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HEMOSTASIS, THROMBOSIS, AND VASCULAR BIOLOGY
Functional endothelial cells derived from rhesus monkey embryonic stem cells
Dan S. Kaufman, Rachel L. Lewis, Eric T. Hanson, Robert Auerbach, Johanna Plendl, and James A. Thomson
We have used rhesus monkey embryonic
stem (ES) cells to study endothelial cell
development. Rhesus ES cells (R366.4
cell line) exposed to medium containing
vascular endothelial growth factor (VEGF),
basic fibroblast growth factor (bFGF), insulin-like growth factor (IGF), and epidermal growth factor (EGF) assumed a relatively uniform endothelial cell morphology
and could be propagated and expanded
with a consistent phenotype and normal
karyotype. When placed in Matrigel, these
rhesus ES cell–derived endothelial cells
(RESDECs) formed capillary-like structures characteristic of endothelial cells.
Immunohistochemical and flow cytometric analysis of RESDECs showed that
they take up acetylated low-density lipoprotein (LDL), express CD146, von Willebrand factor, and the integrin ␣v␤3, and
bind the lectin ulex europaeus agglutinin-1. These cells also express the VEGF
receptor Flk-1 and secrete VEGF. When
introduced in a Matrigel plug implanted
subcutaneously in mice, RESDECs
formed intact vessels and recruited new
endothelial cell growth. In vivo function
was demonstrated by coinjection of
RESDECs with murine tumor cells subcutaneously into immunocompromised
adult mice. RESDECs injected alone did
not form measurable tumors. Tumor cells
grew more rapidly and had increased
vascularization when coinjected with the
RESDECs. Immunohistochemical staining demonstrated that the RESDECs participated in forming the tumor neovasculature. RESDECs provide a novel means
to examine the mechanisms of endothelial cell development, and may open up
new therapeutic strategies. (Blood. 2004;
103:1325-1332)
© 2004 by The American Society of Hematology
Introduction
Embryonic stem (ES) cells serve as an excellent model to study
early mammalian development.1,2 Studies with mouse ES cells
have characterized phenotypic and genotypic pathways of many
cell lineages, including hemangioblast cells that are thought to
serve as a common precursor for both blood and endothelial
cells.3-5 Deletion of several specific genes such as Flk-1, SCL, and
Runx1 in mouse ES cells leads to defects in both blood and
endothelial cells, suggesting that common genetic pathways regulate development of both cell types.6-11 Lineage analysis based on
surface antigen expression and response to the addition of specific
growth factors also shows that Flk-1⫹ common precursor cells
derived from mouse ES cells can form both endothelial cells and
smooth muscle cells.12 Therefore, these ES cell–based studies
suggest that blood, endothelium, and smooth muscle cells are all
closely linked during embryogenesis.11,13
In addition to mice, pluripotent stem cells have also been
derived from preimplantation blastocysts of nonhuman primates
(common marmosets and rhesus monkeys) and humans.14-16 Like
mouse ES cells, these human and nonhuman primate ES cells can
be propagated indefinitely in culture as undifferentiated cells, yet
likely retain the potential to form any cell type of the body (though
this capacity cannot be formally tested with human ES cells).
Recent studies with human ES cells have defined methods to derive
and characterize many cell types of clinical interest, including
hematopoietic, neural, cardiomyocyte, pancreatic (insulin⫹ cells), endothelial, and trophoblast cells.17-23 Research on hematopoiesis using
rhesus ES cells also clearly demonstrates the utility of this system.24
Relatively few studies show in vivo function of cells derived
from ES cells. Recently, blood cells that can sustain long-term
engraftment of multiple hematopoietic lineages and be serially
transplanted between mice have been characterized.25,26 Also,
intact blood vessels composed of endothelium and smooth muscle
cells produced from a single precursor cell can be derived from
mouse ES cells.12 Other investigators have shown that neuronal and
pancreatic cells derived from ES cells can have some in vivo
function.27-30 These results make more viable the potential for ES
cells to serve as the starting point for novel therapies to replace or
repair cells or tissues damaged by disease, trauma, or other
degenerative processes.
Rhesus monkeys offer an excellent preclinical model for
potential human therapies, including models of gene therapy,
hematopoietic stem cell transplantation, and HIV therapies.31,32
Because of their similarity to human ES cells, the characterization
of rhesus monkey ES cells provides an excellent opportunity to use
these cells to devise new cell-based therapies that take advantage of
the unique properties of ES cells. Rhesus monkey ES cells are
similar to human ES cells in terms of morphology, growth
characteristics, and developmental potential.33 While resembling
each other, human and rhesus monkey ES cells are distinctly
different from mouse ES cells.15,16,33
Here, we use rhesus monkey ES cells to analyze endothelial cell
development. We show that treatment of undifferentiated ES cells
with defined growth factors known to support endothelial cell
growth leads to a homogenous population of endothelial-appearing
From the Wisconsin National Primate Research Center, the Laboratory of
Developmental Biology, and the Department of Anatomy, University of
Wisconsin, Madison; and the Freie Universität Berlin, Fachbereich
Veterinärmedizin, Institut für Veterinär-Anatomie, Germany.
Reprints: Dan S. Kaufman, University of Minnesota Stem Cell Institute, 420
Delaware St SE, MMC 716, Minneapolis, MN 55455; e-mail:
[email protected].
Submitted March 14, 2003; accepted October 7, 2003. Prepublished online as
Blood First Edition Paper, October 16, 2003; DOI 10.1182/blood-2003-03-0799.
BLOOD, 15 FEBRUARY 2004 䡠 VOLUME 103, NUMBER 4
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.
© 2004 by The American Society of Hematology
1325
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1326
BLOOD, 15 FEBRUARY 2004 䡠 VOLUME 103, NUMBER 4
KAUFMAN et al
cells. These cells express many of the same cell-surface antigens
and genes as endothelial cells isolated from other sources. Moreover, these cells are shown to produce functional blood vessels
which contribute directly to angiogenesis of tumor implants.
Materials and methods
Cell culture
Undifferentiated rhesus monkey ES cells (R366.4 cell line) were cultured as
previously described.14 Briefly, R366.4 cells were cocultured with irradiated mouse embryonic fibroblast (MEF) cells in media containing Dulbecco
modified Eagle medium (DMEM), 20% fetal bovine serum (FBS; Hyclone,
Ogden UT), 2 mM L-glutamine (Sigma, St Louis, MO), 0.1 mM 2-mercaptoethanol (Sigma), and 1% MEM nonessential amino acids (Invitrogen,
Carlsbad, CA). Undifferentiated cells were fed daily with fresh media and
passaged onto new MEFs approximately every 5 to 7 days. To promote
endothelial cell differentiation the medium was removed from the ES cells
24 hours after plating and replaced with medium consisting of endothelial
cell basal medium-2 (EBM2), 5% FBS, vascular endothelial growth factor
(VEGF), basic fibroblast growth factor (bFGF), insulin-like growth factor 1
(IGF-1), epidermal growth factor (EGF), and ascorbic acid (EGM2-MV
Bullet Kit; Clonetics/BioWhittaker, Walkersville, MD). The ES cells were
differentiated for 29 days in the EGM2 medium, which was changed every
3 to 5 days. Differentiated rhesus ES cells were dissociated with 0.05%
trypsin/0.53 mM EDTA (ethylenediaminetetraacetic acid; Gibco/Brl, Carlsbad, CA) for 5 minutes, centrifuged, and replated in EGM2 in 10-cm
tissue-culture dishes without irradiated MEF cells. After 24 hours, nonadherent cells were removed and adherent cells were fed fresh medium. The
rhesus ES cell–derived endothelial cells (RESDECs) could be grown to
confluence and serially passaged and expanded in the EGM2 medium.
Human umbilical vein cells (HUVECs; Clonetics/Biowhittaker) were also
grown and passaged in EGM2.
Tube formation on Matrigel
Matrigel (0.2 mL; Hynda Kleinman, Bethesda, MD; Becton Dickinson, San
Diego, CA) was added to each well of a 24-well tissue-culture plate and
allowed to solidify at 37°C for at least 30 minutes. Following gelation, 0.2
mL of a cell suspension containing 5 ⫻ 104 to 1 ⫻ 105 RESDECs was
placed on top of the Matrigel. The cultures were incubated at 37°C, 5%
CO2, and observed at 24, 48, and 72 hours for rearrangement of cells into
capillary-like structures.34 Individual experiments were performed in
triplicate and representative wells recorded by photomicrography.
Electron microscopy
Cells were detached from monolayer cultures by brief EDTA treatment,
rinsed with cacodylate buffer, and centrifuged at 1000g for 5 minutes.
Resulting pellets were fixed by immersion in Karnovsky fixative at 4°C for
2 hours. After 2 washes in cacodylate buffer (pH 7.4, 0.1 M) and subsequent
centrifugation, pellets were postfixed with 1% osmium tetroxide and 1.5%
potassium ferrocyanide in distilled water at 4°C for 2 hours.
Subsequently, samples were washed in cacodylate buffer, transferred to
20% bovine serum albumin (BSA) in cacodylate buffer and centrifuged
again. Two drops of 25% glutaraldehyde were layered on the pellet to
achieve gelling of BSA. Then samples were washed in cacodylate buffer,
dehydrated in a graded series of ethanol, and embedded in epoxy resin
(Polysciences, Warrington, PA). Polymerization was achieved at 60°C for
24 hours. Ultrathin sections (50 nm-60 nm) were cut on an LKB
ultramicrotome (LKB Pharmacia, Uppsala, Sweden), mounted on copper
grids, and contrasted with uranylacetate (10 minutes) and lead citrate (5
minutes). Sections were examined on a Zeiss 10CR electron microscope
(Weltzer, Germany).
VEGF and bFGF ELISA
RESDECs were cultured for 3 days in the absence of VEGF or bFGF in
EGM2, EGM2 supplemented with 10% Knockout serum replacer (Gibco/
Brl) instead of FBS, or DMEM supplemented with 10% FBS. After 72
hours the conditioned medium (CM) was collected and centrifuged to
remove dead cells. EGM2 medium alone served as a negative control. The
amount of VEGF or bFGF in the CM was analyzed by colorimetric
enzyme-linked immunosorbent assay (ELISA; R&D Systems, Minneapolis, MN).
Flow cytometry
RESDECs were washed with Ca2⫹- and Mg2⫹-free phosphate-buffered
saline (PBS) and detached from the monolayer with 0.05% trypsin/0.53
mM EDTA for 5 minutes. The dissociated cells were centrifuged and
washed with fluorescence-activated cell sorter (FACS) medium consisting
of PBS supplemented with 2% FBS and 0.1% sodium azide. After filtration
through 80-␮m mesh Nitex, the single-cell suspension was aliquoted and
stained with either isotype control or antigen-specific antibodies diluted to
appropriate concentrations in FACS medium. Cell-surface antigen expression was analyzed using antigen-specific primary antibody followed by
fluorescent-tagged secondary antibodies (indirect staining), or fluorescently
conjugated antigen-specific antibodies (direct staining). Appropriate unconjugated mouse and goat immunoglobulin G (IgG; both Sigma) as well as
fluorescein isothiocyanate (FITC)–conjugated mouse IgG (Pharmingen,
San Diego, CA) were used as isotype controls. Unconjugated antigenspecific antibody against flk-1 (Research Diagnostics, Flanders, NJ) was
detected with an FITC-labeled antigoat IgG antibody (Sigma). Unconjugated antibodies against VEGF receptor 1 (Flt-1) and VEGF receptor 2
(flk-1) (both Sigma) were detected with an FITC-labeled goat antimouse
IgG (Caltag, Burlingame, CA). Unconjugated P1H12 antibody that recognizes CD146 (mouse IgG1, kindly provided by Dr Robert Hebbel,
University of Minnesota), and anti–VE-cadherin antibody (Pharmingen)
were detected with rat antimouse IgG-FITC–conjugated secondary antibody (Caltag). The cells were also tested for expression of the VEGF
receptor using a biotinylated VEGF Kit (R&D Systems). Direct conjugated
antibodies were used against HLA-A, -B, -C–FITC (Pharmingen), ␣V␤3FITC (clone LM609, Chemicon). Ulex europaeus agglutinin-1–FITC
(Sigma) was used to directly measure binding to cell. HUVECs (Clonetics)
served as a positive control. Cells were analyzed without fixation on a
FACScan or FACS Calibur (Becton Dickinson) using propidium iodide to
exclude dead cells. Data analysis was carried out using CellQuest software
(Becton Dickinson).
Immunostaining
Analysis for the acetylated LDL receptor was performed by diluting
diIAcLDL (Molecular Probes, Eugene, OR) in serum-free EGM2 media.
Cells were washed twice and incubated overnight in EGM2 medium
containing diI-acetylated low-density lipoprotein (diIAcLDL). After washing, the cells were observed by fluorescence microscopy (UV, rhodamine
filter). HUVECs were used as a positive control. Expression of von
Willebrand factor (VWF) protein (Dako, Carpinteria, CA) was detected
with a goat antirabbit IgG-FITC secondary antibody (Sigma). Cells were
fixed, permeabilized, then incubated at room temperature for 1 hour with
VWF, washed, and incubated for 30 minutes in the secondary antibody.
After a final wash, cells were observed by fluorescence microscopy (UV,
FITC filter). HUVECs again served as a positive control.
Reverse transcriptase–polymerase chain reaction (RT-PCR)
Total RNA was extracted from HUVECs, RESDECs, rhesus bone marrow,
and rhesus umbilical cord (RNeasy kit; Qiagen, Valencia, CA) with
homogenization using a Qiashredder (Qiagen) according to the manufacturer’s instructions. RNA was quantified with a UV spectrophotometer and
cDNA was synthesized from 1 ␮g total RNA for each RT reaction. Reverse
transcription was performed using an Omniscript RT kit (Qiagen) with
RNase inhibitor (Promega, Madison, WI), and oligo(dT)15 primers (Promega) according to the manufacturer’s instructions. Simultaneous RT
reactions were performed without the addition of reverse transcriptase to
control for the transcription of contaminating genomic DNA. RT product (2
␮L) was used to perform PCR with a HotStarTaq PCR kit (Qiagen)
following the manufacturer’s instructions. An initial 15-minute, 95°C
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BLOOD, 15 FEBRUARY 2004 䡠 VOLUME 103, NUMBER 4
hotstart was used, followed by cycles consisting of 1 minute denaturation at
94°C, 1 minute annealing at variable temperature as noted below, and 1
minute extension at 72°C. A 10-minute extension was done at 72°C after the
final cycle. Thirty-five cycles were done for CD31, VWF, CD34, and VEcadherin, and 38 cycles for Flk-1. The amplified products were separated on
1.5% agarose gels and visualized using ethidium bromide staining.
Oligonucleotide primer sequences, annealing temperature (Ta), and
predicted product size were as follows: FLK-1: forward: 5⬘-ATGCACGGCATCTGGGAATC-3⬘, reverse: 5⬘-TGACTTCTCCTGCATGCACT-3⬘,
Ta: 58°C, product size: 537 bp; CD31: forward: 5⬘-CAACGAGAAAATGTCAGA-3⬘, reverse: 5⬘-GGAGCCTTCCGTTCTAGAGT-3⬘, Ta: 53°C, product size: 256 base pair (bp); VWF: forward: 5⬘-AGGCAGCTCCACTCGGCACAT-3⬘, reverse: 5⬘-AGCAAACAGTGGTAAGAGGAGGAC-3⬘, Ta:
58°C, product size: 582 bp; CD34: forward: 5⬘-GCACCCTGTGTCTCAACATGG-3⬘, reverse: 5⬘-GCACAGCTGGAGGTCTTATTTTGC-3⬘, Ta:
58°C, product size: 380 bp; VE-cadherin: forward: 5⬘-ACGGGATGACCAAGTACAGC-3⬘, reverse: 5⬘-ACACACTTTGGGCTGGTAGG-3⬘, Ta:
58°C, product size: 597 bp.
Matrigel plugs
In one experiment, severe combined immunodeficient (SCID) mice (Harlan
Sprague Dawley) were injected subcutaneously with 0.5 mL Matrigel
(Becton Dickinson) containing 5 ⫻ 105 to 1 ⫻ 106 RESDECs mixed evenly
in suspension within the plug.35 A second experiment was performed with a
polyvinyl sponge (Rippey, Eldorado Hills, CA) containing RESDECs
implanted into the solidified Matrigel.36 In each case, the Matrigel plugs
were removed after 14, 21, 28, 35, and 42 days. Vessels were observed by
injecting high-molecular-weight FITC-dextran (Sigma) intravenously a few
minutes before removing and fixing the plugs. Matrigel plugs were also
fixed and stained by standard hematoxylin and eosin (H&E) staining, as
well as immunohistochemistry using either an anti–HLA-A, –HLA-B,
–HLA-C antibody (W6/32) or IgG2a isotype control, followed by goat
antimouse IgG-FITC secondary antibody.
Tumor growth
Adherent RESDECs were harvested by trypsinization and mixed with the
mouse mammary carcinoma, C755 cell line. In 2 separate experiments
SCID mice (Jackson Laboratory, Bar Harbor, ME) were injected subcutaneously with either 1 ⫻ 106 C755 tumor cells or 1 ⫻ 106 RESDECs, or with a
mixture of 1 ⫻ 106 tumor cells and 1 ⫻ 106 RESDECs. After initial growth,
tumors were measured by caliper every 2 to 3 days. Approximately 3 weeks
after transplantation all mice were killed for histochemical analysis of the
grown tumors. Mice injected with RESDECs alone failed to grow tumors.
Statistical analysis was done by paired, 2-tailed Student t test using
Microsoft Excel software (Redmond, WA).
RHESUS MONKEY ES CELL–DERIVED ENDOTHELIAL CELLS
1327
shaped endothelial cells. In contrast, ES cells grown in medium
supplemented with FBS alone differentiated into a heterogeneous
cell population with no distinct endothelial-appearing cells. The
potential endothelial cells could be serial passaged and expanded
for approximately 20 population doublings while grown in EGM2,
with maintenance of a homogeneous appearance (Figure 1A-B). As
an initial test of endothelial cell identity, these cells were placed in
Matrigel-based media where they rapidly formed capillary-like
structures similar to those formed by HUVEC or other endothelial
cell populations when placed in Matrigel (Figure 1C-D).37 Cytogenetic studies showed all cells to have a normal rhesus monkey 40
XY karyotype (data not shown). These cytogenetic results are
important to demonstrate that these cells were not transformed after
prolonged culture, nor were they derived from potentially contaminating mouse embryonic fibroblast cells that are used for the
growth of undifferentiated ES cells.
Electron micrographs of the rhesus ES cell–derived cells
demonstrated typical endothelial cell features. These included
multiple dense round or rod-shaped structures that resemble
Weibel-Palade bodies, tight junctions between cells, and endocytic/
exocytic vesicles. Also, multilamellated bodies were identified
(Figure 2A-D). These structures appear similar to those seen in the
electron micrograph of human microvascular endothelial cells
(Figure 2E).
Immunophenotyping
Next, immunohistochemical staining of these rhesus ES cell–
derived endothelial-like cells demonstrated the presence of VWF,
the ability to rapidly take up acetylated LDL (acLDL) and to bind
the lectin UEA-1 (Figure 3). VWF and acLDL staining of the
rhesus-derived cells is similar to HUVECs (Figure 3A compared
with 3B, and Figure 3E compared with 3F). Undifferentiated rhesus
ES cells do not express VWF (Figure 3D). However, MEFs do
show trace staining by acLDL and VWF (Figure 3C,G), likely due
to the presence of some mouse endothelial cells within this
heterogeneous feeder-cell population. Importantly, since MEFs
used for ES cell culture are mitotically inactivated, no MEFs are
present within the rhesus-derived cell population (as also confirmed by cytogenetic analysis and surface antigen staining).
Immunohistochemical staining of tumors
In the first experiment tumors were isolated and fixed in 10% formalin for
the preparation of paraffin sections. To prepare frozen sections tumors were
fixed in 2% paraformaldehyde. All sections were mounted onto Fisher
Superfrost slides (Fisher Scientific, Pittsburgh, PA). Using the standard
ABC technique (VectaStain Elite ABC kit; Vector Laboratories, Burlingame, CA) sections were processed for expression of HLA class I A, B, C
(W6/32 antibody, IgG2a). Mouse IgG (Southern Biotechnology, Birmingham, AL) was used as isotype controls. The peroxidase activity was
visualized with a diaminobenzidine (DAB) substrate (Vector Laboratories)
and sections were counterstained with hematoxylin.
Results
Derivation of endothelial cells from rhesus monkey ES cells
Rhesus monkey ES cells were grown in EGM2 medium containing
VEGF, bFGF, EGF, and IGF as described in “Materials and
methods.” After approximately 5 to 10 days, these ES cells
assumed a uniform morphology similar to elongated or stellate-
Figure 1. Morphology of rhesus monkey ES cell–derived endothelial cells
(RESDECs). (A-B) Rhesus ES cells were grown in EGM2 media and then transferred
to a new tissue-culture plate where uniform flat, adherent, stellate-appearing cells
were demonstrated. (C-D) RESDEC cells were made into a single-cell suspension
and transferred to a Matrigel-coated plate where capillary tube–like structures were
seen within 18 hours after transfer. Original magnifications: ⫻ 100 (A,D); ⫻ 200 (B);
and ⫻ 40 (C).
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BLOOD, 15 FEBRUARY 2004 䡠 VOLUME 103, NUMBER 4
Figure 2. Electron photomicrographs of RESDECs. (A) Image showing
RESDEC with eccentric nucleus, vesicles, and electron-dense WeibelPalade bodies (original magnification, ⫻ 10 000). (B) Intercellular connection (tight junction) between adjacent cells (arrow) (original magnification,
⫻ 100 000). (C) Rod-shaped structures similar to Weibel-Palade body
(white arrow), electron-dense, multilamellated structures (black arrows)
and exocytotic vesicle (arrowhead) (original magnification, ⫻ 63 000). (D)
Endocytotic, exocytotic, and intracellular vesicles (arrows) (original magnification, ⫻ 40 000). (E) As comparison, electron photomicrograph of
human neonatal microvascular endothelial cell demonstrates many similar
structures including Weibel-Palade bodies (arrows) and endocytotic/
exocytotic vesicles (arrowheads) (original magnification, ⫻ 20 000).
Flow cytometric studies (Figure 4) demonstrate high levels of
UEA-1 binding, as well as expression of the integrin ␣v␤3 and the
surface antigen CD146 recognized by the P1H12 antibody. CD146
has been shown to be important in endothelial cell-cell interactions.38,39 These results led us to call these RESDECs. Surprisingly,
antibodies against the VEGF receptors Flk-1 and Flt-1 did not bind
RESDECs, though binding of these antibodies to HUVECs was
also weak. However, an assay using biotinylated VEGF protein and
secondary streptavidin-FITC showed binding to RESDEC and
HUVEC cells. Specificity of this binding was demonstrated by
blocking with an anti-VEGF antibody. RT-PCR studies (Figure 5)
did show expression of Flk-1 mRNA in the RESDECs, suggesting
that the antibodies may not cross-react. Alternatively, it is possible
that protein derived from this mRNA is not properly expressed on
the cell surface.
RESDECs do differ from HUVECs by lack of expression of
CD31 and VE-cadherin, two surface antigens commonly, but not
uniformly, found on the surface of endothelial cells (Figure 4).
RT-PCR studies failed to detect mRNA expression of these genes in
RESDECS. Importantly, RT-PCR for CD31 and VE-cadherin was
positive in both HUVECs and rhesus tissue (bone marrow aspirate
and umbilical cord tissue; Figure 5 and data not shown). Therefore,
Figure 3. Immunohistochemical staining of RESDECs demonstrating endothelial cell features. (A-D) von Willebrand factor staining demonstrates positive staining only
in RESDECs and HUVECs. (A) RESDECs; original magnification, ⫻ 100. (B) HUVECs; original magnification, ⫻ 40. (C) MEFs; original magnification, ⫻ 100. (D)
Undifferentiated rhesus ES cells; original magnification, ⫻ 100. (E-G) Uptake of acetylated LDL demonstrates specific staining of RESDEC and HUVEC. (E) RESDECs;
original magnification, ⫻ 100. (F) HUVECs; original magnification, ⫻ 40. (G) MEFs; original magnification, ⫻ 100. (H) Binding to ulex europaeus agglutinin-1 to RESDECs
grown as tubes in Matrigel; original magnification, ⫻ 100.
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BLOOD, 15 FEBRUARY 2004 䡠 VOLUME 103, NUMBER 4
RHESUS MONKEY ES CELL–DERIVED ENDOTHELIAL CELLS
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Figure 4. Single-color flow cytometric analysis of RESDECs and HUVECs. (A) RESDECs. (B) HUVECs. Appropriate isotype control is indicated by an open plot, and
indicated antibody by filled plot. Antibodies were either directly FITC-conjugated, or unconjugated antibodies were followed by staining with an FITC-conjugated goat
antimouse (GAM-FITC) secondary antibody, as indicated. In the last panels, VEGF binding was measured using biotinylated VEGF followed by avidin-FITC (black plot).
Specificity for VEGF staining is demonstrated by blocking with an anti-VEGF antibody (gray plot).
the negative PCR results for CD31 and VE-cadherin in RESDECs
was not due to nonreactive primers. Other genes commonly
expressed by endothelial cells including VWF, Flk-1, and CD34
were detected in both RESDECs and HUVECs (Figure 5).
Angiogenesis from RESDEC cells
In vivo function of the RESDECs was first assessed by a
sponge/Matrigel plug assay.36 Here, RESDECs were imbedded in a
sponge that was suspended into solidified Matrigel. This Matrigel
plug was then implanted subcutaneously in a SCID mouse.
Previous studies of this sponge/Matrigel assay demonstrate that
mouse-derived vessels will grow in response to proangiogenic
stimuli implanted in the sponge. This response is demonstrated
here when, after approximately 28 days, the mouse was injected
intravenously with an FITC-dextran solution, followed by plug
removal and imaging. The plug demonstrated intense vascular
localization toward the RESDEC-containing sponges (Figure 6A).
This angiogenic response was reminiscent of the directional
vascularization that arises from murine vessels that has been
previously demonstrated to occur in this sponge/Matrigel assay
when the sponge is impregnated with either angiogenic proteins
(such as VEGF or bFGF), or cell lines that promote an angiogenic
response (such as LnCap or 4T1 tumor cells, or C166 endothelial
cells).36 Based on this result, we tested RESDECs for their ability
to produce angiogenic proteins. Analysis of RESDEC supernatant
by ELISA demonstrated a significant level (600 pg/mL-1200
pg/mL) of VEGF produced by these cells (data not shown).
Figure 5. RT-PCR analysis of RESDECs and HUVECs. Total cellular RNA was
isolated from RESDECs and HUVECs. Sequence-specific primers for the indicated
genes were used for PCR analysis. RESDECs express RNA for VWF, flk-1, and
CD34 as indicated, though no expression of CD31 and VE-cadherin is detected. All 5
genes are expressed in HUVECs. To control for contaminating genomic DNA,
reactions were done using cellular RNA with no reverse transcriptase and these
reactions did not demonstrate bands for any of the primers in either cell type.
However, bFGF, another angiogenic protein often produced by
endothelial cells, was not detected in RESDEC culture supernatant
(data not shown). Therefore, like other endothelial cell lines,
RESDECs appear to be angiogenic. This angiogenic effect is likely
due at least in part to the ability of these cells to secrete VEGF.
To demonstrate that neo-vessels could be produced directly by
the RESDECs, 0.5 ⫻ 106 to 1.0 ⫻ 106 cells were evenly suspended
in a Matrigel plug implanted subcutaneously in a SCID mouse.35
Again, the animals were injected with FITC-dextran, followed by
plug removal and imaging. Here, larger vascular structures were
seen (Figure 6B). Subsequent histologic examination of the plug
showed vessel formation by RESDECs, as demonstrated by vessels
that were stained with antihuman HLA class I–specific antibodies
(that cross-react with rhesus monkey; Figure 6C-D). The lumen of
these vessels contained circulating red blood cells, and appeared
similar to vessels derived from other endothelial cells studied by
this Matrigel plug assay.35
Next, to demonstrate the ability of RESDEC cells to contribute
to active angiogenesis within tumors in vivo, another SCID mouse
model was used. Cells of the mouse mammary carcinoma line
C755 were injected subcutaneously either alone or coinjected with
an equivalent number of RESDECs. Tumor growth was measured
at regular intervals. Tumors grew significantly faster when coinjected with RESDECs (Figure 7A). Importantly, 106 RESDECs
injected alone did not lead to any measurable tumor growth,
suggesting that these cells are not themselves tumorigenic. The
C755 cell plus RESDEC-derived tumors were highly vascular
(Figure 7B). Again, immunohistochemical staining of the tumors
with antihuman–specific antibodies (that cross-react with rhesus
monkey but not mouse cells) demonstrated a contribution to the
endothelium from the RESDEC cells. Thus, staining of endothelial
cells was positive using antihuman MHC class I antibodies in C755
tumors coinjected with RESDEC cells (Figure 7C) but endothelial
cells in the vasculature of tumors derived from C755 cells alone
were negative for these antigens (Figure 7D).
Discussion
Here we demonstrate an efficient method to derive a nearly pure
population of endothelial cells from rhesus monkey ES cells. To
firmly establish endothelial cell identity, multiple complementary
methods were employed to characterize these cells. Morphology
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BLOOD, 15 FEBRUARY 2004 䡠 VOLUME 103, NUMBER 4
Figure 6. RESDECs induce and form vessels in Matrigel plug. (A) RESDECs were imbedded in a permeable sponge (arrow) that was suspended in Matrigel and implanted
subcutaneously. After 28 days, the mouse was injected intravenously with FITC-dextran and the Matrigel plug was removed and imaged to demonstrate neovascularization
centered toward the RESDEC sponge. Original magnification, ⫻ 10. (B) RESDEC cells were uniformly resuspended in a Matrigel plug and implanted subcutaneously. After 32
days, FITC-dextran was injected intravenously and the plug was removed and imaged to demonstrate intact vessels formed from the RESDEC cells in the plug. Original
magnification, ⫻ 40. (C-F) IHC demonstrates vessels in Matrigel plug (B) are RESDEC derived. Panels C, E, and G are phase contrast images of cross-sectioned Matrigel with
RESDEC-derived vessels, and panels D, F, and H are corresponding fluorescent IHC images using either isotype control (IgG2a) antibody (D) or anti–HLA class I antibody,
followed by secondary FITC-labeled antibody (F,H). Original magnifications: ⫻ 200 (C-D); and ⫻400 (E-H).
(both by light microscopy and electron microscopy), cell-surface
antigens, expression of VWF, and uptake of acetylated LDL were
all consistent with an endothelial cell phenotype. Moreover, these
cells played an active role in promoting angiogenesis. This was
shown first by recruitment of neo-vessels into a Matrigel plug, most
likely due to secretion of VEGF (and possibly other factors).
Second, active vessel formation by the RESDECs was seen when
the cells were suspended uniformly within a Matrigel plug. Finally,
coinjection of RESDECs with C755 mouse mammary tumor cells
resulted in more rapid tumor growth and showed a contribution to
the endothelial cells within the tumor by RESDECs. However, to
what degree the accelerated tumor growth is due to direct
contribution to the vasculature by the RESDECs or due to secretion
of angiogenic factors such as VEGF leading to recruitment of
mouse endothelial cells was not determined in our experiments.
Endothelial cell populations isolated at different developmental
stages, and from various tissues, can have significant phenotypic
differences. For example, endothelial cells isolated from mouse
brain, thoracic duct, and ovary each express a unique panel of
cell-surface antigens.40 Tumor cell adhesion to endothelial cells can
differ depending on site of origin of the endothelial cell.41
Developmentally primitive endothelial cells isolated from mouse
and pig yolk sacs express surface antigens that differ from more
developmentally mature endothelial cells.42-44 Interestingly, coculture of embryonic mouse yolk sac endothelial cells with embryonic
mouse brain rudiments leads to expression and functional upregulation of glucose transporter-1 and p-glycoprotein on the
endothelial cells.43 Since these antigens are typically expressed on
brain-derived endothelial cells, some endothelial cell heterogeneity
is likely acquired during organogenesis. Other molecular studies
have also characterized unique characteristics of venous, arterial,
lymphatic, and tumor endothelial cells.45-47
This known endothelial cell diversity can make characterization
and comparison of endothelial cell populations quite challenging.37,48,49 A panel of antigenic and functional assays is required to
provide optimal characterization of endothelial cell populations.
We demonstrate RESDECs function as endothelial cells based on
multiple criteria described above. Phenotypically, RESDECs bind
UEA-1, and express CD146, ␣v␤3, and VEGF receptors, as typical
of most other endothelial cells. Interestingly, the absence of CD31
and VE-cadherin suggest that we have identified a novel embryonic
endothelial cell population. Initial studies to look for these
antigenic markers by flow cytometry could have been falsely
negative due to an inability of the antihuman monoclonal antibodies to cross-react with the rhesus antigens. However, subsequent
RT-PCR analysis confirmed lack of expression of these genes. It is
important to note that while these antigens are commonly identified
on endothelial cells, various studies have also characterized
populations of endothelial cells that fail to express CD31 and/or
VE-cadherin.50,51 Moreover, expression of these surface antigens is
not unique to endothelial cells since both can be expressed on
hematopoietic precursor cells,52-55 and VE-cadherin can be expressed on some aggressive melanoma cells that act as vascular
“mimics.”56 Indeed, some studies that have used CD31 to select for
endothelial cells from bone marrow, peripheral blood, or ES cells
may unfairly skew the resulting expanded population of cells that
are derived solely from CD31⫹ progenitors.21 In contrast, in the
method used here, ES cells were allowed to grow and differentiate
in media optimized for endothelial cells, to induce outgrowth of
endothelial cells without bias toward a more specific phenotype.
Further studies are needed to examine if RESDECs and human ES
cell–derived endothelial cells21 represent early, developmentally
primitive endothelial cells and if they have the capacity to
differentiate into one or more specific types of endothelial cell
population. If so, these cells become an excellent resource to
study cellular and genetic mechanisms that regulate endothelial
cell diversity.
Most studies that have used mouse, human, or rhesus monkey
ES cells as the starting point to characterize developmental
pathways toward specific lineages use methods of differentiation
that lead to only a small percentage of cells being the cell type of
interest. For example, differentiation of any of these 3 types of ES
cells into hematopoietic cells by methods such as embryoid body
formation or coculture with stromal cells results in only about 2%
From www.bloodjournal.org by guest on June 17, 2017. For personal use only.
BLOOD, 15 FEBRUARY 2004 䡠 VOLUME 103, NUMBER 4
Figure 7. RESDECs lead to more rapid tumor growth and form vessels in
tumors. (A) 1 ⫻ 106 RESDEC cells (E), 1 ⫻ 106 C755 tumor cells (‚), or 1 ⫻ 106 of
each cell type (䡺) were injected subcutaneously in SCID mice. Tumors developed
earlier in mice coinjected with both cell types. RESDECs alone do not form detectable
tumors. Five mice were injected in each group and similar results were obtained in 2
independent experiments. Error bars represent standard deviation of tumor diameter
measurements for each group. The asterisk labels time points with statistically
significant (P ⬍ .01) increased tumor growth at the indicated time point. (B)
Hematoxylin and eosin–stained section of C755 tumor coinjected with RESDECs to
demonstrate vascularity. (C) C755-derived tumor without coinjection of RESDECs
stained for anti–HLA class I antibody that cross reacts with rhesus, but not mouse
cells. Minimal staining demonstrates lack of RESDEC-derived vessels. (D) C755
tumor coinjected with RESCDECs stained for anti–HLA class I antibody demonstrates tumor vessels derived from RESDECs. (E) Isotype control (IgG2a) staining of
same C755 tumor coinjected with RESDECs as in panel D. Original magnifications:
⫻ 200 (B); ⫻ 400 (C-E)
to 5% CD34⫹ cells.17,24,57 However, the method to derive endothelial cells in the studies presented here is unique in that virtually all
cells have the same phenotype, based on surface antigen expression.
Since all antigens are uniformly positive or negative in this cell
population (Figure 4), this suggests an essentially pure population of
endothelial cells. It is important to note that undifferentiated rhesus
monkey and human ES cells express Flk-1.17,24 Therefore, growth of
these cells in media containing VEGF may directly select or promote the
proliferation of endothelial cells.
Hematopoietic and endothelial cell development are intricately
intertwined.11,58 Several studies using mouse ES cells identify an
Flk-1⫹ hemangioblast cell that serves as a common precursor for
both blood and endothelial cells.3,4 Other studies using dissection
RHESUS MONKEY ES CELL–DERIVED ENDOTHELIAL CELLS
1331
of timed mouse embryos suggest that common hematopoietic and
endothelial cell precursors arise within the developing yolk sac and
aorta-gonad-mesonephric region.54,58-60 Specific transcription factors such as SCL and Runx1 required for hemangioblast development have been identified and absence of these genes prevents
normal development of both lineages.5,61 Similar characterization
of human (or nonhuman primate) ES cell–derived hemangioblasts
has not yet been done. Because the early embryologic stages of
yolk sac and early blood development are different between mice
and humans (although similar between humans and non-human
primates),60 it is unclear to what extent the phenotype and genotype
of hemangioblasts, hematopoietic, and endothelial cell precursors
are similar or different between these species. Effective methods to
derive both blood and endothelial cells from both human and
rhesus monkey ES cells have now been clearly identified.17,21,24
Further studies to dissect more carefully the developmental events
that occur during differentiation of these earliest precursor cells
should now be undertaken.
ES cell–derived endothelial cells may be suitable for clinical
studies to revascularize ischemic tissues. Flk-1⫹ cells derived from
mouse ES cells can form functional vasculature in vivo.12 Human
endothelial cell progenitors isolated from peripheral blood and
expanded in vitro will form vessels and improve blood flow in
mouse models of limb and myocardial ischemia.62,63 Other progenitor cells isolated from human bone marrow form endothelial cells
that contribute to regions of active angiogenesis.64 Since human
and nonhuman primate ES cells can be grown in virtually unlimited
numbers, these cells may serve as a stable source of therapeutic
endothelial cells.
Finally, the ability for RESDECs to contribute actively to
angiogenesis of developing tumors may lead to novel therapeutic
strategies to inhibit tumor growth. Human ES cells can now be
engineered to express exogenous genes65,66 and similar strategies
should be possible with rhesus monkey ES cells. It will now be
feasible to derive endothelial cells that stably express genes that
will inhibit tumor growth by blocking angiogenesis. For example,
antiangiogenic proteins or peptides such as endostatin, or a
dominant-negative form of Raf-1,67 could be secreted, perhaps in
an inducible fashion. In this manner, injection of these engineered
ES cell–derived endothelial cells might directly target regions of
active angiogenesis and inhibit tumor growth. Another intriguing
study demonstrates the ability of fixed xenogeneic endothelial cells
to act as a vaccine to induce tumor regression due to antiangiogenic
effect.68 The availability of a nonhuman primate model of ES
cell–based treatments will be very advantageous in development
and preclinical testing of these various novel therapeutic strategies.
Acknowledgments
We gratefully acknowledge Hynda Kleinman for providing Matrigel, Robert Hebbel for providing P1H12 antibody, Charles Harris
for karyotype analysis, and Andrew MacLean for providing the
rhesus umbilical cord sample.
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2004 103: 1325-1332
doi:10.1182/blood-2003-03-0799 originally published online
October 16, 2003
Functional endothelial cells derived from rhesus monkey embryonic
stem cells
Dan S. Kaufman, Rachel L. Lewis, Eric T. Hanson, Robert Auerbach, Johanna Plendl and James A.
Thomson
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