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Keywords:

  • Pericytes;
  • Angiogenesis;
  • Immunomodulation;
  • Myocardial infarction;
  • Stem cell therapy

Abstract

  1. Top of page
  2. Abstract
  3. INTRODUCTION
  4. MATERIALS AND METHODS
  5. RESULTS
  6. DISCUSSION
  7. CONCLUSION
  8. Acknowledgements
  9. DISCLOSURE OF CONFLICTS OF INTEREST
  10. REFERENCES
  11. Supporting Information

Human microvascular pericytes (CD146+/34/45/56) contain multipotent precursors and repair/regenerate defective tissues, notably skeletal muscle. However, their ability to repair the ischemic heart remains unknown. We investigated the therapeutic potential of human pericytes, purified from skeletal muscle, for treating ischemic heart disease and mediating associated repair mechanisms in mice. Echocardiography revealed that pericyte transplantation attenuated left ventricular dilatation and significantly improved cardiac contractility, superior to CD56+ myogenic progenitor transplantation, in acutely infarcted mouse hearts. Pericyte treatment substantially reduced myocardial fibrosis and significantly diminished infiltration of host inflammatory cells at the infarct site. Hypoxic pericyte-conditioned medium suppressed murine fibroblast proliferation and inhibited macrophage proliferation in vitro. High expression by pericytes of immunoregulatory molecules, including interleukin-6, leukemia inhibitory factor, cyclooxygenase-2, and heme oxygenase-1, was sustained under hypoxia, except for monocyte chemotactic protein-1. Host angiogenesis was significantly increased. Pericytes supported microvascular structures in vivo and formed capillary-like networks with/without endothelial cells in three-dimensional cocultures. Under hypoxia, pericytes dramatically increased expression of vascular endothelial growth factor-A, platelet-derived growth factor-β, transforming growth factor-β1 and corresponding receptors while expression of basic fibroblast growth factor, hepatocyte growth factor, epidermal growth factor, and angiopoietin-1 was repressed. The capacity of pericytes to differentiate into and/or fuse with cardiac cells was revealed by green fluorescence protein labeling, although to a minor extent. In conclusion, intramyocardial transplantation of purified human pericytes promotes functional and structural recovery, attributable to multiple mechanisms involving paracrine effects and cellular interactions. STEM CELLS2013;31:305–316


INTRODUCTION

  1. Top of page
  2. Abstract
  3. INTRODUCTION
  4. MATERIALS AND METHODS
  5. RESULTS
  6. DISCUSSION
  7. CONCLUSION
  8. Acknowledgements
  9. DISCLOSURE OF CONFLICTS OF INTEREST
  10. REFERENCES
  11. Supporting Information

Coronary heart disease (CHD) is the leading cause of death in the United States, affecting 16.3 million people and accounting for one of every three deaths in 2007 [1]. Prolonged pathological interference with the coronary blood supply, such as atherosclerosis and thromboemboli, results in ischemic cardiomyopathy and/or myocardial infarction (MI) [2]. MI often leads to heart failure (HF) due to the limited capacity of the human heart to repair/regenerate its damaged myocardium [2, 3]. As an alternative to heart transplantation, stem/progenitor cell therapy (SCT) is deemed promising for restoration of cardiac function and prevention of progressive HF [3, 4]. In particular, human bone marrow precursor cells, endothelial progenitor cells, skeletal myoblasts, and endogenous cardiac progenitor cells have been intensively investigated with uneven success in preclinical and clinical trials [3–6]. Given the vascular origin of CHD pathology, stem/progenitor cells capable of reconstituting host vascular networks, in addition to other merits, will be ideal cell sources for SCT.

Microvascular pericytes (aka mural cells or Rouget cells) that tightly encircle capillaries and microvessels (arterioles and venules) and regulate microvascular physiology have recently been shown to contain precursor cells endowed with mesodermal differentiation potential [7]. Pericytes (CD146+/34/45/56) purified by cell sorting from human skeletal muscle, adipose, placenta, pancreas, and other organs repair and regenerate damaged/defective tissues [8–12] and represent the CD146-positive developmental origin of the heterogeneous mesenchymal stem/stromal cells (MSCs) [12–16]. Owing to their wide distribution in the microvasculature, pericytes are regarded as a promising and attractive source of precursor cells for regenerative medicine [17–19]. We hypothesize SCT with purified pericytes to be a suitable approach for the treatment of ischemic heart disease (IHD) [20].

Besides cardiomyogenesis, cardioprotective mechanisms, including antifibrosis, anti-inflammation, and neovascularization, play critical roles in SCT-mediated cardiac repair following ischemic insults [21–23]. SCT reduces myocardial fibrosis and induces favorable tissue remodeling in the ischemic heart, which in turn increases myocardial compliance/strength and prevents the progressive, pathological decline toward HF [24, 25]. This antifibrotic feature was attributed in part to increased collagen degradation by matrix metalloproteinases (MMPs) and inhibition of fibroblast activation, possibly through a paracrine mechanism [26, 27]. Additionally, the immunosuppressive/anti-inflammatory capacity of MSCs through secretion of immunoregulatory molecules has recently attracted clinical attention in organ transplantation and immune regulation [28, 29]. Whether pericytes possess similar antifibrotic and immunoregulatory capacities within the ischemic microenvironment remains to be addressed.

To relieve the underlying cause of IHD, SCT-based approaches toward myocardial revascularization have been extensively pursued [30, 31]. Due to their native vascular localization and secretion of trophic factors that are associated with tissue repair and vascular growth/remodeling, pericytes may restore injured vascular networks more efficiently [18]. Proangiogenic signaling molecules released by stem/progenitor cells stimulate neovascularization in ischemic tissues [31]. Additionally, cell–cell interaction between vascular mural cells and endothelial cells was lately suggested to play essential roles in blood vessel remodeling and maturation [32, 33]. Whether pericytes mediate revascularization of the ischemic myocardium through any of these two mechanisms has yet to be tested.

In this study, we investigated the therapeutic potential of purified human skeletal muscle pericytes in IHD, using an acute MI model in immunodeficient mice. Transplantation of pericytes not only reversed cardiac dilatation but also improved cardiac contractility. Major repair mechanisms were investigated, including reduction of fibrosis, inhibition of chronic inflammation, promotion of angiogenesis, and regeneration of the myocardium. We further describe putative mediators used by pericytes. Green fluorescence protein (GFP) labeling was used to track perivascular homing and lineage fate of transplanted pericytes. Our results demonstrate that the overall benefit of pericyte treatment is collectively attributed to multiple cardioprotective mechanisms that involve paracrine and direct cell–cell interactions.

MATERIALS AND METHODS

  1. Top of page
  2. Abstract
  3. INTRODUCTION
  4. MATERIALS AND METHODS
  5. RESULTS
  6. DISCUSSION
  7. CONCLUSION
  8. Acknowledgements
  9. DISCLOSURE OF CONFLICTS OF INTEREST
  10. REFERENCES
  11. Supporting Information

Human Tissue Biopsies and Cell Isolation

In total, three independent human skeletal muscle specimens (one adult and two fetal) were used for cell isolation. The procurement of adult muscle biopsies from the National Disease Research Interchange was approved by the Institutional Review Board (IRB) at the University of Pittsburgh. Muscle biopsies (male subject, 57 years old) were preserved in Dulbecco's modified Eagle's medium (DMEM) containing 1% antibiotics and transported to the laboratory on ice. Human fetal tissues (21 and 23 weeks of gestation) were obtained following voluntary pregnancy interruptions performed at Magee Womens Hospital, Pittsburgh, in compliance with IRB protocol 0506176. Informed consents for the use of fetal tissues were obtained from all patients. Cells were mechanically and enzymatically dissociated from muscle biopsies following the reported protocol [12]. Details are described in Supporting Information.

Fluorescence-Activated Cell Sorting and Flow Cytometry Analysis

Fluorescence-activated cell sorting (FACS) and flow cytometry were used to purify microvascular pericytes (CD146+/34/45/56) and examine cell lineage marker expression, respectively, as we previously reported [12]. Details are documented in Supporting Information.

Cell Culture and Cell Labeling

Sorted pericytes were expanded in reported culture conditions [12]. Single donor-derived human umbilical cord vein endothelial cells (HUVECs, Lonza) were cultured in endothelial cell growth medium-2 (EGM-2, Lonza, Allendale, NJ, http://www.lonza.com). Cultured pericytes were labeled with GFP following a published protocol [12], using a lentivirus-based Cytomegalovirus (CMV)-driven eGFP-expression vector. For short-term experiments, cells were labeled with cell membrane dyes, PKH26 (red) and PKH67 (green) (both from Sigma-Aldrich, St. Louis, MO, http://www.sigmaaldrich.com), and used immediately without further expansion.

Cell Transplantation in an Acute MI Model

The Institutional Animal Care and Use Committee at Children's Hospital of Pittsburgh and University of Pittsburgh approved the animal usage and surgical procedures (Protocol#37-04, 55-07, 0901681A-5). A total of 78 male : non-obese diabetic/severe combined immune deficiency (NOD/SCID) mice (Jackson Laboratory, Bar Harbor, ME, http://www.jax.org) were used. MI induction (by ligation of left anterior descending coronary artery) and intramyocardial cell injection (3 × 105 cells per heart) were performed by a blinded surgeon as previously reported [34]. Control mice received injections of 30 μl phosphate-buffered saline (PBS).

Evaluation of Cardiac Function by Echocardiography

Mice were anesthetized with isoflurane, and transthoracic echocardiography was performed by a blinded investigator repeatedly before and after surgery (at 2 and 8 weeks), using a high resolution ultrasound system (Vevo 770, Visual Sonics, Inc., Toronto, Ontario, Canada, http://www.visualsonics.com), as described previously [34]. Mice which died prior to 8 weeks postinjection were excluded. Echocardiographic measurements are listed in Supporting Information.

Histological and Immunohistochemical Analyses

At 1, 2, and 8 weeks after surgery, hearts were harvested and processed as previously described [34]. Cryosections at 6–8 μm thickness were used for histological and immunohistochemical analyses following published protocols [34]. Anti-GFP immunofluorescent staining was performed on 4% paraformaldehyde-fixed sections. Donor cell engraftment and perivascular homing were quantified on serial sections stained with anti-GFP antibody (Abcam, Cambridge, U.K., http://www.abcam.com) and dual-stained with anti-GFP/anti-mouse CD31 (BD Biosciences, San Diego, CA, http://www.bdbiosciences.com) antibodies, respectively, using Image J. The engraftment ratio was defined as the extrapolated total number of engrafted GFP-positive cells to the initial 3 × 105 cells injected. Perivascular homing ratio was defined as the extrapolated number of GFP-positive cells juxtaposing CD31-positive mouse endothelial cells to the extrapolated total number of engrafted cells. Using Masson's trichrome staining (IMEB, Inc., San Marcos, CA, http://www.imebinc.com), fibrotic area fraction and infarct wall thickness were estimated from six randomly selected sections at comparable infarct levels per heart as previously described [34]. Quantification of host angiogenesis and chronic inflammation was computed from 6–10 randomly selected images taken from the designated area in sections stained with anti-mouse CD31 and anti-mouse CD68 (Abcam) at the midinfarct level, respectively. Experimental details are documented in Supporting Information.

Hypoxia Assay and Enzyme-Linked Immunosorbent Assay

To simulate the lower oxygen tension at the tissue level, physiologically or pathologically, pericytes were cultured under 2.5% O2 hypoxic conditions (with 5% CO2 and 92.5% N2) as formerly described [34]. Cells were washed twice before defined, serum-free DMEM medium was added upon the transition to low O2 conditions. Culture supernatant and cell lysates were collected 24 hours later. Cells cultured under 21% O2 (normoxia) served as controls. The secretion of vascular endothelial growth factor (VEGF), angiopoietin-1 (Ang-1), Ang-2, and transforming growth factor (TGF)-β1 in the culture supernatant was measured by respective enzyme-linked immunosorbent assay (ELISA) with human VEGF (Invitrogen, Carlsbad, CA, http://www.invitrogen.com), Ang-1, Ang-2, and TGF-β1 (all from R&D Systems, Minneapolis, MN, http://www.rndsystems.com) ELISA Kits.

Real-Time Quantitative Polymerase Chain Reaction and Semi-Quantitative Real Time Polymerase Chain Reaction

Real-time quantitative polymerase chain reaction (rt-qPCR) was performed as previously reported [9]. Total RNA (n = 6) was extracted for cDNA synthesis. The quantitative analyses were performed in the core facility at the University of Pittsburgh. All data are presented as expression level normalized to human cyclophilin (in arbitrary fluorescence units). For semi-quantitative RT-PCR (sqRT-PCR), total RNA (n = 3) was extracted using RNeasy plus-mini-kits (Qiagen, Hilden, Germany, http://www1.qiagen.com). From each sample, 500 ng of total RNA was reverse transcribed, followed by PCR. The intensity of the product bands was calculated using Quantify-One software and normalized to β-actin. The primer sequences are listed in Supporting Information Table T1 and T2.

In Vitro Vascular Network Formation

Cell culture/coculture experiments using two-dimensional (2D) and three-dimensional (3D) Matrigel systems were performed to observe the capillary-like network formation. For 2D culture, 5 × 104 pericytes or HUVECs were seeded onto Matrigel-coated well and incubated for 24 hours. For 3D culture, 25 × 104 pericytes or HUVECs were resuspended in EGM-2 medium and mixed with Matrigel in a 3:1 ratio. The Matrigel plug was subsequently incubated for 72 hours. Equal numbers of dye-labeled pericytes and HUVECs were well mixed in 2D or 3D coculture for 72 hours to observe pericyte–HUVEC interactions.

Measurement of Cell Proliferation

Murine RAW264.7 monocyte/macrophage cells American Type Culture Collection (ATCC), Manassas, VA, USA, http://www.atcc.org/ or primary murine skeletal myoblasts, muscle fibroblasts, and cardiac fibroblasts were cultured with normoxic or hypoxic pericyte-culture conditioned medium, or with serum-free control medium, for 72 hours. Cell proliferation was measured by the absorbance at 490 nm after incubation with CellTiter Proliferation Assay Reagent (Promega, Madison, WI, http://www.promega.com) for 3 hours. Experiments were performed in quadruplicates and repeated three times independently.

Statistical Analysis

All measured data are presented as mean ± SE. Kaplan-Meier survival curve estimation with log-rank test was performed to compare the animal survival rate between treatment groups. Statistical differences were analyzed by Student's t test (two groups), one-way ANOVA (multiple groups), or two-way repeated ANOVA (repeated echocardiographic measurements) with 95% confidence interval. Statistical significance was set at p < .05. Student-Newman-Keuls multiple comparison test (or Bonferroni test if specified) was performed for ANOVA post hoc analysis. Statistical analyses were performed with SigmaStat 3.5 (Systat Software, Inc., Chicago, IL, http://www.systat.com) and SPSS19 (IBM Corp., Armonk, NY, http://www.ibm.com) statistics software.

RESULTS

  1. Top of page
  2. Abstract
  3. INTRODUCTION
  4. MATERIALS AND METHODS
  5. RESULTS
  6. DISCUSSION
  7. CONCLUSION
  8. Acknowledgements
  9. DISCLOSURE OF CONFLICTS OF INTEREST
  10. REFERENCES
  11. Supporting Information

Isolation and Transplantation of Human Pericytes

As reported in our previous studies [12], FACS was used to purify human microvascular pericytes (CD146+/34/45/56) to homogeneity from skeletal muscle biopsies of three donors (one adult and two fetal, designated as AP, FP1, and FP2, respectively), by their differential expression of cell lineage markers, including CD34 (endothelial/stem cells), CD45 (hematolymphoid cells), CD56 (myogenic cells), and CD146 (pericyte/endothelial cells) (Supporting Information Fig. S1). No phenotypical difference between adult and fetal pericytes was noted, consistent with our previous observations [12]. After in vitro expansion (25–35 cell doublings) and prior to transplantation, in all three pericyte populations, we have observed no alteration to their distinctive morphology as well as classic antigenic profile, including robust expression of CD146, alkaline phosphatase, and typical MSC markers: CD44, CD73, CD90, CD105 with the absence of CD34, CD45, and CD56 (Supporting Information Fig. S2A, S2B). Additionally, cell labeling (in subsequent experiments) did not alter pericyte phenotype (data not shown). Cells (3.0 × 105 cells per heart) resuspended in 30 μl PBS were injected into the acutely infarcted myocardium of immunodeficient mice. The control group received injections of 30 μl PBS following the induction of MI.

Human Pericyte Transplantation Improves Cardiac Function

The survival of animals receiving pericyte treatment or PBS injection was monitored over the course of 8 weeks (Kaplan-Meier survival curve, log-rank test p = .529, Fig. 1A). Cardiac function was assessed by echocardiography performed repeatedly before (healthy) and at 2 and 8 weeks after surgery (Supporting Information Fig. S3). Ischemic hearts injected with either of the three pericyte populations (n = 8 per donor) had significantly smaller left ventricular (LV) chamber size, as measured by LV end-diastolic area (LVEDA, Fig. 1B) and end-systolic area (LVESA, Fig. 1C), than the control group (n = 8) (all p < .05), suggesting the reversal of progressive heart dilatation. Moreover, all pericyte-transplantation groups displayed significantly better LV contraction, evaluated by LV fraction shortening (LVFS, Fig. 1D), LV fractional area change (LVFAC, Fig. 1E), and LV ejection fraction (LVEF, Fig. 1F), than the control group (all p < .05). Collectively, when compared to vehicle treatment, pericyte treatment not only resulted in considerably smaller LV chamber dimension (p < .001, two-way repeated ANOVA) but also notably improved LV contractility (p < .001, two-way repeated ANOVA) for up to 2 months. Dimensional and functional echocardiographic parameters are documented in Supporting Information Table T3. In a separate experiment, we compared cardiac function of acutely infarcted hearts injected with either APs or CD56+ myogenic progenitors, sorted from a single adult muscle biopsy. The echocardiographic results at 2 weeks postinfarction showed that pericyte-treated hearts (n = 6) have significantly better LV function than CD56+ progenitor-treated ones (n = 6) in five parameters examined, including LVEDA (p = .004), LVESA (p = .002), LVFS (p = .003), LVFAC (p = .003), and LVEF (p ≤ .001) (Supporting Information Fig. S4).

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Figure 1. Survival rate and cardiac functional assessment. (A): Cumulative survival rate of the animals over 8 weeks after surgery (Kaplan-Meier survival curve, log-rank test p = .529). Echocardiographic analyses revealed a significant reduction of left ventricular (LV) dilatation by transplantation of all three pericyte populations (AP, FP1, and FP2), as shown by the smaller LV area in end-diastole (B) and end-systole (C) of hearts at both time points. Injection of pericytes also resulted in substantial improvement in LV contractility, as indicated by greater fractional shortening (D), fractional area change (E), and ejection fraction (F), at both time points. (, p ≤ .001; §, p ≤ .005; #, p ≤ .01; *, p ≤ .05; vs. PBS control group at each time point). Abbreviation: PBS, phosphate-buffered saline.

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Transplantation of Pericytes Reduces Cardiac Fibrosis

To understand the influence of pericyte treatment on cardiac fibrosis, we evaluated scar tissue formation using Masson's trichrome histological staining. At 2 weeks postinfarction, pericyte-treated hearts displayed less collagen deposition (stained in blue/purple) at the ischemic area (Fig. 2A). Estimation of the total fibrotic tissue ratio unveiled a 45.3% reduction of cardiac fibrosis in the pericyte-injected myocardium (n = 5, 22.03% ± 1.81%) when comparing to saline-injected controls (n = 5, 40.28% ± 2.15%) (Fig. 2B, p ≤ .001), suggesting the antifibrotic efficacy of pericytes. Measurement of LV wall thickness at the center of the infarct indicated no significant difference (p > .05) between the pericyte group (0.255 ± 0.026 mm) and the PBS group (0.202 ± 0.040 mm), suggesting that pericytes had limited beneficial effects to reduce transmural infarct thinning following MI (Fig. 2C).

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Figure 2. Attenuation of myocardial fibrosis by pericyte treatment. (A): Masson's trichrome-stained transverse sections of hearts injected with pericytes or PBS (collagen in blue/purple, cardiac muscle in red; scale bar = 1 mm). (B): The fibrotic area fraction was dramatically decreased in pericyte-injected hearts (p ≤ .001). (C): Pericyte group had no significant increase in the infarct wall thickness. (D): When culturing with hypoxic pericyte-conditioned medium (P-CM), the proliferation of murine cardiac fibroblasts was significantly reduced (, p = .019, vs. normoxic P-CM) while muscle fibroblast proliferation exhibited the same pattern (*, p < .001). Normoxic P-CM had a pro-proliferative effect over control medium in muscle fibroblasts but not in cardiac fibroblasts (#, p < .05). Skeletal myoblast proliferation was not significantly affected by either of the P-CMs. (E): Expression of MMP-2 in cultured pericytes was higher than that in skeletal muscle lysates. Conversely, MMP-9 expression in pericytes was nearly 10 times less (logarithmic scale of 10 in arbitrary fluorescence units). (F): Expression of both MMP-2 and -9 in pericytes did not change significantly under hypoxia (p > .05, logarithmic scale of 10 in arbitrary fluorescence units). (G): Immunohistochemistry revealed MMP-2 expression (red arrows) by some of the GFP-labeled donor pericytes (green arrows) within the infarct area at 2 weeks postinfarction (a) merge (b) anti-GFP in green (c) MMP-2 in red (d) DAPI nuclei staining in blue (scale bar = 20 μm). Abbreviations: DAPI, 4′,6-diamidino-2-phenylindole (DAPI) ; GFP, green fluorescent protein; MMP, matrix metalloproteinase; PBS, phosphate-buffered saline.

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Paracrine Antifibrotic Effects of Pericytes Under Hypoxia

Oxygen tension within tissues, physiologically or pathologically, is considerably lower than the ambient oxygen concentration in vitro. To elucidate the mechanism involved in pericyte-mediated reduction of fibrosis, we mimicked, at least in part, the hostile hypoxic microenvironment that donor cells encounter within the ischemic myocardium by culturing pericytes under 2.5% oxygen for 24 hours in defined, serum-free medium. Pericytes cultured under 21% oxygen (normoxia) served as controls. We then performed a cell proliferation assay using primary murine skeletal muscle and cardiac fibroblasts as well as skeletal myoblasts cultured with pericyte-conditioned medium (P-CM). Cardiac fibroblast proliferation was significantly reduced when cultured in hypoxic P-CM, compared to normoxic P-CM (p = .019) (Fig. 2D). Muscle fibroblast proliferation exhibited the same inhibitory pattern (p < .001, hypoxic vs. normoxic P-CM) with normoxic P-CM showing a pro-proliferative effect over control serum-free medium (p < .05) (Fig. 2D). Neither of the two P-CMs had significant influence over skeletal myoblast proliferation (p = .76). One-way ANOVA with Bonferroni multiple comparisons was performed for statistical analysis. This suggests a paracrine antifibrotic effect by pericytes in hypoxia. We further proposed a fibrolytic role of pericyte-derived MMPs and examined gene expression of MMP-2 and MMP-9 by real-time qPCR. Cultured pericytes expressed more MMP-2 but nearly 10 times less MMP-9 than total skeletal muscle lysates (tissue origin control) (Fig. 2E). We then explored MMP expression in hypoxia-cultured pericytes and demonstrated that MMP-2 expression in pericytes was well sustained under 2.5% oxygen, compared to normoxic culture (Fig. 2F, p > .05), while MMP-9 expression remained extremely low without significant change (Fig. 2F, p > .05). Immunohistochemical study revealed that some of the transplanted pericytes within the infarct region expressed MMP-2 (Fig. 2G, a–d).

Transplantation of Pericytes Inhibits Chronic Inflammation

Histological analysis of pericyte- and PBS-injected hearts after hematoxylin and eosin staining indicated an increased focal infiltration of inflammatory cells (cluster of cells with dark blue-stained nuclei) within the infarct region in the latter (Fig. 3A). To more precisely evaluate the immunomodulatory effect of pericyte transplantation, we detected host CD68-positive monocytes/macrophages by immunohistochemistry. Pericyte-injected hearts exhibited diminished infiltration of host phagocytic cells within the infarct region at 2 weeks postinfarction (Fig. 3B). Districts of the myocardium unaffected by the ischemic insult (posterior and septal walls) contained few CD68-positive cells in either group, similar to healthy hearts (Fig. 3C). Quantitatively, injection of pericytes (n = 5) resulted in a 34% reduction in infiltration of CD68-positive cells at 2 weeks postinfarction when compared to PBS controls (n = 5) (Fig. 3D, p < .001).

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Figure 3. Reduction of host phagocytic cell infiltration by pericyte transplantation. (A): Hematoxylin and eosin staining revealed a greater focal infiltration of leukocytes (dark blue-stained nuclei) within the infarct region in PBS-injected controls at 2 weeks postinfarction (scale bar = 100 μm). (B): Anti-mouse CD68 immunostaining showed that the infarct region of pericyte-injected hearts contains less host phagocytic cells (scale bar = 50 μm). (C): Host CD68-positive cells were locally attracted to the infarct region but not to the unaffected myocardium (posterior ventricular wall) in both groups (scale bars = 50 μm). (D): Host monocytes/macrophage infiltration at the infarct site was significantly reduced (p < .001). (E): The proliferation of murine macrophages was significantly inhibited when culturing with pericyte-conditioned media (*, p = .018; #, p < .001, vs. control medium), an effect more prominent with hypoxic pericyte-conditioned medium (, p = .002, hypoxia vs. normoxia). (F): Cultured pericytes exhibited sustained, high expression of genes regulating the inflammatory responses, even under 2.5% O2. Little expression of IL-1α and no expression of IL-4, IL-10, iNOS, 2,3-IDO, TNF-α, and IFNγ were detected. (G): No statistically significant difference in expression of genes of immunoregulatory molecules between normoxic- and hypoxic-cultured pericytes except MCP-1, which notably decreased in hypoxic cultures (semiquantitative real time polymerase chain reaction analysis, p = .027). Abbreviations: COX-2, cyclooxygenase-2; HMOX-1, heme oxygenase-1; HIF-1α, hypoxia-inducible factor-1α; H, hypoxia) IL-6, interleukin-6; iNOS, inducible nitric oxide synthase; 2,3-IDO, indoleamine 2,3-dioxygenase; IFNγ, interferon-γ; LIF, leukemia inhibitory factor; MCP-1, monocyte chemotactic protein-1; N, normoxia; PBS, phosphate-buffered saline.

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Paracrine Immunomodulation by Pericytes

To understand the underlying mechanism of pericyte-induced inhibition of phagocytic cell infiltration, we analyzed the proliferation of murine macrophages cultured with P-CM. Murine macrophage proliferation was significantly inhibited when culturing with both normoxic (p = .018) and hypoxic (p < .001) P-CM, compared to control medium (Fig. 3E). Furthermore, hypoxic P-CM exhibited a more prominent immunomodulatory effect than the normoxic counterpart (Fig. 3E, p = .002). We then investigated by sqRT-PCR the differential expression of genes encoding immunoregulatory molecules that are potentially accountable for this paracrine immunomodulation by pericytes. Under either normoxia or hypoxia, pericytes indeed expressed a considerable array of anti-inflammatory cytokines: interleukin-6 (IL-6), leukemia inhibitory factor (LIF), cyclooxygenase-2 (COX-2/prostaglandin endoperoxide synthase-2), and heme oxygenase-1 (HMOX-1) (Fig. 3F). Similarly, monocyte chemotactic protein-1 (MCP-1) and hypoxia-inducible factor-1α (HIF-1α) were highly expressed by pericytes (Fig. 3F). Conversely, we detected very low to no expression of proinflammatory cytokines including IL-1α, tumor necrosis factor-α (TNF-α), and interferon-γ (IFNγ) (Fig. 3F). No expression of IL-4, IL-10, inducible nitric oxide synthase (iNOS), and indoleamine 2,3-dioxygenase (2,3-IDO) was observed (Fig. 3F). Quantitatively, there was no significant alteration of expression under hypoxia of immunoregulatory genes investigated, except MCP-1, whose expression was notably decreased in hypoxia-cultured pericytes (Fig. 3G, all p > .05; MCP-1, p = .027).

Transplanted Pericytes Promote Host Angiogenesis

We examined whether intramyocardial transplantation of pericytes restores the host vascular network postinfarction. Capillaries in the peri-infarct areas (Fig. 4A) and within the infarct region (Fig. 4B) were revealed by anti-mouse CD31 (platelet endothelial cell adhesion molecule-1) immunofluorescent staining and subsequently quantified. Capillary structure density in the peri-infarct areas of pericyte-injected hearts (n = 5) was increased by 45.4% when compared to PBS-injected controls (n = 5) (Fig. 4C, p = .01). Higher microvascular density was also observed within the infarct region, with 34.8% more capillaries in the pericyte-treated hearts (Fig. 4C, p = .002). Detection of the proliferating host endothelial cells (ECs) by Ki-67, a cell proliferation marker, and CD31 showed that pericyte-injected hearts (n = 3) had significantly more proliferating ECs than PBS-injected controls (n = 3) in both the infarct region (p = .034) and peri-infarct areas (p = .025) (Supporting Information Fig. S5A-S5C). These findings suggest that transplanted pericytes promote host angiogenesis not only in the peri-infarct areas, where blood vessels were generally better preserved after the ischemic injury, but also within the blood vessel-deprived infarct region.

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Figure 4. Promotion of host angiogenesis by pericyte treatment. Representative images of anti-mouse CD31 immunostaining (A) in the peri-infarct area and (B) within the infarct region of hearts injected with pericytes or PBS (scale bar = 50 μm). (C): Pericyte-treated hearts displayed significantly higher capillary densities in the peri-infarct area (p < .05) and within the infarct region (p < .001). Abbreviation: PBS, phosphate-buffered saline.

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Pericytes Support Microvascular Structures

To demonstrate that pericytes benefit host vascular networks through their support of microvascular structures, we developed 2D and 3D Matrigel cultures/cocultures using pericytes and HUVECs. HUVECs seeded onto Matrigel-coated wells formed typical capillary-like networks after 24 hours (Fig. 5A). Pericytes, however, formed similar structures within 6–12 hours of seeding (Fig. 5B). To illustrate the reciprocal influence between pericytes and endothelial cells (ECs), dye-labeled pericytes (PKH67, green) and HUVECs (PKH26, red) were mixed and cocultured in 2D Matrigel, which resulted in the formation within 6–12 hours of capillary-like structures that included both cell types (Fig. 5C). Pericytes (green) were observed to collocate with HUVECs (red) in the co-formed three-dimensional structures after incubation for 24 hours (Fig. 5C, inset). Additionally, HUVECs (red) appeared to morphologically align with pericytes (green) (Fig. 5D). To further unveil the vascular supportive role of pericytes, an in vitro 3D Matrigel system designed to simulate native capillary formation was used. HUVECs evenly distributed within the 3D Matrigel plug were unable to form organized structures after 72 hours (Fig. 5E). To the contrary, pericytes started to form capillary-like networks 24 hours after gel casting, with structural remodeling over time (Fig. 5F). The dynamic interaction between pericytes and ECs was best depicted by encapsulating dye-labeled pericytes (green) and HUVECs (red) in a 3D Matrigel plug. Together these two types of cells formed capillary-like structures after incubation for 72 hours (Fig. 5G) with pericytes surrounding HUVECs (Fig. 5H). These data suggest that pericytes retained vascular cell features and formed structures supportive of microvascular networks even after purification and long-term culture, while pericyte–EC association may play a role in the pericyte-facilitated angiogenic process.

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Figure 5. Pericytes support microvascular structures. (A): While HUVECs seeded onto Matrigel-coated wells formed typical capillary-like networks after 24 hours, (B) pericytes formed similar structures within 6–12 hours (scale bar = 1 mm). (C): When cocultured on Matrigel, dye-labeled pericytes (green) and HUVECs (red) coformed capillary-like networks within 6–12 hours, (C, inset) with collocations of pericytes and HUVECs in three-dimensional structures formed 24 hours after seeding (scale bars: main = 200 μm; inset = 100 μm). (D): HUVECs (red) appear to line and spread out on top of the pericyte-formed structures (green) (scale bar = 100 μm). (E): To simulate native capillary formation, HUVECs were evenly encapsulated into 3D Matrigel plug for 72 hours but unable to form any organized structure (scale bars = 1 mm). (F): Pericytes instead formed capillary-like networks in Matrigel plug with structural organization and maturation over time (scale bar = 1 mm). (G): When dye-labeled pericytes (green) and HUVECs (red) were cocasted into the 3D-gel plug, the two types of cells formed microvessel-like networks within 72 hours, (H) with pericytes surrounding HUVECs (scale bars: (G) = 200 μm; (H) = 50 μm). Abbreviation: HUVEC, human umbilical cord vein endothelial cell.

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Differential Expression of Proangiogenic Factors and Associated Receptors by Pericytes Under Hypoxia

The paracrine angiogenic potential of pericytes in the ischemic heart was investigated using the simulated hypoxic environment in vitro. Expression of genes encoding proangiogenic factors and corresponding receptors was assessed by real-time qPCR. VEGF-A, platelet-derived growth factor-β (PDGF-β), and transforming growth factor (TGF)-β1 were notably upregulated by 307% (p ≤ .001), 437% (p = .067), and 178% (p = .037), respectively, in pericytes cultured under hypoxic conditions (Fig. 6A). Expression of other proangiogenic factors, including basic fibroblast growth factor (bFGF), hepatocyte growth factor (HGF), and epidermal growth factor (EGF), was downregulated to 44% (p < .05), 23% (p ≤ .001), and 60% (p > .05) of their expression levels in normoxia (Fig. 6A). On the other hand, VEGF receptor-1 (VEGFR-1/Flt-1) and -2 (VEGFR-2/KDR/Flk-1) were substantially upregulated by 458% (p = .004) and 572% (p ≤ .001), respectively, under 2.5% oxygen (Fig. 6B). PDGF receptor-β (PDGF-Rβ) expression was not significantly changed (161%, p > .05) (Fig. 6B). VEGF secretion by pericytes, measured by ELISA, significantly increased over threefold (p ≤ .001) under hypoxic culture conditions while Ang-1 secretion reduced by 35% (p > .05) (Fig. 6C). Very little secretion of Ang-2 by pericytes was detected under both conditions (p > .05) (Fig. 6C). Additionally, TGF-β1 secretion increased by 30.1% under hypoxia (p = .028) (Fig. 6C), consistent with its upregulated gene expression. The expression of human VEGF165 by engrafted pericytes within the infarct region was confirmed by immunohistochemistry (Fig. 6D, a–c).

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Figure 6. Expression of proangiogenic factors and associated receptors under hypoxia. (A): Pericytes dramatically upregulated VEGF-A, PDGF-β, TGF-β1 gene expression under hypoxic conditions (2.5% O2) while expression of other proangiogenic factors, including bFGF, HGF, and EGF were distinctively repressed. (B): Simultaneously, VEGFR-1 (Flt-1) and -2 (Flk-1) were substantially upregulated, and PDGF-Rβ expression was moderately increased. All expression levels are normalized to human cyclophilin and presented in arbitrary fluorescence units on an expanded logarithmic scale (#, p < .05, *, p ≤ .001; †, p < .01, hypoxia vs. normoxia). (C): Significantly increased secretion of VEGF (p ≤ .001) and TGF-β1 (p = .028) by pericytes under hypoxic culture conditions was detected by ELISA while secretion of Ang-1 was reduced by 35% (p > .05). Very little secretion of Ang-2 was detected under both conditions (p > .05). (D): Immunohistochemistry revealed human VEGF165 expression by GFP-labeled donor pericytes within the infarct area at 2 weeks postinfarction (a) merge (b) hVEGF165 in red (c) anti-GFP in green (scale bar = 50 μm). Abbreviations: Ang-1, angiopoietin-1; bFGF, basic fibroblast growth factor; EGF, epidermal growth factor; GFP, green fluorescent protein; HGF, hepatocyte growth factor; PDGF, platelet-derived growth factor; PDGFR, PDGF receptor; TGF, transforming growth factor; VEGF, vascular endothelial growth factor.

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Transplanted Pericytes Home to Perivascular Locations

It is not known whether purified pericytes home back to perivascular areas in vivo. To reveal their engraftment and homing pattern, cultured pericytes were transduced with a GFP reporter gene at near 95% efficiency (Fig. 7A) and injected (3.0 × 105 cells) into acutely infarcted hearts. GFP-labeled pericytes engrafted throughout the ventricular myocardium (Fig. 7B), particularly in the peri-infarct area (Supporting Information Fig. S6). Many donor pericytes retained expression of NG2, a pericyte marker (Supporting Information Fig. S7). Confocal microscopy showed that a fraction of pericytes were identified in perivascular positions, adjacent to host CD31-positive endothelial cells (Fig. 7C). Indeed, pericytes were aligned with (Fig. 7C, main) or surrounding (Fig. 7C, inset) CD31-positive microvessels, suggestive of perivascular homing. The number of engrafted GFP-positive pericytes was approximately 9.1% ± 1.3% of total injected cells at the first week and declined over time to 3.4% ± 0.5% at 8 weeks post-infarction (n = 3 per time point) (Fig. 7D, dash-dot line). The perivascular homing rate instead increased from 28.6% to 40.1% over the course of 8 weeks, implicating the merit of niche-homing for long-term donor cell survival (Fig. 7D, solid line). To demonstrate cellular interactions between donor pericytes and host ECs, we performed immunohistochemical studies for ephrin type-B receptor 2 (EphB2) and connexin43, a gap junction protein. Confocal images revealed that some GFP-positive pericytes juxtaposing host ECs expressed human-specific EphB2 (Fig. 7E) or formed gap junctions with ECs (Fig. 7F). These results suggest that cellular interactions between host ECs and donor pericytes homed to perivascular locations.

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Figure 7. Transplanted pericytes home to perivascular locations. (A): Pericytes were transduced with GFP reporter at nearly 95% efficiency. Fluorescence (A, main) and bright-field (A, inset) images were taken from the same low-power field (scale bar = 200 μm). (B): Engraftment of GFP-labeled pericytes within host myocardium was revealed by anti-GFP immunostaining at 1 week postinjection (scale bar = 500 μm, infarct site encircled by dotted lines). (C): Pericytes were lining with (C, main) or surrounding (C, inset) host CD31-positive microvasculature (scale bar = 20 μm). (D): The engraftment efficiency of pericytes at 1 week (9.1% ± 1.3%) and 8 weeks (3.4% ± 0.5%) postinfarction was depicted (dash-dot line). The perivascular homing ratio instead increased from 28.6% to 40.1% and was delineated separately (solid line). Some GFP-positive pericytes juxtaposing host ECs (E) expressed human-specific EphB2 (green/white arrows) or (F) formed connexin43-positive gap junctions with endothelial cells (red arrow heads) (scale bar = 10 μm). Abbreviation: GFP, green fluorescent protein.

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Cell Lineage Fate of Transplanted Pericytes

GFP-labeled pericytes were used to track cell lineages developed from donor pericytes and investigate the capacity of human muscle pericytes to reconstitute major cardiac cell types after injury. Immunohistochemistry was performed to simultaneously detect GFP and cell lineage markers: the cardiomyocyte marker, cardiac troponin-I (cTn-I); the smooth muscle cell marker, smooth muscle myosin heavy chain (SM-MHC); the endothelial cell (EC) marker, CD31. Confocal microscopy revealed that in the peri-infarct area, a minor fraction of transplanted pericytes coexpress GFP and cTn-I (Supporting Information Fig. S8A–S8C, main), a few of which appear single-nucleated (Supporting Information Fig. S8A–S8C, inset). Some GFP-positive cardiomyocytes were identified within the remaining myocardium (Supporting Information Fig. S8D–S8F) with organized sarcomeric patterns (Supporting Information Fig. S8G–S8I). A very small number of donor pericytes coexpressed GFP and human-specific CD31 (<1%) (Supporting Information Fig. S8J-S8L). Similarly, coexpression of GFP and human-specific SM-MHC was detected in very few transplanted cells (<0.5%) (Supporting Information Fig. S8M–S8O). Negative control images were stained only with matching fluorescence-conjugated secondary antibodies (Supporting Information Fig. S8P–S8R).

DISCUSSION

  1. Top of page
  2. Abstract
  3. INTRODUCTION
  4. MATERIALS AND METHODS
  5. RESULTS
  6. DISCUSSION
  7. CONCLUSION
  8. Acknowledgements
  9. DISCLOSURE OF CONFLICTS OF INTEREST
  10. REFERENCES
  11. Supporting Information

Pericytes constitute a major structural component of small blood vessels, regulating vascular development, integrity, and physiology. The recent identification of microvascular pericytes as one of the native sources of MSC ancestors raised the possibility that these cells participate in the repair of injured/ageing organs [11–16, 35]. The therapeutic potential of microvascular pericytes was indicated by structural and functional regeneration of skeletal muscle involving direct pericyte differentiation into regenerative units as well as applications in lung repair and vascular tissue engineering [8, 11, 12, 35, 36]. Pericytes can also repair tissue via secretion of trophic factors, implying broad usage in clinical settings [8, 18]. Recent studies indicated a possible developmental hierarchy among different stem/progenitor cell populations residing in the blood vessel walls [15, 37]. Katare et al. [38] reported that transplantation of adventitial progenitor cells repairs infarcted hearts through angiogenesis involving microRNA-132. Herein we demonstrate that transplantation of human FACS-purified microvascular pericytes contributes to the functional and structural repair of the ischemic heart, albeit unequally, through both paracrine effects and cellular interactions. A major goal of SCT, the prevention of progressive LV dilatation and consequent HF, was largely achieved by pericyte treatment, implicating the attenuation of deleterious remodeling. We also observed significant improvement of cardiac contractility in an acute infarction milieu, with up to 70% of healthy contractile function consistently maintained for at least 2 months. No significant difference was observed between adult and fetal pericytes in terms of heart repair. The therapeutic benefits observed could be explained, at least in part, by antifibrotic, anti-inflammatory, angiogenic, and to a lesser extent, cardiomyogenic properties of pericytes.

The antifibrotic action of mesodermal stem/progenitor cells in the injured heart has been reported [25–27]. MSC-conditioned medium diminished viability, proliferation, collagen synthesis, and α-Smooth Muscle Actin (SMA) expression in cardiac fibroblasts but stimulated MMP-2/-9 activities, indicating a paracrine antifibrotic property of MSCs [26, 27]. Our results demonstrated a near 50% reduction of myocardial fibrosis following pericyte injection. Along with the attenuation of progressive LV dilatation, pericyte treatment appears to result in propitious remodeling, leading to improved myocardial compliance and strengthening of the ischemic cardiac tissue.

We speculated that decreased fibrosis/scar formation is, at least partially, associated with a reduced number of fibrotic cells resulting from the administration of pericytes. Interestingly, pericyte-treated hearts contained significantly less cells within the infarct area than PBS-injected controls (p < .05, Supporting Information Fig. S9A) with no statistical difference in Ki-67-positive proliferating cell density (p = .808, Supporting Information Fig. S9B). Additionally, Terminal deoxynucleotidyl transferase dUTP nick end labeling (TUNEL) staining revealed no significant difference between pericyte- and PBS-injected hearts in the number of apoptotic cells within the infarct region (p = .296, Supporting Information Fig. S10A, S10B). Due to the highly fibrotic nature of MI, we were unable to quantitate fibrotic cells in vivo. Nevertheless, murine fibroblast proliferation was notably inhibited when cultured in hypoxic P-CM in vitro, indicating the paracrine fibrosuppressive effect of pericytes under hypoxia. MMPs were suggested to play important roles in postinjury scar remodeling, angiogenesis, and vascular cell proliferation/migration [27, 39]. In particular, a preponderant role of MMP-2 in preventing collagen accumulation by cardiac fibroblasts was proposed [27]. Consequently, we postulated the existence of a fibrolytic activity from donor pericytes, involving MMPs, which contributes to the attenuation of cardiac fibrosis. Indeed, high expression of MMP-2, but not MMP-9, by pericytes, even under hypoxic conditions, was observed. The expression of MMP-2 was confirmed in some, but not all, transplanted pericytes, implying a minor role of pericyte-mediated fibrolysis. Overall, our data suggest that the gross amelioration of fibrosis presumably involves decreased collagen deposition, reduced proliferation of fibroblasts, and altered remodeling of the extracellular matrix (ECM). Yet whether there exists one or more determining mechanism(s) remains to be investigated.

The immunosuppressive potential of MSCs, demonstrated by inhibiting T-lymphocyte proliferation in culture and counteracting graft-versus-host reaction in recipients of allogeneic blood stem cells, is currently exploited in clinical trials [28, 29]. In the cardiac milieu, MSC transplantation in a rat model of acute myocarditis mitigated the increase in CD68+ phagocytic cells [40]. In this study, pericyte treatment significantly diminished host monocyte/macrophage infiltration in the infarcted myocardium, suggesting an anti-inflammatory potential which contributed to the reduction of fibrosis, amelioration of adverse remodeling, and improvement of cardiac function. Nevertheless, whether pericytes inhibit the acute-phase inflammation occurring soon after the incidence of MI is unknown. Inhibition of murine macrophage proliferation in culture by P-CM suggests a paracrine mechanism of their immunomodulatory capacity.

The immunosuppressive and anti-inflammatory capacities of MSCs are primarily attributed to soluble factors/molecules, as IL-6, LIF, and HMOX-1 were shown to exercise beneficial immunosuppressive effects [28, 29]. The attenuation of intense inflammation and mitigation of multiorgan damage by MSCs during sepsis are dependent on monocyte/macrophage-derived cytokines and regulated via PGE2 signaling [41]. The immunoregulatory and cardioprotective functions of these molecules appear to be similar in the cardiac milieu [22, 42]. Our data demonstrate that pericytes express high levels of IL-6, LIF, COX-2, HMOX-1, and HIF-1α, which are sustained under hypoxic conditions. MCP-1 expression, however, notably decreased under hypoxia, corresponding with the reduced CD68+ cell infiltration in vivo. Little to no expression of proinflammatory cytokines including IL-1α, TNF-α, and IFNγ was detected. Virtually no expression of IL-4, IL-10, iNOS, and 2,3-IDO was observed in pericytes, suggestive of an immunoregulatory cytokine secretome that is unique to human microvascular pericytes [29, 43]. Intriguingly, TGF-β1, also an anti-inflammatory yet fibrogenic cytokine, was strongly expressed by pericytes [22, 44]. Our data suggested increased TGF-β1 expression/secretion by pericytes under hypoxic conditions (Fig. 6A, 6C). Given the multiple functions each proposed growth factor/cytokine possesses, it is likely that a dynamic, interactive, and intricately orchestrated balance of trophic factors between donor and host cells holds the key to a successful ischemic tissue remodeling and regeneration.

A linear correlation between secretion of proangiogenic factors, angiogenesis, and cardiac restoration was illustrated by blocking the bioactivity of VEGF secreted from transplanted murine muscle-derived stem cells in a mouse MI model, which not only abolished their stimulation of neovascularization but in turn negatively influenced LV contractility and infarct size [45]. Okada et al. [34] further delineated the superior angiogenic properties of human myoendothelial progenitor cells and increased secretion of VEGF in response to hypoxia. Given the indigenous vascular association of pericytes, we hypothesized that pericytes are able to repair the damaged host vasculature. Indeed, upon pericyte treatment, we observed a significantly larger host microvascular network not only in the peri-infarct collateral circulation but also within the infarct region. Cultured pericytes secrete growth factors/cytokines/chemokines related to vascular physiology and remodeling [18]; among which, only VEGF-A, PDGF-β, and TGF-β1 were substantially upregulated under hypoxia, suggesting their role in pericyte-enhanced angiogenesis [39, 46].

Angiogenesis may follow cell–cell contact between donor pericytes and host ECs, in addition to stimulation by angiogenic factors. Recent studies reported that MSCs and vascular mural/adventitial cells support ECs in small blood vessel formation and maturation in culture and in vivo [33, 47]. We did observe the perivascular homing of donor pericytes in the ischemic heart. Some donor pericytes juxtaposing host ECs expressed interactive molecules including EphB2 and connexin43, suggestive of cellular interactions [48, 49]. Planar Matrigel culture confirmed the vascular cell characteristics of pericytes and their capability to enhance the angiogenic behavior of ECs. We further demonstrated microvessel formation and vascular support by pericytes in 3D cultures, indicating that associations between pericytes and ECs may contribute to revascularization. Nevertheless, the vibrant angiogenic response of pericytes observed in vitro may be reduced in vivo because of the harsh microenvironment caused by post-MI, ischemia, and inflammation. Altogether, these results demonstrate that the angiogenic properties of pericytes may result from indirect paracrine effects and, albeit minor, direct cellular interactions.

Compared to other types of stem/progenitor cells, pericytes appear to engraft well in the infarcted heart initially, presumably attributable to several factors [50]. We did not observe apparent cell death of pericytes cultured under 2.5% O2 for up to 48 hours, implying their resistance to hypoxia (data not shown). The increased proliferation and migration of pericytes in response to low oxygen concentration and ECM degradation products have important implications for ischemic injury repair [10]. The perivascular niche-homing capacity may further benefit the long-term survival of pericytes. Nonetheless, it remains unclear whether pericytes actively migrated to perivascular locations or served as a revascularizing center inducing/recruiting angiogenic proliferation/migration of host ECs. Future studies are needed to reveal the kinetics of pericyte-EC interaction and migration in vivo.

The potential of human muscle pericytes to reconstitute major cell types in the injured myocardium, although to a small extent, was hereby demonstrated. Cell fate tracking suggests that a minor fraction of donor pericytes differentiated into and/or fused with cardiomyocytes. Given the small number of GFP-cTnI dual-positive cells present, it is unlikely that these cells contributed significantly to functional recovery [21]. Pericytes, all of which were α-SMA-positive during culture expansion, lost α-SMA expression once homing to host microvasculature (data not shown), consistent with our finding that a subset of native microvascular pericytes do not express α-SMA in situ. [12]

CONCLUSION

  1. Top of page
  2. Abstract
  3. INTRODUCTION
  4. MATERIALS AND METHODS
  5. RESULTS
  6. DISCUSSION
  7. CONCLUSION
  8. Acknowledgements
  9. DISCLOSURE OF CONFLICTS OF INTEREST
  10. REFERENCES
  11. Supporting Information

In summary, FACS-purified human microvascular pericytes contribute to anatomic and functional cardiac improvement postinfarction through multiple cardioprotective mechanisms: reverse of ventricular remodeling, reduction of cardiac fibrosis, diminution of chronic inflammation, and promotion of host angiogenesis. Vessel-homing and small-scale regenerative events by pericytes partially reconstitute lost cardiac cells and contribute to the structural recovery. These cardioprotective and cardioregenerative activities of a novel stem cell population that can be purified to homogeneity and expanded in vitro await further research and exploitation in ischemic cardiovascular diseases.

Acknowledgements

  1. Top of page
  2. Abstract
  3. INTRODUCTION
  4. MATERIALS AND METHODS
  5. RESULTS
  6. DISCUSSION
  7. CONCLUSION
  8. Acknowledgements
  9. DISCLOSURE OF CONFLICTS OF INTEREST
  10. REFERENCES
  11. Supporting Information

We thank Alison Logar for excellent technical assistance with flow cytometry, Dr. Bin Sun for expert assistance in real-time qPCR, Dr. Simon Watkins for confocal microscopy, Dr. Bing Wang for lentiviral-GFP vectors, James H. Cummins for editorial assistance, as well as Bolat Sultankulov and Damel Mektepbaeva for insightful discussion. This work was supported by grants from the Commonwealth of Pennsylvania (B.P.), National Institute of Health R01AR49684 (J.H.) and R21HL083057 (B.P.), the Henry J. Mankin Endowed Chair (J.H.), the William F. and Jean W. Donaldson Endowed Chair (J.H.), and the Ministry of Education and Science of the Republic of Kazakhstan (A.S.). C.W.C. was supported in part by the American Heart Association predoctoral fellowship. M.C. was supported by the California Institute for Regenerative Medicine training grant (TG2-01169).

DISCLOSURE OF CONFLICTS OF INTEREST

  1. Top of page
  2. Abstract
  3. INTRODUCTION
  4. MATERIALS AND METHODS
  5. RESULTS
  6. DISCUSSION
  7. CONCLUSION
  8. Acknowledgements
  9. DISCLOSURE OF CONFLICTS OF INTEREST
  10. REFERENCES
  11. Supporting Information

J.H. received remuneration from Cook MyoSite, Inc. for consulting services and for royalties received from technology licensing during the period that the above research was performed. All other authors have no conflict of interest to disclose.

REFERENCES

  1. Top of page
  2. Abstract
  3. INTRODUCTION
  4. MATERIALS AND METHODS
  5. RESULTS
  6. DISCUSSION
  7. CONCLUSION
  8. Acknowledgements
  9. DISCLOSURE OF CONFLICTS OF INTEREST
  10. REFERENCES
  11. Supporting Information

Supporting Information

  1. Top of page
  2. Abstract
  3. INTRODUCTION
  4. MATERIALS AND METHODS
  5. RESULTS
  6. DISCUSSION
  7. CONCLUSION
  8. Acknowledgements
  9. DISCLOSURE OF CONFLICTS OF INTEREST
  10. REFERENCES
  11. Supporting Information

Additional Supporting Information may be found in the online version of this article.

FilenameFormatSizeDescription
sc-12-0465_sm_SupplFigure1.pdf23KSupplemental Figure S1. Fluorescence-activated cell sorting (FACS) of human muscle pericytes. Human muscle biopsies were mechanically dissociated and digested with collagenase. After labeling, single cells suspension was subjected to FACS to purify the pericyte population based on their differential expression of cell lineage markers, including robust expression of the pericyte/endothelial cell marker CD146 and absence of the myogenic, endothelial/stem, and hematopoietic cell markers: CD56, CD34 and CD45, respectively. Skeletal muscle-derived pericytes (CD146+/34-/45-/56-) sorted to homogeny were collected and expanded in vitro.
sc-12-0465_sm_SupplFigure2A.pdf62KSupplemental Figure S2. (A) Morphology of purified pericytes in long-term culture. Flow cytometry analysis revealed that cultured pericytes retain original cell surface marker expression, including the robust expression of CD146 and alkaline phosphatase (ALP) with the absence of CD34, CD45, and CD56. (B) Flow cytometry analysis showed that long-term cultured pericytes strongly express classic MSC markers: CD90, CD44, CD73 and CD105.
sc-12-0465_sm_SupplFigure2B.pdf39KSupplemental Figure S2. (A) Morphology of purified pericytes in long-term culture. Flow cytometry analysis revealed that cultured pericytes retain original cell surface marker expression, including the robust expression of CD146 and alkaline phosphatase (ALP) with the absence of CD34, CD45, and CD56. (B) Flow cytometry analysis showed that long-term cultured pericytes strongly express classic MSC markers: CD90, CD44, CD73 and CD105.
sc-12-0465_sm_SupplFigure3.tif2672KSupplemental Figure S3. Representative echocardiographic M-mode images of left ventricle (LV). End-systolic dimension (ESD) and end-diastolic dimension (EDD) were indicated.
sc-12-0465_sm_SupplFigure4.pdf40KSupplemental Figure S4. Echocardiographic measurement of cardiac function in acutely infarcted hearts injected with either APs (CD146) or CD56+ myogenic progenitors (CD56) at 2 weeks post-infarction. APs and CD56+ myogenic progenitors were simultaneously sorted from a single adult muscle biopsy. Pericyte-treated hearts (N=6) exhibited significantly better LV function than CD56+ progenitor-treated ones (N=6) in multiple categories, including LVEDA (p=0.004), LVESA (p=0.002), LVFS (p=0.003), LVFAC (p=0.003), and LVEF (p≤0.001).
sc-12-0465_sm_SupplFigure5.pdf50KSupplemental Figure S5. Detection of host EC proliferation at 2 weeks post-infarction. Representative images of Ki-67 (green) and mouse CD31 (red) co-immunostaining (A) within the infarct region and (B) in the peri-infarct area of hearts injected with pericytes or PBS (scale bars=50μm). Proliferating host ECs were identified as Ki-67/CD31 dual-positive cells (green/red arrows). (C) Pericyte-treated group had a significantly larger number of proliferating ECs both within the infarct region (p=0.034) and in the peri-infarct area (p=0.025) than PBS control group (N=3 per group).
sc-12-0465_sm_SupplFigure6.pdf358KSupplemental Figure S6. (A-D) Donor pericytes (stained in red by anti-GFP) were particularly abundant in the peri-infarct area where host CD31(+) capillaries (mouse CD31 stained in green) remained largely intact (scale bar=50μm).
sc-12-0465_sm_SupplFigure7.tif2320KSupplemental Figure S7. Expression of the pericyte marker NG2 (chondroitin sulphate) was detected in the majority of, but not all, GFP-positive donor pericytes at 2 weeks post-infarction (scale bar=10μm).
sc-12-0465_sm_SupplFigure8-1.pdf273KSupplemental Figure S8. Tracking cardiac cell lineage fates of donor pericytes. Confocal microscopy revealed that in the peri-infarct area (A-C) a minor fraction of GFP-labeled pericytes co-expressed a mature cardiomyocyte marker, cardiac troponin-I (cTn-I), while additional GFP(+) cells remained in the interstitium ([A], main, scale bar=50μm); a few of them appear single-nucleated ([A], inset, scale bar=10μm) (cTn-I in red, Anti-GFP in green). (D-F) Immunofluorescent detection of GFP-cTn-I dual-positive cardiomyocytes integrating within the residual myocardium (scale bar=50μm) with dotted area enlarged in (G-I) showing sarcomeric patterns (scale bar=20μm). (J-L) A very small number of GFP(+) pericytes (<1%) co-expressed human-specific CD31 (hCD31 in red, Anti-GFP in green; scale bar=10μm). (M-O) Few donor pericytes (<0.5%) expressed human-specific SM-MHC (hSM-MHC in red, Anti-GFP in green; scale bar=10μm). (P-R) Negative control images were taken from sections immunostained only with matching fluorescence-conjugated secondary antibodies (no primary antibody) (Cy3 in red, AlexaTM488 in green; scale bar=10μm).
sc-12-0465_sm_SupplFigure8-2.pdf301KSupplemental Figure S8. Tracking cardiac cell lineage fates of donor pericytes. Confocal microscopy revealed that in the peri-infarct area (A-C) a minor fraction of GFP-labeled pericytes co-expressed a mature cardiomyocyte marker, cardiac troponin-I (cTn-I), while additional GFP(+) cells remained in the interstitium ([A], main, scale bar=50μm); a few of them appear single-nucleated ([A], inset, scale bar=10μm) (cTn-I in red, Anti-GFP in green). (D-F) Immunofluorescent detection of GFP-cTn-I dual-positive cardiomyocytes integrating within the residual myocardium (scale bar=50μm) with dotted area enlarged in (G-I) showing sarcomeric patterns (scale bar=20μm). (J-L) A very small number of GFP(+) pericytes (<1%) co-expressed human-specific CD31 (hCD31 in red, Anti-GFP in green; scale bar=10μm). (M-O) Few donor pericytes (<0.5%) expressed human-specific SM-MHC (hSM-MHC in red, Anti-GFP in green; scale bar=10μm). (P-R) Negative control images were taken from sections immunostained only with matching fluorescence-conjugated secondary antibodies (no primary antibody) (Cy3 in red, AlexaTM488 in green; scale bar=10μm).
sc-12-0465_sm_SupplFigure9.pdf19KSupplemental Figure S9. Comparison of total and proliferating cell density within the infarct area at 2 weeks post-infarction. (A) Pericyte-injected hearts had a significantly less number of cells within the infarct area than the PBS-injected controls (p<0.05, N=5 per group). (B) A cell proliferation marker, Ki-67, was used to detect proliferating cells within the infarct area. No statistical difference in Ki-67(+) proliferating cell density was observed between pericyte- and PBS-injected hearts (p=0.808, N=3 per group).
sc-12-0465_sm_SupplFigure10.pdf27KSupplemental Figure S10. (A) Terminal dUPT nick end-labeling (TUNEL) staining revealed the apoptotic cells within the infarct region of pericyte- or PBS-injected hearts (scale bars=100μm). (B) Quantification of the apoptotic cell number (N=5 per group) showed no statistical difference between pericyte treatment (12.55±1.87 cells/mm2) and PBS injection (9.01±2.55 cells/mm2) at 2 weeks post-infarction (p=0.296).
sc-12-0465_sm_SupplTable1.pdf49KSupplementary Table 1
sc-12-0465_sm_SupplTable2.pdf56KSupplementary Table 2
sc-12-0465_sm_SupplTable3.pdf9KSupplementary Table 3
sc-12-0465_sm_Supplinfor.pdf172KSupplementary Data

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