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Mutational inactivation of the tumor suppressor tuberous sclerosis complex 2 (TSC2) constitutively activates mTORC1, increases cell proliferation, and induces the pathological manifestations observed in tuberous sclerosis (TS) and in pulmonary lymphangioleiomyomatosis (LAM). While the role of mTORC1 in TSC2-dependent growth has been extensively characterized, little is known about the role of mTORC2. Our data demonstrate that mTORC2 modulates TSC2-null cell proliferation and survival through RhoA GTPase and Bcl2 proteins. TSC2-null cell proliferation was inhibited not only by reexpression of TSC2 or small interfering RNA (siRNA)-induced downregulation of Rheb, mTOR, or raptor, but also by siRNA for rictor. Increased RhoA GTPase activity and P-Ser473 Akt were inhibited by siRNA for rictor. Importantly, constitutively active V14RhoA reversed growth inhibition induced by siRNA for rictor, siRNA TSC1, reexpression of TSC2, or simvastatin. While siRNA for RhoA had a modest effect on growth inhibition, downregulation of RhoA markedly increased TSC2-null cell apoptosis. Inhibition of RhoA activity downregulated antiapoptotic Bcl2 and upregulated proapoptotic Bim, Bok, and Puma. In vitro and in vivo, simvastatin alone or in combination with rapamycin inhibited cell growth and induced TSC2-null cell apoptosis, abrogated TSC2-null tumor growth, improved animal survival, and prevented tumor recurrence by inhibiting cell growth and promoting apoptosis. Our data demonstrate that mTORC2-dependent activation of RhoA is required for TSC2-null cell growth and survival and suggest that targeting both mTORC2 and mTORC1 by a combination of proapoptotic simvastatin and cytostatic rapamycin shows promise for combinational therapeutic intervention in diseases with TSC2 dysfunction.
Mutational inactivation of the tumor suppressor gene tuberous sclerosis complex 2 (TSC2) occurs in hamartoma syndrome tuberous sclerosis (TS) and pulmonary lymphangioleiomyomatosis (LAM). TS manifests as multiple tumors in the brain, kidney, heart, and skin (33). LAM, a rare lung disease predominantly affecting women, which can be sporadic or associated with TS, manifests by neoplastic growth of atypical smooth muscle LAM cells, lung cyst formation, obstruction of lymphatics, and spontaneous pneumothoraces (33, 46, 85). A major advance in understanding TS and LAM occurred with the discovery of TSC2 as a negative regulator of the mammalian target of rapamycin complex 1 (mTORC1) (30, 32, 50, 54, 59, 93). TSC2 forms a tumor suppressor complex with TSC1 and regulates mTORC1 by directly controlling the activity of the small GTPase Rheb via the GTPase-activating protein (GAP) domain of TSC2 (9, 24, 43, 95). Rheb binds to raptor and controls the activity of mTORC1, which directly phosphorylates p70 S6 kinase (S6K1) and 4E-BP1 (57, 63). Importantly, the constitutive activation of S6K1 promotes a negative-feedback loop that also inhibits phosphatidylinositol 3-kinase (PI3K) activity (37, 80, 88). Activation of mTORC1 is sensitive to inhibition by rapamycin, which has been used in the treatment of TS and LAM (35). Rapamycin treatment induced regression of astrocytomas in TS (21) and markedly improved pulmonary functions and reduced the size of angiomyolipomas (AML) in LAM-TS patients (3). Unfortunately, cessation of rapamycin therapy resulted in the regrowth of subependymal giant cell astrocytomas (21) and the return of diminished pulmonary function and AML tumors to the state observed prior to rapamycin treatment (3). Accordingly, alternative or combinational therapies are needed to treat diseases with mutational inactivation of TSC2.
The mTOR forms two functionally distinct multiprotein complexes, mTORC1 and mTORC2, that control cell growth, proliferation, and survival. Both mTORC1 and mTORC2 are critical for embryonic development (36, 81). Further, the rapamycin-sensitive mTORC1 has an evolutionarily conserved function in regulating cell growth, cell cycle progression, and cell proliferation (20). While both mTORC1 and mTORC2 are required for cell growth in yeast (64), little is known about the role of mTORC2 in increased cell proliferation due to TSC2 loss. The first established function of TORC2 was the rapamycin-insensitive cell cycle-dependent regulation of actin cytoskeleton through activation of Rho GTPase (76, 77). Identification of mTORC2 in mammals confirmed its rapamycin-insensitive regulation of actin cytoskeleton through the Rho GTPases RhoA and Rac1 (44, 75). The Rho family of small GTPases consists of three major members, RhoA, Rac1, and Cdc42 (16). In its GTP-bound state, activated RhoA promotes stress fiber and focal-adhesion formation. TSC1 forms a tumor suppressor complex with TSC2 and promotes stress fiber formation through activation of RhoA GTPase (56). We demonstrated that loss of TSC2 induces a rapamycin-insensitive increase in stress fiber formation due to activation of RhoA GTPase that is inhibited by reexpression of TSC2 or small interfering RNA (siRNA) for TSC1 (27, 31). Whether TSC2 regulates the actin cytoskeleton through mTORC2 activation and/or whether TSC2-dependent activation of Rho is required for increased TSC2-null cell proliferation remains unknown.
The biochemical functions of Rho GTPases in regulating the actin cytoskeleton and cell adhesion translate into biological functions, including regulation of cell proliferation, migration, and morphogenesis (45). Geranylgeranylation of Rho GTPase ensures membrane localization and activation (74) that can be affected by statins, small-molecule inhibitors of 3-hydroxy-3-methylglutaryl coenzyme A (HMG-CoA) reductase. Statins modulate lipid metabolism and geranylgeranylation of Rho GTPase (58, 73) and induce apoptosis and inhibit proliferation in various cancer cell lines and human smooth muscle cells (22, 23, 48, 66, 71, 84, 91). Importantly, statins synergize with rapamycin and its analogs in inhibiting proliferation and inducing apoptosis in leukemia, prostate and breast cancer, and rat smooth muscle cells (6, 8, 17, 70). Interestingly, atorvastatin had little effect on syngeneic TSC2−/− mouse embryonic fibroblast (MEF) subcutaneous tumors in nude mice (61). In this study, we utilized TSC2-null ELT3 cells derived from spontaneous uterine leiomyoma (38) of the Eker rat, a naturally occurring animal model with germ line TSC2 gene mutation (51). In our published studies, we have also demonstrated that in ELT3 cells, loss of TSC2 increased proliferation due to constitutively active mTORC1 (30) and activated Rho GTPase due to TSC1-dependent inhibition of Rac1 (27). Whether TSC2-dependent Rho activation acts through mTORC2 and is required for increased TSC2-null cell proliferation has not been investigated.
In this study, we present evidence that mTORC2-dependent RhoA GTPase activation is necessary for increased TSC2-null cell proliferation and survival. We found that siRNA for rictor inhibits both increased RhoA activity and increased P-Ser473 Akt in TSC2-null ELT3 cells and smooth-muscle-like cells derived from LAM lungs. Additionally, siRNA for rictor and siRNA for RhoA inhibit TSC2-null cell proliferation, and expression of constitutively active RhoA rescued TSC2- and rictor siRNA-dependent inhibition of DNA synthesis. We also show that combined targeting of RhoA GTPase with simvastatin and mTORC1 with rapamycin inhibits TSC2-null cell proliferation, induces apoptosis, abrogates TSC2-null tumor growth, and prevents tumor recurrence after simvastatin or the combination of simvastatin with rapamycin treatment was withdrawn. Our data suggest that combined inhibition of RhoA by simvastatin and mTORC1 by rapamycin may be beneficial for inhibiting TSC2-related tumorigenesis and for preventing posttreatment tumor recurrence in LAM and TS.
TSC2-null ELT3 cell, derived from the Eker rat uterine leiomyoma (38) were generously provided to us by Cheryl Walker, M. D. Anderson Cancer Center, University of Texas, Smithville, TX, and maintained as previously described (18, 28, 30). A littermate-derived pair of Tsc2+/+ p53−/− and Tsc2−/− p53−/− MEFs with p53 deleted for immortalization and isogenic Tsc2−/− and Tsc2+/+ MEFs with reconstituted TSC2 were generously provided by D. J. Kwiatkowski (Brigham and Women's Hospital, Boston, MA) and were maintained in Dulbecco's modified Eagle's medium (DMEM) with 10% fetal bovine serum (FBS) (94). LAM-derived (LAMD) cells were dissociated from LAM nodules from the lungs of LAM patients who had undergone lung transplantation, as described previously (30), and obtained from the National Disease Research Interchange (Philadelphia, PA) according to approved protocol. LAMD cells in subculture during the 3rd through 12th passages were used. All experiments were performed with a minimum of three different LAMD cell cultures. Prior to the start of experiments, cells were serum deprived for 24 h.
Microinjection was performed using the Eppendorf Microinjection System (Hamburg, Germany) as described previously (27, 30). Briefly, specific siRNA from Dharmacon, Inc. (Lafayette, CO), directed against Rheb, mTOR, raptor, or rictor or scrambled siRNA was comicroinjected with green fluorescent protein (GFP) or glutathione S-transferase (GST) to identify microinjected cells; 30 to 42 h postinjection, the cells were serum deprived overnight, followed by immunocytochemical or bromodeoxyuridine (BrdU) incorporation analysis.
Cells were washed 3 times with phosphate-buffered saline (PBS), fixed with 3.7% paraformaldehyde for 15 min, treated with 0.1% Triton X-100 for 30 min at room temperature, and blocked with 2% bovine serum albumin (BSA) in PBS for 60 min. After incubation with rhodamine-phalloidin (Molecular Probes, Eugene, OR) or primary and then secondary antibodies conjugated with either Alexa Fluor 488, Alexa Fluor 594, or Alexa Fluor 633, the cells were mounted in Vectashield mounting medium (Vector Laboratories, Burlingame, CA). Immunostaining was analyzed using a Nikon Eclipse TE2000-E microscope equipped with an Evolution QEi digital video camera or using a Nikon Eclipse E400 microscope equipped with a Nikon Coolpix 995 digital camera under ×200 magnification.
pEGFP, pEGFP-TSC2, pEBG, and pEBG-V14RhoA expression vectors were prepared using the EndoFree Plasmid Maxi kit (Qiagen, Valencia, CA). siRNA GLO, scrambled siRNA, and siRNA for Rho, mTOR, rictor, or raptor were purchased from Dharmacon RNA Technologies (Lafayette, CO). Transient transfection was performed using Effectene or RNAiFect transfection reagent (Qiagen) according to the manufacturer's protocols as described previously (27, 29). For siRNA transfection, serum-deprived cells were incubated for 48 or 72 h with specific or control siRNA, followed by the DNA synthesis assay or the analysis of apoptosis. Reexpression of GFP-TSC2 and depletion of RhoA, rictor, or raptor were verified by immunoblot analysis with anti-tuberin (Santa Cruz Biotechnology, Inc., Santa Cruz, CA), anti-RhoA, anti-raptor (Cell Signaling Technology), anti-rictor (Affinity Bioreagents), or anti-total actin (Sigma-Aldrich, Inc., St. Louis, MO) antibodies. For immunoblot analysis, siRNA for Rheb, mTOR, raptor, or rictor was introduced using RNAiFect transfection reagents (Qiagen). As a control, we used scrambled and siGLO RISC-Free siRNA (Dharmacon, Inc., Lafayette, CO). Apoptotic immunoblot analyses were performed using phospho-Ser46 p53, total p53, cleaved caspase 3 antibodies, and a proapoptotic Bcl-2 family antibody kit purchased from Cell Signaling Technology, Inc.
DNA synthesis was examined using a BrdU incorporation assay (28, 30, 34). The cells were examined using the Nikon Eclipse TE2000-E microscope at ×200 magnification with the appropriate fluorescence filters to identify microinjected or transfected cells. The mitotic index was defined as the percentage of BrdU-positive microinjected or transfected cells per field divided by the total number of microinjected or transfected cells per field. A total of 200 cells were counted for each condition in each experiment.
Preconfluent LAMD cells, plated on 6-well plates, were serum deprived for 24 h and then maintained for 14 days in serum-free medium, which was changed daily and supplemented with diluent, 0.3 or 0.5 μM simvastatin, or 20 nM rapamycin alone or in combination with 0.5 μM simvastatin. Cell counts were performed on days 0, 2, 4, 6, 8, 10, 12, and 14. A minimum of three repetitions were performed for each condition in each experiment.
Apoptosis analysis of TSC2-null ELT3 cells that were serum deprived for 24 h and incubated with 0.1, 1, or 10 μM simvastatin alone or in combination with 20 nM rapamycin, 0.2 or 2 μg/ml C3 transferase, or diluent for 18 h was performed as described previously (29). Analysis of the apoptosis of ELT3 cells transfected with siRNA for RhoA or scrambled siRNA was performed using an Alexa Fluor Annexin V/Dead Cell Apoptosis kit (Molecular Probes, Inc., Eugene, OR) according to the manufacturer's protocol. Briefly, serum-deprived cells transfected with siRNA for RhoA or control siRNA for GLO were washed with ice-cold PBS, resuspended in annexin-binding buffer, and then incubated for 15 min with annexin V conjugate. Then, the cells were washed with annexin-binding buffer, deposited on glass slides, mounted in Vectashield mounting medium, and visualized on the Nikon Eclipse E400 microscope using the appropriate filters. Data are represented as the percentage of apoptotic cells per total number of cells, taken as 100%. A total of 200 cells were counted for each condition in each experiment.
Tissue sections of subcutaneous tumors from NCRNU-M athymic female mice treated with diluent, simvastatin, or rapamycin or cotreated with simvastatin and rapamycin were subjected to analysis of apoptosis using the In Situ Cell Death Detection Kit based on terminal deoxynucleotidyltransferase-mediated dUTP-biotin nick end labeling (TUNEL) technology (Roche, Nutley, NJ) according to the manufacturer's protocol. Tumors from a minimum of five animals for each treatment condition were analyzed.
Tsc2+/+ p53−/− and Tsc2−/− p53−/− MEFs and isogenic Tsc2−/− and Tsc2+/+ MEFs were maintained in serum-free medium with 1 μg/ml C3 transferase or diluent for 18 h, and then flow cytometry analysis with primary annexin V and secondary fluorescein isothiocyanate (FITC)-conjugated antibodies was performed as we described previously (28). The negative control included diluent-treated cells incubated with matched IgG and secondary FITC-conjugated antibody.
All animal procedures were performed accordingly to a protocol approved by the University of Pennsylvania Animal Care and Use Committee (IACUC). Six- to 8-week-old NCRNU-M athymic nude female mice (Taconic) were injected subcutaneously in both flanks with 5 × 106 TSC2-null ELT3 cells (see Fig. 9A for the experimental scheme). When tumors reached 5 mm in diameter, the mice were transferred to a diet supplemented with simvastatin (250 mg/kg of body weight/day) or injected with rapamycin (intraperitoneal [i.p.] injections, 1 mg/kg 3 times a week) alone or in combination with a simvastatin diet for 50 days. These doses of simvastatin and rapamycin were chosen based on published observations (41, 48, 90). Chow containing simvastatin was prepared by Animal Specialties & Provision (Quakertown, PA) (JL Rat & Mouse/4F [5LG6] diet based on regular chow, received by the control group, and supplemented with simvastatin [Zocor; Merck]). Animals from each group were euthanized at days 0, 10, and 20 of treatment or when tumors reached 10 mm in diameter; the tumors were removed and evaluated by histological (hematoxylin and eosin [H&E]) and immunohistochemical analyses. Survival analysis was performed using the termination time, when the tumor diameter reached 10 mm. During the treatment time, the general behavior of the mice was monitored three times a week. Tumor size was monitored three times a week and measured using calipers. Tumor volumes were calculated using the following formula, as described previously (87): length × width2 × 0.5. Mouse weight was monitored throughout the experiment, and no significant differences in weight were detected between mice receiving the simvastatin-supplemented diet and mice receiving a regular diet; the average weight of simvastatin-treated animals after 50 days of treatment was 25.488 ± 0.407 g, and that of animals receiving a regular diet was 25.980 ± 1.352 g. During the treatment period, 4 animals were sacrificed: 1 control mouse, 1 rapamycin-treated mouse, and 1 simvastatin-treated mouse were sacrificed due to tumor-related weight loss at days 15, 50, and 22 of treatment, respectively; 1 rapamycin-treated mouse was sacrificed at day 24 of treatment because of an open sore. While these deaths did not cluster in either the rapamycin or the simvastatin treatment groups and thus were not attributed to drug toxicity, these mice were excluded from the analysis. To determine whether rapamycin or simvastatin withdrawal induced the tumor recurrence, treatment of tumor-free animals was terminated after 50 days, and the animals were monitored three times a week; when tumors reached 10% of the animal's body weight, the animals were sacrificed.
Immunohistochemical analysis was performed using sections of tumor tissues, which were snap-frozen in OCT embedding compound (Tissue-Tek, Tokyo, Japan) as described previously (28, 29). Tissue sections were immunostained with anti-Ki-67 (Fisher Scientific International Inc., Hampton, NH) and anti-phospho-S6 (anti-P-S6) antibodies (Cell Signaling Technology, Beverly, MA). Staining with 4′,6-diamidino-2-phenylindole (DAPI) was performed to detect cell nuclei. Tumors from a minimum of five animals per treatment condition were analyzed. Staining was visualized using a Nikon Eclipse TE2000-E microscope with appropriate filters. S6 phosphorylation levels were analyzed by optical density (OD) using Gel-Pro Analyzer software; the P-S6 OD at day 0 was taken as 100%, as we previously described (27, 30). Proliferation levels were determined as a percentage of Ki67-positive cells per total number of cells, taken as 100%. A minimum of 300 cells were analyzed per condition in each tumor.
Data points from individual assays represent mean values ± standard errors (SE). Statistically significant differences among groups were assessed by analysis of variance (ANOVA) (Bonferroni-Dunn), with P values of <0.05 sufficient to reject the null hypothesis for all analyses. All experiments were designed with matched control conditions within each experiment to enable statistical comparison as paired samples.
Activation of mTOR, which forms the catalytic core of multiprotein complexes mTORC1 and mTORC2, is modulated by raptor and rictor, respectively. Loss of TSC2 increases cell proliferation due to the constitutive activation of mTORC1 through the small GTPase Rheb (65), while the role of mTORC2 in TSC2-dependent cell proliferation remains unknown. We examined whether mTORC2 is required for cell proliferation due to TSC2 loss using TSC2-null ELT3 cells derived from the Eker rat uterine leiomyoma (38). As seen in Fig. 1, siRNA-induced depletion of mTOR, Rheb, and raptor inhibited S6 phosphorylation (P-S6) (Fig. 1A and B) and TSC2-null cell proliferation (Fig. 1D) compared to control scrambled siRNA. In contrast, siRNA depletion of rictor had little effect on P-S6, confirming that mTORC2 is not involved in the constitutive activation of S6K1 signaling in TSC2-null cells (Fig. 1A and B). These data are consistent with our and other's published data (9, 30, 32, 95) showing that mTOR, Rheb, and raptor are required for TSC2-null cell proliferation. To test the direct involvement of mTORC2 signaling in cell proliferation, we compared the effects of siRNA for rictor to those of siRNA for mTOR and siRNA for raptor on TSC2-null cell proliferation. Not surprisingly, raptor and mTOR depletion inhibited cell proliferation (Fig. 1D). Importantly, siRNA for rictor also significantly inhibited TSC2-null cell DNA synthesis (Fig. 1D), demonstrating that both raptor and rictor are required for TSC2-null cell proliferation. Thus, our data show that in cells deficient in TSC2, Rheb and mTORC1, but not mTORC2, are required for the activation of S6K1 signaling. Notably, siRNA for rictor and siRNA for raptor attenuated only TSC2-null cell proliferation, suggesting that both mTORC1 and mTORC2 are required for TSC2-null cell proliferation.
In both yeast and mammals, mTORC2 regulates actin stress fiber formation through regulation of RhoA GTPase (44, 75). We demonstrated that TSC2 loss upregulates RhoA activity and stress fiber formation through TSC1 and Rac1 (27, 31). Consistent with our published data (27, 31), TSC2 loss activated RhoA GTPase (Fig. 2A) that manifested as increased stress fiber formation (data not shown) and was inhibited by TSC2 reexpression (Fig. 2A).
To determine whether mTORC2 acts downstream of TSC2 in regulating RhoA activity, TSC2-null and LAMD cells were transfected with siRNA for rictor (Fig. 2B, top). As shown in Fig. 2B, cells transfected with control scrambled siRNA had marked Akt phosphorylation at Ser 473 (P-Ser473 Akt), indicating that mTORC2 is activated in TSC2-null and LAMD cells. Interestingly, in Tsc2−/− MEFs, P-Ser473 Akt was shown to be reduced (40), suggesting that TSC2 loss could have different effects on mTORC2 activity depending on the cell type. In TSC2-null ELT3 and LAMD cells, siRNA for rictor inhibited P-Ser473 Akt compared to control siRNA but had little effect on phospho-S6 levels.
Analysis of RhoA GTPase activity demonstrated that serum-deprived TSC2-null and LAMD cells transfected with control scrambled siRNA had marked RhoA activation (Fig. 2C), as demonstrated by our published studies (27, 31). In contrast, RhoA activity was significantly downregulated in rictor siRNA-transfected cells (Fig. 2C and D), demonstrating that mTORC2 is required for RhoA GTPase activation in cells with TSC2 loss or deficiency. Surprisingly, siRNA for rictor had little effect on P-Ser2481 mTOR in TSC2-null and LAMD cells (data not shown). In contrast, in TSC2-positive rat PAVSM cells used for comparison to human smooth-muscle-like LAMD cells, siRNA for rictor decreased both P-Ser2481 mTOR and P-Ser473 Akt, while it had little effect on P-S6 (55 and data not shown). Since P-Ser2481 mTOR manifests mTOR intrinsic catalytic activity in both mTORC1 and mTORC2 (82), mTORC1 hyperactivation in TSC2-null cells may mask the effects of siRNA for rictor on P-Ser2481 mTOR due to the experimental limitations of using siRNA in contrast to the approaches utilized in the published studies (12, 82).
To examine whether RhoA activation is required for increased proliferation of TSC2-null cells, we used C3 toxin, a cell-penetrating form of Clostridium botulinum exoenzyme that catalyzes the specific inactivation of RhoA (11). Inhibition of RhoA with C3 transferase significantly attenuated TSC2-null cell proliferation (Fig. 3A). C3 also inhibited the proliferation of primary human LAMD cells (Fig. 3B), which have increased RhoA activity (31). To directly address the role of RhoA in increased proliferation of TSC2-null cells, RhoA was downregulated with specific siRNA, followed by DNA synthesis analysis. Immunoblot analysis showed that endogenous levels of RhoA were reduced by 67 to 68% at 48 and 72 h posttransfection (Fig. 3C). The BrdU incorporation assay also showed that siRNA for RhoA had a modest but significant inhibitory effect in both TSC2-null and LAMD cells (Fig. 3D). In these experiments, DNA synthesis was measured in all cells, including both cells transfected with siRNA for RhoA and untransfected cells. This limitation of the experimental approach may account for the modest inhibitory effect of siRNA for RhoA on DNA synthesis in TSC2-null and LAMD cells. Interestingly, pharmacological inhibition of the Rho-associated kinase (ROCK), an established downstream effector of RhoA GTPase (16), with its specific inhibitor HA1077 reduced TSC2-null and LAMD cell proliferation in a concentration-dependent manner (Fig. 3E and F, respectively), demonstrating that RhoA and its downstream effector ROCK contribute to the increased proliferation of TSC2-null and LAMD cells.
Collectively, our data demonstrate that increased proliferation of cells with TSC2 loss or dysfunction is, in part, due to RhoA activation. Notably, siRNA for RhoA and C3 transferase attenuated DNA synthesis, and siRNA-dependent depletion of RhoA had little effect on mTORC1 activity, as shown by the unchanged S6 phosphorylation levels (Fig. 3C), suggesting that RhoA and mTORC1 act in parallel to promote TSC2-null cell proliferation.
Since mTORC2 is required for TSC2-null cell proliferation (Fig. 1D) and RhoA is downstream of mTORC2 (Fig. 2C and D), we next determined whether mTORC2 modulates TSC2-null cell proliferation via RhoA. TSC2-null cells were comicroinjected with siRNA for rictor and the constitutively active RhoA mutant GST-V14RhoA or control GST plasmids, and then DNA synthesis was assessed using a BrdU incorporation assay. As shown in Fig. 4A, siRNA for rictor markedly inhibited DNA synthesis in TSC2-null cells; importantly, activated V14RhoA rescued rictor siRNA-induced inhibition of DNA synthesis. These data demonstrate that RhoA acts downstream of mTORC2 in modulating TSC2-null cell proliferation. Interestingly, we also found that V14RhoA only partially reversed TSC2-induced inhibition of DNA synthesis due to reexpression of TSC2 or siRNA-induced downregulation of TSC1 (Fig. 4B and C). This evidence and our data showing that either siRNA for rictor or siRNA for raptor alone only partially inhibits DNA synthesis in TSC2-null cells (Fig. 1D) suggest that both mTORC2 and mTORC1 are necessary to rescue growth inhibition induced by TSC2 reexpression.
Because RhoA GTPase modulates cell survival in some cell types (74) and because of the modest inhibitory effect of siRNA for RhoA on TSC2-null cell proliferation, we examined whether RhoA is required for the survival of TSC2-null cells. Using two complementary approaches, depletion of endogenous RhoA with specific siRNA (Fig. 3C) and specific inhibition of Rho activity with C3 transferase, we examined TSC2-null cell apoptosis. Analysis of apoptosis with an Alexa Fluor Annexin V/Dead Cell Apoptosis kit demonstrated that control (scrambled) siRNA had little effect on cell apoptosis (Fig. 5A). In contrast, siRNA for RhoA markedly promoted apoptosis at 48 h and 72 h of transfection (Fig. 5A). Inhibition of RhoA GTPase activity with C3 transferase also promoted apoptosis of TSC2-null cells in a concentration-dependent manner (Fig. 5B). Interestingly, HA1077-dependent inhibition of ROCK, while inhibiting cell proliferation (Fig. 3E), had little effect on apoptosis in TSC2-null cells (data not shown), suggesting that downregulation of RhoA induces apoptosis in a ROCK-independent manner. Taken together, these data demonstrate that downregulation of RhoA protein levels or activity induces apoptosis and suggest that RhoA activity is required for TSC2-null cell survival.
Studies demonstrate that RhoA GTPase may regulate cell survival via Bcl2 family proteins in a p53-independent manner (13, 19, 26). We next, examined whether suppression of RhoA activity with C3 transferase modulates the levels and/or phosphorylation of Bcl2 family proteins in TSC2-null and LAMD cells. As shown in Fig. 5C, inhibition of RhoA GTPase in TSC2-null ELT3 cells markedly reduced levels of the prosurvival protein Bcl2 and increased the levels of the proapoptotic BH3-only proteins Bim and Bok. In LAMD cells, inhibition of RhoA activity also decreased Bcl2 protein levels while increasing proapoptotic Bim and Puma protein levels (Fig. 5D). Interestingly, C3 transferase had little effect on the phosphorylation of Bad in both TSC2-null and LAMD cells. Importantly, Bok, which is expressed predominantly in reproductive tissues (39), was increased in TSC2-null ELT3 cells, which are derived from rat uterine leiomyomas, suggesting that RhoA-dependent apoptosis can be cell type specific. Notably, inhibition of RhoA had little effect on mTORC1-dependent P-S6 and mTORC2-dependent P-Ser473 Akt, suggesting that RhoA acts downstream of mTORC2 in modulating Bcl2 family protein levels, potentially in parallel with mTORC2-Akt signaling. Taken together, these data demonstrate that RhoA GTPase modulates the survival of cells with TSC2 loss and deficiency via Bcl2 family proteins.
To determine whether p53 is involved in RhoA-dependent modulation of TSC2-null cell survival, we performed immunoblot analysis of LAMD cells treated with C3 transferase with phospho-Ser46 p53 (P-Ser46 p53) antibody and found that inhibition of RhoA GTPase had little effect on P-p53 and p53 protein levels (data not shown). Next, we examined C3 transferase-induced apoptosis in Tsc2−/− and Tsc2+/+ MEFs compared to Tsc2−/− p53−/− and TSC2+/+ p53−/− MEFs with p53 deleted (94) using flow cytometry analysis with annexin V antibody. We found that all diluent-treated cells had low levels of apoptosis (Fig. 5E). Inhibition of RhoA increased apoptosis by 15 to 20% in Tsc2−/− and Tsc2+/+ MEFs at levels comparable to those of Tsc2+/+ p53−/− MEFs (Fig. 5E), suggesting that either TSC2 or p53 loss alone has little effect on basal and C3 transferase-induced apoptosis. In contrast, inhibition of RhoA in Tsc2−/− p53−/− MEFs markedly increased apoptosis compared to Tsc2−/−, Tsc2+/+, and Tsc2+/+ p53−/− MEFs (Fig. 5E), demonstrating that loss of both TSC2 and p53 augments cell sensitivity to RhoA inhibition. These data suggest that p53 is necessary for RhoA-dependent Tsc2−/− p53−/− MEF survival but that these effects maybe cell type specific.
Simvastatin inhibits proliferation and induces apoptosis in different cell types through inhibition of RhoA GTPase activity by preventing RhoA geranylgeranylation (4, 23, 84). To directly examine the effect of simvastatin on RhoA GTPase activity, we performed a RhoA activation assay in TSC2-null and LAMD cells treated with different concentrations of simvastatin. As seen in Fig. 6A and B, respectively, simvastatin inhibited RhoA GTPase activity in TSC2-null and primary human LAMD cells in a concentration-dependent manner. Because RhoA activity is required for cell migration and to confirm that simvastatin-dependent RhoA inhibition is functionally significant for LAMD cells, the migration assay was performed under the same experimental conditions. We found that simvastatin inhibited LAMD cell migration in a concentration-dependent manner that correlated with its effects on RhoA activity (data not shown). Since simvastatin inhibits proliferation in different types of cells, including cancer and smooth muscle cells (1, 4, 7), and our data show that RhoA is required for TSC2-null cell proliferation (Fig. 3), we examined the effects of simvastatin alone and in combination with rapamycin on the proliferation of TSC2-null and LAMD cells that were treated with different concentrations of simvastatin, 20 nM rapamycin, or diluent separately or in combination. As shown in Fig. 7A, simvastatin inhibited DNA synthesis in TSC2-null cells in a concentration-dependent manner. In agreement with our previous studies (28), 20 nM rapamycin alone markedly inhibited the DNA synthesis. Combined treatment with simvastatin and rapamycin further inhibited DNA synthesis of TSC2-null cells compared to treatment with rapamycin alone (Fig. 7A, black bars). Similarly, in primary human LAMD cells, simvastatin alone attenuated DNA synthesis in a concentration-dependent manner and markedly enhanced the growth-inhibitory effect of rapamycin (Fig. 7B). Similar results were obtained using cell counts: LAMD cell growth in the presence of 0.3 and 0.5 μM simvastatin was markedly attenuated in a concentration-dependent manner compared to diluent-treated cells (Fig. 7C). Maintenance in media supplemented with 20 nM rapamycin inhibited LAMD cell growth, and simvastatin-rapamycin cotreatment significantly enhanced the inhibitory effects of either rapamycin or simvastatin given separately (Fig. 7D). Taken together, these data demonstrate that simvastatin alone attenuates TSC2-null and LAMD cell proliferation and cooperates with rapamycin in the inhibition of cell growth caused by TSC2 loss or dysfunction.
Next, we examined whether simvastatin promotes apoptosis in TSC2-null cells, since our data demonstrate that RhoA is required for TSC2-null cell survival (Fig. 5) and that simvastatin inhibits RhoA activation (Fig. 6). As shown in Fig. 8A and B, simvastatin induced apoptosis in TSC2-null cells in a concentration-dependent manner. Rapamycin, as we reported previously (29), had little effect on the number of apoptotic cells compared to diluent-treated cells (Fig. 8B). Interestingly, cotreatment with rapamycin modestly increased simvastatin-induced apoptosis (Fig. 8B). These data demonstrate that simvastatin induces apoptosis in TSC2-null cells.
To determine whether simvastatin promotes apoptosis by inhibiting RhoA GTPase, we transfected TSC2-null cells with constitutively active V14RhoA, and then the cells were treated with simvastatin, followed by assessment of cell apoptosis using immunostaining with cleaved caspase 3 antibody. As seen in Fig. 8C and D, simvastatin promoted a marked increase in cleaved caspase 3 immunostaining. Importantly, cells transfected with constitutively activated V14RhoA stained negative for cleaved caspase 3 antibody, demonstrating that activated RhoA rescues simvastatin-induced apoptosis in TSC2-null cells (Fig. 8D). Collectively, these data suggest that simvastatin induces apoptosis via inhibition of RhoA GTPase through cleavage of caspase 3 in TSC2-null cells.
To examine whether our in vitro findings have in vivo relevance, we examined the effects of simvastatin alone or in combination with low-dose rapamycin on xenographic TSC2-null tumor growth in nude mice. NCRNU-M athymic nude mice were injected subcutaneously with 5 × 106 TSC2-null ELT3 cells. Tumors reached 5 mm in diameter in 4 to 6 weeks; then, the animals were treated with simvastatin (250 mg/kg/day; special diet), rapamycin (1 mg/kg; i.p. injections 3 times a week), or a combination of simvastatin and rapamycin for 50 days. A schematic representation of the experimental design is shown in Fig. 9A. The dose levels and treatment schedule for rapamycin were selected on the basis of previously verified pharmacokinetics and the doses of rapamycin approved for immunosuppression after organ transplantation (67), clinical trials (3), and dose-response rodent studies (41). Notably, the doses of rapamycin used in this study were 4 to 10 times lower than those in previous reports (59–61, 68, 72). The treatment dose and administration for simvastatin were selected based on published studies as the highest well-tolerated dose that had also shown biological activity in rodent models (48, 79, 90). No significant changes in general behavior and weight were observed during the treatment period and after treatment was terminated (animal weight by day 50 of the experiment was 25.49 ± 0.41 g for simvastatin-treated animals and 25.98 ± 1.35 g for animals on a regular diet). To determine the effects of simvastatin and rapamycin on TSC2-null tumor growth, tumor size was monitored three times a week by measurement with calipers. Animals were sacrificed when the tumors reached 10 mm in diameter; the date of sacrifice was also used as the date of termination for survival analysis.
Analysis of tumor-bearing mouse survival demonstrated that only 15% of control animals survived by day 50 of the experiment (Fig. 9B). Simvastatin and rapamycin used alone increased animal survival by 47.64% and 50.0%, respectively (Fig. 9B). Importantly, combined treatment with rapamycin and simvastatin markedly improved survival compared to animals treated with simvastatin and rapamycin singly. Thus, after 50 days of combined treatment, animal survival was 67.0%, which was more than 4 times higher than the control group (Fig. 9B).
Analysis of TSC2-null tumor growth demonstrated marked tumor growth in control (untreated) mice. Thus, tumor volumes doubled by day 6 and showed an approximately 3-fold increase by day 13 from the beginning of the experiment (Fig. 9C). During the first 40 days of treatment, rapamycin and simvastatin had marked inhibitory effects on tumor growth compared to control animals, showing an approximately 3.5-fold increase in tumor volume during the same period (Table 1). The combination of simvastatin and rapamycin reduced tumor volume by 4 weeks of treatment. Interestingly, both simvastatin and rapamycin as single agents abrogated tumor growth at approximately day 45 (Fig. 9C).
Taken together, these data demonstrate that rapamycin and simvastatin alone inhibited TSC2-null tumor growth in nude mice and increased animal survival and that combined treatment with simvastatin and rapamycin abrogated tumor growth and improved animal survival compared to control and single-agent treatments.
To examine the effects of rapamycin and simvastatin withdrawal on TSC2-null tumor growth, mice that were treated with rapamycin and simvastatin separately or in combination for 50 days as described above and had visible tumor disappearance (5 animals per treatment condition) were subjected to treatment withdrawal and were monitored three times a week. Three of five animals from the rapamycin-treated group demonstrated flank tumor recurrence after treatment withdrawal. Initial tumor recurrence was observed within a week after final rapamycin administration, and during the next 41 days, tumors reached 10% of animal body weight (Fig. 9D and E). Importantly, no tumor recurrence was detected in simvastatin- and simvastatin-rapamycin-treated animals during the 9 months after treatment withdrawal (Fig. 9D and E). Collectively, these data demonstrate that the combination of rapamycin with simvastatin prevents tumor regrowth after treatment withdrawal.
Because our in vitro data demonstrate that simvastatin promotes apoptosis in TSC2-null ELT3 and LAMD cells, we performed apoptosis analysis of tumor tissues collected at days 0, 10, and 20 of treatment using a TUNEL-based In Situ Cell Death Detection Kit. As seen in Fig. 10A and B, little apoptosis was detected in control and rapamycin-treated tumors. In contrast, marked apoptosis was detected in the simvastatin- and simvastatin-rapamycin-treated tumors at days 10 and 20 of treatment (Fig. 10A and B). Similarly, marked apoptosis was detected in simvastatin- and simvastatin-rapamycin-treated tumors at days 34 and 32 of treatment, respectively, while tumors from mice treated with rapamycin alone for 41 days had low levels of apoptosis that were comparable to those of untreated tumors collected at day 45 of the experiment (Fig. 11). Taken together, these data demonstrate that simvastatin promotes apoptosis in TSC2-null tumors.
To examine the effects of simvastatin and rapamycin on TSC2-null cell growth in vivo, proliferation of TSC2-null cells within tumors was examined by immunohistochemical analysis with anti-Ki-67 antibodies. As shown in Fig. 10C and D and and11,11, tumor tissues of control animals had marked proliferation rates, which persisted to days 10, 20, and 45 of the experiment. At day 10 of treatment, rapamycin significantly inhibited cell proliferation, while simvastatin had little effect. At day 20, however, both simvastatin and rapamycin inhibited tumor cell growth (Fig. 10C and D). Importantly, combined treatment of simvastatin with rapamycin markedly reduced proliferation in a time-dependent manner compared to rapamycin or simvastatin treatment alone (Fig. 10C and D). Our data demonstrate that simvastatin and rapamycin cooperate in inhibiting the proliferation of TSC2-null cells in vivo.
Because rapamycin inhibits mTORC1 activity in different cell and animal models, we examined the activation state of mTORC1 by the phosphorylation levels of ribosomal protein S6, the molecular marker of mTORC1 activation, using two different techniques, immunocytochemistry and immunoblot analysis. Analysis of tumor sections with P-S6 antibodies demonstrated that TSC2-null tumors had marked P-S6 levels prior to treatment (day 0) and control (untreated) tumors maintained high P-S6 levels through days 1, 10, 20, and 45 of the experiment (Fig. 11 and and12A12A and B). A low dose of rapamycin (1 mg/kg) induced a marked reduction in P-S6 compared to untreated tumors in a time-dependent manner (Fig. 11 and and12).12). Simvastatin alone had little effect on P-S6 and did not modulate the rapamycin-dependent inhibition of P-S6 in cotreated tumors (Fig. 11 and and12).12). Immunoblot analysis performed using tissue samples collected at days 15, 30, and 40 of the experiment also demonstrated a marked increase in P-S6 in control tumors at days 15 and 30 (Fig. 12C). Consistent with immunohistochemical data, rapamycin decreased P-S6 at days 15, 30, and 40; simvastatin had little effect on P-S6; and combined rapamycin- and rapamycin-simvastatin-treated tumors showed decreases in P-S6 comparable to those of tumors treated with rapamycin alone (Fig. 12C and D, left). These data demonstrate that rapamycin, but not simvastatin, inhibits mTORC1 signaling in TSC2-null tumors.
Since simvastatin may promote apoptosis in an Akt-dependent manner (25, 42, 52), we examined P-Ser473 Akt in tumor tissue samples. As shown in Fig. 12C, P-Ser473 Akt was detected in control untreated tumors, which correlates with Akt activation in vitro (Fig. 2B). In agreement with previously published studies (5, 83), rapamycin increased P-Ser473 Akt in both rapamycin and rapamycin-simvastatin-treated tumors (Fig. 12C and D, right). Levels of P-Ser473 Akt in simvastatin-treated tumors, however, were comparable to those in control tumors (Fig. 12C and D, right), demonstrating that simvastatin has little effect on P-Ser473 Akt. These data suggest that the growth-inhibitory and proapoptotic effects of simvastatin are likely Akt independent.
In summary, these data collectively demonstrate that simvastatin and rapamycin inhibit TSC2-null tumor growth through different signaling mechanisms. Rapamycin inhibits cell proliferation through the suppression of mTORC1 signaling but has little effect on apoptosis; simvastatin inhibits TSC2-null cell proliferation and induces apoptosis but has little effect on mTORC1 activity. Importantly, combined treatment with simvastatin and rapamycin induces apoptosis, abrogates proliferation, inhibits mTORC1 signaling in TSC2-null tumors, abrogates TSC2-null tumor growth, and prevents tumor recurrence after simvastatin and rapamycin withdrawal.
Increased proliferation of atypical smooth muscle cells in pulmonary LAM and in kidney tumors in TS occurs due to mutational inactivation of tumor suppressor TSC2 and mTORC1 activation (54). This study demonstrates that TSC2-dependent increased cell proliferation and survival also requires mTORC2 and its downstream effector RhoA GTPase. Thus, inhibition of mTORC1 and mTORC2 signaling with siRNAs for raptor and rictor, respectively, had comparable inhibitory effects on TSC2-null cell growth, demonstrating that both mTORC1 and mTORC2 are required for TSC2-null cell proliferation. Importantly, siRNA for rictor inhibited both increased P-Ser473 Akt and RhoA activity, demonstrating mTORC2-dependent regulation of Akt and RhoA GTPase in TSC2-null and LAMD cells. We also show that RhoA activity is necessary for TSC2-null cell survival and that inhibition of RhoA promotes apoptosis through downregulation of Bcl2 and upregulation of the proapoptotic proteins Bim, Bok, and Puma. In vivo, the combination of the proapoptotic activity of simvastatin with the cytostatic effect of rapamycin abrogated xenographic TSC2-null tumor growth in nude mice, improved animal survival, and prevented tumor recurrence after treatment withdrawal.
While the critical role of mTORC1 signaling in TSC2-deficient cell and tumor growth makes it a logical therapeutic target for LAM and TS, recent preclinical and clinical studies have demonstrated that the mTORC1 inhibitor rapamycin and its analogs have a primarily cytostatic effect in TSC2-deficient cells and tumors and that termination of treatment results in disease reversal (3, 72). Thus, data from rapamycin (sirolimus) clinical trials for patients with pulmonary LAM and TS (3, 14) show marked regression in renal angiomyolipoma volume during rapamycin therapy (3, 14). However, approximately 1 year after treatment withdrawal, tumors reappeared and reached approximately 86% of their original volume (3). Similarly, the rapamycin analog RAD001 attenuated renal tumor development in TSC2+/− mice, and its withdrawal led to marked tumor regrowth (72). Our data demonstrate that 50-day rapamycin treatment in doses that were 4 to 10 times lower than those used in previously published reports (1 mg/kg versus 4 to 10 mg/kg) (59–61, 68, 72) had growth-inhibitory effects on TSC2-null tumors in nude mice and suppressed mTORC1 signaling without induction of apoptosis. Not surprisingly, rapamycin withdrawal resulted in tumor recurrence within 1 week after the last treatment, with tumor regrowth reaching 10% of the animal body weight by day 41. Thus, our findings further demonstrate that rapamycin as a single agent has limitations in inhibiting TSC2-null tumor growth and suggest that induction of apoptosis in TSC2-null tumors should have therapeutic benefits.
Lamb and colleagues showed that TSC1, which forms a tumor suppressor complex with TSC2, promotes stress fiber formation through activation of RhoA GTPase (56). We demonstrate that TSC2 loss induces TSC1-dependent stress fiber formation due to activation of RhoA GTPase that is rapamycin insensitive (27, 31). Rapamycin-insensitive mTORC2 regulates the actin cytoskeleton and Rho GTPase activity (44, 75), suggesting that mTORC2 may play a role in modulating Rho activity and Rho-dependent stress fiber formation due to TSC2 loss. Here, we demonstrate that the mTOR and rictor, but not raptor, are required for stress fiber formation in TSC2-null cells and that rictor regulates the activity of RhoA GTPase. Activated Rho rescued rictor-induced stress fiber disassembly in TSC2-null cells, demonstrating that rictor acts upstream of RhoA in regulating its activity and stress fiber formation due to TSC2 loss.
Rho GTPases have prosurvival and antiproliferative effects in different diseases, including human cancers (47, 74). We show that RhoA GTPase is necessary for TSC2-null cell proliferation and survival, and either RhoA knockdown or specific inhibition of Rho activity significantly inhibits proliferation and induces apoptosis in TSC2-null and LAMD cells in an mTORC1-independent manner. Importantly, inhibition of mTORC2 signaling with siRNA for rictor markedly inhibited TSC2-null cell proliferation at levels comparable to those of siRNA for raptor, while it had little effect on mTORC1-dependent S6 phosphorylation. Importantly, constitutively active Rho rescued rictor siRNA-induced inhibition of proliferation, suggesting that mTORC2 modulates TSC2-null cell proliferation via Rho GTPase. Thus, our data show that TSC2 loss, in addition to activation of mTORC1, leads to mTORC2-dependent activation of RhoA GTPase, which acts as a prosurvival molecule and is required for the proliferation of TSC2-null cells. Thus, these data suggest that therapeutic targeting of RhoA may inhibit abnormal cell growth due to TSC2 loss and promote apoptosis in TSC2-null cells.
Extensive efforts have been made to develop selective Rho inhibitors; however, to date, none have been approved for clinical use (74). Rho GTPases require prenylation (geranylgeranylation) for membrane binding and activation. Statins, which are HMG-CoA reductase inhibitors, suppress GGPP production, leading to nonselective inhibition of Rho GTPases. Statins, which are widely used clinically to reduce cholesterol levels (89), also have an oncoprotective effect in humans (15, 49, 70, 91) and inhibit proliferation and promote apoptosis in different cancer cells (4, 10, 23); a natural statin, simvastatin, induces apoptosis in numerous human cancer cell lines via suppression of RhoA GTPase activity (10, 23, 92).
We show that simvastatin inhibited Rho GTPase activity, attenuated proliferation, induced cleavage of caspase 3, and promoted apoptosis in TSC2-null cells. Importantly, the constitutively active Rho prevented simvastatin-induced caspase 3 cleavage, suggesting that simvastatin inhibits TSC2-related cell proliferation and promotes apoptosis via inhibition of Rho GTPase. In TSC2-null and LAMD cells, the RhoA-dependent apoptotic response involves downregulation of antiapoptotic Bcl2 and upregulation of proapoptotic Bim, Bok, and Puma. Further, simvastatin attenuated TSC2-null subcutaneous tumor growth in nude mice and prolonged animal survival. Interestingly, simvastatin-dependent inhibition of tumor growth, which was observed after 12 days of simvastatin treatment, correlates with statin pharmacokinetic studies (62). Importantly, simvastatin promoted apoptosis and attenuated proliferation while it had little effect on mTORC1 signaling, confirming that the effects of simvastatin are mTORC1 independent. Although a relatively high dose of simvastatin was used in this study, because mice metabolize simvastatin more rapidly than humans, lower doses may be effective in humans (48).
Consistent with our in vivo findings, simvastatin inhibited proliferation, induced apoptosis, and cooperated with rapamycin in the inhibition of TSC2-null ELT3 cell growth in vitro. Importantly, similar data were obtained for primary LAMD cells dissociated from the nodules from the lungs of LAM patients (30, 32), suggesting that our findings may be applicable to LAM disease. The concentrations of simvastatin that inhibited growth and induced apoptosis in TSC2-null ELT3 and LAMD cells (0.3 to 1 μM) were comparable to the plasma levels of statins achieved in humans in clinical trials (2.32 ± 1.27 μM at peak concentration) (86) and did not result in severe drug toxicity in the patients (91). Because rapamycin elevates triglycerides and total cholesterol levels (69), statins are routinely used in rapamycin-treated transplant patients to decrease the risk of cardiovascular disease and prolong survival (2, 78), and studies show no significant pharmacokinetic drug-drug interactions between rapamycin and statins (53, 96).
We demonstrate that simvastatin and rapamycin cooperated in inhibiting TSC2-null and LAMD cell proliferation in vitro, in reducing TSC2-null tumor size, and in improving tumor-bearing mouse survival in vivo. The combination of simvastatin and rapamycin induced apoptosis, inhibited mTORC1 signaling, and abrogated DNA synthesis in TSC2-null tumors. Importantly, in contrast to rapamycin-treated mice, which demonstrated marked tumor recurrence after rapamycin withdrawal, no tumor regrowth was detected in simvastatin- and simvastatin-rapamycin-treated mice during 9 months of observation following the last treatment. Taken together, these data demonstrate that the combination of cytostatic rapamycin with proapoptotic simvastatin improves survival, suppresses tumor growth, and prevents tumor regrowth upon treatment withdrawal.
Collectively, our data demonstrate that mTORC2-dependent RhoA GTPase activation is necessary for TSC2-null cell growth and survival. In vivo targeting of RhoA GTPase with simvastatin and mTORC1 with rapamycin abrogates TSC2-null tumor growth, induces apoptosis, increases tumor-bearing mouse survival, and prevents posttreatment tumor regrowth. Our findings have important implications, because they show RhoA GTPase is a potential therapeutic target for combinational therapy in diseases associated with TSC2 dysfunction.
We thank our colleagues Yvonne Paterson, Reynold A. Panettieri, Jr., and Andrew Eszterhas for critical reading of the manuscript and helpful discussions; Margaret M. Chou for generously providing us with GST-V14Rho or control GST plasmids; Cheryl L. Walker, M. D. Anderson Cancer Center, University of Texas, Smithville, TX, for ELT3 cells; David Kwiatkowski, Brigham and Women's Hospital, Harvard University, Boston, MA, for Tsc2−/−, Tsc2+/+, Tsc2+/+ p53−/−, and Tsc2−/− p53−/− MEFs; and the National Disease Research Interchange (NDRI) for providing us with LAM tissue.
This work is supported by grants from the NIH/NHLBI (2RO1HL071106 [V.P.K.], RO1HL090829 [V.P.K.], and ALA CI-9813-N [V.P.K.]) and the LAM Foundation (V.P.K.), by Abramson Cancer Center Core Support Grant NIH P130-CA-016520-34 (V.P.K.), and by ATS/LAM Foundation research grant LAM-07-001 (E.A.G.) and in part by a Gilead Science Research Scholars grant (E.A.G.). E.A.G. is a Parker B. Francis Fellow in Pulmonary Research.
Published ahead of print on 11 April 2011.