1 Program in Human Molecular Biology and Genetics, University of Utah, Salt Lake
City, UT, USA
2 Department of Medicine and Oncological Science, University of Utah, Salt Lake
City, UT, USA
3 Hospital Sant Pau, Barcelona, Spain
4 Minnesota Cardiovascular Research Institute, Minneapolis, MN, USA
5 Howard Hughes Medical Institute, Department of Cell Biology, Harvard Medical
School, Department of Cardiology, Children's Hospital, Boston, MA, USA
Author for correspondence (e-mail:
dean.li{at}hmbg.utah.edu)
Accepted 19 October 2002
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SUMMARY |
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Key words: Elastin matrix, Vascular smooth muscle, Morphogenesis, G-protein signaling, Vascular proliferative diseases
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INTRODUCTION |
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Vascular smooth muscle cells are not terminally differentiated and can
alternate between a quiescent, contractile state and a proliferative
non-contractile state (Raines and Ross,
1993; Owens, 1998
;
Thyberg, 1998
). In a healthy
artery, vascular smooth muscle cells are quiescent and largely comprise a
contractile apparatus that functions to dilate and constrict the lumen as
required by physiological demands. Actin stress fibers serve as the scaffold
for the contractile apparatus, and are a hallmark of mature and quiescent
vascular smooth muscle cells (Burridge and
Chrzanowska-Wodnicka, 1996
;
Small and Gimona, 1998
). Under
circumstances of injury, repair and regeneration, vascular smooth muscle cells
lose their contractile apparatus and dedifferentiate into an immature
phenotype capable of proliferating and depositing extracellular matrix
(Owens, 1998
;
Thyberg, 1998
). This
fibrocellular response plays an important role in all forms of vascular
proliferative diseases. In atherosclerotic lesions, the major components of
the fibrous plaque are vascular smooth muscle cells, the matrix products
deposited by these cells, and extracellular cholesterol
(Ross, 1995
;
Lusis, 2000
;
Raines and Ross, 1993
). In
restenosis, transplant arteriopathy and vascular graft disease; smooth muscle
cells are the predominant component of the occlusive lesion
(Raines and Ross, 1993
). The
phenotypic modulation of vascular smooth muscle cells offers a tempting target
for preventing vascular proliferative diseases.
Elastin is the dominant extracellular matrix protein deposited in the
arterial wall and can contribute up to 50% of its dry weight
(Parks et al., 1993). The
protein product of the elastin gene is synthesized by vascular smooth muscle
cells and secreted as a monomer, tropoelastin. After post-translational
modification, tropoelastin is crosslinked and organized into elastin polymers
that form concentric rings of elastic lamellae around the arterial lumen. Each
elastic lamella alternates with a ring of smooth muscle, and provides the
compliance that arteries need to absorb and transmit hemodynamic force
(Wolinski et al., 1967
). There
is a growing body of evidence that implicates elastin in vascular development
and disease. We previously demonstrated that loss-of-function mutations of one
elastin allele cause supravalvular aortic stenosis (SVAS) and Williams-Beuren
Syndrome (Li et al., 1997
;
Curran et al., 1993
;
Ewart et al., 1993
). These
disorders are characterized by discrete fibrocellular stenoses in the aorta,
coronary arteries, carotid arteries, pulmonary arteries and other peripheral
arteries. Often, individuals with these diseases are young children who are
susceptible to peripheral vascular disease, myocardial infarctions or stroke
in the absence of other risk factors such as high serum cholesterol, diabetes
and cigarette smoking (van Son et al.,
1994
). In subsequent experiments, we showed that the deposition of
elastin matrix in the arterial wall during late fetal development is essential
to arterial morphogenesis (Li et al.,
1998
). Mice that lack elastin (Eln-/-) died
from occlusive fibrocellular pathology caused by subendothelial proliferation
and accumulation of vascular smooth muscle cells in early neonatal life. In
Eln-/- vessels, there is no evidence of abnormal
endothelial structure, activation or proliferation. Furthermore, the occlusive
vascular phenotype occurred in the absence of hemodynamic stress and
inflammation. This work demonstrated that elastin is required for arterial
development. However, because the absence of elastin undoubtedly distorts the
presentation and stability of other important elements of the matrix
architecture, these studies failed to define the specific role for elastin in
establishing and maintaining a mature artery.
Here we show that elastin regulates the phenotypic modulation, proliferation and migration of vascular smooth muscle cells in culture. We confirm a direct signaling effect by demonstrating that elastin regulates vascular smooth muscle cells via a G-protein-coupled signaling pathway. Finally, we establish that in vivo, exogenous elastin reduces the vascular proliferative response of an injured artery. These results demonstrate that elastin is a crucial signaling molecule that directly controls vascular smooth muscle biology and stabilizes arterial structure.
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MATERIALS AND METHODS |
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Cellular assays
Cellular experiments of proliferation, actin polymerization and migration
described below were performed on at least three independently isolated
primary cell lines. Proliferation was assayed by cell count and
[3H]thymidine incorporation. Confluent cultures of
Eln+/+ and Eln-/- vascular smooth
muscle cells from the fourth passage were seeded at a density of
2x103 cells/well on a plastic 24-well plate (Corning Costar,
Corning, NY), and stimulated to proliferate in AmnioMAX C-100 growth medium
(Gibco-BRL) treated with 100 µg/ml recombinant tropoelastin or left
untreated. Cell numbers for each culture were assayed by hemocytometer after
24, 48 and 72 hours of incubation. For [3H]thymidine incorporation,
Eln+/+ and Eln-/- cells were seeded at
a density of 20,000 cells/well on a 24-well plate. After attachment, cells
were starved in 0.1% BSA (Fisher) in Amniomax Basal Medium (Gibco) for 24
hours. Cells were then grown in whole Amniomax medium treated/untreated with
100 µg/ml tropoelastin (Grosso et al., 1991a). Cells were assayed for
[3H]thymidine incorporation using a scintillation counter after 24
hours by precipitating with 5% TCA, followed by NaOH solubilization. Data were
calculated as mean±s.d. (n=6).
Subconfluent cultures of vascular smooth muscle cells were evaluated and
scored for the presence of actin stress fibers in the cytoplasm following
immunofluorescent staining for SM -actin (Clone 1A4, Sigma), vinculin
(Clone hVin-1, Sigma), desmin (Sigma) and tubulin (Clone DM-1A, Calbiochem).
Cells were treated with tropoelastin (1 µg/ml) (Grosso et al., 1991a) or
not, and assays were performed in serum-free media. Cells were scored as
contractile if they had distinct actin stress fibers continuous throughout the
cytoplasm, or at least 10 well-defined focal adhesions distributed throughout
the cytoplasm and cell periphery. Scoring was performed by three separate
observers blinded to the cell genotypes, and their numbers averaged. At least
100 cells were scored in three separate cultures for each genotype.
Additionally, identical results were obtained for cells cultured on glass
slides, 24 plastic tissue culture plates (Corning) coated with matrigel, and
another brand of plasticware (Falcon).
The chemotactic activity of cells was assayed in a modification of the Boyden chamber method using 6.5 mm transwell polycarbonate chemotaxis filter inserts in a plastic 24-well tissue culture plate (Corning Costar, Corning, NY). Various concentrations of recombinant tropoelastin (Grosso et al., 1991a) were placed in both the upper or lower compartment and covered with a polycarbonate membrane filter (8 µm porosity). PDGF-BB (30 ng/ml) was added to the lower chamber and served as a positive control. Medium alone was used as the negative control. Cells to be tested were placed in the upper wells of the chamber and incubated at 37°C/5% CO2 for 3 hours. After incubation, the cells adhering to the upper surface of the filter were scraped off and the cells that had migrated to the lower surface were fixed with 10% buffered formalin, stained with DAPI (Molecular Probes, Eugene, OR) and viewed under a mercury lamp. Fifteen randomly selected high power (x200) fields were counted on each filter by observers blinded to their conditions. Chemotactic response was measured as the number of cells that had transversed the filter in response to tropoelastin, PDGF and collagen type I (10 µg/ml), and expressed as fold increase over baseline. Each condition was assayed in triplicate wells and each experiment was repeated at least three times. Actin polymerization and migration assays experiments were also performed in triplicate in the presence of 100 µM Actinomycin D (Calbiochem), 10 µg/ml Cycloheximide (Calbiochem), 20 µM Y27632 (Welfide), 20 µM EDTA, 100 ng/ml Ptxn (Calbiochem) or 100 ng/ml B-protomer (Calbiochem).
Measurement of cAMP and activated Rho A
All biochemical measurements were performed in cells that were serum
starved overnight. cAMP assay levels were detected using an RIA kit (RPA538,
Amersham Pharmacia). Briefly, cells were plated at a density of 80,000 to
100,000 cells on a plastic 12-well cell-culture plate, pretreated with 100
ng/ml cholera toxin (Calbiochem) for 3 hours, treated with 1 µg/ml
recombinant tropoelastin for 20 minutes, and cAMP subsequently isolated.
Experimental cAMP levels were quantified through use of a standard curve where
known amounts of cAMP were added to radioimmunoassay.
RhoA-GTP was detected using a Rho Activation Kit (Upstate Biotechnology). Briefly, Eln-/- cells were grown to 70% confluence in a six-well plate, and pretreated with 100 ng/ml pertussis toxin or 100 ng/ml B-protomer for 3 hours. Control experiments with no pretreatment were also performed. Cells were then treated with 1 µg/ml recombinant tropoelastin for 3 hours and activated Rho A was isolated by immunoprecipitations with Rhotekin coated beads. RhoA-GTP was detected by western blot analysis after samples were run on a 10% acryllamide gel and transferred to nitrocellulose membranes. Equal amounts of total cellular lysate were run out for each sample to ensure that equivalent amounts of protein were used in all experiments.
Porcine coronary model of in-stent restenosis
Elastin sheaths were prepared from dissected common carotid arteries of
adult domestic swine (40 to 60 kg) using a variation of a previously published
protocol (Malone et al.,
1984). Briefly, these vessels were sequentially treated with a 1%
SDS solution supplemented with doxycycline (10 mg/l) and EDTA (5 mM),
potassium hydroxide (5 N) at 60°C, collagenase solution (0.5 mg/ml,
Collagenase D, Roche), autoclaved and
irradiated. The product of these
extractions, a tubular elastin matrix sheath, was cut to size and fitted over
14 mm long stainless steel expandable stents (Medtronic AVE, S670) mounted on
3.5 mm diameter angioplasty balloons and secured by a metal coil. Control
stents deployed were identical to elastin covered stents except for the
absence of elastin sheaths.
Nineteen domestic pigs (30 to 40 kg) on a normal diet were pretreated with
oral aspirin (625 mg), ticlopidine (250 mg) and verapamil (120 mg), and placed
under general anesthesia. Animals underwent placement of 32 stents in the left
anterior descending, circumflex or right coronary artery. The methods of stent
implantation have been previously described
(Schwartz et al., 1994).
Briefly, control stents or elastin sheath-covered stents were advanced under
fluoroscopic guidance to an appropriate site in the coronary vasculature and
deployed at a 1.2:1 to 1.4:1 stent-to-artery ratio compared with the baseline
vessel diameter. After the procedure, the wounds were closed and the pigs were
returned to their quarters on a normal diet. Pigs were sacrificed at day 3
(elastin-covered stent, n=2) and day 14 (elastin-covered stent,
n=2) to assess biocompatibility of the elastin sheath. To study
neointima formation, pigs were sacrificed at day 28 (control stents,
n=15; elastin-covered stents, n=14) and instrumented
arteries were removed. Coronary arteries were fixed by pressure perfusion (100
mm Hg) with 10% buffered formalin for 24 hours, dissected free, washed,
dehydrated through graded alcohols and infiltrated with methylmethacrylate
(MMA) at 4°C. After polymerization, embedded specimens were
cross-sectioned at 5 µm thickness using a tungsten-carbide microtome blade.
Each arterial segment was stained with Hematoxylin and Eosin and elastic Van
Gieson stains. Three cross-sections taken from the two ends and the middle of
each artery were used for histomorphometric analysis. Histological examination
confirmed that the elastin sheaths were retained in the arterial wall of all
specimens.
Histomorphometric analysis
Mean neointimal thickness, percent stenosis and mean injury score were
measured on Van Gieson-stained sections using calibrated digital microscopic
planimetry as previously described
(Schwartz et al., 1992;
Schwartz et al., 1994
).
Briefly, the mean neointimal thickness was measured by drawing a radial line
from the lumen border to the point maximum penetration for each stent strut,
and averaging the measurements. Percent area stenosis was calculated as %
stenosis=100 x [1-(stenotic lumen area/original lumen area)]. The
original lumen area was measured as the area subtended within the internal
elastic lamina. The injury score measures the physical penetration of the
stent strut into the vessel wall and was calculated using a graded scale from
0-3 as previously described (Schwartz et
al., 1992
). Briefly, a score of 1 indicated that only the internal
elastic lamina was lacerated; a score of 2 indicated laceration of the
arterial media; and a score of 3 indicated that the external elastic lamina
was lacerated. The mean injury score for each histological section was
calculated as mean injury score=
injury scores for every strut/number
of struts present per section.
Statistics
For all in vitro experiments, mean and standard deviations were calculated
and statistical analysis were carried out by analysis of variance (ANOVA). For
in vivo experiments, statistical analysis was performed on injury score,
average neointimal thickness and percent lumen stenosis using regression
modeling (Schwartz et al.,
1994). Linear regression modeling accounts for injury (a strong
covariate) and the injury-dependent neointimal thickening between control
arteries and elastin treated arteries. Three models were used to establish
whether there were: (1) differences in intercepts, (2) differences in slopes
allowing any intercept or (3) differences in slopes when the intercepts are
fixed. Differences between the elastin sheath-stent and control stent at each
injury level were analyzed using the Tukey-Kramer multiple comparisons
t test for all three regression models.
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RESULTS |
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To evaluate the role of elastin in vascular smooth muscle cell proliferation, we compared the growth rate of Eln+/+ and Eln-/- cells. Cells were seeded at the same density and growth rates were assayed by counting cells after 24, 48 and 72 hours. The number of Eln-/- cells was greater than Eln+/+ cells at all time points, and was increased over twofold at 72 hours (Fig. 1D). These data indicate that in the absence of elastin synthesis, vascular smooth muscle cells proliferate at an increased rate.
To prove the specificity of the effect of elastin on vascular smooth muscle
cell growth, we tested whether adding exogenous recombinant tropoelastin to
the culture media inhibits cellular proliferation. Recombinant tropoelastin
was synthesized using a bacterial expression system (Grosso et al., 1991a) and
determined to be pure by gel electrophoresis and amino acid composition.
Tropoelastin had a dose-dependent inhibitory effect on proliferation, with a
maximal effect at 100 µg/ml (Fig.
1D; data not shown). At this dose, the number of
Eln-/- cells was nearly identical to
Eln+/+ cells at 72 hours. At each time point, more than
98% of all cells were viable as determined by Trypan Blue staining, indicating
that tropoelastin was not cytotoxic. Similar results were observed when acid
hydrolyzed elastic fibers, elastin, were used for this assay. The
responses of Eln+/+ and Eln-/-
cellular proliferation to tropoelastin
(Fig. 1E) and
elastin
(data not shown) were reproduced in [3H]thymidine incorporation
assays. By comparison, there was no significant inhibitory effect on
proliferation when Eln-/- cells were treated with type I
collagen (100 µg/ml) (data not shown). These data demonstrate that elastin
regulates vascular smooth muscle cell proliferation.
Elastin induces a mature contractile phenotype in vascular smooth
muscle cells
The primary function of mature differentiated vascular smooth muscle cells
is contraction. This ability requires highly organized actin myofilaments,
often referred to as actin stress fibers
(Burridge et al., 1996;
Small and Gimona, 1998
). To
determine if elastin modulates the phenotype of vascular smooth muscle cells,
we evaluated actin organization in Eln+/+ and
Eln-/- cells using direct immunofluorescent staining with
FITC-conjugated SM
-actin antisera. Well-defined actin myofilaments
were apparent in more than 95% of Eln+/+ cells scored by
observers blinded to genotype (Fig.
2A,E). By contrast, only 23% of Eln-/- cells
had actin stress fibers (Fig.
2B,E). Furthermore, Eln-/- cells with actin
stress fibers were qualitatively different than Eln+/+
cells, with a distinctive, rounder morphology (data not shown). These data
indicate that vascular smooth muscle cells lacking elastin fail to organize
their contractile apparatus.
|
To confirm that the failure to form an organized contractile apparatus in
Eln-/- cells resulted from the inability to synthesize and
secrete elastin, we treated these cells with recombinant tropoelastin protein.
Within 3 hours of exposure to tropoelastin, the percentage of
Eln-/- cells with organized actin stress fibers increased
threefold to 74% (Fig. 2D,E).
This response was does dependent and was also seen by phallodin staining for
F-actin (data not shown). By comparison, tropoelastin treatment had not effect
on actin stress fiber organization in Eln+/+ cells
(Fig. 2C,E). Similar results
were observed when Eln-/- cells were treated with
crosslinked elastin (data not shown). However, no change in actin
stress fiber organization was observed when Eln-/- cells
were treated with an equivalent concentration of type I collagen. In addition,
the induction of actin polymerization by tropoelastin was unaffected by the
presence or absence of serum in culture (data not shown). Thus, extracellular
elastin is an important mediator of actin polymerization and contractile
apparatus organization in vascular smooth muscle cells.
The Rho signal transduction pathway is known to be a central converging
step in the formation of actin stress fibers through a post-translational
mechanism (Mack et al., 2001;
Bishop and Hall, 2000
). We
investigated the role of this pathway in the regulation by tropoelastin of
actin polymerization. The addition of actinomycin D
(Greenburg et al., 1986
), an
inhibitor of gene transcription, or cycloheximide
(Greenburg et al., 1986
), an
inhibitor of protein translation, did not block tropoelastin-induced actin
polymerization in Eln-/- cells
(Fig. 2E). Furthermore, a
specific inhibitor of the Rho signaling pathway that targets Rho kinase,
Y27632 (Mack et al., 2001
;
Ushata et al., 1997
), blocked
tropoelastin mediated actin polymerization of Eln-/- cells
(Fig. 2E). Additionally,
tropoelastin treatment of Eln-/- vsmc did not alter the
amount of SM
actin transcript or protein levels as quantified by
northern and western blot analysis but did shift the filamentous:globular
(F:G) actin ratio from 1:1 to 3.1:1 (data not shown). These experiments, in
combination with the failure of either transcription or translation inhibitors
to inhibit tropoelastin-induced actin stress fiber formation in
Eln-/- vsmc, suggest that elastin modulates vsmc phenotype
by regulating actin treadmilling via a signal transduction pathway involving
Rho GTPases and their effector proteins.
To evaluate further the effect of elastin on the contractile phenotype in
vascular smooth muscle cells, we examined the organization of vinculin,
tubulin and desmin in Eln+/+ and
Eln-/- cells (Burridge
et al., 1996). Vinculin is concentrated in focal adhesion plaques
that bind the actin cytoskeleton and connect with the cell membrane. Indirect
immunofluorescent staining for vinculin revealed abundant focal adhesion
plaques in 94% of Eln+/+ cells when scored by observers
blinded to genotype (Fig.
2F,J). By contrast, only 12% of Eln-/- cells
had a normal distribution of defined focal adhesion plaques
(Fig. 2G,J). Within 3 hours of
exposure to recombinant tropoelastin protein, the percentage of
Eln-/- cells with defined focal adhesions increased 6-fold
to 79% (Fig. 2I,J). Treatment
had no effect on focal adhesion organization in Eln+/+
cells (Fig. 2H,J).
Immunostaining for other cytoskeletal proteins, tubulin
(Fig. 2K,L) and desmin (data
not shown) showed no difference between Eln+/+ and
Eln-/- cells. Thus, the loss of elastin in vascular smooth
muscle cells does not lead to a broad disruption of cytoskeletal architecture.
These results indicate that elastin induces a mature, contractile phenotype in
vascular smooth muscle cells by regulating the organization of specific
cytoskeletal proteins.
Elastin controls the migration of vascular smooth muscle cells
Vascular proliferative disease involves the migration of smooth muscle
cells from the arterial media to the subendothelial space, forming a neointima
(Schwartz, 1997). The same
phenomenon is observed during the development of Eln-/-
arteries. We postulated that the elastin matrix surrounding each concentric
layer of vascular smooth muscle cells provides a cue that localizes these
cells to the arterial media, and prevents their migration to the neointima. To
test this hypothesis, we used a modified Boyden chamber chemotaxis assay. We
discovered that vascular smooth muscle cells migrate to tropoelastin in a
dose-dependent manner (Fig.
3A). Cell migration depended on the concentration gradient of
tropoelastin, not the total amount used in the assay. Chemotaxis was minimal
for cells in control (0/0 ng/ml) and static fields of tropoelastin (200/200
ng/ml), but increased significantly as the concentration gradient of
tropoelastin rose to 200 ng/ml. A similar but less dramatic effect was seen
with cells that produce elastin (Fig.
3A), suggesting that the synthesis and secretion of elastin by
Eln+/+ cells reduced migration to external stimuli. These
data demonstrate that elastin is a potent chemoattractant for vascular smooth
muscle cells.
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Cytokines and growth factors, such as platelet-derived growth factor
(PDGF), are thought to mediate the subendothelial migration of vascular smooth
muscle cells in occlusive vascular lesions
(Lusis, 2000;
Bornfeldt et al., 1993
). To
determine if elastin matrix deposited in the arterial media can counteract the
potent chemotactic activity of PDGF, we repeated the Boyden chamber
experiments. At a PDGF concentration of 30 ng/ml, Eln-/-
cells showed a fourfold increase in chemotactic activity
(Fig. 3B). Increasing
concentrations of tropoelastin prevented Eln-/- cell
migration towards PDGF (Fig.
3B). Similar results were observed in Eln+/+
cells. These data suggest that elastin provides a local cue that prevents
vascular smooth muscle cells from migrating away from the arterial media to
the subendothelial space. Together, these in vitro experiments demonstrate
that elastin is a potent and specific regulator of vascular smooth muscle cell
maturation, migration and proliferation.
Elastin signals via a non-integrin, G-protein coupled signaling
pathway
To determine whether elastin has a direct signaling effect, we examined the
molecular mechanism of the regulation by elastin of vascular smooth muscle
cells. Though various binding proteins, chaperones and other matrix elements
that interact with elastin have been identified and cloned, the mechanism of
elastin signaling remains to be elucidated
(Mecham and Hinek, 1986;
Hinek et al., 1988
;
Mecham et al., 1991
; Grosso et
al., 1991b; Hinek, 1995
;
Hinek, 1996
;
Privitera et al., 1998
). Using
in vitro assays of migration and actin polymerization, we examined the
signaling cascade stimulated by elastin in Eln-/- vascular
smooth muscle cells. Above, we demonstrate that elastin mediates actin
polymerization through a Rho mediated signal transduction pathway
(Fig. 2E). Cell-surface
receptors that are known to regulate the Rho signaling pathway include the
integrins and G-protein-coupled receptors
(Seasholtz et al., 1999
;
Wei et al., 2001
). Integrins
are a well-characterized family of receptors that recognize matrix proteins
such as collagen, vitronectin, fibulin and fibronectin. Although elastin is
not a known ligand for integrins, recent work suggests that the integrins may
be involved in elastin signaling through an intermediary, fibulin 5
(Nakamura et al., 2002
;
Yanagisawa et al., 2002
).
Fibulin 5 interacts directly with elastin, and serves as a ligand for
cell-surface integrins. Integrins require extracellular divalent cations to
bind their matrix ligands, and low doses of chelators such as EDTA block these
cell-matrix interactions (Brockdorff et
al., 1998
). As expected, control experiments demonstrated that
integrin mediated migration of vascular smooth muscle cells to collagen was
EDTA sensitive (Fig. 4A).
However, EDTA did not interfere with tropoelastin-mediated migration or actin
polymerization of Eln-/- cells
(Fig. 4A,B). In addition,
blocking antibodies to integrins known to bind other matrix proteins such as
collagen and fibulin 5 did not perturb elastin-mediated actin polymerization
or chemotaxis (data not shown). Thus, these data strongly suggest that the
integrin family of receptors is not involved in elastin signaling.
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G-protein-coupled receptors (GPCR) are the largest family of transmembrane
receptors and are known to activate Rho kinase
(Seasholtz et al., 1999;
Marinissen and Gutkind, 2001
).
GPCRs signal through heterotrimeric G proteins classified into four protein
families: Gs, Gi, Gq and G12/13
(Marinissen and Gutkind,
2001
). Pertussis toxin, a specific inhibitor of Gi
(Thomas et al., 2000
), blocked
tropoelastin-mediated migration and actin polymerization of vascular smooth
muscle cells (Fig. 4C,D). The
specificity of pertussis toxin inhibition was demonstrated in a series of
control experiments. First, migration of vascular smooth muscle cells to
platelet-derived growth factor is not dependent on a G-protein-coupled
signaling pathway and was not disrupted by pertussis toxin
(Fig. 4C). Second, pertussis
toxin is composed of two subunits, A and B
(Thomas et al., 2000
). The A
protomer inhibits Gi by ADP ribosylation, while the B protomer
facilitates the entry of the toxin into the cell. In the presence of B
protomer alone, there was no disruption of either migration or actin
polymerization (Fig. 4C,D). Using a Ca2+-sensitive dye, fura-2, and immunoprecipitations with
specific antibodies, we detected no evidence that rapid Ca2+
influx, tyrosine kinase receptors or the Map kinase pathway was involved in
elastin signaling (data not shown). Together, these experiments indicate that
elastin signaling is mediated via a non-integrin and pertussis toxin-sensitive
G-protein-coupled signaling pathway.
The pertussis toxin-sensitive G protein, Gi, inhibits adenylate
cyclase, the enzyme responsible for generating cAMP
(Thomas et al., 2000). To
obtain direct biochemical evidence that tropoelastin signals through
Gi, we measured the level of cAMP in tropoelastin-treated cells. To
measure a reduction in cAMP level, it was necessary to increase the baseline
cellular levels of cAMP using either forskolin or cholera toxin, agents that
activate adenylate cyclase (Thomas et al.,
2000
; Coward et al.,
1999
). When tropoelastin was added to either forskolin or cholera
toxin pretreated cells, the reduction in cAMP levels was between 40 and 60%,
indicating that Gi was activated and inhibited adenylate cyclase
activity (Fig. 4E). These
reductions in cAMP levels are comparable with those observed with other
ligands known to activate Gi via known G-protein-coupled receptors
(Thomas et al., 2000
;
Coward et al., 1999
). The
reduction in cAMP was pertussis toxin sensitive, which confirmed the role of
Gi. Thus, tropoelastin activates Gi and reduces cAMP
levels. Finally, consistent with the activation of Gi, we found no evidence
for rapid Ca2+ influx, a prominent feature of Gq
activation, in our cellular system. Although GPCRs activate the Gi
pathway and ligands for many of these receptors have not been identified, our
data does not rule out the possibility that elastin might indirectly activate
the GPCR pathway at the level of the heterotrimeric G proteins.
Others have shown that the GPCRs pathway can trigger the Rho signaling
cascade (Seasholtz et al.,
1999; Wei et al.,
2001
; Kabarowski et al.,
2000
). In this cascade, activated RhoA stimulates Rho kinase. We
examined whether tropoelastin stimulated the RhoA signaling pathway via
activation of a G-protein-coupled signaling pathway. Immunoprecipitation
experiments demonstrate that tropoelastin activates RhoA through a pertussis
toxin-sensitive mechanism (Fig.
4F). Together, our pharmacological and biochemical data lead us to
propose a molecular mechanism for elastin signaling
(Fig. 4G). In vascular smooth
muscle cells, elastin activates a pertussis toxin-sensitive G-protein-coupled
pathway that stimulates Gi, inhibits adenylate cyclase, reduces
cAMP levels and stimulates Rho induced actin polymerization. In the absence of
elastin synthesis, this mechanism is disrupted and vascular smooth muscle
cells lose their contractile phenotype. Thus, there is a direct role for
elastin in controlling vascular smooth muscle cells.
Elastin reduces the vascular proliferative response to arterial
injury in vivo
The in vitro experiments described above demonstrated that elastin is an
autocrine factor that induces a contractile state, inhibits proliferation and
localizes vascular smooth muscle cells to the vessel wall. These data suggest
that disruption of a crucial morphogenic signal in the vessel wall may release
smooth muscle cells to dedifferentiate, proliferate and occlude mature
arteries. To test this hypothesis in vivo, we used a porcine model to
determine whether the application of exogenous elastin to a site of vascular
injury would reduce the neointimal accumulation of smooth muscle cells and
arterial stenosis.
Porous sheaths of elastin matrix were generated from porcine carotid
arteries using established methods (Malone
et al., 1984). The purity of the elastin was confirmed by scanning
electron microscopy (Fig. 5A),
immunohistochemistry, amino acid composition and desmosine content
(Starcher, 1977
) (data not
shown). The amino acid composition and the concentration of the crosslinking
amino acids desmosine, isodesmosine and lysinonorleucine revealed no
microfibrillar proteins or other impurities. Elastin sheaths were secured to
intracoronary stents (Fig. 5B)
and successfully deployed in porcine coronary arteries using standard
catheterization techniques. The biocompatibility of elastin sheaths was
assessed 3 days and 14 days after placement in the porcine coronary artery.
Elastin sheaths did not evoke an inflammatory or thrombotic response (data not
shown). Moreover, elastin sheaths were biologically stable and did not degrade
within the arterial wall during the course of animal studies. These
experiments demonstrate that elastin sheaths can be used to restore elastin
matrix to sites of vascular injury.
|
To determine if elastin sheaths would reduce vascular smooth muscle
accumulation and neointimal formation, we used an established porcine coronary
injury model of in-stent restenosis
(Schwartz et al., 1992;
Schwartz et al., 1994
). Two
major coronary arteries of domestic pigs received either a control stent or an
elastin sheath-covered stent following vascular injury caused by balloon
overexpansion. Four weeks after injury and stent placement, representative
cross-sections taken from control arteries displayed a thick fibrocellular
neointima (Fig. 5C). By
contrast, neointimal formation was substantially reduced in elastin
sheath-treated arteries (Fig.
5D). Standard measurements of mean neointimal thickness (NIT) and
percent stenosis were measured and correlated with the degree of injury
induced by stent placement. The well-established correlation between the
severity of injury and the amount of neointimal accumulation
(Schwartz et al., 1994
) is
reproduced in our experiments with the control stents. Throughout the range of
injury scores, there is a significant reduction in the NIT and percent
stenosis in elastin sheath-treated arteries, with the greatest benefit at the
highest injury scores (Fig.
5E,F). At a mean injury score of 2, when stent placement disrupts
internal elastic lamella and lacerates the arterial media, elastin sheaths
reduced the mean neointimal thickness by 52% as compared with controls. The
specificity of the effect of elastin as a stent coating is supported by the
work of our group and others with non-elastin sheaths. Sheaths made from
collagen, fibrin or synthetic biopolymers failed to reduce neointimal
thickness compared with bare stents and were frequently associated with a
worse outcome (Goodwin et al.,
2000
; McKenna et al.,
1998
; van der Giessen et al.,
1996
; van Beusekom et al.,
1998
). Thus, consistent with the in vitro results, our in vivo
experiments indicate that restoration of elastin matrix to a site of injury
reduces vascular smooth muscle accumulation and limits neointimal
formation.
![]() |
DISCUSSION |
---|
![]() ![]() ![]() ![]() ![]() ![]() ![]() |
---|
Elastin-VSMC signaling is a critical morphogenic signal that is
disrupted during the pathogenesis of vascular disease
Vascular smooth muscle cells are able to exist in a quiescent contractile
state or a proliferative non-contractile state
(Raines and Ross, 1993;
Owens, 1998
). This plasticity
enables the vascular system to regenerate and grow. However, plasticity must
be balanced by the need to maintain a mature and stable structure capable of
circulating blood throughout a whole animal. Because vascular smooth muscle
cells modulate their phenotype readily, external factors must instruct them to
remain in a mature state if homeostasis is to be achieved. Our data indicate
that the elastin matrix is a potent autocrine factor that regulates arterial
morphogenesis by instructing vascular smooth muscle cells to localize around
the elastic fibers in organized rings and remain in a quiescent, contractile
state (Fig. 6A). This
cell-matrix interaction is mediated via a heterotrimeric G-protein signaling
pathway that activates downstream rho GTPases and appears to be crucial for
stabilizing and maintaining the structure of the mature artery. When this
morphogenic signal is absent during arterial development, the unregulated
migration and proliferation of vascular smooth muscle cells results in
occlusion of the arterial lumen (Li et
al., 1998
) (Fig.
6B). This leads us to propose that focal disruption or destruction
of the elastin matrix in the mature artery by factors such as mechanical
injury or inflammation play an important and direct role in the fibrocellular
response characteristic of proliferative vascular diseases
(Fig. 6C). Previously, emphasis
was placed on the pivotal role of inflammatory cells in regulating vascular
smooth muscle cells through the secretion of cytokines and growth factors
(Lusis, 2000
;
Ross, 1995
). In our model, the
degradation of elastin by macrophages, T-cells and their proteases acts to
release vascular smooth muscle cells from their mature contractile state to
migrate, proliferate and form a neointima. Thus, the disruption of an
essential morphogenic signal contributes to the pathogenesis of vascular
disease.
|
Four lines of evidence support our model of elastin-vascular smooth muscle signaling in the pathogenesis of vascular proliferative diseases.
Pathology
Disruption of elastin matrix is consistently associated with vascular
proliferative diseases in human pathological specimens
(Sandberg et al., 1981;
Sims, 2000
;
Sims et al., 1989
). Moreover,
the severity of occlusive vascular pathology increases in proportion to the
magnitude of defects and discontinuities in the elastin matrix
(Sims et al., 1989
). It has
also been noted that the programmed intimal hyperplasia and arterial occlusion
required for the closure of the ductus arteriosis is associated with impaired
elastic fiber formation (Hinek et al.,
1991
).
Genetic
We previously showed in human genetic studies that loss-of-function
mutations in elastin is sufficient to cause a human vascular proliferative
disease, supravalvular aortic stenosis and Williams-Beuren syndrome
(Li et al., 1997;
Curran et al., 1993
; Ewart,
1993). In these diseases, there is an aggressive occlusive pathology that
develops throughout the arterial tree of affected children in the absence of
common risk factors for vascular disease.
Experimental
In murine gene-targeting experiments, we demonstrated that loss of elastin
is sufficient to cause occlusive vascular pathology
(Li et al., 1998). This
pathology was caused by the unregulated proliferation, migration and
accumulation of vascular smooth muscle cells in the subendothelial space
(Fig. 6B). In this model,
severe arterial obstruction occurs in the absence of an inflammatory response
or hemodynamic stress.
Therapeutic
In this manuscript, we show that exogenous tropoelastin can control the
proliferation, migration and maturation of vascular smooth muscle cells in
vitro, and reduce the development of fibrocellular pathology in vivo. Other
investigators have shown that matrix metalloproteinase inhibitors, which
prevent the degradation of matrix components, can reduce neointimal formation
(Zaidi et al., 2000).
Together, these data indicate that the destruction of the elastin matrix is a
critical step in the fibrocellular response characteristic of vascular
proliferative disorders. Although strongly indicative, definitive proof of a
role for elastin signaling in a vascular proliferative response requires
further studies.
Elastin is unique among vascular extracellular matrix proteins
The extracellular matrix is known to play a crucial function in the
regulation of vascular smooth muscle cell biology. The myriad associations and
interaction between the many structural proteins, proteoglycans and growth
factors of the vascular matrix makes it difficult to distinguish the effects
of each element from one another. However, numerous in vitro studies have
demonstrated the ability of matrix proteins such as collagen, fibronectin and
laminin to affect vascular smooth muscle cell activity, including phenotypic
modulation, migration and proliferation
(Raines, 2000;
Hedin et al., 1999
). These
data might suggest that there is overlap and redundancy with regard to the
function of different vascular matrix proteins. Elastin, however, is unique
among matrix elements in that the disruption of this gene leads to a vascular
proliferative pathology in human and animal models. Disruption of other genes
encoding vascular matrix proteins, including fibulin, fibrillin and collagen
is associated with either arterial tortuosity, dissection or aneurysm
formation in vivo, not proliferative or occlusive vascular pathology
(Nakamura et al., 2002
;
Yanagisawa et al., 2002
;
Arteaga-Solis et al., 2000
;
Dietz and Mecham, 2000
).
Moreover, the specificity of the effect of elastin in vivo is supported by
studies showing that collagen matrix-covered stents do not reduce neointimal
formation in a porcine model of restenosis
(Goodwin et al., 2000
). Thus,
elastin, when compared with other matrix proteins, is effective in both in
vitro and in vivo experimental models. From these studies one would expect
that targeted disruption of the elastin signaling mechanism would replicate
phenotypes observed in Eln-/- cell and mice.
Restoration of elastin for the treatment of proliferative vascular
diseases
Our work suggests that understanding the link between vascular development
and disease may provide an alternative and potentially complementary strategy
for the treatment of vascular proliferative diseases. Previously, emphasis has
been placed on improving the outcome of vascular disease by inhibiting smooth
muscle cell proliferation with coronary stents coated with either
radioactivity or chemotherapeutic drugs such as rapamycin, actinomycin D and
paclitaxil (Heldman et al.,
2001; Leon et al.,
2001
; Sousa et al.,
2001
). These treatments use a common strategy of disrupting
fundamental pathways such as microtubule assembly, DNA stability, and
regulatory cell cycle proteins that are required in virtually all actively
dividing cells. We present work that suggests restoring a natural morphogenic
signal to the vessel wall may also be therapeutically beneficial. Clearly,
additional work is needed to verify that results with murine and porcine
models are recapitulated in humans.
![]() |
ACKNOWLEDGMENTS |
---|
![]() |
Footnotes |
---|
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