© 1999 American Heart Association, Inc.
Vascular Biology |
Angiostatin Binds to Smooth Muscle Cells in the Coronary Artery and Inhibits Smooth Muscle Cell Proliferation and Migration In Vitro
From the Section of Cardiology, Wake Forest University School of Medicine, Winston-Salem, NC.
Correspondence to David C. Sane, MD, Section of Cardiology, Wake Forest University School of Medicine, Medical Center Boulevard, Winston-Salem, NC 27157-1045. E-mail dsane{at}wfubmc.edu
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Abstract |
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Abstract—Angiostatin is an inhibitor of angiogenesis that is known to reduce endothelial cell proliferation and consequently prevent the progression of tumor metastases. However, the modest effect of angiostatin on endothelial cell proliferation raises the possibility that angiostatin might exert its effects on other cells. To determine the cellular distribution of angiostatin binding in tissues with neovasculature (atherosclerotic coronary arteries), we developed a fusion protein consisting of placental alkaline phosphatase and the first 3 kringles of plasminogen. Angiostatin binding colocalized with smooth muscle cells and could be inhibited by a 50-fold molar excess of plasminogen and 10 mmol/L
-amino-n-caproic acid. The fusion protein also bound
to smooth muscle cells in culture. Angiostatin inhibited
hepatocyte growth factor–induced proliferation and
migration of smooth muscle cells, suggesting that they are a target for
the antiangiogenic effect of angiostatin.
Key Words: angiostatin • angiogenesis • plasminogen • hepatocyte growth factor • smooth muscle cells
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Introduction |
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Angiogenesis, the elaboration of blood vessels by endothelial cell outgrowth from preexisting vessels,1 plays a vital role in embryogenesis and physiological processes such as menses formation and wound healing. The pathogenesis of a variety of diseases, including tumor growth and metastasis, diabetic retinopathy, arthritis, psoriasis, and atherosclerosis, is also influenced by either excessive or deficient angiogenesis.1 2
A variety of stimulators and inhibitors of angiogenesis have been described. One of the most novel antiangiogenesis compounds is angiostatin, a 38-kDa protein containing the first 4 of the 5 plasminogen kringles.3 Although angiostatin is derived from plasminogen, reportedly through the activity of either a serine proteinase4 or metalloelastase,5 the antiangiogenic activity of angiostatin is unique to this derivative. Angiostatin has been shown to dramatically suppress the growth and metastases of experimental tumors in animal models.6 This effect is thought to be mediated via the inhibition of endothelial cell proliferation, resulting in suppressed tumor angiogenesis. Although endothelial cell growth is a vital component of the process of angiogenesis,1 2 the near-total inhibition of tumor metastasis is paralleled by a much more modest effect on endothelial cell proliferation.4 5 This apparent discrepancy raises the possibility that angiostatin might exert some of its effects by an alternate mechanism, or that other cells, in addition to endothelial cells, could be its target. Localizing the distribution of angiostatin binding could potentially lead to a further understanding of its mechanism of action.
Because the atherosclerotic coronary artery is a site of intense neovascularization,7 we examined the cellular distribution of angiostatin binding to coronary artery sections. In this study, we report that the smooth muscle cell (SMC) is the principal binding site in the coronary artery for binding angiostatin and further demonstrate that angiostatin inhibits SMC proliferation and migration. Because there is evidence that SMCs participate in angiogenesis,8 these affects could contribute to the antiangiogenic effect of angiostatin.
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Methods |
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Materials
The full-length plasminogen cDNA and the APtag-4 vector were generous gifts from L.O. Heden9 (Department of Microbiology, University of Lund, Lund, Sweden) and J. Flanagan,10 (Department of Cell Biology, Harvard Medical School, Boston, Mass), respectively. Competent Escherichia coli cells (MAX efficiency DH10B/P3) were purchased from GIBCO/BRL. The restriction enzyme BglII was purchased from Promega. Ampicillin and tetracycline were purchased from Sigma Chemical Co. COS-1 cells were obtained from the American Type Culture Collection (ATCC, Manassas, Va). Opti-MEMI and LipofectAMINE were purchased from GIBCO/BRL. All cell culture reagents, including Dulbecco's modified Eagle's medium (DMEM), FCS, the serum-free medium AIMV, L-glutamine, penicillin/streptomycin (pen/strep), and trypsin/EDTA were purchased from GIBCO/BRL. Placental alkaline phosphatase (PAP) activity was detected by using either the soluble substrate p-nitrophenyl phosphate with diethanolamine buffer from Bio-Rad or, for in situ analyses, the ProtoBlot NBT (nitro blue tetrazolium) and BCIP (5-bromo-4-chloro-3-indolyl phosphate) color development system (Promega). L-Homoarginine, an inhibitor of cellular phosphatases but not of PAP, was purchased from Sigma. Centricon-30 concentrators were purchased from Amicon. The polyclonal antibody to plasminogen was purchased from Calbiochem, and the monoclonal antibody to PAP was purchased from Genzyme Diagnostics. A monoclonal antibody to smooth muscle actin (HHF-35) was purchased from ENZO Diagnostics, Inc. A monoclonal antibody to the endothelial cell antigen CD31 was purchased from Dako Corp. The enhanced chemiluminescence kit, which contained peroxidase-conjugated anti-rabbit IgG and peroxidase-conjugated anti-mouse IgG, was purchased from Amersham.
-Amino-n-caproic acid (EACA), hepatocyte
growth factor (HGF), and BSA were purchased from Sigma. Rb-1 cells
(rabbit aortic SMCs) were a gift from M. Nachtigal11
(Department of Pathology, University of South Carolina School of
Medicine, Columbia), and human aortic SMCs (CRL 1999) were obtained
from ATCC. Purified angiostatin, used in the proliferation and
migration assays, was purchased from Angiogenesis Research Industries.
The Transwell apparatus was purchased from Corning
Costar Corp, and staining was performed with Diff-Quick from Dade
International. Oligodeoxynucleotide primers were
synthesized on an ABI 394 DNA/RNA synthesizer by a standard cyanoethyl
phosphoramidite procedure in the DNA Synthesis Core Laboratory of the
Comprehensive Cancer Center of Wake Forest University.
Development of Fusion Protein Containing PAP and
Plasminogen Kringles 1 through 3 (PAP/K1-3)
The fusion vector was constructed by amplifying the human
plasminogen cDNA using the polymerase chain reaction (PCR)
primers listed as follows: sense, 5'-GA AGA TCT
(BglII) GGC (Gly) GGC (Gly) GTG TAT
CTC TCA GAG TGC-3'; antisense, 3'-GG TGC CTT GTT AAC CGA ACT
(stop) TCT AGA (BglII) AG-5'.
These primers create 5' and 3' BglII sites, Gly-Gly linkers between PAP and kringle 3, and amplify the first 3 kringles of plasminogen (residues Lys78 to Ala344). The 3-kringle form of angiostatin has been found to be more potent than the 4-kringle derivative form.12 PCR amplification was performed using a denaturing temperature of 94°C for 1 minute, annealing temperature of 60°C for 1 minute, and extending temperature of 72°C for 2 minutes. The PCR product was cloned into the BglII site of APtag-4.10 The recombinant vector (PAP/K1-3) was then subcloned into MAX efficiency DH10B/P3 competent cells and selected with ampicillin and tetracycline. The recombinant plasmid (PAP/K1-3) was digested with the BglII restriction enzyme to verify the correct size of the insert and sequenced on both strands by using an ABI Prism automated sequencer (with primers derived from APtag-4 and the plasminogen sequence) to verify construction and orientation and to eliminate the possibility of PCR-induced errors.
Expression of PAP/K1-3 in COS-1 Cells
Transient expression of PAP/K1-3, as well as the wild-type
vector APtag-4, was achieved by transfecting COS-1 cells that were
grown in DMEM with 10% FCS and pen/strep. The cells were transfected
in Opti-MEMI at 80% confluence with 2 µg of PAP/K1-3 or APtag-4 and
8 µL of LipofectAMINE. After 1 hour, fresh Opti-MEMI plus 10% FCS
was added for 4 hours, followed by fresh DMEM, FCS, and pen/strep.
After 24 hours, the medium was changed to AIMV serum-free medium with
BSA. The medium from transfected COS-1 cells was harvested at 4 days
and assayed for PAP activity.
PAP Assay
A sample of the harvested medium (10 µL) was heated to 65°C
for 10 minutes to eliminate background alkaline phosphatase activity
(PAP is resistant to heat inactivation). The
OD405 was measured after incubating the heated
sample for 1 hour at room temperature in 1 mol/L diethanolamine (pH
9.8), 0.5 mmol/L MgCl2, 10 mmol/L
L-homoarginine (a cellular phosphatase
inhibitor that does not inhibit PAP), 0.5 mg/mL BSA, and
12 mmol/L p-nitrophenyl phosphate.
Analysis of Secreted Protein
The medium from the transfections was concentrated 10-fold by
using a Centricon-30 concentrator. A 10% SDS–polyacrylamide
gel electrophoresis (PAGE) of the concentrated media was performed,
followed by transfer of the proteins to nitrocellulose and blocking
with 3% nonfat milk in 1x Tris-buffered saline, pH 7.4, and 0.1%
Tween-20. A polyclonal antibody to plasminogen (1:500) was
used, followed by a peroxidase-conjugated anti-rabbit IgG (1:5000). For
detecting PAP, a monoclonal antibody to human PAP (1:2000) was used,
followed by a peroxidase-conjugated anti-mouse IgG (1:5000).
Chemiluminescence detection was performed with an
enhanced chemiluminescence kit.
In Situ Binding Assays
The medium from the transfections was concentrated so that it
had equal alkaline phosphatase activity (OD405
APtag-4=1.37, OD405 PAP/K1-3=1.32). The
unfused-PAP medium was used as a control for nonspecific binding and
for background alkaline phosphatase activity that was not destroyed by
heating.
The in situ binding analysis of the fusion protein to the frozen coronary arteries was performed by following the method of Flanagan and Leder.10 OCT-embedded frozen sections were thawed briefly at room temperature and then immediately washed with HBHA buffer (Hanks' balanced salt solution with 0.5 mg/mL BSA, 0.1% NaN3, and 20 mmol/L HEPES, pH 7.0). The slides were then incubated with the 10x-concentrated PAP/K1-3 protein or unfused PAP, with or without a 50-fold molar excess of plasminogen or 10 mmol/L EACA, for 90 minutes. The slides were rinsed 7 times with HBHA buffer and fixed for 30 seconds with 60% acetone, 3% formaldehyde, and 20 mmol/L HEPES, pH 7.5. The slides were then washed in 150 mmol/L NaCl, 20 mmol/L HEPES (pH 7.5) twice for 5 minutes each and floated on a 65°C water bath for 60 minutes to inactivate cellular phosphatases. The slides were then rinsed with 100 mmol/L Tris-HCl, pH 9.5, 100 mmol/L NaCl, and 5 mmol/L MgCl2. To stain the slides, the ProtoBlot NBT and BCIP color development system containing 0.17 mg/mL BCIP and 0.33 mg/mL NBT (in 100 mmol/L Tris-HCl, pH 9.5; 100 mmol/L NaCl; and 5 mmol/L MgCl2) was added to each slide for up to 72 hours.
Conditions for SMCs in Culture
Rabbit aortic (Rb-1) SMCs were grown in DMEM supplemented with
L-glutamine and 10% FCS. Human aortic SMCs (CRL 1999) were
cultured in Ham's F12K supplemented with 2 mmol/L glutamine,
10 mmol/L HEPES, 10 mmol/L TES, 50 µg/mL ascorbic acid, 10
µg/mL insulin, 10 µg/mL transferrin, and 10 ng/mL
Na2SeO3 with 10% FCS. The
cells were cultured in 75-cm2 plastic
tissue-culture flasks and incubated at 37°C in a humidified
atmosphere of 5% CO2 in air. The cells were
subcultured at a 1:4 split ratio at confluence with the use of 0.02%
trypsin/EDTA.
Binding of PAP/K1-3 to Human Aortic SMCs
Human aortic SMCs were grown to confluence in 96-well plates.
The 10x-concentrated medium from the transfection of PAP/K1-3 or
APtag-4 was incubated at the concentrations shown for 90 minutes at
room temperature. Unbound material was rinsed with HBHA, and
endogenous alkaline phosphatase activity was
heat-inactivated as described for the in situ binding
assay. Alkaline phosphatase activity was detected using
p-nitrophenyl phosphate, and the OD405
was measured in an ELISA plate reader.
Proliferation Assays
Rb-1 cells were seeded at 1000 cells per well in a 96-well plate
and grown to 10% to 20% confluence. Cells were incubated for 24 hours
in serum-free DMEM to allow synchronization of the cell cycle.
Preliminary experiments established optimum proliferation conditions as
DMEM with 10% FCS and 20 ng/mL HGF for a 96-hour incubation period.
The cells were therefore incubated in DMEM plus 10% FCS and HGF (20
ng/mL), with or without varying concentrations of angiostatin (0 to
2.0 µmol/L) or plasminogen (2.0 µmol/L), for
96 hours. The cells were harvested by trypsinization, followed by
counting with a Coulter counter. The results were compared with the
proliferation observed in the presence of DMEM and 10% FCS, which was
defined as baseline (100%). An additional cell proliferation assay
(Cell Titer 96 AQ, Promega) was performed by using the NBT compound
that is reduced by living cells to a colored formazan product that
can be detected at 490 nm. Each experiment was performed 6 times. The
effect of angiostatin on apoptosis was determined using an in
situ cell death detection kit (AP, Boehringer Mannheim;
essentially a terminal deoxynucleotidyl
transferase–mediated dUTP-biotin nick end-labeling assay).
Migration Assays
Migration assays were performed as previously
described13 14 15 using a Transwell
apparatus with an 8-µm pore size polycarbonate membrane
separating the 2 chambers. Trypsinized cells were pelleted by
centrifugation at 1000 rpm for 5 minutes and
resuspended in fresh medium to eliminate the trypsin. Preliminary
experiments established optimum migration conditions as a combination
of DMEM, 10% FCS, and 50 ng/mL HGF. The DMEM/10% FCS/HGF (50 ng/mL)
mixture, with or without angiostatin (0 to 2.0 µmol/L) or
plasminogen (2 µmol/L), was added to the lower
chamber while 5x104 cells in DMEM/10% FCS were
added to the upper chamber. Migration was allowed to proceed for 6
hours, a time frame in which preliminary experiments had demonstrated
that there was no significant effect on cell proliferation. After the
cells adherent to the upper side of the membrane were scraped away, the
cells on the lower side were fixed in 10% neutral buffered formalin
overnight and stained with Diff-Quick. Three random, nonoverlapping
high-powered (200x) fields (HPFs) were manually counted, and the mean
of 6 HPFs was analyzed. The results were compared with the
number of cells that migrated in the presence of DMEM/10% FCS, which
was defined as baseline (100%).
Calculations
The data are expressed as mean±SEM. Differences were compared
using the 2-tailed Student's t test for paired
samples.
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Results |
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Analysis of the Fusion Protein PAP/K1-3
PAP activity was measured from the medium of COS-1 cells transfected with the APtag-4 and PAP/K1-3 vectors by using p-nitrophenyl phosphate substrate. At 4 days after transfection, there was
5 µg/mL of recombinant protein expressed
in the serum-free medium.
To verify the construction of the secreted fusion protein, an
immunoblot analysis was performed. Detection of the
97-kDa fusion protein with both anti-plasminogen (Figure 1A
) and anti-PAP (Figure 1B)
verified the structure of our fusion protein containing both PAP and
the first 3 kringles of plasminogen. As expected, the
medium from APtag-4 transfections did not have a visible band with
anti-plasminogen but did have a 67-kDa protein that was
detected by anti-PAP, which represented nonfused PAP (not
shown). Construction of PAP/K1-3 was also verified by sequencing both
strands of the plasmid construct.
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PAP/K1-3 Binds to Endothelial Cells and SMCs in the
Coronary Artery
In situ binding assays of the PAP/K1-3 fusion protein to human
coronary arteries localized angiostatin binding sites to both
the intimal and medial regions (Figure 2B), whereas the protein obtained from
the transfection of the vector APtag-4 showed minimal binding (Figure 2A).
When compared with endothelial cells
(anti-CD31, Figure 2C) and SMCs (anti–
-actin, Figure 2D)
detected by immunohistochemistry, it is apparent that most
of the binding of the fusion protein occurred in a distribution
mirroring that of SMCs and was not restricted to
endothelial cells. Similar binding patterns of the
fusion protein were observed in arterioles and capillaries of the
adventitia (Figures 3 and 4).
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The binding of the fusion protein was nearly completely blocked with
10 mmol/L EACA (not shown), as well as with a 50-fold molar excess
of plasminogen (Figure 5).
These results indicate that angiostatin binds to a
plasminogen binding site and that the interaction is
mediated via lysine binding sites.
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In situ binding analysis performed on atherectomy tissue also
revealed binding of the fusion protein to SMCs, as identified by the
anti–-actin pattern (Figure 6A and 6B), and was inhibited by excess plasminogen (Figure 6C).
The fusion protein did not bind to cellular components
other than SMCs, as shown in the hematoxylin-and-eosin–stained
sections (Figure 6D). Binding of concentrated medium from
APtag-4 transfections showed that there was no background cellular
phosphatase activity (Figure 6E).
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PAP/K1-3 Binds to Human Aortic SMCs
The binding of various concentrations of the fusion protein
PAP/K1-3 to human aortic SMCs in vitro was determined. Figure 7 shows a concentration-dependent
increase in fusion protein binding, whereas binding of PAP alone was
minimal.
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Effect of Angiostatin on the Proliferation and Migration of Rabbit
Aortic (Rb-1) SMCs
Rb-1 cells were exposed to various concentrations of HGF, with or
without FCS, for 24, 48, 72, or 96 hours. From these preliminary
experiments, it was found that the most significant induction of
proliferation occurred at a concentration of 20 ng/mL HGF, in the
presence of FCS, at 96 hours (not shown). Similarly, preliminary
experiments demonstrated that a concentration of 50 ng/mL HGF produced
a maximal effect on Rb-1 cell migration. These cells were allowed to
migrate for 6 hours, a time frame in which preliminary experiments had
demonstrated there was no significant cell proliferation.
The effect of angiostatin (Angiogenesis Research Industries; 0.25 to
2.0 µmol/L) and plasminogen (2.0 µmol/L) on
HGF-induced cell proliferation was determined (Figure 8). HGF (20 ng/mL) induced a 41%
increase in cell number at 96 hours (P<0.01). Angiostatin
at 2.0 µmol/L significantly inhibited HGF-induced Rb-1 cell
proliferation (P<0.01). Plasminogen (2.0
µmol/L) had no effect on HGF-induced Rb-1 cell proliferation. In the
absence of HGF, angiostatin at 2.0 µmol/L produced a 37%
inhibition of proliferation (compared with DMEM/10% FCS alone,
P<0.05; data not shown). The inhibitory effect
of angiostatin on cell proliferation was confirmed by using the
reduction of NBT (data not shown). No effect of angiostatin (2.0
µmol/L) on Rb-1 cell apoptosis was observed (data not
shown).
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Similarly, Figure 9 demonstrates the
effect of angiostatin on HGF-induced Rb-1 cell migration. HGF (50
ng/mL) increased SMC migration by 239% (P<0.01).
Angiostatin significantly inhibited HGF-induced migration at a
concentration of 0.5 µmol/L (P<0.05), with maximal
inhibition at 2.0 µmol/L (P<0.01).
Plasminogen (2.0 µmol/L) had no effect on
HGF-induced migration. Angiostatin had no significant effect on
migration that occurred in the absence of HGF (data not shown).
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Discussion |
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Our study demonstrates that angiostatin, a novel inhibitor of angiogenesis, binds to SMCs and endothelial cells in the coronary artery. The binding to endothelial cells is consistent with the previous observation that the antiangiogenic effect of angiostatin is at least partially mediated through its inhibition of endothelial cell proliferation.6 This relatively modest effect (35% at 10 µg/mL),4 however, does not appear adequate to account for the dramatic effect that this agent has on inhibiting tumor growth and metastases. This apparent discrepancy led us to consider other potential cellular targets and mechanisms for the effect of angiostatin. The demonstration that the angiostatin fusion protein binds to SMCs and that angiostatin inhibits the HGF-induced proliferation and migration of SMCs may lead to an appreciation of novel cellular targets and molecular mechanisms for the antiangiogenic effect of angiostatin.
The role of SMCs in vasculogenesis, the embryological development of blood vessels from progenitor cells,16 has been defined in mice. The recruitment of SMCs to the wall of developing blood vessels is mediated by angiopoietin-1 binding to the TIE2 receptor17 and inhibited by angiopoietin-2.18 In contrast, the contribution of SMCs to angiogenesis, or the sprouting of new vessels from an existing network, is less clearly established. One potentially important role for SMCs in this process could be the elaboration of angiogenic factors that act in a paracrine manner to promote endothelial cell proliferation. SMCs are known to secrete a variety of direct and indirect angiogenesis factors, including HGF,19 vascular endothelial growth factor (VEGF),20 and transforming growth factor-ß1 (TGF-ß1).21 It has been suggested that pericytes and myofibroblasts, cells that play a supporting role in angiogenesis,22 23 may be derived from a population of SMCs.24 25 Although there is no direct evidence that SMCs are necessary for the formation of capillaries in the early stages of angiogenesis, the histological detection of proliferating SMCs in vessels elicited from a variety of angiogenic stimuli26 27 28 suggest that the formation of larger, mature, conduit-caliber vessels requires the participation of SMCs. This finding also supports the possibility that SMCs could be a target for antiangiogenic therapy. In addition to the present study showing that angiostatin inhibits SMC proliferation and migration, octreotide29 and TNP-470,30 known angiogenesis inhibitors, have both been demonstrated to have an antiproliferative effect on SMCs.
The inhibition of endothelial cell proliferation by
angiostatin, but not plasminogen,3 suggests
that there could be a specific angiostatin receptor on
endothelial cells. Although preliminary studies are
consistent with a specific, saturable
endothelial receptor,31 this putative
receptor has yet to be identified [note added in proof: Moser et al
(Proc Natl Acad Sci U S A. 1999;96:2811–2816) have
shown that angiostatin binds to the /ß ATP synthase in HUVECS].
Our studies suggest that angiostatin binds to a plasminogen
receptor on SMCs, because the binding of the fusion protein could be
inhibited by excess plasminogen and EACA.32
The effective concentration of angiostatin in our studies was 1 to
2 µmol/L, which is similar to its concentration in
plasma.33 Even higher levels could be achieved during
therapeutic administration of angiostatin. However, it should be noted
that our studies do not exclude the possibility of additional binding
interactions with other types of receptors or even with extracellular
matrix proteins.
By occupying plasminogen receptor(s), such as annexin
II34 and -enolase,35 on the SMC surface,
angiostatin might block the binding of plasminogen and its
subsequent activation to plasmin. Plasmin is an important proteinase in
angiogenesis by virtue of its ability to directly degrade matrix
components36 and by its ability to activate matrix
metalloproteinases.37 38 Plasmin releases proteolytically
processed soluble products of VEGF-165 and -189 isoforms that
retain mitogenic and vascular permeability-enhancing
activity.39 Plasmin appears to cleave these forms of VEGF
at the carboxy termini, thereby eliminating the heparin binding sites
that are responsible for their retention to heparan sulfate
proteoglycan reservoirs in the extracellular matrix or on plasma
membranes. Plasmin can also release other direct or indirect modulators
of angiogenesis from the extracellular matrix, such as basic fibroblast
growth factor,40 or from cell surface receptors, like
TGF-ß1.41 In addition to releasing angiogenic growth
factors from the extracellular matrix, plasmin also proteolytically
activates 2 precursor forms of growth factors to the active
state, HGF42 and TGF-ß1.43 As an example of
the reciprocal cooperation between growth factors and plasmin formation
in angiogenesis, VEGF,44 basic fibroblast growth
factor,45 and angiogenin 46 induce
plasminogen activator expression and enhance
pericellular plasmin generation by endothelial cells.
Plasmin generation appears necessary for SMC
migration,47 48 49 50 and therefore, the lack of pericellular
plasmin activity could limit the ability of SMCs to participate in
angiogenesis.
It is also possible that angiostatin inhibits both endothelial cell and SMC proliferation and migration by mechanisms that are independent from its ability to displace binding to plasminogen receptors. One potential mechanism would involve binding to c-met, the receptor for HGF, which is known to be expressed on both endothelial cells and SMCs.19 This theory is attractive because HGF is a potent angiogenic factor, inducing endothelial cells and SMCs to proliferate and migrate.51 52 53 Furthermore, HGF is strikingly similar to plasminogen, with 38% identity at the amino acid level and conservation of the kringle motifs, as well as the presence of a nonfunctional serine proteinase–like domain.54 In addition, the internal cleavage site that gives rise to the disulfide-linked, 2-chain, active form of HGF is identical to that of plasminogen.51 Thus, it is possible that angiostatin displaces the structurally homologous angiogenic factor HGF from binding to its cell surface receptor c-met on both endothelial cells and SMCs. The finding in the present study that angiostatin inhibits HGF-induced SMC proliferation and migration is consistent with this theory. Because HGF and c-met are expressed in vascular tissue19 where they interact in an autocrine and paracrine fashion,51 angiostatin might inhibit the proliferation of SMCs in response to endogenous vascular levels of HGF. Indeed, in the present study, there was a trend toward angiostatin-induced inhibition of SMC proliferation and migration, even in the absence of exogenously added HGF.
In summary, these data demonstrate that angiostatin binds to SMCs in the coronary artery. Furthermore, angiostatin inhibits the effects of HGF, a stimulant of angiogenesis, on SMC proliferation and migration in vitro. These studies suggest that the SMC should be considered a potential cellular target for the effects of angiostatin and other inhibitors of angiogenesis.
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Acknowledgments |
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The DNA Core Synthesis Laboratory of the Comprehensive Cancer Center of Wake Forest University was supported in part by National Institutes of Health ([NIH], Bethesda, Md) grant CA12197. This work was supported by a Wake Forest University Comprehensive Cancer Center pilot study and by NIH grants CA81233 and R21DK55061 (to D.C.S.). J.J.W. was supported in part by NIH training grant HL07868 "Pathobiology of Vascular Disease."
Received May 7, 1998; accepted January 26, 1999.
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