Endostatin Inhibits Human Tongue Carcinoma Cell Invasion and Intravasation and Blocks the Activation of Matrix Metalloprotease-2, -9, and -13*

Pia Nyberg {ddagger}, Pia Heikkilä §, Timo Sorsa §, Jani Luostarinen ¶, Ritva Heljasvaara ¶, Ulf-Håkan Stenman ||, Taina Pihlajaniemi ¶ and Tuula Salo {ddagger} ** {ddagger}{ddagger}

From the {ddagger}Department of Diagnostics and Oral Medicine, Institute of Dentistry, University of Oulu, FIN-90014 Oulu, the §Department of Oral and Maxillofacial Diseases, Helsinki University Central Hospital, Institute of Dentistry, University of Helsinki, Orton Research Institute and the Orthopedic Hospital of Invalid Foundation, FIN-00014 Helsinki, the Collagen Research Unit, Biocenter and Department of Medical Biochemistry and Molecular Biology, University of Oulu, FIN-90014 Oulu, the ||Department of Clinical Chemistry, Helsinki University Central Hospital, FIN-00029 Helsinki, and the **Institute of Dentistry, University of Helsinki and Helsinki University Dental Clinic, FIN-00014 Helsinki, Finland

Received for publication, October 9, 2002 , and in revised form, March 4, 2003.


    ABSTRACT
 TOP
 ABSTRACT
 INTRODUCTION
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 REFERENCES
 
Endostatin, a 20-kDa collagen XVIII fragment, inhibits angiogenesis and tumor growth in vivo, but the mechanisms are still unclear. Matrix metalloproteases (MMPs), a family of extracellular and membrane-associated endopeptidases, collectively digest almost all extracellular matrix and basement membrane components, and thus play an important role in tumor progression. We studied the effects of recombinant human endostatin on human MMP-2, -9, -8, and -13. We found that endostatin inhibited the activation and catalytic activity of pro-MMP-9 and -13 as well as recombinant pro-MMP-2. It prevented the fragmentation of pro-MMP-2 that was associated with reduction of catalytic activity. Endostatin had no effect on MMP-8 as shown by collagenase activity assays. An in vitro migration assay and an in vivo chicken chorioallantoic membrane intravasation assay with the human tongue squamous cell carcinoma cell line HSC-3 revealed the biphasic nature of endostatin; low endostatin concentrations inhibited intravasation and migration of these cells in a dose-dependent manner, but at increased concentrations, the inhibitory effect was far less efficient. The results show that endostatin blocks the activation and activities of certain tumor-associated pro-MMPs, such as pro-MMP-2, -9, and -13, which may explain, at least in part, the antitumor effect of endostatin. Our results also suggest that endostatin inhibits tumor progression by directly affecting the tumor cells and not just acting via endothelial cells and blockage of angiogenesis.


    INTRODUCTION
 TOP
 ABSTRACT
 INTRODUCTION
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 REFERENCES
 
Neoangiogenesis, the formation of new capillaries, is among the key factors in various tissue destructive pathological processes, such as tumor growth and metastasis (1). The expansion of solid tumors is critically dependent on the neoangiogenesis, and the inhibition of vascular supply to tumors can suppress their growth (2). Angiogenesis is thought to depend on a delicate balance between endogenous stimulators and inhibitors (1). A number of anti-angiogenic substances have been described, of which many are fragments of naturally occurring proteins (3).

Endostatin, an angiogenesis inhibitor identified and purified from a murine hemangioendothelioma cell line (4), corresponds to a 20-kDa fragment derived from the carboxyl-terminal noncollagenous NC11 domain of type XVIII collagen (46). Recombinant endostatin efficiently blocks angiogenesis and suppresses primary tumor and metastasis growth in experimental animal models without any apparent side effects, toxicity, or developmental drug resistance (4, 7, 8). However, the molecular mechanisms of endostatin in inhibition of tumor growth are not yet clear (8). Some recent studies have suggested that endostatin interferes with fibroblast growth factor-2-induced signal transduction blocking endothelial cell motility (9), causes G1 arrest of endothelial cells through inhibition of cyclin D1 (10), and blocks vascular endothelial growth factor-mediated signaling via a direct interaction with the kinase-insert domain-containing receptor/Flk-1 receptor tyrosine kinase in human umbilical vein endothelial cells (11).

Matrix metalloproteases (MMPs), a family of secreted or transmembrane enzymes, can collectively digest almost all extracellular matrix and basement membrane components (12). Thus, MMPs are largely implicated in promoting angiogenesis and tumor metastasis (13, 14). In particular, the activities of gelatinases (MMP-9 and -2) and collagenase-3 (MMP-13) correlate with the invasive potential of cancer, e.g. in head and neck squamous cell carcinomas (15, 16) and in breast cancer (17). MMPs are secreted in latent proforms, and they require activation in the extracellular milieu or on the cell surface to be catalytically competent. Activation of MMPs can be achieved in vitro with various agents, such as organomercurials (aminophenylmercuric acetate (APMA)) (18), or in vivo in a network with other proteases, e.g. tumor-associated trypsinogen-2 (TAT-2), which has been shown to be an efficient activator of MMP-9 (1921). Kim et al. (22) have recently shown that endostatin significantly reduces invasion of endothelial as well as tumor cells into a reconstituted basement membrane by inhibiting the activation of pro-MMP-2 and the catalytic activities of MT1-MMP and MMP-2 (22). Furthermore, they showed that endostatin binds to the catalytic domain of MMP-2 (23). It has been found previously that certain MMPs can generate endostatin-containing peptides differing in molecular size (20–30 kDa) from human type XVIII collagen (24).2 These fragments inhibit the proliferation and migration of human umbilical vein endothelial cells in a similar fashion as native 20-kDa endostatin.2

Since the effect of endostatin on MMPs other than MMP-2 is not known, the aim of the present study was to determine the ability of endostatin to inhibit the activation and enzymatic activity of human pro-MMP-9 as well as pro-MMP-2, -8, and -13. Furthermore, we studied the efficacy of endostatin in inhibiting human tumor cell migration, intravasation, and invasion.


    MATERIALS AND METHODS
 TOP
 ABSTRACT
 INTRODUCTION
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 REFERENCES
 
Cell Cultures—A human tongue squamous cell carcinoma cell line HSC-3 (JCRB Cell Bank 0623, National Institute of Health Sciences, Japan) was grown in 1:1 Dulbecco's modified Eagle's medium (Invitrogen) and Ham's Nutrient Mixture F-12 (Invitrogen) supplemented with 10% heat-inactivated fetal calf serum, 100 units/ml penicillin, 100 µg/ml streptomycin, 50 units/ml nystatin, 250 ng/ml fungizone, 1 mM sodium pyruvate, 2 mM L-glutamine (all supplements from Invitrogen), and 0.4 ng/ml hydrocortisone (Diosynth, Oss, The Netherlands).

Expression and Purification of Recombinant Human Endostatin— The cloning of the recombinant human endostatin used in this work has been described earlier (25). A fragment of human collagen XVIII that corresponds to the mouse endostatin sequence (4) was cloned to an expression vector and transformed into the Escherichia coli strain M15(pRep4). Recombinant proteins were expressed according to the protocol suggested by Qiagen (Hilden, Germany). The His-tagged recombinant proteins were purified according to a protocol described previously (25).

Inhibition of APMA-mediated Pro-MMP-9 Activation by Endostatin in HCS-3 Cells—18,000, 28,000, or 38,000 HSC-3 cells per well were plated on 24-well plates (Nunclon, Roskilde, Denmark), and the cells were allowed to grow in normal media for 24 h. Then the HSC-3 cells were incubated with serum-free medium containing 0, 1, 5, 10, 12.5, 15, 17.5, 20, 30, or 40 µg/ml recombinant endostatin for 48 h. After collection, the media were concentrated 4-fold and treated with 1 mM APMA (Sigma) at 37 °C for 25 min to observe the effect of endostatin on pro-MMP-9 and pro-MMP-2. The APMA activation was stopped by adding nonreducing Laemmli buffer. MMP-9 processing was detected by gelatin zymography and at least four separate experiments were quantitated by densitometric scanning with ScionImage (Scion Corp.).

Zymography—Gelatin zymography was performed in 10% SDS-PAGE that had been cast in the presence of 1 mg/ml fluorescently (2-methoxy-2,4-diphenyl-3-[2H]furanone; Fluka, Ronkonkoma, NY) labeled gelatin (26). After electrophoresis, SDS was removed by 2.5% Triton X-100 to renature the gelatinases. Gels were then incubated in 50 mM Tris-HCl buffer (pH 7.8, 150 mM NaCl, 5 mM CaCl2, 1 µM ZnCl2) overnight at 37 °C. The degradation of gelatin was visualized under long wave UV light. Gels were also stained with 0.5% Coomassie Blue R-250. The intensities of the separate bands from at least four separate experiments were measured quantitatively by ScionImage software.

Gelatin Degradation Assays—The degradation of gelatin was also assayed using 125I-labeled gelatin as a substrate (27). 50 ng of recombinant MMP-9 (Oncogene Research Products, Boston, MA), MMP-2 purified from human gingival fibroblast cultures, or MMP-9 purified from human gingival keratinocyte culture media (28) was incubated with 0, 0.05, 5, 10, or 20 µg of recombinant endostatin in 20 µl of buffer (50 mM Tris-HCl, pH 7.8, 200 mM NaCl, 1 mM CaCl2) for 1 h at 37 °C. Then the samples were treated with or without 1.3 mM APMA at 37 °C for 1 h and incubated with soluble 125I-labeled gelatin (1.5 µM)for1hat 37 °C. Undegraded gelatin was precipitated with 20% trichloroacetic acid. The radioactivity in the supernatants, containing the degraded gelatin and reflecting the gelatinase activity, was counted with a {gamma}-counter (Clinigamma, LKB Wallac, Turku, Finland). The radioactivity reflected gelatinase activity (28).

Western Blotting—Various amounts of recombinant endostatin (0, 120, 360, 600, or 960 ng) were incubated with 50 ng of recombinant MMP-2 and -9 at 37 °C for 45 min with or without APMA treatment (1 mM at 37 °C for 1 h). Pro-MMP-9 was also activated using 10 ng of TAT-2 (purified from serum-free conditioned medium of COLO 205 colon carcinoma cells or cyst fluid of ovarian tumors as described in Refs. 29 and 30) for 45 min at 37 °C (19), and the reaction was stopped with 80 ng of the specific tumor-associated trypsinogen inhibitor, purified from urine of cancer or pancreatitis patients as described previously (31, 32). Protein samples were separated on 12% SDS-polyacrylamide gels under reducing conditions and electrotransferred to nitrocellulose membranes (Schleicher & Schüll). Nonspecific binding was blocked with 5% non-fat dry milk for 90 min at 37 °C. The membranes were incubated with the monoclonal anti-MMP-2 (1:400 dilution; Chemicon) and polyclonal anti-MMP-9 (1:500 dilution; Ref. 33) antibodies for 3 h at 37 °C and followed by incubation with peroxidase-conjugated goat anti-rabbit (for polyclonal antibodies) or anti-mouse (for monoclonal antibodies) immunoglobulins (1:200 dilution; DAKO A/S, Glostrup, Denmark) for 1 h at room temperature. After extensive washing, the immunoreactive proteins were visualized with ECL Western blotting detection reagents (Amersham Biosciences) or with 60 mg/ml diaminobenzidine tetrahydrochloride in 50 mM Tris-HCl, pH 8.0, and 0.003% H2O2 (34).

Immunomagnetic Precipitation—Various concentrations (0, 0.5, and 5 µg/ml) of recombinant endostatin carrying a histidine tag were preincubated with 200 ng of MMP-9 (purified from human gingival keratinocyte culture media (28)) or with 200 ng of recombinant MMP-9 (Oncogene Research Products, Boston, MA) for2hat room temperature in PBS. 50 µl of IgG-coated magnetic beads (Dynabeads® M-280 Sheep anti-Mouse IgG, Dynal, Oslo, Norway) were conjugated with 5 µg of a monoclonal penta-His antibody (Qiagen, Hilden, Germany) for 2 h in 0.1% BSA/PBS, pH 7.4, at room temperature. The beads were washed twice with 500 µl of 0.1% BSA/PBS, suspended with the preincubated MMP-9-endostatin mix, and incubated 2 h at room temperature. Unbound proteins were removed by washing the beads three times for 5 min with 1% Triton X-100, 0.1% SDS-PBS. The bound protein precipitates were removed from the beads by boiling in loading buffer for 5 min, fractionated on a 12% SDS-PAGE, and identified by Western blotting using penta-His antibody and polyclonal MMP-9 antibody (33).

Collagen Degradation Assay—The degradation of native type I collagen by MMP-8 and -13 was determined by a collagen degradation assay. 50 ng of recombinant human MMP-8 or -13 (Invitrotek GmbH, Berlin, Germany) was incubated with 600 ng of recombinant endostatin at 37 °C for 45 min with or without APMA treatment (1 mM at 37 °C for 1 h), after which 1.5 µM type I collagen substrate (34) was added and incubated for 5 h at room temperature. The proteins were electrophoresed by 12% SDS-PAGE and stained with Coomassie Brilliant Blue.

In Vitro Cell Migration Assay—HSC-3 cell migration was studied using 8.0-µm pore size and 6.5-mm diameter Transwell inserts (Costar, Cambridge, MA) that were coated with type I collagen (BD Biosciences, 1 µg/cm2 in 10 mM acetic acid for 1 day) and equilibrated in serum-containing medium for 2 h before use. Cells were preincubated at 37 °C in a humidified 5% CO2 atmosphere for 30 min in the presence of 5 or 20 µg/ml endostatin. For migration assay, 600 µl of the serum-containing medium was added to the lower compartment of the migration apparatus, and 20,000 cells in a volume of 100 µl of serum-containing medium were plated on a type I collagen-coated Transwell filter. After culturing for 6 h, the cells were fixed in methanol, washed, and stained in toluidine blue. The cells were removed from the upper surface of the membrane with a cotton swab, and the cells that migrated to the underside of the membrane were counted under the microscope (35, 36).

CAM Assay—The chorioallantoic membrane (CAM) assay was done according to Kim et al. (37) except for a few modifications. Chick embryos were maintained at 37 °C in a humidified incubator and turned manually at least three times a day. HSC-3 cells used for the invasion experiment were detached from the culture dish, counted, and resuspended in PBS++ solution (13.7 mM NaCl, 2.7 mM KCl, 8 mM Na2HPO4, 1.5 mM KH2PO4, 700 µM CaCl2 2H2O, 490 µM MgCl2 6H2O). 2 x 106 cells in 50 µlofPBS++ with different amounts of endostatin (0, 0.5, 5, 20, and 50 µg/CAM) were inoculated onto a CAM of 10-day-old chick embryos. After 50 h of incubation, the eggs were cut along the long circumference, and the upper halves (with the inoculum) and the contents of the lower halves of the eggs were discarded. The CAMs lining the cavity of the eggshell were removed, snap-frozen, and used for extraction of genomic DNA. The frozen CAMs were crushed to fine powder, suspended in digestion buffer (100 mM NaCl, 10 mM Tris-Cl, pH 8.0, 25 mM EDTA, pH 8.0, 0.5% SDS, 0.1 mg/ml proteinase K), and incubated at 50 °C for 18 h. The samples were extracted with phenol/chloroform/isoamyl alcohol (25:24:1) and centrifuged 10 min at 1700 x g. The DNA in the aqueous phase was precipitated with 0.5 volumes of 7.5 M ammonium acetate and 2 volumes of ethanol, centrifuged 2 min at 1700 x g, washed, dried, and resuspended in sterile water. The radioactive PCR, which produced an Alu band of 224 bp, was done according to Kim et al. (37) except that the enzyme used in the PCR was Dynazyme (Finnzymes, Helsinki, Finland). The PCR products were electrophoresed on a 6% polyacrylamide gel at 1500 V for 1.5 h and exposed to film at –70 °C. The bands from four separate experiments were quantitated by densitometric scanning using ScionImage software.


    RESULTS
 TOP
 ABSTRACT
 INTRODUCTION
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 REFERENCES
 
Endostatin Inhibits the Activation of Pro-MMP-9 and Fragmentation of Pro-MMP-2 in Tongue Carcinoma Cell Cultures—To determine whether the anti-invasive activity of endostatin is associated with the inhibition of MMP-9, we analyzed the effect of endostatin on the levels of and degree of activation of the MMP-9 secreted by the cultured HSC-3 tongue carcinoma cells. Gelatin zymography of the serum-free culture medium revealed that endostatin clearly blocked the APMA-mediated pro-MMP-9 activation at endostatin concentrations from 12.5 to 40 µg/ml in a dose-dependent manner (Fig. 1A, lanes 5–10). The molecular sizes of pro-MMP-9 and active MMP-9 were 92 and 77 kDa, respectively. Without APMA activation endostatin had no effect on MMP-9 (data not shown). Endostatin had no apparent effect on the barely detectable 72-kDa pro-MMP-2 levels and activation present in HSC-3 cell media, but it very clearly inhibits the formation of smaller (~40 kDa) activation products of MMP-2 (Fig. 1A). Fig. 1B represents the scanned intensities of the gelatinolytic bands, and the calculated ratios of pro-MMP-9 versus active MMP-9. The ratio increased slightly even with lower concentrations of endostatin (1–10 µg/ml). With endostatin concentrations of 12.5 and 15 µg/ml the ratio was 1.6-fold, and at endostatin concentrations of 17.5, 20, 30, and 40 µg/ml the ratios were 1.9-, 2.1-, 2.4-, and 3.1-fold, respectively, in relation to the controls (Fig. 1B). We further studied the effect of endostatin on the catalytic activities of gelatinases by a gelatin degradation assay. Endostatin decreased the activity of APMA-activated MMP-9 (purified from human gingival keratinocyte culture media) from 100 to 92% (S.D., ±2.8%), 85% (S.D., ±2.8%), 76% (S.D., ±5.8%), and 61% (S.D., ±4.3%) with endostatin concentrations of 0, 0.05, 5, 10, and 20 µg, respectively (Fig. 1C). Endostatin also inhibited the activity of MMP-9 without treatment with APMA but slightly less efficiently; gelatin degradation was decreased from 100 to 98% (S.D., ±5%), 97% (S.D., ±8.9%), 82% (S.D., ±5.6%), and 70% (S.D., ±4.3%) with endostatin concentrations of 0, 0.05, 5, 10, and 20 µg, respectively (Fig. 1D).



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FIG. 1.
Inhibition of pro-MMP-9 activation and MMP-2 fragmentation by endostatin in the culture medium of HSC-3 tongue carcinoma cells. A, 28,000 HSC-3 tongue carcinoma cells per 24 wells were grown overnight and then incubated in serum-free medium for 48 h in the presence of various concentrations of recombinant endostatin. To activate pro-MMP-9 and pro-MMP-2, the collected media were concentrated 4-fold and treated with APMA (1 mM 37 °C for 25 min), and samples were analyzed with gelatin zymography. APMA treatment without endostatin (C, lane 1), 1 µg/ml endostatin + APMA (lane 2), 5 µg/ml endostatin + APMA (lane 3), 10 µg/ml endostatin + APMA (lane 4), 12.5 µg/ml endostatin + APMA (lane 5), 15 µg/ml endostatin + APMA (lane 6), 17.5 µg/ml endostatin + APMA (lane 7), 20 µg/ml endostatin + APMA (lane 8), 30 µg/ml endostatin + APMA (lane 9), and 40 µg/ml endostatin + APMA (lane 10). B, the bands in the zymography were quantitated with ScionImage software, and pro-MMP-9 versus active MMP-9 ratios were calculated (mean ± S.D., n is at least 3). C and D show enzyme activity assay results with MMP-9 purified from cell culture media and recombinant endostatin. 50 ng of pro-MMP-9 was incubated with endostatin (0.05, 5, 10, or 20 µg) for 1 h at 37 °C and after that with (C) or without (D) 1.3 mM APMA for 1 h at 37 °C and finally with 125I-labeled gelatin for 1 h at 37 °C. After precipitation of the undegraded gelatin, the degraded gelatin in the supernatants reflecting the MMP-9 activity was counted with a {gamma}-counter (mean ± S.D., n = 3).

 

The HSC-3 Cell Density Affects the Ability of Endostatin to Inhibit MMP-9 —The density of HSC-3 cells per well (28,000 cells/well in 24-well plates) was found to be rather critical for the inhibitory effect of endostatin in the cell culture experiments (Fig. 2A). At lower cell densities (18,000 cells/well), the effect of endostatin was much less clear, and at higher densities (38,000 cells/well), the effect of endostatin on pro-MMP-9 activation disappeared almost completely (Fig. 2A). Fig. 2B represents the scanned intensities of the gelatinolytic MMP-9 bands and the calculated ratios of pro-MMP-9 versus active MMP-9 with increasing concentration of endostatin.



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FIG. 2.
The effect of HSC-3 cell density on MMP-9 activation by endostatin. A, 18,000 (upper panel), 28,000 (middle panel), or 38,000 (lower panel) HSC-3 tongue carcinoma cells per 24 wells were grown overnight and then incubated in serum-free medium for 48 h in the presence of various concentrations of recombinant endostatin. To activate pro-MMP-9, the collected media were concentrated 4-fold and treated with APMA (1 mM 37 °C for 25 min), and samples were analyzed with gelatin zymography. APMA treatment (lane 1), 1 µg/ml endostatin + APMA (lane 2), 5 µg/ml endostatin + APMA (lane 3), 10 µg/ml endostatin + APMA (lane 4), and 40 µg/ml endostatin + APMA (lane 5). B, the bands in the zymography were quantitated with ScionImage software. Pro-MMP-9 versus active MMP-9 ratios were calculated with endostatin concentrations of 0, 5, and 40 µg/ml. White bars represent the cell density of 18,000 cells/well; black bars represent 28,000 cells/well, and striped bars represent 38,000 cells/well.

 

Endostatin Inhibits APMA and TAT-2-mediated Activation of Recombinant Human Pro-MMP-9 —Recombinant human endostatin was incubated with recombinant pro-MMP-9 that was activated with APMA or TAT-2. The processed forms of MMP-9 were then separated by SDS-PAGE and identified by Western blotting. The Western band intensities of pro- and active MMP-9 were quantitated by scanning. As was expected based on the results of zymography, endostatin decreased the APMA-mediated activation of MMP-9 in a dose-dependent manner (Fig. 3A). The ratios of pro/active MMP-9 increased 1.6-fold when endostatin was present compared with the control treated only with APMA (Fig. 3B). The naturally occurring MMP-9 activator TAT-2 was also incubated with pro-MMP-9 and different amounts of endostatin. Endostatin decreased TAT-2-mediated conversion of the 92-kDa pro-MMP-9 to the 77-kDa active form (Fig. 3C). The ratio of the pro/active MMP-9 band intensity increased up to 7-fold with the highest amount of endostatin (1000 ng) compared with the control treated with only TAT-2, and even with 200 ng of endostatin the ratio was 2.7-fold higher than the pro/active MMP-9 ratio with only TAT-2 and without endostatin (Fig. 3D).



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FIG. 3.
The effect of endostatin on APMA- or TAT-2-activated MMP-9. Recombinant human pro-MMP-9 (50 ng) was incubated with endostatin at 37 °C for 45 min and then activated either with 1 mM APMA (60 min at 37 °C) or 10 ng of TAT-2 (60 min at 37 °C). Native and processed MMP-9 were separated by SDS-PAGE and identified with Western blotting with polyclonal MMP-9 antibody. A, 50 ng of recombinant pro-MMP-9 (lane 1), MMP-9 treated with APMA (lane 2), MMP-9 treated with APMA and 120 ng of endostatin (lane 3), MMP-9 with APMA and 360 ng of endostatin (lane 4), MMP-9 with APMA and 600 ng of endostatin (lane 5), and MMP-9 with APMA and 960 ng of endostatin (lane 6). B, the zymographic bands in A were quantitated with ScionImage software and pro-MMP-9 versus active MMP-9 ratios were calculated. C, 50 ng of recombinant pro-MMP-9 (lane 1), MMP-9 treated with purified 10 ng of TAT-2 (lane 2), MMP-9 treated with TAT-2 and 200 ng of endostatin (lane 3), MMP-9 with TAT-2 and 400 ng of endostatin (lane 4), MMP-9 with TAT-2 and 600 ng of endostatin (lane 5), MMP-9 with TAT-2 and 800 ng of endostatin (lane 6), and MMP-9 with TAT-2 and 1000 ng of endostatin (lane 7). D, the zymographic bands in C were scanned with ScionImage, and pro-MMP-9 versus active MMP-9 ratios were calculated. The data represent mean ± S.D. (n = 3).

 

Endostatin Binds to Pro-MMP-9 —Since endostatin inhibited the activation of pro-MMP-9 by both APMA and TAT-2, it might be possible that endostatin forms a complex with MMP-9. To examine this, we incubated recombinant MMP-9 and MMP-9 purified from human gingival keratinocyte culture media with different amounts of endostatin carrying a His tag. The complex was immunoprecipitated with the anti-His antibody. A Western immunoblot shows that pro-MMP-9 was co-precipitated in the presence of His-tagged endostatin in a dose-dependent manner (Fig. 4, lanes 6 and 7). However, a trace amount of MMP-9 was detectable even in the absence of endostatin (Fig. 4, lane 5). The result was similar with recombinant MMP-9 (data not shown).



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FIG. 4.
Interaction of endostatin with pro-MMP-9. MMP-9 purified from cell culture media was incubated with different amounts of recombinant endostatin carrying a His tag, immunoprecipitated with an anti-His antibody, and detected with Western immunoblotting. 200 ng of MMP-9 (lane 1), 200 ng of MMP-9 and 0.5 µg/ml endostatin (lane 2), 200 ng of MMP-9 and 5 µg/ml endostatin (lane 3), and 20 ng endostatin (lane 4) were detected with the anti-His antibody. 200 ng of MMP-9 (lane 5), 200 ng of MMP-9 and 0.5 µg/ml endostatin (lane 6), 200 ng of MMP-9 and 5 µg/ml endostatin (lane 7), and 12.5 ng MMP-9 (lane 8) were detected with a polyclonal MMP-9 antibody. Direct elution of proteins into SDS-PAGE sample loading buffer results in disruption the antibody-protein complex. Mouse IgG light and heavy chain polypeptides, 25 and 55 kDa in size, respectively, can be seen on Western blot when goat anti-mouse secondary antibody is used for detection (left panel).

 

The Effect of Endostatin on Recombinant Human Pro-MMP-2—The effect of endostatin on recombinant pro-MMP-2 was identified with Western immunoblotting. Endostatin reduced the fragmentation of APMA-treated 72-kDa pro-MMP-2 to ~40-kDa species, but it did not affect the conversion to the 62-kDa form (Fig. 5A). Endostatin seemed to have no effect on the degree of activation or on the level of the 72-kDa pro-MMP-2 in zymography, although the amount of pro-MMP-2 is very low and barely detectable (Fig. 1A). A gelatin degradation assay with MMP-2 purified from human gingival fibroblast cell media showed that endostatin inhibited APMA activation of MMP-2 (Fig. 5B) and in high concentrations also reduced the MMP-2 activity in the absence of APMA (Fig. 5C). Gelatin degradation was decreased from 100 to 82% (S.D., ±5.3%), 60% (S.D., ±13.8%), 41% (S.D., ±5.5%), and 47% (S.D., ±16.8%) in the presence of APMA and from 100 to 95% (S.D., ±5.4%), 97% (S.D., ±21.3%), 63% (S.D., ±15.5%), and 74% (S.D., ±16.6%) in the absence of APMA with endostatin concentrations of 0, 0.05, 5, 10, and 20 µg, respectively (Fig. 5, B and C).



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FIG. 5.
The effect of endostatin on MMP-2. Recombinant human pro-MMP-2 (50 ng) was incubated with recombinant endostatin at 37 °C for 45 min and activated with 1 mM APMA (60 min at 37 °C). Native and processed MMP-2 were detected by Western blotting with monoclonal MMP-2 antibody. A, 50 ng of recombinant pro-MMP-2 (1st lane), MMP-2 treated with APMA (2nd lane), MMP-2 treated with APMA and 120 ng of endostatin (3rd lane), MMP-2 with APMA and 360 ng of endostatin (4th lane), MMP-2 with APMA and 600 ng of endostatin (5th lane), and MMP-2 with APMA and 960 ng of endostatin (6th lane). B and C, enzyme activity assay results with MMP-2 purified from cell culture media and recombinant endostatin. 50 ng of pro-MMP-2 was incubated with endostatin (0.05, 5, 10, or 20 µg)for1hat37 °C and after that with (B) or without (C) 1.3 mM APMA for 1 h at 37 °C and finally with 125I-labeled gelatin for 1 h at 37 °C. The degraded gelatin reflecting the MMP-2 activity was counted with a {gamma}-counter. The results are from 3 separate experiments and represent mean ± S.D.

 

The Effect of Endostatin on Human Interstitial Collagenases MMP-8 and -13—The activities of MMP-8 and -13 were studied by assaying type I collagen degradation with SDS-PAGE. Recombinant endostatin (20:1 molar ratio to MMP-13) inhibited the MMP-13-mediated formation of {alpha}1A, {alpha}2A, {alpha}1B, and {alpha}2B degradation products from native type I collagen {alpha}1 and {alpha}2 chains, when MMP-13 was treated with APMA (Fig. 6A, lanes 2 and 4). However, endostatin had no effect on MMP-8 (20:1 molar ratio)-mediated type I collagen degradation with or without APMA (Fig. 6B).



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FIG. 6.
The effect of endostatin on collagenases MMP-13 and -8. 50 ng of recombinant MMP-13 or MMP-8 was incubated with 600 ng of recombinant endostatin (45 min at 37 °C) in the presence or absence of 1 mM APMA (60 min at 37 °C) following an incubation with 1.5 µM native type I collagen (5 h at room temperature). The cleavage products resulting from catalytic collagenase action on native collagen I were separated by 12% SDS-PAGE and stained with Coomassie Blue. A, control, 1.5 µM type I collagen without collagenase treatment (lane 1), collagen I incubated with MMP-13 (lane 2), collagen I incubated with MMP-13 and APMA (lane 3), collagen I with MMP-13 and endostatin (lane 4), and collagen I with MMP-13, endostatin and APMA (lane 5). B, control, 1.5 µM type I collagen without collagenase treatment (lane 1), collagen I incubated with MMP-8 (lane 2), collagen I incubated with MMP-8 and APMA (lane 3), collagen I with MMP-8 and endostatin (lane 4), and collagen I with MMP-8, endostatin, and APMA (lane 5). C, control. {alpha}1 and {alpha}2 indicate intact native type I collagen monomer. {alpha}1A, {alpha}2A, {alpha}1B, and {alpha}2B indicate the characteristic 3/4 and 1/4 cleavage products resulting from the action of catalytically competent collagenases on native type I collagen.

 

Endostatin Inhibits Human Tumor Cell Migration in Vitro— The effect of endostatin on migration of human oral squamous cell carcinoma HSC-3 cells was studied using an in vitro Transwell assay, which measures random cell migration. Endostatin concentrations of 0–20 µg/ml prevented migration of HSC-3 cells in a dose-dependent manner. Migration was 89% (S.D. ±3.4%), 80% (S.D., ±3.0%), and 55% (S.D., ±2.1) of controls (100%) with endostatin concentrations of 3, 5, and 20 µg/ml, respectively. With the highest concentration of endostatin (40 µg/ml), the inhibitory effect on migration was partially lost (66%, S.D., ±8.6%) (Fig. 7).



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FIG. 7.
Inhibition of tumor cell migration by endostatin in vitro. HSC-3 cells were plated on Transwell filters in the presence of different concentrations of recombinant endostatin (0, 3, 5, 20, and 40 µg/ml) and allowed to migrate for 16–24 h. Cells that traversed to the underside of the filter were stained and counted microscopically. Data represent mean ± S.D.; n = 6.

 

Endostatin Inhibits Carcinoma Cell Intravasation in the CAM Model—Because intravasation, when tumor cells invade through blood vessel walls into the bloodstream, is the key step of the carcinoma process leading to metastasis, we studied the effect of recombinant endostatin to the intravasative capacity of the HCS-3 carcinoma cells using the in vivo CAM assay. Intravasation decreased in a dose-dependent manner to half with low concentrations of endostatin (0.5 and 5 µg/CAM), but with higher concentrations of endostatin (20 and 50 µg/CAM) the inhibitory effect on intravasation efficiency was partially lost, and dose dependency was not observed any longer (Fig. 8, A and B). The intravasation efficiencies were 66% (S.D., ±12), 55% (S.D., ±16), 63% (S.D., ±18), and 71% (S.D., ±18) compared with untreated controls with endostatin concentrations of 0.5, 5, 20, and 50 µg/CAM, respectively (Fig. 8B).



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FIG. 8.
Biphasic effect of endostatin on the HSC-3 carcinoma cell intravasation in the CAM model. A, 2 x 106 HSC3 cells with different amounts of recombinant endostatin (0, 0.5, 5, 20, and 50 µg/CAM) were inoculated in duplicate on upper CAMs of 10-day-old chicken embryos. 50 h after inoculation, the human DNA content in the lower CAM was determined using radioactive human-specific Alu-PCR. The resulting 220-bp PCR band intensity and thus the number of intravasated cells decreased in the presence of endostatin compared with untreated controls. B, the CAM assay results were quantitated with ScionImage software. Alu PCR bands from four experiments were scanned, and the results were expressed as percentage of change in band intensity compared with untreated control band intensities.

 


    DISCUSSION
 TOP
 ABSTRACT
 INTRODUCTION
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 REFERENCES
 
The present study demonstrates that endostatin inhibited the activation of native and recombinant pro-MMP-9 and recombinant pro-MMP-13 but did not affect pro-MMP-8. Endostatin further reduced the formation of the ~40-kDa activation products and gelatinase activity of MMP-2. By immunoprecipitation we demonstrated that endostatin directly binds to MMP-9. We also showed that endostatin decreased carcinoma cell migration in vitro and in vivo using the CAM model. These results may point to a role for endostatin in intravasation, the first step of invasion, when cancer cells invade through blood vessel walls into the bloodstream.

It was quite surprising to find that endostatin had no detectable effect on the conversion of the 72-kDa pro-MMP-2 to the 62-kDa active form in HUV-EC-C endothelial cells (not shown) or HSC-3 carcinoma cells when analyzed by gelatin zymography. However, the previous reports on the effects of endostatin on pro-MMP-2 activation have been somewhat conflicting. Kim et al. (22) reported that endostatin inhibited the activation and activity of pro-MMP-2 in endothelial cells. There is also a study reporting no changes in MMP-2 activation events in response to endostatin treatment (38). With Western blotting we showed the formation of two ~40-kDa fragmentation products in the presence of endostatin and APMA, and endostatin reduced the formation of the smaller one of these fragments in a dose-dependent manner. Furthermore, also in zymography we showed that endostatin inhibited the formation of different ~40-kDa activation products of MMP-2. This endostatin-associated molecular processing of pro-MMP-2 correlated with the reduction in the activity and activation of MMP-2 in the catalytic gelatin degradation assay. It has been demonstrated previously that during the autoactivation of MMP-2, multiple smaller activation products are formed in addition to the 62-kDa active enzyme (39), and our fragments probably correspond to these 38–43-kDa MMP-2 autoactivation products. Furthermore, pro-MMP-2 lacking the hinge region and hemopexin-like domain has a molecular size of 48 kDa, which is converted to the 42-kDa form upon APMA activation. The 42-kDa form has the same kind of activity as the native 62-kDa active form (23). Therefore, our results seem to support the report by Kim et al. (22), indicating an inhibitory effect of endostatin on MMP-2.

Endostatin affected differently the two interstitial collagenases, MMP-8 and -13. It inhibited the APMA-mediated collagen I-digesting activity of MMP-13 but did not affect MMP-8 activity. Endostatin and MMPs seem to form a complex network with respect of regulation with each other. In this regard, we and others (24)2 have recently shown that certain but not all MMPs degrade endostatin-containing fragments from type XVIII collagen that possess the activity of native endostatin.2 Noteworthy, it seems that most of the MMPs that were able to digest endostatin-containing fragments appeared to be the same that were also inhibited by endostatin, suggesting product inhibition in regulation of these MMP activities. As observed here, MMP-9 and -13 both can generate endostatin-containing fragments, and their activities are inhibited by endostatin. MMP-8 does not cleave endostatin-containing fragments from collagen XVIII, and its activity is unaffected by endostatin. Although according to our previous findings MMP-2 does not cleave endostatin-containing fragments,2 another study (24) has reported that MMP-2 is able to generate endostatin-containing fragments after prolonged incubation, although much less efficiently than the other MMPs. In this study, an endostatin-induced effect on the activation and activity of purified MMP-2 as well as fragmentation of recombinant MMP-2 and MMP-2 secreted by oral HSC-3 cells was observed. Surprisingly, no such effects on the formation of the active 62-kDa form of MMP-2 could be seen. However, also the smaller ~40-kDa fragments of MMP-2 possess proteolytic activity. Together, this and previous studies indicate that MMP-2 and endostatin may have an interactive relationship similar to other MMPs, but the effects in both directions are weaker. Further studies specifically directed to evaluation of MMP-2-endostatin interactions must be conducted before final conclusions can be drawn in this matter.

TAT-2, a matrix-degrading extrapancreatic tumor-associated matrix serine proteinase, is produced by many cells, especially by various malignant human cell lines, and it is a natural and efficient activator of MMP-9 (1921). The amount of TAT-2 correlates with the metastatic potential of tumors (31), and TAT-2 has been suggested to participate in tumor growth and metastasis (21, 40). Therefore, it was very interesting that we could show that not only APMA-mediated activation but also naturally occurring TAT-2 activation of MMP-9 could be inhibited by endostatin, and the inhibitory effect was even more efficient with TAT-2 activation than with APMA activation.

The concentrations of endostatin, which significantly inhibited the APMA activation of pro-MMP-9 in zymography (12.5–40 µg/ml), were quite high compared with previous studies, where clear inhibitory effects on MMP-2 activation could be observed already at endostatin concentrations of 3 µg/ml (22). However, in the Western immunoblot (Fig. 2), the concentrations of endostatin were lower and in the same scale as shown previously (22) and showing markedly reduced MMP-9 activation already with 360 ng of endostatin. There might also be some substances in the HSC-3 cell culture conditions, not present in human umbilical vein endothelial cell culture (22) or in in vivo studies with recombinant MMP-9, that interfered with the actions of endostatin. The endostatin concentrations affecting MMP activation were higher than the physiological concentrations of endostatin. It has been shown with collagen XVIII null mice and wild type mice that there is no difference in the growth rates of fibrosarcomas and melanomas, suggesting that physiological levels of endostatin (40–100 ng/ml in serum) are insufficient to exert anti-tumor effects in mice (41). Based on the observations, it has also been suggested that the antitumor and anti-angiogenic effects of endostatin might be pharmacological effects at high doses (42).

In addition to MMPs, endostatin has been shown to interfere with the actions of other proteases. In the plasminogen activator system, endostatin decreases the levels and modulates the distribution of urokinase-type plasminogen activator and plasminogen activator inhibitor, type 1, in cultured human endothelial cells. Furthermore, endostatin treatment caused redistribution of receptor-bound urokinase-type plasminogen activator from focal contacts and a rapid disassembly of focal adhesions (38).

Previous reports have suggested that endostatin can directly interact with pro-MMP-2 (22) or the catalytic domain of active MMP-2 (23). Since MMPs share a highly homologous catalytic domain (43), binding of endostatin solely to this region would result in inactivation of all MMPs. In our studies endostatin inhibited the activity of MMP-9 and -13 and partially MMP-2, but not MMP-8, suggesting that additional interactions are required to accomplish the inhibitory effect of endostatin on certain MMPs. We examined the possibility of a direct interaction between endostatin and pro-MMP-9 by immunocoprecipitation, and we found that they indeed form a complex. This direct binding of endostatin and MMP-9 implies that such an interaction can be important for the prevention of pro-MMP-9 activation. MMP-9 seemed to also bind slightly to the anti-His antibody even without endostatin, most likely because of a group of histidine residues in the MMP-9 sequence. Endostatin has been suggested to interact also with some other extracellular matrix components and cell membrane receptors. Rehn et al. (25) showed that recombinant human endostatin interacts with {alpha}5 and {alpha}V integrins on the surface of human endothelial cells. Furthermore, the endostatin-integrin interaction is of functional significance in vitro, as immobilized endostatin promotes and soluble endostatin inhibits integrin-dependent endothelial cell functions (25). HSC-3 cells express {alpha}V{beta}6 integrins on their surface, and the blocking of this integrin inhibited the invasive growth of HSC-3 cells (44). Endostatin also binds to heparin (45) and with low affinity to the heparan sulfate proteoglycans glypican-1 and -4 and with high affinity to an as yet unidentified molecule on endothelial cells (46).

Previous studies have examined the capability of endostatin to regulate biological events induced by treatment of cells with various growth factors. In many cases endostatin alone had no effect, and its action was detectable only in the presence of certain growth factors. Endostatin inhibits vascular endothelial growth factor- and basic fibroblast growth factor-induced cell migration (47, 48). Endostatin increased apoptosis only in the presence of fibroblast growth factor-2 but not alone (45). Thus, in quiescent cells the effects of endostatin can be very different from the effects in tumors where increased amounts of different growth factors are present. In addition, we found that the cell density seemed to be quite critical for the inhibitory effect of endostatin, as densities lower (18,000 cells/well) or higher (38,000 cells/well) than 28,000 HSC-3 cells per well (in 24-well plates) did not respond to endostatin treatment in the same manner with MMP-9 inhibition. Exponentially growing cells produce different amounts of various growth factors and receptors than confluent cells and that might be a reason why HSC-3 cells at different densities and thus in different phases of growth do not respond to endostatin treatment in the same way. In endothelial cells the same phenomenon is observed; endostatin has no effect on endothelial cell proliferation when the endothelial cells are almost confluent due to the contact inhibition of the cells (49). On the other hand, squamous cell carcinoma cell growth is not contact-inhibited (50). Further studies are needed in the future to clarify the reason why endostatin works better in certain cell densities.

The anti-angiogenic effects of endostatin appear to involve inhibition of cell growth mediated via both reduced G1/S phase transition and increased apoptosis (45, 51). The mechanism of the endothelial G1 arrest involves inhibition of cyclin D1 RNA and protein expression. This suppression is mediated by transcriptional inhibition through the lymphoid enhancer factor-1 site in the cyclin D1 promoter (10). Endostatin also rapidly down-regulates many other genes in growing endothelial cells, including immediate early response genes, cell cycle-related genes, and genes regulating apoptosis inhibitors, mitogen-activated protein kinases, focal adhesion kinases, G-protein-coupled receptors mediating endothelial growth, a mitogenic factor, adhesion molecules, and cell structure components (52). In our studies endostatin did not affect the total level of gelatinase production as observed in zymography. We are currently studying the effect of endostatin on regulation of genes in human oral carcinoma cells.

In this study, endostatin seemed to have a biphasic effect on intravasation and migration. Lower concentrations of endostatin inhibited intravasation and migration in a dose-dependent manner, but high endostatin concentrations were not as efficient. We also observed the same phenomenon in the catalytic gelatin degradation activity assay for MMP-2, in which lower concentrations of endostatin inhibited MMP-2 more effectively than higher ones. One possible reason for these observations is the fact that endostatin exists in two forms, as a monomer or as an NC1 trimer, that exert different or even opposite effects (53, 54). For example, trimeric endostatin in the NC1 domain of collagen XVIII is needed for cell migration, but monomeric endostatin inhibits the migratory activity (53). We consider it possible that with high endostatin concentrations, aggregates may occur resulting in different effects on migration and intravasation. In fact, it has been shown recently that endostatin indeed exists in two forms as follows: as a soluble globular form and as an insoluble form with abundant cross-{beta}-sheets aggregating into amyloid deposits. These different endostatin conformations have distinct effects on at least tissue-type plasminogen activator-mediated plasminogen activation. Insoluble endostatin stimulates plasminogen activation, but globular endostatin has no effect (55). Another explanation for such a biphasic effect might be negative feedback (8).

In conclusion, we show here that endostatin affects tumor cell migration, intravasation, and invasion. Our data suggest that the antitumor effect of endostatin is, at least in part, associated with the ability of endostatin to inhibit the activation and activity of certain MMPs, especially MMP-9 and MMP-13, and to some extent also MMP-2 but not MMP-8.


    FOOTNOTES
 
* This work was supported by Oulu University Hospital KEVO grants, the Cancer Society of Finland, the Cancer Society of Northern Finland, Helsinki University Research Funds, the Wilhelm and Else Stockman Foundation, the Helsinki University Central Hospital-EVO Grant TI020Y0002, Finnish Dental Society Apollonia, European Commission Grant QLK3-2000-00084, and by Grant 44843 from the Finnish Centre of Excellence Program (2000-2005) of the Academy of Finland. 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. Back

{ddagger}{ddagger} To whom correspondence should be addressed: Dept. of Diagnostics and Oral Medicine, Institute of Dentistry, University of Oulu, P. O. Box 5281, FIN-90014 Oulu, Finland. Tel.: 358-8-537-5592; Fax: 358-8-537-5560; E-mail: tuula.salo{at}oulu.fi.

1 The abbreviations used are: NC1, carboxyl-terminal noncollagenous domain 1; MMP, matrix metalloproteinase; APMA, p-aminophenylmercuric acetate; TAT-2, tumor-associated trypsinogen-2; HSC-3, human tongue squamous cell carcinoma; TATI, tumor-associated trypsinogen inhibitor; PBS, phosphate-buffered saline; CAM, chorioallantoic membrane; BSA, bovine serum albumin; MSP, matrix serine proteinase; PAI, plasminogen activator inhibitor. Back

2 R. Heljasvaara, J. Luostarinen, P. Nyberg, M. Parikka, M. Rehn, T. Sorsa, T. Salo, and T. Pihlajaniemi, submitted for publication. Back


    ACKNOWLEDGMENTS
 
We thank Maija-Leena Lehtonen, Sirpa Kangas, and Aila White for their excellent technical assistance and professor Juha Risteli for 125I-labeled gelatin.



    REFERENCES
 TOP
 ABSTRACT
 INTRODUCTION
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 REFERENCES
 

  1. Folkman, J. (1995) Nat. Med. 1, 27–31[Medline] [Order article via Infotrieve]
  2. Hanahan, D., and Folkman, J. (1996) Cell 86, 353–364[Medline] [Order article via Infotrieve]
  3. Cao, Y. (2001) Int. J. Biochem. Cell Biol. 33, 357–369[CrossRef][Medline] [Order article via Infotrieve]
  4. O'Reilly, M. S., Boehm, T., Shing, Y., Fukai, N., Vasios, G., Lane, W. S., Flynn, E., Birkhead, J. R., Olsen, B. R., and Folkman, J. (1997) Cell 88, 277–285[Medline] [Order article via Infotrieve]
  5. Rehn, M., Hintikka, E., and Pihlajaniemi, T. (1994) J. Biol. Chem. 269, 13929–13935[Abstract/Free Full Text]
  6. Oh, S. P., Kamagata, Y., Muragaki, Y., Timmons, S., Ooshima, A., and Olsen, B. R. (1994) Proc. Natl. Acad. Sci. U. S. A. 91, 4229–4233[Abstract]
  7. Boehm, T., Folkman, J., Browder, T., and O'Reilly, M. S. (1997) Nature 390, 404–407[CrossRef][Medline] [Order article via Infotrieve]
  8. Marneros, A. G., and Olsen, B. R. (2001) Matrix Biol. 20, 337–345[CrossRef][Medline] [Order article via Infotrieve]
  9. Dixelius, J., Cross, M., Matsumoto, T., Sasaki, T., Timpl, R., and Claesson-Welsh, L. (2002) Cancer Res. 62, 1944–1947[Abstract/Free Full Text]
  10. Hanai, J., Dhanabal, M., Karumanchi, S. A., Albanese, C., Waterman, M., Chan, B., Ramchandran, R., Pestell, R., and Sukhatme, V. P. (2002) J. Biol. Chem. 277, 16464–16469[Abstract/Free Full Text]
  11. Kim, Y.-M., Hwang, S., Kim, Y.-M., Pyun, B., Kim, T.-Y., Lee, S.-T., Gho, Y. S., and Kwon, Y.-G. (2002) J. Biol. Chem. 277, 27872–27879[Abstract/Free Full Text]
  12. Chambers, A. F., and Matrisian, L. M. (1997) J. Natl. Cancer Inst. 87, 1260–1270[CrossRef]
  13. Coussens, L. M., and Werb, Z. (1996) Chem. Biol. 3, 896–904
  14. Stetler-Stevenson, W. G. (1999) J. Clin. Invest. 103, 1237–1241[Free Full Text]
  15. Juarez, J., Clayman, G., Nakajima, M., Tanabe, K. K., Saya, H., Nicolson, G. L., and Boyd, D. (1993) Int. J. Cancer 55, 10–18[Medline] [Order article via Infotrieve]
  16. Kawamata, H., Uchida, D., Hamano, H., Kimura-Yanagawa, T., Nakashiro, K. I., Hino, S., Omotehara, F., Yoshida, H., and Sato, M. (1998) Int. J. Oncol. 13, 699–704[Medline] [Order article via Infotrieve]
  17. Freije, J. M., Diez-Itza, I., Balbin, M., Sanchez, L. M., Blasco, R., Tolivia, J., and Lopez-Otin, C. (1994) J. Biol. Chem. 269, 16766–16773[Abstract/Free Full Text]
  18. Stricklin, G. P., Jeffrey, J. J., Roswit, W. T., and Eisen, A. Z. (1983) Biochemistry 22, 61–68[Medline] [Order article via Infotrieve]
  19. Sorsa, T., Salo, T., Koivunen, E., Tyynelä, J., Konttinen, Y. T., Bergmann, U., Tuuttila, A., Niemi, E., Teronen, O., Heikkilä, P., Tschesche, H., Leinonen, J., Osman, S., and Stenman, U. H. (1997) J. Biol. Chem. 272, 21067–21074[Abstract/Free Full Text]
  20. Paju, A., Sorsa, T., Tervahartiala, T., Koivunen, E., Haglund, C., Leminen, A., Wahlström, T., Salo, T., and Stenman, U. H. (2001) Br. J. Cancer 84, 1363–1371[CrossRef][Medline] [Order article via Infotrieve]
  21. Nyberg, P., Moilanen, M., Paju, A., Sarin, A., Stenman, U. H., Sorsa, T., and Salo, T. (2002) J. Dent. Res. 81, 831–835[Abstract/Free Full Text]
  22. Kim, Y.-M., Jang, J.-W., Lee, O.-K., Yeon, J., Choi, E.-Y., Kim, K.-W., Lee, S.-T., and Kwon, Y.-G. (2000) Cancer Res. 60, 5410–5413[Abstract/Free Full Text]
  23. Lee, S.-L., Jang, J.-W., Kim, Y.-M., Lee, H. I., Jeon, J. Y., Kwon, Y.-G., and Lee, S.-T. (2002) FEBS Lett. 519, 147–152[CrossRef][Medline] [Order article via Infotrieve]
  24. Ferreras, M., Felbor, U., Lenhard, T., Olsen, B. R., and Delaisse, J.-K. (2000) FEBS Lett. 486, 247–251[CrossRef][Medline] [Order article via Infotrieve]
  25. Rehn, M., Veikkola, T., Kukk-Valdre, E., Nakamura, H., Ilmonen, M., Lombardo, C., Pihlajaniemi, T., Alitalo, K., and Vuori, K. (2001) Proc. Natl. Acad. Sci. U. S. A. 98, 1024–1029[Abstract/Free Full Text]
  26. O'Grady, R. L., Nethery, A., and Hunter, N. (1984) Anal. Biochem. 140, 490–494[Medline] [Order article via Infotrieve]
  27. Risteli, L., and Risteli, J. (1987) Methods Enzymol. 145, 391–411[Medline] [Order article via Infotrieve]
  28. Mäkelä, M., Salo, T., Uitto, V. J., and Larjava, H. (1994) J. Dent. Res. 73, 1397–1406[Abstract]
  29. Koivunen, E., Huhtala, M. L., and Stenman, U. H. (1989) J. Biol. Chem. 264, 14095–14099[Abstract/Free Full Text]
  30. Koivunen, E., Saksela, O., Itkonen, O., Osman, S., Huhtala, M. L., and Stenman, U. H. (1991) Int. J. Cancer 47, 592–596[Medline] [Order article via Infotrieve]
  31. Koivunen, E., Itkonen, O., Halila, H., and Stenman, U. H. (1990) Cancer Res. 50, 2375–2378[Abstract]
  32. Koivunen, E., Ristimäki, A., Itkonen, O., Osman, S., Vuento, M., and Stenman, U. H. (1991) Cancer Res. 51, 2107–2112[Abstract]
  33. Kjeldsen, L., Johnsen, A. H., Sengelov, H., and Borregaard, N. (1993) J. Biol. Chem. 268, 10425–10432[Abstract/Free Full Text]
  34. Sorsa, T., Ding, Y.-L., Salo, T., Lauhio, A., Teronen, O., Ingman, T., Ohtani, H., Andoh, N., Takeha, S., and Konttinen, Y. T. (1994) Ann. N. Y. Acad. Sci. 732, 112–131[Abstract]
  35. Koivunen, E., Arap, W., Valtanen, H., Raininsalo, A., Penate Medina, O., Heikkilä, P., Kantor, C., Gahmberg, C. G., Salo, T., Konttinen, Y. T., Sorsa, T., Ruoslahti, E., and Pasqualini, R. (1999) Nat. Biotechnol. 17, 768–774[CrossRef][Medline] [Order article via Infotrieve]
  36. Heikkilä, P., Teronen, O., Moilanen, M., Konttinen, Y. T., Hanemaaijer, R., Laitinen, M., Maisi, P., Pluijm, G., Bartlet, T. J., Salo, T., and Sorsa, T. (2002) Anti-Cancer Drugs 13, 245–254[CrossRef][Medline] [Order article via Infotrieve]
  37. Kim, J., Yu, W., Kovalski, K., and Ossowski, L. (1998) Cell 94, 353–362[Medline] [Order article via Infotrieve]
  38. Wickström, S. A., Veikkola, T., Rehn, M., Pihlajaniemi, T., Alitalo, K., and Keski-Oja, J. (2001) Cancer Res. 61, 6511–6516[Abstract/Free Full Text]
  39. Bergmann, U., Tuuttila, A., Stetler-Stevenson, W. G., and Tryggvason, K. (1995) Biochemistry 34, 2819–2825[Medline] [Order article via Infotrieve]
  40. Miyata, S., Miyagi, Y., Koshikawa, N., Nagashima, Y., Kato, Y., Yasumitsu, H., Hirahara, F., Misugi, K., and Miyazaki, K. (1998) Clin. Exp. Metastasis 16, 613–622[CrossRef][Medline] [Order article via Infotrieve]
  41. Fukai, N., Eklund, L., Marneros, A. G., Oh, S. P., Keene, D. R., Tamarkin, L., Niemelä, M., Ilves, M., Li, E., Pihlajaniemi, T., and Olsen, B. R. (2002) EMBO J. 21, 1535–1544[Abstract/Free Full Text]
  42. Olsen, B. R. (2002) Matrix Biol. 21, 309–310[CrossRef][Medline] [Order article via Infotrieve]
  43. Bode, W., Fernandez-Catalan, C., Tschesche, H., Grams, F., Nagase, H., and Maskos, K. (1999) Cell. Mol. Life Sci. 55, 639–652[CrossRef][Medline] [Order article via Infotrieve]
  44. Xue, H., Atakilit, A., Zhu, W., Li, X., Ramos, D. M., and Pytela, R. (2001) Biochem. Biophys. Res. Commun. 288, 610–618[CrossRef][Medline] [Order article via Infotrieve]
  45. Dixelius, J., Larsson, H., Sasaki, T., Holmqvist, K., Lu, L., Engström, Å., Timpl, R., Welsh, M., and Claesson-Welsh, L. (2000) Blood 95, 3403–3411[Abstract/Free Full Text]
  46. Karumanchi, S. A., Jha, V., Ramchandran, R., Karihaloo, A., Tsiokas, L., Chan, B., Dhanabal, M., Hanai, J. I., Venkataraman, G., Shriver, Z., Keiser, N., Kalluri, R., Zeng, H., Mukhopadhyay, D., Chen, R. L., Lander, A. D., Hagihara, K., Yamaguchi, Y., Sasisekharan, R., Cantley, L., and Sukhatme, V. P. (2001) Mol. Cell 7, 811–822[CrossRef][Medline] [Order article via Infotrieve]
  47. Yamaguchi, N., Anand-Apte, B., Lee, M., Sasaki, T., Fukai, N., Shapiro, R., Que, I., Lowik, C., Timpl, R., and Olsen, B. R. (1999) EMBO J. 18, 4414–4423[Abstract/Free Full Text]
  48. Dhanabal, M., Ramchandran, R., Volk, R., Stillman, I. E., Lombardo, M., Iruela-Arispe, M. L., Simons, M., and Sukhatme, V. P. (1999a) Cancer Res. 59, 189–197[Abstract/Free Full Text]
  49. Chavakis, E., and Dimmeler, S. (2002) Arterioscler. Thromb. Vasc. Biol. 22, 887–893[Abstract/Free Full Text]
  50. Croce, M. V., Colussi, A. G., Zambelli, A., Price, M. R., and Segal-Eiras, A. (2001) Int. J. Oncol. 18, 729–735[Medline] [Order article via Infotrieve]
  51. Dhanabal, M., Ramchandran, R., Waterman, M. J., Lu, H., Knebelmann, B., Segal, M., and Sukhatme, V. P. (1999) J. Biol. Chem. 274, 11721–11726[Abstract/Free Full Text]
  52. Shichiri, M., and Hirata, Y. (2001) FASEB J. 15, 1044–1053[Abstract/Free Full Text]
  53. Ackley, B. D., Crew, J. R., Elamaa, H., Pihlajaniemi, T., Kuo, C. J., and Kramer, J. M. (2001) J. Cell Biol. 152, 1219–1232[Abstract/Free Full Text]
  54. Kuo, C. J., LaMontagne, K. R., Jr., Garcia-Gardena, G., Ackley, B. D., Kalman, D., Park, S., Christofferson, R., Kamihara, J., Ding, Y. H., Lo, K. M., Gillies, S., Folkman, J., Mulligan, R. C., and Javaherian, K. (2001) J. Cell Biol. 152, 1233–1246[Abstract/Free Full Text]
  55. Kranenburg, O., Bouma, B., Kroon-Batenburg, L. M. J., Reijerkerk, A., Wu, Y. P., Voest, E. E., and Gebbink, M. F. B. G. (2002) Curr. Biol. 12, 1833–1839[CrossRef][Medline] [Order article via Infotrieve]