1:100 (#M7254; DakoCytomation, Carpinteria, CA); fibrinogen, 1:2000 (#A0080; DakoCytomation); vimentin, 1:800 (#0725; DakoCytomation); desmin, 1:1600 (#0760; DakoCytomation); and Ki-67, 1:250 (#M7240; DakoCytomation). Antibodies were pretreated with either citrate at 10 mM of citrate buffer (pH 6) or EDTA for 15 min in a microwave oven and 20 min in a resting solution. ApoTag (Millipore; Billerica, MA) was used as a marker for apoptosis.

The following antibodies and dilutions were used for antigen detection in fresh frozen sections: anti–α-sarcomeric actinin, 1:20,000 (#A7811; Sigma-Aldrich); α-smooth muscle actin, dystrophin 1 (#NCL-DYS1; Novacastra, Norwell, MA); von Willebrand factor VIII, 1:8000 (DakoCytomation); vimentin; and desmin. All antibodies were diluted with 0.02% BSA/PBS. All frozen specimens were cut on positively charged slides at 4 μm and allowed to air dry for 2 hr at room temperature. Sections were fixed in 4C acetone for 2 min, allowed to air dry at room temperature for 15 min, and placed in PBS (pH 7.5) for 5 min. Tissue sections were removed from PBS and placed in 1.5% normal horse serum (#S-2000; Vector, Burlingame, CA) for 10 min. The horse serum was gently tapped off, and the sections were incubated with the primary antibody for 1 hr at room temperature. The sections were rinsed with PBS for 5 min and placed in fluorescein solution for 10 min in a dark chamber at room temperature. For polyclonal antibody factor VIII, goat anti-rabbit fluorescein (#Fl-1000; Vector) was used. For monoclonal antibodies, smooth muscle actin, anti–α-sarcomeric actinin, and horse anti-mouse fluorescein (#Fl-2000; Vector) were used. The sections were thoroughly rinsed in PBS and aqueous mounted with Aquaperm (#484985; Thermo-Shandon, Waltham, MA).

Image Analysis

Formalin-fixed, paraffin-embedded sections were evaluated by quantitative morphometry, performed with Olympus MicroSuite software on an Olympus BX61 microscope (Olympus America; Melville, NY). For the quantification analysis of unresolved myocyte necrosis within the infarct, we measured all sections that contained infarcted tissue from all slices of the heart. These sections were stained with H&E and Masson's trichrome, digitally imaged, and traced with the above-mentioned software. Results from the determination of individual areas of unresolved infarct and their respective distance from the infarct border were expressed in square millimeters and millimeters, respectively. For quantification of ECM components, we analyzed sections from the equatorial slice of each heart that contained infarcted tissue. For collagen quantification, sections were stained with picrosirius red, imaged entirely, and subsequently analyzed by color threshold with Olympus Microsuite software. Laminin, fibronectin, and fibrinogen were quantified on sections IHC stained for their respective antibodies (see IHC). Three to five high power magnification (×20) random fields per section were analyzed (a total of 16 sections designated as regions of interest) by using predetermined color thresholds. The results obtained from each ECM component were considered representative of the infarct as a whole and were expressed as a percentage of the infarct.

Fluorescence Microscopy

Frozen sections immunostained against the above-mentioned antibodies were visualized on an Olympus BX61 microscope (Olympus America) and scanned with an Applied Precision DeltaVision scanning deconvolution fluorescence microscope (Issaquah, WA). Stacked image reconstructions were made, and image analysis was performed on a Silicon graphics workstation using SoftWoRx software (Applied Precision) (Davis et al. 2006).

Statistical Analyses

The data obtained from measurements and the quantification process mentioned in the section above were pooled for each group, and analyses were performed with appropriate software. All values are expressed as mean ± SD. Comparisons were performed using ANOVA. Comparisons between individual groups were performed using an unpaired Student's t-test. A value of p<0.05 was considered statistically significant.

Results

Infarct Size, Bed at Risk, and Unresolved Infarct

There were no significant differences in the size of the infarct or the bed-at-risk among the three groups (data not shown). Nevertheless, notable differences were seen in the amount of unresolved necrotic myocardium present within the infarcts. In this dog model, very large transmural infarcts were seen at 3 weeks after occlusion and reperfusion, and all dogs except three (one from each group) showed various quantities of unresolved necrotic myocardium within the healing area of the infarct. Quantification studies showed significantly less unresolved infarct per heart in the IC group than in the control group, but the difference between TE and IC groups was not significant [TE, 56.1 ± 53 mm2; IC, 32.16 ± 14 mm2; control, 125.6 ± 82 mm2; data not shown (IC vs control, p<0.05)]. In addition, the average size (area) of each individual foci of unresolved infarct was smaller in the IC group than in the control and TE groups [Figures 1A–1E; TE, 9.7 ± 17.2 mm2; IC, 3.1 ± 3.1 mm2; control, 16.7 ± 16.7 mm2 (IC vs control, p<0.01; IC vs TE, p<0.05)]. In contrast, the distance between any focus of unresolved necrotic infarct and the infarct border was noticeably smaller in the control group than in both the IC and TE MSC-treated groups [Figure 1F; control, 0.9 ± 0.3 mm; IC, 1.9 ± 1.1 mm; TE, 2.0 ± 1.2 mm (IC vs control and TE vs control, p<0.01); Figures 1A, 1B, and 1F].

Figure 1.

Figure 1

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Unresolved necrotic myocardium within healing infarcts in dogs. Trichrome stain of sections showing the border and center of an infarct in a dog that underwent intracoronary (IC) delivery of mesenchymal stem cells (MSCs) (A and C, respectively) compared with those of a control dog (B and D, respectively). Note the large confluent area of unresolved necrotic myocardium (outlined in black) in the control dog compared with the multiple small foci seen in the IC dog, in between which collagen has been deposited. (E) Average size (area in mm2) of individual foci of unresolved necrotic myocardium within healing infarcts in dogs. (F) Average distance (in mm) of unresolved necrotic myocardium from the infarct border. *p<0.01 vs control; **p<0.01 IC vs control; p<0.05 IC vs transendocardial (TE). Bar = 2 mm.

Extracellular Matrix

Collagen deposition within the infarct was greater in the infarct of MSC-treated dogs than in control dogs (Figures 2A–2C). Both the TE and IC groups had significantly more dense collagen content in their infarcts than did control dogs, as reflected by the quantification of picrosirius red staining. Comparison of the collagen content between TE and IC groups was not significant [Figure 2C; TE, 64.5 ± 15.3%; IC, 56.4 ± 12.4%; control, 40.2 ± 8.37% (TE vs control, p<0.01; IC vs control, p<0.05)].

Figure 2.

Figure 2

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Extracellular matrix components in healing infarcts in dogs 3 weeks after occlusion and reperfusion and 2 weeks after MSC delivery. Picrosirius red stain showing collagen deposition in sections corresponding to the infarct center of a dog treated with TE delivery of MSCs (A) and a control dog (B). Note the large area of unresolved necrotic myocardium (n) lacking collagen in the control dog compared with a more advanced collagen network in the MSC-treated dog. (C) Quantification of collagen content within the infarct. Laminin content at the infarct of a dog treated with IC delivery of MSCs (D) vs a control dog (E). Note the more intense staining in the granulation tissue and abundant neovascularization (arrows) of the IC dog, whereas the control dog presents a large necrotic zone (n) devoid of laminin, which corresponds to dead, non-phagocytosed cardiomyocytes, with small amounts of incipient granulation tissue beginning to form in the surroundings (asterisks). (F) Quantification of laminin within the infarct. Fibronectin staining within the infarcts of an IC dog (G) and a control dog (H). Significant portions of the infarct have been replaced by dense collagen (dc). The control dog, however, still shows intense staining, particularly at areas containing granulation tissue that surrounds the necrotic myocardium (n) at the infarct center. (I) Fibronectin quantification within the infarct. Positive staining for fibrinogen at the infarct border of an IC dog (J) and in the unresolved necrotic myocardium of a control dog (K). The nuclei of the numerous inflammatory cells that surround the necrotic myofibers appear as a purple rim (asterisks). (L) Quantification of fibrinogen within the infarct. *p<0.05 vs control; #p<0.01 vs control. Bars: A,B = 2 mm; D,E,G,H,J,K = 200 μm.

Laminin, a basement membrane component of cardiomyocytes and vascular cells, was detected in greater amounts in the infarcts of MSC-treated dogs than in controls (Figures 2D and 2E). No significant difference in laminin content between the TE and IC groups was seen [TE, 35.7 ± 8.9%; IC, 38.1 ± 6.1%; control, 22.5 ± 16.1% (IC and TE vs control, p<0.01); Figure 2F].

Fibronectin was observed mostly in areas of incipient neoangiogenesis and as part of the provisional matrix (Figures 2G–2I). Significantly more fibronectin was seen in the infarcts of both the TE and IC groups than in controls; however, no significant difference was observed between the TE and IC groups [TE, 19.1 ± 5.2%; IC, 26.4 ± 3.7%; control, 41.3 ± 0.8% (TE and IC vs control, p<0.01)].

Positive immunostaining for fibrinogen was seen in necrotic myofibers within the unresolved regions of infarct centers and at the infarct border zones (Figures 2J–2L). However, comparison of the fibrinogen content among the three groups did not reach statistical significance (TE, 60.4 ± 9.7%; IC, 73.6 ± 12.5%; control, 60.5 ± 18.9%).

Identification and Characterization of Injected MSCs

MSC survival and engraftment was identified by the presence of DAPI+ cells in frozen sections under fluorescence microscopy. Frozen sections were immunostained for antibodies against α-sarcomeric actinin, α-smooth muscle actin, dystrophin 1, von Willebrand factor VIII, vimentin, and desmin and screened with an Olympus BX61 microscope (Olympus America) for colocalization with mature phenotypic markers. Sections that stained positive with the above antibodies were scanned with an Applied Precision DeltaVision scanning deconvolution fluorescence microscope to confirm the findings. Stacked image reconstructions failed to unequivocally link DAPI with any of the common mature phenotypical markers tested, such as α smooth muscle actin, factor VIII, or α-sarcomeric actinin (Figures 3A and 3B). DAPI+ nuclei observed in clusters at injection sites were positive for vimentin (Figure 3C). In addition, paraffin-embedded sections containing demonstrable injection sites that were immunostained for adult phenotypic markers and observed under bright-field microscopy also showed positive staining for vimentin (Figures 4D and 4F).

Figure 3.

Figure 3

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Deconvolution fluorescence microscopy of MSCs delivered through the TE route in a dog after experimental infarction. (A) Host myocardium displaying 4′-6-diamidino-2-phenylindole (DAPI)+ nuclei. The delivered cells are seen in the interstitium, outside of native cells. Red, dystrophin outlining the host cardiomyocytes; green, factor VIII; blue, DAPI. (B) Pooling of MSCs along the tissue planes of the uninfarcted myocardium, near an injection site. Green, α-actinin; blue, DAPI. (C) Image of a cluster of DAPI-labeled MSCs corresponding to an injection site. Yellow, vimentin; blue, DAPI. Bars: A,C = 30 μm; B = 100 μm.

Figure 4.

Figure 4

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IHC panel of an MSC cluster (cl) corresponding to an injection site located in the normal myocardium near the infarct border (ib) in the left ventricular wall of TE dog. (A) Hematoxylin and eosin. Boxed area is magnified in B–I. (B) Trichrome stain. (C) Apoptosis marker. (D) Desmin. (E) Vimentin. (F) α-smooth muscle actin. (G) Ki-67. (H) CD3. (I) Macrophages. Bars: A = 2 mm; B–I = 200 μm.

Focal Response to MSC Delivery

Sections displaying DAPI+ cells under fluorescence microscopy were examined under bright-field microscopy in H&E and trichrome stains and with various IHC markers (Figure 4).

A mild infiltration of mononuclear inflammatory cells was seen only at sites in which cells had been injected at higher densities, such as sites where cells were found in clusters, in needle tracks, or at areas where the delivered cells had pooled along the tissue planes. IHC analysis showed that the infiltrating cells stained positive for the T-cell marker CD3 and for macrophages (Figures 4H and 4I, respectively). Some eosinophils were also observed in the larger clusters of injected MSCs. The injection sites also stained scarcely positive for apoptosis, and staining for the proliferation marker Ki-67 was negative (Figures 4C and 4G, respectively). Inflammation was not detected in sections that had diffuse or scattered DAPI+ cells in either the IC or TE groups.

Sites showing clusters or pooling of injected cells had minimal signs of focal or replacement fibrosis (Figure 4B) and stained strongly positive for vimentin, a feature congruent with their mesenchymal origin (Figure 4E). These same sites were negative for desmin (Figure 4D), with the exception of small vascular structures forming at the periphery of larger clusters. α-Smooth muscle actin (Figure 4F) stained mostly interspersed myofibroblasts.

Discussion

Effects of MSC Administration on Infarct Healing

Our study showed various features that indicate a hastened healing process in dogs that had been treated with MSCs shortly after myocardial infarction compared with control dogs. Although infarct sizes were similar among all groups, one of our most noticeable findings was the vast area of unresolved necrotic myocardium and hemorrhage seen within the infarcted areas of dogs that had not received MSCs 1 week after the ischemic event (Figures 1B and 1D). In contrast, the dogs that received MSCs had smaller foci of non-phagocytosed, necrotic myocyte remnants surrounded by collagen deposition (Figures 1A, 1C, 1E, and 2A). This finding was particularly significant in the IC group and may represent an advantage of IC delivery in regard to allowing delivered cells to reach the infarct center. A comparative study by Freyman et al. (2006) reported similar findings of IC diffusion efficacy compared with TE and IV routes in pigs, with the caveat of decreased blood flow during IC MSC delivery.

In addition, in our study, both the TE and IC groups showed broader areas of dense collagen deposition at the infarct borders than did control dogs, and any foci of unresolved infarct was restricted to the most central parts of the infarct (Figures 1A and 1F). In contrast, control animals had necrotic myocardium extending almost to the edges of the infarct (Figure 1B). This finding is important because it suggests that MSC administration may confer the nascent scar with the necessary mechanical characteristics to prevent or ameliorate negative remodeling (Holmes et al. 2005) and possibly prevent complications such as wall rupture in large transmural infarcts. The schematic in Figure 5 shows the observed healing pattern in MSC-treated and control dogs.

Figure 5.

Figure 5

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Schematic of healing patterns at 3 weeks after the onset of a transmural infarct in a model of infarction and reperfusion in dogs where A represents the findings in the hearts of control animals and B shows the healing observed in MSC-treated dogs.

During the normal healing of an infarct in dogs, collagen tends to increase over time as scar maturation ensues. Laminin is initially low because of cardiomyocyte death but becomes more prominent during the second week because of angiogenesis. Fibronectin content peaks by the end of the first week and decreases progressively thereafter. Fibrinogen immunoreactivity remains high throughout the first 2–3 weeks after infarction but decreases considerably after 4 weeks (Dobaczewski et al. 2006). Given the normal healing process, our findings of differences among the ECM components we studied indicate hastened healing in MSC-treated dogs compared with controls. Dogs in both IC and TE groups showed more collagen and laminin content in their infarcts and less fibronectin than in the infarcts of control dogs (Figure 2). The decreased amount of fibronectin present in MSC-treated dogs (Figures 2G and 2H) was caused primarily by the replacement of significant portions of the infarct by dense collagen. The large areas of necrotic, non-phagocytosed cardiomyocytes seen within the infarcts of control dogs contributed to their reduced laminin content (Figure 2E). The lack of a significant difference in fibrinogen content between treated and control dogs was mostly attributable to intense staining of necrotic myofibers in the infarcts of control dogs, combined with marked staining of the infarct borders in MSC-treated dogs (Figures 2J and 2K).

Focal Response to MSC Delivery

Allogenic MSCs have been reported to have a low inherent immunogenicity and favorable immunoregulatory properties (Le Blanc 2006; Rasmusson 2006; Nauta and Fibbe 2007). The immunosuppressive characteristics of MSCs have been studied in vitro, with potential extrapolation to in vivo applications (Bartholomew et al. 2002; Di Nicola et al. 2002; Tse et al. 2003; Aggarwal and Pittenger 2005). However, recent in vivo studies, in both small and large animal models, have shown evidence contradicting the immunoprivilege of MSCs in the allogenic (Eliopoulos et al. 2005; Nauta et al. 2006; Poncelet et al. 2007) and xenogeneic (Grinnemo et al. 2004) settings.

The complex interactions and mechanisms that govern the interaction of MSCs with host cells cannot be determined from our analysis; however, our histological examination showed mild to moderate macrophage and T-cell infiltration, mostly at the periphery of cell clusters and needle tracks from injection sites. Similar findings have been reported by two other groups (Poncelet et al. 2007; Hashemi et al. 2008). Importantly, we found no overt fibrosis at the cluster and needle track sites (Figure 4B), indicating that the injected MSCs produced more of a displacement of myocardial fibers without direct cytotoxicity or damage. Furthermore, we did not find signs of active proliferation of the host immune cells.

MSC Retention, Distribution Pattern, and Engraftment

The pattern of MSC distribution according to route of delivery has been previously described in greater detail (Perin et al. 2008). In this study, the pattern observed in staining for proliferation and apoptotic markers suggests that most injected cells found in the injection clusters are not proliferating (Figures 4C and 4G). Cell density is a critical element in vitro; each cell has an ideal level of confluency that is optimal to its growth, expansion, senescence, and death. We cannot determine whether the higher cell densities in MSC clusters produced by TE delivery affect the ability of the MSCs to proliferate in situ. Our lack of additional time points of study (earlier and later than 2 weeks after MSC delivery) precludes a definitive answer.

We used detailed three-dimensional fluorescence analysis of tissue and multiple markers for mature phenotypes of cells found in mature cardiac tissue. We found no unequivocal evidence of DAPI+ nuclei inside cardiomyocytes, cardiac vascular endothelium, or smooth muscle cells. Our findings showed that DAPI+ nuclei that appeared to be inside native cardiac cells when viewed by standard fluorescence microscopy were really “pseudonuclei” when examined by deconvoluted fluorescence microscopy. Other researchers have reported similar findings (Taylor et al. 2002).

Several studies have suggested that MSCs were capable of migrating through tissues (Barbash et al. 2003; Abbott et al. 2004; Kawada et al. 2004; Nagaya et al. 2004); however, our observations seem to indicate that a substantial portion of the cells delivered or deposited by the TE route remain at the site as a cluster. The dispersion pattern seen in many of the injected segments and previously described by Perin et al. (2008) most likely results from a combination of migration and of some of the cells reaching small vessels and being distributed by the circulation.

MSCs have been reported to incorporate into newly formed vessels and to most likely differentiate into vascular endothelium and smooth muscle (Tomita et al. 2002; Tang et al. 2006; Schuleri et al. 2008), thus contributing to improved coronary blood flow. In this analysis, the lack of definitive evidence for MSC differentiation and signs of minimal proliferation in situ, at 2 weeks after delivery, support the work of other researchers whose data suggest MSCs produce a paracrine effect, by which they may alter the tissue microenvironment and modulate biological activities, such as angiogenesis, wound repair, and the inflammatory response (Phinney et al. 2006; Dai et al. 2007; Feygin et al. 2007; Prockop 2007; Zhang et al. 2007; Chen et al. 2008). Either or both of these mechanisms may directly enhance neovascularization and facilitate cardiac healing after AMI.

Study Limitations

One limitation of this study was the unavailability of samples from different time points after myocardial infarction and MSC delivery. Having samples from these time periods would have helped assess whether proliferative activity occurs within injection sites at earlier times. In addition, a larger number of dogs within each group would have strengthened our conclusions.

In conclusion, our results suggest that MSCs may improve outcome by mechanisms other than differentiation into native cardiac cells. Delivery of MSCs to the infarct border 1 week after AMI may help fortify the nascent infarct scar by hastening healing and modulating deposition of extracellular matrix components. In doing so, MSCs may prevent the undesirable effects of negative remodeling of the ventricle wall. However, long-term follow-up is necessary to ensure that favorable healing and effective remodeling are maintained.

Acknowledgments

This work was funded by Ronald MacDonald Research Fund Grant 06RDM011.

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