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Coronary Vessel Development
The Epicardium Delivers
Harold E. Olivey, Leigh A. Compton, and Joey V. Barnett*
Coronary artery disease accounts for 54% of all cardiovascular disease
in the United States. Understanding how coronary vessels develop is
likely to uncover novel drug targets and therapeutic strategies that
will be useful in directing the repair or remodeling of coronary vessels in
adults. Recent insights have identified the importance of cells derived
from the proepicardium and epicardium in the formation of coronary
vessels. This article reviews the basic steps in coronary vessel development, the molecules implicated in these steps, and the pressing questions awaiting answers. (Trends Cardiovasc Med 2004;14:247–251)
D 2004, Elsevier Inc.
The origin of the coronary vessels has
been debated in the scientific literature
for over a century. Development of
coronary vessels coincides temporally
and spatially with formation of the
epicardium (EP), not only with respect
to individual embryos during organogenesis, but also phylogenetically, with
respect to the increasing oxygen demand of hearts during vertebrate evolution (Ostadal et al. 1975). Three
distinct mechanisms have been proposed for coronary vascular formation:
(1) migration of endocardial cells to the
subepicardial space or trapping in sinusoids formed by trabeculation of the
ventricular muscle (Grant 1926, Viragh
and Challice 1981), (2) formation by
the process of angiogenesis as an outgrowth of the proximal aorta (Bennett
1936, Goldsmith and Butler 1937), and
Harold E. Olivey, Leigh A. Compton, and Joey
V. Barnett are at the Department of Pharmacology, Vanderbilt University Medical Center,
Nashville, Tennessee, USA.
* Address correspondence to: Joey V.
Barnett, PhD, Department of Pharmacology,
Vanderbilt University Medical Center, Room
476 RRB, 2220 Pierce Avenue, Nashville, TN
37232-6600, USA. Tel.: (+1) 615-936-1722;
fax: (+1) 615-343-6532; e-mail: joey.barnett@
vanderbilt.edu.
D 2004, Elsevier Inc. All rights reserved.
1050-1738/04/$-see front matter
TCM Vol. 14, No. 6, 2004
(3) derivation from the proepicardium
(PE) and EP (Mikawa and Fischman
1992, Poelmann et al. 1993). Although
each of these mechanisms has enjoyed
varying degrees of acceptance by the
scientific community, the importance of
epicardially derived cells in coronary
vessel formation has now been demonstrated in a number of experimental
systems. Before detailing specific experiments and hypotheses, we begin with
an overview of epicardial and coronary
vessel formation.
EP and Origin of Coronary Vessels
The EP originally was thought to be
derived from the myocardium, but
increasing evidence has identified the
PE as the source for the majority of the
mature EP that covers the myocardium
(Manasek 1968, Manner 1993). In chick
embryos, the PE arises from mesothelial cells along the caudal border of the
pericardial cavity (Figure 1A, Figure 2A)
(Ho and Shimada 1978). These mesothelial cells initially form villi, but soon
develop into a small, bulbous mass adjacent to the sinus venosus. The PE
enlarges, contacts the heart at the atrioventricular (AV) groove, and migrates
to the heart, assisted by glycosylated
microfibrils (Figure 1B) (Nahirney et al.
2003) that bridge the gap between the
myocardium and the PE. Cells of the PE
maintain polarity as they migrate over
the heart as an intact epithelium with
the formerly luminal surface in contact
with the myocardium (Figures 2B–D). In
mammals, clusters of PE cells detach as
vesicles that are transferred to the heart
via the pericardial fluid (Figure 1B)
(Komiyama et al. 1987, Kuhn and Liebherr 1988). In both avians and mammals, subpopulations of cells of the PE
undergo epithelial–mesenchymal transformation (EMT) soon after contacting
the myocardium, and cells migrate into
the subepicardial space (Figure 1C). A
subset of these cells migrates further
into the compact zone of the myocardium. The fate of these transformed cells
is intimately linked to coronary vessel
development (Mikawa and Fischman
1992, Poelmann et al. 1993). If the PE
is prevented from interacting with
the heart, coronary vessel development
is absent (Gittenberger-de Groot et al.
2000, Kwee et al. 1995, Yang et al. 1995).
Coronary vessel formation begins as
angioblasts coalesce to form a primitive
vascular plexus in the subepicardial
space and myocardium (Figure 1D).
These endothelial tubes join to form
larger vessels that are recognizable as
coronary arteries and veins. Once established, the coronary vessels link to the
ascending aorta and the right atrium
and recruit PE-derived cells to form the
smooth muscle and fibroblast components of the vascular network. Therefore,
coronary vessels form by a process of
vasculogenesis after precursor cells are
delivered to the heart by the PE (MunozChapuli et al. 2002). However, whether
the PE contributes precursor cells for all
cell lineages in the coronary vessels—
endothelial, smooth muscle, and fibroblast—or is required for subsequent
delivery and support of these precursors
has been hotly debated.
Proepicardial and Epicardial
Contributions to the Vasculature:
Are Endothelial, Smooth Muscle,
and Fibroblast Precursors Derived
from the PE?
The origin of coronary vessel endothelial
cells remains controversial. Quail-tochick PE transplants have been interpreted to suggest that coronary vascular
endothelial cells do not arise in the
PE. When the quail PE is removed
and transplanted isochronically into an
247
Figure 1. Key stages in coronary vasculogenesis. (A) Formation of the proepicardium (PE). The
PE (blue) forms adjacent to the SV and opposite the AV groove of the developing heart tube
(orange) by embryonic day (ED) 9.0 in the mouse and stage 15 in the chick. (B) PE transfer and
attachment to the myocardium. Images are magnifications of the area enclosed by the dashed
box in Figure 1A. Attachment and transfer occur through different mechanisms in mammals
and avians. Starting at ED 9.25 in the mouse, clusters of cells detach from the PE and travel
across the pericardial space to the AV myocardium. Upon attachment, clusters flatten into a
monolayer and coalesce during initial epicardial formation. In avians, PE cells migrate across
an extracellular matrix bridge between the PE and myocardium. By stage 17, PE cells have
traversed the bridge, contacted the AV myocardium, and migrated radially from the point of
attachment. (C) Epicardial migration and epithelial–mesenchymal transformation (EMT). PEderived cells migrate and proliferate across the surface of the myocardium to form the
epicardium. The spatiotemporal pattern of migration is similar in mammals and avians. The
initial migration of the epicardium from the AV groove at ED 9.5 and stage 18 is depicted in a
left lateral view. Progression of epicardial migration at ED 10.5 and stage 21 is depicted in a
ventral view. Migration is complete by ED 11.0 and stage 24. Epicardial EMT begins soon after
contact with the myocardium. In cross section, epicardially derived mesenchymal cells are
depicted invading the subepicardial space and the myocardium. (D) Vessel assembly.
Angioblasts delivered by the PE coalesce to form vesicles ( purple) comprised of endothelial
cells between ED 11–12 and stage 23–26 as depicted in the ventral view. In cross section, both
subepicardial and intramyocardial vesicles are shown surrounded by epicardially derived
mesenchyme (blue). (E) Vessel maturation. Endothelial vesicles coalesce to form nascent
coronary vessels (purple) beginning at ED 11.5 and stage 27. Coronary vessels attach to the
systemic circulation by ED 13.5 and stage 32. After attachment to the aorta, smooth muscle
progenitors derived from the epicardium are recruited to the artery walls in a proximal to distal
fashion with respect to the aorta. Nascent vessels are concentrated along the AV and IV surface
of the heart as depicted in the ventral view. In cross section, endothelial tubule (purple)
formation and smooth muscle (yellow) recruitment in the subepicardial and intramyocardial
spaces are depicted. A, atrium; AV, atrioventricular; CT, conotruncus; epi, epicardium; IV,
interventricular; LA, left atrium; LV, left ventricle; myo, myocardium; RA, right atrium; RV,
right ventricle; SV, sinus venosus; V, ventricle.
248
intact chick embryo adjacent to the endogenous chick PE and sinus venosus,
PE-derived structures arise as chimeras
containing both chick and quail cells.
Poelmann et al. (1993) observed that
grafted quail PE supplied smooth muscle
cells and fibroblasts to the host heart,
but did not result in quail-derived endothelial cells in the coronary vasculature.
However, quail PE grafted with liver contributed quail-derived endothelial cells
to the host embryo (Poelmann et al.
1993). Liver alone grafted into the pericardial space also contributed endothelial cells to the coronary vessels. These
data suggested that the PE did not
contribute endothelial cells to the developing coronary vessels.
Labeling cells of the PE before migration to the heart has generally supported
the view that endothelial cells derive
from the PE itself. Mikawa and Fischman (1992) used both vital dye and
a replication-incompetent retrovirus
expressing h galactosidase (h-gal) to label
PE cells. Viral labeling allowed for infected cells and their progeny to be identified from a time shortly after infection
until hatching. Discrete h-gal-positive
colonies of either smooth muscle cells
or endothelial cells along short segments
of the coronary arteries were noted in
hatched chicks. Injections using lowtiter virus resulted in labeling of either
smooth muscle cells or endothelial cells,
but never both, demonstrating that endothelial cells and smooth muscle cells
originated from precursor cells committed before the PE contacted the heart.
Endothelial cell labeling was most common in embryos in which virus was
targeted near the dorsal mesocardium,
which is continuous with the liver. These
data could be interpreted to support the
chimera studies (Poelmann et al. 1993)
that suggested the liver as a source of
coronary vascular endothelium or, alternatively, to support the hypothesis that
endothelial cell precursors are found in
the most proximal portion of the PE,
which lacks a well-defined border separating it from the liver. The more proximal location of angioblasts in the PE
may also explain why angioblast or endothelial cell markers are absent from the
PE until well after the PE has contacted
the heart (Perez-Pomares et al. 2002a),
suggesting that angioblasts arrive relatively late after other mesenchyme have
begun seeding the myocardium.
TCM Vol. 14, No. 6, 2004
Figure 2. Coronary vasculogenesis in the chicken embryo. (A) Proepicardium (PE) location.
The PE is located adjacent and caudal to the looped heart tube. Fast green was injected into the
PE of a stage-17 chicken embryo to aid in imaging. (B) PE attachment. At attachment, the PE
cells evert, allowing contact between the formerly luminal surface of the cells of the PE and
the myocardium. These cells migrate along the myocardial surface (curved arrows) to cover
the myocardium. Bar represents 25 Am. (C) Epicardium formation. After contacting the
atrioventricular groove, cells of the PE form the primitive epicardium, migrating craniolaterally
as an intact epithelium. The dotted line demarcates the boundary of the migrating epithelium at
stage 19. Bar represents 200 Am. (D) Epicardium formation. By stage 24, the epicardium covers
the ventricles, atria, and all but a small area of the outflow tract, as denoted by the dotted line.
Bar represents 200 Am. A, atrium; O, outflow tract; M, myocardium; V, ventricle. Panel B
adapted with permission from Nahirney et al. 2003. Copyright 2003, Wiley-Liss, Inc., a
subsidiary of John Wiley & Sons, Inc. Panels C and D reprinted with permission from Manner
et al. 2001. Copyright 2001, S. Karger AG.
In a subsequent study, Mikawa and
Gourdie (1996) injected very low titers of
h-gal virus (10 or fewer infectious particles) into the PE. In this instance, virus
labeled only coronary vascular smooth
muscle cells, whereas higher-titer virus
labeled coronary vascular endothelium
and fibroblasts as well. These data
strongly support the hypothesis that
smooth muscle cell precursors exist in
the PE and argue against a common
progenitor of smooth muscle cells and
fibroblasts. In total, these studies demonstrate that smooth muscle cell and
fibroblast progenitors are found within
the PE. Further, they suggest that angioblasts reside at the proximal border of
the PE in close association with the
liver and arrive at the myocardium
after other progenitor cells have begun
to seed the myocardium.
Molecular Signals that Direct
Coronary Vessel Formation
Studies using experimental embryology,
genetic manipulation, and in vitro assays
have been useful tools in revealing the
TCM Vol. 14, No. 6, 2004
roles of specific molecules during coronary vessel development. This section
discusses a subset of molecules whose
functions have been investigated.
Although several molecules have been
found to be expressed in the PE, functional studies have failed to identify those
required for PE formation. In contrast,
several molecules have been shown to be
required for EP formation and maintenance. Vascular cell adhesion molecule 1
(VCAM-1) is expressed throughout the
myocardium (Kwee et al. 1995) and
becomes localized to the outer compact
layer that abuts the newly formed EP
by embryonic day (ED) 11.5. VCAM-1deficient mice lack an EP at ED 11.5. The
VCAM-1 counterreceptors, a4h-integrin
heterodimers, are expressed in the PE
and EP (Sengbusch et al. 2002, Yang et al.
1995). Mice deficient in a4 form an EP
by ED 10.5 that degenerates (Yang et al.
1995), suggesting that VCAM-1 and a4
integrin are required for maintenance
of the EP. A second line of a4 null
mice has fewer vesicles released from
the PE that are less likely to attach to
the myocardium and fail to form an EP
(Sengbusch et al. 2002). Together, these
data suggest multiple roles for a4 integrin during PE and epicardial development, including budding of the PE,
PE cell attachment, cell migration, and
maintenance of PE-derived cells.
Two zinc finger transcription factors
have been implicated in epicardial EMT.
Wilms’ tumor 1 (WT-1) is expressed in the
PE, EP, and mesenchyme (Moore et al.
1999, Perez-Pomares et al. 2002b). Mice
homozygous null for WT-1 form a partial EP at the AV groove and the caudal
aspect of the heart by ED 12.5, with
fewer subepicardial cells present (Moore
et al. 1999), suggesting that WT-1 is required for epicardial formation, maintenance, and EMT. Friend of GATA 2
(FOG-2) is a cofactor for GATA 4 (Lu
et al. 1999, Svensson et al. 1999) required
for epicardial EMT. FOG-2 null mice
form a complete EP, but lack coronary
vessels, because the EP fails to undergo
EMT (Tevosian et al. 2000). EMT is
rescued by overexpression of FOG-2 in
the myocardium, demonstrating the importance of a myocardially derived signal in the regulation of epicardial EMT.
Mice homozygous for a GATA-4 allele
with an inactivated FOG-binding domain phenocopy the FOG-2 null mice
(Crispino et al. 2001), suggesting that
a FOG-2/GATA-4 complex is required
for epicardial EMT.
Studies of EMT in explanted PE and
EP have identified candidate factors that
regulate EMT. Both vascular endothelial
growth factor (VEGF) and fibroblast
growth factor (FGF) stimulate EMT of
epicardial cells in vitro (Morabito et al.
2001). Whereas transforming growth factor h (TGF-h) has been noted to inhibit
EMT in epicardial explants (Morabito
et al. 2001), TGF-h stimulates EMT in
PE explants (H.E. Olivey, N.A. Mundell,
and J.V. Barnett, unpublished observations). Recent evidence identifying functionally antagonistic TGF-h signaling
pathways involving the activation of different activin receptor-like kinases in
mediating endothelial cell transformation, migration, and proliferation
(Goumans et al. 2003, Lai et al. 2000) and
cardiac myocyte gene expression (Ward
et al. 2002) may be one mechanism to
explain these apparently contradictory
effects of TGF-h. FGF, VEGF, and TGFh ligand expression patterns support
roles during epicardial transformation
(Molin et al. 2003, Morabito et al. 2001,
249
Tomanek et al. 1999). How these and
other factors interact to regulate transformation, and if they act on specific
populations of precommitted cell lineages, remain to be determined.
Final steps in coronary vessel development include vessel patterning,
attachment of the vascular network to
the systemic circulation, and recruitment of smooth muscle. Connexin
43 (Cx43) mRNA is expressed abundantly in PE cells, and Cx43 / mice
display defects in coronary vessel patterning (Li et al. 2002). Although neural
crest cell ablation results in similar
defects in coronary vessel patterning,
these are not seen in neural crest specific loss of Cx43 (Li et al. 2002, Sullivan
et al. 1998). These data suggest a primary role for Cx43 and PE cells in
coronary vessel patterning. Little is
known about the molecular regulation
of the attachment of the coronary vessels to the systemic circulation. Formation of the coronary orifice has been
demonstrated to be dependent on
the proper formation of the parasympathetic ganglia (Waldo et al. 1994).
Expression of VEGFR-2 and -3 in the
truncus arteriosus prior to coronary
artery ingrowth suggests a role for
VEGF in this process (Tomanek et al.
2002). Penetration of the coronaries into
the aorta is accompanied by apoptosis of
cells at the site of attachment (Velkey
and Bernanke 2001).
Smooth muscle recruitment and differentiation occurs in a proximal to
distal fashion after attachment of the
coronary arteries to the aorta. Molecules known to direct smooth muscle
cell differentiation outside of the heart
are likely to play similar roles during
coronary vessel development. Serum
response factor (SRF), a MADS box
transcription factor, is expressed in vivo
in the PE and subepicardial mesenchyme but is absent from the EP,
although expression has been noted in
EP-derived cells in vitro (Landerholm
et al. 1999, Nelson et al. 2004). Misexpression of dominant negative SRF in
PE explants reduced the expression of
smooth muscle markers without affecting EMT, demonstrating that SRF is
required for smooth muscle cell differentiation in vitro. Platelet-derived
growth factor (PDGF-BB)-stimulated
smooth muscle differentiation was found
to require the activity of rhoA and
250
p160rho kinase (Lu et al. 2001). Inhibition of p160rho kinase decreases SRF
transcription and, in vivo, the EP and
subepicardium form apparently normally, but mesenchyme is lacking from
the myocardium (Lu et al. 2001). These
data suggest that p160rho kinase
is required for the migration or survival
of mesenchyme in the myocardium.
Mice homozygous null for PDGF-B, or
the cognate receptor PDGFR-h, have
generalized vascular smooth muscle
defects, including lack of smooth muscle
in intramyocardial vessels, whereas subepicardial vessels are only partially ensheathed by smooth muscle (Hellstrom
et al. 1999). Smad6 null mice also display
distended subepicardial vessels deficient
in smooth muscle (Galvin et al. 2000),
suggesting that bone morphogenetic
protein and TGF-h signaling, as well as
PDGF, play a role in smooth muscle differentiation or recruitment.
Pressing Questions
The determination of the lineage of PEderived endothelial cells, smooth muscle
cells, and fibroblasts is of major importance. How are these cells specified and
fated to the PE? Of particular interest is
the origin of endothelial cells. Are coronary endothelial cells specified prior to
PE cells entering into the heart? Why are
these cells delivered late relative to mesenchyme seeding the heart? When does
commitment of smooth muscle progenitor cells in the PE occur? Given that
most studies have focused on arterial
development, how closely does development of the venous system follow that
of the arterial system? Finally, is the developmental program that generates precursor cells and vessels retained in the
EP or mesenchyme of the adult? Can this
program be reactivated in adults? The
answers to these questions promise general insight into organogenesis and may
suggest novel therapeutic approaches to
coronary vessel repair in humans.
Acknowledgments
The authors have attempted to distill
the major ideas and discoveries that
have shaped this area. Inevitably, owing
to limited space, some insights have not
been discussed and some references
have been omitted. They apologize for
this and encourage the reader to explore
the area fully. J.V.B. wishes to acknowledge the support of HL67105, HL52922,
the March of Dimes, and the American
Heart Association. The authors thank
Drs. Jorg Manner and Patrick Nahirney
for supplying figures and Drs. David
Bader and Christopher Brown for critically reading the manuscript.
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