Download Embryonic Cell Lines with Endothelial Potential: An In Vitro System

Embryonic Cell Lines with Endothelial Potential: An In
Vitro System for Studying Endothelial Differentiation
Yijun Yin, Jianwen Que, Ming Teh, Wei Ping Cao, Reida Menshawe El Oakley, Sai-Kiang Lim
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Objective—Endothelial differentiation is a fundamental process in angiogenesis and vasculogenesis with implications in
development, normal physiology, and pathology. To better understand this process, an in vitro cellular system that
recapitulates endothelial differentiation and is amenable to experimental manipulations is required.
Methods and Results—Embryonic cell lines that differentiate exclusively into endothelial cells were derived from early
mouse embryos using empirical but reproducible culture techniques without viral or chemical transformation. The cells
were not pluripotent and expressed reduced levels of Oct 4 and Rex-1. They were non-tumorigenic with a population
doubling time of ⬇15 hours. When plated on matrigel, they readily differentiated to form patent tubular structures with
diameters of 30 to 150 ␮m. The differentiated cells endocytosed acetylated low-density lipoprotein (LDL) and began
to express endothelial-specific markers such as CD34, CD31, Flk-1, TIE2, P-selectin, Sca-1, and thy-1. They also
expressed genes essential for differentiation and maintenance of endothelial lineages, eg, Flk-1, vascular endothelial
growth factor (VEGF), and angiopoietin-1. When transplanted into animal models, these cells incorporated into host
vasculature.
Conclusions—These cell lines can undergo in vitro and in vivo endothelial differentiation that recapitulated known
endothelial differentiation pathways. Therefore, they are ideal for establishing an in vitro cellular system to study
endothelial differentiation. (Arterioscler Thromb Vasc Biol. 2004;24:691-696.)
Key Words: endothelial 䡲 progenitor cells 䡲 differentiation 䡲 embryonic cell line
T
issue vascularization plays pivotal roles in normal physiological processes such as embryonic development and
adult tissue repair, as well as pathophysiological processes
such as tumor growth. It was previously thought that vasculogenesis or the in situ formation of blood vessels from
endothelial progenitor cells (EPCs) or angioblasts was restricted to the embryo, whereas neovascularization in adults
occurred from pre-existing mature endothelial cells or angiogenesis.1 However, with the identification of EPCs in several
adult tissues such as peripheral blood, bone marrow, and cord
blood, it was demonstrated that EPCs or angioblasts also
contribute substantially to neovascularization in adult tissues.2 Therefore, most current therapeutic strategies to enhance or inhibit neovascularization have incorporated endothelial progenitor cells (EPC) either as a therapeutic agent to
facilitate neovascularization or as a therapeutic target in
antiangiogenic therapy of malignant tumors3–5
To better-facilitate the applications of EPC, it is therefore
imperative to understand the molecular mechanisms in endothelial differentiation, and detailed molecular dissections of
differentiation pathways often require an in vitro cellular
system. At present, the cell types commonly used in the study
of endothelial cell biology are endothelial cells isolated from
blood vessels such umbilical veins and aortas. However,
these differentiated endothelial cells cannot be used to study
endothelial progenitor cell differentiation. Although putative
EPCs have been isolated from embryonic tissues, embryonic
stem (ES) cells, adult bone marrow, or peripheral blood using
various markers, such as a combination of markers associated
with hematopoietic stem cells such as Lin⫺, CD34⫹, c-Kit⫹
and Sca⫹, flk-1, VEGF and vascular endothelial (VE) cadherin, tie-2, and so on,2,6 the isolation procedures are usually
laborious and low-yielding. These cell preparations are often
heterogenous and variable. For example, different subsets of
bone marrow (BM) cells such as Lin⫺, c-Kit⫹; Lin⫺, CD34⫺,
low c-Kit⫹, Sca⫹; CD14⫺, CD34⫹; or human G-CSF–mobilized Lin⫺ CD34⫹lo BM cells have been shown to have
angiogenic potential, and when transplanted in experimental
models of acute coronary occlusion result in significant
recovery of the heart muscle.7–12 However, none of the
subsets has been demonstrated to be more superior in inducing angiogenesis. There is some confusion with the identity of
the angiogenic BM subfraction, eg, both CD34⫹ and CD34៮
subfractions have been used with success in inducing neovascularization.11–13 In isolating EPCs from ES cells, the
general method is to induce formation of embryoid bodies
Received December 23, 2003; revision accepted January 16, 2004.
From the Department of Obstetrics and Gynaecology (Y.Y., S.-K.L.); Department of Surgery (J.Q., R.M.E.O.); Department of Pathology (M.T.);
National University Medical Institute (P.C.), National University of Singapore; and Genome Institute of Singapore (S.-K.L.), Singapore.
Correspondence to Dr Sai-Kiang Lim, Genome Institute of Singapore, 60 Biopolis Street, Singapore 138672, Singapore. E-mail [email protected]
© 2004 American Heart Association, Inc.
Arterioscler Thromb Vasc Biol. is available at http://www.atvbaha.org
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DOI: 10.1161/01.ATV.0000120375.51196.73
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and isolating cells from embryoid bodies at different stages of
maturation to form blast colonies.14,15 Colonies with endothelial potential are identified by culturing them in the presence
of VEGF, c-kit ligand, and conditioned medium from an
endothelial cell line, D4T. Alternatively, ES-derived EPCs
have been isolated by culturing ES cells on collagen IVcoated plates and then purifying Flk-1⫹ E-cadherin⫺ cells by
fluorescence-activated cell sorter (FACS).16 Therefore, preparation of EPCs from ES cells will require several procedures
over a period of several days and impose a constraint on the
scale of preparation.
In some aspects, the study of endothelial differentiation is
limited by this lack of a suitable cell system with homogenous and expandable cells that can undergo endothelial
differentiation in vitro and in vivo. To overcome this obstacle, we describe a simple empirical culture protocol using
mouse embryos to derive embryonic cell lines that had the
potential to differentiate into vascular tissues with the appropriate endothelial markers when plated on matrigel or when
transplanted into suitable animal models. These cell lines are
not embryonic stem cells and do not express any endothelial
specific markers before differentiation.
Methods
Derivation of RoSH2 Cells
All animal experimentation protocols were approved by National
University of Singapore Animal Ethics Research Committee.
B6.129S7-GtRosa26 mice were purchased from Jackson Laboratory
(Bar Harbor, Me). The 5.5-days post coitum (dpc) delayed blastocysts and 6- to 7.5-dpc embryos were prepared as previously
described.17 For 6- to 7.5-dpc embryos, the egg cylinders were
dissected out and were placed next to the extra-embryonic tissues
during culture. All the embryos were cultured on embryonic fibroblast feeder in ES cell media.17 For the older embryos, the extraembryonic tissues were removed after the embryos attached and
started proliferation. The cultures were fed every 3 or 4 days. After
1 to 2 weeks, colonies of adherent, short, spindle-shaped cells with
loosely attached round cells were observed in some of the wells.
These colonies were picked when they reached a size visible to the
naked eye and were expanded. The colonies were picked by using a
pipet tip on Gilson P20 pipet to gently scrape the colony. The
detached colony was then transferred into a drop of trypsin and
incubated at 37°C for 5 minutes before plating onto a 3-cm feeder
plate. The pipetting of large colonies was facilitated by using a
large-bore pipet tip or a regular tip that had ⬇2 to 3 mm of the tip
cut off. Once the colony had expanded to a confluent 10-cm tissue
culture plate, the cells were adapted to growing on gelatinized plates
without feeder. Monoclonal cell line was established by plating
2⫻103 single cells in 100 ␮L ES media in methylcellulose-based
media consisting of 3.9 mL methylcellulose (MethoCult M3134;
StemCell Technologies, Vancouver, Canada), 4.2 mL DMEM, 1.5
mL serum, 100 ␮L diluted monothioglycerol solution (37.8 ␮L in10
mL PBS) (Sigma, St Louis, Mo), and 100 ␮L 100⫻L-glutamine/
penicillin/streptomycin stock solution (Life Technologies, Rockville,
Md). After 1 week, the colony that had the least number of ring-like
cells would be used to establish a line. After the colony expanded,
they were designated RoSH1, RoSH2, and so on, in the order of their
derivation.
Genomic DNA and Total RNA Analysis by
Polymerase Chain Reaction
Genomic DNA and total RNA were prepared using standard protocols and were quantified using, respectively, the RiboGreen RNA
Quantification kit and the Pico Green dsDNA Quantification kit
(Molecular Probes, Eugene, Ore). Primer set for amplification of the
following genes or genomic sequences and the expected amplified
fragment size were: Flk-1 5⬘-ATTGCACACACGGGATTCTG-3⬘
and 5⬘- CATACAGTACGACACTGACG-3⬘ and 744 bp; VEGF
5⬘-CCTATGACCACCCACATCCG-3⬘ and 5⬘-GATGAGGACCAGAATGAGAGAC-3⬘, 702 bp; TPI 5⬘-CCCTGGCATGACAAAGA CTT-3⬘ and 5⬘-CCTTGCTCCAGTCTTTCACAT-3⬘, 242 bp;
Rex-1 5⬘-ATGTCACTGCTGGTGCTGGA-3⬘ and 5⬘-TGCTAGCCAATTCCTCCCAG-3⬘, 409 bp; and Oct 4 5⬘ GGAGCACGAGTGGAAAGCAAC-3⬘ and 5⬘-TTCCTCCACCCACTTCTCCAG-3⬘, 327
bp. Primer sets for nestled polymerase chain reaction (PCR) of
angiopoietin-1 (ang-1) were: first primer set, 5⬘-GGAGGAAA
AAGAGAAGAAGAG-3⬘ and 5⬘-ACTTGCTGCTGCAACGGAGAC3⬘, 770 bp; and second primer set, 5⬘-GGAGGAAAAAGAGAAGAAGAG-3⬘ and 5⬘-TGAAATCAGCACCGTGTAAG-3⬘, 458 bp.
PCR and reverse-transcriptase PCR (RT-PCR) was performed as
previously described.18 All RT-PCR primers span at least 1 intron.
For nestled PCR, the second PCR was performed using the nestled
PCR primers and 2 ␮L of a 10-times diluted amplified reaction
sample.
In Vitro Differentiation
To differentiate RoSH2 cells in vitro, cells from confluent RoSH2
cell culture were plated on a matrigel-coated plate at 106 cells per
6-cm tissue culture dish. To coat the plate with matrigel, matrigel
(BD Laboratories, Torrey Pines, Calif) was diluted 10 times with ES
media before spreading on a tissue culture dish, and excess matrigel
was removed. The cells were fed every 3 to 4 days. The formation of
tubular structures was highly dependent on batches of matrigel. On
average, it took ⬇1 to 2 weeks before distinct tubular structures
became obvious. If we were very careful with the culture, then the
tubular structures could be maintained for ⬇4 weeks, reaching a
diameter of ⬇150 to 200 ␮m before falling apart, possibly from
dying through apoptosis. RoSH cells or RoSH-derived tubular
structures were labeled with fluorescent ␤-galactosidase (␤-gal)–
specific substrate, Imagene Green, according to the manufacturer’s
protocol (Molecular Probe, Eugene, Ore), and counterstained with
propidium iodide for nuclear staining. Acetylated low-density lipoprotein (LDL) uptake was determined by incubating the tubular
structures with diI-labeled acetylated LDL (Molecular Probe) for 24
hours at 37°C, fixed with formalin, and counterstained with SYTOX
Green fluorescent stain as per the manufacturer’s protocol.
Immunohistochemistry
Immunofluorescence and immunohistochemistry was performed using standard procedures. Cells and tissues are fixed in 4% paraformaldehyde. Tissues were either embedded in paraffin and sectioned
at 4-␮m thickness or subjected to whole-mount staining. The
primary antibodies used were goat anti-mouse P-selectin, goat
anti-mouse CD34, rabbit anti-mouse TIE2, and rabbit anti-mouse
Thy-1 (Santa Cruz Biotechnology, Santa Cruz, Calif), rat anti-mouse
CD31, rat anti-mouse Ly-6A/E (Sca-1), and rat anti-mouse Flk-1
(BD Pharmingen, San Diego, Calif). For paraffin-embedded tissue
sections, the primary antibodies were detected using biotinylated
secondary antibodies and streptavidin-conjugated horseradish peroxidase and DAB (Sigma, St Louis, Mo). The sections were counterstained with Mayer hematoxylin. For antibodies that did not react
with paraffin-embedded sections, whole-mount in situ immunofluorescence was performed. Briefly, after fixation, the tubular mesh was
incubated in sequential order: the first primary antibody, a biotinylated secondary antibody, and then avidin-fluorescein isothiocyanate.
The tissues were then counterstained with propidium iodide. The
tubular mesh was analyzed by confocal microscopy.
FACS Analysis
Cells were fixed in 2% paraformaldehyde, stained with the antibodies listed, and analyzed on a FACStarplus (Becton Dickinson, San
Jose, Calif).
Yin et al
Endothelial Progenitor Cell
693
Figure 1. Derivation of RoSH cell lines from mouse blastocysts/embryos. Mouse blastocysts/embryos were harvested and cultured on
embryonic fibroblast feeder cells. Some cultures had putative RoSH cell colonies and, if present, the colonies were present at a frequency of 1 per culture. When the colony expanded to a size visible to the naked eye, it was picked and expanded further on embryonic fibroblast feeder cells. The cells were then adapted to grow on gelatinized plates before they were plated singly on
methylcellulose-based media. When the cells had grown to form colonies visible to the naked eye, 5 individual colonies were picked
and expanded further in ES media on gelatinized plates. The colony that had the least number of ring-like cells was selected and used
to establish RoSH line.
Vascularization of ES Cell-Derived Teratomas
To determine vascularization by RoSH2 in ES cell-derived teratoma,
a cellular mix of 1⫻104 RoSH2 cells and 1⫻106 CS ES cells (a gift
from CS Lin) was injected subcutaneously into Rag1⫺/⫺ mice. After
3 weeks, the mice were sacrificed and the tumors were removed and
cryosectioned at 12 ␮m for immunohistochemistry staining with
rabbit anti-␤-gal antibodies (ICN, Aurora, Ohio).
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Incorporation of RoSH2 Cells Into Liver
Vasculature During Liver Injury
To demonstrate incorporation of RoSH2 cells into the vasculature
during liver injury, mice were anesthetized and 1⫻106 RoSH2 cells
were injected intrasplenically in 20 ␮L of saline. In shamtransplanted mice, saline was injected in the place of cells. The mice
were then injected intraperitoneally with 0.2 ␮g per gram body
weight of hamster anti-fas antibody (BD Pharmingen). After 5 days,
the livers were harvested and stained for the presence of ␤-gal as
previously described.19
Results
Derivation of RoSH Cell Lines
To isolate cells with endothelial potential, we postulated that
because vasculogenesis are initiated early in mouse embryogenesis, early embryos would be a good source of these cells.
Therefore, 3.5-dpc blastocysts, 5.5-dpc delayed blastocysts,
and 6.0- to 7.0-dpc embryos were harvested from pregnant
transgenic B6.129S7-GtRosa26 mice. The blastocysts and
embryos were cultured as outlined in Figure 1 and in
Methods.20 After 2 to 4 weeks, colonies of adherent fibroblastic cells with loosely attached, rapidly dividing round
cells and characteristic ring-like cells were observed in some
of the cultures and, when present, were present at 1 per
culture. Some colonies grew to a size visible to the naked eye
and, of these, ⬎95% survived to eventually establish a line.
The frequency of these colonies in cultures of delayed
blastocysts was ⬇1 in 30, and in that of E6.0 and older
embryos was ⬇1 in 10 (data not shown). We did not observe
these colonies in cultures of 3.5-dpc blastocysts or embryos
older than E8.0. After expanding these colonies as described
in Methods, they were plated as single cells in
methylcellulose-based media. When the cells had grown to
form colonies visible to the naked eye, 5 individual colonies
were picked and expanded further in ES media on gelatinized
plates. The colony that had the least number of ring-like cells
was used to establish RoSH line. The RoSH lines were
numerated as RoSH1, RoSH2, and so on, in the order of
derivation. Besides having similar cellular morphology, all
RoSH cell lines were tested and found to be able to form
tubular structures on matrigel.
To date, we have isolated 13 monoclonal lines, namely
RoSH1 to RoSH13, 4 from 5.5-dpc delayed blastocysts, 3
from 6.5-dpc embryos, 3 from 7.0-dpc embryos, and 3 from
7.50-dpc embryos. On gelatinized-coated plates, RoSH cells
have a fibroblastic morphology at subconfluency, with 1% to
5% of the cells assuming a von Willebrand factor-reactive
ring-like structure (Figure 2); at high confluency, the cells
formed a meshwork of cord-like structures (Figure I, available online at http://atvb.ahajournals.org). The cells have a
population doubling time of 12 to 15 hours, are not LIFdependent, and have little morphological resemblance to ES
cells. At the mRNA level, they expressed much reduced
levels of Oct4 and Rex-121,22 but increased levels of
brachyury23 (Figure II, available online at http://atvb.ahajournals.org). Because these cells were derived from B6.129S7GtRosa26 embryos, they expressed ␤-gal in the cytoplasm, as
verified by incubating the cells with Imagene Green, a green
fluorescent substrate for ␤-gal (Figure I).
Differentiation of RoSH2 Cells
When plated on matrigel, the RoSH cells terminally differentiate to form a mesh of patent tubular structures with
extensive branching and sprouting from the main tubular
structures and with diameters ranging from an average of 30
to 50 ␮mol/L to a maximum of 100 to 150 ␮mol/L (Figure
3A). These tubular structures can be maintained for a few
weeks before falling apart, possibly from dying through
apoptosis. The cells endocytosed acetylated LDL, a property
unique to endothelial cells and macrophages (Figure 3A).24
As typical of vascular structure, flattened monolayer cells
lining the lumen were in tight apposition on an amorphous
Figure 2. Characterization of RoSH2 cell line. A, Subconfluent
culture of RoSH2 cells on gelatinized tissue culture plate. Cell
morphology includes adherent fibroblast-like cells and ring-like
structures (arrowheads). B, Immunohistochemistry staining of
confluent RoSH2 cell culture for vWF. Arrows indicate brown
positive staining of ring-like structures. The cells were counterstained with eosin.
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Figure 3. In vitro endothelial differentiation of RoSH2 cells.
RoSH2 cells were induced to differentiate by plating the cells on
matrigel. A, Confocal microscopy of tubular structure and its
uptake of acetylated LDL. Left panel, The tubular mesh was
labeled with Imagene Green, a green fluorescent ␤-gal substrate
and propidium iodide. Right panel, The tubular mesh was incubated with diI-labeled acetylated LDL for 24 hours and counterstained with SYTOX Green, a green fluorescent nuclear dye. B,
Electron microscopy of the tubular structures. i, Endothelial cell
resting on acellular matrix with polarized plasma membrane. ii,
Filamentous structures on the luminal surface of endothelial cell
with microvesicles (arrowheads). iii, Tight apposing neighboring
cells (arrowheads). iv, Electron-nascent Weibel-Palade bodies
(arrowhead).
matrix and had polarized plasma membranes, with filamentous structures on the luminal surface and microvesicles
underlying the plasma membrane (Figure 3A). The larger
structures (100 to 200 ␮m) also had electron-dense organelles
reminiscent of nascent Weibel-Palade bodies, as described in
endothelial cells generated from ES cell-derived hemangioblast (Figure 3B).14 They also expressed typical endothelial
surface markers such as CD34, PECAM-1 (or CD31), Flk-1,
TIE2, Sca-1, Thy-1, and P-selectin (Figure 4). These antigens
were, however, not detected on undifferentiated cells by
FACS analysis (Figure III, available online at http://atvb.
ahajournals.org). Important factors in the regulation of vasculogenesis such as VEGF, VEGF receptor, Flk-1, and
angiopoietic factors were also expressed before and after
induction of differentiation (Figure IV, available online at
http://atvb.ahajournals.org).25–29 However, some of these
genes, eg, Flk-1 was not detected on the cell surface before
differentiation (Figure 4). We have observed that in the
undifferentiated RoSH cells, several endothelial-specific
genes such as Flk-1 and c-kit were transcribed but not
translated into proteins (unpublished data). To verify that
tubular structures formed by other RoSH lines were also
endothelial structures, we tested those formed by RoSH1 and
RosH3 lines and determined that these were also positive for
Figure 4. Immunohistochemical analysis of surface antigens. A,
Tie-2, Thy-1, CD34, and P-selectin staining by horseradish peroxidase– based standard immunohistochemistry on paraffinembedded sections of RoSH2-derived tubular structures. Brown
precipitates indicate positive staining. The nuclei were stained
with Mayer hematoxylin. B, Flk-1, Sca-1, and CD31 staining by
immunofluorescence using FITC-conjugated antibodies and
viewed under confocal microscope. The nuclei were counterstained with propidium iodide.
endothelial-specific markers, eg, Flk-1 and Tie-2 (data not
shown). RT-PCR analysis of RNA from RoSH1 and RosH3
lines also indicated that VEGF, VEGF receptor, Flk-1, and
angiopoietic factors were also expressed in these lines before
and after induction of differentiation (data not shown).
Transplantation of RoSH2 Cells in Mice to
Generate Vascular Tissues
To demonstrate endothelial potential of RoSH cells in vivo, 2
mouse models were used. In the first model, RoSH cells and
ESCs were co-transplanted into immunodeficient B6.Rag1⫺/⫺
mice, with the rationale that the developing ESC-derived teratoma will provide a suitable microenvironment for differentiation of RoSH cells into the entire repertoire of potential lineages.
RoSH cells were identified by the presence of ␤-gal. At 3 weeks,
4 of 6 mice had highly vascularized hybrid teratomas (Figure 5)
whereas mice injected with ES cells alone had similar-size
tumors after only 4 to 6 weeks (data not shown). Similar
injections of only RoSH cells from RoSH1 to RoSH6 lines into
isogenic C57BL6/J or B6.Rag1⫺/⫺ mice did not cause any tumor
formation during a 6-months observation period. Many of the
blood-filled vascular spaces were enclosed by a layer of RoSHderived endothelial cells, as evident by immunohistochemical
staining (Figure 5). Based on 20 random high-power (400⫻)
magnification fields per tumor section and 3 tumor sections per
tumor (n⫽4), we estimated that 41.5⫾19% (SD) of the vascular
tissues were derived from RoSH cells. Tumors formed from ES
cells alone were smaller, at ⬇50% of the size of those formed
from ES and RoSH cells. Not unexpectedly, the smaller tumors
were less vascularized. With less vascular spaces, they had a
harder, more solid macroscopic and histologic appearance (data
not shown). In the second model, the cells were injected
intrasplenically into C57BL6/J mice that were subsequently
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Figure 5. Formation of ES/RosH2 hybrid teratoma. Mouse ES
cells and RoSH2 cells were co-injected subcutaneously into
Rag1⫺/⫺ to generate teratomas. A, After 3 weeks, the mice had
highly vascularized tumors. They were euthanized, perfusionfixed, and the tumors were cryosectioned. B, Cryosections of
the tumors were immunostained with HRP-based anti-␤-gal
antibody. RoSH2-derived cells were identified by the presence
of ␤-gal, as indicated by dark brown staining (arrowheads).
treated with anti-fas antibody to induce liver injury. Five days
later, RoSH cells were shown to incorporate into the vasculature
of the liver, including the endothelium of hepatic central vein, as
demonstrated by X-gal staining (Figure 6).
Discussion
In this study, we described an empirical protocol using mouse
embryos to isolate and establish cell lines that were capable of
undergoing endothelial differentiation in vitro and in vivo.
Because the cardiovascular system is the first functioning
system to be formed during mammalian embryonic development, using delayed blastocysts and early mouse embryos in
our isolation protocol essentially increased the probability of
isolating cells with endothelial potential.20 These lines were
not ES cell lines and expressed much reduced levels of Rex-1
and Oct4, which are genes that are associated with pluripotent
stem cells.21,22 It is therefore unlikely that RoSH cells were
derived from the inner cell mass as ES cells, but we have yet
to confirm the parental embryonic cells from which RoSH
cells were derived. We hypothesized that they are likely to be
embryonic stem cells with endothelial potential and possibly
of mesodermal origin, because they expressed brachyury.23
Consistent with the apparent absence of pluripotency, RoSH
cells did not form teratomas when transplanted into immunodeficient Rag⫺/⫺ mice or isogenic C57BL/6J newborn or
adult mice during a 6-month observation period.
The RoSH cells did not display any typical endothelial and
hematopoietic markers before differentiation. On differentiation, they displayed endothelial markers such as CD34,
Endothelial Progenitor Cell
695
Figure 6. Anti-fas–induced liver injury. RoSH2 cells were
injected intrasplenically into mice treated with ␣-fas antibody to
induce apoptosis in hepatocytes, resulting in global liver hepatoxicity. After 5 days, mice were perfused, fixed, and the livers
were either stained with X-gal (A) or cryosectioned (B) at
15 ␮mol/L or 4 ␮mol/L (Figure V, available online at http://
atvb.ahajournals.org) before staining with X-gal. The cells were
counterstained with eosin.
PECAM-1 (or CD31), Flk-1, Tie-2, Sca-1, Thy-1, and
P-selectin, endocytosed acetylated LDL, and expressed genes
essential for differentiation and maintenance of endothelial
lineages, eg, Flk-1, VEGF, and angiopoietin-1. In particular,
the expression of Flk-1, Tie-2, VEGF, and angiopoietin-1
underscored the recapitulation of some of the known endothelial differentiation pathways. Flk-1, a receptor for VEGF,
and VEGF, a critical endothelial growth factor, are essential
for vasculogenesis and angiogenesis during embryo development. 26 –28,30 Tie-2 is a receptor tyrosine kinase for
angiopoietin-1 and is expressed almost exclusively in endothelial cells and early hematopoietic cells.29 Although
angiopoietin-1 does not directly promote the growth of
cultured endothelial cells, it is required for the later stages of
vascular remodeling and definitive hematopoiesis.31,32 Its late
expression during RoSH cell differentiation was consistent
with the branching and sprouting of patent tube-like structures to form a three-dimensional structure that is highly
reminiscent of vascular tissue.
Our in vitro data were further verified by in vivo transplantation studies. Using 2 different experimental models, we
showed that RoSH2 cells participated in angiogenesis during
anti-fas–mediated liver injury and healing and vascularization
of ES cell-derived teratomas. It is noteworthy that RoSH2
cells will proliferate and differentiate into endothelial cells
only under microenvironment of tissue injury.
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In summary, RoSH cell lines can differentiate to form
endothelial cells using pathways that recapitulated some of
the known endothelial differentiation pathways. They can be
easily maintained and manipulated in culture and can be
readily induced to form endothelial cells by plating on
matrigel. These characteristics fulfilled many of the requirements of an in vitro cell-based model system to understand
endothelial differentiation. In particular, they will be important in exploiting recent technological developments in highvolume gene expression and proteomic profiling to understand the roles of different genes and proteins in endothelial
differentiation. The rapid proliferation of RoSH cells and
their propensity to differentiate almost exclusively into endothelial cells would ensure an abundant supply of RNA and
protein from undifferentiated EPCs and endothelial cells for
analysis.
To some extent, ES cells also fulfilled many of these
requirements. They can differentiate into the endothelial
lineage by successive maturation steps recapitulating in vivo
events33 and have been used as a model system to study
vascular development during embryogenesis.14 However, unlike RoSH cells that differentiate almost exclusively into
endothelial cells when plated on matrigel, ES cells often
differentiate into complex multi-lineages. Protocols for generating ES cell-derived endothelial cells generally entail
several maturation and isolation steps aimed at the isolation
of rare endothelial progenitor cells.6 As such, ES cells are
generally more useful in studying molecular events in early
lineage commitments during embryogenesis rather than in
terminal endothelial differentiation. Therefore, we propose
that endothelial differentiation of RoSH cell lines represent a
more relevant model for in vitro studies on endothelial
differentiation.
12.
13.
14.
15.
16.
17.
18.
19.
20.
21.
22.
23.
24.
25.
Acknowledgments
We thank Dr Bing Lim (Genome Institute of Singapore/Harvard
Medical School) for his critique of the manuscript and for his advice.
26.
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Embryonic Cell Lines with Endothelial Potential: An In Vitro System for Studying
Endothelial Differentiation
Yijun Yin, Jianwen Que, Ming Teh, Wei Ping Cao, Reida Menshawe El Oakley and Sai-Kiang
Lim
Arterioscler Thromb Vasc Biol. 2004;24:691-696; originally published online February 5, 2004;
doi: 10.1161/01.ATV.0000120375.51196.73
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Supplementary figure I
(A) At confluency, the cells formed cord-like structures. (B) RoSH2 cells stained
positive for E.coli β-gal activity. The cells were incubated with Imagene Green, a
green fluorescent substrate for β-gal activity and counterstained with DAPI, a blue
fluorescent nuclear stain.
Supplementary figure II
Relative gene expression in RoSH2 cells vs ES cells. Total RNA from undifferentiated
RoSH2 cells and E14 ES cells were reverse transcribed and amplified for Rex-1, Oct4
and brachyury. A housekeeping gene, triose phosphate isomerase, TPI, was used as a
control for RNA quantitation.
Supplementary figure III
FACS analysis of surface antigens on undifferentiated cells. Postive control for each
antibody was mouse bone marrow mononuclear cells (not shown).
Supplementary figure IV
Endothelial gene expression. Total RNA was prepared from undifferentiated RoSH2
cells, RoSH2 cells that were plated on matrigel for 24 hours and mesh of RoSH2-derived
tubular structures. The total RNA was reverse transcribed into cDNA and the cDNA
amplified using gene-specific probes. Triose phosphate isomerase, a housekeeping gene
was used as a control.
Supplementary figure V
Liver samples were sectioned at 4 µM for morphological assessment at a higher
resolution. A representative section as viewed at 160× and 400× magnification,
respectively.