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Regular Articles |
From the Departments of Surgery,*
Cardiovascular
Surgery,
and
Cardiology,
Children’s Hospital, Boston; and
the Department of Pathology,
Brigham and
Women’s Hospital, Boston, Massachusetts
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Abstract |
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 Top Abstract IntroductionMaterials and MethodsResultsDiscussionReferences |
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-smooth muscle actin (-SMA), an established
marker for this process, two distinct pathways of
transdifferentiation were identified in clonally derived endothelial
cell populations isolated from ovine aortic valve leaflets. -SMA
expression was induced by culturing clonal endothelial cells in medium
containing either transforming growth factor-ß or low levels of serum
and no basic fibroblast growth factor. Cells induced to express -SMA
exhibited markedly increased migration in response to platelet-derived
growth factor-BB, consistent with a mesenchymal phenotype. A
population of the differentiated cells co-expressed CD31, an
endothelial marker, along with -SMA, as seen by
double-label immunofluorescence. Similarly, this co-expression
of endothelial markers and -SMA was detected in a subpopulation of
cells in frozen sections of aortic valves, suggesting the
transdifferentiation may occur in vivo. Hence,
the clonal populations of valvular endothelial cells described here
provide a powerful in vitro model for dissecting
molecular events that regulate valvular endothelium.
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Introduction |
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TopAbstractIntroductionMaterials and MethodsResultsDiscussionReferences |
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During development, the heart begins as a tube
consisting of two layers, the endocardium and the myocardium, separated
by an extracellular matrix known as cardiac jelly.3
Studies of avian heart development indicate that endothelial cells
(ECs) from the endocardium are initially activated by an unknown factor
secreted by the myocardium.4
Sequential signaling by
transforming growth factor (TGF)-ß2 and -ß3 then promotes
transdifferentiation of a subset of ECs to mesenchymal
cells.5
These cells migrate into the cardiac
jelly4,6
and further develop into mature valves. A
biochemical marker for this transdifferentiation is the expression of
-smooth muscle actin (-SMA), which is not normally expressed by
ECs. -SMA has been shown to be important for mesenchymal formation
from ECs in embryonic explants of endocardial cushion
tissue,7
suggesting that -SMA may play a functional
role in this process. Alterations in cell phenotype, such as that
occurring in the developing valve, are referred to by various terms
including phenotypic modulation, transformation, or
transdifferentiation.8
We use transdifferentiation here to
denote an irreversible alteration from one cell lineage to that
characteristic of a different cell lineage.
In contrast to embryological studies, little is known about the cellular properties of mature valves. Valve leaflets are composed of an outer layer of ECs that cover interstitial mesenchymal cells located throughout the leaflet. Both the ECs9 and interstitial cells10 of valves have been isolated and cultured in vitro. ECs from human cardiac valves appear spindle-shaped in culture, but like all ECs, they express inducible intercellular adhesion molecule-1 and E-selectin.9 The interstitial cells, although not well characterized, have been reported to share some characteristics with both fibroblasts and smooth muscle cells.11
Because valvular disease leads to 60,000 valve replacement surgeries every year in the United States and current replacement valves are not optimal,2 understanding the cells that make up valves is essential if better options for correcting valvular defects are to be developed. To pursue this, we isolated ECs from the aortic valve leaflets of healthy mature sheep. Although the cells in primary culture formed typical cobblestone endothelial monolayers, other cellular morphologies, reminiscent of mitral valve interstitial cells,10 appeared within 2 to 3 weeks of subculture. We therefore isolated clonal EC populations from adult aortic valve leaflets that express the endothelial markers CD31/PECAM-1 and E-selectin. Herein we demonstrate that clonal ECs can transdifferentiate to a mesenchymal cell phenotype in a manner that resembles one of the early events during valve development.
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Materials and Methods |
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TopAbstractIntroductionMaterials and MethodsResultsDiscussionReferences |
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Material used were endothelial basal medium (EBM) (CC-3121;
Clonetics, San Diego, CA); fetal bovine serum (FBS) (Hyclone, Logan,
UT); 100x GPS (29.2 mg/ml L-glutamine, 10,000 U/ml
penicillin G, 10,000 µg/ml streptomycin sulfate); gentamicin sulfate
and 100x PSF (10,000 U/ml penicillin G, 10,000 µg/ml streptomycin
sulfate, 25 µg/ml amphotericin B) (Life Technologies, Inc., Grand
Island, NY); collagenase A (Boehringer Mannheim, Indianapolis, IN);
Immobilon-P membrane (Millipore, Bedford, MA); Hyperfilm ECL,
fluorescein-streptavidin, and Texas Red-streptavidin (Amersham Life
Sciences, Arlington Heights, IL); Lumiglo (KPL); human TGF-ß1,
recombinant human TGF-ß2 and -ß3, recombinant human
platelet-derived growth factor (PDGF)-BB, and anti-PDFG-BB (R&D
Systems, Minneapolis, MN); Vectastain Elite ABC kit, avidin/biotin
blocking kit, fluorescein anti-mouse IgG, Texas red anti-rabbit IgG,
peroxidase-conjugated anti-goat IgG, biotinylated horse anti-mouse IgG,
avidin-peroxidase, peroxidase-conjugated anti-mouse IgG, and
3,3'5,5'-tetramethylbenzidine (Vector Laboratories, Burlingame, CA);
3-amino-9-ethyl carbazol and mouse anti-human -SMA (clone
1A412
) (Sigma Chemical Co., St. Louis, MO); goat
anti-human CD31/PECAM-1 IgG (Santa Cruz Biotechnology, Santa Cruz, CA);
rabbit anti-human von Willebrand factor (vWF) and mouse anti-human
CD31/PECAM-1 (DAKO, Carpinteria, CA); polycarbonate PVP-F membranes
(Neuro Probe, Inc, Gaithersburg, MD). Recombinant human bFGF was kindly
provided by Scios Nova Inc., Mountain View, CA; soluble recombinant
TGF-ß type II receptor, prepared as described,13
was
kindly provided by Philip Gotwals, Biogen, Cambridge, MA; SM1 antibody
was kindly provided by Masanori Aikawa, Brigham and Women’s Hospital,
Boston; rabbit anti-bovine CD31/PECAM-1 was kindly provided by Steven
Albelda, University of Pennsylvania. Figures were prepared from scanned
images using Adobe Photoshop version 5.5.
Tissue Procurement
Ovine tissues from animals weighing 20 to 25 kg and 8 to 10 months of age were obtained under approved guidelines for animal experimentation at Children’s Hospital, Boston. Human valvular cells were isolated from pulmonary valve leaflets obtained from children undergoing open-heart surgery at Children’s Hospital, Boston. Valve tissue was obtained in accordance with the Committee on Clinical Investigation, Children’s Hospital, Boston. Adult human aortic valve tissue was obtained in accordance with the Human Investigation Review Committee, Brigham and Women’s Hospital, Boston, MA.
Cell Culture
Ovine and human valve cells were grown on 1% gelatin-coated dishes in EBM, 10% heat inactivated FBS, 1x GPS, and 2 ng/ml bFGF (growth medium). Cells were passaged 1:3 or 1:4 every 6 to 14 days and used between passages 8 to 14. Human dermal microvascular ECs (HDMECs) and human umbilical vein ECs (HUVECs) were isolated and cultured as described.14,15
Isolation of Valve ECs
Valve leaflets were incubated in media with 5% FBS, PSF, 2 mmol/L L-glutamine, and 100 µg/ml gentamicin sulfate for 1 to 4 hours, then minced into 2 mm2 pieces, incubated with 0.2% collagenase A in EBM for 5 minutes at 37°C, and diluted with Hanks’ balanced salt solution containing 5% FBS, 1.26 mmol/L CaCl2, 0.8 mmol/L MgSO4, and 1x PSF (wash buffer). The supernatant containing released cells was sedimented at 200 x g, resuspended in growth medium, and plated. The following day, primary cultures were washed to remove unattached cells and refed. ECs from primary cultures of human pulmonic valve were isolated using Ulex europaeus I-coated Dynabeads.16
Clonal Cell Populations
Primary cultures were trypsinized, resuspended in growth medium at 3.3 cells/ml, and 100 µl plated in each well of a 96-well plate, yielding a distribution, on average, of one cell in every third well. Each well was then checked to confirm that only one cell colony grew up in each well. When the colonies covered two-thirds of the well, cells were split into 24-well dishes.
Indirect Immunofluorescence
Cells on gelatin-coated glass coverslips were fixed with -20°C methanol, incubated with primary antibodies followed by species-specific fluorescein-conjugated and Texas Red-conjugated secondary antibodies, and then analyzed using a Zeiss Axiophot II fluorescence microscope. The rabbit polyclonal antibody against bovine E-selectin described previously17 was shown to cross-react with ovine E-selectin (data not shown). A rabbit polyclonal against bovine CD31 or a goat anti-human CD31/PECAM-1 was used to detect ovine CD31 by immunofluorescence.
Western Blots
Cells were lysed with 4 mol/L urea, 0.5% sodium dodecyl sulfate, 0.5% Nonidet P-40, 100 mmol/L Tris, and 5 mmol/L ethylenediaminetetraacetic acid, pH 7.4, containing 100 µmol/L leupeptin, 10 mmol/L benzamidine, 1 mmol/L phenylmethyl sulfonyl fluoride, and 12.5 µg/ml aprotinin. Lysates were subjected to 10% sodium dodecyl sulfate-polyacrylamide gel electrophoresis (10 µg of protein per lane) and transferred to Immobilon-P membrane. Membranes were incubated with primary antibody diluted in 1x phosphate-buffered saline (PBS), 2% dry milk, 0.1% Tween-20, and then with secondary antibody (peroxidase-conjugated anti-mouse or anti-goat). Antigen-antibody complexes were visualized using Lumiglo and chemiluminescent sensitive film.
Migration Assay
Cellular migration was measured using the Boyden chamber assay.18 Briefly, cells in EBM and 0.1% bovine serum albumin were pipetted into the upper wells of the chamber at 10,000 cells/well. Lower wells contained EBM and 0.1% bovine serum albumin (control) or 5% FBS, or growth factors diluted in EBM and 0.1% bovine serum albumin. Each condition was performed in quadruplicate. To quantitate, the number of cells in four high-power fields, each of which corresponded to 1.25 mm2, was counted. Therefore, the migration data represents the mean ± SE of 16 high-power fields from four different wells. Total area of the well equals 12.5 mm2.
Flow Cytometry
Cells were fixed in 0.1% paraformaldehyde in PBS at 4°C for 16
hours, permeabilized with 0.1% Triton X-100, incubated with either 2.6
µg/ml mouse anti--SMA or 2.6 µg/ml isotype matched
IgG2a for 1 hour at 4°C, and then with
fluorescein-conjugated goat anti-mouse IgG for 30 minutes at 4°C.
Cells were resuspended in 0.1% sodium citrate, 20 µg/ml RNase A,
0.3% Nonidet P-40, and 50 µg/ml propidium iodide at 200,000 cells/ml
and analyzed on a Becton Dickinson FACScan flow cytometer (Becton
Dickinson, Mountain View, CA) for fluorescein (-SMA) and propidium
iodide (DNA).
Immunohistochemistry of Human and Sheep Valves
Hearts were obtained from healthy sheep (n
= 7) and aortic valves were dissected. A portion of each valve
including the aortic wall was embedded in OCT compound. Human aortic
valves were obtained from autopsy cases (n = 3).
Serial frozen sections (6 µm) were prepared, fixed in 4%
paraformaldehyde for 5 minutes, and stained by the
avidin-biotin-peroxidase method. The peroxidase reaction was
visualized with 3-amino-9-ethyl carbazole. Sections were counterstained
with Gill’s hematoxylin solution. Frozen sections were incubated with
anti--SMA, followed by biotinylated secondary horse anti-mouse, and
Texas Red-conjugated streptavidin. Subsequently, specimens were treated
with an avidin-biotin blocking kit and stained with vWF or CD31
antibodies overnight at 4°C, then appropriate secondary antibodies,
and finally fluorescein-conjugated streptavidin was applied for 30
minutes. Specimens of human aortic valves were fixed in 10% formalin
and embedded in paraffin. Sections (6 µm) were stained by the
avidin-peroxidase method as described above. Immune complexes were
visualized using 3,3'5,5'-tetramethylbenzidine and counterstained with
methyl green.
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Results |
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TopAbstractIntroductionMaterials and MethodsResultsDiscussionReferences |
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A limiting dilution technique was used to isolate clonal EC
populations from primary cultures of ovine aortic valve leaflets. We
used immunofluorescence to examine whether the clonal cell populations
expressed the endothelial markers CD31 and E-selectin (Figure 1)
. Clone av 10 cells express CD31 at
cell-cell borders as expected for ECs (Figure 1a)
, but only diffuse
background staining was observed with control serum (Figure 1b)
.
E-selectin was detected in av 10 cells treated with lipopolysaccharide
(Figure 1d)
but only limited expression was detected in untreated cells
(Figure 1c)
, consistent with the known regulation of E-selectin in ECs.
-SMA was not detected in av 10 under normal culture conditions
(Figure 1e)
. A classic endothelial cobblestone morphology was
consistently observed in confluent cells (Figure 1f)
. Based on: 1) CD31
expression at cell-cell borders, 2) lipopolysaccharide-induced
up-regulation of E-selectin, 3) lack of -SMA, and 4) cobblestone
morphology, the clonal cells were designated ECs. Similar results were
obtained with six different clonal populations derived from aortic
valve and additional clones from ovine pulmonary valve leaflets (data
not shown).
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To determine whether clonal aortic valve-derived ECs could be
induced to transdifferentiate to a mesenchymal phenotype, we tested
whether TGF-ß1–3 isoforms could induce -SMA19
, a
marker of valvular transdifferentiation in heart development. A second
approach was to culture cells in low-serum medium, which is known to
promote differentiation in many cell types. Thus, we cultured clonal
cell populations in either growth medium with 1 ng/ml TGF-ß1 or in
medium containing no bFGF and 0.5 to 1% serum (reduced medium) for 6
days. The cells were then assayed for expression of CD31 and -SMA by
double-label immunofluorescence and by Western blot (Figure 2)
. Individual cells co-expressing CD31
and -SMA, appearing as yellow/orange, were seen in clone av 10 cells
treated with TGF-ß1 and in clone av 17 cells cultured in reduced
medium (Figure 2A)
, providing direct evidence for transdifferentiation.
Because TGF-ß1 and -ß3 at 1 or 10 ng/ml had similar effects on av
10 cells, TGF-ß1 at 1 ng/ml was used for subsequent experiments.
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-SMA by the addition of TGF-ß1 but not when cultured in reduced
medium (Figure 2A)
. Clone av 17 is representative of a second type of
response in which -SMA was not induced in response to TGF-ß1
(Figure 2A)
. All three TGF-ß isoforms were tested at 1 and 10 ng/ml
and found to have no effect on inducing -SMA in av 17 (data not
shown). However, av 17 expressed high levels of -SMA after culture
in reduced medium (Figure 2, A and C)
. Time course experiments showed
that TGF-ß-mediated and reduced serum-mediated induction of -SMA
in av 15 and av 17, respectively, peaked at 5 to 7 days (data not
shown).
The TGF-ß-induced up-regulation of -SMA in av 15 could be blocked
by adding a soluble TGF-ß type II receptor antagonist to the medium
concurrently with TGF-ß1 (Figure 2B)
. Similar results were obtained
with av 10 (data not shown). In contrast, the soluble TGF-ß receptor
antagonist had no effect on the induction of -SMA when either clone
av 17 (Figure 2C)
or av 14 (data not shown) was cultured in reduced
medium. This indicates that the induction of -SMA in these clones
does not occur by the generation of extracellular TGF-ß, but instead
occurs by a TGF-ß-independent pathway. It is possible, however, that
signaling pathways induced by culturing the cells in TGF-ß or in
reduced medium converge intracellularly to stimulate expression of
-SMA.
Besides its induction during valve development, -SMA is expressed in
smooth muscle cells, myofibroblasts, and dermal
fibroblasts.20,21
To determine whether induction of
-SMA in valvular ECs reflects a conversion to the smooth muscle
lineage, we examined expression of smooth muscle myosin heavy chain, a
definitive marker of the smooth muscle cell lineage,21
in
valve-derived cells (Figure 2D)
using a monoclonal antibody against the
SM-1 isoform of smooth muscle myosin heavy chain.22
Lysates of freshly resected ovine artery were used to demonstrate that
the antibody cross-reacts with ovine SM-1 (lane 1). Lysates from
cultured human dermal microvascular ECs were used as a negative control
(lane 2). Smooth muscle-specific myosin heavy chain was not detected in
clones av 14 (lanes 3 and 4), av 17 (lanes 5 and 6), or av 10 (lanes 7
and 8) before (-) or after (+) induction of -SMA in reduced medium
for 7 days or up to 21 days (data not shown), indicating that the
valvular -SMA-positive cells obtained using our methods are distinct
from classic arterial or venous smooth muscle cell. This is not
surprising given that smooth muscle myosin heavy chain in valve
interstitial cells has not been reported.
Increased Cellular Migration Coincides with Transdifferentiation
The induction of -SMA and loss of cobblestone morphology
suggested transdifferentiation to a mesenchymal cell phenotype. To
determine whether these observations coincided with alterations in cell
function, or with activities consistent with a mesenchymal phenotype,
we measured cell migration of clonal populations of transdifferentiated
cells. Basic FGF was tested because it is known to stimulate EC
migration, PDGF-BB was tested because of its role in recruitment and
assembly of mesenchymal cells during blood vessel
development,23
and TGF-ß was tested to determine whether
it also plays a role in cellular migration. Figure 3A
shows the results of a migration assay
using av 17 in response to 5% serum, 1 ng/ml bFGF, 10 or 50 ng/ml
PDGF-BB, or 1 ng/ml TGF-ß1. Clone av 17 cultured in reduced medium
exhibited greatly increased migration in response to PDGF-BB, but not
to TGF-ß or bFGF when compared to the control. The PDGF-BB-induced
migration in transdifferentiated av 17 cells was effectively blocked by
a neutralizing antibody against PDGF-BB, whereas control IgG at the
same concentration had no effect (Figure 3B)
. Clone av 15 cells
cultured in growth medium plus TGF-ß1 also exhibited increased
migration in response to PDGF-BB (Figure 3C)
. Results similar to those
shown in Figure 3, A and C
, were obtained with clones av 14 and av 10,
respectively (data not shown). Checkerboard analysis demonstrated that
the PDGF-induced migration of av 17 cells grown in reduced medium for 6
days was because of chemotaxis rather than random cell migration
because cells migrated directionally toward PDGF-BB in the bottom well
(Figure 3D)
. These results demonstrate that increased chemotaxis toward
PDGF-BB coincides with the induction of -SMA and the conversion to a
mesenchymal phenotype. The precise role that PDGF plays in valve
development and in normal ongoing function of postnatal valve leaflets
warrants further investigation.
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To determine whether or not the induction of -SMA was
reversible, cells that had up-regulated -SMA in response to TGF-ß1
or reduced medium were trypsinized and replated on fresh gelatin-coated
plates in normal growth medium for 6 to 8 days. In Figure 4A
, clone av 10 cells were plated in
growth medium and 24 hours later either transferred
to medium containing TGF-ß1 (lanes 2 and 3) or maintained in growth
medium (lane 1) for 7 days. At day 7, cells were trypsinized and
replated in normal growth medium (lanes 1 and 3) or in growth medium
containing TGF-ß (lane 2) for another 6 days. A parallel experiment
was performed using clone av 17 cells cultured in reduced medium
(Figure 4B)
. The induction of -SMA by TGF-ß1 or reduced medium was
not reversible throughout 6 days (Figure 4, A and B
; lane 3) or 8 days
(data not shown). Thus, the induction of -SMA seems to reflect a
permanent alteration in cellular phenotype rather than a transient
modulation of phenotype.
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. After 6 days in reduced medium, the av 17 cells were in
G0/G1 (top left panel). The
portion of quiescent cells that was replated in normal growth medium
with 10% serum and 2 ng/ml bFGF for 26 hours was harvested and
analyzed for -SMA expression. More than 90% of the cells expressed
-SMA (right panel). When these -SMA-positive cells were analyzed
for cell cycle distribution, approximately one third of the cells had
progressed into S and G2/M in 26 hours (bottom
left panel). These results demonstrate that av 17 cells induced to
transdifferentiate can proliferate in response to serum and bFGF. In
additional experiments, we found that mitomycin C did not block the
induction of -SMA (data not shown), indicating that cell
proliferation is not required for transdifferentiation.
To determine whether ECs from other vascular sites could also be
induced to transdifferentiate, we tested ECs derived from human
pulmonic valve, HDMECs, HUVECs, and ovine peripheral vein cultured in
either growth medium (G) or in reduced medium (R) (Figure 5A)
. Increased -SMA was seen in human
pulmonic valve ECs and in the positive control, av 17 cells, but not in
HDMECs, HUVECs, or ECs isolated from ovine peripheral vein (Figure 5A)
.
The modest level of induction of -SMA in human pulmonic valve ECs in
response to reduced medium, compared to av 17, was most likely because
of the fact that the pulmonary valve ECs were not clonal, and therefore
represented a heterogeneous population of ECs in which only some cells
were capable of transdifferentiation. As expected, TGF-ß1 caused no
induction of -SMA in HDMECs or HUVECs (Figure 5B)
. In summary, of
the vascular ECs tested, the induction of -SMA in response to
reduced medium or TGF-ß1 was specific to valve-derived ECs.
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To determine whether evidence of the transdifferentiation process
could be detected in vivo in mature valve leaflets, we
stained frozen sections of aortic valves from mature sheep with
anti-CD31 and anti--SMA (Figure 6)
.
Figure 6, A and B
, are adjacent sections stained with anti-CD31 and
anti--SMA, respectively, that demonstrate co-localization of these
two antigens in focal regions along the leaflet endothelium, suggesting
these ECs may be transdifferentiating. Figure 6, C and D
, show
low-power photomicrographs of the same sections to demonstrate
co-localization of CD31 and -SMA occurs in subregions of the
endothelial layer, and that CD31 was expressed predominantly along one
side of the leaflet, whereas -SMA was most abundant in the
subendothelial region adjacent to the base of the leaflet.
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-SMA on frozen sections of ovine
aortic valve (Figure 6, E and F)
. In Figure 6E
, vWF-positive ECs
(green), double-labeled vWF-positive/-SMA-positive cells
(yellow/orange) and -SMA-positive cells (red) were observed. The
double-labeled cells and cells expressing only -SMA seem to
represent intermediates in the transdifferentiation process. Similar
results were observed in human aortic valve sections (data not shown).
Classic blood vessel architecture was observed in the ovine aortic root
and served as a positive control in this analysis (Figure 6F)
. In
paraffin sections from human aortic valve leaflets, -SMA-positive
cells were seen along the endothelium and also seemed to be migrating
into the subendothelial regions (Figure 6G)
, similar to what was
observed in ovine aortic valve (Figure 6E)
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Discussion |
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TopAbstractIntroductionMaterials and MethodsResultsDiscussionReferences |
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To our knowledge, the only nonvalvular ECs reported to express -SMA
are bovine aortic ECs treated with TGF-ß1.19
In this
study, the -SMA expression induced by TGF-ß1 was reversible.
Endothelial-to-mesenchymal transdifferentiation has also been suggested
to occur in the embryonic quail dorsal aorta, raising the possibility
that transdifferentiation may give rise to subendothelial
-SMA-positive cells in other vascular beds.24
We
cultured ECs from several sources to determine whether any of these
cells would transdifferentiate in response to TGF-ß1 or reduced
medium. We saw transdifferentiation only in valvular ECs and not in ECs
from microvascular or venular sources. Hence, the valve-derived clonal
cell populations described here provide a unique and powerful in
vitro model for elucidating the molecular events that regulate
valvular endothelium.
TGF-ß and the type II and type III TGF-ß receptors have been implicated in the endothelial-to-mesenchymal transdifferentiation that underlies the formation of the endocardial cushions during heart development.3-6,25,26 Furthermore, attenuation of TGF-ß signaling seems to be critical for proper development of the aortic and pulmonary valve leaflets, because mice lacking the inhibitor of TGF-ß signaling, SMAD6, have an overabundance of mesenchymal cells that results in hyperplastic thickening of the valves.27 In addition, a number of genes, such as neurofibromin-1,28 NF-ATc,29,30 and endoglin31 have been found to be essential for normal valve development based on phenotypes observed in mice genetically deficient in these gene products. For the most part, the valvular defects in these mice were not anticipated and underscore how little is known about the molecular basis of valve development.
Based on our observations, we hypothesize that adult valvular endothelium contains a subset of cells capable of undergoing transdifferentiation to a mesenchymal cell phenotype. These cells may serve to replenish the interstitial cells, which in turn synthesize the extracellular matrix needed for maintenance of the leaflet architecture. Such cells would need to be capable of prolonged self-renewal and might, therefore, be thought of as progenitor-like cells. This speculation is supported by the fact that clonal EC populations were readily obtained from ovine pulmonary and aortic valve leaflets and were expanded easily for at least 20 passages. Further studies will be required to determine the significance of these cells for valve function and durability in vivo.
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Acknowledgements |
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Footnotes |
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Supported by the National Heart Lung and Blood Institute (grant R01 HL60490), the Vascular Pathology Core at the Brigham and Women’s Hospital (HL48743 PPG), and a fellowship from the Sarnoff Foundation (to S. K.).
Accepted for publication June 26, 2001.
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References |
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TopAbstractIntroductionMaterials and MethodsResultsDiscussionReferences |
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