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Keywords:

  • Endothelial progenitor cells;
  • Cell transplantation;
  • Bone marrow transplantation;
  • Pulmonary hypertension;
  • Microvasculature;
  • Chronic hypoxia

Abstract

  1. Top of page
  2. Abstract
  3. Introduction
  4. Materials and Methods
  5. Results
  6. Discussion
  7. Conclusion
  8. Disclosure of Potential Conflicts of Interest
  9. Acknowledgements
  10. References
  11. Supporting Information

Circulating endothelial progenitor cells (EPCs) contribute to neovascularization of ischemic tissues and repair of injured endothelium. The role of bone marrow-derived progenitor cells in hypoxia-induced pulmonary vascular remodeling and their tissue-engineering potential in pulmonary hypertension (PH) remain largely unknown. We studied endogenous mobilization and homing of EPCs in green fluorescent protein bone marrow chimeric mice exposed to chronic hypoxia, a common hallmark of PH. Despite increased peripheral mobilization, as shown by flow cytometry and EPC culture, bone marrow-derived endothelial cell recruitment in remodeling lung vessels was limited. Moreover, transfer of vascular endothelial growth factor receptor-2+/Sca-1+/CXCR-4+-cultured early-outgrowth EPCs failed to reverse PH, suggesting hypoxia-induced functional impairment of transferred EPCs. Chronic hypoxia decreased migration to stromal cell-derived factor-1α, adhesion to fibronectin, incorporation into a vascular network, and nitric oxide production (−41%, −29%, −30%, and −32%, respectively, vs. normoxic EPCs; p < .05 for all). The dysfunctional phenotype of hypoxic EPCs significantly impaired their neovascularization capacity in chronic hind limb ischemia, contrary to normoxic EPCs cultured in identical conditions. Mechanisms contributing to EPC dysfunction include reduced integrin αv and β1 expression, decreased mitochondrial membrane potential, and enhanced senescence. Novel insights from chronic hypoxia-induced EPC dysfunction may provide important cues for improved future cell repair strategies.

Disclosure of potential conflicts of interest is found at the end of this article.


Introduction

  1. Top of page
  2. Abstract
  3. Introduction
  4. Materials and Methods
  5. Results
  6. Discussion
  7. Conclusion
  8. Disclosure of Potential Conflicts of Interest
  9. Acknowledgements
  10. References
  11. Supporting Information

Circulating endothelial progenitor cells (EPCs) are mobilized from the bone marrow in response to tissue ischemia or traumatic injury [1]. At sites of vessel injury, EPCs can differentiate into mature endothelial cells and play an important role in tissue repair and endothelial function recovery [2, 3]. The endogenous regenerative capacity of EPCs, however, was significantly impaired in patients with coronary artery disease [4] and type II diabetes [5] but has received little attention in patients with pulmonary vascular disease.

Pulmonary hypertension (PH) is a devastating disease with significant morbidity and mortality, characterized by increased pulmonary artery pressure, right ventricular hypertrophy, and pulmonary vascular remodeling [6, 7]. In rat and canine models of monocrotaline-induced pulmonary endothelial injury, early-outgrowth EPCs have been shown to mitigate toxin-induced PH [8, [9]10], spurring increased interest in culture-expanded EPCs as a promising tissue-engineering tool in PH. Recently, circulating progenitor cells were shown to participate in hypoxic pulmonary vascular remodeling with recruitment of c-Kit+ cells to the adventitia of hypoxic bovine lung vessels [11] and of green fluorescent protein (GFP)-labeled bone marrow-derived EPCs to the endothelium of hypoxic murine lung vessels [12]. The biological significance of endogenous bone marrow cell recruitment to hypoxic lungs remains unknown. Also, whether or not bone marrow-derived progenitor cells can be considered in cell transfer strategies for hypoxia-induced PH is unknown and depends in part on adhesive, migratory, and vasculogenic properties of EPCs under these pathophysiologic conditions.

Therefore, we proposed to study the endogenous bone marrow-derived cellular response to pulmonary vascular remodeling in bone marrow chimeric mice, as well as the potential of exogenous EPC transfer to reduce PH. We observed that chronic hypoxia significantly impairs EPC phenotype compared with normoxic (Nx) mice, limiting therapeutic efficacy of EPCs, and we investigated several mechanisms that could contribute to EPC dysfunction. We identified reduced integrin αv and β1 expression, impaired mitochondrial transmembrane potential, and enhanced cellular senescence in EPCs from mice exposed to chronic hypoxia. These findings could provide novel cues to reverse EPC dysfunction, not only in chronic hypoxia but also, by analogy, in patients with cardiovascular risk factors.

Materials and Methods

  1. Top of page
  2. Abstract
  3. Introduction
  4. Materials and Methods
  5. Results
  6. Discussion
  7. Conclusion
  8. Disclosure of Potential Conflicts of Interest
  9. Acknowledgements
  10. References
  11. Supporting Information

Hypoxia-Induced Pulmonary Hypertension

Pulmonary hypertension was induced by exposing C57Bl6/N mice to chronic hypoxia (FIO2 0.10 for 3 weeks) and assessed by measuring right ventricular systolic pressure (RVSP; 1.4 F Millar catheter; Millar Instruments, Houston, TX, http://www.millarinstruments.com) under urethane anesthesia and right ventricular (RV) hypertrophy (post-mortem RV fractional weight). Hematocrit was determined following centrifugation of heparinized blood. Pulmonary vascular remodeling was assessed following perfusion fixation via the RV using phosphate buffered saline and Z-fix (Anatech Ltd., Battle Creek, MI, http://www.anatechltdusa.com/) at a perfusion pressure of 50 cm H2O. At the same time, the airways were inflated using Z-fix at 25 cm of H2O pressure. Lungs from Nx and chronic hypoxic (CHx) animals were incubated overnight in Z-fix and embedded in paraffin. Sections (5 μm) were incubated with anti-GFP (ab6556; Abcam, Cambridge, MA, http://www.abcam.com), anti-von Willebrand factor (anti-vWF; A0082; Dako, Glostrup, Denmark, http://www.dako.com), or anti-CD45 antibody (553076; BD Pharmingen, San Diego, http://www.bdbiosciences.com/index_us.shtml) diluted 1/500, and specific binding was visualized with diaminobenzidine (Sigma-Aldrich, St. Louis, http://www.sigmaaldrich.com). Pictures were obtained with an Axiovert 200M microscope (Carl Zeiss, Jena, Germany, http://www.zeiss.com) using AxioVision acquisition software.

Generation of Chimeric Mice

Wild-type C57Bl6/N mice, 8 weeks of age, were lethally irradiated (9.5 Gy of total-body irradiation) using a linear accelerator (GE Healthcare, Little Chalfont, U.K., http://www.gehealthcare.com) at 3.9 Gy/minute. Irradiation was followed within 24 hours by injection of 5 × 106 total bone marrow cells harvested from femurs of C57Bl6/N mice ubiquitously expressing GFP. Bone marrow reconstitution was allowed for 8 weeks.

EPC Isolation

After homogenization of the spleen, mononuclear cells (MNCs) were isolated from CHx and age-matched Nx mice by density centrifugation (Histopaque 1083; Sigma-Aldrich). MNCs (40 × 106) were plated on fibronectin-coated (Tebu-Bio, Boechout, Belgium, http://www.tebu-bio.com/) six-well plates in endothelial basal medium 2 (EBM-2; Cambrex, Walkersville, MD, http://www.cambrex.com) supplemented with 5% fetal bovine serum, vascular endothelial growth factor (VEGF), basic fibroblast growth factor (bFGF), recombinant insulin-like growth factor, epidermal growth factor, ascorbic acid, and gentamicin/amphotericin-B. After 4 days of culture, nonadherent cells were removed, and cells were cultured for 3 additional days. These early-outgrowth EPCs were used for in vitro analysis of EPC function.

EPC Transfer During PH

After 1 or 2 weeks of exposure to hypoxia (FIO2 0.10), C57BL6/N mice were randomized to receive either saline or 5 × 105 EPCs via jugular vein injection (n = 6–8 in each group). After a total of 3 weeks of hypoxia, RVSP and RV hypertrophy were measured. In cell tracking experiments, EPCs derived from luciferase-expressing mice (described in the supplemental online Methods) or EPCs labeled with CM-1,1′-dioctadecyl-3,3,3′,3′-tetramethylindocarbocyanine perchlorate (DiI) according to the manufacturer's instructions (Molecular Probes, Eugene, OR, http://probes.invitrogen.com) were transplanted at the indicated time points. Homing of EPCs was determined at different time points using bioluminescence imaging [13, 14] (described in the supplemental online Methods). For detection of CM-DiI-labeled EPCs, lungs and spleens were paraffin-embedded, and sectioned. Sections were stained with 4′,6-diamidino-phenylindole (DAPI; Vector Laboratories, Burlingame, CA, http://www.vectorlabs.com) to detect nuclei or for vWF using an Alexa647-conjugated secondary antibody to detect blood vessels. Images were obtained with a LSM510 confocal microscope (Carl Zeiss). All animal experiments were approved by the ethical committee of the University of Leuven.

EPC Characterization and Enumeration

Characterization of In Vitro-Cultured EPCs.

After 7 days of culture, adherent spleen-derived cells were incubated with 12 μg/ml DiI-labeled acetylated low-density lipoprotein (acLDL) (Molecular Probes) for 4 hours. After fixation with 1% paraformaldehyde, cells were incubated with fluorescein-isothiocyanate-conjugated lectin from Bandeiraea simplicifolia (BS1 lectin; 1 hour; 10 μg/ml; Sigma-Aldrich), and fluorescent images were obtained (LSM510 confocal microscope; Carl Zeiss). Expression analysis was done by flow cytometry using a FACSCalibur (BD Biosciences, San Diego, http://www.bdbiosciences.com). Antibodies for vascular endothelial growth factor receptor-2 (VEGFR-2), Sca-1, c-Kit (CD117), and CXCR-4 were from BD Pharmingen, and isotype controls were run in parallel.

Analysis of Circulating EPC Number.

Blood was collected from the vena cava, and Fc block (BD Pharmingen) was added to the samples prior to antibody incubation. Fluorescently labeled antibodies against CD45/VEGFR-2 and CD45/VEGFR-2/c-Kit (all from BD Pharmingen) were used to enumerate circulating endothelial cells and circulating endothelial progenitors, respectively. After 30 minutes of incubation, red blood cells were lysed (PharmLyse; BD Pharmingen), and samples were evaluated by flow cytometry. Isotype controls were run in parallel. Approximately 50,000 events were recorded. Following culture of spleen-derived MNCs, adhering acetylated LDL- and BS1 lectin-positive EPCs from Nx and CHx animals were counted in 10 high-power fields (HPFs), and the number of EPCs per mouse was determined from the surface area of the culture dish and total number of MNCs isolated.

EPC Function In Vitro

Migration.

Migration toward stromal cell-derived factor-1α (SDF-1α) was determined by resuspending 1 × 105 EPCs derived from Nx and CHx mice in 100 μl of EBM-2 plus 0.5% bovine serum albumin (BSA) in the upper chamber of a modified Boyden chamber (8 μm pore size; Corning Life Sciences, Schiphol-Rijk, The Netherlands, http://www.corning.com/lifesciences). The upper chamber was transferred to a 24-well containing EBM-2/BSA and 150 ng/ml SDF-1α. After 5 hours at 37°C, membrane inserts were fixed with 4% formaldehyde. SDF-1α was kindly provided by M. Tjwa (Center for Transgene Technology and Gene Therapy, Leuven, Belgium). Cell nuclei of migrated EPCs were stained using DAPI, and the difference in migrating EPCs toward EBM-2/BSA with or without SDF-1α was counted in four HPFs.

Adhesion.

To evaluate adhesion, 50,000 EPCs were resuspended in EBM-2; plated onto fibronectin-coated (5 μg/cm2; Tebu-Bio), 1% BSA-blocked 24-wells; and incubated for 30 minutes. The number of adherent cells per 10 HPFs was counted and compared between EPCs derived from Nx and CHx mice.

Incorporation into a Vascular Network.

EPCs from Nx and CHx mice were incubated with DiI-labeled acetylated LDL for 4 hours and coplated with human umbilical vein endothelial cells (HUVECs; Cambrex) in a 1:2 proportion on presolidified Matrigel (BD Biosciences). After 16 hours, the number of EPCs incorporating into the vascular network per HPF were determined. The average of six HPFs per experiment was calculated.

Nitric Oxide Production.

Nitrite, nitrate, and S-nitrosothiol (NOx) levels were determined as stable oxidative products of nitric oxide (NO) in the supernatants of Nx and CHx EPCs using a Sievers 280 chemiluminescent NO analyzer (GE Analytical Instruments, Boulder, CO, http://www.geinstruments.com) [15]. Negative control values (unconditioned medium) were subtracted from the sample values (24 hours of conditioning), and NOx concentrations, normalized to cell number, were calculated. HUVECs and the murine endothelial cell line MS1 (CRL-2279; American Type Culture Collection, Manassas, VA, http://www.atcc.org) were used in control experiments. To demonstrate nitric oxide synthase (NOS)-dependent NO production, experiments were performed in the presence and absence of 2 mM NG-mono-methyl-l-arginine (l-NMA; Sigma-Aldrich).

Integrin and Nitric Oxide Synthase 3 (NOS3) Expression Analysis.

Transcript levels of integrins α5, αv, β1, β2, β3, and NOS3 were determined by reverse transcriptase-polymerase chain reaction using specific primers and fam-labeled probes (supplemental online Table 1). Results were expressed as relative copy numbers standardized to levels of the housekeeping 18S rRNA gene (Applied Biosystems), and the average values for EPCs derived from CHx mice were normalized to the average values in EPCs derived from Nx mice.

Senescence-Associated β-Galactosidase Staining.

Gently fixed EPCs were incubated for 6 hours at 37°C (without CO2) with fresh senescence-associated 5-bromo-4-chloro-3-indolyl-β-d-galactopyranoside (X-gal) staining solution (1 mg/ml X-gal [Immunosource, Halle-Zoersel, Belgium, http://www.immunosource.com], 40 mM citric acid/sodium phosphate, pH 6, 5 mM K3Fe[CN]6, 5 mM K4Fe[CN]6, 150 mM NaCl, 2 mM MgCl2) [16]. After incubation, nuclei were counterstained with nuclear fast red, and the percentage of cells showing senescence was determined on 10 HPFs per condition.

Mitochondrial Membrane Potential Analysis, Lactate Production, and Cellular ATP Content

EPCs cultured from Nx and CHx mice were loaded with the potentiometric mitochondrial dye JC-1 (Molecular Probes) at 4 μM for 20 minutes at 37°C. After two washing steps, green and red mean fluorescence intensities were evaluated using flow cytometry or confocal microscopy. To exclude any influence of different mitochondrial size, shape, or density, we calculated the ratio of red to green fluorescence for each experiment. In control experiments, the mitochondrial membrane potential was dissipated with carbonyl cyanide 4-(trifluoromethoxy)phenylhydrazone (FCCP; 10 μM; Sigma-Aldrich) [17]. Lactate concentration in conditioned supernatants was measured using an enzymatic method on an automatic analyzer (Dimension analyzer; Siemens Healthcare Diagnostics, Brussels, Belgium, http://diagnostics.siemens.com/). In the presence of excess NAD+ and lactate dehydrogenase, lactate was converted to pyruvate and NADH. The formation of NADH results in increased absorbance at 340 nm, and this value was corrected for nonspecific signal by subtracting absorbance at 383 nm. Cellular ATP concentrations in EPCs from CHx and Nx mice were determined by luciferine/luciferase luminometry according to the manufacturer's instructions (ATPLite; PerkinElmer Life and Analytical Sciences, Boston, http://www.perkinelmer.com).

EPC Function In Vivo

To induce limb ischemia, the left femoral artery and vein were excised in 8–10-week-old athymic BALB/cJ nude mice as described previously [18]. Perfusion was measured using a Lisca PIM II camera (Perimed, Stockholm, Sweden, http://www.perimed.se/) and expressed as the ratio of ischemic to normal limb perfusion. After 24 hours, animals with significant impairment of perfusion (Doppler signal ratio <25%) and without signs of severe necrosis were randomized to i.v. injection of 5 × 105 EPCs from Nx or CHx mice or saline vehicle (n = 10 per group). After 14 days, animals were euthanized, and adductor muscles were perfused with phosphate-buffered saline, fixed in Z-fix, and embedded in paraffin. Capillary density was determined in four HPFs after visualization of BS1 lectin (Sigma-Aldrich) with diaminobenzidine.

Statistical Analysis

All data are expressed as mean ± SEM. Normality of data was confirmed using the Kolmogorov-Smirnov test. Since circulating endothelial and circulating EPC numbers were not normally distributed, data from Nx and CHx mice were compared using a Wilcoxon rank-sum test. All other differences in EPC function between Nx and CHx mice were compared using a paired t test, and a p value <.05 was considered statistically significant. For evaluation of EPC transfer in vivo, a one-way analysis of variance was used to compare perfusion ratios or vascular densities among the three treatment groups.

Results

  1. Top of page
  2. Abstract
  3. Introduction
  4. Materials and Methods
  5. Results
  6. Discussion
  7. Conclusion
  8. Disclosure of Potential Conflicts of Interest
  9. Acknowledgements
  10. References
  11. Supporting Information

Bone Marrow Transplantation

To investigate whether bone marrow-derived cells participate in the pulmonary vascular remodeling response during chronic hypoxia, we evaluated the presence and distribution of GFP-expressing cells in the lungs of chimeric mice exposed to chronic hypoxia. We observed GFP-positive cells distributed throughout the lung parenchyma and predominantly associated with alveolar interstitium (Fig. 1A). Staining on 5 μm adjacent sections for the leukocyte common antigen (CD45) showed that most GFP-expressing cells coexpressed CD45 (Fig. 1B). When adjacent sections were stained for an endothelial marker (vWF) and GFP (Fig. 1C, left and right panels, respectively), we were not able to detect GFP-positive mature endothelial cells in remodeling pulmonary vessels, suggesting that bone marrow-derived endothelial cells do not play a predominant role in the pulmonary vascular remodeling response to hypoxia.

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Figure Figure 1.. Immunohistochemical analysis of green fluorescent protein (GFP), CD45, and von Willebrand factor (vWF) in consecutive lung sections from bone marrow chimeric mice exposed to chronic hypoxia. (A, B): GFP staining (A) and CD45 staining (B) on adjacent sections. Low-magnification images are shown on the left (scale bar = 100 μm), and high-magnification images are shown on the right. A round macrophage is shown in the blue square of (A), and cells with a more elongated phenotype are shown in the red square of (A). The corresponding area in (B) shows that these cells are CD45-positive. (C): Example of a lung section stained for vWF (left panel) and an adjacent section stained for GFP (right panel) (scale bars = 25 μm). Arrows indicate pulmonary blood vessels, and arrowheads indicate GFP-positive cells.

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Circulating EPC Numbers

To investigate whether lack of bone marrow-derived EPC engraftment in pulmonary resistance vessels was attributable to deficient mobilization during chronic hypoxia, we measured circulating EPCs using two alternative methods: flow cytometry of circulating EPCs in whole blood [19, 20] and spleen-derived mononuclear cell culture [21].

The percentage of CD45/VEGFR-2+/c-Kit+ cells in the peripheral blood was significantly higher in CHx mice, as determined by three-color flow cytometry (0.33% ± 0.10% of all nucleated cells vs. 0.07% ± 0.01% in Nx mice; n = 14; p < .001; supplemental online Fig. 1). Also, the percentage of CD45/VEGFR-2+ circulating endothelial cells, of which endothelial progenitor cells are a subset, was higher in CHx mice (0.53% ± 0.12% vs. 0.21% ± 0.05% in Nx mice; n = 14; p < .001). The total number of circulating white blood cells did not differ between CHx and Nx mice (data not shown).

In addition, spleen-derived MNCs from both Nx and CHx mice, cultured for 7 days, express VEGFR-2, Sca-1, CXCR-4, and moderate levels of c-Kit (Fig. 2A), with more than 95% of these early-outgrowth EPCs being positive for acLDL and BS1 lectin, regardless of whether EPCs were derived from Nx or CHx mice. The number of acLDL/BS1 lectin double-positive EPCs per mm2 was significantly increased in CHx mice (220 ± 49 vs. 106 ± 25 EPCs per mm2 in Nx mice; n = 7; p < .01; Fig. 2B). The total number of EPCs (taking into account the number of MNCs in the spleen) was also higher in CHx mice (352 ± 87 vs. 240 ± 67 × 103 EPCs per mouse in Nx mice; n = 7; p < .05). Taken together, flow cytometry and cell culture experiments indicate that failure of EPCs to integrate in hypertensive lung vessels is not caused by impaired EPC mobilization during chronic hypoxia.

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Figure Figure 2.. Characterization of cultured endothelial progenitor cells (EPCs). (A): Expression analysis using flow cytometry: 7-day-old EPCs expressed VEGFR-2 (upper left panel), Sca-1 (upper right panel), c-Kit (lower left panel), and CXCR-4 (lower right panel). EPCs incubated with antibody are shown in gray, and isotype controls are shown in black. (B): Representative pictures of EPCs from a normoxic mouse (left panel) or a chronic hypoxic mouse (right panel) stained with acLDL (red) and BS1 lectin (green). Scale bar = 100 μm. Abbreviations: acLDL, acetylated low-density lipoprotein; VEGFR-2, vascular endothelial growth factor receptor-2.

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EPC Transfer in PH Mice

We next investigated whether defective homing of endogenous EPCs to hypertensive lungs could be overcome by direct in vivo cell transfer of culture-expanded early-outgrowth Nx EPCs. Cell tracking experiments using luciferase-expressing EPCs and in vivo bioluminescence imaging demonstrated that luciferase activity was clearly detectable in lungs up to 12 hours after EPC transfer. However, 24 hours after transfer, EPCs were no longer detectable in the lungs, and after 7 days, luciferase activity was detected exclusively in the spleen (Fig. 3A, 3B). Because the sensitivity of in vivo bioluminescence imaging does not allow detection of low amounts of incorporated cells, we also performed transfer of DiI-labeled EPCs and observed only a small number of EPCs around precapillary pulmonary resistance vessels 1 or 2 weeks after transfer into pulmonary hypertensive animals (Fig. 3C), whereas the majority of EPCs were trapped in the spleen (Fig. 3C). Transplanted EPCs reside for only a limited time in hypertensive lungs, which may relate to their failure to mitigate PH (Table 1). Moreover, EPC transfer in splenectomized mice equally failed to reduce hypoxic PH (data not shown). To address the deficient homing of EPCs in vivo, we determined intercellular adhesion molecule-1 (ICAM-1) expression levels by immunoblot and SDF-1α expression levels by enzyme-linked immunosorbent assay. Although SDF-1α was significantly higher in lungs from CHx mice (131 ± 13 vs. 72 ± 11 pg of SDF-1α per mg of protein in Nx lungs; p < .01; n = 9 in each group), we did not observe upregulation of ICAM-1 expression in CHx lungs (n = 6 in each group; supplemental online Fig. 2).

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Figure Figure 3.. Homing of transplanted and labeled early-outgrowth EPCs in lungs and spleen of chronic hypoxic mice. (A): Bioluminescence imaging after i.v. injection of luciferase-positive early-outgrowth EPCs. At each time point, a control mouse (without transplanted EPCs, shown on the left side of each panel) was scanned, together with a mouse receiving EPCs (shown on the right). Two hours after EPC transfer, the majority of EPCs were localized in the lungs. After 1 day, there was no detectable luciferase signal in the lungs, as EPCs are distributed throughout the systemic circulation and this is below the sensitivity of in vivo detection. After 7 days, marked bioluminescence was detected in the left subcostal region, indicating EPC homing to the spleen. (B): Kinetic analysis of bioluminescence imaging. There was clear luciferase activity in the lungs 2–6 hours after i.v. EPC transfer, but 12 hours after injection, this signal started to decrease, and after 1 day, luciferase activity was almost absent in the lungs. In contrast, there was no signal in the spleen immediately after EPC transfer, but after 7 days, majority of luciferase activity was present in the spleen (n = 7 mice). (C): Using fluorescence microscopy, we were able to detect few DiI-labeled EPCs in the vicinity of small pulmonary blood vessels 1 week after i.v. injection (endothelial cells stained with von Willebrand factor are shown in blue, and DiI-labeled EPCs are shown in red). Scale bar = 50 μm. (D): However, most transplanted EPCs were found in the spleen (nuclear 4′,6-diamidino-phenylindole staining of splenocytes is shown in blue, and DiI-labeled EPCs are shown in red). Scale bar = 25 μm. Abbreviations: d, days; EPC, endothelial progenitor cell; h, hours; p, photons; ROI, region of interest; s, second; sr, steradian.

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Table Table 1.. Hemodynamic and morphometric analysis after EPC transfer in chronically hypoxic mice
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EPC Functional Characterization During Chronic Hypoxia

Migration.

In ischemic tissues, SDF-1α is induced and promotes the recruitment of CXCR-4-positive progenitor cells [22]. The ability of EPCs to migrate toward SDF-1α was therefore investigated using a modified Boyden chamber (Fig. 4A). EPCs from CHx mice were less responsive to SDF-1α compared with EPCs from Nx mice (99 ± 28 vs. 169 ± 27 migrating EPCs per HPF, respectively; n = 7; p < .05). In contrast, flow cytometry revealed no significant difference in CXCR-4 expression (mean fluorescence intensity of CHx EPCs was 110 ± 26 vs. 91 ± 20 arbitrary units in Nx EPCs; n = 7; p = not significant).

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Figure Figure 4.. Effect of chronic hypoxia on endothelial progenitor cell (EPC) migration and tube formation. (A): Representative membrane inserts stained with 4′,6-diamidino-phenylindole after migration of EPCs from a normoxic (Nx) (left panel) or a chronic hypoxic (CHx) mouse (right panel). (B): Tube formation with DiI-labeled EPCs coplated with human umbilical vein endothelial cells on Matrigel. Left panel shows EPCs derived from an Nx mouse, right panel from a CHx mouse. Scale bar = 200 μm. Inset shows a high-magnification view of incorporating EPCs.

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Adhesion to Fibronectin and Incorporation into a Vascular Network.

Chronic hypoxia caused a 29% decrease in adhesive capacity of EPCs to fibronectin, an important extracellular matrix component involved in the initial steps during vasculogenesis (52 ± 6 vs. 73 ± 10 adhering EPCs per HPF from Nx mice; n = 7; p < .05). Similarly, incorporation into a developing vascular network in vitro mimics in vivo incorporation of EPCs during vasculogenesis. To study the effect of chronic hypoxia on vasculogenic potential of EPCs, we plated DiI-labeled EPCs on Matrigel in the presence of HUVECs and determined incorporation after overnight incubation (Fig. 4B). The total number of incorporating EPCs per HPF was lower for EPCs derived from CHx mice (46 ± 3 vs. 66 ± 6 incorporating EPCs per HPF from Nx mice; n = 7; p < .05).

NO Production.

To investigate whether chronic hypoxia influences the ability of EPCs to release NO, a downstream effector in VEGF-induced angiogenesis, we measured NO concentrations in 24-hour-conditioned medium. Although NOS3 mRNA expression did not differ between CHx and Nx EPCs (supplemental online Table 2), NOS activity was significantly impaired, as indicated by the approximately 30% lower NOx concentrations in the supernatants of EPCs derived from CHx mice compared with Nx EPCs (2.3 ± 0.4 vs. 3.4 ± 0.4 nmol per 106 EPCs, respectively; n = 9; p < .05). These concentrations are similar to concentrations released from HUVECs (4.6 ± 0.7 nmol per 106 HUVECs) and the murine endothelial MS1 cell line (3.9 ± 1.0 nmol per 106 cells) under identical conditions. NOx concentrations are reduced approximately 80% by the NOS inhibitor l-NMA.

Hind Limb Ischemia.

To investigate whether or not the dysfunctional phenotype of CHx EPCs limits vascular repair capacity in vivo, we performed EPC transfer in mice with hind limb ischemia. Whereas EPCs derived from Nx mice significantly improved blood flow recovery in the ischemic hind limb compared with saline (recovery to 69% ± 2% of flow in the nonoperated limb; n = 10; p < .01), an equal number of EPCs from CHx mice failed to do so (Fig. 5A, 5B). Correspondingly, capillary density in the adductor muscle was significantly higher in mice receiving Nx EPCs than in mice receiving CHx EPCs or saline (Fig. 5C, 5D).

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Figure Figure 5.. Blood flow recovery and capillary density 2 weeks after hind limb ischemia with or without EPC cell transfer. (A): Representative laser Doppler perfusion images 14 days after hind limb ischemia and transfer of EPCs from Nx mice (left panel), CHx mice (middle panel), or saline (right panel). (B): Relative perfusion as a percentage of the contralateral limb in mice following injection of EPCs from Nx or CHx mice or saline. (C): Representative sections of adductor muscles stained with BS1 lectin (brown) from mice treated with EPCs from Nx or CHx mice or saline. Scale bar = 50 μm. (D): Capillary densities (capillaries per mm2) in adductor muscle following EPC transfer from Nx or CHx mice or saline. Abbreviations: CHx, chronic hypoxic; EPC, endothelial progenitor cell; N.S., not significant; Nx, normoxic.

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Characterization of Mechanisms Contributing to Hypoxia-Induced EPC Dysfunction

Integrin Expression Levels.

To investigate potential molecular mechanisms for reduced adhesive capacity, we measured integrin expression levels of CHx and Nx EPCs. As shown in supplemental online Table 2, integrin αv and β1 transcript levels were significantly lower in EPCs derived from CHx compared with Nx mice (−61% and −44% respectively; p < .05 for both). The expression of integrins α5, β2, and β3 was not significantly affected by chronic hypoxia.

Senescence-Associated β-Galactosidase Staining.

To explore whether chronic hypoxia induces senescence of EPCs, we compared senescence-associated β-galactosidase staining of EPCs cultured from Nx and CHx mice. The percentage of dark blue-stained EPCs, indicative of a higher content of the lysosomal hydrolase β-galactosidase, was significantly higher in EPCs from CHx mice (9.6% ± 1.4%) compared with EPCs from Nx mice (5.2% ± 1.4%; n = 7; p < .01; Fig. 6A).

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Figure Figure 6.. Senescence and mitochondrial dysfunction in endothelial progenitor cells (EPCs). (A): Senescent, β-galactosidase-expressing cells are dark blue (arrows) and were more predominant in EPCs from chronic hypoxic (CHx) mice (right panel) compared with EPCs from normoxic (Nx) mice (left panel). Scale bars = 50 μm. (B): JC-1 staining of EPCs from an Nx mouse (left panel) and a CHx mouse (right panel). Arrows indicate normally polarized mitochondria (red), and arrowheads point to cells with depolarized mitochondria (green + red = yellow). Scale bar = 25 μm.

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Mitochondrial Membrane Potential Analysis and Cellular ATP Content.

To evaluate mitochondrial membrane potential of EPCs, an important rheostat in determining EPC survival and function, we compared JC-1 aggregate fluorescence in EPCs cultured from Nx or CHx mice. Ratiometric analysis of JC-1 aggregated forms (red fluorescence) versus the monomeric form (green fluorescence) indicated a 30% lower ratio in CHx EPCs (0.61 ± 0.08 vs. 0.86 ± 0.12 in Nx EPCs; n = 10; p < .05; Fig. 6B). In EPCs treated with the uncoupler FCCP, this ratio decreased to 0.30 for both Nx and CHx EPCs. Since anaerobic ATP synthesis is coupled to the formation of lactate, we measured lactate concentrations in conditioned medium from CHx and Nx EPCs. CHx EPCs did not upregulate glycolysis for enhanced ATP generation, as lactate production was not increased in CHx EPCs (3.0 ± 0.8 vs. 5.2 ± 1.1 μmol per 106 Nx EPCs; n = 6; p = .11). Total cellular ATP concentrations were 10% lower in EPCs from CHx mice (1.99 ± 0.21 vs. 2.17 ± 0.24 μmol per 106 EPCs; n = 9; p < .05).

Discussion

  1. Top of page
  2. Abstract
  3. Introduction
  4. Materials and Methods
  5. Results
  6. Discussion
  7. Conclusion
  8. Disclosure of Potential Conflicts of Interest
  9. Acknowledgements
  10. References
  11. Supporting Information

The critical role of endothelial dysfunction in many forms of pulmonary hypertension has spurred increased interest in bone marrow-derived EPCs, not only as pathophysiologic modulators but also as potential tissue-engineering tools in PH. We report here that chronic hypoxia, a common stimulus in both experimental and human pulmonary hypertension, significantly alters mobilization and functional characteristics of early-outgrowth endothelial progenitor cells. Our experiments using GFP bone marrow chimeric mice clearly indicate the failure of a reparative endogenous EPC response, as virtually no endogenous bone marrow-derived cells were associated with the remodeling vasculature. Moreover, cell transfer of culture-expanded normoxic early-outgrowth EPCs failed to reverse or mitigate the hypoxia-induced hypertensive phenotype because of defective homing and incorporation into the hypertensive pulmonary vasculature.

During prolonged hypoxia/hypoxemia, greater numbers of EPCs are mobilized into peripheral blood, as determined by flow cytometry (CD45/VEGFR-2+/c-Kit+ MNCs) and cell culture (acetylated LDL/BS1 double-positive cells). However, EPCs from these hypoxic mice demonstrated reduced migration toward SDF-1α, a critical ligand for EPC homing; reduced adhesive capacity to the subendothelial extracellular matrix component fibronectin (a first necessary step in vascular integration); and reduced angiogenic potential and NO release. These data suggest a sustained effect of chronic hypoxia on the functionality of early-outgrowth EPCs and have important therapeutic consequences. Indeed, functional impairment of hypoxic EPCs was also obvious in a well-standardized in vivo neovascularization assay, where CHx EPCs failed to improve perfusion in the ischemic hind limb model, contrary to identically cultured Nx EPCs. Taken together, these data provide evidence that the hypoxic milieu significantly impairs the tissue repair capacity of EPCs.

Our data of EPC function in chronic hypoxia are in marked contrast with the traditional view of the vasculoprotective or reparative role of EPCs at sites of vascular injury. EPC transfer accelerates re-endothelialization in the denuded vessel wall, improves endothelium-dependent vasorelaxation, and reduces neointima formation in rabbits [23] and mice [3]. EPCs also appear to have a promising tissue repair capacity in monocrotaline-induced PH in rats [8, 9]. Contrary to the transient engraftment in precapillary vessels in this inflammatory model of PH, we failed to observe a similar endogenous homing response of EPCs to the remodeling hypoxic pulmonary circulation.

The lack of an endogenous tissue repair response was not attributable to impaired mobilization of EPCs from the bone marrow compartment to the pulmonary circulation, as evidenced by the significant increase in circulating EPC numbers during chronic hypoxia. Moreover, i.v. transfer of fully functional Nx EPCs also failed to incorporate in remodeling vessels and to reduce PH. These observations suggest that either the extent of pulmonary endothelial damage or activation is insufficient to elicit effective engraftment of EPCs or, alternatively, that EPCs become dysfunctional during sustained exposure to chronic hypoxia and lose their ability to adhere and transmigrate.

To investigate the former possibility, we compared ICAM-1 and SDF-1α expression in CHx and Nx lungs. The interactions between integrin β2 on EPCs and ICAM-1 in vascular tissue and between CXCR-4 on EPCs and SDF-1α in tissue are important determinants for EPC homing [22, 24, [25]26]. We observed increased SDF-1α generation but no upregulation of ICAM-1 in CHx lungs compared with Nx lungs. This contrasts with the significant upregulation of ICAM-1 in ischemic skeletal muscles [26]. These findings suggest that failure to upregulate ICAM-1 expression might contribute to the impaired homing and incorporation of circulating EPCs in chronic hypoxia.

To test the latter hypothesis (that EPCs become dysfunctional during sustained exposure to CHx), we performed several in vitro functionality assays, corresponding to essential steps during vasculogenesis: migration of chronic hypoxia EPCs toward the chemokine SDF-1α, a critical ligand for EPC homing, was impaired; adhesion to the important extracellular matrix component fibronectin was decreased; and the cells were less likely to incorporate into a developing vascular network. In addition, EPCs from CHx mice showed reduced NO release. Importantly, the dysfunctional phenotype did not result from specific in vitro conditions or tissue culture artifacts during cell expansion, as simultaneously expanded Nx EPCs significantly improved neovascularization after hind limb ischemia.

The exact mechanisms of EPC dysfunction remain unknown, despite extensive studies in atherosclerosis and ischemic heart disease. Persistent dysfunction after 7 days of in vitro culture has also been described for EPCs derived from type I diabetics (reduced secretion of angiogenic growth factors) [27], type II diabetics (reduced incorporation into a developing vascular network) [5], and patients with chronic renal failure (reduced migration and incorporation into a developing vascular network) [28]. Moreover, a higher percentage of EPCs from patients with cardiovascular risk factors are senescent compared with EPCs from healthy persons after 7 days of culture [29]. In contrast to the extensive description of EPC dysfunction in cardiovascular diseases, animal models of EPC dysfunction are limited. EPCs from cathepsin L knockout mice have impaired in vivo neovascularization capacity, yet they preserved migration and growth factor secretion [30]. In contrast, EPCs from glutathione peroxidase 1-deficient mice incorporate less in a vascular network and have an impaired migratory response [31]. In our study, we have characterized EPC dysfunction associated with chronic hypoxia, which is relatively easy to use and well standardized in a genetically homogeneous population of inbred mice.

Our data indicate that chronic hypoxia is a useful experimental condition to investigate basic mechanisms of EPC dysfunction, including altered integrin expression, enhanced senescence, and perturbed energy metabolism. First, integrins regulate important cellular interactions of progenitor cells, including transendothelial migration toward SDF-1α (integrin β1) [32], bFGF- or VEGF-induced angiogenesis (integrin αv) [33], and retention of progenitor cells in the bone marrow compartment (integrin β1 via very-late antigen 4 or integrin α4β1) [34]. We observed significantly reduced expression levels of αv and β1 integrins in CHx EPCs, which contribute to reduced adhesion to fibronectin and migration toward SDF-1α and to increased circulating EPC numbers. In contrast, NOS3 expression was not different between Nx and CHx EPCs, whereas NOS activity was impaired, which is in accordance with previously published results of unaltered expression but impaired post-transcriptional activation during chronic hypoxia [35]. Second, the impaired function of CHx EPCs may be attributable in part to enhanced senescence, as evidenced by a twofold increase in the percentage of β-galactosidase-positive EPCs. Other environmental risk factors, including smoking, have been shown to promote a similar degree of EPC senescence [29]. Finally, the ratiometric analysis of the mitochondrial membrane potential in Nx and CHx EPCs demonstrated a 30% reduction in transmembrane potential. A similar depolarization of the mitochondrial membrane has recently been documented in isolated cells exposed to ischemic preconditioning [36, 37] and is thought to represent a cell defense mechanism against a lethal insult. Indeed, cellular ATP levels in EPCs from CHx mice were only moderately reduced despite the significant loss of mitochondrial membrane potential, which is normally required for maintaining respiratory ATP generation. Preservation of cellular ATP concentrations could be accounted for by previously described adaptive survival mechanisms of cells in response to chronic hypoxia in vitro and moderate ischemia in vivo (i.e., downregulation of ATP requiring processes, including ion pump activity and protein synthesis, to induce a hypometabolic state [38, [39], [40]41]). Our results are therefore consistent with the view that EPCs from chronically hypoxic mice are less metabolically active and consume less ATP, which in turn could account for their dysfunctional phenotype.

Several limitations of our study should be mentioned. First, we used a standardized study protocol using spleen-derived EPCs and cannot rule out differential in vitro behavior of spleen-derived EPCs versus circulating EPCs. The latter are routinely studied in patients, but limited numbers preclude a similar approach in mice. Second, although our molecular and cellular studies clearly establish altered integrin expression, senescence, and mitochondrial function in EPCs from CHx mice, the intracellular downstream signaling pathways remain to be determined. Third, although hypoxia-induced EPC dysfunction represents a useful paradigm to develop improved cell-based therapeutic strategies, we need to recognize that multiple, complex disease pathways likely cooperate in patients.

Conclusion

  1. Top of page
  2. Abstract
  3. Introduction
  4. Materials and Methods
  5. Results
  6. Discussion
  7. Conclusion
  8. Disclosure of Potential Conflicts of Interest
  9. Acknowledgements
  10. References
  11. Supporting Information

Chronic hypoxia increases the number of bone marrow-derived circulating EPCs, which have a critically impaired function and engraft poorly in hypertensive lung vessels. Transplantation of normoxic early-outgrowth EPCs in a hypoxic milieu does not result in effective cell-mediated repair in the hypertensive lung, because of insufficient homing and incorporation. Reduced integrin expression, enhanced senescence, and decreased mitochondrial membrane potential contribute to the dysfunctional EPC phenotype. Our data may have important implications for early-outgrowth EPC transfer protocols in PH, myocardial infarction, or stroke, since these EPCs will similarly be exposed to reduced oxygen tensions. Our observations in chronic hypoxia may therefore help to develop innovative targeted strategies to improve early-outgrowth EPC function for tissue repair.

Acknowledgements

  1. Top of page
  2. Abstract
  3. Introduction
  4. Materials and Methods
  5. Results
  6. Discussion
  7. Conclusion
  8. Disclosure of Potential Conflicts of Interest
  9. Acknowledgements
  10. References
  11. Supporting Information

We thank Christopher Contag and Yu-An Cao (Molecular Biophotonics and Imaging Laboratory, Stanford University) for providing luciferase-expressing L2G85 mice and Julian Aragones Lopez and Peter Fraisl (Center for Transgene Technology and Gene Therapy, VIB, K.U. Leuven) for scientific assistance and discussion. Research was funded by a Ph.D. grant of the Institute for the Promotion of Innovation through Science and Technology in Flanders (IWT-Vlaanderen) (to G.M.). S.J. is a Clinical Investigator of the Fund for Scientific Research-Flanders and holder of a chair in cardiology sponsored by AstraZeneca.

References

  1. Top of page
  2. Abstract
  3. Introduction
  4. Materials and Methods
  5. Results
  6. Discussion
  7. Conclusion
  8. Disclosure of Potential Conflicts of Interest
  9. Acknowledgements
  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. Disclosure of Potential Conflicts of Interest
  9. Acknowledgements
  10. References
  11. Supporting Information
FilenameFormatSizeDescription
SC-07-0562_Supplemental_Figure_1.eps1607KSupplemental Figure 1
SC-07-0562_Supplemental_Figure_2.eps1376KSupplemental Figure 2

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