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HEMOSTASIS, THROMBOSIS, AND VASCULAR BIOLOGY
Human endothelial cells derived from circulating progenitors display specific
functional properties compared with mature vessel wall endothelial cells
Heidi Bompais, Jalila Chagraoui, Xavier Canron, Mihaela Crisan, Xu Hui Liu, Aurora Anjo, Carine Tolla-Le Port, Marylene Leboeuf,
Pierre Charbord, Andreas Bikfalvi, and Georges Uzan
Endothelial progenitor cells (EPCs) were
shown to be present in systemic circulation and cord blood. We investigated
whether EPCs display specific properties
compared with mature endothelial cells.
Human cord blood CD34ⴙ cells were isolated and adherent cells were amplified
under endothelial conditions. Expression
of specific markers identified them as
endothelial cells, also called endothelial
progenitor-derived cells (EPDCs). When
compared to mature endothelial cells, human umbilical vein endothelial cells
(HUVECs) and human bone marrow endo-
thelial cells (HBMECs), endothelial markers, were expressed to the same extent
except for KDR, which is expressed more
in EPDCs. They display a higher proliferation potential. Functional studies demonstrated that EPDCs were more sensitive
to angiogenic factors, which afford these
cells greater protection against cell death
compared with HUVECs. Moreover,
EPDCs exhibit more hematopoietic supportive activity than HUVECs. Finally,
studies in nonobese diabetic/severe combined immunodeficiency (NOD/SCID) mice
demonstrated that human circulating
EPCs are able to colonize a Matrigel plug.
EPDCs display the morphology and phenotype of endothelial cells. Their functional features indicate, however, that although these cells have undergone some
differentiation steps, they still have the
properties of immature cells, suggesting
greater tissue repair capabilities. Future
use of in vitro amplified peripheral blood
EPDCs may constitute a challenging strategy for cell therapy. (Blood. 2004;103:
2577-2584)
© 2004 by The American Society of Hematology
Introduction
Until recently, it was believed that neovascular formation in adults
resulted exclusively from proliferation, migration, and remodeling
of preexisting vessels, a process referred to as angiogenesis,
whereas, in contrast, vasculogenesis was defined as the formation
of new blood vessels from endothelial progenitor cells (EPCs)
during embryogenesis. The discovery of EPCs in adults1 has led to
the new concept that vasculogenesis and angiogenesis may occur
simultaneously. Extensive data support the existence of these cells,
their bone marrow origin, and their contribution to the formation of
new blood vessels in adults.2 Circulating EPCs (CEPCs) differ
from the circulating endothelial cells that are randomly detached
from the vessel walls and enter the circulation as a result of
vascular injury. When incubated in vitro with appropriate medium
in the presence of specific growth factors, CEPCs have a high
proliferating potential and within 10 to 20 days produce colonies of
cells expressing endothelial cell markers. Previous studies have
established that CEPCs express the cell surface markers CD34,2,3
CD133,4 and KDR.5
These cells may be recruited and incorporated into sites of
active neovasularization2,6 and have the ability to be mobilized
from bone marrow in response to ischemia or cytokine therapy.7
Moreover, human EPCs isolated from peripheral blood and expanded ex vivo have been shown to be activated in different models
of vascular diseases. For example, these cells have been shown to
promote successfully neovascularization of ischemic tissues in
nude mice8 and to develop into neovessels in vivo when seeded in
decellularized grafts.9
The physiologic role of CEPCs is not yet clearly established,
but several studies suggest that these cells are involved in the
protection against tissue degeneration due to vascular dysfunction.
The number of CEPCs is inversely correlated with the risk factor
for coronary artery disease.10 These results open up prospects for
the use of CEPCs as a prognostic tool for cardiovascular diseases.
In contrast, an increase in angiogenesis could have critical consequences in several diseases. Indeed, the hypothesis that tumor
growth was angiogenesis dependent11 and its subsequent confirmation by genetic methods12 provided a strong incentive to try to
target the vascular bed of tumors. Id mutant mice that do not
develop tumors because of defective angiogenesis have been
shown to develop tumors after bone marrow transplantation from
wild-type mice,13 suggesting that CEPCs also participate in tumor
vascularization. More studies are necessary to measure the real risk
of tumor development after CEPC injection. Meanwhile, the
potential presence of microtumors should be scrutinized with
peculiar attention before treating patients with highly angiogenic
cell therapy products.
Until now, the bone marrow stem cells that produce CEPCs
have not been clearly identified. Over recent years, much attention
From INSERM U506, Hôpital Paul Brousse, Villejuif, France; INSERM EPI
0113 Bordeaux, France; Institut du Cancer et d’Immunogénétique, Hôpital Paul
Brousse, Villejuif, France; and Laboratoire d’Hématopoı̈èse, Faculté de
Médecine, Tours, France.
Myopathie/Institut National de la Santé et de la Recherche Médicale
(AFM/INSERM No. 4CS14F).
Submitted August 20, 2003; accepted November 4, 2003. Prepublished online
as Blood First Edition Paper, November 20, 2003; DOI 10.1182/blood-200308-2770.
The publication costs of this article were defrayed in part by page charge
payment. Therefore, and solely to indicate this fact, this article is hereby
marked ‘‘advertisement’’ in accordance with 18 U.S.C. section 1734.
Supported by research grants from the Association Française contre la
© 2004 by The American Society of Hematology
BLOOD, 1 APRIL 2004 䡠 VOLUME 103, NUMBER 7
Reprints: Georges Uzan, Bâtiment Lavoisier, Hôpital Paul Brousse, 12, Av
P Vaillant-Couturier, 94807, Villejuif Cedex, France; e-mail: [email protected].
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BLOOD, 1 APRIL 2004 䡠 VOLUME 103, NUMBER 7
BOMPAIS et al
has focused on bone marrow stem cell potentiality. It has been
reported that bone marrow contains cells termed multipotent adult
progenitor cells (MAPCs), which, at a single-cell level, can be
differentiated into a large number of cell types, including endothelial cells.14 The existence of a common precursor for both
hematopoietic and endothelial cells (hemangioblast) was thought to
be restricted to embryonic development. However, recent studies
have identified a postnatal hemangioblast.15 Other studies support
the existence of a bipotent progenitor giving rise to both endothelial
cells and smooth muscle cells.16 The relationship between these
different bone marrow stem cells is not clear. It is also possible that
CEPCs mobilized from the bone marrow are in fact progenitors or
stem cells with broader differentiation potential, among which
endothelial differentiation is revealed at sites of neovascularization
that provide an adequate microenvironment.
The hypothesis that bone marrow is the reservoir of EPCs
prompted several groups to test the potential therapeutic effect of
autologous cells of medullar origin in human ischemic disease.
Bone marrow mononuclear cells injected locally in patients with
lower limb or cardiac ischemia seems to contribute to the regeneration of the vasculature.17-19 However, these studies constitute
pioneering clinical trials and it is likely that more purified cell
therapy product will be used in the near future, thus limiting
possible toxicity and side effects. In this context, autologous
CEPCs purified from peripheral blood and expanded in vitro
appear to be the best alternative to crude bone marrow mononuclear cells.
Before envisaging the use of differentiated EPCs for therapeutic
purposes, it is first essential to investigate whether they have
particular in vitro behavior compared with mature vessel wall cells.
The aim of our study was to increase knowledge of endothelial
progenitor-derived cells (EPDCs) by characterizing their specific
phenotypic and functional properties.
Materials and methods
Isolation and culture of CD34ⴙ mononuclear cells from cord
blood, HUVECs, HBMECs, and L87-4
Human umbilical cord blood samples were collected in blood packs
(Maco-Pharma, Tourcoing, France) from donors in compliance with French
legislation. Mononuclear cells (MNCs) were isolated by centrifugation on
Pancoll (Dutscher, Burmath, France). CD34⫹ MNCs were isolated by a
magnetic beads separation method following the manufacturer’s instructions (MACS; Miltenyi Biotec, Paris, France). CD34⫹ cells were plated
onto gelatin-coated wells (Sigma-Aldrich, Poole, United Kingdom) and
maintained in endothelial basal medium-2 (EBM2) supplemented with
EGM2-MV SingleQuots (Clonetics Cambrex, Ermerainville, France). Human umbilical vein endothelial cells (HUVECs) were isolated according to
the method of Jaffe et al.20 HUVECs and human bone marrow endothelial
cells (HBMECs),21 both used as endothelial cell positive controls, were
cultured in EGM2-MV. The fibroblastic cell line L87-422 was used as
negative control.
FACS analysis
Cells were detached with nonenzymatic cell dissociation medium to avoid
the destruction of cell membrane markers. Staining was performed with
monoclonal antibodies against CD31-phycoerythrin (PE; PharMingen, San
Diego, CA), CD146-fluorescein isothiocyanate (FITC; Chemicon, Temecula, CA), CD45-FITC (Immunotech, Marseille, France), and CD34peridinin chlorophyll protein (PerCP; Becton Dickinson, Heidelberg,
Germany), or with unconjugated antibodies against intercellular adhesion
molecule 2 (ICAM-2), endoglin (Immunotech), KDR (Sigma-Aldrich),
CD144 (Immunotech), integrin subunits ␤1, ␤2, ␣1, ␣2, ␣5, ICAM-1 (Becton
Dickinson, Le Pont de Claix, France), complex ␣v␤3, complex ␣v␤5
(Chemicon), vascular cell adhesion molecule 1 (VCAM-1; Diaclone),
FLT-1, fibroblast growth factor receptor 2 (FGFR-2), nitric oxide synthetase
III (NOSIII; Santa Cruz Biotechnology, Santa Cruz, CA) followed by
FITC- or PE-conjugated secondary antibodies (Sigma-Aldrich). Quantitative fluorescence analysis was performed using a fluorescence-activated
cell sorting (FACS) Calibur flow cytometer and WinMDI software (Becton
Dickinson).
Immunocytochemistry analysis
Cells seeded and fixed onto gelatin-coated 8-chamber Lab-Tek slides
(Nunc, Roskilde, Denmark) were sequentially incubated with primary
antibodies against CD31, von Willebrand factor (VWF; Dako, Glostrup,
Denmark), CD144, or collagen IV (SBA, Birmingham, AL) and with
appropriate secondary antibodies. To detect acetylated low-density lipoprotein (LDL) uptake, cells were incubated with Dil-ac-LDL. Slides were
examined and photographed under a fluorescence microscope (Leica,
Rueil-Malmaison, France).
Transmission electron microscopy analysis
Cells were fixed in 2.5% (vol/vol) glutaraldehyde, postfixed in 2% (wt/vol)
osmium tetroxide, dehydrated in graded ethanol, and embedded in epoxy
resin. Thin sections (76 nm) were cut and examined with a JEOL 100C
electron microscope (Jeol, Croissy sur Seine, France), after uranyl acetate
and lead citrate staining.
Semiquantitative RT-PCR analysis
RNA extraction was performed by the Trizol method, according to the
manufacturer’s instructions (Gibco, Karlsruhe, Germany). The reverse
transcription–polymerase chain reaction (RT-PCR) products were hybridized with an internal 32P-labeled probe to allow a semiquantitative analysis.
The oligonucleotides used were: CD133, 5⬘ probe: 5⬘GCCACCGCTCTAGATACTGC3⬘; 3⬘ probe: 5⬘TGTTGTGATGGGCTTGTCAT3⬘; internal
probe: 5⬘CGGCTCTAATTTTTGCGGTA3⬘. KDR, 5⬘ probe: 5⬘AATACCAGTGGATGTGATGC3⬘; 3⬘ probe: 5⬘CTGGCATGGTCTTCTGTGAAG3⬘; internal probe: 5⬘TCCCAGTTGAAGTCAATCCC3⬘. CD144, 5⬘
probe: 5⬘CCCACCGGCGCCAAAAGAGA3⬘; 3⬘ probe: 5⬘CTGGTTTTCCTTCAGCTGGA3⬘; internal probe: 5⬘GCCAGGTATGAGATCGTGGT3⬘.
Tie-2, 5⬘ probe: 5⬘CCAAACGTGATTGACACTGG3⬘; 3⬘ probe: 5⬘TGTGAAGCGTCTCACAGGTC3⬘; internal probe: 5⬘TGAGCCTTACTTTGGGGATG3⬘. Flt-1 5⬘ probe: 5⬘GTCACAGAAGAGGATGAAGG3⬘; 3⬘
probe: 5⬘CACAGTCCGGCACGTAGGTG3⬘; internal probe: 5⬘GGCTCTGTGGAAAGTTCAGC3⬘. GAPDH, 5⬘ probe: 5⬘CCACCCATGGCAAATTCCAT3⬘; 3⬘ probe: 5⬘GGTGGACCTGACCTGCCGTC3⬘; internal probe:
5⬘TGTGGTCATGAGTCCTTCCA3⬘.
Cell proliferation potential
At subconfluence, cells were detached and counted. The number of
divisions was estimated using the following equation: (Ln (number of cells
counted/number of cells at the beginning of the assay)/Ln2).
Cell proliferation assay
EPDCs or HUVECs were seeded and incubated overnight in EGM2-MV
medium (day 0). At day 1, cells were incubated in EGM2-MV medium
depleted in fibroblast growth factor 2 (FGF-2) and vascular endothelial
growth factor (VEGF) with or without 2.5% fetal bovine serum (FBS) as
control or supplemented with 2, 4, 8, 12.5, 25, or 50 ng/mL VEGF,
FGF-2, or placental growth factor (PlGF; R&D Systems, Minneapolis,
MN). At day 3, the medium was changed. At day 4, cells were counted in
a particle counter (CoulterZ1; Coultronics, Margency, France).
Cell survival assay
Cells were stained with annexin V–FITC and propidium iodide (PI)
according to the manufacturer’s protocol (R&D Systems) and subjected to
FACS analysis. To exclude necrotic cells, only PI⫺ cells were counted.
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ENDOTHELIAL PROGENITOR CELL–SPECIFIC PROPERTIES
2579
Wound healing assay
Cell migration was performed by using the method described by Sato and
Rifkin23 with modifications. One linear scar was drawn in a confluent cell
monolayer. A set of digital photographs was taken. The border between cells
and dish surface was then marked by a line on a computer using LuciaG of
Metech version 4.61 (Nikon, Paris, France). Dishes were incubated for 18
hours in EGM2-MV medium containing 0.1% bovine serum albumin
(BSA), without FBS, FGF-2, and VEGF as control, or supplemented with
10 ng/mL of VEGF, FGF-2, or PlGF. A second set of photographs was
taken. Photographs were superimposed on the computer screen, and nuclei
of cells that migrated across the line drawn at the border of the scar in the
first set of photographs were counted.
Capillary tube formation assay in Matrigel
For analysis of capillary tube formation, 150 ␮L Matrigel (Becton
Dickinson) was laid into a 48-well plate (Falcon, Heidelberg, Germany) and
incubated at 37°C for 30 minutes. EPDCs or HUVECs were trypsinized and
4 ⫻ 104 cells were suspended in 300 ␮L EGM2-MV medium and plated
onto Matrigel. Twenty-four hours later, the medium was removed and the
cells were incubated under various conditions: EGM2-MV medium depleted in FBS, FGF-2 and VEGF as control conditions, or control
conditions supplemented with 10 ng/mL VEGF, FGF-2, PlGF. Capillary
tube formation in Matrigel was observed under an inverted microscope after
6 hours of incubation.
Cobblestone area–forming cell assay
CD34⫹ sorted cells were seeded at limiting dilutions (ranging from 200 to
6.25 cells/well) on a preestablished confluent monolayer of EPDCs or
HUVECs in a long-term culture medium consisting of 50% (vol/vol)
Myelocult (StemCell Technologies, Vancouver, BC, Canada), 35% (vol/
vol) ␣-minimal essential medium (␣-MEM), 15% (vol/vol) FBS with 10⫺6
M hydrocortisone. Wells were screened weekly for the presence of
hematopoietic colonies. After 5 weeks, the incidence of cobblestone
area–forming cells (CAFCs) were estimated by plotting the proportion of
negative wells versus the number of cells per well. As control, to ensure that
the long-term culture medium used did not affect the proliferation of
HUVECs or EPDCs, we cultivated both types of cells in this medium for 5
weeks and compared their proliferation to that of HUVECs and EPDCs in
EGM2-MV medium.
Results
CD34ⴙ MNCs differentiate into cells with a typical
endothelial phenotype
Cord blood CD34⫹ cells were isolated (purity 93%-98%) and
cultivated under endothelial conditions, yielding a monolayer of
cells displaying morphology similar to that of endothelial cells.
We first characterized by flow cytometry the expression of a
selected set of markers and compared it to that obtained in
HUVECs and HBMECs and to the L87-4 cell line. Endothelial
cells derived from CD34⫹ cells, HUVECs, and HBMECs are
strongly positive for endothelial markers such as CD31, ICAM-2,
endoglin, and FGFR-2. All these cells express CD146, KDR,
and CD144, but at a low level. The culture no longer contained
any hematopoietic cells at this stage (day 40) as attested by the
absence of CD45⫹ cells (Figure 1). These cells are thus
endothelial cells. One of the important functions of endothelial
cells is to produce nitric oxide (NO). This NO production is
mediated by NOS. Among these, NOSIII is constitutively
expressed in endothelial cells. We showed that NOSIII is
expressed in EPDCs, HUVECs, and HBMECs (Figure 1). This
suggests that EPDCs are functional in terms of NO production.
Relative mean fluorescence intensity (MFI) was compared in
EPDCs, HUVECs, HBMECs, and L87-4 cells (Table 1). This
revealed that EPDCs expressed KDR at a significantly higher level
than HUVECs and HBMECs (4.9 ⫾ 0.3 versus 2.3 ⫾ 0.1 and
1.5 ⫾ 0.2, P ⬍ .001, n ⫽ 3). CD31, CD144, CD146, Flt-1, endoglin, ICAM-1, ICAM-2, ␣v␤3, ␣ v␤5, and integrin ␤1, ␣2, and ␣5 were
uniformly expressed on the 3 endothelial cell types. VCAM-1 and
integrins ␣1 and ␤2 were not expressed. CD133, a marker of
Matrigel plug assay
Nonobese diabetic/severe combined immunodeficiency (NOD/SCID) mice
were engrafted in the retro-orbital vein with 5 ⫻ 105 cord blood CD34⫹ or
5 ⫻ 105 cord blood CD34⫺. Four weeks later, Matrigel maintained at 4°C
was mixed with 1 ␮g FGF-2 and injected subcutaneously into NOD/SCID
mice. The experiment was performed 3 times with 5 mice under each set of
conditions. Ten days after injection, tissues containing the Matrigel plug
were removed and frozen. Matrigel cryosections were prepared. At the
same time, spleen, liver, lung, and heart of each mouse were frozen and
cryosections were prepared. May-Grünwald-Giemsa staining was performed to visualize the Matrigel invasion. Human cells were visualized by
using the MOM kit (AbCys, Paris, France) following the manufacturer’s
instructions on Matrigel sections and with primary antibody against human
CD31 (Chemicon) and then with an appropriate secondary antibody.
Finally, an FITC-labeled antibody against human CD45 (Immunotech) or
against CD144 (Bender MedSystems, Vienna, Austria) was added. Cryosections of organs were stained using same antibodies. Staining with ␣-smooth
muscle actin (ASMA) was also performed as positive control. 4,6-diamino2-phenylindole (DAPI) staining was used to visualize the cell nuclei.
Sections were examined and photographed under a fluorescence microscope (Leica).
Statistical analysis
Results are given as mean ⫾ SEM. Comparisons between means were made
using Student t test and the nonparametric Mann-Whitney U test.
Figure 1. FACS analysis of CD31, ICAM-2, endoglin, CD146, KDR, CD144, CD45,
FGFR-2, Flt-1, and NOSIII in EPDCs, HUVECs, and HBMECs at 40 days of
culture. Plots show isotype control IgG staining (white histograms) versus specific
antibody staining (gray histograms). Each analysis shown is a representative profile.
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BLOOD, 1 APRIL 2004 䡠 VOLUME 103, NUMBER 7
BOMPAIS et al
Table 1. MFI of EPDC, HUVEC, HBMEC, and L87-4 phenotypic markers analyzed by flow cytometry
EPDCs, MFI ⴞ SEM
HUVECs, MFI ⴞ SEM
HBMECs, MFI ⴞ SEM
KDR
4.9 ⫾ 0.3
2.3 ⫾ 0.1
1.5 ⫾ 0.2
L87-4, MFI ⴞ SEM
1.5 ⫾ 0.4
CD31
31.1 ⫾ 3.2
23.7 ⫾ 1.3
27.8 ⫾ 10.8
1.1 ⫾ 0.1
CD144
6.9 ⫾ 2.4
5.7 ⫾ 0.2
2.0 ⫾ 0.9
1.2 ⫾ 0.2
CD146
5.43 ⫾ 1.7
8.9 ⫾ 1.3
10.9 ⫾ 0.3
0.9 ⫾ 0.2
8 ⫾ 0.3
7.8 ⫾ 0.5
9.6 ⫾ 0.7
1.1 ⫾ 0.1
17.9 ⫾ 5.2
25.3 ⫾ 5.8
15.7 ⫾ 2.8
9.2 ⫾ 3.2
Flt-1
Endoglin
ICAM-1
7.7 ⫾ 3.6
14.2 ⫾ 4.5
9.2 ⫾ 1.6
7.2 ⫾ 0.3
ICAM-2
8.3 ⫾ 3.3
7.6 ⫾ 0.2
11.0 ⫾ 3.9
1.3 ⫾ 0.2
␣v␤3
7.1 ⫾ 1.0
12.5 ⫾ 3.4
4.3 ⫾ 0.6
2.4 ⫾ 0.2
␣ v ␤5
7.7 ⫾ 0.4
15.0 ⫾ 2.1
6.4 ⫾ 1.5
1.9 ⫾ 0.3
VCAM-1
1.6 ⫾ 0.5
1.0 ⫾ 0.1
1.6 ⫾ 0.6
1.2 ⫾ 0.1
Integrin ␤1
28.9 ⫾ 3.8
25.6 ⫾ 8.9
32.1 ⫾ 7.4
18.1 ⫾ 0.9
Integrin ␤2
1.1 ⫾ 0.1
1.0 ⫾ 0.1
1.1 ⫾ 0.1
1.3 ⫾ 0.4
Integrin ␣1
1.0 ⫾ 0.1
1.0 ⫾ 0.1
2.3 ⫾ 1.3
1.7 ⫾ 0.9
Integrin ␣2
12.9 ⫾ 6.3
16.2 ⫾ 6.5
12.0 ⫾ 5.8
1.7 ⫾ 0.5
Integrin ␣5
20.5 ⫾ 7.2
14.4 ⫾ 5.0
18.4 ⫾ 2.3
4.1 ⫾ 0.1
1.2 ⫾ 0.1
1.0 ⫾ 0.1
1.2 ⫾ 0.2
1.3 ⫾ 0.2
CD133
MFI was considered positive at 2.
immature hematopoietic cells and EPCs, was not detected on
EPDCs, HUVECs, or HBMECs, confirming previous observations.
The expression of all these markers was measured in L87-4
cells. These cells did not express KDR, CD31, ICAM-2, CD144,
CD146, ␣1, ␣2, ␤2 integrins, ␣v␤5, VCAM-1, Flt1, or CD133.
However, integrin ␤1 was markedly expressed, whereas endoglin,
ICAM-1, ␣v␤3, and integrin ␣5 were detected (Table 1).
Immunocytochemistry analysis was performed to characterize
further the phenotype of EPDCs. Figure 2 demonstrates that at day
40 of culture, EPDCs stained positively for CD144 and CD31. Both
have a membrane expression and have been observed in lamellipods and at cellular junctions. All cells incorporate Dil-Ac-LDL,
one of the characteristic functions of endothelial cells. The same
results were observed in HUVECs and HBMECs (Figure 2A). The
expression of different matrix molecules was also analyzed.
EPDCs and both positive control cells were stained with collagen
IV (Figure 2A) as well as the matrix network. L87-4 control cells
did not exhibit positive staining for CD31 and CD144 and did not
incorporate Dil-Ac-LDL (data not shown). Without primary antibody, no fluorescence was observed (data not shown). The granular
staining obtained with VWF antibody suggested the presence of
Weibel-Palade bodies, typical secretion granules present in endothelial cells. This was confirmed by electron microscopy analysis.
Figure 2B shows that EPDCs, as well as HUVECs, cultured for 40
days displayed organelles morphologically corresponding to WeibelPalade bodies. Furthermore, EPDCs exhibited a higher density of
endoplasmic reticulum network in comparison with HUVECs,
indicating an increased metabolic activity.
Variations in marker expression during EPDC differentiation
The expression of markers was monitored by RT-PCR followed by
hybridization of PCR products by an internal labeled probe. As
shown in Figure 3, the expression of CD133, a marker of immature
EPDCs, was still detectable at day 14 and decreased thereafter. This
marker was not detectable in HUVECs even at the beginning of the
culture (data not shown). Expression of endothelial markers such as
CD144, KDR, and Tie-2 was detected from day 14 and increased
throughout maturation, reaching levels comparable to that of
Figure 2. Immunofluorescence analysis and ultrastructural studies. (A) Immunofluorescence analysis of EPDCs, HUVECs, and HBMECs. Cells were stained with
antibody against VWF, CD144, CD31, or collagen IV. EPDCs as well as HUVECs and HBMECs take up Dil-Ac-LDL and were positively stained with VWF, CD144, CD31, or
collagen IV antibodies. Scale bar equals 5 ␮m. (B) Ultrastructural studies of EPDCs and HUVECs. EPDCs and HUVECs were fixed and prepared for transmission electron
microscopy. Scale bar equals 1 ␮m. Arrowheads indicate Weibel-Palade bodies.
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BLOOD, 1 APRIL 2004 䡠 VOLUME 103, NUMBER 7
ENDOTHELIAL PROGENITOR CELL–SPECIFIC PROPERTIES
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doublings under the same culture conditions. Outgrowth subsequently slowed down. HUVECs could only be cultured for 2
months under our conditions.
Effect of angiogenic factors on EPDC and HUVEC proliferation
in the presence of serum
We first tested to what extent VEGF, FGF-2, and PlGF induce cell
proliferation when incubated with EPDCs or HUVECs in the
presence of 2.5% FBS. Figure 5A shows that VEGF induced EPDC
proliferation from 4 ng/mL. Proliferation increased up to 50 ng/mL
in a concentration-dependent manner. In contrast, HUVECs were
less sensitive to VEGF induction because they did not respond at
concentrations below 12.5 ng/mL. Higher concentrations of VEGF
did not further increase their proliferation. Furthermore, EPDCs
and HUVECs responded strongly to 2 ng/mL FGF-2. The maximal
response was reached at 4 ng/mL FGF-2. PlGF stimulated EPDC
proliferation only at 50 ng/mL and was not efficient in inducing a
significant increase in HUVEC proliferation.
Figure 3. RT-PCR analysis of EPDCs and HUVECs. Expression of CD133, CD144,
KDR, Tie-2, Flt-1, and GAPDH mRNA was determined with a 32P-labeled internal
probe at days 14, 28, and 40 of culture for EPDCs and at day 21 of culture for
HUVECs. RT⫺ was the control realized without reverse transcription before PCR. D
indicates day.
HUVECs at day 40. The expression of Flt-1 was detected at day 14
and had decreased slightly by day 40, but was still detectable.
Effect of angiogenic factors on EPDCs and HUVECs in
serum-free conditions
Important differences between EPDCs and HUVECs were revealed
when cells were cultured with the same growth factors in the
absence of FBS (Figure 5B). Under these conditions, the number of
HUVECs moderately increased in the presence of VEGF. The
maximal effect (1.63-fold compared with unstimulated cells) was
observed at 8 ng/mL. In contrast, EPDCs were more sensitive to
growth factors. In the presence of 4 ng/mL VEGF, cell number was
EPDCs display a higher proliferation potential than HUVECs
Figure 4 compares EPDC proliferation potential to that of HUVECs.
The population doubling curve demonstrates that EPDCs have a
higher proliferation potential. After a period of latency (day 1 to
day 5), the cells grew in clusters. Serial dilutions showed that these
clusters appeared at a frequency of 1:4 ⫻ 105 to 5 ⫻ 105 seeded
CD34⫹ cells. The number of EPDCs rapidly increased to 6.9 ⫻ 013
cells per outgrowth progenitor cell, corresponding to up to 45
population doublings at day 94. Outgrowth of the EPDCs then
slowed down. EPDCs reached 49 divisions and could be maintained in culture for more than 3 months. In contrast, HUVECs
started proliferating immediately after seeding. The number of cells
reached 5.1 ⫻ 106/seeded cell, corresponding to 22 population
Figure 4. Proliferation potential of EPDCs compared with HUVECs under
EGM2-MV conditions. Cells were enumerated at each passage.
Figure 5. Effect of VEGF, FGF-2, or PlGF on EPDC and HUVEC proliferation,
survival, and mobility. Cell proliferation was analyzed (A) in the presence of 2.5% of
FBS or (B) under serum starvation conditions. (C) The percentage of living cells
compared with the control (EGM2-MV medium depleted in FBS, FGF-2, and VEGF)
was estimated by annexin V/PI staining. The annexin V⫺/PI⫺ population was
representative of living cells. (D) Effect of 10 ng/mL FGF-2, VEGF, or PlGF on EPDC
and HUVEC mobility compared with control conditions (EGM2-MV medium containing 0.1% BSA, depleted in FBS, FGF-2, and VEGF).
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BOMPAIS et al
2.32-fold higher than in the control. This effect was even more
striking at 12.5 ng/mL VEGF (7.39-fold increase). The maximal
response of HUVECs to FGF-2 was obtained with 2 ng/mL
(2.72-fold). At higher concentrations, the effect of FGF-2 decreased. FGF-2 was much more active on EPDCs (5.05-fold
increase at 2 ng/mL). This effect was even more exacerbated at
12.5 ng/mL (6.31-fold increase). Finally, PlGF was effective only
on EPDCs. The maximum effect was observed at 25 ng/mL
(3.32-fold increase).
We compared the response of subconfluent cells cultivated
under serum starvation conditions for 72 hours in the absence or
presence of increasing concentrations of VEGF, FGF-2, or PlGF.
The annexin V⫺/PI⫺ population represented the surviving cells. In
the concentration range tested, a protective effect of each growth
factor was seen in EPDCs as well as in HUVECs against serum
starvation–induced cell death. This effect was 2-fold higher in
EPDCs, whatever the growth factor used (Figure 5C).
Effect of angiogenic factors on EPDC and HUVEC mobility and
vascular tube formation
This was tested by incubating EPDCs or HUVECs with 10 ng/mL
VEGF, FGF-2, or PlGF. VEGF and FGF-2 significantly induced
EPDC and HUVEC mobility (Figure 5D). The effect was greater
for EPDCs than for HUVECs (262% for EPDCs versus 162% for
HUVECs). PlGF had no effect (Figure 5D).
We analyzed vascular tube formation in the presence of VEGF
or FGF-2 or PlGF. The EPDC network was denser than that
observed for HUVECs, especially with PlGF (data not shown).
Effect of EPDCs on human hematopoiesis
Between day 10 and 20 of culture, we observed in 15% to 20% of
our primary culture of EPDCs that hematopoietic cells spontaneously formed cluster structures underneath the endothelial cell
monolayer (Figure 6A), resembling CAFCs observed when hematopoietic progenitor cells were cocultured with stromal cells. These
structures maintain hematopoietic progenitors in long-term culture.
We followed the generation of CAFCs from human cord blood
CD34⫹ cells seeded at limiting dilutions and cocultured for 5
weeks with EPDCs or HUVECs as feeder layers. Figure 6B shows
a typical representative experiment of limiting dilutions for CAFCs
generated on these different feeders. After 5 weeks, the CAFCs
observed correspond to early progenitors. Their frequency was
1/21.2 ⫾ 1/2.69 (P ⬍ .005) for EPDCs versus 1/95.8 ⫾ 1/10.32
(P ⬍ .005) for HUVECs. This difference was significant (P ⬍ .03)
Figure 6. Hematopoietic supportive activity of EPDCs and HUVECs. (A)
CAFC-like structure observed in primary culture of CEPDCs. Original magnification,
⫻ 250. (B) Cord-blood CD34⫹ cells were seeded at limiting dilutions onto a
preestablished EPDC or HUVEC monolayer. EPDCs and HUVECs were seeded at
passage 2. CAFC frequencies (determined at 37% of negative wells) were measured
at week 5 of culture. Data shown are the means of 3 experiments.
and was observed from the first week, suggesting that EPDCs were
better able than HUVECs to support all kinds of CAFCs.
Even though the medium used for coculture with hematopoietic
cells is not fully optimized for endothelial cell cultures, we verified
that this medium does not affect HUVEC and EPDC proliferation
and survival (data not shown).
Human EPDCs migrate toward an angiogenic site and
participate in neovascularization in NOD/SCID mice
NOD/SCID mice engrafted with cord blood CD34⫹, CD34⫺, or not
engrafted were then injected with Matrigel plug and killed 10 days
later. May-Grünwald-Giemsa staining of Matrigel cryosections
showed vessel-like structures in the invaded Matrigel (Figure 7A).
The presence of human endothelial cells was analyzed by the
immunofluorescence of cryosections of the Matrigel plugs. No
human CD31⫹ cells or CD45⫹ cells were detected in the Matrigel
plug of the mice that were not injected (used as negative staining
control; Figure 7B) or were injected with the CD34⫺ population
(data not shown). However, the presence of human CD31⫹ cells
was detected in vascular structure present in the Matrigel plug.
These CD31⫹ cells were CD45⫺ as evidenced by double staining.
However, these CD31⫹ cells coexpressed the CD144 marker,
confirming their endothelial phenotype. This demonstrates that the
injected CD34⫹ population contained functional EPCs that had the
capacity to reach an angiogenic site and to participate in a
neoangiogenic process (Figure 7C). The staining of the spleen
cryosections did not reveal the presence of human endothelial cells
in this organ. No positive staining was obtained with antibody
against human CD31. Murine endothelial cells were detected by
CD144 staining (Figure 7D). As positive control, cells stained with
ASMA were detected (Figure 7D). Staining of liver, lung, and heart
cryosections did not reveal the presence of human endothelial cells
(data not shown). The same results were obtained with mice
injected with CD34⫺ cells.
Discussion
The use of EPCs as a tool for cell therapy in diseases involving
defects of angiogenesis, such as ischemia, was envisioned as soon
as these cells were discovered. Many experiments performed in
different models confirmed that these cells were very potent in the
revascularization of ischemic tissues and their specific tropism for
the diseased tissues was very encouraging.17-19 However, the
feasibility of using these cells as a cell therapy product in patients
with ischemic diseases was hampered by the rarity of the cells in
the peripheral blood. Expansion of these cells is thus necessary, but
until now all the ensuing technical problems were unresolved.
Among these problems is the determination of the extent to which
these cells could be expanded without losing their migration/
vascularization capacities. Meanwhile, increasing numbers of
clinical trials have started, using bone marrow mononuclear cells in
cell therapy of ischemic diseases.24 It is likely that EPCs play a
major role in the beneficial effect of cell therapy, but until now this
hypothesis was unproven.
It is therefore important to enhance our knowledge of the
specific features of EPDCs. We thus undertook a systematic study
of the phenotypic and functional properties of EPDCs and compared them to those of mature endothelial cells (HUVECs or
HBMECs). We found that from 14 to 40 days of culture, EPDCs
express all the markers of mature endothelial cells, whereas CD133
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BLOOD, 1 APRIL 2004 䡠 VOLUME 103, NUMBER 7
Figure 7. Presence of human endothelial cells in Matrigel plug. (A) May-GrünwaldGiemsa staining of Matrigel cryosection. (B) DAPI (blue) and human CD31 (red)
staining of Matrigel cryosections from negative control mice (no CD34⫹ transplants).
(C) DAPI (blue), CD144 (green), human CD31 (red), and double CD144 and human
CD31 (yellow) staining of Matrigel cryosection from mice receiving CD34⫹ transplants. (D) DAPI (blue), CD144 (green), human CD31 (red), and ASMA (red) staining
of spleen cryosections. Scale bars equal 10 ␮m.
expression decreases gradually during maturation. At day 40,
quantitative analysis of these markers in EPDCs, HUVECs, and
HBMECs did not reveal striking differences. However, KDR was
expressed at higher levels in EPDCs than in the other endothelial
cells. Like mature endothelial cells, EPDCs express NOSIII,
suggesting that these cells have the capacity to produce NO, one of
the important functions of endothelial cells. The morphology of
EPDCs is typical of endothelial cells, as revealed by immunofluorescence and electron microscopy. EPDCs also express high levels
of VWF, which is localized in granular structures resembling
Weibel-Palade bodies. The presence of these granules was further
confirmed by electron microscopy. These phenotypic similarities
between EPDCs and mature endothelial cells of different origins
confirm that EPCs are able to be fully differentiated in vitro into
endothelial cells, thus validating the differentiation culture conditions that have been used.
More striking differences between EPDCs and mature endothelial cells were revealed by functional studies. First, EPDCs could
be maintained in culture for more than 3 months, whereas
senescence of HUVECs was observed after 55 to 65 days of
culture. We observed that EPDCs were able to perform more
divisions than HUVECs. Thus, we confirm previous observations
showing that EPDCs are more proliferative than HUVECs, and
quantified this difference. We then investigated the individual
influence of VEGF, FGF-2, and PlGF on the proliferation of
EPDCs and HUVECs. In the presence of serum, HUVECs and
EPDCs responded to FGF-2 to a comparable extent. However,
EPDCs were more sensitive to VEGF than HUVECs. This
ENDOTHELIAL PROGENITOR CELL–SPECIFIC PROPERTIES
2583
observation correlates with the higher KDR expression observed in
EPDCs and may contribute to their higher proliferative capacities
compared with HUVECs. More striking differences were revealed
when the cells were cultured without serum. When 12.5 ng/mL
FGF-2, 25 ng/mL VEGF, or 25 ng/mL PlGF was added to the
culture, the number of EPDCs increased 6.5-, 7.5-, and 3.2-fold,
respectively, whereas the number of HUVECs was almost stable at
the same concentrations. Because the experiment was performed
over 72 hours, it is possible that this differential effect of growth
factor could be explained by a survival advantage of EPDCs
compared with HUVECs. Indeed, the percentage of living EPDCs
at 72 hours (annexinV⫺/PI⫺) was twice that of HUVECs. The
number of apoptotic EPDCs (annexinV⫹/PI⫺) was also half that of
HUVECs (data not shown). This greater sensitivity to growth
factors displayed by EPDCs compared with HUVECs confirms
that, even when differentiated, they still retain the properties of
immature cells. Another important function assumed by endothelial
cells is tissue repair. This function needs both migrating and
proliferative capacities. We measured the ability of the cells to
repair a linear scar cut in a confluent monolayer of endothelial cells.
This ability reflects both motility and proliferation potential. We
show that in the presence of FGF-2, the repair of the artificial
wound was more efficient with EPDCs than with HUVECs.
Converging evidence indicates that hematopoietic and endothelial
cells establish interactions in the bone marrow.25 In this context, we
observed cobblestone-forming structures in the EPDC culture that were
very similar to those obtained with hematopoietic progenitors cultured
on a stromal cell monolayer. Hematopoietic cells niche underneath the
stromal monolayer forming CAFCs. This interaction results in the
long-term maintenance of the immature and proliferative properties of
the hematopoietic progenitors. This observation prompted us to investigate whether EPDCs display the property of “stromal-like cells” that
could maintain the long-term survival of hematopoietic immature
progenitors. We used a classical test that has been established for stromal
cells and demonstrated that the stromal ability was much greater for
EPDCs than for HUVECs. This may reflect an intrinsic property and
suggests that, under certain conditions, endothelial progenitors may
cooperate with stromal cells in supporting hematopoiesis. Likewise, in
the context of leukemia, malignant hematopoietic cells could stimulate
the formation of new vessels in the bone marrow by producing high
levels of angiogenic growth factors such as VEGF.26
Moreover, it has been shown that endothelial and hematopoietic
progenitors share common markers suggesting a common progenitor. In particular, circulating endothelial cells and chronic myeloid
leukemia blasts may be of clonal origin because they both carry the
Philadelphia chromosome.27
Finally, we checked in NOD/SCID mice the ability of human
EPCs contained in the engrafted cord blood CD34⫹ population to
colonize a Matrigel plug diffusing FGF-2, constituting an in vivo
model of neoangiogenesis. We found that EPCs contained in the
CD34⫹ engrafted cell population were able specifically to reach the
Matrigel plug and to participate in vascular-like structures. We
analyzed whether EPCs were localized only in the Matrigel plug,
thus confirming the specific affinity of these cells for angiogenic
targets. No human cells were detected in the vascular structures of
organs such as liver, lung, heart, or spleen.
In summary, we show that in vitro–differentiated EPDCs display
the morphologic, phenotypic, and functional features of endothelial cells. In addition, even differentiated, these cells still retain
properties of immature cells as compared to mature endothelial
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2584
BLOOD, 1 APRIL 2004 䡠 VOLUME 103, NUMBER 7
BOMPAIS et al
cells isolated from blood vessels. Injected EPCs have the capacity
to reach a neoangiogenic site and there to differentiate into
endothelial cells, suggesting tissue repair capabilities. Another
specific feature of EPDCs is their ability to support long-term
survival of hematopoietic progenitors. This capacity could be used
to target pharmacologic agents to the angiogenic bone marrow
environment of the leukemic clone.
Acknowledgments
The authors thank the Centre Hospitalier de Longjumeau and the
Clinique Ambroise-Paré of Bourg-la-Reine for providing us
with cord blood samples, and Sophie Darquenne for her
technical advice.
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2004 103: 2577-2584
doi:10.1182/blood-2003-08-2770 originally published online
November 20, 2003
Human endothelial cells derived from circulating progenitors display
specific functional properties compared with mature vessel wall
endothelial cells
Heidi Bompais, Jalila Chagraoui, Xavier Canron, Mihaela Crisan, Xu Hui Liu, Aurora Anjo, Carine
Tolla-Le Port, Marylene Leboeuf, Pierre Charbord, Andreas Bikfalvi and Georges Uzan
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