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INTRODUCTION |
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 (1-9). The UTI gene encodes a
Kunitz-type protease inhibitor of molecular mass 40 kDa (9), which is
composed of a ligand-binding domain (amino terminus) for
cell-associated UTI-binding sites (10, 11) and protease inhibitor
domain (carboxyl terminus) (9), 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 (12), 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 (13). 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-
, interleukin-1
,
interferon-
, tumor growth factor-
, fibroblast growth factor-2,
okadaic acid, retinoic acid, UV, and oncogene products v-Src and v-Ras
(14). 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 (15).
Agents that modulate uPA have been shown to alter the rate of
metastasis in in vitro experiments and in some animal models
(16). 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 (12); 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 of
uPA 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.
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EXPERIMENTAL PROCEDURES |
Materials--
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 (9). 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.
Cell Line and Culture Conditions--
Human ovarian cancer cell
line HRA was obtained from Dr. Y. Kikuchi (17). 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 (18) 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.
Determination of 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 the
x axis and their corresponding absorbance values on the
y axis. The uPA activities were quantitatively obtained by
reading from the calbration curve.
Lactate Dehydrogenase Activity--
Viability of the cells was
assessed by measuring lactate dehydrogenase release in the medium as
described previously (19). Lactate dehydrogenase release is expressed
as % of total enzyme content determined after cell disruption with
Triton X-100.
Zymography and Caseinolysis--
Zymography was performed as
described previously (20). 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 (21), using
plasminogen-rich casein-agarose plates. Secreted uPA activity was
referenced to a standard uPA curve and normalized to 106
cells 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.
SDS-Polyacrylamide Gel Electrophoresis and Western
Blot--
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 mM
sodium vanadate, 2 mM sodium pyrophosphate, 1 mM phenylmethylsulfonyl fluoride, 25 mM
-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.
Determination and Cellular Localization of PKC Activity--
PKC
was determined essentially as described by Walton et al.
(22). Aliquots of both cytosolic and membrane extracts were assayed for
PKC activity by a PKC enzyme assay system (PepTagTM
Non-Radioactive Protein Kinase Assays; Promega, Madison, WI), according
to the instructions of the manufacturer. Two micrograms of PepTagTM 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
-subtype-specific monoclonal antibodies. Immunoreactivity was
analyzed using ECL detection kit as described above.
Northern Blot Analysis--
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 labeled
uPA oligonucleotide probe. A 1.0-kilobase
EcoRI-PstI fragment of a human uPA
cDNA (23) 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 (24).
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--
Invasion assays were performed essentially
as described previously (25). 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 (4). For chemotaxis assay,
the upper filters were not coated with Matrigel.
Statistical Analysis--
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.
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RESULTS |
PMA Specifically Stimulates HRA Cell Expression of uPA
Activity--
It has been established that the cell-associated
receptor-bound uPA activity is important for tumor cell invasiveness
(26). 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).

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Fig. 1.
Induction of PA activity by PMA, calcium
ionophore A23187, or 8-Br-cAMP. Cells (5 × 105
cells/well) exposed to PMA (100 nM, for 4 h), A23187
(5 µM, for 6 h), or 8-Br-cAMP (1 mM, for
2 h) were preincubated with anti-uPA IgG (50 µg) or nonimmune
IgG (NI-IgG) overnight at 4 °C. Anti-uPA antibody used is a
neutralizing antibody. After washing, the cell associated PA activity
in the samples was determined as described under "Experimental
Procedures." Inset, dose-response. Cells were treated for
5 h with different concentrations of PMA (0, 1, 10, 100, and 1000 nM). Optical density was read at 405 nm. Each point
represents the mean (A405) and S.D. of
measurements made on three independent samples. a-c,
mean ± S.D. with unlike superscripts are different
(p < 0.05).
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Suppression of PMA-induced uPA Expression and Secretion by
UTI--
We investigated whether UTI could inhibit PMA-induced
stimulation of PA activity expressed on the cell surface. We initially reported (2) 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. 2A). 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. 2B). 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.

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Fig. 2.
Suppression of PMA-induced stimulation of PA
activity by pretreatment with UTI and HI-8. A,
dose-response. Cells (106/well) were preincubated with
0-10 µM UTI for 30 min. The preincubation media were
replaced with medium containing UTI and PMA (100 nM) for
5 h. B, effect of UTI and HI-8 on PA activity in the
HRA cells stimulated during 5 h by PMA. Mean PA activity in
control stimulated cells amounts to 121 ± 9.0 nmol of
p-nitroaniline/h/106 cells. Values represent
mean ± S.D. of three experiments. a-d, mean ± S.D. with unlike superscripts are different (p < 0.05).
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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.

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Fig. 3.
Effect of UTI on HRA cell uPA production
during a 24-h treatment. uPA activity in the conditioned media is
expressed in IU/106 cells/24 h for control or treated
groups. A, zymography; B, Western blot with
anti-uPA antibody; and C, uPA activity in conditioned media.
Lane 1, control high molecular weight uPA (0.1 µg/lane).
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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.

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Fig. 4.
Effect of exposure of cells to UTI before,
during, and after stimulation by PMA. Mean PA activity in control
stimulated cells amounts to 153 ± 11.3 nmol of
p-nitroaniline/h/106 cells. Values represent
mean ± S.D. of three experiments. a, 10 nM
UTI; b, 100 nM UTI; and c, 1000 nM UTI. a-g, mean ± S.D. with unlike
superscripts are different (p < 0.05).
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The Effect of UTI and Several Types of PKC Inhibitor on
PMA-stimulated Expression of PA Activity--
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.

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Fig. 5.
Effects of inhibitors on PMA-induced HRA
cell-dependent PA activity. We examined the ability of
UTI (1 µM) to reduce PMA (1 µM)-stimulated
expression of PA activity by cells pretreated with PKC inhibitors or
other types of specific inhibitors. Cell associated uPA activity was
assayed. Working concentrations: H7, 10 µM; staurosporin
(Sta), 50 nM; aspirin, 2 µg/ml; and
eicosatetraenoic acid (ET), 0.1 mM. Experiments
were performed twice with similar results.
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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.

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Fig. 6.
Time-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 bars
that are significantly different from the 1-h incubation
(p < 0.05).
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The Effect of UTI and Several Types of PKC Inhibitors on
PMA-induced uPA mRNA Expression--
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 of
uPA. 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 the
uPA gene was increased by ~12-fold at 100 nM
PMA for 3 h. This stimulation was abrogated in cells pretreated
with UTI or PKC inhibitors.

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Fig. 7.
Induction of uPA mRNA by
PMA in the absence or presence of either UTI or PKC inhibitor.
Northern blot analysis of total RNA (10 µg) from HRA cells for
uPA mRNA levels. B, the bar graph
was derived from the ratio of uPA and
glyceraldehyde-3-phosphate dehydrogenase (GAPDH)
densitometric measurements for each condition. Experiments were
performed twice with similar results.
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The time-dependent effect of UTI on PMA-stimulated
expression of uPA mRNA was determined. As illustrated in
Fig. 8, an increase in HRA cell
uPA 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.

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Fig. 8.
Time-dependent effect of UTI or
H-7 on steady state levels of PMA-stimulated uPA
mRNA expression. Cells (2 ± 107/flask)
were exposed to PMA (100 nM, ) for 1 to 9 h in the
absence or presence of UTI ( ) or H-7 ( ). Total RNA was isolated,
and uPA mRNA levels were determined by Northern blot
hybridization utilizing a human uPA cDNA probe.
GAPDH, glyceraldehyde-3-phosphate dehydrogenase.
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Inhibition of PKC Translocation by UTI--
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 nM
PMA for 30 min at 37 °C resulted in the translocation of PKC-
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).

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Fig. 9.
Subcellular localization of PKC
activity. A, the PKC activity is expressed as picomoles
of phosphate transferred per minute per 106 cells. ,
cytosol; , plasma membrane. PMA, 100 nM; UTI and HI-8, 1 µM. Data reported are the mean ± S.D.
(n = 4-6). B, cells were grown in 10-cm
dishes to confluency and then incubated for 30 min with 100 nM PMA. Cytosolic and membrane extracts were prepared, and
100 µg of each extracts were separated by polyacrylamide gel
electrophoresis, blotted, and subjected to Western analysis using PKC
-subtype-specific monoclonal antibodies. Experiments were performed
twice with similar results. C, cytosol; M,
membrane
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UTI Suppresses PMA-triggered Phosphorylation of MEK1, ERK2, and
c-Jun--
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 (27). Recent
publication demonstrated that activation of a PMA-dependent
signal pathway involves a relay of phosphorylation of several proteins
making up the pathway (27). Therefore, we investigated the effect of UTI on the phosphorylation of MEK1, ERK2, and c-Jun in HRA cells stimulated with PMA. This may be detected by Western blot analysis as a
mobility shift in SDS-polyacrylamide gel electrophoresis (Fig.
10). As expected, PMA could activate a
signaling pathway involving MEK1/ERK2/c-Jun. In nonstimulated cells the
expression of phosphorylated proteins was weak, whereas PMA
significantly raised the levels of the phosphorylated form of these
proteins. We found that phosphorylation of these proteins is modified
within 30 min of induction by PMA and then returned to the uninduced state after 5 h (not shown). When cells were preincubated with UTI, however, we could detect suppression of phosphorylation of these
proteins. The results, based on densitometric scanning, show that UTI
significantly inhibits PMA-triggered phosphorylation of MEK1, ERK2, and
c-Jun, by ~80, 60, and 50%, respectively. Moreover, HI-8 did not
change significantly the expression of phosphorylated proteins (not
shown). Therefore, these results show that UTI inhibits phosphorylation
of MAP kinase proteins at the concentration (1 µM) that
prevents earlier translocation of PKC-
.

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Fig. 10.
Western blot analysis of MEK1, ERK2, and
c-Jun proteins. Modification of these proteins was shown. Whole
cell extracts (20 µg of total protein) from cells treated for 30 min
with 100 nM PMA in the absence or presence of 1 µM UTI were analyzed by Western blot. Western blots were
first incubated with antibodies against MEK1, ERK2, and c-Jun and then
with biotin-conjugated secondary antibodies and avidin-peroxidase,
followed by enhanced chemiluminescence. The "m" shows
the modified isoforms. Phosphorylation-dependent mobility
shift of MEK1 and ERK2 proteins in response to PMA induction was
detected. This is a representative experiment selected from two
performed.
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The Effect of PMA, PKC Inhibitors, and UTI on
Invasiveness--
The effects of agents that alter the activity of PKC
on the invasiveness of HRA cells were determined by measuring the
ability of cells treated with these agents to pass through a layer of the extracellular matrix extract Matrigel coating a filter using chemoinvasion chambers as described under "Experimental
Procedures." Fig. 11 shows that
treatment with PMA produced a significant stimulation of the
invasiveness of the cells in a dose-dependent manner. The stimulation was maximal at 100 nM PMA (7-fold). This
stimulation was reversed by concurrent treatment with either PKC
inhibitor or UTI. Fig. 12 shows the
effect of adding increasing concentrations of PKC inhibitors
(staurosporin and H-7) or UTI on the invasiveness of cells stimulated
with 100 nM PMA. Calphostin C was also able in our studies
to inhibit the stimulatory effects of PMA on invasiveness (not shown).
Although concurrent treatment with UTI produced a dose-dependent reversal of the PMA-induced stimulation, the
migration of cells treated with 100 nM PMA was reduced to
about 60% by the addition of 1 µM UTI, indicating that
UTI may exhibit less inhibitory to abrogate PMA-induced cell migration
compared with the PKC inhibitors. The inhibition seen is not due to
nonspecific cytotoxicity produced by these agents, as the
concentrations used do not significantly inhibit the cell
viability.

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Fig. 11.
Induction of HRA cell invasiveness by
treatment with PMA. HRA cells were trypsinized and counted, and
5 × 104 cells were placed in the top wells of a
chemoinvasion chamber apparatus with the indicated concentration of
PMA. After 18 h the cells that had passed through the
Matrigel-coated filter toward the bottom wells containing
fibroblast-conditioned medium as a chemoattractant were quantitatied.
Each point represents the mean and S.D. of measurements made
on three independent wells. Asterisks indicate bars that are
significantly different from the vehicle control (p < 0.05).
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Fig. 12.
Suppression of PMA-induced HRA cell
invasiveness by treatment with UTI or PKC inhibitor H-7. HRA cells
were trypsinized and counted, and 5 × 104 cells were
placed in the top wells of a chemoinvasion chamber apparatus with the
indicated concentration of UTI or a PKC inhibitor, H-7, in the presence
of 100 nM PMA. Each point represents the mean and S.D. of
measurements made on three independent wells. a-g, mean ± S.D. with unlike superscripts are different (p < 0.05).
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In addition, the cell chemotactic response was also tested to determine
whether the inhibitory effect of UTI on cell invasion of basement
membranes was due to an inhibition of chemotaxis. The cells tested
showed good chemotactic migration in the presence of UTI (data not
shown; see Ref. 4). In addition, we examined the effects of UTI on cell
attachment. No inhibition of attachment to Matrigel (or fibronectin)
was seen with UTI (data not shown). Thus, we confirmed that UTI has no
ability to inhibit cell migration or chemotaxis, which is consistent
with our previous data (4).
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DISCUSSION |
A variety of signal transduction pathways for uPA gene
expression have been described for the action of cytokines and growth factors. Multiple signal transduction pathways, for example, PKC and
cAMP protein kinase pathways, converge at the level of uPA gene regulation. The uPA promoter contains functional
binding sites for transcription factors AP-1, PEA3, and NF-
B (28). The induction of uPA is regulated at least in part by a
c-Jun-dependent MAP kinase pathway (28). Agents that
regulate signaling pathways involved in uPA expression
appear to be a useful probe to help dissect cell invasion process.
One may speculate that PMA can promote cell invasion possibly through
an increase in cell-associated uPA expression. This by-passes the cell
surface events involved in the phosphatidylinositol system. In HRA
cells used in this study, PKC-dependent signal transduction
needs to be activated to achieve uPA expression, whereas neither
increase in intracellular calcium nor cAMP activation was able to
stimulate uPA expression. It follows then that inhibition of PKC
signaling pathway could result in the inhibition of uPA expression and
cell invasion. Moreover, preincubation of cells with inhibitors of
cyclooxygenase or lipoxygenase had no effect on PMA-induced uPA
expression. These data suggest that the effect of PMA on uPA expression
is not likely due to alterations in the permeability to calcium,
adenylate cyclase activity, or arachidonic acid metabolism. Therefore
we tested the effects of UTI on PKC-dependent signaling
pathway only.
We have previously shown that in human umbilical vein endothelial cells
and promyeloid leukemia cell line U937 activation of PKC represents a
critical event in tumor necrosis factor-triggered signaling cascade
leading to uPA secretion (12). Furthermore, we had shown that induction
of uPA expression by tumor necrosis factor-
was inhibited when these
cells were incubated with UTI and that UTI may influence the
PKC-dependent protein kinase pathway in uPA expression.
However, the molecular mechanisms of UTI-mediated changes in signaling
pathways regulating uPA gene expression is not fully
understood. Therefore, we asked whether the inhibition was due to
interference with the PKC signaling system which cooperates with the
MAP kinase pathway.
We detected changes in the PMA-stimulated uPA expression by UTI. HRA
cells normally only express low levels of this enzyme, and upon
stimulation with PMA expression and secretion were each induced well.
This induction was inhibited by pre- or concurrent treatment with UTI
or PKC inhibitors. The UTI's action is relatively rapid and
irreversible; it occurs after 30 min of contact and persists after UTI
is removed and the cells are washed. One h after stimulation by PMA,
however, UTI has no effect on uPA expression. We show that this
reduction occurs at the level of gene transcription (or mRNA
degradation) in that the mRNA levels for uPA were
reduced in the presence of UTI in the Northern analysis. Present data do not indicate whether UTI is selectively inhibiting the
uPA gene or also inhibits other gene transcription as well.
Notwithstanding these limitations, we have also shown in HRA cells that
the PMA-stimulated cell motility (invasiveness) can be suppressed by
UTI. Increased uPA activity might be contributing to the increase in
motility produced by PMA treatment, indicating that the level of uPA
expression closely parallels the degree of cell motility. Thus, the
reduction in uPA expression by UTI might appear to explain the degree
of inhibition of cell motility.
Although UTI does not directly inhibit the catalytic activity of PKC,
UTI inhibited the translocation of PKC from cytosol to membrane when
stimulated with PMA. Thus UTI may prevent PKC from being up-regulated
by PMA and PKC remained in an inactivated state. Therefore, the
inhibition of PMA-stimulated uPA expression by UTI must be occurring at
a point beyond the diacylglycerol activation event. Activated PKC also
causes the phosphorylation of a wide variety of intracellular proteins
(29). The MAP kinase pathway is activated via the stimulation of Ras
Raf
MAPKK (MEK1/2)
ERK1/2. It was suggested that PKC
phosphorylates and activates Raf-1 and ERK1/2 (27). More recent data
support the implication of different PKC isoenzymes in the phorbol
ester-induced activation of the ERKs pathway in different cell types
(30, 31). If phorbol ester leads phosphorylation and assembly of the
MAP kinase pathway, it remains possible that UTI is inhibiting one or
more of these phosphoprotein products. The present results show for the
first time that UTI's function on PMA-induced production of uPA in
tumor cells involves the PKC regulatory signaling pathway which may
cooperate with the MEK/ERK/c-Jun pathway. We report that UTI blocks
PMA-triggered phosphorylation of MAP kinases, when used at a
concentration of 1 µM. Because this concentration of UTI
affects cytosol-to-membrane translocation of PKC-
isoform, it is
likely that at least in part PKC-
participates in the activation of
MAP kinases in our system. These data allow us to conclude that the
inhibitory effect of UTI for uPA expression involves an interference
with the PKC second messenger system, and appears to occur downstream
of diacylglycerol activation and upstream of MEK phosphorylation, which
are critical in the uPA expression and subsequently in cell motility.
Transcription of human genes encoding uPA is a complex
process that requires participation of several transcription factors.
It is also well documented that NF-
B activity could be strongly
induced by PMA (32). We cannot exclude the possibility that UTI may
directly affect MAP kinase phosphorylation or that UTI may inhibit DNA
binding activity of transcription factors. Also, there is no evidence
whether UTI binds to PKC at a catalytic site to which PMA binds or a
site other than the catalytic one.
Suppression by PKC inhibitors of uPA stimulation by PMA is complete.
Contrary to several PKC inhibitors, inhibition by UTI of
down-regulation of uPA expression is partial. PKC actually exists not
as a single entity, but in a family of isoforms; their activation
mechanisms are very complex and they may exhibit significant biochemical differences (33, 34). It is possible that the mechanism of
action of both UTI and PKC inhibitors is partly different, although a
PMA-sensitive subtype of PKC could be involved in UTI's function.
On the basis of our previous data obtained in our laboratories, we had
shown that UTI binds to tumor cells via specific UTI-binding sites (10,
11). Cell-associated UTI-binding proteins (UTI-BPs) may be critical
targets for UTI, since HI-8, which does not possess a ligand for
UTI-BPs (11), failed to prevent the PMA-stimulated uPA expression.
Membrane-associated UTI-BPs are believed to represent the rate-limiting
step for UTI-dependent signal transduction or cellular
uptake of the UTI. Taken together, we hypothesize the following
mechanism in the UTI-dependent regulation: the UTI/UTI-BP interaction is followed by modulation of a PKC system, which inhibits translocation of certain PKC isoform(s) including PKC-
from the microsome to the plasma membrane compartment. UTI-dependent
suppression of activation and redistribution of PKC-
results in the
reduction of uPA expression possibly via inhibition of phosphorylation
of the MEK/ERK/c-Jun proteins. Thus, UTI makes cells refractory to stimulation with exogenously added PMA.
Recent publication demonstrated that one of the UTI-BPs is identical to
link protein (11). Link protein has been implicated in the
stabilization of the hyaluronic acid-rich matrix on the cell surface in
a variety of cell types (35, 36). Link protein specifically binds to
hyaluronic acid and sits on the outer lamellae of the cell surface
membrane and is not able to transduce a signal into the cell interior,
unless it associates with other cell surface molecules able to
transduce such a signal. Again, the mechanism involved in the
ligand-receptor interaction remains unknown.
In previous studies (12), we had shown that UTI was as effective as PKC
inhibitors in inhibiting uPA expression by tumor necrosis factor in
both human umbilical vein endothelial and U937 cells. Treatment of the
cells with UTI alone failed to alter uPA production. Incubation of the
cells with UTI, however, had no effect on the ability of PMA to
stimulate cell-associated uPA expression. The mechanism of the
discrepancy remains to be explained. One candidate mechanism is
difference in the PKC isoenzymes that mediate uPA signal transduction.
In conclusion, we describe the in vitro regulation of the
PMA-induced expression of uPA gene and cell motility by UTI
in ovarian cancer cells and the signaling pathways involved in the
UTI-dependent modulation of the PMA induction. We found
that cytosol-to-membrane translocation of PKC-
isoenzyme may
represent early steps in signaling cascades that lead to uPA production
in human ovarian cancer cells and that UTI prevents translocation of
PKC-
, and subsequently inhibits MEK1, ERK2, and c-Jun
phosphorylation and inhibits cell motility at the concentrations
relevant to reported IC50 values for uPA production. It is
unlikely that UTI directly inhibits catalytic activity of PKC-
. At
this point, it is not possible to implicate any UTI-specific targets
(protease, adhesive molecule, or other determinant of motility).
Nevertheless, as reported here, UTI exhibited no cytotoxic effects even
at concentrations of 10 µM. The present effect of UTI
occurs at concentrations in the micromolar range; it is thus likely
that such effects may occur in vivo at therapeutically
relevant concentrations.