Harvard Medical School and Children's Hospital, Boston, Massachusetts 02115
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Abstract |
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We aimed to determine if and how endothelial cells (EC) recruit precursors of smooth muscle cells
and pericytes and induce their differentiation during
vessel formation. Multipotent embryonic 10T1/2 cells
were used as presumptive mural cell precursors. In an
under-agarose coculture, EC induced migration of
10T1/2 cells via platelet-derived growth factor BB.
10T1/2 cells in coculture with EC changed from polygonal to spindle-shaped, reminiscent of smooth muscle
cells in culture. Immunohistochemical and Western blot analyses were used to examine the expression of
smooth muscle (SM)-specific markers in 10T1/2 cells
cultured in the absence and presence of EC. SM-myosin, SM22, and calponin proteins were undetectable in
10T1/2 cells cultured alone; however, expression of all
three SM-specific proteins was significantly induced in 10T1/2 cells cocultured with EC. Treatment of 10T1/2
cells with TGF-
induced phenotypic changes and
changes in SM markers similar to those seen in the cocultures. Neutralization of TGF-
in the cocultures
blocked expression of the SM markers and the shape change. To assess the ability of 10T1/2 cells to contribute to the developing vessel wall in vivo, prelabeled
10T1/2 cells were grown in a collagen matrix and implanted subcutaneously into mice. The fluorescently
marked cells became incorporated into the medial layer
of developing vessels where they expressed SM markers. These in vitro and in vivo observations shed light
on the cell-cell interactions that occur during vessel development, as well as in pathologies in which developmental processes are recapitulated.
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Introduction |
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THE vasculature is among the first organ systems to develop, and is vital for the distribution of nutrients and oxygen as well as removal of waste products. Blood vessels in general are composed of distinct cell layers. The intima, the innermost layer, is made up of a single cell type: endothelial cells (EC).1 The media is composed of layers of mural cells, smooth muscle cells (SMC) in large vessels, and pericytes in microvessels. The outermost layer of large vessels, the adventitia, consists of loose connective tissue containing smaller blood vessels and nerves.
Blood vessel assembly during embryogenesis, a process termed vasculogenesis, begins with clustering of primitive vascular cells also known as hemangioblasts (13) into tubelike endothelial structures that define the pattern of the vasculature (41). Mural cells become associated with the forming vessels at later stages of development (26), which has led to the suggestion that the EC may govern vessel layer acquisition. Although EC are thought to be a homogeneous population of cells derived entirely from mesodermal tissue (8, 48), SMC are locally derived from one of two sources: cardiac neural crest (i.e., ectoderm) or lateral mesenchyme (i.e., mesoderm; reference 28). Recruitment of pericytes to nascent capillaries during angiogenesis is likely to involve similar processes.
The exact mechanism(s) by which EC recruit mural cell
precursors and induce their differentiation into mural cells
during vessel formation is unknown, and is difficult to examine using in situ developmental models. There has been
intense interest in understanding the regulation of muscle
cell differentiation. Much progress has been made in elucidating the molecular regulators of skeletal and cardiac
muscle differentiation (42, 43). These rapid advances have
been facilitated by the availability of culture systems that
can be used to model the differentiation process in vitro.
Although the identification of a number of smooth muscle (SM)-specific proteins, including SM-myosin, calponin,
and SM22 (14, 19) has enabled the description of SM development in vivo, understanding the molecular regulation
of SM differentiation has been hindered by the lack of an
appropriate in vitro system.
Thus, we have developed in vitro coculture systems in order to dissect and analyze the cellular interactions and potential mediators involved in SMC recruitment and differentiation. We have used the multipotent mouse embryonic 10T1/2 cells as presumptive mural cell precursors. These cells have been used in other systems, and are induced by various means to express markers of adipocyte (33), osteoblast (17), and myoblast cell lineages (10). We report a novel and biologically relevant coculture system in which we demonstrate recruitment and differentiation of a multipotent precursor toward a SMC lineage in response to EC.
The studies described herein demonstrate that growth factors released from EC induce migration of undifferentiated mesenchymal cells toward EC, and that, upon contact with the EC, the mesenchymal cells differentiate into SM-like cells. We hypothesize that these processes operate in vivo during vessel assembly via vasculogenesis and angiogenesis. We further postulate that contact and communication between EC and mural cells is important not only for establishing new vessels, but also for maintaining the differentiated quiescent vessel.
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Materials and Methods |
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Tissue Culture
EC were isolated from bovine aortas by collagenase digestion as previously described (18). The cells were grown on uncoated tissue culture plastic in DME with 10% calf serum (CS) supplemented with penicillin, streptomycin, and glutamine, and were used through passage 20. 10T1/2 cells (CCL 226; American Type Culture Collection, Rockville, MD) were grown and maintained in DME with 10% FCS and 4.5 g/liter glucose. All cells were maintained at 37°C in a humidified atmosphere. DME with 2% CS was used in all coculture experiments. All tissue culture experiments were performed at least three times. Representative experiments are shown.
Cocultures
Cocultures of bovine aortic EC (BAE) and 10T1/2 cells were established in one of two ways: (a) an under-agarose system assay was conducted with modifications of a previously described assay (40). EC and mural cell precursors were plated into two 5-mm wells created 1-2 mm apart in 1% agarose/1% BSA in DME. 10T1/2 cells were plated in one well, and EC, a control vehicle, or growth factors were added to the other well. Cells were incubated for up to 6 d at 37°C; or (b) direct cocultures in which equal numbers of both cell types were plated simultaneously into the same plate and incubated for up to 48 h. This assay was used when a large number of interacting cells was needed (e.g., for Western blot analyses).
Migration Assay
BAE and 10T1/2 cells were plated in the under-agarose assay at 2 × 104 cells/well in 25 µl DME with 2% CS as described above, and were allowed to incubate for 48 h. The cells were then fixed with 2% paraformaldehyde in PBS for 30 min and stained with 0.1% Coomassie blue for 30 min. The distance migrated by each cell type was quantified by projecting the image of each well on a Profile Projector Model 6C-2TM (Nikon, Inc., Melville, NY) under a low-power objective fitted with a grid. In some experiments, neutralizing antibodies against PDGF-A or -B (1:200; Genzyme Corp., Cambridge, MA) or basic fibroblast growth factor (bFGF; provided by Dr. Michael Klagsbrun, Children's Hospital, Boston, MA; used at 1:100) were incorporated into the agarose before plating. In all experiments, the distance migrated by 10T1/2 cells cultured alone (background) was subtracted from the distances migrated by the cells under experimental conditions.
Immunocytochemistry
EC and 10T1/2 cells were cocultured in the under-agarose assay established in four-chamber culture slides (Lab-tek, Naperville, IL). In some
cases, neutralizing antiserum against TGF- was added at the initiation of
the cocultures. Before plating, EC were labeled with 5 µg/ml Di-I-acetylated low density lipoprotein (Ac-LDL; Biomedical Technologies, Inc.,
Stoughton, MA) for 4-18 h at 37°C. After coculture for up to 6 d, the cells
were fixed with 4% paraformaldehyde and immunostained for the presence of SMC-specific markers, including
SM-actin, SM-myosin, calponin, and SM22
. Anti-
SM-actin (mouse monoclonal antibody; DAKO
Corp., Carpinteria, CA) was used at 1:50 in blocking buffer consisting of
4% normal goat serum/3% BSA/0.1% Triton X-100 in PBS. Anti-SM-myosin heavy chain mouse monoclonal antibody (9A9 used at 1:200) provided
by Dr. Gary Owens (University of Virginia Medical School, Charlottesville, VA) has been shown to cross-react with both isoforms of SM myosin
heavy chain (50). This antibody specifically recognized SMC, and not EC,
in bovine aorta (data not shown). SMC specificity is further demonstrated
in Fig. 8, where antibody labels only vascular SMC, and not surrounding
skeletal muscle. Similar labeling patterns were obtained using a polyclonal
SM myosin antisera from Biomedical Technologies, Inc. Anti-calponin
(mouse monoclonal antibody provided by Dr. Marina Glukhova, Curie
Institute, Paris, France) was used at 1:10. Affinity-purified rabbit polyclonal SM22
antisera was generously provided by Dr. Mario Gimona
(Salzburg, Austria), and was used at 1:500. All antibody-antigen complexes were visualized using the Vectastain Elite ABC Kit (Vector Labs,
Inc., Burlingame, CA), and the biotinylated secondary antibodies (1:250)
were provided by the manufacturer.
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Western Blot Analysis
EC and 10T1/2 cells were cultured alone (6 × 105 cells/100 mm dish) or in
coculture (3 × 105 of each cell type/100 mm dish) for 48 h with or without
a monoclonal anti-TGF- antibody that recognizes TGF-
1, 2, and 3 (10 µg/ml; generously provided by Genzyme Corp., Boston, MA). Protein
was isolated from cells using a method previously described (53); 10 µg of
protein from solo cultures and 20 µg of protein from cocultures were electrophoresed on SDS-PAGE minigels (Bio-Rad Laboratories, Hercules,
CA) and electrophoretically transferred to Immobilon-P membranes
(Millipore Corp., Bedford, MA). Membranes were blocked with 5% nonfat dry milk/1% BSA in PBS, and then incubated for 1-2 h with primary
antibody (
SM-actin, 1:50; SM-myosin, 1:150; SM22
, 1:1,000; calponin,
1:10,000) diluted in blocking buffer. Mouse monoclonal antibodies against
calponin were purchased from DAKO Corp. Similar results were obtained with a rabbit polyclonal antisera against SM-myosin (Biomedical
Technologies, Inc.) and mouse monoclonal antibodies (9A9). Antibody-
antigen complexes were revealed using the ECL detection system (Amersham Life Sciences, Arlington Heights, IL). Quantitation was performed
using a Digital Imaging SystemTM (Alpha Innotech Corp., San Leandro, CA).
Integration of 10T1/2 Cells into Vessels In Vivo
These studies were conducted to determine if undifferentiated 10T1/2 cells would become incorporated into newly forming vessels and display a SM phenotype in a complex in vivo environment. 10T1/2 cells were permanently labeled with a fluorescent dye and implanted subcutaneously onto the backs of 6-8-wk-old C57 mice.
Permanent Labeling of 10T1/2 Cells. 10T1/2 cells were prelabeled with PKH26 (Sigma Chemical Co., St. Louis, MO), a permanent red fluorescent dye that is retained in the cells for up to 100 cell doublings (25). Cells were trypsinized to a single-cell suspension, rinsed with PBS, and counted using a Coulter Counter (Coulter Corp., Miami, FL). Cells were gently resuspended at 2 × 106 cells/100 µl of Diluent C (provided with the PKH26 dye). An equal volume of 40 µM PKH26 dye was added, followed by incubation for 2-3 min with gentle agitation. The labeling reaction was terminated by adding 2 vol of FCS and 7 ml of 0.1% BSA in PBS. The cells were then layered onto 3 ml of FCS and pelleted. After centrifugation, the cells were rinsed with DME/10% FCS, resuspended in fresh media, and incubated overnight at 37°C. Cells were then trypsinized, counted, and plated as needed.
Mouse Implantation Assay.
The fluorescently-labeled 10T1/2 cells were
seeded at various densities (4-20 × 104 cells/cm2) onto 0.3-mm2 pieces of
cross-linked collagen matrix (InstatTM; Johnson & Johnson Medical, Inc.,
Arlington, TX) in the presence or absence of 30 ng bFGF and incubated
for 3 d at 37°C. Cell/matrix preparations were implanted subcutaneously
onto the backs of 6-8-wk-old C57 mice and incubated for 7-14 d. Tissue
from the experimental area was subsequently excised, fixed in 4%
paraformaldehyde, embedded, sectioned, and immunostained for SM-actin (1:500), SM-myosin (9A9; 1:2,000) and calponin (1:10). Stained sections were analyzed with an Axiophot epifluorescent microscope (Carl
Zeiss, Inc., Thornwood, NY) or an Odyssey XL confocal microscope with
Intervision (Noran Instruments, Middleton, WI).
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Results |
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Effects of EC on 10T1/2 Cell Migration
10T1/2 cells, a multipotent cell line, were used as presumptive mural cell precursors. EC and 10T1/2 cells were cocultured in the under-agarose assay to examine the effects of EC on migration of the presumptive mural cell precursors. 10T1/2 cells and BAE were plated into two wells created ~2 mm apart in an agarose matrix. The 10T1/2 cells were cocultured for 48 h with either BAE or media (DME with 2% CS) as a control. At the end of the incubation period, the cells were fixed and visualized by staining with 0.1% Coomassie blue, and migration was measured using images cast with a Profile Projector. 10T1/2 cells cultured alone (in the absence of BAE) exhibited a pattern of random movement, migrating uniformly from all edges of the circle. In contrast, 10T1/2 cells cocultured with BAE migrated toward the BAE in a directional manner (Fig. 1 a). A higher magnification view revealed that the 10T1/2 cells at the front of the well closest to the BAE migrated as single cells, extending long pseudopodia in the direction of the EC (Fig. 1 b). 10T1/2 cells at the back of the well furthest from the BAE also migrated outward, but traveled a shorter distance. In addition, these 10T1/2 cells moved as a sheet, and were flat, spread cells (Fig. 1 c) and not at all elongated as in the case of the cells closer to the BAE.
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10T1/2 cells in the presence of BAE migrated ~500-700 µm above the background migration (~100 µm) occurring in the absence of EC (set to zero in Fig. 2). Parallel experiments, conducted using bovine capillary EC, yielded similar results. As a test of the specificity of this effect, 10T1/2 cells were cocultured with Madin-Darby canine kidney (MDCK) cells; 10T1/2 in coculture with MDCK cells were not stimulated to migrate further than the cells cultured alone (data not shown).
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Proliferating EC have been shown to synthesize various diffusible factors, including PDGF and FGF, which can act as chemoattractants for SMC and other mesenchymally derived cells (64, 67). To determine if PDGF was mediating the EC-directed migration of the 10T1/2 cells, we first assessed its direct effects on 10T1/2 cell migration. 10T1/2 cells were plated in the presence of 20 ng/ml of each of the three PDGF isoforms. PDGF-BB was the most potent chemoattractant, followed by PDGF-AB and PDGF-AA; the 10T1/2 cells migrated 1559 ± 62, 1013 ± 36, and 512 ± 5 µm above background, respectively.
We then examined the role of PDGF in the EC-10T1/2 cell cocultures. When neutralizing antibodies against PDGF-B were added to the agarose before establishing the EC-10T1/2 cell coculture, EC-stimulated migration of 10T1/2 cells was completely inhibited. Neutralizing antibodies against PDGF A or bFGF (not shown) had no effect on EC-induced 10T1/2 cell recruitment (Fig. 2). Although in this experiment PDGF-B-neutralizing antisera caused some suppression of background 10T1/2 cell migration, and PDGF-A antisera caused mild stimulation, neither of these effects was reproducible among numerous experiments.
There is obviously an important role for proliferation in vessel formation that may also be mediated by EC-derived PDGF; we are examining this possibility in a separate series of studies. However, to eliminate the possible contribution of EC-induced 10T1/2 cell proliferation to the migration observed in the under-agarose assay, the assay was also conducted with 10T1/2 cells that had been growth- arrested with mitomycin C (45). The magnitude of EC-stimulated 10T1/2 cell migration was equivalent to that observed in assays conducted with proliferating 10T1/2 cells (data not shown).
Effects of EC on 10T1/2 Cell Phenotype
We hypothesized that EC might induce differentiation of
mesenchymal cells into mural cells, that is, SMC and pericytes. To assess the effects of EC on the phenotype of
10T1/2 cells, 10T1/2 cells and BAE were cultured alone or
together in the under-agarose assay for 3-6 d, allowing
the two cell types to come into contact. For these experiments, BAE were prelabeled with the fluorescent molecule Di-I-Ac-LDL so that the EC in the cocultures could
be distinguished from the 10T1/2 cells. All cocultures were
subsequently immunostained for SM-actin, SM-myosin,
calponin, or SM22
. The SM myosin results shown were
obtained with monoclonal antibody 9A9; staining with a
rabbit polyclonal anti-SM-myosin antisera yielded similar
results. Importantly, none of the antibodies used to assess
the SMC phenotype cross-reacted with skeletal muscle. This result is demonstrated in Fig. 8, where the antibodies
specifically recognized vascular SMC and not the surrounding skeletal muscle.
BAE exhibited negligible staining for SM-actin (Fig. 3
a), SM-myosin (Fig. 3 b), calponin (Fig. 3 c), and SM22
(Fig. 3 d). 10T1/2 cells cultured alone exhibited a low but
detectable level of
SM-actin staining (Fig. 3 e) and less labeling for SM-myosin (Fig. 3 f), calponin (Fig. 3 g), and
SM22
(Fig. 3 h). However, when the 10T1/2 cells contacted the BAE, the levels of immunoreactivity for
SM-actin (Fig. 3 i), SM-myosin (Fig. 3 j), calponin (Fig. 3 k),
and SM22
(Fig. 3 l) were significantly increased. Furthermore, in the presence of EC, the 10T1/2 cells underwent a
dramatic shape change from polygonal to spindle-shaped
(Fig. 3, i-l). BAE in the coculture were distinguished from
the 10T1/2 cells by fluorescent visualization of EC-associated Di-I-Ac-LDL (Fig. 3, m-p; BAE are indicated by arrows), and in many cases were seen to lie underneath the
10T1/2 cells.
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Western blot analyses revealed that SM-actin, SM-myosin, SM22
, and calponin proteins are all expressed specifically by SMC (Fig. 4 a). A faint SM-myosin band was occasionally observed in BAE with both the monoclonal
(9A9) and polyclonal (BTI) antibodies, and has been reported elsewhere (6). Changes in SM-specific protein expression were examined by Western blot analyses in experiments in which 10T1/2 cells and BAE were plated
alone or simultaneously in direct cocultures (1:1) for 48 h.
There was a 20-fold increase in
SM-actin protein level in
10T1/2 cells cocultured with BAE over that in 10T1/2 cells
cultured alone (Fig. 4 b). SM-myosin, SM22
, and calponin were undetectable in 10T1/2 cells cultured alone, but
were significantly induced in 10T1/2 cells cocultured with
EC (Fig. 4 b). 10T1/2 cells cocultured with BAE did not
express skeletal muscle actin or type II collagen, and did
not demonstrate lipid accumulation, indicating that under
these coculture conditions, 10T1/2 cells were not induced
toward skeletal muscle, chondrocyte, or adipocyte lineages (data not shown).
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In control studies not shown, increased expression of SM-specific proteins was also seen in 10T1/2 cells cocultured with either bovine or rodent capillary EC, indicating that these cell-cell interactions are neither species- nor vessel size-specific. Similar phenotypic changes, including increased expression of mural cell-specific markers and altered cell shape, occurred in bovine adventitial fibroblasts, human dermal fibroblasts, and human placental fibroblasts cocultured with aortic EC. However, well-differentiated cell types such as MDCK did not express SMC-specific markers when cocultured with EC. Conversely, MDCK cells did not induce a mural cell phenotype in either the 10T1/2 cells or the fibroblasts.
Mechanism of EC-Induced 10T1/2 Cell Differentiation
We were next interested in determining the factor(s) involved in the EC-induced SMC differentiation. TGF- has
been shown to influence
SM-actin expression in myofibroblasts (11), pericytes (66), and SMC (44), and we have
previously shown that cocultures of EC and SMC activate
TGF-
(2). Therefore, we hypothesized that TGF-
was
involved in induction of the SMC phenotype in the EC-10T1/2 cocultures. To test this hypothesis, 10T1/2 cells
were treated with TGF-
to determine whether the phenotypic changes observed in coculture could be mimicked
by exogenous addition of this regulatory factor. Indeed,
treatment of 10T1/2 cells with 1 ng/ml TGF-
1 for 24 h led
to morphological changes similar to those seen in the
BAE-10T1/2 cell cocultures: a change from a flat, polygonal to a polarized and elongated shape (Fig. 5). To determine if changes in SM-specific protein expression accompanied the shape change in 10T1/2 cells, Western analysis
was performed on similarly treated cells. Importantly, we
found that TGF-
-treated 10T1/2 cells exhibited increased
expression of
SM-actin, SM-myosin proteins (Fig. 5, b
and d), and SM22
(data not shown) compared with untreated controls (Fig. 5, a and c). These TGF-
-induced alterations in cell shape and differentiation were not accompanied by a change in cell proliferation.
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Next, neutralizing antibodies against TGF- (10 µg/ml)
were added to the agarose before the coculture of prelabeled BAE and 10T1/2 cells in an under-agarose assay.
The cocultures were allowed to incubate for 4 d followed
by staining for
SM-actin and SM-myosin. The intensity of
immunostaining for both proteins was significantly less in
the 10T1/2 cell-EC cocultures cultivated in the presence of
neutralizing TGF-
antisera (Fig. 6, b and e, respectively), compared with untreated cocultures (Fig. 6, a and d, respectively). The fluorescently-labeled EC of the cocultures
incubated with anti-TGF-
antibodies are visualized in
Fig. 6, c and f. Western blot analyses revealed that the neutralization of TGF-
suppressed the levels of
SM-actin
by 95% (Fig. 7 a), and SM-myosin protein expression by
87% (Fig. 7 b). The specificity of this effect was demonstrated in parallel studies in which PDGF-B-neutralizing antisera were shown to have no effect on 10T1/2 cell differentiation. Cellular toxicity due to the neutralizing antibody to TGF-
was ruled out in related studies (Hirschi et
al., manuscript submitted for publication) showing that the
antibody did not affect the growth of either BAE or 10T1/2
cells.
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Endothelial-Mesenchymal Interactions In Vivo
Our in vitro observations indicated that EC can recruit undifferentiated mesenchymal cells and direct their differentiation into smooth muscle. To determine if mesenchymal cells can become recruited toward, and incorporated into, newly forming vessels in a more complex in vivo environment, fluorescently-labeled 10T1/2 cells embedded in a cross-linked collagen matrix were implanted subcutaneously onto the backs of mice.
After 10 d in the mouse, the experimental tissue was excised, and the newly formed vessels were examined for the
presence of the labeled 10T1/2 cells. Examination of the
sections by fluorescent microscopy revealed that the fluorescently-labeled 10T1/2 cells had become associated with
vessels in tissue from the back of the mouse; red fluorescent labeling is specifically seen in cells associated with the
vessel wall (arrowhead; Fig. 8). Furthermore, once incorporated into the vessel, the prelabeled cells stained strongly for SM-actin, SM-myosin, and calponin. Importantly, none of the SM-specific antisera cross-reacted with
skeletal muscle tissue seen in these sections, demonstrating the specificity of these reagents for vascular SMC. Interestingly, in parallel studies conducted in the chick
chorioallantoic membrane assay, prelabeled 10T1/2 were
observed to become incorporated into other nonvascular portions of the chorioallantoic membrane, but did not express SM markers at these sites (data not shown).
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Discussion |
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Development of the vascular system is a complex process that begins with the formation of simple tubes. In vasculogenesis, endothelial precursors called angioblasts associate to form vessel tubes. Major axial vessels such as the aorta, and the vascular plexus of endodermal organs such as the spleen, are formed by vasculogenesis (8, 48). In angiogenesis, vessels form by sprouting from pre-existing vessels. Vascularization of neural and mesodermal tissues, like the brain and limb buds, occurs by angiogenesis (15, 47, 61). In both cases, the primitive vessel tubes then remodel, forming the more complex architecture of the adult vasculature. Developmental studies suggest that the EC in these nascent tubes may govern the acquisition of additional vessel layers (23, 38). Knowledge of the mechanism(s) by which EC recruit mural cell precursors and enable their differentiation remains unknown, and would significantly add to the understanding not only of blood vessel development, but of pathological conditions such as atherosclerosis or vascular malformations, in which these basic processes appear to be deregulated.
Our studies demonstrate that EC do indeed recruit undifferentiated mesenchymal cells. Furthermore, EC-derived
PDGF-BB mediates the mesenchymal movement. We
found that the ability of BAE as well as bovine or rodent
capillary EC to recruit 10T1/2 cells (presumptive mural cell-precursors) was totally blocked by a neutralizing antibody
to PDGF-B. PDGF has been shown in previous in vitro
studies to act as a mitogen and chemoattractant for mesenchymal cells (67). Neutralizing antibodies to PDGF-A or to
bFGF had no effect on EC recruitment of 10T1/2 cells, even
though these factors have been shown to be mitogenic and
chemotactic for SMC and fibroblasts. These findings suggest that EC-derived PDGF-B specifically exerts a paracrine effect on undifferentiated mesenchymal cells during vascular development in vivo. Observations by Holmgren
et al. (24) on the expression of PDGF ligand and receptors
in forming blood vessels of human placenta, support this
concept. They found that EC of developing blood vessels
express the mRNA and protein for PDGF-B, but not the
PDGF- receptor, whereas PDGF-
receptor mRNA was
detectable in fibroblast-like cells and SMC surrounding intermediate and large blood vessels.
The phenotypes of mice deficient for components of the
PDGF system lend further support to our hypothesis.
Patch mice that have a spontaneous mutation deleting the
PDGF- receptor are embryonic lethal, and have cardiovascular defects characterized by reduced numbers of SMC
(56). Mice lacking PDGF-B also exhibited severe cardiovascular and renal abnormalities (31). The lack of mesangial cells in the kidneys of PDGF-B null mice is of particular
interest, since mesangial cells are considered to be specialized pericytes and developmentally related to SMC (57).
Indeed, more recently, PDGF-B null mice have been reported to lack pericytes (32).
Our studies show that once undifferentiated mesenchymal cells are recruited to EC, they are induced to become
SMC-like. Mature SMC and pericytes in intact vessels express a series of proteins that define the SMC lineage (for
review see reference 58). These include, but are not limited to, SM-actin (46, 65), SM-myosin (29, 52), calponin
(3, 63), SM22
(14, 30, 58), h-caldesmon (59), and desmin
(39). A variety of cells including astrocytes (20) and myofibroblasts (11) have been reported to express
SM-actin in
culture. Therefore, induction of
SM-actin in an otherwise
undifferentiated cell is not evidence for differentiation towards an SMC lineage. Furthermore, a few of these proteins expressed specifically in adult SMC are also transiently expressed in other cell types (e.g., cardiac muscle)
during embryogenesis (36, 37). However, simultaneous expression of multiple SM-specific proteins is indicative of a
SM phenotype. Hence, simultaneous expression of
SM-actin, SM-myosin, SM22
, and calponin in 10T1/2 cells in
culture with EC, along with a dramatic change in cell shape, provides strong evidence that these mesenchymal
cells are truly induced toward the smooth muscle lineage.
Certainly it is possible that this coculture system will have
limitations and will not reproduce all aspects of differentiated SMC function. However, the biologically relevant induction of SM-specific protein expression in 10T1/2 cells,
which can be easily genetically manipulated, make this an
ideal system in which to investigate the molecular aspects
of SM-specific gene expression.
We demonstrated that induction of 10T1/2 to an SMC
phenotype occurs only after the mesenchymal cells come
into close proximity with the EC. This sequence of events
has been shown to occur during development of the aorta
in the quail. Hungerford et al. found that mesodermal cells
become associated with the developing aorta, beginning at
the ventral surface, and that they express SM-actin only
after associating with the EC tube (26). By recreating aspects of vessel assembly in vitro, we have begun to investigate directly the mechanisms involved in this EC-mesenchymal interaction. We have found that the induction of
the SMC phenotype in this model is mediated at least in
part by TGF-
. In addition to showing that direct treatment of 10T1/2 cells with TGF-
1 induces expression of
SM-specific markers, we have demonstrated that neutralizing antibodies against TGF-
blocks induction of SM-specific markers in the cocultures.
Although the cells in these cocultures make contact, it is
not clear if physical contact between the cells is necessary
for the inductive event. Speculation regarding the source
of the TGF- in the cocultures is based on previous studies
from our lab and others, showing that both EC and mural
cells, when grown separately, produce a latent form of
TGF-
that is activated in EC-mural cell cocultures (2,
55). The demonstration that local activation of TGF-
may be involved in directing undifferentiated mesenchymal cells to an SMC lineage in the vessel wall is consistent
with previous observations of elevated TGF-
in mesenchymal remodeling, and at sites of epithelial-mesenchymal interactions (22, 49).
An increasing body of evidence indicates roles for members of the TGF- family in regulating muscle differentiation in general. TGF-
has been implicated in the regulation of both cardiac and skeletal muscle differentiation
(for review see references 42 and 43). The epithelial-mesenchymal transformation in embryonic heart can be mimicked by TGF-
(49), and antisense against TGF-
3 blocks
epithelial-mesenchymal transformation of cardiac endothelial cells (49). Expression of a dominant-negative form
of the type II TGF-
receptor suppressed myogenic differentiation in a culture model (16). Furthermore, data from
gene disruption studies suggest that a new member of the
TGF-
family, growth/differentiation factor-8, acts as a
negative regulator of skeletal muscle growth (35).
Whether all of the effects of TGF- on SM-specific gene
expression in this model are direct remains to be determined. To date, regulatory regions identified as conferring
responsiveness to TGF-
have been found within the 5'
flanking sequences of
SM-actin (21). The 5' and 3' regulatory regions of other SMC-specific genes, including SM-myosin and SM22
, are under investigation (27, 68). In addition, some of the effects of TGF-
on vascular wall cell
differentiation may be due to TGF-
-induced changes in
the extracellular matrix (34). It is well known that the matrix can influence cell growth, polarity, and organization as well as differentiation (1, 62). In fact, the interaction of
presumptive SMC and pericytes with the abluminal EC
surface are temporally associated with deposition of a
basement membrane (9). In this regard, it is interesting to
speculate that TGF-
, activated when mural cells associate
with EC, may serve several functions, all aimed at establishing a mature vessel. These include inhibition of endothelial proliferation (45) and migration (54), stimulation of
SMC/pericyte differentiation (our present observations), and induction of basement membrane assembly (34).
Observations of mice with a targeted disruption of TGF-
support this concept. Fifty percent of TGF-
1 null mice
die in utero from a defect in yolk sac vasculogenesis, which
is thought to be due to improper interactions between epithelial cells and mesenchymal cells, and may be the result
of altered cell-matrix interactions (12). This observation
also suggests that local activation of TGF-
, resulting from
such cell-cell interactions, may occur widely throughout
development. Whether TGF-
activation is a constitutive event at the site of EC-mural cell contact is not known.
The reversibility of the SMC phenotype (4, 7) and the concept that differentiation may require constant signaling,
at least under some circumstances (5), leads us to suspect
that local activation of TGF-
is an ongoing process. In
fact, it is well-documented that the endothelium makes frequent contacts with SMC and pericytes throughout the vasculature (51, 60).
Although we have shown that 10T1/2 cells are capable
of becoming SMC-like in vitro in response to EC, we
wanted to confirm that 10T1/2 cells have the capacity to
become incorporated into developing vessels in vivo.
Therefore, undifferentiated 10T1/2 cells were permanently
labeled with a fluorescent dye, and were placed in the
proximity of developing vessels in collagen matrices subcutaneously in the mouse. Not only did these cells become
incorporated into the medial layers of the newly forming
vessels (Fig. 8), but once associated with the vessel, simultaneously expressed SM-actin, SM-myosin, and calponin,
reflective of a SM phenotype. The resolution was not sufficient to comment on the role of contact in the induction of
the SMC phenotype in these vessels. However, the 10T1/2
cells expressing SM markers usually comprised the innermost layer of cells in the vessel wall.
In summary, we have developed coculture systems to identify potential regulators of vessel formation, and to elucidate their relative contributions. A complimentary in vivo system was established, and observations of developing vessels in the model corroborate our tissue culture observations. We believe that the information gained from this system regarding the cellular and molecular regulation of vessel formation will be important in understanding not only developmental regulation, but also pathophysiological processes such as atherosclerosis and vascular malformations, where there appear to be defects in the normal control mechanisms.
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Footnotes |
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Received for publication 26 November 1997 and in revised form 2 March 1998.
Address all correspondence to Patricia A. D'Amore, Laboratory for Surgical Research, Children's Hospital, Enders 1061, 300 Longwood Ave., Boston, MA 02115. Tel.: 617-355-8377. Fax: 617-355-7043. E-mail: damore_p{at}a1.tch.harvard.eduThe authors thank Dr. Shmuel Ben-Sasson, Dr. Diane Darland, Laurence Beck, and Sandra Smith for their critical reading of the manuscript. We gratefully acknowledge the contribution of Evelyn Flynn in immunohistochemistry of the mouse tissue. We thank Dr. Gary Owens, Dr. Marina Glukhova, and Dr. Mario Gimona for their generosity with their antibody reagents.
This work was supported by National Institutes of Health grants EY05318 and CA45548 to P.A. D'Amore. Dr. Karen Hirschi is supported by a National Institutes of Health Postdoctoral Fellowship (HL09037), and Dr. Stephanie Rohovsky was supported by the Harvard-Longwood Research Training Program Grant in Vascular Surgery (HL 07734).
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Abbreviations used in this paper |
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Ac-LDL, acetylated low density lipoprotein; BAE, bovine aortic EC; CS, calf serum; EC, endothelial cells; SM, smooth muscle; SMC, smooth muscle cells.
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