Suppression of Urokinase Expression and Invasiveness by Urinary Trypsin Inhibitor Is Mediated through Inhibition of Protein Kinase C- and MEK/ERK/c-Jun-dependent Signaling Pathways
2001; Elsevier BV; Volume: 276; Issue: 3 Linguagem: Inglês
10.1074/jbc.m007650200
ISSN1083-351X
AutoresHiroshi Kobayashi, Mika Suzuki, Yoshiko Tanaka, Yasuyuki Hirashima, Toshihiko Terao,
Tópico(s)Enzyme Production and Characterization
ResumoUrinary trypsin inhibitor (UTI), a Kunitz-type protease inhibitor, interacts with cells as a negative modulator of the invasive cells. Human ovarian cancer cell line, HRA, was treated with phorbol ester (PMA) to evaluate the effect on expression of urokinase-type plasminogen activator (uPA), since the action of uPA has been implicated in matrix degradation and cell motility. Preincubation of the cells with UTI reduced the ability of PMA to trigger the uPA expression at the gene level and at the protein level. UTI-induced down-regulation of PMA-stimulated uPA expression is irreversible and is independent of a cytotoxic effect. Down-regulation of uPA by UTI is mediated by its binding to the cells. We next asked whether the mechanism of inhibition of uPA expression by UTI was due to interference with the protein kinase C second messenger system. An assay for PKC activity demonstrated that UTI does not directly inhibit the catalytic activity of PKC and that PMA translocation of PKC from cytosol to membrane was inhibited by UTI, indicating that UTI inhibits the activation cascade of PKC. PMA could also activate a signaling pathway involving MEK1/ERK2/c-Jun-dependent uPA expression. When cells were preincubated with UTI, we could detect suppression of phosphorylation of these proteins. Like several types of PKC inhibitor, UTI inhibited PMA-stimulated invasiveness. We conclude that UTI markedly suppresses the cell motility possibly through negative regulation of PKC- and MEK/ERK/c-Jun-dependent mechanisms, and that these changes in behavior are correlated with a coordinated down-regulation of uPA which is likely to contribute to the cell invasion processes. Urinary trypsin inhibitor (UTI), a Kunitz-type protease inhibitor, interacts with cells as a negative modulator of the invasive cells. Human ovarian cancer cell line, HRA, was treated with phorbol ester (PMA) to evaluate the effect on expression of urokinase-type plasminogen activator (uPA), since the action of uPA has been implicated in matrix degradation and cell motility. Preincubation of the cells with UTI reduced the ability of PMA to trigger the uPA expression at the gene level and at the protein level. UTI-induced down-regulation of PMA-stimulated uPA expression is irreversible and is independent of a cytotoxic effect. Down-regulation of uPA by UTI is mediated by its binding to the cells. We next asked whether the mechanism of inhibition of uPA expression by UTI was due to interference with the protein kinase C second messenger system. An assay for PKC activity demonstrated that UTI does not directly inhibit the catalytic activity of PKC and that PMA translocation of PKC from cytosol to membrane was inhibited by UTI, indicating that UTI inhibits the activation cascade of PKC. PMA could also activate a signaling pathway involving MEK1/ERK2/c-Jun-dependent uPA expression. When cells were preincubated with UTI, we could detect suppression of phosphorylation of these proteins. Like several types of PKC inhibitor, UTI inhibited PMA-stimulated invasiveness. We conclude that UTI markedly suppresses the cell motility possibly through negative regulation of PKC- and MEK/ERK/c-Jun-dependent mechanisms, and that these changes in behavior are correlated with a coordinated down-regulation of uPA which is likely to contribute to the cell invasion processes. urinary trypsin inhibitor a carboxyl-terminal domain of UTI phorbol myristate acetate urokinase-type plasminogen activator urinary trypsin inhibitor-binding protein protein kinase C plasminogen activator mitogen-activated protein mitogen-activated protein kinase extracellular signal-regulated kinase A number of studies have indicated that exogenously applied UTI,1 also known as bikunin, to tumor cells could suppress their invasiveness and metastatic formation in an in vitro assay system and in an in vivo animal model (1Kobayashi H. Shinohara H. Takeuchi K. Itoh M. Fujie M. Saitoh M. Terao T. Cancer Res. 1994; 54: 844-849PubMed Google Scholar, 2Kobayashi H. Shinohara H. Ohi H. Sugimura M. Terao T. Fujie M. Clin. Exp. Metast. 1994; 12: 117-128Crossref PubMed Scopus (37) Google Scholar, 3Kobayashi H. Fujie M. Shinohara H. Ohi H. Sugimura M. Terao T. Int. J. Cancer. 1994; 57: 378-384Crossref PubMed Scopus (48) Google Scholar, 4Kobayashi H. Gotoh J. Shinohara H. Moniwa N. Terao T. Thromb. Haemost. 1994; 71: 474-480Crossref PubMed Scopus (64) Google Scholar, 5Kobayashi H. Gotoh J. Hirashima Y. Fujie M. Sugino D. Terao T. J. Biol. Chem. 1995; 270: 8361-8366Abstract Full Text Full Text PDF PubMed Scopus (73) Google Scholar, 6Kobayashi H. Gotoh J. Kanayama N. Hirashima Y. Terao T. Sugino D. Cancer Res. 1995; 55: 1847-1852PubMed Google Scholar, 7Kobayashi H. Shinohara H. Gotoh J. Fujie M. Fujishiro S. Terao T. Br. J. Cancer. 1995; 72: 1131-1137Crossref PubMed Scopus (52) Google Scholar, 8Kobayashi H. Shinohara H. Fujie M. Gotoh J. Itoh M. Takeuchi K. Terao T. Int. J. Cancer. 1995; 63: 455-462Crossref PubMed Scopus (45) Google Scholar, 9Kobayashi H. Sugino D. She M.Y. Ohi H. Hirashima Y. Shinohara H. Fujie M. Shibata K. Terao T. Eur. J. Biochem. 1998; 253: 817-826Crossref PubMed Scopus (66) Google Scholar). The UTI gene encodes a Kunitz-type protease inhibitor of molecular mass 40 kDa (9Kobayashi H. Sugino D. She M.Y. Ohi H. Hirashima Y. Shinohara H. Fujie M. Shibata K. Terao T. Eur. J. Biochem. 1998; 253: 817-826Crossref PubMed Scopus (66) Google Scholar), which is composed of a ligand-binding domain (amino terminus) for cell-associated UTI-binding sites (10Kobayashi H. Gotoh J. Fujie M. Terao T. J. Biol. Chem. 1994; 269: 20642-20647Abstract Full Text PDF PubMed Google Scholar, 11Kobayashi H. Hirashima Y. Sun G.W. Fujie M. Shibata K. Tamotsu S. Miura K. Sugino D. Tanaka Y. Kondo S. Terao T. Biochim. Biophys. Acta. 1998; 1383: 253-268Crossref PubMed Scopus (27) Google Scholar) and protease inhibitor domain (carboxyl terminus) (9Kobayashi H. Sugino D. She M.Y. Ohi H. Hirashima Y. Shinohara H. Fujie M. Shibata K. Terao T. Eur. J. Biochem. 1998; 253: 817-826Crossref PubMed Scopus (66) Google Scholar), which effectively inhibits trypsin, plasmin, and granulocyte elastase. In addition to its protease inhibiting effects, UTI plays a role in suppressing urokinase-type plasminogen activator (uPA) production responsible for the invasiveness of tumor cells (12Kobayashi H. Gotoh J. Terao T. Eur. J. Cell Biol. 1996; 71: 380-386PubMed Google Scholar), although UTI does not inhibit directly the catalytic activity of uPA. uPA converts plasminogen into plasmin, a serine protease with broad substrate specificity toward components of the basement membrane and the extracellular matrix including laminin, vitronectin, and fibronectin (13Dano K. Andreasen P.A. Grondahl-Hansen J. Kristensen P. Nielsen L.S. Skriver L. Adv. Cancer Res. 1985; 44: 139-266Crossref PubMed Scopus (2306) Google Scholar). These proteolytic functions facilitate the migration of tumor cells through the extracellular matrix and basement membrane barriers. Therefore, UTI apparently plays a key role in regulation of cell invasiveness and metastatic formation possibly through down-regulation of uPA expression. Expression of uPA is controlled by a variety of extracellular signals such as phorbol ester, protein kinase C (PKC), and Fos/Jun-dependent signals, cAMP, cytoskeletal reorganization, tumor necrosis factor-α, interleukin-1β, interferon-γ, tumor growth factor-β, fibroblast growth factor-2, okadaic acid, retinoic acid, UV, and oncogene products v-Src and v-Ras (14Nagamine Y. Lee J.S. Menoud P-A. Nanbu R. Glas-Grenwalt P. Fibrinolysis in Disease. CRC Press, Boca Raton, FL1995Google Scholar). uPA activity of malignant cells is induced during the promotion stage of the carcinogenic process and phorbol myristate acetate (PMA) is one of the best characterized, tumor promoting agent. PMA is generally recognized to modulate cellular functions by activating a Ca2+-phospholipid-dependent PKC (15Castagna M. Takai Y. Kaibuchi K. Sano K. Kikkawa U. Nishizuka Y. J. Biol. Chem. 1982; 257: 7847-7851Abstract Full Text PDF PubMed Google Scholar). Agents that modulate uPA have been shown to alter the rate of metastasis in in vitro experiments and in some animal models (16Ways D.K. Kukoly C.A. deVente J. Hooker J.L. Bryant W.O. Posekany K.J. Fletcher D.J. Cook P.P. Parker P.J. J. Clin. Invest. 1995; 95: 1906-1915Crossref PubMed Scopus (269) Google Scholar). Recent publication demonstrated that UTI with anti-inflammatory and anti-tumor promoting properties can influence the PKC-dependent signal pathway in uPA expression in cultured human umbilical vein endothelial cells and in the promyeloid leukemia cell line U937 (12Kobayashi H. Gotoh J. Terao T. Eur. J. Cell Biol. 1996; 71: 380-386PubMed Google Scholar); exogenous UTI inhibits a rapid increase in membrane-associated PKC activity, and a decrease in cytosolic PKC activity. However, the precise molecular mechanisms of the UTI-mediated changes occurring downstream of the PKC signal transduction have remained unclear. In the present study, we have sought to define the UTI-dependent regulatory mechanisms involved in PMA-induced uPA expression and cell motility. First, we have determined the effect of UTI on PMA-induced uPA expression, as well as quantitating time- and dose-dependent alterations in the steady state levels ofuPA mRNA and uPA activity. Second, we ask whether the inhibition was due to interference with the PKC second messenger system. For this, we have compared the effect of UTI and several types of PKC inhibitor on PMA-induced uPA expression, PKC translocation, and signal pathway involving a relay of phosphorylation of several proteins. Third, we have investigated the possibility that UTI binding to tumor cells might be involved in the down-regulation of uPA expression. Finally, we have determined if there is a relation between UTI-dependent alterations in the uPA expression and cellular motility. UTI was purified to homogeneity from human urine. A highly purified preparation of human UTI was kindly supplied by Mochida Pharmaceutical Co., Tokyo, Japan. The COOH-terminal fragment of UTI (HI-8) was purified as described previously (9Kobayashi H. Sugino D. She M.Y. Ohi H. Hirashima Y. Shinohara H. Fujie M. Shibata K. Terao T. Eur. J. Biochem. 1998; 253: 817-826Crossref PubMed Scopus (66) Google Scholar). Polyclonal antibodies raised against MEK1, ERK2, and c-Jun were obtained from Santa Cruz Biotechnology (Santa Cruz, CA). Monoclonal antibodies raised against human uPA and human high molecular weight uPA were supplied by Yoshitomi Pharmaceutical Co., Ltd. (Osaka, Japan). PMA, calcium ionophore A23187, H-7, calphostin C, staurosporin, aspirin, and 5,8,11,14-eicosatetraenoic acid were purchased from Sigma, and all other chemicals were of reagent grade or better and were purchased from major suppliers. Ethanol was used as the solvent for PMA, and the final concentration of ethanol was 0.1% in all experimental points. Human ovarian cancer cell line HRA was obtained from Dr. Y. Kikuchi (17Kikuchi Y. Kizawa I. Oomori K. Miyauchi M. Kita T. Sugita M. Tenjin Y. Kato K. Cancer Res. 1987; 47: 592-596PubMed Google Scholar). The HRA was cultured in RPMI 1640 with 10% fetal calf serum (Life Technologies, Inc., Rockville, MD). Cells were disaggregated routinely with 0.1% trypsin/EDTA solution and replated at a split ratio of 1:10. The cells were harvested and aliquoted into 12-well tissue culture plates (0.5–1.0 × 106 cells/well) in RPMI 1640 supplemented with penicillin (100 units/ml), streptomycin (100 μg/ml), and 10% fetal calf serum. On the next day, the cells were washed three times with phosphate-buffered saline to remove serum, and the medium was replaced with RPMI 1640 supplemented with antibiotics. Serum-free medium plus the test drugs were added and incubation was continued for different time lapses. The cells were incubated with various concentrations of UTI during three different periods, i.e.(a) during 30 min preceding the stimulation phase, (b) during the PMA-stimulation phase, and (c) 60 min after the stimulation, UTI being then added to the expression medium. Conditioned media were individually harvested, one of the remaining monolayers were trypsinized and hemocytometer cell counting or protein content determinations were performed. The protein concentration was determined by the method of Bradford (18Bradford M.M. Anal. Biochem. 1976; 72: 248-254Crossref PubMed Scopus (218536) Google Scholar) using bovine serum albumin as the standard and reagents purchased from Bio-Rad. Conditioned media were used for measurement of plasminogen activator activity by chromogenic and zymographic analyses. Following the recovery of conditioned media, monolayers were washed and used for determination of cell-associated plasminogen activator activity. Plasminogen activator (PA) activity in the cell-conditioned media (100 μl) was quantitated utilizing a functional assay for plasmin. Medium (100 μl) was then incubated for 3 h in buffer A (phosphate-buffered saline containing plasminogen (0.165 units/ml) and S-2251 (0.5 mm)) as the chromogenic substrate of plasmin. In a parallel experiment, after the cells were washed, the medium was replaced with buffer A and incubated for 5 h to determine the cell associated PA activity. The amount of p-nitroaniline released was determined spectrophotometrically at 405 nm. Each assay was run with a plasminogen-free blank. In some experiments, using log-log graph paper, a calibration curve was drawn by plotting the calibrator values on thex axis and their corresponding absorbance values on they axis. The uPA activities were quantitatively obtained by reading from the calbration curve. Viability of the cells was assessed by measuring lactate dehydrogenase release in the medium as described previously (19Dubois P. Lison D. Lauwerys R. Biochem. Pharmacol. 1988; 37: 2139-2143Crossref PubMed Scopus (9) Google Scholar). Lactate dehydrogenase release is expressed as % of total enzyme content determined after cell disruption with Triton X-100. Zymography was performed as described previously (20Granelli-Piperno A. Reich E. J. Exp. Med. 1978; 148: 223-234Crossref PubMed Scopus (645) Google Scholar). To confirm uPA activity present in zymograms, plasminogen-casein-agarose underlays were prepared with an anticatalytic anti-human uPA antibody or without plasminogen. uPA activity was quantified by a caseinolysis assay (21Saksela O. Anal. Biochem. 1981; 111: 276-282Crossref PubMed Scopus (120) Google Scholar), using plasminogen-rich casein-agarose plates. Secreted uPA activity was referenced to a standard uPA curve and normalized to 106cells and to a 24-h incubation period, unless otherwise indicated. Of note that, in the present study, zymographic analysis only detected an uPA (molecular mass 55 kDa) in HRA cells. Cells (3 × 106) were plated in 100-mm dishes with 10 ml of RPMI 1640 containing 10% fetal calf serum. The next day, the cells were treated as described above. Cells were washed with phosphate-buffered saline and lysed in 500 μl of lysis buffer (20 mm Tris-HCl, pH 7.4, 137 mm NaCl, 2 mm EDTA, 1% Triton X-100, 10% glycerol, 1 mmsodium vanadate, 2 mm sodium pyrophosphate, 1 mm phenylmethylsulfonyl fluoride, 25 mmβ-glycerophosphate, and 1 μg/ml leupeptin). Lysates were centrifuged for 10 min at 15,000 × g, and the protein concentrations were determined. Total protein (20 μg) was diluted in SDS sample buffer, heated for 5 min at 100 °C, and subjected to 10% SDS-polyacrylamide gel electrophoresis. Proteins were blotted onto a polyvinylidene difluoride membrane with the use of a semidry electroblotter apparatus. The blots were finally incubated with ECL chemiluminescence substrate mixture according to the manufacturer's instructions (Amersham Pharmacia Biotech, Tokyo) for 1 min and exposed to Fuji x-ray film (Fuji Photo Film). Anti-uPA antibodies were used as the primary antibodies. For quantification, computerized scanning and densitometry (Power Macintosh 7600/200-assisted FAS-II and Electronic U.V. transilluminator; Toyobo Co. Ltd., Tokyo) were used. PKC was determined essentially as described by Walton et al.(22Walton G.M. Bertics P.J. Hudson L.G. Vedvick T.S. Gill G.N. Anal. Biochem. 1987; 161: 425-437Crossref PubMed Scopus (79) Google Scholar). Aliquots of both cytosolic and membrane extracts were assayed for PKC activity by a PKC enzyme assay system (PepTagTMNon-Radioactive Protein Kinase Assays; Promega, Madison, WI), according to the instructions of the manufacturer. Two micrograms of PepTag™ C1 peptide were incubated as in the standard reaction with varying amounts of PKC in a final volume of 25 μl for 30 min at 23 °C. The reactions were stopped by heating to 95 °C for 10 min. The samples were loaded on a 0.8% agarose gel and run at 100 V for 15 min. Phosphorylated peptide migrated toward the anode, while nonphosphorylated peptide migrated toward the cathode. Using a razor blade, the nagatively charged phosphorylated bands were excised from the gel and assayed for PKC activity according to the manufacturer's instructions. Western blot analysis of PKC was performed using α-subtype-specific monoclonal antibodies. Immunoreactivity was analyzed using ECL detection kit as described above. Total RNA was isolated from cells by lysis in Trizol reagent according to the manufacturer's instructions (Life Technologies, Inc.); 10 μg of RNA were separated in 1.2% agarose gels and blotted onto Hybond N+ membranes.uPA mRNA was detected by a radioactively labeleduPA oligonucleotide probe. A 1.0-kilobaseEcoRI-PstI fragment of a human uPAcDNA (23Medcalf R.L. Van den Berg E. Schleuning W.D. J. Cell Biol. 1988; 106: 971-978Crossref PubMed Scopus (123) Google Scholar) was used as a probe in the hybridization experiments.uPA cDNA was labeled with [32P]dCTP by the random primed DNA labeling technique as described before (24Niiya K. Taniguchi T. Shinbo M. Ishikawa T. Tazawa S. Hayakawa Y. Sakuragawa N. Thromb. Haemostasis. 1994; 72: 92-97Crossref PubMed Scopus (16) Google Scholar). Following hybridization with uPA, blots were stripped and rehybridized with glyceraldehyde-3-phosphate dehydrogenase(GAPDH) as a semi-quantitative control by densitometry. After each hybridization, the membranes were washed and exposed on Kodak BioMax MS-1 film at −70 °C. Invasion assays were performed essentially as described previously (25Johnson M.D. Torri J.A. Lippman M.E. Dickson R.B. Cancer Res. 1993; 53: 873-877PubMed Google Scholar). Assays were conducted using a 24-well Boyden chamber apparatus. For motility assays, 8-μm pore sized, polycarbonate filters were placed in the apparatus separating the upper from the lower wells. The lower wells contained 25 μl of fibroblast-conditioned media prepared by incubating confluent monolayers of NIH 3T3 fibroblasts for 24 h with Dulbecco's modified Eagle's medium containing 0.1% bovine serum albumin and 0.05 mg/ml ascorbic acid. HRA cells were harvested with trypsin/EDTA, washed twice with Dulbecco's modified Eagle's medium containing 10% fetal calf serum, resuspended in media containing the appropriate treatment, and added to the top well (5 × 104 cell/well). The filters were coated with 0.375 mg of Matrigel per filter. The apparatus was incubated in a humidified incubator at 37 °C in 5% CO2, 95% air for 18 h, after which the cells that had traversed the membrane and spread on the lower surface of the filter were stained with Diff-Quick and quantified electronically with the image analysis system. This system analyzes 32 independent fields for each filter. Cells are identified on the basis of nuclear staining and a count of cells per field is generated. The chemotactic assay was conducted as described previously (4Kobayashi H. Gotoh J. Shinohara H. Moniwa N. Terao T. Thromb. Haemost. 1994; 71: 474-480Crossref PubMed Scopus (64) Google Scholar). For chemotaxis assay, the upper filters were not coated with Matrigel. All experiments were performed using at least three different cell preparations. Data are presented as mean ± S.D.. All statistical analysis was performed using StatView for Macintosh. The Mann-Whitney U test was used for the comparisons between two groups. In cases in which significant interactions were detected, Duncan's multiple range test was used for group comparisons. p less than 0.05 was considered significant. It has been established that the cell-associated receptor-bound uPA activity is important for tumor cell invasiveness (26Reuning U. Magdolen V. Wilhelm O. Fischer K. Lutz V. Graeff H. Schmitt M. Int. J. Oncol. 1998; 13: 893-906PubMed Google Scholar). The effect of various concentrations of PMA on the uPA expression was determined by a chromogenic assay. The HRA cells were treated with PMA, calcium ionophore A23187, or 8-Br-cAMP. A23187 and cAMP were included for comparison because they have been shown to induce the uPA in other cell types. Fig. 1 shows that PMA strongly induced PA activity on HRA cell surface. Plasminogen activator activity on the surface of the cells incubated with PMA (100 nm) was increased about 7-fold as compared with the control cells. In contrast, A23187 and cAMP showed a negligible effect by themselves or in combination with PMA (not shown). When cells were preincubated with anti-uPA IgG, PA activity was inhibited more than 85%, indicating that most of the PA activity expressed by HRA cells is uPA. As shown in Fig. 1, inset, induction by PMA reached maximal and 50% values at the concentrations 100 and ∼10 nm, respectively. Time course analyses showed that PMA induction reached a maximum at 9 h (see Fig. 6).Figure 6Time-dependent effect of UTI or H-7 on steady state levels of PMA-stimulated PA activity. Cells (1 × 106/well) were exposed to PMA (100 nm, ●) for 1 to 9 h in the absence or presence of UTI (100 nm, ■) or H-7 (10 μm, ○). Cell associated uPA activity was assayed. UTI was added to the cells 30 min before PMA stimulation. Asterisks indicate barsthat are significantly different from the 1-h incubation (p < 0.05).View Large Image Figure ViewerDownload Hi-res image Download (PPT) We investigated whether UTI could inhibit PMA-induced stimulation of PA activity expressed on the cell surface. We initially reported (2Kobayashi H. Shinohara H. Ohi H. Sugimura M. Terao T. Fujie M. Clin. Exp. Metast. 1994; 12: 117-128Crossref PubMed Scopus (37) Google Scholar) that UTI is able to reduce cell-associated protease activity directly via inactivation of plasmin and trypsin, while UTI fails to inhibit uPA activity. In the present study, to examine if UTI could be directly modifying uPA catalytic activity, purified human uPA activity was assayed with or without UTI (0.01–1 μm). We again confirmed that there was no inhibitory effect of UTI on uPA catalytic activity. The dose-dependent ability of UTI to inhibit PMA-induced expression of uPA activity by cells is clearly demonstrated (Fig.2 A). In cell monolayers treated with PMA, cell associated PA activity was significantly decreased in the presence of 100 nm UTI. The maximal suppression of PMA-induced PA expression was obtained at 1000 nm UTI. Constitutive uPA expression without stimulation by PMA was also affected to a lesser degree by 1000 nm UTI (Fig. 2 B). Cells exposed to not less than 100 nm(PMA-stimulated) or 1000 nm UTI (constitutive, nonstimulated) exhibited a significant decrease in PA activity. Contrary to UTI, the COOH-terminal domain of UTI (HI-8), which is active fragment for protease inhibitor but is not recognized by the cell-associated UTI-binding sites, failed to suppress PMA stimulated PA activity at concentrations of HI-8 as high as 5000 nm. Furthermore, to determine whether the increased expression of cell-associated uPA produced by treatment with PMA resulted in an increase in the secretion of this enzyme, conditioned media were prepared from cells treated with PMA and UTI and analyzed by zymography and Western blotting (Fig. 3). Western blot with anti-human uPA antibody confirmed and correlated the decrease in PMA-stimulated uPA secreted activity with a lower uPA antigen expression. Zymographic assays also showed that cell monolayers produced a PA activity corresponding to a main band of 55 kDa. This activity was completely abolished by anti-catalytic antibodies to human uPA, confirming uPA identity (not shown). By zymography, plasminogen-independent protease activity was not detected in HRA cells (not shown). Thus, we confirmed that treatment of cell monolayers with UTI showed an inhibitory effect on PMA-stimulated expression and secretion of enzymatically active uPA. Results obtained after exposing the cells to UTI before, during, and after stimulation by PMA are presented in Fig.4. Preincubation of the cells with UTI during 30 min before 100 nm PMA stimulation results in a concentration-dependent inhibition of the induction of cell associated PA activity. At concentrations of 100 and 1000 nm, PA activity is inhibited by 55 and 75%, respectively. This inhibition is irreversible since it cannot be reversed by washing the cells before PMA stimulation. The presence of UTI during stimulation by PMA (concurrent treatment) does not cause a dramatic reduction of PA activity. At a concentration of 1000 nm, PA activity is inhibited by ∼25%. In contrast, no significant decrease of PA activity is observed when UTI is added to the medium 1 h after stimulation by PMA. More than 90% of the control value still remains at the highest UTI concentration (1000 nm) tested. Cell viability, monitored by lactate dehydrogenase leakage in the culture medium and trypan blue dye exclusion test, is not altered under the different exposure conditions (data not shown). These experiments demonstrated that a marked and a slight, but significant, decrease of PA activity are observed when UTI is added to the medium before and during stimulation by PMA. The cell signaling pathways associated with UTI-induced down-regulation of PA expression were estimated utilizing several types of specific inhibitors (Fig.5). The cells exposed to 100 nm PMA exhibited a 7.5-fold increase in PA activity on the cell surface. Preincubation of the cells with either aspirin (2 μg/ml) or eicosatetraenoic acid (0.1 mm) had no effect on the ability of PMA to stimulate PA activity. This experiment demonstrated that it is unlikely that cyclooxygenase or lipoxygenase products of arachidonic acid are involved. We next compared the ability of UTI to reduce PMA-stimulated expression of PA activity by cells pretreated with each PKC inhibitor. When cells were incubated with 10 μm H-7, the ability of PMA to stimulate the expression of cell associated PA activity was inhibited about 90%. When cells were preincubated with 1.0 μm UTI, the ability of PMA to stimulate the expression of PA activity was inhibited about 60%. Higher concentrations of UTI (10 μm) gave similar results on inhibition of PMA-dependent stimulation of PA activity (see Fig. 2). Although UTI, like H-7, was effective to reduce PMA-stimulated PA activity, induction of PA activity was not synergistically reduced. This shows that component(s) of UTI action are mediated dependently of PKC. Other PKC inhibitors such as calphostin C (250 nm), not shown, and staurosporin (50 nm) gave similar results. These data suggest that UTI may suppress PA activity in a manner analogous to PKC inhibitor. The time-dependent accumulation of PA activity on the cells exposed to PMA is presented in Fig. 6. Cell associated PA activity did not increase until after 2 h of PMA stimulation and continued to increase over 9 h. As expected, PMA stimulated PA activity was significantly suppressed in a time-dependent manner when the cells were preincubated with 1 μm UTI or 10 μm H-7, respectively. Of note, both UTI and H-7 did not directly inhibit uPA activity. The effect of UTI or PKC inhibitors on the PMA-induced expression of uPA mRNA was studied. RNA was prepared from cells treated with PMA and inhibitors and hybridized with probes derived from human cDNA clones ofuPA. Fig. 7 shows the results from a blot probed for uPA mRNA. PMA produced a marked increase in uPA expression at the gene level. The expression of theuPA gene was increased by ∼12-fold at 100 nmPMA for 3 h. This stimulation was abrogated in cells pretreated with UTI or PKC inhibitors. The time-dependent effect of UTI on PMA-stimulated expression of uPA mRNA was determined. As illustrated in Fig. 8, an increase in HRA celluPA mRNA levels was observed after 1 h and peaked after 3 h. uPA mRNA levels dropped sharply over the next 6 h. The increase in levels of uPA mRNA observed in cells treated with PMA for 9 h was inhibited more than 50% (UTI) or 95% (H-7) when cells were preincubated with UTI or H-7, respectively. We have investigated whether UTI can suppress PMA-mediated PKC translocation. HRA cells were grown to confluency and then maintained overnight in serum-free conditions to keep the cells in a quiescent state. PKC enzymatic assay and immunoblot analysis showed that stimulation with 100 nmPMA for 30 min at 37 °C resulted in the translocation of PKC-α in the membrane fraction, with a concomitant decrease in the cytosolic pool, while total PKC activity did not significantly change (Fig.9). Abolition of PMA effect was achieved in the presence of UTI, but not of HI-8. UTI did not directly inhibit the catalytic activity of partially purified porcine brain PKC even at the concentration of 1 μm (data not shown). The phosphorylation of proteins in tyrosine/threonine residues is a prerequisite for the activation of these enzymes. In some systems these events are PKC-dependent (27Widmann C. Gibson S. Jarpe M.B. Johnson G.L. Physiol. Rev. 1999; 79: 143-180Crossref PubMed Scopus (2304) Google Scholar). Recent publication demon
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