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Cancer Research 68, 4736, June 15, 2008. doi: 10.1158/0008-5472.CAN-07-6612
© 2008 American Association for Cancer Research
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Tumor Microenvironment

Tissue Inhibitor of Metalloproteinase 3 Suppresses Tumor Angiogenesis in Matrix Metalloproteinase 2–Down-regulated Lung Cancer

Chandramu Chetty1, Sajani S. Lakka1, Praveen Bhoopathi1, Sateesh Kunigal1, Roger Geiss2 and Jasti S. Rao1,3

1 Program of Cancer Biology, Department of Cancer Biology and Pharmacology, and Departments of 2 Pathology and 3 Neurosurgery, University of Illinois College of Medicine at Peoria, Peoria, Illinois

Requests for reprints: Jasti S. Rao, Department of Cancer Biology and Pharmacology, University of Illinois College of Medicine at Peoria, One Illini Drive, Peoria, IL 61605. Phone: 309-671-3445; Fax: 309-671-3442; E-mail: jsrao{at}uic.edu.


   Abstract
Top
 Abstract
Introduction
 Materials and Methods
 Results
 Discussion
 Disclosure of Potential...
 References
 
Matrix metalloproteinase-2 (MMP-2) expression is often up-regulated in advanced cancers and known to play an important role in tumor angiogenesis. We previously showed that adenoviral-mediated delivery of siRNA for MMP-2 (Ad-MMP-2-Si) inhibited lung cancer growth, angiogenesis, and metastasis. In this study, we investigated the signaling mechanisms involved in Ad-MMP-2-Si–mediated inhibition of angiogenesis. Ad-MMP-2-Si treatment inhibited neovascularization in vivo as determined by mouse dorsal air sac model, and conditioned medium from Ad-MMP-2-Si–infected A549 lung cancer cells (Ad-MMP-2-Si-CM) inhibited endothelial tube formation in vitro. Ad-MMP-2-Si-CM decreased proliferation as determined by Ki-67 immunofluorescence and induced apoptosis in endothelial cells as determined by terminal deoxynucleotidyl-transferase–mediated dUTP nick-end labeling (TUNEL) assay. Furthermore, Ad-MMP-2-Si-CM inhibited AKT phosphorylation and induced phosphorylation of extracellular signal-regulated kinase (ERK)/mitogen-activated protein kinase in endothelial cells. Overexpression of constitutively active AKT reversed the Ad-MMP-2-Si-CM–mediated inhibition of tube formation and induction of ERK phosphorylation. Conversely, Ad-MMP-2-Si-CM induced tissue inhibitor of metalloproteinase (TIMP) 3 expression, and the interaction of vascular endothelial growth factor 2 and TIMP-3 was determined by coimmunoprecipitation experiments. TIMP-3 induction was mediated by ERK activation. In addition, electrophoretic mobility shift and chromatin immunoprecipitation assays show that Sp1 transcription factor mediated Ad-MMP-2-Si-CM–stimulated increase of TIMP-3. Vasculature destruction was confirmed with colocalization studies with TUNEL and an endothelial marker, CD31, in tumor sections of Ad-MMP-2-Si–treated mice. Our data collectively suggest that MMP-2 inhibition induces endothelial apoptosis in vivo and inhibits endothelial tube formation. These experiments provide the first evidence that inhibition of p-AKT and induction of p-ERK1/2 are crucial events in the induction of TIMP-3–mediated endothelial apoptosis in MMP-2 inhibited lung tumors. [Cancer Res 2008;68(12):4736–45]


   Introduction
Top
 Abstract
 Introduction
Materials and Methods
 Results
 Discussion
 Disclosure of Potential...
 References
 
The initiation and development of adequate neovascularization from existing blood vessels to furnish nutrients and oxygen to tumor cells is crucial for tumor growth. Angiogenesis, a process of the origination of functional microcapillaries from existing vasculature, plays a pivotal role in tumor survival, growth, and metastasis (1). The angiogenic switch, a discrete step in tumor development, implies a shift in equilibrium between angiogenesis inducers and countervailing inhibitors in the local environment of the tumor (2). A common line of attack for shifting the balance involves altered gene transcription of positive and negative regulators of angiogenesis. Among these, vascular endothelial growth factor (VEGF) is the most potent and specific of the endothelial cell mitogens, and acts as both an endothelial cell survival factor and a key factor in mobilizing circulating endothelial cell precursors to nascent blood vessels (3–6). Not only does VEGF promote the vascularization and growth of the primary tumor, but it also seems to play a key role in the early stages of establishing new metastatic foci (5). Malignant pleural effusions from lung cancer have high levels of VEGF, and VEGF secretion has also been associated with the development of ascites (7). Moreover, many tumors including lung tumors evidence increased expression of VEGF compared with their normal tissue counterparts (8, 9). VEGF receptors are also expressed on lung cancer cells and mediate antiapoptotic and metastatic signals (10).

Matrix metalloproteinases (MMP), a family of extracellular proteolytic enzymes, and their tissue inhibitors (TIMP) are known to play significant roles in tumor invasion, angiogenesis, and metastasis (11). The possible roles of MMP-2 and MMP-9 (also known as gelatinases) in tumor angiogenesis have been explored using different in vivo and in vitro experimental models. Human umbilical vein endothelial cells (HUVEC) undergoing tubulogenesis within a three-dimensional fibrin matrix have exhibited increased expression of several MMPs (MMP-2, MT1-MMP, MT2-MMP, and MT3-MMP) compared with HUVECs growing in monolayer cultures (12). MMP-2 was expressed mostly by the microvascular cells of the blood vessels within and surrounding the tumor and by fibroblasts adjacent to the tumor stroma, whereas MMP-9 was expressed in the tissue macrophages in the vicinity of tumor nodules (13, 14). MMP inhibition studies support the notion that angiogenesis is dependent at least in part on the actions of MMPs because both TIMPs and synthetic MMP inhibitors, such as BB94 (Batimastat) and BMS-275291, display antiangiogenic properties in vitro and in vivo (15–17). The antiangiogenic signaling capacity of TIMPs was shown to be distinct from their MMP inhibitory activity, and more specifically, the ability of TIMP-3 to interact with VEGFR2, competed with VEGF for binding to its receptor (18). MMP-2–deficient mice displayed reduced tumor-induced angiogenesis as measured by the dorsal air sac assay (19). In a Swarm rat chondrosarcoma model, in which MMP-2 activation was associated with the angiogenic phenotype, suppression of MMP-2 activity by antisense oligonucleotides inhibited angiogenic potential and tumor growth (20). However, the mechanisms underlying decreased angiogenesis via MMP-2 inhibition remain largely unknown.

We have previously shown that down-regulation of MMP-2 by adenovirus-mediated delivery of MMP-2 siRNA (Ad-MMP-2-Si) reduced spheroid invasion and angiogenesis in vitro and metastasis and tumor growth in vivo (21). Hence, we hypothesized that MMP-2 inhibition in the tumor cells causes a shift in the balance between positive and negative regulators of neovascularization, thereby resulting in the inhibition of tumor-induced angiogenesis and subsequent inhibition of tumor growth. These studies, as detailed below, document that MMP-2 inhibition in lung cancer cells resulted in decreased induction of VEGF in endothelial cells and the subsequent cascade of signaling mechanisms that culminated in TIMP-3 induction leading to inhibition of angiogenesis and endothelial apoptosis.


   Materials and Methods
Top
 Abstract
 Introduction
 Materials and Methods
Results
 Discussion
 Disclosure of Potential...
 References
 
Cells and reagents. A549 cells were cultured on RPMI 1640 (American Type Culture Collection) supplemented with 10% fetal bovine serum, 50 units/mL penicillin, and 50 µg/mL streptomycin (Life Technologies, Inc.). In addition, glutamine, epidermal growth factor, and hydrocortisone (Stem Cell Technologies) were added to advanced MEM medium (Invitrogen) for human dermal microvascular endothelial cells (HMEC). Cells were incubated in a humidified 5% CO2 atmosphere at 37°C. We used antibodies specific for VEGF, VEGFR2, phospho-VEGFR2, extracellular signal-regulated kinase (ERK), phospho-ERK, Ki-67, Sp1 (Santa Cruz Biotechnology), phosphatidylinositol-3-OH kinase (PI3K), AKT, phospho-AKT (Cell Signal), glyceraldehyde-3-phosphate dehydrogenase (GAPDH; Novus Biologicals, Inc.), TIMP-3, CD31, and horseradish peroxidase (HRP)-conjugated secondary antibodies (Biomeda). Chromatin Immunoprecipitation (ChIP) Assay kit (Upstate Biotechnology), diffusion chambers (Fisher Scientific), constitutively expressed AKT (myr-AKT; Addgene, Plasmid 10841), Protein A/G PLUS-Agarose, TIMP-3 siRNA, and control siRNA oligos (Santa Cruz Biotechnology) were used in this study.

Adenoviral siRNA constructs and infection. Adenoviral siRNA for MMP-2 (Ad-MMP-2-Si) and scrambled vector (Ad-SV) were constructed and amplified as described by us previously (21). Viral titers were quantified as plaque-forming unit (pfu) per milliliter after infection of 293 cells. Titers were obtained for the viruses used in this work are Ad-SV (7.6 1011 pfu/mL) and Ad-MMP-2-Si (5.0 1011). The specificity of this virus to inhibit MMP-2 was documented in our previous studies (21, 22). The amount of infective adenoviral vector per cell (pfu per cell) in culture medium was expressed as multiplicity of infection (MOI). Virus constructs were diluted in serum-free culture medium to the desired concentration, added to cells, and incubated at 37°C for 1 h. Then, the necessary amount of complete medium was added and cells were incubated.

Preparation of tumor-conditioned medium. A549 cells (1.5 x 106) were seeded in 100-mm Petri dishes and left overnight. Cells were infected with mock (PBS), 100 MOI of either Ad-SV (adenovirus carrying a scrambled sequence) or Ad-MMP-2-Si (adenovirus carrying siRNA against MMP-2) and incubated for a further 24 h. The medium was replaced with serum-free DMEM/F-12 50/50 medium and incubated for 16 h. This conditioned medium (CM) was removed, centrifuged for 5 min, and used to grow endothelial cells. CM collected from mock, Ad-SV, and Ad-MMP-2-Si–infected A549 cells were designated as mock-CM, Ad-SV-CM, and Ad-MMP-2-Si-CM, respectively. All the experiments were performed in the presence of serum-free endothelial culture medium to see if CM by itself had any effect on endothelial cells.

In vitro angiogenic assay. Angiogenesis was performed as described earlier (23) with little modification. HMECs (2 x 104 cells per well) were grown in the presence of tumor-conditioned medium (TCM) in 96-well plates coated with Matrigel and incubated for 16 h at 37°C. The formation of capillary-like structures was captured. The degree of angiogenesis was measured as the number of branch points per view and tube length of branches was counted.

Mouse dorsal air sac model. Athymic nude mice were maintained within a specific-pathogen, germ-free environment. The implantation technique of the dorsal skin fold chamber model has been described previously (24). Briefly, diffusion chambers with mock or Ad-SV or Ad-MMP-2-Si–infected A549 cells (2 x 106) were placed underneath the skin into the superficial incision made horizontally along the edge of the dorsal air sac. After 10 d, the mice were carefully skinned around the implanted chambers and the skin fold covering the chambers was photographed under a visible light microscope. The number of blood vessels within the chamber in the area of the air sac fascia was counted and quantitated.

Immunocytochemical analysis for Ki-67 index. HMECs (5 x 103 cells per well) seeded in 8-well chamber slides were allowed to grow for 72 h on A549-CM. The effects of the TCM on HMEC cellular proliferation were measured by analysis for Ki-67 immunoreactivity. Cells were fixed in cold methanol and permeabilized in 0.1% Triton X-100 in PBS. After blocking with 1% bovine serum albumin in PBS for 1 h at room temperature, cells were incubated overnight with anti–Ki-67 (1:100 dilution). Mouse IgG was used as a negative control. After incubation with HRP-conjugated secondary antibody (1:200 dilution) for 1 h, 3,3-diaminobenzidine solution (Sigma) was used for developing chromogen and counterstained with hematoxylin and mounted. The bright field images were captured with an Olympus BX-60 research fluorescence microscope attached to a CCD camera.

Terminal deoxy nucleotidyl transferase–mediated nick labeling assay and immunohistochemistry. We detected apoptosis in HMECs cultured in TCM as well as the s.c. tumor tissue sections of Ad-MMP-2-Si–treated mice using terminal deoxynucleotidyl transferase–mediated dUTP nick-end labeling (TUNEL) enzyme reagent as per the manufacturer's instructions. Briefly, 5x103 HMECs were cultured in TCM for the indicated time periods (fresh TCM was added every 24 h), fixed in 4% paraformaldehyde in PBS for 1 h at room temperature, and permeabilized in 0.1% Triton-X100 in 0.1% sodium citrate in PBS for 2 min (for cells) or 10 min (for tissue sections) on ice. The samples were incubated in TUNEL reaction mixture in a humidified atmosphere at 37°C for 1 h in the dark. For tissue sections, after TUNEL, immunohistochemical analysis was performed using anti-CD31 antibody as described previously (22). Images were captured with an Olympus BX 60 research fluorescence microscope attached to a charge-coupled device (CCD) camera, and cells were counted. The apoptotic index was defined as follows: apoptotic index (%) = 100 x (apoptotic cells/total cells).

Extraction of extracellular matrix protein. After 16 h in the presence of TCM, HMEC cells were washed and extracellular matrix (ECM) was extracted as described previously (22). The extracts were subjected to immunoblotting.

Immunoprecipitation and immunoblotting. HMECs were grown in TCM for 16 h. Then, cells were collected, and whole cell lysates were prepared by lysing cells in radioimmunoprecipitation assay lysis buffer containing 1 mmol/L sodium orthovanadate, 0.5 mmol/L phenylmethylsulfonyl fluoride (PMSF), 10 µg/mL aprotinin, and 10 µg/mL leupeptin. Equal amounts of protein fractions or immunoprecipitates of lysates with the indicated antibodies were resolved by SDS-PAGE and transferred to polyvinylidene difluoride membrane. Proteins were detected with primary antibodies followed by HRP-conjugated secondary antibodies. Comparable loading of proteins on the gel was verified by reprobing the blots with an antibody specific for the housekeeping gene product GAPDH.

Electrophoretic mobility shift assay. Nuclear extracts were prepared from HMECs grown in TCM for 16 h as described previously (25). The protein was quantified and 5 µg nuclear extract were incubated in 20 µL of buffer [20 mmol/L HEPES (pH 7.9), 0.4 mmol/L EDTA (pH 8.4), 0.4 mmol/L DTT, 10% glycerol, 100 µg/mL poli-dI/dC, and 1% NP40] with 32P-labeled, double-stranded oligonucleotides containing the Sp1 binding site with 80,000 cpm for 15 min at room temperature. For super shift, Sp1 antibodies were used. The reaction mixture was electrophoresed through a 5% polyacrylamide gel, and the gel was dried and subjected to autoradiography using phosphor screens at –70°C.

ChIP assay. ChIP assays were performed as per the manufacturer's instructions. Briefly, HMECs were growm in TCM for 16 h, fixed with formaldehyde (final concentration of 1%), and incubated for 10 min at 37°C. The cells were washed twice with ice-cold PBS containing protease inhibitors (1 mmol/L PMSF, 1 µg/mL aprotinin, and 1 µg/mL pepstatin-A), harvested, and lysed with SDS lysis buffer for 10 min on ice. The lysates were sonicated to shear the DNA to fragment lengths ~1,000 bp (amplitude 60%, 4 bursts of 10 s; Fisher Sonic Dismembrator 60). From each sonicated sample, 5% was used as the input control for immunoprecipitated fragments. The complexes were immunoprecipitated with antibodies specific for Sp1. Complexes were collected using salmon sperm DNA/protein-A agarose slurry and washed as per the manufacturer's instructions. The immune complexes were eluted with 1% SDS and 0.1 mol/L NaHCO3, and the crosslinks were reversed by incubation at 65°C for 4 h in the presence of 200 nmol/L NaCl. After treatment with proteinase-K at 45°C for 1 h, the DNA was purified by phenol/chloroform extraction, ethanol precipitation, and resuspended in 3 µL of nuclease-free water. PCR was carried out for 30 cycles with primers specific for the TIMP-3 promoter. PCR products were resolved on 2% agarose gels and analyzed using ethidium bromide staining.

Xenograft animal models. Severe combined immunodeficient (SCID) mice were maintained and experiments were performed as described previously (21). S.c. tumors were fixed in 10% phosphate-buffered formaldehyde.

Statistical analysis. All data are presented as means ± SE of at least three independent experiments, each performed at least in triplicate. One way ANOVA combined with the Tukey post hoc test of means was used for multiple comparisons in cell culture experiments. Statistical differences are presented at probability levels of P values of <0.05, <0.01, and <0.001.


   Results
Top
 Abstract
 Introduction
 Materials and Methods
 Results
Discussion
 Disclosure of Potential...
 References
 
Ad-MMP-2-Si inhibits angiogenesis. We have shown previously that the down-regulation of MMP-2 with Ad-MMP-2-Si inhibited tumor growth and lung metastasis in a mouse model (21). In the present study, TCM from A549 cells infected with mock (mock-CM) or 100 MOI of either Ad-SV (Ad-SV-CM) or Ad-MMP-2-Si (Ad-MMP-2-Si-CM) was used to induce HMEC to form a capillary network in an in vitro angiogenic assay. Figure 1A shows that MMP-2 activity is inhibited by >75% in the CM of A549 cells infected with 100 MOI of Ad-MMP-2 Si compared with mock and Ad-SV controls. Mock-CM and Ad-SV-CM elicited a strong angiogenic response and induced HMECs to differentiate into capillary-like structures within 16 hours. In contrast, Ad-MMP-2-Si-CM inhibited microvessel morphogenesis. Cells grown the presence with endothelial culture medium without serum (no serum) were just beginning to differentiate into capillaries at 16-hour time point (Fig. 1B). Quantification indicated a ~85% decrease in branch points and a ~90% decrease in vessel length in HMECs cultured with Ad-MMP-2-Si-CM compared with mock-CM and Ad-SV-CM (Fig. 1B).


Figure 1
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  		Figure 1. Ad-MMP-2-Si inhibits tumor-induced angiogenesis. A and B, A549 cells were infected as described in Materials and Methods. Briefly, A549 cells were infected with 100 MOI of either Ad-SV or Ad-MMP-2-Si, medium was aspirated after 36 h of incubation and 3 mL of serum-free medium were added, and cells were incubated overnight. A, gelatin zymographic analysis of TCM. The band intensities of MMP-2 activity were quantified by densitometry and normalized with the intensity of the mock-CM band. Columns, mean of triplicate experiments; bars, SE; *, P < 0.01, significant difference from mock-CM. B, in vitro angiogenesis (top): The TCM was added into 96-well plates, which were coated with Matrigel and preseeded with HMEC (2 x 104 cells per well). After overnight incubation at 37°C, cells were observed under the bright field microscope for formation of capillary-like structures. Bottom, the degree of angiogenic induction by Mock-CM, Ad-SV-CM, and Ad-MMP-2-Si-CM was quantified for the numerical value of the product of the relative capillary length and number of branch points per field. The capillary length and number of branch points were reduced drastically in HMECs grown in CM from Ad-MMP-2-Si–infected A549 cells. Columns, mean of quadruplicate experiments; bars, SE; *, P < 0.01, significant difference from mock-CM. C, in vivo angiogenic assay using the dorsal skin fold model as described in Materials and Methods. Briefly, the animals were implanted with diffusion chambers containing mock, 100 MOI of either Ad-SV or Ad-MMP-2-Si–infected cells in a dorsal cavity. Ten days after implantation, the animals were sacrificed and the skin fold covering the diffusion chamber was observed under bright field light microscope for the presence of tumor-induced neovasculature (TN) and preexisting vasculature (PV) and photographed (top). The mean number of neovasculature (>20 µm) per field was quantified (bottom). Columns, mean of quadruplicate experiments; bars, SE; *, P < 0.01, significant difference from mock.

 
We also examined whether Ad-MMP-2-Si could inhibit tumor angiogenesis in vivo as assessed by the dorsal window model. Implantation of a chamber containing mock or Ad-SV–infected A549 cells in the dorsal skin fold chamber resulted in the development of microvessels (arrows) with curved thin structures and many tiny bleeding spots. In contrast, implantation of A549 cells infected with Ad-MMP-2-Si resulted in the development of a few additional microvessels (Fig. 1C).

Ad-MMP-2-Si-CM inhibits endothelial cell proliferation and survival. To elucidate the cellular mechanisms underlying the inhibitory effect of TCM from tumor cells infected with Ad-MMP-2-Si on angiogenesis, we investigated the effect of TCM on the proliferation and survival of endothelial cells. We examined cell proliferation indices in endothelial cells grown in Ad-MMP-2-Si-CM. We assayed for the presence of the Ki-67 molecular marker (expressed in the G1-S-G2 phases of the cell cycle), and after immunocytochemical analysis, we quantified the number of positive cells in all treatment groups at x400 magnification. Figure 2A indicates that Ad-MMP-2-Si-CM caused a decrease in the fraction of cells expressing the proliferation marker Ki-67 at 72 hours. The controls had Ki-67 labeling indices of 45.01% ± 4.81% in mock-CM and 44.59% ± 5.26% in Ad-SV-CM, whereas Ad-MMP-2-Si-CM had a significantly lower index at 15.7% ± 2.11%. Having observed a marked reduction in the Ki-67 index in endothelial cells grown in Ad-MMP-2-Si-CM, we determined the presence of apoptosis by TUNEL assay. Figure 2B indicates a time-dependent increase in apoptotic index in endothelial cells grown in Ad-MMP-2-Si-CM. The number of cells staining positive for apoptosis were counted in 5 randomly selected fields at x400 magnification. TUNEL labeling indices in endothelial cells grown in the presence of mock-CM and Ad-SV-CM showed hardly any apoptosis for up to 72 hours (Fig. 2B). However, endothelial cells grown in the presence of Ad-MMP-2-Si-CM displayed a time-dependent increase in apoptosis. At the 24-hour time point, 7% to 8% of the cells were TUNEL positive; 31.1% ± 2.83% at the 48-hour time point; and 62.1% ± 5.15% at the 72-hour time point.


Figure 2
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  		Figure 2. Ad-MMP-2-Si-CM inhibits endothelial cell survival and proliferation. A, proliferating HMECs were identified by Ki-67 indirect immunofluorescence. Values were calculated as the number of Ki67-positive cells per number of cells with intact nuclei (hematoxylin staining) at x400 magnification. Columns, mean of quadruplicate experiments; bars, SE; *, P < 0.05, significant difference from mock-CM. B, determination of apoptosis in endothelial cells grown in the presence of Ad-MMP-2-Si-CM by TUNEL staining up to 72 h. The percentage of apoptotic cells were shown over time. Columns, mean of quadruplicate experiments; bars, SE; *, P < 0.001, significant difference from mock-CM.

 
Ad-MMP-2-Si-CM reduces VEGF expression and VEGFR2 phosphorylation in endothelial cells. To determine the effect of Ad-MMP-2-Si-CM on the expression of angiogenesis-inducing molecules, we used Western blotting and immunoprecipitation to assess the VEGF protein content and VEGFR2 phosphorylation of endothelial cells grown in TCM for 16 hours. Figure 3A indicates that VEGF protein levels were decreased in HMECs grown in Ad-MMP-2-Si-CM compared with HMECs grown in mock-CM and Ad-SV-CM. The phosphorylation state of VEGFR2 was assessed by immunoprecipitation of the receptor followed by phospho-tyrosine immunodetection in endothelial cells. Decreased tyrosine phosphorylation of VEGFR2 was observed in endothelial cells grown in the presence Ad-MMP-2-Si-CM (Fig. 3B). Densitometric normalization of VEGFR2 phosphorylation against VEGFR2 band density indicated that there was a 55% decrease in VEGFR phosphorylation.


Figure 3
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  		Figure 3. Ad-MMP-2-Si-CM inhibits expression of VEGF and phosphorylation of VEGFR2 in endothelial cells. A, HMECs grown on the Mock-CM, Ad-SV-CM, or Ad-MMP-2-Si-CM for 16 h were collected and lysed and immunoblotted for VEGF, PI3K, AKT, and pAKT using specific antibodies. The experiments were carried out thrice and representative immunoblot was shown. The blots were stripped and reprobed with GAPDH antibody to detect total amounts of the respective proteins. B, VEGFR2 was immunoprecipitated from solubilized cell extracts with a polyclonal anti-VEGFR2 antibody, and VEGFR2 phosphorylation was determined by immunoblotting using an anti–phospho-tyrosine antibody. The experiments were carried out thrice, and representative autoradiograms are presented. Densitometric normalization of VEGFR2 phosphorylation against VEGFR2 band density is shown. Columns, mean of triplicate experiments; bars, SE; *, P < 0.01, significant difference from mock-CM. C, endothelial cells were transfected with a plasmid expressing myr-AKT and were grown in presence of CM as described above. Cell lysates were immunoblotted for pAKT and VEGF. Densitometric normalization of VEGF and phospho-AKT is shown. Columns, mean of quadruplicate experiments; bars, SE; *, P < 0.05, significant difference from Ad-MMP-2-Si-CM treatment alone. D, an in vitro angiogenic assay was performed under the same conditions as described in Fig. 1. The experiments were carried out thrice, and representative pictures are shown. The degree of angiogenic induction was quantified for the numerical value of the product of the relative capillary length and number of branch points per field. Columns, mean of triplicate experiments; bars, SE; *, P < 0.05, significant difference from Ad-MMP-2-Si-CM treatment alone.

 
Ad-MMP-2-Si-CM suppresses AKT activation and enhances ERK phosphorylation. Because Ad-MMP-2-Si-CM inhibited VEGFR2 phosphorylation, we sought to determine whether the downstream mediators of VEGFR2 signaling were potential targets of Ad-MMP-2-Si-CM. VEGFR2 is characterized by tyrosine kinase activity and phosphorylation of secondary messengers. These secondary messengers seem to regulate endothelial cell proliferation via activation of ERK1/2 mitogen-activated protein kinases (MAPK; ref. 26) as well as endothelial cell survival via AKT activation (27). To assess the effect of Ad-MMP-2-Si-CM on these pathways, endothelial cells were grown in the presence of the TCM for 16 hours and cell lysates were subjected to immunodetection using antibodies against PI3K, AKT, phosphorylated AKT (Ser473), ERK, and phosphorylated ERK1/2. There is an increase in AKT phosphorylation in endothelial cells grown in the presence of TCM from mock-treated cells compared with no serum medium. However, Ad-MMP-2-Si-CM significantly decreased PI3K expression and phosphorylation of AKT in endothelial cells compared with the controls (Fig. 3A). The serine/threonine protein kinase AKT is an important component in migratory and prosurvival signaling pathways, and Akt-mediated signaling is stimulated by several angiogenic factors and plays a pivotal role for endothelial cell survival, migration, and proliferation, all of which are intimately involved in angiogenesis and vascular repair (28).

To assess this hypothesis directly, we transfected cells with constitutively active AKT, myr-AKT (29), and examined the implications for Ad-MMP-2-Si-CM–mediated inhibition on VEGF expression and endothelial cord formation. As shown in Fig. 3C, myr-AKT transfection reversed the inhibitory effect of Ad-MMP-2-Si-CM on phosphor-AKT and VEGF expression in endothelial cells. Moreover, overexpression of AKT in the presence of Ad-MMP-2-Si-CM reversed the inhibitory effect of Ad-MMP-2-Si-CM on endothelial tube formation (Fig. 3D). Quantification of tube length and branch points indicated that there was a ~5-fold increase in tube length and a ~2-fold increase in the number of branch points in myr-AKT–overexpressed cells grown on Ad-MMP-2-Si-CM compared to cells grown on Ad-MMP-2-Si-CM alone.

Conversely, an unexpected increase in ERK1/2 phosphorylation was observed in endothelial cells grown in Ad-MMP-2-Si-CM compared with mock-CM and Ad-SV-CM (Fig. 4A ). The ratio between phosphorylated ERK and total ERK indicated a 2-fold increase in p-ERK levels in Ad-MMP-2-Si-CM–treated endothelial cells compared with the controls. Moreover, overexpression of AKT inhibited Ad-MMP-2-Si-CM–induced phosphorylation of ERK, thereby confirming that this effect was a result of AKT (Fig. 4B). Taken together, these results suggest that inhibition of AKT phosphorylation by Ad-MMP-2-Si-CM plays a key role the inhibition of angiogenesis. In addition, Ad-MMP-2-Si-CM not only inhibited the downstream effects of the AKT survival pathway but also allowed overactivation of the ERK pathway via AKT blockade.


Figure 4
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  		Figure 4. Ad-MMP-2-Si-CM induces ERK-mediated expression of TIMP-3. Endothelial cells were grown in the presence of mock-CM, Ad-SV-CM, or Ad-MMP-2-Si-CM for 16 h. A, cellular extracts were subjected to immunoblotting with antibodies for phosphorylation-state–specific ERK1 and total ERK1 followed by chemiluminescence. Densitometric normalization of ERK phosphorylation against total protein band density is shown. *, P < 0.01, significant difference from mock-CM. B, endothelial cells were transfected with a plasmid expressing myr-AKT and were grown in presence of CM as described above. Cell lysates were immunoblotted for pERK, total ERK, and TIMP-3. Densitometry normalization of ERK phosphorylation against total protein band density is shown. Columns, mean of quadruplicate experiments; bars, SE; *, P < 0.01, significant difference from control; **, P < 0.01, significant difference from Ad-MMP-2-Si-CM treatment. C, endothelial cells were cultured as described above and extracellular matrix protein were analyzed by immunoblotting for Ad-MMP-2-Si-CM–induced TIMP-3 in endothelial cells. Densitometric analysis of TIMP-3 band density is shown in columns. Columns, mean of quadruplicate experiments; bars, SE; *, P < 0.001, significant difference from mock-CM. D, effect of ERK pathway on Ad-MMP-2-Si-CM–stimulated expression of TIMP-3 in endothelial cells. Endothelial cells were transfected with Dn-ERK followed by culturing on the tumor-CM from A549 cells infected with mock, Ad-SV, or Ad-MMP-2-Si. Cell lysates were immunoblotted with antibodies for phosphorylation-state–specific ERK1/2 and total ERK. ECM protein was analyzed for TIMP-3 immunoblotting. Densitometric normalization of ERK phosphorylation against total protein band density is shown. The quantification of TIMP-3 bands is shown in arbitrary units. Columns, mean of quadruplicate experiments; bars, SE; *, P < 0.01, significant difference from mock-CM; **, P < 0.01, significant difference from Ad-MMP-2-Si-CM treatment.

 
Ad-MMP-2-Si-CM–mediated ERK activation leads to TIMP-3 expression in endothelial cells. TIMP-3 has been reported to inhibit capillary morphogenesis of endothelial cells in vitro and basic fibroblast growth factor–induced angiogenesis in vivo (15). Therefore, we examined whether Ad-MMP-2-Si-CM induced TIMP-3 expression in endothelial cells. Figure 4C indicates a 4-fold increase in the 24-kDa band equivalent to nonglycosylated TIMP-3 in endothelial cells grown in Ad-MMP-2-Si-CM compared with the controls. To explore the requirement of ERK pathway stimulation for Ad-MMP-2-Si-CM–induced TIMP-3 expression, endothelial cells were transiently transfected with a dominant negative mutant of ERK (Dn-ERK) to inhibit ERK signaling and then grown in the presence of Ad-MMP-2-Si-CM for 16 hours. To ascertain if Dn-ERK affected ERK phosphorylation, we assessed ERK phosphorylation under these conditions. As shown in Fig. 4D, Dn-ERK transfection inhibited ERK phosphorylation and inhibited Ad-MMP-2-Si-CM–induced TIMP-3 expression. Similarly there was decrease in Ad-MMP-2-Si-CM–induced TIMP-3 expression in Myr Akt–transfected cells (Fig. 4B). Because p42/p44 (Erk1/2) MAPKs can phosphorylate Sp1 in vitro and in vivo (30) and the promoter region of the human TIMP-3 gene contains four Sp1 transcription factor–binding sites (31), we determined Sp1 binding activity in endothelial cells grown in the presence of Ad-MMP-2-Si-CM. As shown in Fig. 5A , Sp1 activity increased in endothelial cells grown in Ad-MMP-2-Si-CM compared with cells grown in CM from mock and Ad-SV–transfected cells. Preincubation of nuclear extracts with Sp1 antibody indicated a shift, thereby confirming the specificity of the band.


Figure 5
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  		Figure 5. Sp1 is a transcription factor for TIMP-3. A, endothelial cells were grown as described above and nuclear extracts subjected to electrophoretic mobility shift assay for Sp1 binding to its consensus sequence. B, HMECs were grown, fixed in formaldehyde, and immunoprecipitated with Sp1 antibody as described in Materials and Methods. Purified DNA was subjected to PCR with TIMP-3 primers. Densitometric analysis of the intensity of the bands is shown. Columns, mean of triplicate experiments; bars, SE; *, P < 0.01, significant difference from mock-CM. C, immunoprecipitation (IP) and immunoblot (IB) analyses from HMECs cultured as described in Materials and Methods. Immunoprecipitation of cell lysates with antibodies specific for VEGFR2 followed by immunoblotting with TIMP-3 and VEGF. For loading controls, bolts were reprobed with VEGFR2 antibody. D, after 24 h of treatment with either TIMP-3 siRNA or control siRNA transfection, HMECs were grown in CM for 72 h, and the apoptosis in endothelial cells grown in the presence of Ad-MMP-2-Si-CM was detected using the TUNEL assay. Columns, mean of triplicate experiments; bars, SE; *, P < 0.001, versus cells grown on Mock-CM; **, P < 0.001, versus cells grown on Ad-MMP-2-Si-CM.

 
To assess whether Sp1 interacts with the TIMP-3 promoter, we performed ChIP assays. Using an antibody specific for Sp1, protein chromatin complexes were immunoprecipitated, and the amount of TIMP-3–specific DNA that was released from the immunoprecipitates was analyzed by PCR. As shown in Fig. 5B, ChIP assay results indicated that Sp1 is strongly associated with TIMP-3 in HMECs grown in Ad-MMP-2-Si-CM compared with the controls. Quantitative analysis indicated that there was a 2-fold increase in Sp1 binding to the TIMP-3 promoter in endothelial cells grown in the response of AD-MMP-2-SI-CM compared with Mock-CM and Ad-SV-CM.

TIMP-3 expression contributes to decreased angiogenesis and endothelial apoptosis. TIMP-3 was shown to block the binding of VEGF to VEGFR2, thereby inhibiting downstream signaling and angiogenesis (18). We performed immunoprecipitation experiments to analyze VEGFR2 interactions with TIMP-3 and VEGF in endothelial cells. Figure 5C indicates that binding of VEGF to VEGFR2 was reduced in endothelial cells grown in Ad-MMP-2-Si-CM compared with mock-CM and Ad-SV-CM. We also observed increased binding of TIMP-3 to VEGFR2 in endothelial cells cultured with Ad-MMP-2-Si-CM compared with mock-CM and Ad-SV-CM (Fig. 5C). Because TIMP-3 induces apoptosis in various types of cells (32–35), we further evaluated whether the elevated TIMP-3 levels were responsible for the apoptosis in HMECs grown in Ad-MMP-2-Si-CM. Endothelial cells were transfected with TIMP-3 siRNA and grown in the TCM for 72 hours. Ad-MMP-2-Si-CM–induced apoptosis induction was partially reversed by TIMP-3 siRNA, thereby confirming that the apoptosis was mediated via up-regulation of TIMP-3 (Fig. 5D).

Ad-MMP-2-Si treatment suppresses angiogenesis in vivo. Having shown that Ad-MMP-2-Si infection in lung cancer cells inhibited angiogenesis using capillary formation assay and an in vivo dorsal air sac assay, we next sought to determine whether the tumor growth inhibition observed in a spontaneous metastasis model after Ad-MMP-2-Si treatment (21) was due to an extension of the antiangiogenic activity. The tumor cores of the mock, Ad-SV, and Ad-MMP-2-Si–treated SCID mice were analyzed immunohistologically with anti-CD31. The immunoreactivity of CD31 in the tissue sections from the tumors derived from mice that received each treatment provided some measure of vascularity as affected by Ad-MMP-2-Si treatment (Fig. 6 ). Compared with mock and Ad-SV–treated mice, tumor sections from Ad-MMP-2-Si–treated mice showed decreased CD31. To confirm endothelial cell apoptosis, we performed TUNEL staining on the same sections. Endothelial cell apoptosis as determined by TUNEL and CD31 double staining was significantly higher in the tumor sections from mice treated with Ad-MMP-2-Si compared with the mock and Ad-SV–treated mice (18 ± 2.1 versus 0.5 ± 1.5, respectively). These results suggest that inhibition of MMP-2 inhibited angiogenesis through stimulation of apoptosis in endothelial cells.


Figure 6
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  		Figure 6. MMP-2 inhibition using Ad-MMP-2-Si decreased angiogenesis and induced endothelial cell apoptosis in lung tumors. Representative sections (magnification, x400) from mock and Ad-SV–treated tumors and those treated with Ad-MMP-2-Si. Ad-MMP-2-Si–treated mice showed a decrease in angiogenesis as determined by CD31 staining. Also shown is a negative staining control for CD31 where the antibody was replaced by nonimmune serum (negative control). Dual staining for TUNEL-positive and CD31-positive cells indicating endothelial apoptosis in Ad-MMP-2-Si–treated mice compared with mock and Ad-SV–treated mice.

 

   Discussion
Top
 Abstract
 Introduction
 Materials and Methods
 Results
 Discussion
Disclosure of Potential...
 References
 
Treatment for cancer is focused on developing treatments based on inhibiting tumor angiogenesis, as tumors require a blood supply for nutrition and growth. Such therapy, termed "antiangiogenic" therapy, has now been documented to prolong the survival of patients with lung cancer in large randomized clinical trials. In the present study, we show the role of MMP-2 inhibition in tumor cells by targeting angiogenic components of tumor growth and identifying particular molecules and/or signaling pathways involved in the angiogenic process. We provide evidence that MMP-2 inhibition in tumor cells triggers signaling pathways that ultimately lead to endothelial cell death.

As an initial step of our study, we observed that TCM from lung cancer cells infected with Ad-MMP-2-Si inhibited capillary tube formation in endothelial cells compared with the controls. We also observed decreased proliferative index Ki-67 and increased apoptosis in endothelial cells grown in the presence of tumor-CM from lung cancer cells infected with Ad-MMP-2-Si (Ad-MMP-2-Si-CM). We next carried out experiments designed to identify signaling molecules secreted by endothelial cells that might be responsible for the observed effect on endothelial cells. Endothelial cell differentiation leading to tubule formation is primarily established by activation of VEGF and its receptor. VEGFR2 is the main human receptor responsible for both physiologic and pathologic vascular development, and the VEGF-VEGFR2 signaling pathway has become an important target for the development of antiangiogenic agents (36). In a study of the significance of the various VEGF isoforms in non–small cell lung cancer (NSCLC), Yuan and colleagues (37) reported that a high VEGF-189 level correlated with high intratumoral microvessel density, short survival, and early postoperative relapse, whereas VEGF-121 was correlated with short survival and relapse. Another group also reported a similar correlation between poor prognosis and localized elevated microvessel density at the advancing front of NSCLC tumors accompanied by increased localized staining of VEGF and its receptor (38). In this study, we show that a mechanism in which the paracrine effect of MMP-2–down-regulated lung tumor cells inhibited VEGF expression in endothelial cells, thereby leading to inhibition of the PI3K/AKT pathway. Activated PI3K and AKT are strong inducers of neovascularization and endothelial cell proliferation (39). We also show that TIMP-3 induction occurred under the same conditions and binds to VEGFR2. Previous studies have also shown that TIMP-3 binding to VEGFR2 inhibited downstream signaling and angiogenesis (18).

To determine the functional consequences of the up-regulation of ERK pathway, we manipulated Ad-MMP-2-Si-CM–induced ERK pathway up-regulation and determined the implications for endothelial cell apoptosis. The inhibitory effect of Dn-ERK on TIMP-3 expression implies that the ERK1/2 signal pathway is critical for TCM-induced TIMP-3 expression. To begin to understand the underlying mechanisms of the correlation between levels of ERK and TIMP-3, we evaluated the activity of Sp1, a known transcriptional regulator of TIMP-3 (40). Our results show that Sp1 was induced in endothelial cells grown in the presence of Ad-MMP-2-Si-CM. However, we cannot rule out the possible involvement of other transcription factors in TIMP-3 expression. Nevertheless, it is certain that the Sp1 transcription factor is dominantly involved in the induction of TIMP-3 as determined by ChIP assay. Our studies indicate that TCM from lung cancer cells infected with Ad-MMP-2-Si activates the ERK-MAPK pathway, which in turn, could increase Sp1 phosphorylation and its DNA binding activity leading to TIMP-3 induction. Our results support previous findings that Sp1 is an important link between increased activation of the ERK pathway and TIMP-3 expression (40). We further show the significance of TIMP-3 expression in the induction of endothelial apoptosis using siRNA. These results suggest that the induction of TIMP-3 in Ad-MMP-2-Si–treated mice inhibited in vivo angiogenesis by inhibition of VEGF/VEGFR2 signaling and induction of endothelial apoptosis. Induction of apoptotic cell death by TIMP-3 overexpression is well-documented and has been shown in vascular cells and cancer cells in vitro and in vivo (41–44). Intratumoral injection with TIMP-3–expressing adenovirus inhibits growth of melanomas and squamous cell carcinomas by inducing apoptosis and inhibiting tumor angiogenesis (45). Furthermore, Mino and colleagues (46) observed that TIMP-3 expression status is significantly correlated with pathologic stage and nodal involvement in resected NSCLC, and that the overall survival rate of patients with elevated TIMP-3 expression is high.

In summary, we show that MMP-2 inhibition in lung tumor cells inhibits VEGF-mediated AKT activation in endothelial cells, leading to increased ERK activation that contributes to TIMP-3 induction. We have also established the importance of these pathways in the induction of endothelial cell apoptosis. These data provide insights about the mechanisms of MMP-2 inhibition in tumor cells, which led to decreased angiogenesis and resulted in tumor growth inhibition.


   Disclosure of Potential Conflicts of Interest
Top
 Abstract
 Introduction
 Materials and Methods
 Results
 Discussion
 Disclosure of Potential...
References
 
No potential conflicts of interest were disclosed.


   Acknowledgments
 
Grant support: National Cancer Institute Grant CA75557, CA92393, CA95058, CA116708 and National Institute of Neurological Disorder and Stroke NS47699, NS57529, NS61835, and Caterpillar, Inc., OSF Saint Francis, Inc., Peoria, IL (J.S. Rao) and American Cancer Society Grant #06-03 (S.S. Lakka).

The costs of publication of this article were defrayed in part by the payment of page charges. This article must therefore be hereby marked advertisement in accordance with 18 U.S.C. Section 1734 solely to indicate this fact.

We thank Noorjehan Ali for technical assistance, Shellee Abraham for assistance in manuscript preparation, Diana Meister and Sushma Jasti for review of this article, and Professor Richard A. Roth for the construct encoding the constitutively active myristoylated AKT (myr-AKT {delta} 4-129).

Received 12/11/07. Revised 3/13/08. Accepted 3/21/08.


   References
Top
 Abstract
 Introduction
 Materials and Methods
 Results
 Discussion
 Disclosure of Potential...
 References
 

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