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Molecular basis of endothelial cell morphogenesis in three-dimensional extracellular matrices.

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THE ANATOMICAL RECORD 268:252–275 (2002)
Molecular Basis of Endothelial Cell
Morphogenesis in Three-Dimensional
Extracellular Matrices
Department of Pathology, Texas A&M University System Health Science Center,
College Station, Texas
Although many studies have focused on blood vessel development and new blood vessel
formation associated with disease processes, the question of how endothelial cells (ECs)
assemble into tubes in three dimensions (i.e., EC morphogenesis) remains unanswered. EC
morphogenesis is particularly dependent on a signaling axis involving the extracellular
matrix (ECM), integrins, and the cytoskeleton, which regulates EC shape changes and
signals the pathways necessary for tube formation. Recent studies reveal that genes regulating this matrix-integrin-cytoskeletal (MIC) signaling axis are differentially expressed
during EC morphogenesis. The Rho GTPases represent an important class of molecules
involved in these events. Cdc42 and Rac1 are required for the process of EC intracellular
vacuole formation and coalescence that regulates EC lumen formation in three-dimensional
(3D) extracellular matrices, while RhoA appears to stabilize capillary tube networks. Once
EC tube networks are established, supporting cells, such as pericytes, are recruited to further
stabilize these networks, perhaps by regulating EC basement membrane matrix assembly.
Furthermore, we consider recent work showing that EC morphogenesis is balanced by a
tendency for newly formed tubes to regress. This morphogenesis-regression balance is controlled by differential gene expression of such molecules as VEGF, angiopoietin-2, and PAI-1,
as well as a plasmin- and matrix metalloproteinase-dependent mechanism that induces tube
regression through degradation of ECM scaffolds that support EC-lined tubes. It is our hope
that this review will stimulate increased interest and effort focused on the basic mechanisms
regulating capillary tube formation and regression in 3D extracellular matrices. Anat Rec
268:252–275, 2002. © 2002 Wiley-Liss, Inc.
Key words: endothelial cell morphogenesis; extracellular matrix; integrins;
Rho GTPases; vacuoles and lumens; differential gene expression;
matrix metalloproteinases; plasmin; endothelial cell tube
The molecular control of blood vessel formation has
been a major topic of investigation over the past several
decades. Two major processes—vasculogenesis and angiogenesis—are responsible for blood vessel formation in vivo
(Hanahan, 1997; Carmeliet and Jain, 2000; Patan, 2000;
Conway et al., 2001). Models of these events using in vitro
systems are illustrated in Figure 1. Vasculogenesis refers
to the de novo development of capillaries from individual
endothelial cell (EC) precursors within tissues, or delivered from the circulation (Risau and Flamme, 1995; Drake
et al., 1997; Drake and Little, 1999; Drake and Fleming,
2000; Carmeliet and Luttun, 2001). Angiogenesis refers to
new blood vessel formation from preexisting vessels (Folkman, 1995; Hanahan, 1997; Carmeliet and Jain, 2000;
Conway et al., 2001). Major cytokines that regulate these
events are VEGFs, FGFs, angiopoietins, placental growth
factor, various chemokines (e.g., SDF-1␣) and other
growth factors such as TGF-␤ and insulin-like growth
Grant sponsor: NIH (NHLBI); Grant number: HL59373; Grant
sponsor: Texas Higher Education Coordinating Board; Grant
number: 89-57-2001; Grant sponsor: NRSA; Grant number: F32
*Correspondence to: George E. Davis, M.D., Ph.D., Department
of Pathology, 208 Reynolds Medical Building, College Station, TX
77843-1114. Fax: (979) 862-1299. E-mail:
Received 5 February 2002; Accepted 4 June 2002
DOI 10.1002/ar.10159
Published online 10 October 2002 in Wiley InterScience
Fig. 1. In vitro EC morphogenesis assay systems in collagen matrices that mimic blood vessel formation through vasculogenesis or angiogenic mechanisms. A: ECs were seeded within collagen matrices as
single cells and were allowed to develop interconnecting networks of
tubes for 48 hr (left panels) (Davis and Camarillo, 1996). In other assays,
ECs were seeded on the surface of collagen matrices and were allowed
to invade and undergo morphogenesis for 72 hr (right panels) (Davis et
al., 2000b). Cultures were photographed from a side or bottom view at
a focal plane beneath the EC monolayer. Bar ⫽ 100 ␮m. B: ECs were
seeded within collagen matrices as single cells and allowed to develop
vacuoles (8 hr) and tubes (24 – 48 hr) over time. Bar ⫽ 50 ␮m. In all cases,
the cultures were fixed with glutaraldehyde and stained with toluidine
factors (Pepper, 1997; Yancopoulos et al., 2000; Carmeliet
et al., 2001; Hellstrom et al., 2001b). Although many of the
factors and their receptors that control these processes are
well understood (Yancopoulos et al., 2000), little information exists concerning the molecular mechanisms by
which ECs physically assemble into capillary tube structures in three-dimensional (3D) extracellular matrix
(ECM) environments. EC morphogenesis is defined here
as the process whereby ECs assemble into tubes in 3D
extracellular matrices. These events require EC interactions with ECM through integrins, and signaling events
involving cytoskeletal elements that control EC shape and
cell– cell interactions that dictate the 3D structure of
tubes. How these interactions lead to the ability of ECs to
assemble into tubes with a fluid-filled lumen, an abluminal surface in contact with basement membrane matrix,
and cell– cell junctional contacts remains unclear (Figs. 1
and 2A). However, recent advances in the ability to con-
struct EC-lined tubes using defined in vitro systems, and
the ability to manipulate the expression of individual
genes within primary ECs now allow for a comprehensive
examination of how EC morphogenesis is controlled at a
molecular level (Davis and Camarillo, 1996; Bell et al.,
2001; Bayless and Davis, 2002 ; Davis and Bayless, 2003).
These latter issues concerning the molecular control
of EC tube assembly in 3D extracellular matrices are
the subject of this review. We investigate how a matrixintegrin-cytoskeletal (MIC) signaling axis regulates EC
morphogenesis, and how critical cytoskeletal signaling
molecules (the Rho GTPases) regulate these events. In
addition, mechanisms underlying EC lumen formation
and sprouting, which represent the major morphologic
changes regulating EC morphogenesis, are discussed.
Additional topics include how differential gene expression regulates EC morphogenesis, and how a balance of
genes, molecules, and cells (such as pericytes) regu-
Fig. 2. Electron micrographs of developing capillary tubes in 3D
collagen matrices. A: EC-collagen gel cultures were established, and at
various times cultures were fixed with glutaraldehyde and embedded in
plastic. Thin sections were prepared and examined by electron microscopy. After 72 hr of culture, EC cell– cell junctions were identified; arrows
point to two junctional contacts. Higher-power images of these two
junctions (JXN-1 and 2) are shown above and below the figure. B:
Cultures were fixed at 24 hr. A C-shaped EC is observed surrounding a
lumenal space. Arrowheads: EC processes available for junction formation, as illustrated in part A. Arrows: interface between the collagen
matrix and the lumenal compartment. Bar ⫽ 5 ␮m.
late capillary tube formation and maintenance by controlling the processes of EC morphogenesis vs. regression.
cular morphogenesis, including ␣2␤1, ␣5␤1, ␣1␤1, ␣6␤1,
and ␣v␤3 (Bauer et al., 1992; Brooks et al., 1994; 1995,
1996; Bloch et al., 1997; Senger et al., 1997, 2002;
Bayless et al., 2000; Rupp and Little, 2001). The involvement of particular integrins is dependent on the
matrix environment to which ECs are exposed, and
suggests that signaling pathways common to multiple
integrins are capable of directing the MIC axis required
for capillary tube assembly in three dimensions. The
␣2␤1 and ␣1␤1 integrins regulate EC morphogenesis in
collagen-rich ECM (Davis and Camarillo, 1996; Senger
et al., 1997, 2002), ␣5␤1 and ␣v␤3 in fibronectin-fibrinrich ECM (e.g., in wound tissue matrix) (Bayless et al.,
2000; Kim et al., 2000), and ␣6␤1 in laminin-rich ECM
environments (Bauer et al., 1992; Davis and Camarillo,
1995). The ability of multiple integrins to regulate this
process creates an important integrin signaling redundancy that may be necessary for blood vessel assembly
in different ECM environments. ECM environments
vary considerably between tissues (e.g., skin vs. brain)
and during development or various types of tissue injury wherein new blood vessel formation occurs (Senger,
1996; Sage, 1997; Davis et al., 2000a).
MIC Signaling Axis Controls EC Morphogenesis
in Three Dimensions
It is apparent from recent work that the MIC signaling axis is a major pathway regulating EC morphogenesis in 3D extracellular matrices (Fig. 3) (Davis and
Camarillo, 1996; Salazar et al., 1999; Bayless et al.,
2000; Bell et al., 2001; Rupp and Little, 2001; Bayless
and Davis, 2002; Davis and Bayless, 2003). As shown in
Figure 3, this pathway depends not only on EC exposure
to exogenous ECM environments conducive for vascular
formation such as collagen type I or fibrin matrices, but
also on endogenous synthesis of ECM by ECs (Nicosia
and Madri, 1987; Ingber and Folkman, 1988; Maragoudakis et al. 1988; Iruela-Arispe et al., 1991; Sephel
et al., 1996; Bonanno et al., 2000; Bell et al., 2001). One
of the important conclusions drawn from these studies
is that multiple integrins, as well as a number of ECM
environments, are permissive for EC morphogenesis. It
is clear that both ␤1 and ␣v integrins can support vas-
Fig. 3. The MIC signaling axis regulates capillary tube morphogenesis and regression in 3D matrix
environments. Molecular regulators of these events are indicated next to where they are thought to act during
these events.
Signaling events downstream of ECM–integrin interactions clearly are of critical importance to EC morphogenesis. This signaling affects EC survival, proliferation, migration, shape, and differentiation (Shattil and Ginsberg,
1997; Sastry and Burridge, 2000; Schwartz and Shattil,
2000; Mettouchi et al., 2001). All of these processes are
required during EC morphogenesis. Many of the molecules regulating integrin signaling are associated with the
actin or microtubule cytoskeletal scaffold (Kiosses et al.,
1999; Lee et al., 1999; Schwartz and Shattil, 2000; Paik et
al., 2001). In addition, these cytoskeletal molecules control
cell shape and are centrally relevant to the question of
how capillary tubes form in three dimensions. Our laboratory has identified three actin regulatory proteins that
are coordinately upregulated during EC morphogenesis:
gelsolin, vasoactive-stimulated phosphoprotein (VASP),
and profilin (Salazar et al., 1999). Gelsolin is well known
to sever actin filaments, thereby participating in actin
assembly/disassembly reactions; profilin binds to VASP (a
major phosphoprotein target of protein kinases A and G)
(Reinhard et al., 2001), and also sequesters actin monomers. In addition, profilin is involved in Rho GTPasestimulated actin polymerization (Bishop and Hall, 2000).
An interesting question concerns the extent to which actin
microfilaments or microtubules are responsible for the 3D
shape of capillary tubes in ECM. It is notable that both
actin and microtubule structures are directly affected by
integrin–ECM interactions (Sastry and Burridge, 2000;
Goode et al., 2000). A direct influence of integrins on
microtubules has only recently been appreciated (Volkov
et al., 2001; Zhou et al., 2001). To assess the relative
contribution of actin microfilaments vs. microtubules in
the 3D structure of capillary tubes (Bayless et al., unpublished results), established capillary tube networks in collagen matrices were treated with either cytochalasin B (to
disrupt actin) or nocodazole and/or colchicine (to disrupt
microtubules) (Fig. 4). The microtubule disrupting drugs
induced rapid collapse of tubes, while actin depolymerization caused minimal changes in the overall tube structure
(Fig. 4) (Bayless et al., unpublished results). These data
suggest that microtubules are critical to the overall shape
of capillary tubes in 3D ECM, and that actin is not as
important in regulating this overall structure.
Major signaling molecules downstream of integrin–
ECM interactions are the Rho GTPases, a family of small
GTPases in the Ras superfamily that regulate cytoskeletal
structure and function (Kaibuchi et al., 1999; Bishop and
Hall, 2000; Hall and Nobes, 2000; Schwartz and Shattil,
2000; Takai et al., 2001; Ridley, 2001a). These molecules
regulate one of a series of signaling pathways resulting
from integrin–ECM interactions (Giancotti and Ruoslahti,
1999; Sastry and Burridge, 2000; Geiger et al., 2001;
Schwartz and Assoian, 2001). Rho GTPases initially were
shown to control specific events in actin cytoskeletal dynamics, as reported in the seminal studies of Ridley and
Hall (Ridley et al., 1992; Ridley and Hall, 1992). In addition, these GTPases regulate not only the function of the
actin cytoskeleton, but also the microtubule and intermediate filament cytoskeletons (Nobes and Hall, 1995; Inada
et al., 1999; Meriane et al., 2000; Daub et al., 2001;
Palazzo et al., 2001; Ridley, 2001a). They affect many
critical steps in cell behavior that are characteristic of EC
morphogenesis (i.e., cell migration, cell proliferation and
regulation of gene expression, cell shape, permeability,
and polarity) (Kaibuchi et al., 1999; Hall and Nobes, 2000;
Ridley, 2001a; Settleman, 2001; Takai et al., 2001; Wojciak-Stothard et al., 2001). However, until recently, no
direct link between their function and EC morphogenesis
in 3D ECM has been made (Bayless and Davis, 2002). The
RhoA GTPase has been reported to induce actin stress
Fig. 4. Disruption of microtubules induces rapid collapse of capillary
tubes in 3D collagen matrices. EC-collagen gel cultures were established, and after 48 hr the cultures were left untreated or were treated
with cytochalasin B (10 ␮M), colchicine (10 ␮M), and nocodazole (10 ␮M)
for 30 min. Cultures were then fixed with glutaraldehyde and photographed. Bar ⫽ 50 ␮m.
fiber formation and focal adhesions, and to stimulate actomyosin contractility and microtubule elongation (Palazzo
et al., 2001, Ridley, 2001a). The Rac1 GTPase induces
lamellipodia formation, while Cdc42 induces filopodia.
Both Rac1 and Cdc42 have been reported to increase microtubule stability by inducing the phosphorylation of
stathmin through PAK-1 (Daub et al., 2001). Furthermore, Rho GTPases have been reported to regulate a variety of other cellular processes, such as vesicular trafficking events (including endocytosis, macropinocytosis, and
phagocytosis) (Greenberg, 1995; Swanson and Watts,
1995; Garrett et al., 2000; Cardelli, 2001; Ridley, 2001b).
In this latter case, RhoA was observed to regulate integrin-mediated phagosome formation while Rac1 and
Cdc42 regulate Fc-receptor-mediated phagosome formation (Caron and Hall, 1998). The processes of macropinocytosis and phagocytosis appear to be related to EC intracellular vacuole formation and coalescence, which is an
important step in the lumen formation pathway (Davis
and Camarillo, 1996; Bayless and Davis, 2002; Davis and
Bayless, 2003) (see below). Furthermore, we have recently
shown that Cdc42 and Rac1, which regulate both macropinocytosis and phagocytosis, also control the formation of
capillary lumens in collagen or fibrin matrices (Bayless
and Davis, 2002).
In Vitro Models of Capillary Morphogenesis in
3D Extracellular Matrices
In order to determine how ECs assemble into capillary
tubes in 3D space, models of this process are necessary.
Thus far, it has been difficult to elucidate how capillary
tubes form in vivo, so there is an increasing requirement
for utilization of in vitro models that permit a molecular
analysis of these events. In vitro systems have the following advantages: 1) defined experimental conditions can be
achieved to test hypotheses and define genetic, protein,
and morphologic changes in a coordinated fashion; 2) the
EC population is relatively uniform; and 3) the function of
individual genes during particular events in EC morphogenesis can be directly addressed. In vitro models represent a rapid, defined, and efficient experimental strategy
to elucidate the molecular events required for tube formation in 3D extracellular matrices. Although similar experiments can be performed in vivo, it is difficult to define
morphogenic steps or assess the role of individual genes/
molecules in a complex tissue wherein ECs represent only
a small fraction of the total cells. It is increasingly evident
that a balanced experimental approach using both in vitro
and in vivo morphogenic systems is necessary to elucidate
the molecular basis of capillary tube assembly. Some important questions are raised: What is the minimum molecular machinery necessary for EC tube formation in
three dimensions? Are EC-specific molecules necessary for
these events, and what are these molecules? How are EC
lumen formation and EC sprouting/branching events controlled at a molecular level?
In the early to mid-1980s, Montesano and colleagues
established capillary morphogenesis assays in vitro that
strongly recapitulated the appearance of capillary tube
structures in vivo using ECs alone (Montesano and Orci,
1985; Montesano, 1992; Montesano et al., 1992). In these
assays, primary ECs were seeded as monolayers onto the
surface of collagen or fibrin gels, and over a period of days
the ECs were observed to invade these matrices to form
capillary tube structures. Histologic and electron microscopic analyses revealed capillary tubes with a central
fluid-filled lumenal compartment, with a defined abluminal surface surrounded by ECM. Later, Pepper and Montesano (1990) showed that known angiogenic cytokines
such as VEGF and FGF-2 (which together showed strong
synergism) stimulated the invasion and formation of capillary tubes from EC monolayers, further demonstrating
the relevance of these assays (Pepper et al., 1992). Other
models, such as that described by Nicosia and colleagues
(Nicosia et al., 1982; Nicosia and Madri, 1987; Nicosia and
Ottinetti, 1990; Nicosia and Villaschi, 1999), showed similar capillary tube sprouting events in fibrin or collagen
matrices, using the aortic ring model, in response to angiogenic cytokines. Other morphogenic assays developed
by Vernon et al. (1992) and Vernon and Sage (1995), and
separately by Davis and Camarillo (1995), studied planar
EC morphogenesis, whereby ECs were allowed to rearrange to form interconnecting networks of EC cords on the
surface of a reconstituted basement membrane matrix.
These assays demonstrated how mechanical forces exerted by ECs on ECM (i.e., through the MIC signaling
axis) can rapidly induce the assembly of EC network formation by distortion of ECM into “matrical pathways”
(Vernon and Sage, 1995) or “matrix guidance pathways”
(Davis and Camarillo, 1995) that allow directed EC migration toward neighboring cells. These matrix guidance
pathways may play a key role in how ECs locate each
other in 3D space and rapidly form multicellular networks
in tissues.
Fig. 5. Cellular events during distinct stages of EC morphogenesis in
vivo and their relationships to in vitro morphogenesis systems. A: ECs
convert from a quiescent to an activated state during angiogenesis;
then, following morphogenesis and differentiation, they convert back to
a quiescent state. B: Time-lapse photographs of EC morphogenesis
showing EC lumen development. Two ECs were photographed at the
indicated times using phase-contrast microscopy. Arrowheads: EC intracellular vacuoles. Bar ⫽ 30 ␮m.
More recent assays, such as those developed by our
laboratory, have utilized ECs suspended as single cells in
3D extracellular matrices (Davis and Camarillo, 1996;
Salazar et al., 1999; Bayless et al., 2000; Bell et al., 2001;
Bayless and Davis, 2002) (Figs. 1 and 5). In Figure 5B, a
series of time-lapse images reveal two ECs forming intracellular vacuoles, and then the cells interconnecting to
form a lumenal structure. Related assays were also developed by Marx et al. (1994) and Ilan et al. (1998). These
assays may more closely reflect the process of vasculogenesis as opposed to angiogenesis, since single cells in 3D
space assemble into tubes. Examples of the assay systems
that mimic vasculogenesis (Davis and Camarillo, 1996) vs.
angiogenesis (Davis et al., 2000b) are shown in Figure 1.
An experimental advantage of these “vasculogenic” assays
is that essentially all of the ECs participate in capillary
tube formation, while only a fraction of the ECs in the
assays described above (i.e., more similar to angiogenesis,
in which subsets of cells invade from a monolayer to undergo morphogenesis) proceed through the morphogenic
process. The “vasculogenic” assays have allowed for a comprehensive analysis of differential gene expression during
morphogenesis in three dimensions, and the isolation of
differentially expressed novel genes (i.e., capillary morphogenesis genes (CMGs)) (Bell et al., 2001) (see below)
since the entire population of ECs undergoes morphogenesis during a similar time course. In addition, this analysis of differential gene expression can be correlated with
distinct steps in the EC morphogenic process (Fig. 6). Also,
EC gene expression can be manipulated using recombinant adenoviruses to address the role of particular genes
during these events (Bell et al., 2001; Bayless and Davis,
An interesting question is, why does only a subset of
ECs participate in sprout formation from an EC monolayer (i.e., “angiogenesis” assay) (Fig. 1A, right panel)?
This result is in marked contrast to that observed in the
“vasculogenesis” type of assay, wherein essentially all of
the ECs participate in the morphogenic response (Fig. 1A,
left panel). The phenomenon whereby only a small subset
of ECs sprout and form tubes from a monolayer appears
similar to a phenomenon observed in neural or hair development, called lateral inhibition (Lindsell et al., 1996;
Lanford et al., 1999). Lateral inhibition refers to a situation in which subsets of cells produce factors that inhibit
their neighboring cells (subsets of cells that express receptors for the inhibitory factor), which results in selective
differentiation of cells in defined spatial areas. For example, these types of interactions regulate the density and
spatial arrangement of hair or ciliated cells in the skin
and cochlea (Deblandre et al., 1999; Lanford et al., 1999;
Renaud and Simpson, 2001). It is interesting to consider
that hair or ciliated cell density is analogous to vascular
sprout density during angiogenesis. Varying the levels of
these inhibitory factors directly influences the density of
hair follicles in vivo (Lanford et al., 1999). Thus, these
types of molecules could play a role in the control of EC
sprout density (as illustrated in Fig. 1A, right panels)
through such a lateral inhibition mechanism. Molecules
that regulate lateral inhibition include Notch receptors
and Notch ligands, such as Jagged and Delta (Lindsell et
al., 1996). Zimrin et al. (1996) initially showed that
Fig. 6. Steps in EC morphogenesis in 3D extracellular matrices. ECs were suspended in collagen
matrices for the times indicated. The cultures were fixed, stained with toluidine blue, and photographed. The
role of Rho GTPases and pericytes/VSMCs in regulating different stages of morphogenesis are indicated.
Bar ⫽ 50 ␮m.
Fig. 7. EC intracellular vacuole membranes label with GFP-Rac1
during EC morphogenesis in 3D collagen matrices. ECs were infected
with a recombinant adenovirus carrying a GFP-Rac1V12 fusion gene.
After 48 hr they were cultured in 3D collagen matrices. After 24 hr,
cultures were fixed with paraformaldehyde and examined by confocal
microscopy. A single optical section of a developing EC lumenal (L)
structure is shown. Arrows: GFP-Rac1-labeled vacuole membranes.
Arrowhead: the lumenal membrane. Bar ⫽ 10 ␮m.
Jagged-1 was upregulated in ECs during morphogenesis,
and negatively regulated the degree of EC sprouting and
invasion. More recently, Jagged-1 was shown to be highly
upregulated during EC morphogenesis in our “vasculogenic” assay (it was the most upregulated gene at 8 hr in
more than 7,000 genes screened in a DNA microarray
analysis) (Bell et al., 2001). Other studies have shown that
the Notch receptors (Notch-4 and Notch-1) are present in
ECs during morphogenic events (Zimrin et al., 1996; Uyttendaele et al., 2000; Linder et al., 2001). This pathway
regulates the related process of Drosophila tracheal tube
development (which is related to EC morphogenesis)
(Metzger and Krasnow, 1999) through a lateral inhibition
mechanism (Ikeya and Hayashi, 1999; Llimargas, 1999).
Further work is necessary to investigate how these molecules regulate EC morphogenesis, and to determine
whether they play a role in distinguishing the molecular
events that characterize angiogenesis vs. vasculogenesis.
In vitro EC morphogenesis models are also useful for
studying how ECs physically assemble into tubes during
this process. Currently, little information exists concerning these EC assembly events. Several assay systems appear to be particularly relevant. The first, as described by
Vernon and Sage (1999), places beads containing ECs into
collagen gels and allows EC sprouting and morphogenesis
to proceed radially. Images obtained from this assay
should allow for confocal microscopic imaging of sprouts
and tubes over time, without optical interference from
cells at different focal planes above or below the sprout.
Tube formation could be analyzed with time to determine
how ECs interconnect with each other and change shape
to form tubes. We have recently described a horizontal EC
invasion assay that allows for observation of EC sprouting
and tube formation without optical interference from the
EC monolayer (Davis et al., 2000b). Another assay that
appears promising for this purpose is that described by
Bayless and Davis (2002). In this assay, fluorescentlylabeled individual ECs are suspended in collagen gels, and
1-␮l dots of cell-gel mix are placed on coverslips and inverted onto chambers containing culture media. Under
these conditions, ECs form interconnecting networks of
tubes, as in other 3D assays. These cultures can be visualized as living cultures or following fixation using multiphoton confocal microscopy (Fig. 7) (Bayless and Davis,
2002). To facilitate these experiments, we recently developed GFP-Rho GTPase fusion targeting reagents that label various intracellular components within ECs, including intracellular vacuole membranes (i.e., which regulate
lumen formation) (Bayless and Davis, 2002). The development of these new technologies makes it possible to determine how groups of ECs physically assemble into capillary
tube networks over time.
Critical Steps in the Formation of
Interconnecting Networks of EC-Lined Tubes
During Capillary Morphogenesis
In vitro models can be utilized to directly test the role of
particular molecules in defined steps of capillary tube
network formation (Fig. 6). Data from many laboratories
indicate a critical role for integrin–ECM interactions, the
cytoskeleton, and membrane-type matrix metalloproteinases (MT-MMPs) in these events. Two major morphologic
changes that regulate EC tube development include lumen formation and branching/sprouting, which controls
how ECs interconnect into networks in three dimensions
(Fig. 6). Our data, as well as those of others, suggest that
these distinct morphologic steps may be separable. For
example, during EC morphogenesis on the surface of basement membrane matrix gels, dramatic branching and network formation occurs with little evidence of lumen formation (Vernon et al., 1992; Davis and Camarillo, 1995).
Also, increased expression of the Rac1 GTPase in either its
wild type or constitutively active form in ECs using recombinant adenoviruses increases vacuole and lumen formation, but there is little branching and sprouting (Bayless and Davis, 2002). In contrast, increased expression of
Cdc42 stimulates vacuole and lumen formation (Bayless
and Davis, in press), as well as branching and sprouting,
while increased expression of RhoA results in marked
increases in EC sprouting with little lumen formation
(Bayless and Davis, unpublished results). Using a different experimental approach, another recent study (Kiosses
et al., 2002) proposed a role for PAK1 (a Rac1 and Cdc42
downstream effector) in branching phenomena during EC
morphogenesis. Much work is needed to elucidate the
roles of individual Rho GTPases and their downstream
effectors at different stages of morphogenesis (Fig. 6).
It would be interesting to determine the temporal relationship of EC sprouting to lumen formation. During angiogenesis, it is believed that EC sprouts occur first, providing provisional EC networks which then progress to
form lumens. In our in vitro model, which more closely
mimics a vasculogenic response, the opposite sequence of
events occurs. Individual ECs develop vacuoles and lu-
mens first, followed by sprouting events that lead to interconnected tubes (Figs. 5B, 6 – 8). However, it should be
pointed out that vacuoles remain visible in sprouting ECs,
and we believe they contribute to the process of lumens
extending in the direction of EC sprouts during branching
morphogenesis (Davis and Camarillo, 1996; Davis et al.,
2000b). Recent experiments using our horizontal invasion
assay system reveal that intracellular vacuoles are
present and participate in lumen formation during initial
sprouting events (Bayless and Davis, unpublished results). These data suggest that the initiation of lumen
formation may occur concurrently with sprouting.
An important question that has been addressed over the
years concerns the role of ECM-degrading proteinases in
EC morphogenesis (Pepper and Montesano, 1990; Haas
and Madri, 1999; Werb et al., 1999; Pepper, 2001; Sternlicht and Werb, 2001). Recent work (Hiraoka et al., 1998;
Hotary et al., 2000) has provided very compelling evidence
of a requirement for MT-MMPs in capillary morphogenesis. In addition, MMP-2 and MMP-9 knockout experiments have revealed roles for these proteinases during
angiogenesis in vivo (Vu et al., 1998; Bergers et al., 2000).
Hiraoka et al. (1998) and Hotary et al. (2000) have reported a critical role for MT-MMP-1 and other MT-MMPs
as pericellular proteinases which control EC and epithelial cell invasion and morphogenesis in 3D ECM gels. In
contrast, many of the secreted MMPs, and even mutated
MT-MMPs without a membrane anchor, were unable to
support invasion and morphogenesis (Hiraoka et al., 1998;
Hotary et al., 2000). Experiments from our laboratory also
support these conclusions. When ECs are suspended as
single cells in collagen gels, MMP inhibitors that block
MT-MMPs (such as TIMP-2 and the chemical inhibitor
GM6001) markedly block EC morphogenesis while
TIMP-1, PAI-1, aprotinin, and other inhibitors have no
effect (Davis et al., unpublished data). In our “angiogenesis”-like assay model (Davis et al., 2000b) (see Fig. 1A,
right panels), GM6001 completely inhibited invasion and
morphogenesis, while TIMP-2 blocked morphogenesis
(i.e., lumen and tube formation) but did not fully block
invasion of individual ECs (Bayless et al., unpublished
results). In contrast, TIMP-1 did not block this process.
The effect of TIMP-2 may relate to the discussion above
regarding how EC sprouting/invasion may be regulated
separately from lumen formation, and suggests a complex
role for MMPs in EC morphogenesis. Another intriguing
aspect of the role of MMPs in morphogenesis is that they
can expose matricryptic sites in ECM proteins that can
regulate cell responses (Sage, 1997; Davis et al., 2000a).
Matricryptic sites are biologically active cryptic domains
in ECM molecules that are not exposed in mature ECM
molecules but are exposed following conformational or
enzymatic modifications (Davis et al., 2000a). Recently,
Xu et al. (2001) reported that matricryptic sites in collagen
type IV regulate injury- and tumor-induced angiogenic
responses. Antibodies that recognized these matricryptic
epitopes in collagen type IV were able to block in vivo
angiogenic responses.
Another question concerns the possible role of secreted
proteinases in EC morphogenesis. In addition to their role
in degrading ECM, many of these proteinases have been
found to have alternate substrates, which could create
new roles for these molecules in this process (Sternlicht
and Werb, 2001). For example, MMP-9 was recently observed to liberate ECM-associated VEGF, which induces
Fig. 8. Time course of EC lumen formation through intracellular vacuole formation and coalescence during morphogenesis in 3D collagen
matrices. ECs were placed into collagen matrices and at the indicated
times were fixed with glutaraldehyde, embedded in plastic, thin-sec-
tioned, and stained with toluidine blue. Representative fields were photographed. Arrowheads: open lumenal structures (C-shaped ECs) at 24
hr. Arrows: lumenal structures with a continuous EC lining. Bar ⫽ 25 ␮m.
angiogenesis (Bergers et al., 2000). Another role for secreted EC proteinases is to regulate the process of capillary tube regression (Davis et al., 2001), a process that
normally follows the wound-healing angiogenic response
(Clark, 1996). As discussed below, under appropriate conditions, ECs are capable of degrading the ECM scaffold in
which they are suspended, causing regression of capillary
tubes (Davis et al., 2001).
Figures 5B and 6 – 8, EC vacuole formation and coalescence occurs over time during capillary morphogenesis to
regulate the lumen formation process. In Figure 8, crosssections of fixed cultures at various times illustrate the
formation of intracellular vacuoles and lumens in collagen
matrices. An electron micrograph of an EC with intracellular vacuoles during morphogenesis is shown in Figure 9.
As shown in the figure, collagenous matrix is observed on
the preliminary abluminal surface, while the interior of
the vacuoles are devoid of ECM. These findings are supported by previous work that suggested a role for an
intracellular lumen formation mechanism regulating EC
or epithelial cell lumen formation. Wolff and Bar (1972)
and Guldner and Wolff (1973) described “seamless” ECs in
vivo that contain a lumen and form junctional contacts
with adjacent ECs, but contain no junctional site in crosssection. This type of lumen has to form through an intracellular mechanism, such as that observed during intracellular vacuole formation and coalescence (Davis and
Camarillo, 1996). Similar “seamless” epithelial cells are
observed in the distal tips of the Drosophila tracheal tube
system (Samakovlis et al., 1996). Another example of intracellular lumen formation is that observed in the excretory epithelial cell in C. elegans, which is a single cell with
a lumen (Buechner et al., 1999).
The in vitro systems developed in our laboratory enable
a detailed molecular analysis of the vacuole and lumen
Regulation of Capillary Lumen Formation in
Three Dimensions by Intracellular Vacuole
Formation and Coalescence: An Integrin and
Rho GTPase-Dependent Pinocytic Event
A major direction of our current work is to elucidate how
ECs form lumens in 3D ECM environments. We have
shown in a series of studies that a major mechanism of
lumen formation of ECs in either 3D collagen or fibrin
matrices involves intracellular vacuole formation and coalescence (Davis and Camarillo, 1996; Bayless et al., 2000;
Davis et al., 2000; Bayless and Davis, 2002; Davis and
Bayless, 2003). EC intracellular vacuoles have also been
observed both in vivo and in vitro by many investigators
(Speidel, 1933; Clark and Clark, 1939; Wolff and Bar,
1972; Guldner and Wolff, 1973; Dyson et al., 1976; Wagner, 1980; Folkman and Haudenschild, 1980; Montesano
and Orci, 1988; Konerding et al., 1992; Meyer et al., 1997;
Yang et al., 1999; Dvorak and Feng, 2001). As shown in
Fig. 9. Electron micrograph of intracellular vacuoles regulating lumen
formation during EC morphogenesis in 3D collagen matrices. ECs were
placed into collagen matrices, and after 24 hr cultures were fixed and
processed for electron microscopy (Davis and Camarillo, 1996). Aster-
isks: the fluid-filled intracellular vacuole space. Arrowheads: small EC
processes observed on the abluminal surface and in the interior of EC
vacuoles, indicating pinocytosis of plasma membrane. Arrows: collagen
fibrils. Bar ⫽ 5 ␮m.
formation process. Thus far, our findings indicate that
vacuole formation depends on integrin interactions with
ECM, and occurs through a pinocytic mechanism that
requires both the actin and microtubule cytoskeletons
(Fig. 10). The addition of cytochalasin B or nocodazole,
which disrupt the actin and microtubule cytoskeletons,
respectively, completely blocks vacuole formation and subsequent lumen formation. We have previously shown that
integrin–ECM interactions are required for intracellular
vacuole and lumen formation. The ␣2␤1 integrin regulates
vacuole formation in collagen matrices, and a combination
of ␣v␤3 and ␣5␤1 regulates these events in fibrin matrices
(Davis and Camarillo, 1996; Bayless et al., 2000). We
originally showed that vacuole formation occurs through a
pinocytic mechanism. Plasma membrane markers were
detectable in vacuole membranes, and when membraneimpermeant fluorescent dyes (i.e., dextran-fluorescein)
were added to the culture media, they strongly labeled the
Fig. 10. Pinocytosis of fluorescent dyes into EC intracellular vacuoles
with vacuole membrane-associated GFP-Rac1 GTPase. ECs were infected with a recombinant adenovirus carrying a GFP-Rac1V12 fusion
gene. After 48 hr they were cultured in 3D collagen matrices in the
presence of carboxyrhodamine in the culture medium (Bayless and
Davis, 2002). After 24 hr, cultures were fixed and then photographed
using fluorescence microscopy. Arrows: GFP-Rac1 labeling of EC vacuole membranes. Arrowheads: carboxyrhodamine labeling of EC vacuoles, indicating that they arise through pinocytosis. Bar ⫽ 10 ␮m.
pinocytosed vacuole compartment (Davis and Camarillo,
1996). Plasma membrane markers present in EC intracellular vacuole membranes include PECAM, caveolin-1 (Fig.
11), ICAM-1, and VCAM-1 (Davis and Bayless, 2003).
Also, ␤-catenin, a VE-cadherin and PECAM-associated
molecule (Ilan et al., 2000), is associated with vacuole
membranes, and von Willebrand factor is detectable
within many vacuoles (Fig. 11). This suggests that WeibelPalade bodies (i.e., which contain von Willebrand factor)
fuse with the developing vacuole compartment and contribute membrane and their fluid contents. The presence
of caveolin-1 also suggests the contribution of another
intracellular vesicular trafficking compartment, caveolae,
to these vacuoles (Anderson, 1998). Thus far, we have
unable to detect clathrin, early endosomal markers such
as EEA-1 or rab5, or lysosomal markers such as LAMP in
vacuole membranes.
An important consideration is, how do EC intracellular
vacuoles enlarge and fuse during the lumen formation
process? Do they solely enlarge by fusion with other vesicles, or do they enlarge by accumulation of ions such as
Na⫹ or H2O? DNA microarray or differential display studies have shown that a series of genes associated with
sodium and water transport are upregulated during EC
morphogenesis (Bell et al., 2001). These genes include
NaHCO3 cotransporters 2 and 3, and melanin-concentrating hormone, which have been reported to regulate ion or
water transport (Hawes et al., 2000; Soleimani and Burnham, 2001). It appears that EC intracellular vacuoles
represent a novel pinocytic intracellular compartment
that is utilized to regulate the lumen formation process. It
remains to be determined how EC intracellular vesicles,
such as Weibel-Palade bodies and caveolae, contribute to
the developing lumen by fusing with pinocytic vesicles
that arise through integrin-dependent signaling. It is also
important to identify which vesicular fusion pathways,
Fig. 11. Immunocytochemical characterization of EC intracellular
vacuoles. ECs were cultured in collagen matrices, and after 8 hr they
were digested out of the collagen matrices, plated on glass coverslips,
fixed with paraformaldehyde, and immunofluorescently stained with antibodies directed to PECAM, caveolin-1, and von Willebrand factor
(vWF). Also, some cells were stained with phalloidin-fluorescein to detect F-actin. Arrows: intracellular vacuole membrane staining. Arrowhead: area of increased caveolin-1 staining between EC vacuoles. Bar ⫽
25 ␮m.
such as those controlled by Rab GTPases (Zerial and
McBride, 2001), regulate the fusion of pinocytic vacuoles
to each other or to other EC vesicles.
Another event we observe during EC morphogenesis
is a presumed exocytic event in which intracellular vacuole fusion occurs with the plasma membrane. This
event creates ECs with an open or C-shaped appearance, with processes surrounding an ECM-free space.
Such images are frequently observed in electron micrographs and cross-sections of cultures after 24 hr of
morphogenesis (Figs. 2B and 8). The EC processes that
are generated following exocytosis would be free to interact with each other to form an intra- or intercellular
junctional contact with adjacent ECs (Fig. 2). This
mechanism allows ECs to form a lumenal compartment
through pinocytosis and coalescence of vacuoles, and
then, following exocytosis, creates a mechanism for ECs
to form a reversible junctional contact site (Davis and
Bayless, 2003). Our observation that single ECs can
form a lumen and junctional contact are confirmed by
the presence of similar cells present during Drosophila
tracheal tube morphogenesis (Samakovlis et al., 1996).
The junctional site could remain with a single EC or be
a site where new ECs are recruited to form multicellular
lumenal structures. In the image shown in Figure 2, it
appears that some retraction of the processes may have
occurred following this exocytic event, as ECM is
present (arrows) which is not directly in contact with
the EC processes (Fig. 2B, arrowheads). Extension of
these processes along the ECM-fluid interface would
occur until the processes contact each other or encounter similar processes from adjacent ECs. Thus, this
mechanism could create either single ECs with junctions or multicellular capillary tube structures with
junctional contacts, as shown in Figure 2A. One of the
elegant features of these mechanisms is that the lumenal spaces are already fluid-filled and free of ECM.
These mechanisms described in EC morphogenesis may
be of general utility for lumen formation for either endothelial or epithelial cells.
Regulation of EC Vacuole and Lumen
Formation by Rho GTPases
As discussed above, the EC intracellular vacuole formation process (Davis and Camarillo, 1996) is reminiscent of
other pinocytic events, such as macropinocytosis and
phagocytosis. Like vacuole formation, both of these latter
processes are dependent on actin and microtubules
(Greenberg, 1995; Davis and Camarillo, 1996; Cardelli,
2001). In addition, the inner surfaces of the pinocytosed
membranes contain F-actin (Greenberg, 1995; Davis and
Camarillo, 1996; Cardelli, 2001) (Fig. 11). Recent work
has elucidated some of the molecular components involved
in the cytoskeletal signaling pathways regulating macropinocytosis and phagocytosis. In both cases, Rho
GTPases regulate these events. Rac and Cdc42 GTPases
have been found to be required for macropinocytosis
(Cardelli, 2001; Ridley, 2001a) and Fc receptor-mediated
phagocytosis (Caron and Hall, 1998). In contrast, the Rho
GTPase regulates integrin-dependent phagocytosis (Caron and Hall, 1998; Chimini and Chavrier, 2000). Because
of the similarities of these events to EC vacuole formation,
we tested the hypothesis that Rho GTPases are required
for EC vacuole and lumen formation in 3D ECM environments.
These studies were recently described by Bayless and
Davis (2002) using human ECs cultured in either collagen
or fibrin matrices. Blockade of all three Rho GTPases (Rho, Rac, and Cdc42) with C. difficile toxin B completely blocked EC vacuole and lumen formation in collagen and fibrin gels. In contrast, selective blockade of Rho
with the C3 exoenzyme did not block vacuole and lumen
formation. These data strongly indicate a role for Rac and
Cdc42 in these events. This conclusion is supported by our
data using recombinant adenoviruses that induce expression of dominant negative Rho GTPases in ECs during the
morphogenic process. Dominant negative Rac1 and Cdc42
markedly blocked EC vacuole and lumen formation in
collagen and fibrin matrices, while dominant negative
RhoA had no effect. In addition, constitutively active
Cdc42 also blocked morphogenesis. Previous studies have
shown that either dominant negative or constitutively
active forms of these GTPases can in some instances block
Rho GTPase-mediated cellular events (Tzuu-Shuh and
Nelson, 1998; Tzuu-Shuh et al., 1998). N-WASP, a downstream effector of Cdc42, is known to be activated by
Cdc42 and PIP2. The verprolin cofilin activation (VCA)
domain of N-WASP then binds the Arp2/3 complex to
stimulate actin polymerization (Rohatgi et al., 1999; Prehoda et al., 2000). Expression of the N-WASP VCA domain
alone (i.e., which mimics N-WASP activation and stimulates Arp2/3 binding and actin polymerization) blocks EC
vacuole formation (Bayless and Davis, 2002) like constitutively active Cdc42. These data indicate a role for Cdc42
and Rac1 in regulating EC lumen formation through intracellular vacuole formation and coalescence in 3D ECM.
Additional experiments examined the distribution of
Rac1, Cdc42, and RhoA within cells during the EC morphogenic process. A series of recombinant adenoviruses
were constructed to express either constitutively active or
wild-type Rac1, Cdc42, or RhoA fused to GFP (Bayless and
Davis, 2002). ECs were induced to express these chimeric
proteins, and morphogenesis assays were performed. Cultures at various times of morphogenesis were examined by
confocal or conventional fluorescence microscopy (Figs. 7
and 10). In some cases, the fluorescent dye carboxyrhodamine was added to the culture medium to label pinocytic EC vacuoles. In Figure 10, an EC cell is shown with
multiple intracellular vacuoles that are labeled with carboxyrhodamine. The vacuole membranes are labeled with
GFP-Rac1, showing that Rac1 targets to the pinocytosed
membranes (Fig. 10). In addition, we have also shown that
GFP-Cdc42 targets to vacuole membranes. Furthermore,
we have evidence that increased expression of Cdc42 wildtype protein in ECs induces an increase in intracellular
vacuoles within ECs compared to control vectors. In addition, Cdc42 protein is upregulated during EC morphogenesis when the majority of lumenal development and expansion occurs (after 24 – 48 hrs of morphogenesis)
(Bayless and Davis, 2002). Collectively, our data indicate
that Cdc42 and Rac1 regulate intracellular vacuole formation and coalescence and localize to vacuole membranes
during the process. Our ability to label the intracellular
vacuole membranes with either GFP-Rac1 or GFP-Cdc42
will enable future studies using real-time confocal microscopy to analyze vacuole development and fusion during
EC lumen formation. As shown in the static confocal microscopic image (Fig. 7), GFP-Rac1 targets to vacuoles
which appear to be moving from the EC abluminal surface
toward the lumenal membrane, where they fuse to expand
the lumenal surface. We have observed images similar to
the image shown which indicate that small vacuoles may
be arising from areas of EC process formation. The known
involvement of Rac1 and Cdc42 in cell process formation
and vacuole formation is consistent with this observation.
These vacuoles could then fuse with the lumen and extend
the lumenal space in the direction of the EC sprout. In
previous works we have made similar conclusions based
on other images (Davis and Camarillo, 1996; Davis et al.,
Potential Mechanisms for Creating a Lumenal
Space During Endothelial and Epithelial Cell
Morphogenesis in 3D Extracellular Matrices
Intracellular vacuole formation and coalescence is one
mechanism by which EC lumens can form in 3D matrices.
Vacuole structures similar to those described in ECs have
been reported in epithelial cells. Original studies by VegaSalas et al. (1987, 1988) showed that apical domains of
polarized epithelial cells can be pinocytosed into vacuoles,
which may represent an attempt to form an intracellular
lumenal space. These vacuole membranes contain apicalselective markers. A recent study by Akhtar and Hotchin
(2001) demonstrated targeting of the Rac1 GTPase to apical pinocytosed membrane-bound vacuoles following disruption of E-cadherin mediated cell– cell contacts in
MDCK cells. This result appears similar to our findings of
Rac1 GTPase targeting to vacuoles during EC lumen formation (Bayless and Davis, 2002).
A number of other mechanisms have been described
that may regulate either EC or epithelial cell lumen formation. Well-described models of epithelial cell morphogenesis include assays using either MDCK cells or breast
epithelial cells suspended within or on the surface of various ECM substrates (Montesano et al., 1991, 1997; Pollack et al., 1997; Weaver and Bissell, 1999; Yeaman et al.,
1999; Lipschutz et al., 2000; O’Brien et al., 2001). How
lumens form in epithelial cell clusters remains unclear. A
number of possible mechanisms have been proposed for
epithelial and EC lumen formation, including apoptosis of
centrally placed cells within clusters (Lin et al., 1999),
intracellular vacuole formation (such as that described
above), autophagy (where apically placed portions of cells
are progressively removed through lysosomal degradation
of cellular components (Stromhaug and Klionsky, 2001),
intussusception (where smaller caliber vessels are generated from larger precursor vessels (Djonov et al., 2000),
and membrane-sorting events regulated by endocytic or
exocytic mechanisms. A protein complex called exocyst,
which regulates exocytosis, has been found to regulate
these latter processes during MDCK lumen formation and
morphogenesis in 3D collagen matrices (Lipschutz et al.,
2000; Lipschutz and Mostov, 2002). Further support for
this mechanism is found in studies of Drosophila tracheal
development, wherein exocytic membrane fusion events
regulate lumenal diameter of tubes (Beitel and Krasnow,
2000). Of note, lumenal diameter was not regulated
through changes in cell number. Our findings regarding
the probable role of exocytic mechanisms in EC lumen
formation strongly correlate with these findings, revealing
the importance of exocytic events in lumen formation.
When cells are suspended in ECM or invade into ECM,
they occupy physical space within that ECM. Invagination
of a localized area of EC plasma membrane from cell-ECM
contact sites creates a fluid-filled pocket in the ECM and a
fluid-ECM interface. We have observed ECs forming “suction cup”-like areas with ECM in electron micrographs
(Davis and Bayless, 2003). EC processes can migrate
along the fluid-ECM interface extending from the boundary of the “suction cup” and enclose the fluid pocket, which
creates pinocytic EC vacuoles. As expected, these vacuoles
can be readily labeled with membrane-impermeant fluorescent dyes. Similar events can regulate vacuole formation between adjacent cells. If two or more cells are in
contact with each other, invagination of plasma membrane in an area of cell– cell contact creates an ECM-free
space, which could begin the presumptive lumenal space.
We have observed this type of lumen formation in small
EC clusters. In these clusters, lumens selectively develop
in the contact area between the two cells (Davis and Camarillo, 1996) (see Fig. 5B). Thus, both cell–ECM and cell–
cell sites can initiate pinocytic events leading to vacuole
and lumen formation. Yang et al. (1999) have shown that
blocking antibodies directed to the EC cell– cell adhesion
molecules, VE-cadherin and PECAM, interfere with EC
vacuole and lumen formation.
Similar types of events appear to regulate lumen formation during EC invasion and sprouting, which can be visualized using our “angiogenesis”-like in vitro assays (Fig.
1). As described above for vacuole formation, fluid-filled
spaces in the ECM are generated during invasion events
(i.e., invasion tunnels are visualized trailing invading
ECs) (Davis and Bayless, 2003) (Bayless et al., unpublished results). These appear to be physical tunnels with a
fluid-filled center surrounded by ECM. How such tunnels
are generated remains to be determined. They may be
generated by proteolysis, but also could be created by
physical means or phagocytic removal of ECM by the
invading cell. The ECs appear to use these fluid-ECM
interfaces to migrate on and facilitate the development of
the lumenal space during EC invasion and sprouting.
They also serve as matrix guidance pathways (Davis and
Camarillo, 1995) (Bayless et al., unpublished results) to
facilitate the interconnection of ECs during the tube as-
sembly process. Intracellular vacuoles also appear to play
a critical role during this process to regulate lumen formation during invasion (Bayless et al., unpublished results).
Regulation of Capillary Morphogenesis in 3D
Matrices by Differential Gene Expression
The original assay system of Davis and Camarillo (1996)
has proven to be very useful for analyzing differential
gene expression during capillary morphogenesis in 3D
extracellular matrices (Salazar et al., 1999; Davis et al.,
2001; Bell et al., 2001). The examination of differential
gene expression has been successfully applied to many
biologic systems with the development of new large-scale
genetic screening techniques, such as DNA microarray,
differential display, and serial analysis of gene expression
(Velculescu et al., 1995; DeRisi et al., 1997; Brown and
Botstein, 1999; Martin and Pardee, 1999; Maxwell and
Davis, 2000; Peale and Gerritsen, 2001; Bell et al., 2001).
These approaches have recently been utilized to examine
differential gene expression during capillary morphogenesis in vitro (Glienke et al., 2000; Kahn et al., 2000; Bell et
al., 2001) and to compare tumor angiogenic vs. normal
endothelium in colorectal cancer (St. Croix et al., 2000).
These studies have identified changes in ECM, integrin,
and cytoskeletal regulatory protein gene expression, underscoring the importance of the MIC signaling axis in EC
morphogenesis. In the study by St. Croix et al. (2000),
many of the upregulated ECM genes from tumor-derived
ECs were characteristically produced by mesenchymal
cells such as collagen type I and collagen type III genes.
This suggests the possibility that angiogenic ECs in colorectal carcinomas are undergoing an epithelial-mesenchymal transition, which is observed when epithelial or ECs
invade interstitial matrices (Nakajima et al., 1997; Boyer
et al., 2000; Savagner, 2001). In contrast, Bell et al. (2001)
showed that marked inductions of basement membrane
matrix genes characteristic of EC differentiation were observed during EC morphogenesis in collagen matrices.
These upregulated genes included collagen type IV ␣l
chain, laminin ␥1, laminin ␣4, heparan sulfate N-deacetylase N-sulfotransferase I (the rate-limiting step enzyme in
heparan sulfate biosynthesis), lysyl oxidase (to cross-link
EC synthesized ECM components), and the ␣2 and ␣1
integrin subunits (both of which can bind to collagen type
IV and laminins) (Bell et al., 2001). In addition, another
upregulated gene was the novel, capillary morphogenesis
gene (CMG)-2, which contains a von Willebrand factor A
domain with affinity for collagen type IV and laminin, a
signal peptide, and a transmembrane segment, suggesting
its possible expression on the EC cell surface (Bell et al.,
2001). Preliminary studies have revealed a predominant
localization of CMG-2 to the endoplasmic reticulum (ER)
(Bell et al., 2001). If CMG-2 can be transported to the cell
surface, its cell surface expression may be regulated by an
ER membrane retention signal, which is a common way to
regulate the expression of cell surface receptors (Bermak
et al., 2001; Hardt and Bause, 2002).
The genes that were shown to be differentially expressed in our in vitro morphogenesis model were previously described by Bell et al. (2001). Markedly upregulated genes include jagged-1, stanniocalcin, angiopoietin-2, placental growth factor, sprouty, collagen type
IV ␣1 chain, ␣2 integrin subunit, myosin IC, alpha-2 macroglobulin, egr-1, CD39, and HMG CoA reductase. Mark-
edly downregulated genes include connective tissue
growth factor, RGS-5, RGS-4, frizzled related protein-1,
Id-1, Id-3, fibulin-3, ␤3 integrin subunit, syntenin, PAI-1,
Cdc2, cyclin A1, cyclin B2, Cdc20, and Mcm2, thymidylate
synthetase, and pentaxin. Many of the downregulated
genes are involved in the inhibition of cell cycle progression that occurs during the tube formation process. Further support for this conclusion is that several negative
regulators of cell cycle progression (i.e., p16/INK4a and
Cdc14) increase in expression during the morphogenic
process. The marked downregulation of regulator of Gprotein signaling (RGS) genes, which are GAPs for Gproteins (DeVries et al., 2000), suggests that activation of
G-protein coupled pathways may be occurring. An interesting finding regarding these RGS genes is that they are
known to negatively regulate cell migratory events. This
occurs because many of the receptors that mediate chemotactic events require G-proteins (Bowman et al., 1998).
Thus, downregulation of RGS genes may result in an
increased ability of ECs to sprout and migrate in order to
form interconnecting networks of tubes. Other classes of
genes identified during these screens include upregulation
genes associated with the JAK-STAT pathway, anti-apoptotic signals, EC differentiation genes, cholesterol biosynthesis genes, and genes associated with EC quiescence
(Bell et al., 2001) (Fig. 5). Upregulated genes that may
function to induce EC quiescence include CD39 (an ecto
ATP/ADPase) (Goepfert et al., 2000), CD26 (dipeptidyl
peptidase, which inactivates biological active peptides)
(Mentlein, 1999), A20 zinc-finger protein (which inhibits
TNF-induced NF-␬B-dependent gene transcription) (Lee
et al., 2000; Klinkenberg et al., 2001), and melanomaassociated antigen (MG50), an ECM-like protein containing a peroxidase domain, multiple Ig repeats, and the
precursor sequence of IL-1 receptor antagonist (Mitchell
et al., 2000). The melanoma-associated antigen shows homology with the Drosophila basement membrane matrix
protein, peroxidasin (Nelson et al., 1994). Other induced
genes that may play a role in EC quiescence are alpha
2-macroglobulin and sprouty. Alpha 2-macroglobulin can
sequester and inhibit the function of growth factors such
as TGF-␤, VEGF, and FGF-2 in the extracellular environment (Gonias et al., 2000). Its ability to bind growth factors is induced by conformational changes in ␣2-macroglobulin that occur through proteolytic cleavage of its bait
region. Thus, its potential role in EC quiescence depends
on proteinases synthesized and activated by ECs. We have
shown that cleaved ␣2-macroglobulin is detectable during
EC morphogenesis (Bell et al., 2001). Sprouty was originally shown by Hacohen et al. (1998) to regulate the
development of the Drosophila tracheal system (i.e., networks of epithelial tubes). It was also shown to antagonize
receptor tyrosine kinases (which are directly relevant to
angiogenic signaling) by inducing turnover of internalized
receptors (Reich et al., 1999; Wong et al., 2000). Increased
expression of sprouty by viral gene transfer blocked angiogenic responses, further supporting these conclusions
(Lee et al., 2001). Thus, all of the above genes have in
common the potential ability to suppress EC activation
through biologically active molecules such as ATP, peptides, hydrogen peroxide, IL-1, TNF, and angiogenic cytokines.
An intriguing possibility is that the basement membrane matrix itself (through proteins such as melanomaassociated antigen, laminins, and possibly by the presen-
tation of proteinase inhibitors such as TIMP-3 and TFPI-2
via heparan sulfate proteoglycans (Liu et al., 1999; Yu et
al., 2000; Herman et al., 2001)) also directly contributes to
the development of EC quiescence. A recent study by
Mettouchi et al. (2001) showed that ␣2␤1-dependent interactions with laminin cause EC cell cycle arrest in G1.
In contrast, they also showed that ␣5␤1-dependent EC
interactions with fibronectin lead to cell cycle progression
and EC proliferation. In our system, fibronectin mRNA
expression by ECs was downregulated during EC morphogenesis in collagen matrices (Bell et al., 2001). Also, downregulation of the ␤3 integrin subunit may represent another sign of EC quiescence. This protein has been
described to be associated with angiogenic ECs and to
either be expressed at very low levels, or not at all, in
quiescent ECs (Brooks et al., 1994).
We have also utilized our microassay systems to isolate
mRNA from different time points of EC morphogenesis to
create cDNA libraries representative of these different
stages (Figs. 1B and 6). Using these libraries, we isolated
novel genes that were differentially expressed during EC
morphogenesis and termed them capillary morphogenesis
genes (CMGs) (Bell et al., 2001). To date, four full-length
CMGs have been isolated, and we are in the process of
characterizing their function. The sequences of CMG-1
and CMG-2 were recently reported by Bell et al. (2001),
and the sequences of CMG-3 and CMG-4 will be reported
in forthcoming publications. CMG-2 is a 45 kDa protein
with a signal peptide, a transmembrane segment and an
N-terminal region containing a von Willebrand factor A
domain. These domains have been reported in other proteins to be ECM-binding domains (Colombatti and
Bonaldo, 1991; Perkins et al., 1999). Furthermore, CMG-2
shows considerable sequence homology with a transmembrane protein recently identified in colonic tumor angiogenic ECs (TEM8) (St. Croix et al., 2000). Like CMG-2,
TEM8 contains a von Willebrand factor domain, and this
latter receptor was also recently found to be identical to
the anthrax toxin receptor (Bradley et al., 2001). Bacterial
expression of the CMG-2 von Willebrand factor A domain
yielded a 20 kDa recombinant protein fragment with affinity for the basement membrane matrix proteins, collagen type IV, and laminin (Bell et al., 2001). Since CMG-2
is induced during EC morphogenesis alongside other basement membrane matrix proteins (see above), it is our
hypothesis that CMG-2 participates in basement membrane matrix synthesis/assembly or EC– basement membrane matrix interactions during EC morphogenesis. It
should be noted that blockade of basement membrane
matrix assembly during angiogenesis or EC morphogenesis in vitro can markedly inhibit these events (Ingber and
Folkman, 1988; Maragoudakis et al., 1988; Iruela-Arispe
et al., 1991; Bonanno et al., 2000; Bell et al., 2001).
We also have considerable information concerning another novel gene, CMG-4, a 45 kDa protein, which is
markedly downregulated during EC morphogenesis (Mavila et al., unpublished results) and is selectively expressed in ECs. This gene appears to regulate the EC cell
cycle and shows affinity for the cdc2-cyclin B complex,
which regulates the G2/M transition. Furthermore, the
gene efficiently targets to the EC nucleus and is degraded
by the proteasomal machinery, a characteristic of many
cell cycle regulatory proteins (Koepp et al., 1999). Preliminary work with CMG-3, which is induced during EC
morphogenesis, suggests that it associates with the EC
Fig. 12. Capillary tube formation and maintenance in three dimensions is regulated by a balance of EC morphogenesis (M) and regression
(R). The indicated factors regulate the described steps in a stimulatory
(⫹) or inhibitory (–) manner.
cytoskeleton. CMG-3 contains a pleckstrin homology domain as well as a coiled-coil domain. A GFP-CMG-3 fusion
protein targets to submembranous areas in ECs resembling lamellipodia (Mavila et al., unpublished results).
These data from our laboratory, as well as from others,
reveal the central importance of the MIC signaling axis in
EC tube formation, as well as new insights into how EC
differentiation occurs during this process. This work also
strongly supports the utility of in vitro models for investigating capillary morphogenesis where specific hypotheses can be formulated and then directly tested under defined conditions. These developing technologies allow for a
rapid assessment of the contribution of particular genes to
distinct stages of capillary morphogenesis in 3D ECM
Capillary Tube Formation and Maintenance: A
Balance Between Morphogenesis and
The formation of blood vessels during development and
following tissue injury responses is balanced by signals
regulating both capillary morphogenesis and regression
(Fig. 12). In normal wound-healing responses, the formation of vessels in granulation tissue is followed after several days by a capillary regression response (Clark, 1996).
The final stages of this capillary regression response characteristically overlap with wound contraction (Clark,
1996). We recently described a model of capillary tube
regression in vitro wherein these two processes occur concurrently (Davis et al., 2001). Thus, regression of these
tube structures occurs alongside contraction of the collagen matrices in which they are suspended. This result
describes an understudied ability of ECs to be highly
contractile and capable of exerting strong mechanical
forces on the ECM. As discussed above, this ability helps
ECs locate each other in 3D space by creating “matrix
guidance pathways” that represent physical channels or
aligned ECM fibrils that facilitate directed cell migration
(Vernon and Sage, 1995; Davis and Camarillo, 1995).
The molecular mechanisms regulating capillary tube
regression are for the most part unknown. Elucidation of
this pathway is critical to our ability to induce regression
of tumor-associated angiogenic blood vessels to inhibit
tumor growth and progression (Folkman, 1997). A variety
of studies have identified molecules that appear to induce
tumor vessel regression in experimental animals, including ECM protein fragments, integrin targeting reagents,
and fragments of proteins that regulate hemostasis
(Brooks et al., 1994; O’Reilly et al., 1994, 1997; Sage, 1997;
Browder et al., 2000; Maeshima et al., 2002).
A variety of findings have been made that appear to
address how vascular regression may be regulated during
normal physiologic events such as development, the female menstrual cycle, and wound-healing. Work by Benjamin and Keshet (1997) and Benjamin et al. (1999) has
shown that withdrawal of VEGF induces marked capillary
regression in in vivo models where tumor cells express
VEGF under the control of a tetracycline-regulated promoter. Hyperoxia, which can cause physiological suppression of VEGF expression, is often a problem in the treatment of premature infants, and can cause severe visual
impairment due to decreased VEGF-induced signaling in
the retina (Hellstrom et al., 2001a). In contrast, hypoxia
induces VEGF expression and other hypoxia-regulated
genes, such as PAI-1, a plasminogen activator inhibitor
(Pinsky et al., 1998). PAI-1 appears to play a role along
with VEGF in promoting vascular stability (see below).
Furthermore, findings have shown that angiopoietin-2 can
participate in vascular regression by competing for the
binding of angiopoietin-1 to the tie-2 receptor (Suri et al.,
1996; Holash et al., 1999; Yancopoulos et al., 2000) . Other
studies have revealed elevations of angiopoietin-2 relative
to angiopoietin-1 during physiological capillary tube regression in the ovary and adipose tissue (following leptin
treatment) (Goede et al., 1998; Cohen et al., 2001).
The tendency of EC tubes to regress after formation
appears to be affected by a number of factors. One such
factor appears to be the association of specialized vascular
smooth muscle cells (i.e., pericytes) with EC tubes (Allt
and Lawrenson, 2001; Hellstrom et al., 2001b). Disruption
of this interaction is observed in a variety of genetically
altered mice (e.g., angiopoietin-1, edg-1, endoglin, and
TGF-␤ knockout mice) (Dickson et al., 1995; Suri et al.,
1996; Pepper, 1997; Lee et al., 1999; Li et al., 1999; Liu et
al., 2000; Urness et al., 2000), as well as in human diabetic
retinopathy (Sima et al., 1985; Ruggiero et al., 1997). This
disruption causes marked instability of newly formed
blood vessels or preexisting vessels. In the models described above using tumors expressing VEGF in a regulated fashion, the susceptibility of those vessels to VEGF
withdrawal correlated with the presence of pericytes (Benjamin et al., 1999). Those vessels with pericytes were
much more resistant to VEGF withdrawal than those
without. However, it is not known how pericytes contribute to this stability. Histologic analysis of angiopoietin-1
knockout mice revealed abnormally dilated and tortuous
vessels, which led to embryonic lethality (Suri et al.,
1996). Some ECs in these vessels did not appear to be
adhering well to their underlying ECM. In support of such
conclusions is a recent report describing the ability of
angiopoietin-1 to directly promote EC cell attachment
through integrins (Carlson et al., 2001). Another possibility is that the pericytes are contributing factors in the
specialization of the ECM environment underlying ECs.
One important example of basement membrane matrix
specialization is the presence of unique laminin isoforms
in EC basement membranes. Our recent data suggest the
presence of at least laminin-8 and laminin-10 isoforms
based on the differential regulation of laminin subunit
mRNAs during EC morphogenesis (Bell et al., 2001). This
conclusion is also supported by previous results (Miner et
al., 1998; Lefebvre et al., 1999). These mRNAs were upregulated during later stages of morphogenesis, suggesting that these laminins may play an important role in the
maturation and stabilization of developing tubes. There
are many examples of epithelial–mesenchymal interactions which are necessary for basement membrane matrix
formation and specialization (Kadoya et al., 1997; Smola
et al., 1998; Hedin et al., 1999; Erickson and Couchman,
2000). These interactions contribute different combinations of basement membrane components (laminin and
collagen type IV isoforms, etc.) which are critical to basement membrane assembly. The extent to which pericytes
contribute to EC basement membrane matrix formation
and maintenance is unclear.
Since basement membrane matrix contributes to the
formation, stability, and maintenance of blood vessels, its
proper formation may underlie many of the described
EC–pericyte interaction defects reported in the various
genetic knockout models. Such defects include failure to
properly recruit pericytes to the EC tubes, and defects in
the EC–pericyte interactions necessary to stabilize the
tube. One likely participant in this event is TGF-␤, including its receptors and the signaling pathway molecules
downstream of these receptors (Dickson et al., 1995; Pepper, 1997; Li et al., 1999; Urness et al., 2000). It has been
well described that ECs or pericytes alone are unable to
properly activate secreted TGF-␤ (Sato et al., 1990;
Munger et al., 1997; Hirschi et al., 1998). In contrast,
when both cells are combined, latent TGF-␤ can be activated by a mechanism that is typically mediated through
plasmin-mediated proteolysis. TGF-␤ is well known to
markedly stimulate ECM synthesis and the production of
factors such as PAI-1, which can facilitate ECM stability
by inhibiting matrix proteolysis (Pepper, 1997; Denton
and Abraham, 2001). Another very interesting finding in
this regard is that the EC-derived cytokine, connective
tissue growth factor (CTGF), markedly enhances the ability of TGF-␤ to stimulate matrix production and deposition (Grotendorst, 1997; Denton and Abraham, 2001). In
our DNA microarray work, the mRNAs of CTGF and a
related cytokine (Cyr61) were markedly downregulated
early during morphogenesis but were then returned to
near-baseline levels as EC morphogenesis progressed
(Bell et al., 2001). Other regulated cytokines, such as
PDGF-B, which was induced during EC morphogenesis,
may also play a role in this process by regulating events
such as pericyte recruitment to EC tubes by stimulating
migration and proliferation. CTGF can similarly induce
smooth muscle cell proliferation and migration (Fan et al.,
2000). Thus, EC–pericyte interactions by means of TGF-␤,
CTGF, and PDGF may be responsible for ensuring the
formation, stability, and maintenance of EC-lined capillary tubes through effects on a number of events, such as
EC basement membrane matrix assembly and EC quiescence.
Recent efforts to examine gene expression changes during EC morphogenesis offered some insights into how a
balance between EC morphogenesis vs. regression may be
regulated (Fig. 12). It remains to be determined whether
EC tube regression genes are differentially regulated during EC morphogenesis, and whether genes exist whose
primary purpose is to regulate the process of EC tube
regression rather than EC tube formation. At the moment
it is not clear that such genes exist, but often in studies of
biological systems, molecules that regulate positive events
are identified first, followed by molecules that regulate
negative events. An excellent example of this point are
studies of nervous system development wherein factors
that stimulate neurite outgrowth were discovered first,
while factors that interfere with this process (e.g., collapsins and semaphorins) were discovered later (Yu and
Bargmann, 2001). It has recently become clear that there
is a striking overlap between factors that regulate neural
growth and vascular growth. For example, the recent discovery by Soker et al. (1998) and Miao and Klagsbrun
(2000) that the semaphorin III receptor, neuropilin, is a
VEGF receptor indicates that there are similarities between these systems. These investigators have also shown
that semaphorin III blocks both capillary tube sprouting
and neurite outgrowth (Miao et al., 1999). In addition, we
have detected semaphorin III mRNA in ECs and shown
that its expression is markedly downregulated during the
period of sprouting and branching that occurs during EC
morphogenesis (unpublished observations). Sprouting EC
processes often contain “growth cone”-like projections
which regulate the directional navigation of ECs in three
dimensions (Speidel, 1933; Davis et al., 2000b). The extent
to which other regulators of neural growth cone guidance
may play a role in EC morphogenesis or regression is not
yet known (Yu and Bargmann, 2001).
Differentially regulated genes that could play a direct or
indirect role in regulating the process of EC tube regression
are MMP-1, PAI-1, tissue factor pathway inhibitor (TFPI)-2,
gelsolin, angiopoietin-2, thrombospondin (TSP)-1, and
TSP-2. The latter two ECM proteins have been shown to
negatively regulate blood vessel formation, with TSP-2
appearing to have the most potent activity (Kyriakides et
al., 1998; Streit et al., 1999; Adams, 2001). In a study by
Bell et al. (2001), TSP-1 mRNA was downregulated during
EC morphogenesis while TSP-2 was upregulated. In another work (Davis et al., 2001), PAI-1 (which negatively
regulates plasmin- and MMP-dependent capillary tube
regression) was markedly downregulated as well. In recent studies (Rao et al., 1996; Herman et al., 2001) using
adenoviral gene transfer, upregulated expression of the
serine and MMP proteinase inhibitor, TFPI-2, in ECs
blocks either plasminogen/plasmin or plasma kallikreininduced capillary tube regression (Davis et al., unpublished results). Our model of regression involves not only
ECM degradation but also EC tube collapse and apoptosis
(Davis et al., 2001) (Davis et al., unpublished results). We
previously reported that gelsolin is markedly upregulated
during EC morphogenesis (Salazar et al., 1999). Gelsolin
has been identified as a major caspase-3 target and an
important regulator of apoptosis (Kothakota et al., 1997;
Kamada et al., 1998). In our model of EC capillary tube
regression, caspase-dependent cleavage of gelsolin was
detected (Davis et al., 2001). Thus, gelsolin may be induced in part to regulate the process of EC tube regression
if this process is initiated. The most induced gene (of more
than 7,000 genes profiled) detected at 48 hr of culture
during EC morphogenesis was angiopoietin-2. Angiopoietin-2 has been reported to antagonize the function of
angiopoietin-1 by competing with its binding to the Tie-2
receptor (Suri et al., 1996; Yancopoulos et al., 2000) and it
also has been reported by other groups to be produced by
Fig. 13. The EC–target hypothesis. An analogy between neuron–
target interactions and EC–target interactions is shown. This hypothesis
suggests that EC networks interact with target cells to stabilize and
develop specialized characteristics. These interactions also prevent
capillary tube regression events. Potential EC target cells include pericytes/VSMCs, astrocytes, podocytes, and alveolar epithelial cells.
ECs (Mandriota and Pepper, 1998). The important question here is, why do ECs make angiopoietin-2? The induction of angiopoietin-2 by ECs during morphogenesis could
be envisioned as an attempt by ECs to regress unless some
signal is provided to overcome this inhibitory signal.
An analogy that occurs to us in this regard refers again
to relationships between neural and vascular systems.
During peripheral nervous system development, neurons
extend axonal processes to targets such as skeletal muscle
to form synaptic contacts. These neuron–target interactions provide a trophic role to maintain survival of the
neurons and to stabilize both cell types (Ernfors, 2001;
Sanes and Lichtman, 2001). Do developing vascular networks need an analogous target to provide a similar type
of signal to facilitate EC tube survival and stabilization?
We hypothesize that pericytes/vascular smooth muscle
cells (VSMCs) represent such a target in most tissue environments (Fig. 13). The EC–pericyte interaction would
serve to stabilize and support the survival of the developing tube, and through these effects would facilitate the
development of EC specializations. Such specializations
may include properties characteristic of arterial, capillary,
or venous ECs. The ephrins, which regulate neural development, have been reported to regulate arterial vs. venous
identity for ECs in these locations (Wang et al., 1998;
Wilkinson, 2001). In addition, EC–target interactions may
regulate the development of unique EC properties characteristic of different tissue locations. Also, in other specialized tissues such as the brain, kidney, and lung, ECs are
known to closely interact with astrocytes, podocytes, and
alveolar eptithelial cells, respectively. These other cell
Fig. 14. VSMCs prevent plasminogen-induced capillary tube regression in EC-VSMC cocultures in 3D collagen matrices. ECs were cultured in the
presence or absence of either human coronary artery smooth muscle cells (CASMCs) or dermal fibroblasts (HDFs). The CASMCs and HDFs were added
at 5 ⫻ 105 cells/ml of gel, and the ECs were added at 106 cells/ml of gel. Culture media contained plasminogen (Plg) at 1 ␮g/ml, or contained none, as
previously described (Davis et al., 2001). A: After 72 hr, cultures were fixed, stained, and photographed. The circular dark structures represent contracted
collagen gels. Arrowheads indicate CASMCs. B: Conditioned media samples collected from the indicated cultures were run on SDS-PAGE for
immunoblotting (left panel) with anti-MMP-1 antibodies, or for gelatin zymogram analysis (right panel). Left panel, arrowhead: pro-MMP-1; arrows: activated
MMP-1 forms. Right panel, arrowheads: pro-MMP-9 and pro-MMP-2; arrows: activated forms of these enzymes which directly underlie the proenzyme
bands. C: Indicated conditioned medium (CM) samples were added at a 1:4 dilution to EC-only cultures to determine whether CASMC CM could block
the capillary tube regression and collagen gel contraction response. Cultures were fixed after 48 hr and photographed. Bar ⫽ 25 ␮m.
types may perform functions similar to those of pericytes
to regulate EC stability and create further EC specializations. This EC–target hypothesis may explain many of the
observations regarding the instability of newly formed
tubes in the absence of pericytes or other supporting cells.
Furthermore, it may explain data explaining why factors
such as angiopoietin-2, CTGF, PDGF, and TGF-␤ are produced by ECs during the morphogenic process. These factors are probably synthesized by ECs to facilitate the
development of this EC–target interaction. If the EC–
target interaction does not occur, the EC tubes will regress, which is analogous to what happens when neurons
fail to reach an appropriate target and undergo apoptosis
(Ernfors, 2001). With respect to the EC–pericyte/VSMC
interaction, it is notable that ECs can synthesize neurotransmitters such as acetylcholine and nitric oxide to act
on VSMC targets, similar to what is observed in neuron–
target interactions.
A major new insight into the molecular regulation of
capillary tube regression stems from recent studies showing that ECM proteolysis, which disrupts the matrix scaffold regulating EC morphogenesis, induces capillary tube
regression and EC apoptosis (Zhu et al., 2000; Davis et al.,
2001). This relatively simple model of capillary tube regression has considerable relevance for the development
of strategies and reagents aimed at inducing vascular
regression, and is particularly relevant in the context of
cancer. This work is highly related to the pioneering work
of Werb and colleagues, who studied the molecular control
of mammary gland involution following the cessation of
lactation (Talhouk et al., 1992; Sympson et al., 1994; Werb
et al., 1996). In this model, it is clear that ECM proteolysis
plays a major role in epithelial tube regression as well as
the development of adipose tissue to replace this regressing tissue (Selvarajan et al., 2001). Other studies (Marbaix et al., 1996; Kokorine et al., 1996) have revealed an
important role of MMP-1 in, for example, the menstrual
cycle and endometrial shedding, where it was shown to be
hormonally regulated and to be involved in this cyclical
tissue regression event.
Several laboratories, including ours, have found evidence that vascular regression is controlled by ECM proteolysis. In one set of studies, it was observed that PAI-1
knockout mice demonstrated poor angiogenic responses,
and that, consequently, tumors grew poorly in these mice
(Bajou et al., 1998, 2001). In the absence of PAI-1, an
increase in matrix dissolution occurs via plasmin-dependent degradation of ECM (see below) which destabilizes
ECM environments supporting EC morphogenesis. In addition, PAI-1 has been observed to be a hypoxia-induced
gene (Pinsky et al., 1998). Thus, hypoxia induces ECM
stability to promote angiogenic responses. In our model of
capillary tube regression in vitro, PAI-1 was shown to
negatively regulate plasmin and MMP-dependent regression events (Davis et al., 2001). Blocking antibodies to
PAI-1 markedly accelerates the regression response, and
the addition of PAI-1 to this model blocks capillary tube
regression. Another study (Bacharach et al., 1998) showed
that PAI-1 expression was increased by EC–fibroblast interactions to inhibit proteolysis and stabilize the ECM
environment during angiogenesis. Thus, in some cases,
factors that promote ECM stabilization may be pro-angiogenic while factors that promote ECM dissociation/proteolysis may be anti-angiogenic.
Key regulators of this proteinase-dependent capillary
tube regression response are plasmin and MMPs (such as
MMP-1) (Fig. 3). Plasmin is capable of degrading fibrin, a
major ECM scaffold for EC morphogenesis, and of activating MMPs such as MMP-1, MMP-3, and MMP-9, which
can degrade other critical ECM scaffolds, such as collagen
types I and IV, laminin, and fibronectin (He et al., 1989;
Jeffrey, 1998; Lund et al., 1999; Ramos-DeSimone et al.,
1999). In our serum-free system, the addition of plasminogen leads to the generation of plasmin through EC-derived plasminogen activators. This in turn activates
MMP-1 to initiate collagen type I proteolysis and denaturation, which leads to capillary tube regression (Davis et
al., 2001). Without plasminogen addition, ECs fail to activate MMP-1 (based on assays of conditioned media) and
do not regress. Activated MMP-1, which degrades native
collagen, acts in concert with activated MMP-9 (activated
by plasmin) and MMP-2 (activated by MT-MMPs), which
degrade denatured collagen, to cause capillary tube regression and collagen gel contraction. The addition of proteinase inhibitors such as TIMP-1, PAI-1, and aprotinin,
which block MMPs, plasminogen activators, and plasmin,
respectively, block the regression response. We have also
recently found that the addition of activated plasma kallikrein can independently activate MMP-1 to initiate the
capillary tube regression response (Davis et al., unpublished results). This experiment was performed in response to similar observations recently reported by Selvarajan et al. (2001) that plasma kallikrein can act in
concert with plasmin to facilitate mammary gland regression and adipose tissue development. In our experiments,
plasma kallikrein and plasmin synergistically acted to
induce the capillary tube regression response by initiating
the activation of MMP-1 (Davis et al., unpublished results).
To address the hypothesis that VSMCs may serve as EC
target cells that facilitate the stabilization of capillary
tubes, we performed coculture experiments with ECs and
VSMCs. We sought to determine whether plasminogen/
plasmin-induced capillary tube regression would occur in
the presence of VSMCs. EC-dermal fibroblast cocultures
were used as a control. As shown in Figure 14, the addition of VSMCs to EC cultures completely blocked the
capillary tube regression response, while dermal fibroblasts did not. Capillary tube regression is accompanied
by collagen gel contraction, which indicates that regression has occurred (Fig. 14A). Conditioned medium from
VSMCs suspended in collagen gels also blocked capillary
tube regression and collagen gel contraction, suggesting
that a soluble factor was involved (Fig. 14C). Examination
of MMP-1 and MMP-9 activation revealed that the presence of VSMCs markedly blocks MMP activation, which
explains the inhibitory effect (Fig. 14B). The nature of the
VSMC-derived capillary regression inhibitor is not known
at present. These data show that VSMCs can stabilize
developing tubes that would otherwise regress, suggesting
that inhibition of ECM proteolysis may represent one
mechanism by which pericytes/VSMCs stabilize EC tubes.
Other experiments support the general conclusion that
ECM proteolysis is a key regulator of capillary tube regression responses. In a recent study, transgenic animals
overexpressing TIMP-1 showed increased vascularity
(Yamada et al., 2001), which could be indicative of decreased vascular regression (Fig. 12). TIMP-1 and PAI-1
may be particularly important capillary regression inhib-
itors because of their ability to block EC tube regression
(Davis et al., 2001) and inability to block EC morphogenesis (Davis et al., unpublished observations). Marked
overexpression of TIMP-1 or PAI-1 using recombinant adenoviral vectors in ECs does not block morphogenesis, but
completely inhibits plasminogen/plasmin or plasma kallikrein-induced and MMP-dependent capillary tube regression. Furthermore, recent studies have indicated surprisingly high levels of TIMP-1 in the plasma of patients
with a variety of cancers, including colon, gastric, breast,
and lung cancer (Pellegrini et al., 2000; Yoshikawa et al.,
2001; Yukawa et al., 2001). These data indicate that
TIMP-1 and PAI-1, and perhaps other proteinase inhibitors synthesized by tumors, may block the vascular regression response to facilitate the maintenance of the tumor angiogenic vasculature. Thus, proteinase inhibitors
may be critical positive regulators of tumor angiogenesis
by blocking vascular regression. This new information
suggests that ECM-degrading proteinases, which clearly
play a role in EC morphogenesis, are also critical regulators of capillary tube regression in 3D ECM environments.
In this review, we have addressed many issues regarding how EC morphogenesis and regression are regulated
at a molecular level in 3D extracellular matrices. It is
apparent that this field is still developing, and that considerable information about these processes remains to be
discovered. The molecular tools and cell biological approaches necessary to answer the basic question of how
ECs assemble or disassemble in three dimensions are
currently available. It is our hope that this review will
provide some new insights into how the study of basic
mechanisms regulating EC morphogenesis or regression
will rapidly lead to the identification of novel molecular
targets to modulate these events in the context of human
The authors thank Dr. Scott Bell and Dr. René Salazar,
who made many contributions to our current understanding of EC morphogenesis in 3D ECM environments. This
work was supported in part by grants from the NIH
(NHLBI; HL59373 to G.E.D., and F32 HL69603 to K.J.B.)
and the Texas Higher Education Coordinating Board (8957-2001 to G.E.D.).
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