* Program in Toxicology and Department of Pathology, University of Maryland, School of Medicine, Baltimore, Maryland 21201, and Toxicology and Drug Disposition, Lilly Research Laboratories, A Division of Eli Lilly and Company, P.O. Box 708, Greenfield, Indiana 46140
Received February 2, 2004; accepted May 17, 2004
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ABSTRACT |
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Key Words: Tsc2; protein kinase C; TPA.
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INTRODUCTION |
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The phorbol ester 12-O-tetradecanoylphorbol-13 acetate (TPA) is the active component of croton oil and a potent tumor promoter. TPA mimics the second messenger diacylglycerol to activate protein kinase C (PKC) and alter cell signaling pathways (reviewed in Gottlicher, 1999). TPA has been used extensively to investigate the role of PKC in biologic responses, but recently has been shown to activate Rap1 in neutrophils and fibroblasts (M'Rabet et al., 1998
; Zwartkruis et al., 1998
). The activation of Rap1 by TPA has been shown to be PKC-independent in neutrophils, and is presumably dependent on the ability of the phorbol ester to directly activate diacylglycerol-specific Rap1 guanine nucleotide exchange factors (M'Rabet et al., 1998
). Given the hypothesized role of Tsc2 in Rap1 regulation, we compared the cellular response to TPA between Tsc2-null and Tsc2-expressing cells. We show that TPA rapidly activated PKC
and the extracellular-signal regulated kinase (ERK) in both cell types, but caused weak Rap1 activation in ERC-18 cells only. TPA induced unique focal accumulations of PKC
in ERC-18 cells, and ERK activation was prolonged in ERC-18 cells when compared to NRK-52E. Furthermore, Tsc2-null cells had a prolonged morphologic response to the tumor promoter that was PKC-dependent and Tsc2-modulated.
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MATERIALS AND METHODS |
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Cell culture and treatment. Tsc2-null ERC-18 cells were derived from Eker rat renal tumors (Freed et al., 1990) and were kindly provided by Dr. Cheryl Walker (MD Anderson Cancer Center; Smithville, TX). Eker renal tumors arise from tubular epithelial cells following loss of functional tuberin expression (Everitt et al., 1992
). The NRK-52E cell line was used as a control Tsc2-expressing renal tubular epithelial cell line, and was purchased from American Type Culture Collection (Manassas, VA). ERC18-FLAG-Tsc2 and ERC18-FLAG-2B lines were generated by stably expressing a FLAG epitope-tagged Tsc2 construct (ERC18-FLAG-Tsc2) or the empty expression vector (ERC18-FLAG-2B) in the ERC-18 cell line. A detailed discussion of the development and phenotype of these cell lines is presented elsewhere (manuscript in review). Briefly, a 5357 base pair HindIII fragment of the pcDNA3-Tsc2 vector (provided by Dr. Cheryl Walker) containing the rat Tsc2 sequence was subcloned into the pCMV-Tag-4 vector (Stratagene; La Jolla, CA), to create the pCMV-Tag-Tsc2 plasmid. The plasmid encodes a 1738 amino acid tuberin construct with the C-terminal 42 amino acids removed and replaced with a FLAG-epitope tag. The pCMV-Tag-Tsc2 plasmid sequence was verified by multiple restriction endonuclease digests. A similar Tsc2 construct, lacking 55 amino acids from the C-terminus, completely inhibited N-ethyl-N-nitrosurea-induced renal carcinoma formation in transgenic Eker rats (Momose et al., 2002
). ERC-18 cells were transfected with the linearized pCMV- Tag-Tsc2 plasmid or with the linearized empty pCMV-Tag vector (control) using Effectene transfection reagent (QIAGEN; Valencia, CA) according to the manufacturer's protocol. Individual G418-resistant colonies were screened by immunoblot analysis for expression of FLAG with an anti-FLAG-M2 antibody (Sigma; St. Louis, MO) diluted to a concentration of 1 µg/ml.
ERC-18, ERC18-FLAG-2B (empty plasmid vector) and ERC18-FLAG-Tsc2 (pCMV-Tag-Tsc2) cells were routinely maintained in a 1:1 mixture of Dulbecco's modified Eagle media (DMEM) and Nutrient mixture F12 (Ham) supplemented with 5% fetal bovine serum, 2 mM L-glutamine, 1.6 µM ferrous sulfate, 50 nM sodium selenite, 12 µM vasopressin, 10 nM cholesterol, 200 nM hydrocortisone, 1 nM tri-iodothyronine (T3), 10 pg/ml transferrin, and 25 µg/ml insulin. NRK-52E cells were maintained in DMEM supplemented with 5% fetal bovine serum and 2 mM L-glutamine. All cell lines were grown in a humidified atmosphere of 37°C and 5% CO2/95% room air. Cells were grown to 90% confluence in 60 or 100-mm plastic culture dishes (Western blot, Rap1 activity assay) or 35-mm dishes with glass coverslips (immunocytochemistry), then incubated in low-serum treatment media (DMEM/F12, 1% FBS) for 24 h prior to treatment. Cells were treated with 50 or 100 ng/ml TPA in treatment media for 5 min, 15 min, 1 h, or 24 h, or with an equivalent volume of vehicle (dimethyl sulfoxide) as a control. In PKC inhibitor experiments, cells were co-treated with 1 µM bisindolylmaleimide VIII (Bis VIII), a selective PKC inhibitor with increased specificity for the PKC isoform (Wilkinson et al., 1993
), or with 1 µM bisindolylmaleimide V (Bis V), an inactive structural analogue. At the indicated time points, cells were photographed on a Leitz Diavert inverted microscope.
Immunoblot analysis. Following treatment, cells were lysed on ice with RIPA buffer (50 mM Tris-Cl, pH = 7.4; 150 mM NaCl; 1% NP-40; 0.5% sodium deoxycholate; 0.1% sodium dodecyl sulfate; 1 mM ß-glycerolphosphate; 2.5 mM sodium pyrophosphate) supplemented with protease inhibitor cocktail P8340 and phosphatase inhibitor cocktails P2850 and P5726 (Sigma), sonicated for 10 s, and clarified by centrifugation (17,000 x g, 10 min.). Protein concentrations were measured using the BCA protein assay (Pierce; Rockford, IL), and 20 µg of protein was separated by SDS-PAGE using standard techniques (Laemmli, 1970) and transferred to a PVDF membrane (Bio-Rad Laboratories; Hercules, CA) in 25 mM Tris, 192 mM glycine, and 20% methanol overnight at 100 mA. Proteins were detected by immunoblot analysis with antibodies specific for phospho-ERK (1:1000), PKC
(1:1000), and caspase 3 (1:1000) according to the manufacturer's protocols. All antibodies were purchased from Santa Cruz Biotechnology (Santa Cruz, CA). Band intensities were quantified from scanned images using Molecular Analyst software (Bio-Rad Laboratories; Hercules, CA), and mean values from multiple independent experiments were compared by randomized block ANOVA with Dunnett's multiple comparisons post-test, unpaired t-test, or two-way ANOVA, as described in the text. Statistically significant differences were judged at a p value <0.05. Immunoblots were stripped and reprobed for actin (1:1000; Santa Cruz) to confirm equal protein loading and transfer. Expression of the 200-kDa FLAG-Tsc2 construct was confirmed in 20 µg of protein from nuclear extracts prepared with the Pierce NE-PER kit according to the manufacturer's protocol.
Rap1 activity assay. GTP-bound (active) Rap1 was measured using a previously described method (Franke et al., 1997). Briefly, cells were treated with 100 ng/ml TPA (or vehicle) for 5, 15, or 60 min, then lysed with RIPA buffer on ice and clarified by centrifugation (17,000 x g, 10 min.). Protein concentrations were measured using the BCA assay, and 600 µg of lysate (diluted to 0.6 mg/ml in RIPA buffer) was incubated for 1 h (4°C) with 5 µg of a RalGDS GST-RBD construct (provided by Dr. Johannes Bos; Utrecht University, The Netherlands) pre-coupled to glutathione-agarose beads. The RalGDS GST-RBD construct has a high and specific affinity for GTP-bound Rap1 (Herrmann et al., 1996
). Beads were washed several times with RIPA buffer, and bound Rap1-GTP was purified by boiling in Laemmli sample buffer (62.5 mM Tris-HCl, pH 6.8, 2% SDS, 25% glycerol, 2% ß-mercaptoethanol, 0.01% bromophenol blue). Pulldown products were separated by SDS-PAGE and transferred to PVDF membranes as described above, and Rap1 levels were detected by immunoblotting with a Rap1 specific antibody (Santa Cruz; 1:500). Immunoblots were stripped and reprobed with a glutathione-S-transferase (GST) specific antibody (Santa Cruz; 1:2000) to confirm that equivalents amount of the
38 kDa RalGDS GST-RBD construct were used to precipitate Rap1-GTP from each sample. Individual Rap1 band intensities were quantified and compared from four independent experiments using randomized block ANOVA, with statistically significant differences judged at a p value <0.05.
Immunocytochemistry. Following TPA treatment, cells were fixed with 2% paraformaldehyde at 4°C for 1 h, then permeabilized with 0.1% Triton X-100 for 30 min. Nonspecific antibody binding sites were blocked for 45 min with 5% normal goat serum in phosphate-buffered saline (PBS) supplemented with 0.5% BSA and 0.15% glycine. Cells were incubated with a PKC-specific antibody (Santa Cruz; 2 µg/ml) for 1 h at 37°C, followed by Alexa-488 conjugated anti-rabbit IgG antibody (Molecular Probes; 1 µg/ml) for 1 h at room temperature. As a negative control, cells were subjected to the staining protocol with primary antibody replaced with dilution buffer (PBS, 0.5% BSA, 0.15% glycine). Coverslips were mounted on clean glass slides with MOWIOL mounting medium supplemented with 1 µg/ml DAPI, and photographed on a Nikon TE200 Eclipse inverted microscope.
Cell proliferation assay. NRK-52E and ERC-18 cells were plated in black, clear-bottom 96-well tissue culture dishes at a density of 104 cells/well. Cells were grown in complete media for 24 h, then in low-serum DMEM/F12 treatment media for 24 h prior to treatment. Cells were treated in duplicate wells with TPA (10, 50, 100 ng/ml or the equivalent volume of dimethyl sulfoxide) in the presence of 100 µM bromodeoxyuridine (BrdU) for 24 h. BrdU incorporation was measured using the BrdU Proliferation Assay Kit (Oncogene Research Products) according to the manufacturer's instructions. Mean BrdU incorporation was determined from duplicate wells in three independent experiments, and the percentage change in cell proliferation from control in TPA-treated cells was compared within each cell type using randomized block ANOVA. Statistically significant differences were judged at a p value <0.05.
Measurement of cell death (membrane integrity loss). Cells were plated in six-well tissue culture dishes, then grown to 90% confluence and incubated in DMEM/F12 low-serum treatment media for 24 h. Cells were then treated with 10, 50, 100, 200, or 500 ng/ml TPA (or the equivalent volume of vehicle) in the presence of the membrane-impermeant nuclear dye SYTOX green (0.5 µM, Molecular Probes). Fluorescence was measured after 0, 24, and 48 h of treatment using a Cytofluor 2350 (Millipore; Bedford, MA). After the 48 h reading, all cells were permeabilized with 100 µg/ml saponin, and total fluorescence was determined. At each treatment and time point, fluorescence values were normalized to the total cellular fluorescence (saponin).
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RESULTS |
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TPA-Induced Morphologic Change Does Not Represent Increased Proliferation or Apoptosis
The rounded morphology observed following TPA treatment is similar to cellular rounding observed as an early phenotypic alteration during proliferation or apoptosis. Indeed, TPA has been shown to promote and inhibit both proliferation and apoptosis in a variety of cell types. Therefore, we examined the effects of TPA on cell proliferation using BrdU labeling as a measure of proliferation. We treated NRK-52E and ERC-18 cells with 10, 50, or 100 ng/ml TPA for 24 h in the presence of BrdU. Remarkably, TPA caused a significant decrease in cell proliferation in NRK-52E cells at all doses (p = 0.003; randomized block ANOVA), with no apparent dose-response (Fig. 2). The tumor promoter had no effect on cell proliferation in the ERC-18 cell line (p = 0.97; randomized block ANOVA). Cleavage of caspase-3 was also examined as a measure of apoptosis, and caspase cleavage product was not observed in either cell type, even after treatment with 100 ng/ml TPA for 48 h. We also measured cell permeability as an indicator of cell death in both cell lines after treatment with 10500 ng/ml TPA for 2448 h. No increase in cell death was apparent in either cell type, even after treatment with 500 ng/ml TPA was carried out to 48 h. These data indicate that the TPA did not induce proliferation or apoptosis in either cell type.
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BisVIII cotreatment was able to dramatically reduce TPA-induced ERK phosphorylation in both cell types. Figure 6B shows the relative increase in ERK phosphorylation (compared to control) induced by TPA treatment in the presence of BisVIII and BisV in both cell lines. BisVIII cotreatment significantly reduced TPA-induced ERK activation in NRK-52E cells and ERC-18 cells (p < 0.01; two-way ANOVA and p < 0.001; two-way ANOVA, respectively).
TPA-Induced PKC Degradation Is More Extensive in NRK-52E Cells than in ERC-18 Cells
As PKC appears to be an important regulator of both the morphologic change and increased ERK phosphorylation induced by TPA, we wanted to examine potential differences in the ability of the phorbol ester to down-regulate the kinase between NRK-52E and ERC-18 cells. Cells were treated with 50 or 100 ng/ml TPA for 24 h, and PKC
expression was measured in whole cell lysates. A dose-dependent decrease in PKC
expression was observed in both cell types after 24 h of TPA treatment (not shown). Comparison of PKC
levels after 24 h of treatment with 100 ng/ml TPA showed that this decrease was much more pronounced in NRK-52E cells (Fig. 7A). Indeed, comparison of the relative decrease in PKC
expression (from control) after 24 h treatment with 100 ng/ml TPA showed a statistically significant difference between NRK-52E and ERC-18 cells (p = 0.02; unpaired t-test), with only 6% of control PKC
remaining in NRK-52E cells and 32% remaining inERC-18 cells after TPA treatment (Fig. 7B).
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DISCUSSION |
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Previous studies have shown that cytosolic PKC rapidly translocates to the plasma membrane upon TPA-induced activation (Nakashima, 2002
), and that PKC can activate the MAPK signaling pathway via Raf-1 dependent (Kolch et al., 1993
) and independent (Chao et al., 1994
) mechanisms following phorbol ester activation. In addition, TPA has been shown to activate Rap1 in a PKC-independent manner in neutrophils and fibroblasts (M'Rabet et al., 1998
; Zwartkruis et al., 1998
). In the present study, TPA-induced changes in cell morphology and ERK activation in both cell types appear to be at least partially PKC-dependent. TPA caused translocation of PKC
to the plasma membrane within 5 min in both cell types, temporally consistent with a causal role for the kinase in morphologic change and ERK activation. More importantly, changes in cell morphology and ERK activation were abrogated by cotreatment with the selective PKC inhibitor bisindolylmaleimide VIII. These findings clearly indicate that TPA functions through PKC to induce morphologic changes and ERK activation in NRK-52E and ERC-18 cells. However, while TPA-induced morphologic change appears to be primarily dependent on PKC activation, the prolonged ERK activation observed in TPA-treated ERC-18 cells may be mediated by the effects of the tumor promoter on Rap1. Since Rap1 activity increased in response to TPA in ERC-18 cells, and since Rap1 has been shown to mediate sustained ERK activation (York et al., 1998
), it is possible that basal Rap1 activity is partially responsible for the prolonged ERK activation observed in ERC-18 cells. However, since reexpression of Tsc2 did not abrogate the prolonged ERK activation in ERC-18 cells, it is unclear whether Tsc2 is required for this effect. It may be possible that levels of Tsc2-FLAG expression were not high enough to affect Rap1 activity in the current study. Alternatively, the differential activation of ERK in NRK-52E and ERC-18 based cell lines may be due to cell-type specific differences not involving Tsc2. Regardless, Rap1 activation is not likely to be responsible for the prolonged morphologic response of ERC-18 cells treated with TPA since bisindolylmaleimide VIII completely abrogated TPA-induced morphologic change in both cell types. In addition, expression of Tsc2 in ERC-18 cells abrogated only the prolonged morphologic response of these cells to TPA, implying that the prolonged changes in ERC-18 cell morphology following TPA treatment are not the result of prolonged ERK activation.
Previous studies have indicated that TPA-induced changes in cell morphology may arise from the phosphorylation of key focal adhesion (FA) substrates by PKC. A previous study of TPA-induced changes in renal epithelial cell morphology showed that the tumor promoter caused a disruption of focal contacts, redistribution of the FA protein vinculin, and reorganization of the actin cytoskeleton (Rahilly and Fleming, 1992). PKC
binds the FA proteins vinculin and talin (Hyatt et al., 1994
) and was co-localized with talin at focal contacts in rat embryo fibroblasts (Jaken et al., 1989
). In the current study, PKC
was localized to distinct focal contacts after 1 h of TPA treatment in ERC-18 cells. Focal localization of PKC
was not observed in NRK-52E cells after TPA treatment, and may be an important mediator of the prolonged morphologic response of ERC-18 cells to TPA. Expression of Tsc2 in ERC-18 cells abrogated the prolonged morphologic response, emphasizing the importance of Tsc2 in the regulation of this response.
While TPA rapidly activates PKC, prolonged TPA treatment results in down-regulation of the kinase (Blumberg et al., 2000
). We observed a more pronounced decrease in PKC
in NRK-52E cells compared to ERC-18 cells following prolonged TPA treatment. Since the morphologic response required PKC activation, it may be possible that the prolonged morphologic response of ERC-18 cells to TPA is due to the persistent expression of the kinase in these cells even after 24 h of treatment. In renal epithelial cells, PKC
is normally degraded following prolonged activation via ubiquitination and proteasomal degradation (Lee et al., 1996
). Tsc2 loss may alter the ability of renal epithelial cells to degrade PKC
via normal proteasomal mechanisms. Alternatively, expression of PKC
may be increased in Tsc2-null cells. Indeed, we have consistently observed elevated PKC
expression in ERC-18 cells compared to NRK-52E cells, although it is unclear whether this increase is Tsc2-dependent (unpublished observation). Additional studies are required to directly test these possibilities.
Activation of PKC has been implicated in the regulation of proliferation, apoptosis, differentiation, cell migration, and adhesion (reviewed in Nakashima, 2002
). The morphologic changes observed in the current study, combined with evidence from previous studies, suggest that PKC
-dependent adhesion and/or migration may be altered in Tsc2-null cells. Changes in cell morphology observed in the current study were not the result of increased cell proliferation or apoptosis. In a previous report, nearly identical changes in cell morphology (loss of intercellular adhesion, retraction of cell periphery, increased refractility, rounding) were induced in primary cultures of renal epithelial cells treated with 100 ng/ml TPA and caused cell detachment after 2 h (Rahilly and Fleming, 1992
). In the current study, TPA did not induce detachment in either cell type, even after agitation. This difference may be due to phenotypic differences between the primary cultures used in the previous study and the cell lines used in our study. Alternatively, differences in the culture vessels used (laminin- or fibronectin-coated glass vs. tissue culture-treated plastic) may account for the differences in detachment between studies.
PKC may also regulate cell adhesion and motility by binding and phosphorylating members of the actin-binding ezrin-radixin-moesin (ERM) family of proteins (Ng et al., 2001
). Hamartin, the product of the Tsc1 tumor suppressor gene, binds both tuberin and ERM proteins and appears to be required for cell-matrix adhesion (Lamb et al., 2000
). Over-expression of tuberin in Madine-Darby canine kidney (MDCK) epithelial cells or re-expression of tuberin in Tsc2-null cells has recently been shown to promote cell adhesion, inhibit migration, and activate the Rho GTPase (Astrinidis et al., 2002
). Interestingly, TPA-induced changes in MDCK cell focal adhesion structure and actin organization were shown to be dependent on the activities of Rho and Rab5 GTPases in one study (Imamura et al., 1998
). There is also evidence supporting the possibility that tuberin functions as a GAP for Rab5 (Xiao et al., 1997
). The findings of the current study clearly show that the prolonged morphologic response of renal epithelial cells to TPA is Tsc2-dependent. Taken together, these findings may implicate PKC
as an additional mediator in the Tsc2-dependent regulation of cell adhesion and migration (Fig. 8). Changes in the migratory response of renal epithelial cells to PKC
activation may be particularly important in vivo, as PKC activation appears to have an invasion-promoting role during RCC. The in vitro invasiveness of human renal cell carcinoma lines was reduced by PKC inhibitors, and membrane translocation of the PKC
isoform was correlated with increased invasiveness of the RCC lines used (Engers et al., 2000
).
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ACKNOWLEDGMENTS |
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NOTES |
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1 To whom correspondence should be addressed. Fax: (317) 277-6770. E-mail: davisma{at}lilly.com.
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