CD34−/CD133+/VEGFR-2+ Endothelial Progenitor Cell Subpopulation With Potent Vasoregenerative Capacities
Our goal was to identify functionally important subpopulations within the heterogenous group of endothelial progenitor cells (EPC). Fluorescence-activated cell sorter analysis of CD133+ progenitor cells revealed the presence of CD34+ and CD34− subpopulations. CD34−/133+ progenitors differentiate into CD34+/133+ EPC, adhere more potently than these in response to SDF-1, and rapidly home to sites of limb ischemia in human volunteers. In human coronary atherectomy samples, fewer CD34−/133+ than CD34+/133+ EPC are present in stable plaques, whereas cell numbers increase with a reversion of the ratio in unstable lesions. In CD34−/133+ EPC-injected nude mice, more transplanted cells coexpressing endothelial markers home to carotid artery lesion endothelium than in CD34+/133+-injected mice. In the former, lesions were smaller and reendothelialization higher than in the latter. We identified a new CD34−/133+ EPC subpopulation, which is apparently a precursor of “classical” CD34+/133+ EPC, and functionally more potent than these with respect to homing and vascular repair.
The prevailing dogma, that in adults the formation of new blood vessels occurs only by migration and proliferation of mature endothelial cells in a process termed angiogenesis, was overturned in recent years by the discovery of endothelial progenitor cells (EPC), which differentiate into endothelial cells in a process referred to as vasculogenesis.1,2 In settings of tissue ischemia or endothelial damage, EPC have been shown to be mobilized from the bone marrow into the circulation, home to sites of injury, and incorporate into foci of neovascularization thereby improving blood flow and tissue recovery.3–7 EPC are a heterogeneous group of cells that can be characterized by the expression of surface markers, such as CD34, CD133, and VEGFR-2 (KDR or Flk-1), by the uptake of DiI-Ac-LDL and by binding lectins such as Ulex europaeus, which are thought be endothelial specific. Expression of the stem cell marker CD34 is also found on a lower level on mature endothelial cells, and the search for more specific stem cell markers led to the discovery of CD133 (also termed AC133), which is highly expressed on immature stem cells but whose expression is lost during the differentiation to mature endothelial cells.8–11 Therefore, EPC are thought to be a heterogenous population of progenitors consisting of a more primitive CD133+/CD34+/VEGFR-2+ subpopulation, and a more mature CD133−/CD34+/VEGFR-2+ subpopulation.
The aim of this study was to identify new subpopulations within the diverse EPC population by analysis of surface marker expression patterns and to phenotypically and functionally characterize these progenitors in vitro and in vivo.
Materials and Methods
Ethical Study Approval
All human studies on blood samples obtained under baseline conditions or following limb ischemia as well as on coronary atherectomy specimen were approved by local ethics committees. Written informed consent was obtained from all study subjects (healthy human volunteers and patients undergoing cardiac catheterization for coronary artery disease). Human studies were conducted and human samples were acquired in accordance with institutional guidelines. All animal experiments were approved by local ethics committees and conducted in accordance with institutional guidelines.
Isolation and Culture of Circulating Human EPC Subpopulations
Human mononuclear cells were isolated from buffy-coat layers obtained during erythrocytapheresis or from peripheral blood of healthy volunteers (n=10, age 29±5 years, 6 male, 4 female) by density gradient centrifugation with Biocoll (Biochrom) as previously described.5–7 All blood samples were obtained, processed, and analyzed individually in independent experiments. CD34+/CD133+ and CD34−/CD133+ cells were isolated by 2-step magnetic bead-purification according to the instructions of the manufacturer (Miltenyi Biotec). Purity was determined by fluorescence-activated cell sorter (FACS) analysis and was routinely >96%. When indicated, cells were cultured on fibronectin-coated dishes in endothelial basal medium (Clonetics Cell Systems) supplemented with endothelial growth medium SingleQuots and 20% FCS as previously described.5–7 After 1 and 4 days in culture, adherent cells were incubated with DiI-Ac-LDL (CellSytems) and stained with fluorescein isothiocyanate (FITC)-labeled U europaeus lectin (10 μg/mL; Sigma), as previously described.5–7
Human Coronary Atherectomy Samples
Specimens were obtained from patients scheduled for percutaneus coronary intervention during cardiac catheterization resulting from stable angina (n=6, age 71±10 years, 5 male, 1 female; vascular risk profile: hyperlipidemia 4/6, smoking 2/6, familial predisposition 2/6, arterial hypertension 3/6, diabetes mellitus 1/6), unstable angina (n=6, age 61±14 years, 4 male, 2 female; vascular risk profile: hyperlipidemia 4/6, smoking 3/6, arterial hypertension 5/6), or in-stent restenosis (n=6 per group, age 59±12 years, 5 male, 1 female; vascular risk profile: smoking 4/6, arterial hypertension 3/6, diabetes mellitus 1/6, familial predisposition 2/6). In accordance with actual guidelines from the American Heart Association and the German Cardiac Society, stable angina was defined as reproducible, exercise-induced, troponin-negative chest pain resolving at rest; unstable angina, as troponin-positive chest pain occurring under resting conditions. Samples were fixed with 4% paraformaldehyde, embedded in Tissue Tek OCT embedding medium (Miles Laboratories), snap-frozen, and stored at −80°C. Samples were sectioned on a Leica cryostat (7 μm) and placed on poly-l-lysine-coated slides (Sigma) for immunohistochemical analysis of 30 sections per individual as explained below under Immunohistochemistry.
Human Limb Ischemia Model
Healthy human volunteers (n=5, age 28±5 years, 3 male, 2 female) were subjected to forearm ischemia by inflating a cuff to 20 mm Hg over the diastolic blood pressure for 10 minutes, so that the radial artery pulse was still weakly palpable. Sixty milliliters of blood from the ipsilateral brachial vein draining this ischemic territory were drawn into citrated tubes at this arm before the induction of ischemia (baseline), immediately after deflating the cuff (0-minute time point), and 2 hours later. All blood samples were obtained, processed, and analyzed individually in independent experiments.
EPC were labeled with PKH-26 using the PKH26-GL Red Fluorescent Cell Linker Kit according to the instructions of the manufacturer (Sigma). Nude mice (n=5 per condition; Charles River Laboratories, Sulzfeld, Germany) were transplanted with 1×105 of human EPC subsets (CD34−/CD133+, CD34+/CD133+) resuspended in 0.5 mL of 1× PBS or saline control via intravenous tail vein injection directly after induction of arterial injury, as previously described.6
Carotid Artery Injury Model
Carotid artery injury was induced as described in detail previously.5–7 Following perfusion–fixation with 4% paraformaldehyde, carotid arteries were harvested 14 days after surgery, embedded in Tissue Tek OCT embedding medium (Miles Laboratories), snap-frozen, and stored at −80°C. Samples were sectioned on a Leica cryostat (7 μm) and placed on poly-l-lysine-coated slides (Sigma) for immunohistochemical and morphometric analysis. The former was performed as explained in detail below under Immunohistochemistry. For the latter, hematoxylin/eosin (H&E) staining was performed according to standard protocols, and Lucia Measurement software version 4.6 was used to measure external elastic lamina, internal elastic lamina, neointimal area, and lumen circumference of 25 sections per animal, as previously described.5–7
Disc Angiogenesis Model
Nude mice were subcutaneously implanted with discs of polyvinyl alcohol sponges (Rippey) covered with nitrocellulose cell-impermeable filters (Millipore), allowing capillary growth only through the rim of the discs as previously described.7 Discs were allowed to heal in for 10 days before carotid injury and EPC transplantation. Fourteen days later, space-filling fluorescent microspheres (0.2 μm; Molecular Probes) were injected into the left ventricle to systemically deliver them to the microvasculature. Lucia Measurement software 4.6 was used to assess the disc area invested by fibrovascular growth.
Fluorescence-Activated Cell Sorter Analysis
FACS analysis was performed as described previously in detail.5–7 Viable human EPC were analyzed for CD133-PE (Miltenyi), CD34-APC (Pharmingen), CXCR4-FITC (Pharmingen), CD14-PE (Pharmingen), and VEGFR-2 (Dianova, Hamburg, Germany). The latter was detected with a biotin-conjugated secondary antibody and PerCP-labeled streptavidin (both Pharmingen). Isotype-matched antibodies served as controls in every experiment (Pharmingen).
Tissue sections were stained as previously described in detail.5–7 Human coronary artery cryosections were analyzed for CD133 (Miltenyi) and CD34 (Pharmingen). Murine carotid artery cryosections were analyzed for PKH-fluorescence (Sigma) and CD31 (Pharmingen). Isotype-matched antibodies served as controls (Pharmingen). Sections were counterstained with 4′,6-diamidino-2-phenylindole (DAPI) (Linaris) and mounted with fluorescent mounting medium (Dako) for fluorescent microscopic analysis using a Nikon E600 microscope and Lucia Measurement software 4.6. For each individual patient from which a coronary atherectomy sample was obtained, cells staining triple-positive for CD133, CD34, and DAPI were counted per high-power field (×40) on 30 sections before calculating the corresponding means and summarizing the data over multiple individuals in each group. For each murine carotid artery sample, reendothelialization of injured mice was quantified by counting the total number of CD31+/DAPI+ double-positive cells per ×10 field of 25 sections per animal and expressed as percentage of total number of CD31+/DAPI+ double-positive cells of uninjured controls. In the xenotransplantation model, homing of transplanted stem cells was quantified by counting the number of PKH+/CD31+/DAPI+ triple-positive cells per ×10 field of 25 sections per animal and expressed as percentage of total number of CD31+/DAPI+ double-positive cells along the lumen circumference.
Human EPC (1×105 per condition) were allowed to adhere on fibronectin (Sigma)-coated culture plates for 5 minutes in presence or absence of 100 nmol/L of stromal-derived factor-1 (SDF-1; R&D Systems). Dishes were vigorously washed 3 times with changes of 1× PBS, and adherent cells were counted using a Nikon TS 100 microscope.
Data are presented as the mean±SD. Paired and unpaired Student’s t tests and the Newman–Keuls test for multiple comparisons were used where applicable. The null hypothesis was considered to be rejected at P<0.05.
CD34−/ CD133+/VEGFR-2+ Progenitor Cells Are Precursors of CD34+/CD133+/VEGFR-2+ EPC
With the aim to further characterize the heterogeneous group of EPC, we isolated CD133+ progenitor cells from peripheral blood of healthy human volunteers by means of magnetic bead separation. As seen in Figure 1A, FACS analysis revealed the presence of two subpopulations within this cell fraction: a larger CD34+/CD133+ (69±3% of gated cells) and a smaller CD34−/CD133+ (29±3%) subpopulation. Both of these subpopulations express equal surface levels of VEGFR-2 (>99%) and lack expression of the monocyte/macrophage marker CD14. When these progenitor subpopulations were cultured up to 4 days under conditions favoring endothelial-specific differentiation, CD34−/CD133+/VEGFR-2+ cells progressively downregulated surface expression of CD133, while upregulating CD34 and the endothelial marker CD31 (Figure 1B). These data place CD34−/CD133+/VEGFR-2+ progenitors in the endothelial lineage differentiation at an earlier time point than CD34+/CD133+/VEGFR-2+ cells. Additionally, we analyzed uptake of DiI-Ac-LDL and binding of U europaeus lectin and found that a higher number of CD34−/CD133+/VEGFR-2+ cells stain double-positive for these endothelial-specific markers as compared with CD34+/CD133+/VEGFR-2+ cells (Figure 1C). In contrast to the latter progenitor subpopulation, double-positive staining in the CD34−/CD133+/VEGFR-2+ cells is already evident after only 24 hours of culture, suggesting a higher potential to differentiate into mature endothelial cells that might be attributable to higher adhesive capacity. Therefore, we next performed adhesion assays, which showed that CD34−/CD133+/VEGFR-2+ cells display higher levels of SDF-1–triggered β1-integrin–dependent adhesion as compared with CD34+/CD133+/VEGFR-2+ cells (Figure 1D).
CD34−/CD133+/VEGFR-2 EPC Home to Sites of Ischemia and Are Upregulated in Human Unstable Coronary Lesions
To study EPC trafficking, healthy human volunteers were subjected to limb ischemia and homing of EPC subpopulations was studied 10 to 120 minutes postischemia. As seen in Figure 2A, expression levels of CD34+/CD133+ EPC progressively decreased, whereas expression levels of CD34−/CD133+ EPC increased in parallel, possibly indicating increased homing of the latter more primordial EPC subpopulation to these sites. Expression levels of VEGFR-2 were higher in CD34+/CD133+ progenitors, whereas surface expression of the SDF-1 receptor CXCR4 was consistently higher in CD34−/CD133+ cells (Figure 2B). The latter fact might explain the more potent adhesive response of the latter progenitor subpopulation. There was no significant difference in the expression levels of VEGFR-2 or CXCR4 between baseline and the 10- and 120-minute time points. We next analyzed human coronary atherectomy specimen from stable lesions, unstable lesions, and in-stent restenosis for expression of CD34−/CD133+ and CD34+/CD133+ EPC. As seen in Figure 2C, we detected both progenitor subpopulations in stable plaques with a higher number of CD34+/CD133+ EPC. In unstable lesions, both stem cell subsets were upregulated with a higher number of CD34−/CD133+ EPC. In contrast, no expression of either EPC subset was detectable in in-stent restenosis.
Transplanted CD34−/CD133+/VGFR-2+ EPC Promote Reendothelialization and Reduction of Lesion Size Following Vascular Injury
To study the biological relevance of EPC subpopulations, nude mice with implanted angiogenesis disks were subjected to carotid artery injury and transplanted with equal numbers of PKH-labeled CD34−/CD133+/VEGFR-2+ cells, CD34+/CD133+/VEGFR-2+ cells, or control. As seen in Figure 3 (A and D), in control animals, arterial injury resulted in a prominent neointima formation with a significant reduction of lumen circumference as compared with noninjured controls. In recipients of CD34+/CD133+/VEGFR-2+ EPC, neointima formation was reduced and lumen circumference increased as compared with control. Transplantation of CD34−/CD133+/VEGFR-2+ EPC resulted in an even greater suppression of neointima formation and further augmented lumen circumference. In recipients of EPC subpopulations, PKH+ endothelial cells were detected in the endothelial layer of lesions (Figure 3B, 3C, and 3E). However, in animals transplanted with CD34−/CD133+/VEGFR-2+ EPC more cells homed to lesions, incorporated into the endothelium, and more potently promoted reendothelialization as compared with recipients of CD34+/CD133+/VEGFR-2+ EPC. As seen in Figure 3F, both EPC subpopulations equipotently augmented neoangiogenesis.
This study demonstrates the existence of a peripheral blood–derived CD34−/CD133+ stem cell population, which differentiates into CD34+/CD133+ EPC and then acquires a mature endothelial phenotype. However, this CD34−/CD133+ EPC subpopulation is functionally more active than the supposedly more mature CD34+/CD133+ EPC subpopulation. In vitro, this progenitor subset displays higher levels of SDF-1–triggered adhesion caused by higher expression of CXCR4 and more potently differentiates into mature endothelium as compared with CD34+/CD133+ EPC. In vivo, CD34−/CD133+ EPC are upregulated in unstable human coronary lesions and rapidly home to sites of ischemia and vascular injury, where they more potently promote endothelial regeneration and lesion reduction than CD34+/CD133+ EPC.
Since the initial report by Asahara et al,1 EPC were thought to be defined by surface expression of CD34 and VEGFR-2. The discovery of the stem cell marker CD133,8,9 whose expression is lost as EPC mature to endothelial cells in contrast to CD34 expression, allowed further categorization of the EPC population into a CD133+/CD34+/VEGFR-2+ subpopulation, which presumably consists of more primordial progenitors, and a CD133−/CD34+/VEGFR-2+ subpopulation, which is potentially more mature and further differentiated along the endothelial lineage.3,4 Our data point to the existence of a third, potentially more primitive EPC subpopulation, which is defined by surface expression of CD133 and VEGFR-2 while lacking surface expression of CD34. This conclusion is supported by from Nakamura et al,12 who cultivated murine bone marrow–derived stem cells that were negative for CD34 and lineage markers and converted them to CD34+ cells, thereby demonstrating that CD34− stem cells can be progenitors of CD34+ cells. Likewise, our human peripheral blood–derived CD34−/CD133+ EPC differentiated into CD34+/CD 133+ cells during in vitro expansion under selection pressure favoring endothelial differentiation. The CD34−/CD133+ EPC subpopulation is distinct from the EPC subpopulation described by Rehman et al,13 as the latter is characterized by high surface expression of the monocyte/macrophage marker CD14 (95.7±0.3%) and low expression of CD133 (0.16±0.05%), whereas our data show that CD34−/CD133+ EPC lack expression of CD14. Recently, Kuci et al14 cultured human peripheral blood–derived CD133+ cells with Flt3/Flk2 ligand and interleukin-6, thereby selecting a CD133+/CD34− stem cell population with the potential to repopulate nonobese diabetic/severe combined immunodeficiency (NOD/SCID) mice. Transplantation of these cells induced a substantially higher long-term engraftment compared with that of freshly isolated CD34+ cells. In addition, transplantation of bone marrow–derived CD133+ stem cells has been shown to functionally improve myocardial regeneration from infarction.15 Furthermore, when human cord blood–derived CD133+ progenitors were transplanted into nude mice, they incorporated into capillary networks in ischemic hindlimb, augmented neovascularization, and improved ischemic limb salvage.16 In our limb ischemia model, decreased numbers of CD34+/CD133+ EPC and increased numbers of CD34−/CD133+ EPC might be a consequence of increased homing of the latter to sites of ischemia, increased adherence of the former to the ischemic vasculature, or stem from increased ischemia-induced differentiation of the former cell subpopulation. Future studies are needed to dissect the underlying mechanisms. Additionally, in an acute ischemic setting such as unstable coronary lesions, we found CD34−/CD133+ EPC to be upregulated in a much higher degree than CD34+/CD133+ EPC, which might further speak to the particular importance of the former for vascular endothelial repair. In a different pathophysiological situation, in-stent restenosis, we found neither CD34−/CD133+ nor CD34+/CD133+ EPC, which is in accordance with current line of thinking that in-stent restenosis is mainly attributed to neointima proliferation caused by early smooth muscle cell in-growth from the media of diseased vessels.17 In our xenotransplantation model, transplanted CD34−/CD133+/VEGFR-2 progenitors consistently showed a higher capacity for homing to sites of vascular injury, reendothelialization, and lesion reduction as compared with CD34+/CD133+/VEGFR-2+ cells. We therefore hypothesize that CD34−/CD133+/VEGFR-2+ EPC represent key players for endothelial regeneration and tissue repair. Therapeutic transplantation of this specific stem cell subpopulation may be advantageous because of its higher regenerative capacity.
We conclude that CD34−/CD133+/VEGFR-2+ EPC are precursors of CD34+/CD133+/VEGFR-2+ EPC with a higher potential for vascular repair. These data extend current knowledge about the heterogenous EPC population and may have implications for the treatment of ischemic vascular disease and arterial injury.
This study was supported by the Deutsche Forschungsgemeinschaft.
Original received August 9, 2005; revision received September 9, 2005; resubmission received December 20, 2005; accepted January 13, 2006.
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