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INTRODUCTION |
During angiogenesis, proliferating endothelial cells
(ECs)1 organize to form new
three-dimensional capillary networks. This process has been studied
extensively in the embryo, establishing that an early stage of
angiogenesis involves transition of endothelial precursor cells to a
spindle-shape morphology (1) in combination with alignment into solid,
multicellular, pre-capillary, cord-like structures (2, 3). Moreover,
these cord-like structures are interconnected to form a polygonal
network (1, 4). Solid pre-capillary cords have also been observed
during angiogenesis in the adult (5). These solid cords subsequently
mature into tubes with hollow lumens for the transport of blood (1,
5).
Three-dimensional type I collagen provokes ECs in culture to undergo
marked shape changes that closely imitate pre-capillary formation
during embryonic angiogenesis. Within hours after addition of collagen
I to confluent cultures, ECs partially retract and exhibit a
spindle-shaped morphology, together with re-alignment to form solid
cords organized in a polygonal pattern (6-10). Subsequently, over the
course of several days, these structures mature to form tubes with
hollow lumens through a process involving development and coalescence
of intracellular vacuoles (11).
Consistent with the importance of collagens in regulating EC shape and
multicellular organization into pre-capillary cords, there exists
considerable evidence that interactions between collagens and ECs are
highly relevant in vivo. For example, in the developing embryo, blood vessels arise from the organization of EC precursors within an extracellular matrix rich in collagens. Furthermore, during
angiogenesis in the adult, ECs within existing blood vessels degrade
basement membrane (12) and migrate and proliferate within connective
tissue abundant in interstitial collagens (13). Consistent with the
importance of collagen/EC interactions for angiogenesis, we reported
previously that vascular endothelial growth factor (VEGF) induces
microvascular ECs (MVECs) to express integrins
1
1 and
2
1
(14). Both of these integrins are key collagen receptors on MVECs, and
antagonism of these two integrins inhibits dermal and tumor
angiogenesis in vivo (14, 15), consistent with the
importance of interactions between collagens and ECs. Also, recent
analyses of genes expressed in human tumor endothelium demonstrated
that ECs isolated from tumors express >10-fold more transcripts
encoding collagens type I and III than ECs isolated from corresponding
control tissue, indicating that tumor ECs express their own
interstitial collagens (16). These findings suggest the interesting
possibility that interstitial collagen expression by tumor ECs is
conducive for angiogenesis. In support of this hypothesis, expression
of collagen I by isolated EC clones in vitro closely
correlates with spontaneous multicellular organization into cords (17,
18). Finally, neovascularization was inhibited in animal models both by
proline analogues, which interfere with collagen triple helix assembly,
and by
-aminopropionitrile, which inhibits collagen cross-linking
(19), indicating that collagens play a crucial role in angiogenesis.
Despite the abundant evidence indicating that interstitial collagens
and their receptors are important for angiogenesis, little is known
regarding the signaling events and mechanisms through which collagen
regulates EC morphology and multicellular alignment into cords.
Therefore, we designed experiments to identify key mechanisms involved.
Our studies identify a series of critical steps beginning with collagen
binding to integrins
1
1 and
2
1, followed by
1
1- and
2
1-mediated suppression of cyclic AMP and
cyclic AMP-dependent protein kinase A (PKA), and induction of actin polymerization.
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EXPERIMENTAL PROCEDURES |
Cells and Reagents--
Human dermal MVECs were isolated from
neonatal foreskins and cultured as previously described (9).
Fibroblasts were obtained as outgrowths from human neonatal foreskins,
and they were maintained in Dulbecco's modified Eagle's medium
containing 10% FCS. Cells were used at the fourth to seventh passage.
Purified recombinant human VEGF165 was obtained from the
NCI Preclinical Repository, Biological Resources Branch, Frederick, MD.
Sources of other reagents were as follows: rat tail collagen I (BD
Biosciences, Bedford, MA); mouse laminin-1 (Invitrogen, Carlsbad, CA),
camptothecin, forskolin, adenosine 3',5'-cyclic monophosphate
8-(4-chlorophenylthio)-sodium salt, 3-isobutyl-1-methylxanthine, pertussis toxin, MDL 12,330A, 2',3'-dideoxyadenosine, KT5720, and
cell-permeable PKA inhibitor 14-22 amide (Calbiochem, San Diego, CA);
DAPI, calcein AM, propidium iodide, latrunculin A, FITC-phalloidin, and
Texas Red-DNase (Molecular Probes, Eugene, OR).
The following monoclonal antibodies (mAbs) were used: mouse anti-human
integrin
1 (clone FB12), mouse anti-human integrin a2 (clone P1E6), and rat anti-human
6 (clone
NKI-GoH3), all from Chemicon International, Temicula, CA; isotype IgG1
control mAb (anti-TNP, clone 107.3) was from Pharmingen, La Jolla, CA.
For mAb clustering, either secondary goat anti-mouse or goat anti-rat Fc-specific antibodies (Abs) (Sigma, St. Louis, MO) were employed, as appropriate.
Stimulation of MVECs with Collagen I--
Cells were grown to
confluence in a standard MVEC medium consisting of EBM-2 (Clonetics,
San Diego, CA), 10% FCS, cyclic AMP (cAMP), and hydrocortisone (9).
24 h before use, the culture medium was removed and replaced with
a simplified medium consisting of EBM-2 and 10% FCS to avoid any
complications associated with the presence of exogenous cAMP.
Acid-solubilized rat tail collagen I was neutralized and made isotonic
according to the manufacturer's instructions and diluted in serum-free
medium (EBM-2, Clonetics, San Diego, CA) to a concentration of 500 µg/ml unless indicated otherwise. Culture medium was gently removed
from the cells and carefully replaced with the collagen-containing
serum-free medium. Upon return of the cells to 37 °C, the collagen
typically polymerized within 30 min. We found it important to minimize
disturbance of the cells during the removal and replacement of the
medium, because turbulent medium changes were observed to alter
baseline cAMP. In addition to laminin-1 as a control, other controls
for all of these experiments consisted of replicate wells to which we added the collagen solubilization vehicle (0.02 M acetic
acid), neutralized and made isotonic identically to the collagen I
solution and subsequently diluted identically to collagen in serum-free EBM, and replicate wells without a medium change. When following precautions for careful removal and replacement of the medium, we found
no significant differences among controls regarding all parameters investigated.
Cell Survival Assays--
To detect apoptosis, MVECs were
stained with FITC-Annexin-V (ApoTarget Kit from BioSource, Camarillo,
CA), according to the manufacturer's instructions. Staining was
measured with a SLT Spectrafluor fluorescent plate reader (excitation,
485 nm; emission, 535 nm). For positive controls, apoptosis was induced
by incubating cells with camptothecin (1 µg/ml) for 6 h. Cell
viability was measured with an established method involving the
fluorescent calcein AM substrate (20, 21). Substrate fluorescence,
indicative of live cells, was measured with a fluorescent plate reader
(485-nm excitation, 535-nm emission). Cell death was assessed with
propidium iodide staining (4 µM final concentration),
which identifies loss of membrane integrity. Staining was measured with
a fluorescent plate reader (590-nm excitation, 635-nm emission).
Treatment of cells with Triton X-100 (0.01%) served as a positive
control. To compare cell numbers, cells were fixed in buffered
formalin, permeabilized with 0.01% Triton X-100, and stained with DAPI
(400 ng/ml final concentration), which fluoresces upon binding to DNA. DAPI fluorescence was measured with a fluorescent plate reader (360-nm
excitation, 465-nm emission).
Integrin Clustering and Blocking Experiments--
For
experiments with integrin mAbs, expression of integrins
1
1 and
2
1
on the surface of MVECs was induced maximally by stimulating cells for
3 days with 20 ng/ml VEGF (14). To cluster integrins, integrin mAbs
were incubated with cells in serum-free medium at a concentration of 50 µg/ml for 2 h, and secondary Fc-specific Ab (50 µg/ml) was
added for 4 h. To block integrin ligation of collagen I, integrin
mAbs were added at 10 µg/ml.
Assays for F-actin, G-actin, cAMP, and PKA--
For quantitative
measurement of polymerized filamentous actin (F-actin) and globular
actin (G-actin), cells on 96-well plates were fixed directly by adding
an equal volume of phosphate-buffered formalin to the medium for 30 min. Plates then were washed by immersion in phosphate-buffered saline.
F-actin was measured according to an established method (22) with the
following modifications. Fixed cells were incubated for 45 min with
FITC-phalloidin (200 mM) in phosphate-buffered saline
containing 0.1% Triton X-100. Subsequently, the plates were washed
five times, blotted, and analyzed with a spectrofluorometric plate
reader (excitation, 485 nm; emission, 535 nm). Additional wells lacking
cells were processed in parallel, and they served as background
controls. G-actin was measured according to an established method (23). Plates were processed as described for F-actin assays except that fixed
cells were incubated with Texas Red-DNase (300 nM).
Fluorescent plate reader settings employed were as follows: excitation,
590 nm; emission, 630 nm.
For cAMP analyses (24-well plates), culture medium was removed and
cells were extracted immediately with a
20 °C solution of 95%
ethanol containing 500 µM 3-isobutyl-1-methylxanthine to inhibit phosphodiesterase activity. Samples were evaporated to dryness
in a Speed-vac, and cAMP was measured with a cAMP enzyme-linked immunoassay kit (BioTechnologies, Stoughton, MA). PKA activity was
measured with an assay kit (Calbiochem #539490) that measures incorporation of radiolabeled phosphate from [
-32P]ATP
into a highly specific peptide substrate for PKA (LRRASLG). The
extraction buffer for PKA also was supplemented with 500 µM 3-isobutyl-1-methylxanthine to inhibit
phosphodiesterase activity after cell lysis.
Microscopy--
Fluorescence and phase-contrast images were
collected with a Leica DC200 digital camera and associated software.
For labeling of the actin cytoskeleton, cells were prepared and
incubated with FITC-phalloidin as for the F-actin quantitative assays
described above.
 |
RESULTS |
Through
1
1 and
2
1 Integrins, Collagen I Initiates a
Morphogenetic Program in Dermal MVECs but Not Dermal
Fibroblasts--
Conceivably, there are several strategies for
studying collagen signaling in MVECs, including approaches that begin
with cells in suspension. However, during the early stages of
angiogenesis, adherent MVECs proliferate, invade, and re-organize in a
three-dimensional matrix rich in interstitial collagens. Therefore, we
chose a model system involving the addition of three-dimensional
interstitial collagen I to adherent, confluent MVECs. This model better
represents the conditions encountered by MVECs at the sprouting tips of
blood vessels as they encounter the collagen-rich interstitial matrix.
As illustrated in Fig. 1A,
adherent dermal MVECs responded to addition of collagen I with partial
retraction and re-alignment into polygonal arrays of cord-like
structures within 6 h. As described in the introduction, this
morphogenetic process closely imitates an early organizational step in
the formation of vascular plexuses during embryonic development
in vivo (1, 4). In contrast to collagen I, equivalent
concentrations of laminin-1 were without effect (not shown). Also,
dermal fibroblasts did not respond detectably to collagen I,
establishing that the morphogenetic response of MVECs to collagen I
does not apply generally to all cell types (Fig. 1A).

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Fig. 1.
Through the
1 1 and
2 1 integrins,
collagen I provokes morphogenesis of dermal MVECs but not dermal
fibroblasts. A, representative microscopic fields
illustrating that addition of 500 µg/ml collagen I to confluent human
dermal MVECs provokes cell retraction and reorganization of the
monolayer into cord-like structures within 6 h. By contrast, human
dermal fibroblasts neither retract nor undergo morphogenesis.
Bar = 100 µm. B, collagen-induced
morphogenesis of MVECs is not attributable to apoptosis, as indicated
by the lack of Annexin-V staining (for positive controls, cells were
treated with camptothecin to induce apoptosis). Moreover, staining with
calcein AM, which identifies live cells, indicated that cells were
equally viable in the presence and absence of collagen I. Also,
propidium iodide staining, which identifies cells with permeabilized
membranes, did not distinguish between collagen-treated cells and
controls; cell number, compared by quantifying fluorescence of DAPI,
which binds DNA, was equivalent in both groups. Error
bars = S.D. C, neither 1-blocking
mAb nor 2-blocking mAb alone blocked collagen I-induced
alignment of MVECs into cords; however, both mAbs in combination were
highly effective. Bar = 100 µm.
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Over more extensive time intervals, three-dimensional collagen has been
found to impair EC survival (24). Therefore, we performed a series of
control experiments to test the possibility that morphological changes
we observed were related to induction of apoptosis or loss of cell
viability or cell number. However, as shown in Fig. 1B,
collagen I did not induce apoptosis or cell death in our experiments,
as determined by Annexin-V staining and propidium iodide staining,
respectively. Furthermore, collagen I did not reduce cell viability, as
determined with the live cell marker, calcein AM (20, 21), and the cell
number was not affected (Fig. 1B). Therefore, we conclude
that neither induction of apoptosis nor loss of cell viability or cell
number contributed to the morphological changes induced by collagen I
in our experiments.
Next, we performed experiments with blocking mAbs to test the
involvement of two prominent collagen receptors, the integrins
1
1 and
2
1,
in mediating the shape changes induced in MVECs by collagen I. As shown
in Fig. 1C, antagonism of either of these integrins
individually failed to block collagen-induced changes in cell shape,
but antagonism of both integrins in combination was highly effective.
These findings implicate both
1
1 or
2
1 in mediating the shape changes
provoked by collagen I. Similar to dermal MVECs, dermal fibroblasts
have been shown previously to express both
1
1 and
2
1
(25); we confirmed the presence of these two integrins on our human
dermal fibroblasts with immunohistochemistry (data not shown). Thus,
the failure of dermal fibroblasts to respond to collagen I with shape
changes similar to those observed with dermal MVECs is not attributable
to the absence of either of these two integrins.
Collagen I Induces Actin Polymerization in MVECs but Not
Fibroblasts, and Actin Polymerization Is Required for Formation of
Cords--
To investigate the effects of collagen stimulation on the
actin cytoskeleton, MVECs were stained with FITC-labeled phalloidin, which binds F-actin; and cells were examined with fluorescence microscopy. As shown in Fig.
2A, collagen I
provoked a pronounced reorganization of F-actin in MVEC monolayers, and
cellular alignment corresponded with alignment of prominent stress
fibers. In addition, these experiments suggested that collagen I
induced marked increases in F-actin. Therefore, F-actin was measured
with a quantitative fluorescence assay. As shown in Fig. 2B,
collagen I induced as much as a 300% increase in F-actin, and
induction was dependent on collagen concentration. In addition, the
marked increase in F-actin was paralleled by a corresponding decrease
in G-actin, indicating that collagen drives substantial polymerization
of the free actin pool. Collagen-induced increases in F-actin were routinely observed in numerous experiments with at least five different
isolates of dermal MVECs. Some populations were more responsive than
others, and cells at passage 5 or below were most responsive.
Regardless, in all cases collagen I stimulation increased F-actin
content by at least 100%. In contrast to collagen I and consistent
with the failure of laminin-1 to provoke changes in MVEC shape,
laminin-1 did not increase F-actin (Fig. 2B).

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Fig. 2.
Collagen-1 induces actin polymerization in
MVECs. A, FITC-phalloidin staining of the actin
cytoskeleton of MVECs indicates that collagen I provokes marked changes
in F-actin within 4 h. Top panel: control; bottom panels: 4 h after
addition of 500 µg/ml collagen I. Bars = 100 µm. As
shown in the higher power view (bottom right), actin stress
fibers align within cells forming cords (arrows).
B, quantitative assays for F-actin and free G-actin
establish that collagen I, but not laminin-1, stimulates a marked
increase in actin polymerization by 4 h, resulting in substantial
reduction in the free G-actin pool. Ab-mediated clustering of the
1 1 or 2 1
integrins, but not 6 integrins, also provokes
substantial actin polymerization in MVECs. Finally, in contrast to
dermal MVECs, dermal fibroblasts do not respond to collagen I with
detectable changes in F-actin. Error bars = S.D.
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Because collagen I ligation of integrins
1
1 and
2
1
is required for collagen-induced MVEC alignment into cord-like
structures (Fig. 1C), we also investigated the involvement
of these integrins in regulating actin polymerization. Clustering of
either the
1
1 integrin or the
2
1 integrin on MVECs with functional
integrin mAbs stimulated increases in F-actin (Fig. 2B),
thus implicating both of these integrins in mediating induction of
actin polymerization. Consistent with findings that laminin-1 did not
induce actin polymerization, mAb-mediated clustering of
6 integrins, which bind laminins (26), did not increase
F-actin.
In sharp contrast to dermal MVECs, fibroblasts did not respond to
collagen I by increasing actin polymerization (Fig. 2B, bottom panel). These findings together with findings that
collagen I does not provoke shape changes in fibroblasts (Fig.
1A) suggested the hypothesis that actin polymerization is
critical to the mechanism by which collagen provokes MVEC retraction
and re-organization into cord-like structures. Therefore, to test the
significance of actin polymerization for cord formation by MVECs, we
performed experiments with MVECs exposed to latrunculin A, a potent
inhibitor of actin polymerization (27). As shown in Fig.
3A, latrunculin A blocked
collagen-induced actin polymerization in MVECs, and it blocked
collagen-induced changes in cell shape (Fig. 3B). Thus, these experiments establish that actin polymerization is critical to
the mechanism by which collagen I initiates MVEC alignment into
cords.

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Fig. 3.
Actin polymerization is required for
collagen-induced formation of cords. A, latrunculin A
(250 nM), which interferes with actin assembly by binding
to G-actin, blocks collagen I-induced actin polymerization in MVECs
(error bars = S.D.). B, collagen I was added
to confluent MVEC cultures as in Fig. 1. Latrunculin A blocked collagen
I-induced alignment of MVECs into cords. Bar = 100 µm.
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Collagen I Provokes the Actin Polymerization Required for Cord
Formation through Suppression of cAMP--
We observed that
pharmacological elevation of intracellular cAMP consistently blocked
collagen-induced cord formation by MVECs similar to a combination of
1 and
2 integrin mAbs and similar to
latrunculin A (Fig.
4A).
cAMP-elevating agents with similar blocking effects included forskolin
(20-100 µM), which activates adenylate cyclase, and
3-isobutyl-1-methylxanthine (500 µM), which suppresses
phosphodiesterase activity. Cell-permeable analogues of cAMP (100 µM) also blocked collagen-induced changes in MVEC shape.
In particular, pharmacological elevation of cAMP with forskolin not
only blocked collagen-induced cord formation, but it also blocked
collagen-induced actin polymerization (Fig. 4B).
Collectively, these findings suggested the possibility that the
mechanism through which collagen I induces actin polymerization and
stress fibers and thereby mediates MVEC organization into cords
involves suppression of cAMP. Therefore, we measured intracellular cAMP
in MVECs stimulated with collagen I. As shown in Fig. 4B,
collagen I provoked a substantial decrease in cAMP; however, laminin-1,
which neither provoked changes in cell shape nor induction of actin
polymerization, did not suppress cAMP. Importantly, clustering of
integrins
1
1 and
2
1 with mAbs individually (Fig.
4B) and together (not shown) also produced substantial
decreases in intracellular cAMP, thus implicating both of these
collagen receptors in mediating the action of collagen I. In contrast,
mAb-clustering of
6 integrins, which are laminin receptors, did not suppress cAMP. Also, collagen I did not regulate cAMP in dermal fibroblasts (Fig. 4B). Thus fibroblasts, in
sharp contrast to MVECs, failed to respond detectably to collagen I with either suppression of cAMP, induction of actin polymerization, or
changes in cell shape.

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Fig. 4.
Collagen-induced MVEC morphogenesis is
regulated by cAMP. A, collagen I was added to confluent
MVEC cultures as in Fig. 1, and pharmacological elevation of cAMP by forskolin (100 µM) blocked collagen-induced cellular alignment into
cords. Bar = 100 µm. B, forskolin blocked
collagen-induced actin polymerization (top panel).
Furthermore, quantitative cAMP assays established that collagen I, but
not laminin-1, suppressed cAMP in MVECs and that cAMP is suppressed by
clustering of the 1 1 and
2 1 integrins but not by clustering of
6 integrins (middle panels). Collagen I did
not suppress cAMP in dermal fibroblasts, in marked contrast to MVECs.
Error bars = S.D.
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Collectively, the experiments described above established that collagen
I suppresses cAMP in MVECs, and they suggested that the mechanism
through which collagen I induces actin polymerization in MVECs involves
suppression of cAMP. To test this hypothesis directly, we employed two
synthetic antagonists of cAMP synthesis, 2',3'-dideoxyadenosine and
MDL-12,330A, to achieve suppression of cAMP and thereby imitate the
action of collagen I. Both of these compounds suppressed cAMP in MVECs,
resulting in induction of F-actin (Fig.
5A) and actin stress fibers
(Fig. 5B). Thus, these experiments established a functional
connection between suppression of cAMP by collagen I and induction of
actin polymerization in MVECs.

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Fig. 5.
Suppression of cAMP by collagen I is
functionally linked to induction of actin polymerization in MVECs.
A, adenylate cyclase inhibitors MDL 12,330A (2.5 mM) and 2',3'-dideoxyadenosine
(2',3'-DDA) (90 µM) each
suppressed cAMP in MVECs and induced F-actin within 4 h.
Error bars = S.D. B, FITC-phalloidin
staining indicated that F-actin induced by adenylate cyclase inhibitors
organized into prominent stress fibers, similar to those observed with
collagen I (Fig. 2A). Bar = 100 µm.
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Collagen I Suppression of cAMP Results in Suppression of PKA
Activity, and Suppression of PKA Induces Actin Polymerization in
MVECs--
PKA activity is dependent upon cAMP, and consistent with
our findings that collagen I suppressed cAMP (Fig. 4B),
collagen I also provoked a marked decrease in PKA activity (Fig.
6A). Therefore, we tested the
possibility that loss of PKA activity is a key consequence of cAMP
suppression through which collagen I induces actin polymerization in
MVECs. For these experiments we employed two specific inhibitors of
PKA, KT5720 and a myristoylated synthetic peptide, representing the
active portion of the natural PKA inhibitor PKI (28). Both of these
inhibitors induced actin polymerization (Fig. 6A) and the
appearance of prominent actin stress fibers (Fig. 6B). Thus, these experiments establish that collagen I stimulation of MVECs suppresses cAMP-dependent PKA activity in conjunction with
suppression of cAMP. They also establish a functional connection
between suppression of PKA activity, induction of actin polymerization,
and induction of actin stress fibers.

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Fig. 6.
Suppression of cAMP by collagen I results in
suppression of PKA in MVECs; suppression of PKA links suppression of
cAMP to induction of actin polymerization. A,
quantitative PKA assays established that collagen I suppresses
cAMP-dependent PKA activity in MVECs, consistent with
collagen suppression of cAMP. Specific PKA inhibitors KT5720 and a
myristoylated PKI peptide sequence 14-22, representing the inhibitory
domain of PKI, both induced F-actin thus establishing a functional
connection between suppression of PKA activity and induction of actin
polymerization. B, FITC-phalloidin staining indicated that
F-actin induced by PKA inhibitors organized into prominent stress
fibers, similar to those observed with collagen I (Fig. 2A)
and adenylate cyclase inhibitors (Fig. 5B).
Bar = 100 µm.
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 |
DISCUSSION |
Collagen I provokes MVECs in vitro to assume a
spindle-shaped morphology and to align into solid cord-like assemblies.
These cords, which are organized in polygonal arrays, closely imitate the polygonal patterns of embryonic pre-capillary cords that precede the formation of mature blood vessels with lumens in vivo
(1, 2, 4, 29). Experiments described here identify a previously unrecognized and critical mechanism through which collagen stimulates formation of cords. Specifically, these studies establish that collagen
I drives MVECs to assemble into cords through ligation of integrins
1
1 and
2
1,
followed by suppression of cAMP and suppression of
cAMP-dependent PKA. Suppression of
cAMP-dependent PKA induces actin polymerization and the
formation of prominent F-actin stress fibers, which are required for
cellular re-alignment into cords. Furthermore, collagen-induced actin
polymerization in MVECs coincides with large reductions in G-actin,
indicating that collagen I drives substantial polymerization of the
free actin pool. Thus, by establishing a previously unrecognized and important connection between suppression of cAMP-dependent
PKA and the induction of F-actin and MVEC alignment into cords, studies described here provide new understanding of the collagen-signaling mechanisms that regulate EC morphogenesis and multicellular organization.
Our finding that cAMP negatively regulates actin polymerization and EC
assembly into pre-capillary cords implicates cAMP as a key gatekeeper
of EC organization during angiogenesis. Importantly, cAMP has been
shown to inhibit angiogenesis in vivo (30), however, the
possibility that cAMP functions in regulating multicellular organization of ECs during angiogenesis has not been considered previously. Moreover, our findings suggest the possibility that elevation of cAMP and PKA may provide for clinical control of neovascularization by interfering with the early organizational stages
of blood vessel formation.
Previously, the
2
1 integrin alone had
been implicated in mediating collagen-induced cord formation by
umbilical vein ECs (8) The apparent discrepancy between those findings
and our observations, which also implicate integrin
1
1, is attributable to the fact that
umbilical vein ECs do not express integrin
1
1 and that both
1
1 and
2
1
are prominent collagen receptors on MVECs (14, 31). This distinction is
important, because MVECs are derived from the small capillaries of the
microvasculature, whereas umbilical vein ECs are derived from large
veins. Thus, experiments described here indicate that both
1
1 and
2
1
mediate collagen-induced morphogenesis in microvascular endothelium,
whereas such function may be limited to the
2
1 integrin in large vessel endothelium,
which does not express integrin
1
1.
Although dermal fibroblasts express both the
1
1 and
2
1
integrins, these cells did not respond to collagen I with changes in
cAMP, actin polymerization, or cell shape. These findings illustrate marked differences in collagen signaling among cell types.
Interestingly, such differences in collagen signaling may relate to the
fact that fibroblasts normally reside within a collagen-rich matrix, whereas the MVECs of mature blood vessels are sequestered from interstitial collagen by basement membrane. Moreover, we found that
laminin-1, a member of the laminin family of proteins which are major
components of basement membranes (32, 33), did not suppress cAMP or
initiate cord formation by MVECs, indicating that the responses of
MVECs to collagen and laminin are distinctly different.
Stimulation of vascular smooth muscle cells with collagen I also has
been reported to suppress intracellular cAMP (34). Pertussis toxin,
which prevents inhibition of adenylate cyclase by heterotrimeric
Gi proteins, blocked collagen suppression of cAMP in these
cells, implicating Gi proteins in the mechanism (34).
However, we found that collagen I suppressed cAMP substantially in
MVECs treated with pertussis toxin (50 ng/ml, preincubated overnight)
even though pertussis toxin elevated baseline cAMP in these cells (data
not shown). Thus, we found no evidence that collagen I suppresses cAMP
in MVECs through activation of Gi, and therefore it seems
likely that collagen I regulates cAMP in MVECs differently than in
smooth muscle cells. The possibility remains that collagen I suppresses
cAMP in MVECs through activation of phosphodiesterases. Integrin
6
4 has been shown to activate phosphodiesterase activity in carcinoma cells (35), and other integrins
may also stimulate degradation of cAMP through this mechanism.
Alternatively, collagen receptors may regulate cAMP metabolism through
other mechanisms, consistent with the complexity of integrin signaling
(36-39). Finally, data presented here raise the interesting question
of how cAMP exerts negative control over actin polymerization in MVECs.
The answer to this question may prove complex, because
cAMP-dependent PKA regulates the activities of numerous
proteins, including several that are implicated in cytoskeletal
regulation (40, 41). Regardless, experiments described here identify a
previously unrecognized regulatory role for cAMP in controlling actin
polymerization and cord formation by microvascular endothelium.
In conclusion and summary, experiments described here identify an
important mechanism through which collagen I provokes MVECs to form
solid cord-like structures. These cords closely imitate the solid
pre-capillary cords that appear during the early stages of angiogenesis
in vivo. The signaling events through which collagen induces
cord formation by MVECs begins with ligation of integrins
1
1 and
2
1,
followed by suppression of cAMP and cAMP-dependent PKA. In
turn, suppression of PKA induces actin polymerization, which is
required for EC alignment and organization into cords. In contrast to
dermal MVECs, dermal fibroblasts do not respond to collagen I with
changes in cAMP, F-actin, or cell shape, underscoring important
distinctions in collagen signaling among cell types. Although it has
long been recognized that collagen I provokes ECs to initiate a
morphogenetic program leading to the formation of pre-capillary cords,
underlying signaling mechanisms had not been defined. Collectively,
studies reported here identify a critical mechanism, and they identify
cAMP as a key regulator of EC morphogenesis and multicellular organization.