* Program in Toxicology and Department of Pathology, University of Maryland, School of Medicine, Baltimore, Maryland 21201, and Toxicology and Drug Disposition, Lilly Research Laboratories, Eli Lilly and Company, Greenfield, Indiana 46140
1 To whom correspondence should be addressed at Toxicology and Drug Disposition, Lilly Research Laboratories, A Division of Eli Lilly and Company, P.O. Box 708, Greenfield, IN 46140. Fax: (317) 277-6770. E-mail: davisma{at}lilly.com.
Received July 27, 2005; accepted August 29, 2005
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ABSTRACT |
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Key Words: tumorigenesis; Tsc2; apoptosis; okadaic acid.
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INTRODUCTION |
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Apoptosis plays a key role in the regulation of kidney cell number. Compensatory increases in apoptosis have been correlated with increased proliferation and tumor grade, stage, and size during renal cell carcinoma (RCC) development (Todd et al., 1996). Tuberin, the product of the tuberous sclerosis complex-2 (Tsc2) tumor suppressor gene, also appears to play an important role in renal tumorigenesis. In humans, a genetic mutation in Tsc2 results in tuberous sclerosis, a disease syndrome that consists of hamartomatous tumors in several organs and such renal manifestations as multiple, bilateral angiomyolipomas, cystic disease, and renal cell carcinoma (Cook et al., 1996
). A single germline mutation in the rat homologue of Tsc2 results in the development of renal cell carcinoma with a nearly complete penetrance in the Eker rat model (Everitt et al., 1995
). These rats also develop uterine leiomyomas and splenic hemangiosarcomas (Everitt et al., 1992
, 1995
; Howe et al., 1995
) and are highly susceptible to renal cell carcinoma induced by a variety of toxicants (Hino et al., 1993a
,b
; Horesovsky et al., 1994
; Lau et al., 2001
; Walker et al., 1992
; Wolf et al., 1998
).
Many studies have implicated Tsc2 in the regulation of cell differentiation (Soucek et al., 1998), cell cycle control (Soucek et al., 1997
, 1998
), GTPase activity (Astrinidis et al., 2002
; Wienecke et al., 1995
; Xiao et al., 1997
; Zhang et al., 2003
), transcription (Henry et al., 1998
; Zhang et al., 2003
), polycystin-1 localization (Kleymenova et al., 2001
), and translation initiation (Dan et al., 2002
; Gao et al., 2002
; Goncharova et al., 2002
; Inoki et al., 2002
; Kenerson et al., 2002
; Potter et al., 2002
; Tee et al., 2002
). Tsc2 has recently been shown to be a downstream component of the phosphatidylinositol-3' kinase (PI3 kinase) signaling pathway, which normally functions to regulate cell growth and promote survival (reviewed in Blume-Jensen and Hunter, 2001
; Datta et al., 1999
). A downstream component of this signaling pathway, Akt kinase, promotes survival by phosphorylating multiple substrates. These substrates include the pro-apoptotic proteins BAD (Zha et al., 1996
) and caspase-9 (Cardone et al., 1998
), and the forkhead family of transcription factors (Brunet et al., 1999
). Tuberin is also reported to be an Akt substrate, and phosphorylation of tuberin by Akt promotes degradation (Dan et al., 2002
; Gao et al., 2002
; Goncharova et al., 2002
; Inoki et al., 2002
; Kenerson et al., 2002
; Potter et al., 2002
; Tee et al., 2002
). It is unknown whether Tsc2 is involved in the anti-apoptotic function of Akt via this or any other mechanism. We propose that loss of TSC2 will decrease susceptibility of renal epithelial cells to apoptosis.
To measure the effect of Tsc2 expression on the apoptotic response of renal tumor cells, we expressed Tsc2 in Tsc2-null ERC-18 cells and compared the susceptibility of Tsc2-null and Tsc2-expressing ERC-18 cells to apoptosis induced by the phosphatase inhibitor, okadaic acid (OKA), and the PI3 kinase inhibitor, LY294002. We show that Tsc2 expression increases the susceptibility of ERC-18 cells to death induced by both compounds, and that the unique morphologic response of ERC-18 cells to OKA is Tsc2-dependent. We also show that the apoptosis-promoting function of Tsc2 does not appear to be mediated through mTOR.
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MATERIALS AND METHODS |
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Plasmid construction.
A plasmid encoding the full-length rat Tsc2 cDNA sequence (1765 amino acid splice variant; 36) in the pCDNA3 vector (pcDNA3-Tsc2) was provided by Dr. Cheryl Walker (MD Anderson Cancer Center; Smithville, TX). A 5357 base pair HindIII fragment of pcDNA3-Tsc2 containing the Tsc2 sequence was subcloned into the HindIII site of 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).
Cell line development and culture.
The Tsc2-null ERC-18 cell line was derived from renal tumors of the Eker rat (38), and was provided by Dr. Cheryl Walker. 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. After transfection for 48 h, cells were grown in the presence of 200 µg/ml G418 (BD Biosciences Clontech; Palo Alto, CA) for 11 days. Individual G418-resistant colonies were isolated and propagated in the presence of 200 µg/ml G418 until reaching the 60-mm dish stage, then 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. ERC18-FLAG-2B (empty plasmid vector) and ERC18-FLAG-Tsc2 (pCMV-Tag-Tsc2) cells were routinely maintained in DF-8 media, a 1:1 mixture of Dulbecco's modified Eagle media (DMEM) and Nutrient mixture F12 (Ham) supplemented with 5% fetal bovine serum (FBS), 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. All cell lines were grown in a humidified atmosphere of 37°C and 5% CO2/95% room air. Mycoplasmal contamination assays were not routinely performed. Cells were grown to 90% confluence in 60-mm plastic culture dishes (immunoblot analysis) or in six-well dishes (membrane permeability assay), then incubated in low-serum treatment media (DMEM/F12, 1% FBS) for 24 h prior to treatment with OKA or LY294002. Apoptotic morphology was photographed on a Leitz Diavert inverted microscope.
Determination of clone growth rate.
ERC-18, ERC18-FLAG-2B, and ERC18-FLAG-Tsc2 cells were plated in 12-well culture dishes (1 x 104 cells per well) in complete DF-8 media. On days 1, 3, 5, and 7 after plating, cells in duplicate wells were incubated with 0.5 µM SYTOX Green (Molecular Probes; Eugene, OR) and 100 µg/ml saponin (Sigma; St. Louis, MO) for 10 min at 37°C. SYTOX green fluorescence was measured on a Cytofluor 2350 (Millipore; Bedford, MA), and fluorescence values were averaged for duplicate wells at each time point. Cell number was determined using a standard curve prepared from fluorescence intensities of known cell numbers for each cell type. The mean cell number at each time point was then determined from three independent experiments. One-week growth curves were prepared for each clone and the parent ERC-18 cell line using GraphPad Prism software (Graphpad Software, Inc.; San Diego, CA) by converting to logarithms and plotting against time. Prism software was used to generate sigmoidal concentration-response curves for each cell type using nonlinear regression. Doubling times were calculated between days 3 and 5, when all cells appeared to be in a logarithmic growth phase. Doubling time was calculated for each cell line using the equation Nt = N02tf, where Nt is the number of cells at time = t, N0 is the initial number of cells, t is the time in days, and f is the frequency of cell cycles per day.
Measurement of cell death (membrane permeability assay).
Cell membrane integrity was assessed via SYTOX-green uptake. Viable cells exclude the fluorophore, while nuclei in cells with increased membrane permeability are labeled. Cells were grown to 90% confluence in six-well culture dishes, then incubated in DMEM/F12 low-serum treatment media for 24 h. Cells were treated in low-serum media for 24 h with 0.05, 0.1, and 0.25 µM OKA or 10, 50, and 100 µM LY294002, or with an equivalent volume of vehicle (ethanol or DMSO) as a control. In rapamycin experiments, cells were pre-treated for 1 h with 10 nM rapamycin, and the inhibitor was included (at 10 nM) as a cotreatment with OKA or LY294002. After 24 h of treatment, cells were incubated with 0.5 µM SYTOX-green for 10 min prior to measuring fluorescence using a Cytofluor 2350 (Millipore; Bedford, MA). After measuring death-induced fluorescence, saponin (100 µg/ml) was added for 10 min to permeabilize all cells, and total fluorescence was measured. In each well, death-induced fluorescence values were normalized to the total cellular fluorescence, and the mean change in membrane permeability (fluorescence) from control was determined from three independent experiments. Statistically significant changes in membrane permeability from control (vehicle) were identified using randomized block analysis of variance (ANOVA) with Dunnett's multiple comparisons post-test. With each concentration, the mean change in membrane permeability was compared between cell lines using an unpaired t-test. Statistically significant differences were judged at p < 0.05.
Immunoblot analysis.
Whole cell lysates were prepared from 60-mm cultures after 10 h (OKA) or 24 h (LY294002) of treatment for analysis of caspase-3 cleavage. Floating cells were collected by centrifugation, and lysates were prepared on ice from floating and adherent cells with RIPA buffer (50 mM Tris-Cl, pH = 7.4; 150 mM NaCl; 1% Nonident P-40; 0.5% sodium deoxycholate; 0.1% sodium dodecyl sulfate; 2.5 mM sodium pyrophosphate; 1 mM ß-glycerolphosphate) supplemented with protease inhibitor cocktail P8340 and phosphatase inhibitor cocktails P2850 and P5726 (Sigma). Lysates were sonicated for 10 s, then clarified by centrifugation (17,000 x g, 10 min.). Protein concentrations were measured using the BCA protein assay (Pierce; Rockford, IL), and 25 µg of protein was separated by SDSPAGE using standard techniques (Laemmli, 1970) and transferred to a PVDF membrane (Bio-Rad; Hercules, CA) in 25 mM Tris, 192 mM glycine, and 20% methanol overnight at 100 mA. Full-length and cleaved caspase-3 were detected with a caspase-3 antibody (Santa Cruz Biotechnology; Santa Cruz, CA), used at a 1:1000 dilution according to the manufacturer's protocol.
A separate lysis procedure was used to measure cytosolic BAD accumulation and phosphorylation of Akt (serine 473). Following 30 h of serum starvation (BAD) or 24 h of LY294002 treatment (phospho-Akt), cytosolic extracts were prepared as previously described (Kolb et al., 2002). Briefly, cells were placed in ice-cold MS Buffer (5 mM Tris-Cl, pH = 7.5; 210 mM mannitol; 70 mM sucrose; 1 mM EDTA; 40 µg/ml digitonin) supplemented with protease and phosphatase inhibitors, as above, and scraped into microfuge tubes. After a 10-min incubation on ice, a cytosolic extract was prepared by centrifugation at 17,000 x g for 15 min. The pellet (containing nuclei, mitochondria, and other membranes) was washed once with MS buffer, then centrifuged again (17,000 x g for 15 min ) prior to lysis with 10 volumes of RIPA buffer and sonication (10 s). Cytosolic extracts were separated by SDSPAGE and transferred to PVDF membranes. Immunoblot analysis was performed using antibodies specific for BAD (CST; 1:500) and phospho-Akt (ser473, Santa Cruz; 1:1000) according to the manufacturers' protocol. To measure cytosolic accumulation of BAD, band intensities were quantified from scanned images using Molecular Analyst software (Bio-Rad). Mean cytosolic BAD levels from five independent experiments were compared between 2B-D and Tsc2-D cells using an unpaired t-test with statistically significant differences judged at p < 0.05.
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RESULTS |
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Tsc2 Expression Increases the Susceptibility of ERC-18 Cells to OKA-Induced Apoptosis
In the Tsc2-null ERC-18 cell line, OKA induced extensive apoptosis with a morphologic response that was unique from the response observed in other Tsc2-expressing renal epithelial cell lines. From this observation, we inferred that there may be a difference in the susceptibility of Tsc2-null and Tsc2-expressing renal epithelial cells to OKA. To measure the effect of Tsc2 expression on susceptibility to OKA-induced apoptosis, we compared death induced in Tsc2-D cells after 24 h of exposure to 0.05, 0.1, and 0.25 µM OKA to that induced in a control cell line expressing only the empty vector (2B-D). In both cell lines, OKA induced a concentration-dependent increase in cell death after 24 h (Fig. 3A). Death was significantly increased at all concentrations when compared to cells exposed to vehicle (p < 0.01; Dunnett's multiple comparisons post-test following significant differences in randomized block ANOVA). Comparison between cell lines within each treatment group (concentration) showed that expression of Tsc2 increased susceptibility to OKA-induced death when compared to the control cell line (Fig. 3A). At the lowest concentration (0.05 µM OKA), there was no difference in the susceptibility of 2B-D and Tsc2-D cells to OKA. After 24 h of exposure with 0.1 or 0.25 µM OKA, Tsc2-D cells showed a significant increase in death when compared to the 2B-D cell line (0.1 µM, p = 0.002; 0.25 µM, p = 0.006; unpaired t-test). To confirm that apoptosis was the type of cell death caused by OKA treatment, we measured cleavage of caspase-3 after 10 h of treatment with 0.1µM and 0.25 µM OKA. We have previously shown caspase-3 is cleaved between 6 and 10 h in ERC-18 cells treated with 0.1µM OKA. As shown in Figure 3B, caspase-3 is cleaved in both cell lines after 10 h of exposure with both concentrations. The extent of cleavage was increased in the Tsc2-D cell line when compared to 2B-D cells.
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To determine whether mTOR contributed to the observed Tsc2-dependent increase in apoptotic susceptibility, we inhibited mTOR by pretreating cells with 10 nM rapamycin, then treated cells with 0.25 µM OKA or 100 µM LY294002 in the presence of 10 nM rapamycin. If the pro-apoptotic effects of Tsc2 are mTOR dependent, rapamycin should functionally replace Tsc2 in null cells (2B-D) to inhibit mTOR, thereby increasing apoptotic susceptibility. As shown in Figure 6, mTOR inhibition with rapamycin had no effect on OKA-induced apoptosis in either cell line and actually inhibited the apoptotic response of 2B-D cells to LY294002 (p < 0.01; unpaired t-test).
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DISCUSSION |
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We initially characterized differences in apoptotic susceptibility of Tsc2-null and Tsc2-expressing cells using OKA, a compound that consistently induces apoptosis in renal epithelial cells in vitro (Davis et al., 1994, 1996
), including the ERC-18 tumor cell line (Kolb et al., 2002
). Our results show that expression of Tsc2 in ERC-18 cells increased susceptibility to OKA-induced apoptosis. OKA inhibits the activities of the phosphatases PP1 and PP2A, and OKA-induced apoptosis in renal epithelial cells appears to involve the concurrent activation of multiple kinases, including protein kinase C (PKC
), Raf-1, extracellular signal-regulated kinases 1 and 2 (ERK-1/2), jun N-terminal kinase (JNK), and p38 (Davis and Carbott, 1999
; Davis et al., 1996
). The present studies show that OKA-induced apoptosis in renal epithelial cells can be regulated by Tsc2.
The synthetic quercetin derivative LY294002 inhibits PI3 kinase and may promote cell death via inhibition of Akt activity. Since multiple studies have shown Tsc2 to be an Akt substrate (Dan et al., 2002; Inoki et al., 2002
; Tee et al., 2002
), we wanted to know if expression of the tumor suppressor would influence PI3 kinase inhibitor-mediated cell death. While LY294002 induced apoptosis in a small but significant population of Tsc2-null cells, expression of Tsc2 in ERC-18 cells increased susceptibility to LY294002-induced death. These results imply that Tsc2-expressing cells may be more dependent on PI3 kinase-mediated signaling for survival than Tsc2-null renal tumor cells. The decreased apoptotic response of Tsc2-null cells to PI3 kinase inhibition may have important implications for tumor progression, since survival signals normally activating PI3 kinase may become limiting as cell proliferation and tumor growth continues. Tsc2-null cells would therefore be at a distinct survival advantage under these conditions due to their decreased apoptotic susceptibility.
Although we have not determined the precise mechanistic role played by the Tsc2 in cell death, we have data that imply the mechanism does not involve mTOR. Tuberin has been shown to inhibit mTOR activity (Gao et al., 2002; Inoki et al., 2002
; Kenerson et al., 2002
; Tee et al., 2002
), and tuberin loss results in rapamycin-sensitive increases in p70S6 kinase activity (Goncharova et al., 2002
; Inoki et al., 2002
). Phosphorylation of BAD by p70S6 kinase is rapamycin-sensitive and has been reported to promote cell survival (Harada et al., 2001
). Therefore, it was reasonable to expect that inhibition of mTOR by rapamycin would mimic the pro-apoptotic effects of Tsc2 in null cells if the tumor suppressor promotes the chemical-induced apoptosis observed via inhibition of mTOR. However, rapamycin did not increase the susceptibility of Tsc2-null cells to apoptosis induced by either compound. Although a recent study showed that short-term rapamycin treatment induced apoptosis in Eker rat renal tumors in vivo (Kenerson et al., 2002
), the concentrations of rapamycin used in our study did not induce apoptosis in our cell lines at the times measured. However, both studies strongly support a role for a reduced apoptotic response in the pathogenesis of Tsc2-null renal tumors. Additional study is needed to reveal additional mechanistic links between tuberin and apoptosis in renal epithelial cells.
Our previous characterization of OKA-induced apoptosis in ERC-18 cells showed a unique early morphologic response that was caspase independent (Kolb et al., 2002). In the present study, we show that these early morphologic changes (shrinkage, rounding, detachment) are also Tsc2 dependent. While the Tsc2-null control cell line (2B-D) showed early morphologic changes identical to those observed in ERC-18 cells when exposed to OKA, expression of Tsc2 abrogated this effect and produced a morphological response more characteristic of that observed in other Tsc2-competent renal epithelial cells. These findings indicate that Tsc2 expression may regulate shrinkage, rounding, and detachment in renal epithelial cells. Our results also provide additional support for the hypothesis that Tsc2 plays a role in regulation of cell adhesion. Astrinidis et al. (2002)
recently reported that tuberin regulates cell adhesion, migration, and activation of the Rho GTPase in renal epithelial cells and in a Tsc2-null cell line derived from an Eker rat leiomyoma. These findings may be particularly important in TSC-associated diseases of smooth muscle cell origin like malignant renal angiomyolipomas and pulmonary lymphangioleiomyomatosis (LAM). A model for LAM pathogenesis has been proposed where Tsc2-null smooth muscle cells migrate from the kidney to the lung (Carsillo et al., 2000
; Yu et al., 2001
).
Our results may have important implications for the progression of both spontaneous and chemically-induced Eker rat renal carcinomas. A decreased apoptotic response in Tsc2-null or Tsc2-deficient renal epithelial cells can contribute to the progression of both types of tumors. Furthermore, decreased dependence of Tsc-2 deficient cells on PI3 kinase-mediated survival signals might potentiate renal carcinogenesis or may facilitate tumor progression when growth factors are limiting. A general decrease in the apoptotic response may also contribute to chemical carcinogenesis by preventing deletion of mutated cells following genotoxic exposures or by failing to balance proliferative increases induced by nongenotoxic chemicals. Additional studies on the mechanism of Tsc2-dependent apoptotic regulation may therefore provide a better understanding of Tsc2 tumor suppressor function.
In summary, we have shown that expression of Tsc2 in ERC-18 cells increases susceptibility to apoptosis induced by the tumor promoter OKA and the PI3 kinase inhibitor LY294002. Although the precise mechanism by which Tsc2 promotes apoptotic susceptibility to these two compounds remains unclear, our data indicate that alterations in mTOR signaling pathways are not required. In addition, we have shown that Tsc2 expression abrogates the OKA-inducedcaspase-independent detachment of Tsc2-null ERC-18 cells. Tsc2-dependent changes in cell adhesion properties may have important implications in determining the cellular susceptibility to apoptosis. Additional studies are required to more specifically address the mechanistic relationship between Tsc2 expression, apoptotic susceptibility, and cell adhesion.
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ACKNOWLEDGMENTS |
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