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Agonists Induce Partial Reversion of Epithelial-Mesenchymal Transition in Anaplastic Thyroid Cancer Cells
Dipartimento di Medicina Clinica e Sperimentale (A.A., A.M., A.S., A.B.), Cattedra di Endocrinologia, University of Catanzaro, 88100 Catanzaro, Italy; Dipartimento di Scienze Biomediche (E.C.), University of Catania, 95125 Catania, Italy; and Dipartimento di Medicina Interna e Medicine Specialistiche (G.P., F.F., M.G., R.V.), Cattedra di Endocrinologia, University of Catania, Ospedale Garibaldi, 95123 Catania, Italy
Address all correspondence and requests for reprints to: Antonino Belfiore, M.D., Dipartimento di Medicina Sperimentale e Clinica, Cattedra di Endocrinologia, Università di Catanzaro, v.le Europa, Loc. Germaneto, 88100 Catanzaro, Italy. E-mail: belfiore{at}unicz.it.
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Abstract |
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agonists have recently emerged as potential antineoplastic drugs. To establish whether ATC could be a target of PPAR agonists, we first examined PPAR protein expression in a panel of six ATC cell lines and then studied the biologic effects of two PPAR agonists, ciglitazone and rosiglitazone, that belong to the class of thiazolidonediones. PPAR protein was present and functional in all ATC cell lines. Both ciglitazone and rosiglitazone showed complex biological effects in ATC cells, including inhibition of anchorage-dependent and -independent growth and migration, and increased apoptosis rate. Rosiglitazone-induced growth inhibition was associated with cell cycle arrest and changes in cell cycle regulators, such as an increase of cyclin-dependent kinases inhibitors p21cip1 and p27kip1, a decrease of cyclin D1, and inactivation of Rb protein. Rosiglitazone-induced apoptosis was associated with a decrease of Bcl-XL expression and caspase-3 and -7 activation. Moreover, rosiglitazone antagonized IGF-I biological effects by up-regulating phosphatase and tensin homolog deleted from chromosome 10 with subsequent inhibition of the phosphatidylinositol 3-kinase/Akt signaling pathway. Finally, rosiglitazone increased the expression of thyroid-specific differentiation markers. In conclusions, these data suggest that PPAR agonists induce a partial reversion of the epithelial mesenchymal transition in ATC cells by multiple mechanisms. PPAR agonists may, therefore, have a role in the multimodal therapy currently used to slow down ATC growth and dissemination. |
Introduction |
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Epidemiological and histopathological evidences suggest that ATC may arise from dedifferentiation of long-standing, well-differentiated thyroid tumors. Accordingly, anaplastic cancer cells have lost the ability to uptake iodine, which is a hallmark of normal thyroid follicular cells and well-differentiated thyroid carcinomas. This has therapeutical implications because radiometabolic therapy with 131I is useful for residual or metastatic well-differentiated thyroid cancer but not ATC. It is clear, therefore, that new molecular targets need to be identified for an effective therapeutical strategy of ATC.
Although ATC cells originate from thyroid follicular epithelial cells, they are characterized by a mesenchymal phenotype, as revealed by spindle-shaped cells and absent or reduced levels of E-cadherin and other markers of thyroid cell differentiation [thyroglobulin, sodium-iodine symporter (NIS), thyroperoxidase, and TSH receptor (TSH-R)]. Epithelial-mesenchymal transition (3) is typical of dedifferentiated tumors and is associated with high invasion and dissemination potential (4). Typically, epithelial mesenchymal transition (EMT) is particularly pronounced in ATC.
Also the activation of the IGF system plays a role in EMT (3), and we recently observed that the IGF system is up-regulated in thyroid cancer (5, 6). In ATC this is partially due to the activation of an autocrine loop involving overexpression of the insulin receptor isoform A and enhanced autocrine production of IGF-II, which binds insulin receptor isoform A with high affinity and stimulates growth, resistance to apoptosis, and cell migration (5).
Recently agonists of peroxisomal proliferator-activated receptor (PPAR)- have emerged as promising tumor response modifiers. PPAR is a ligand-activated transcription factor that belongs to the steroid receptor superfamily and is predominantly expressed in adipose cells in which it is a major inducer of differentiation and adipogenesis (7). Specific ligands to this factor have been identified such as the prostanoid 15d-PGJ2 and the antidiabetic agents thiazolidinediones (TZDs) (8). PPAR agonists induce a variety of favorable changes (growth arrest, apoptosis, and/or partial redifferentiation) in certain malignancies, including liposarcoma and cancers of the breast, colon, pancreas, and prostate (9, 10, 11, 12, 13, 14, 15, 16, 17). Interestingly recent studies have shown the presence of a paired box gene 8-PPAR rearrangement in a subgroup of follicular thyroid cancers. The resulting fusion protein inhibits PPAR activation in a dominant-negative manner and has an oncogenic role (18). Moreover, it has been recently shown that PPAR insufficiency promotes thyroid carcinogenesis in mice harboring a knockin mutant thyroid hormone ß-receptor, which spontaneously develop follicular thyroid carcinomas (19).
These studies may suggest, therefore, an antioncogenic role of PPAR in the thyroid. Moreover, PPAR forms heterodimers with the retinoid X receptor. PPAR agonists may therefore synergize with retinoids, such as all-trans retinoic acid, which induces growth arrest and partial redifferentiation in various cancer cells, including poorly differentiated thyroid cancer cells (20). Phase II studies using 13-cis-retinoic acid in patients with poorly differentiated thyroid cancer have shown induction of a partial redifferentiation in approximately 40% of cases (21).
To establish whether PPAR agonists might be a useful adjuvant treatment in ATC, we first examined the expression of PPAR protein in six ATC cell lines and also studied the biologic effects of two PPAR agonists, ciglitazone and rosiglitazone, both belonging to the class of antidiabetic agents TZDs. We used rosiglitazone at doses comparable with those reached during diabetes therapy. We found that both these TZDs have complex biological effects in ATC cells, including reduction of anchorage-dependent and -independent growth and migration, and increased apoptosis rate. Moreover, rosiglitazone antagonized, at least in part, the biological effects of IGF-I by up-regulating phosphatase and tensin homolog deleted from chromosome 10 (PTEN) and consequently inhibiting the phosphatidylinositol 3-kinase (PI3K)/Akt signaling pathway. It also potentiated the antitumor effect of doxorubicin and induced a partial redifferentiation in ATC cells.
Taken together, these effects confirm the antitumor potential of these agents in ATC cells, with a partial reversion of EMT. These observations suggest a possible role of PPAR agonists in the multimodal therapy for ATC.
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Materials and Methods |
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antibody (PA3–821) was obtained by Affinity Bioreagents (Deerfield, IL), and anti-PTEN antibody was obtained by Santa Cruz Biotechnology (Santa Cruz, CA). Antiphospho antibodies to Ser473 and Thr308 of Akt were from Cell Signaling (Beverly, MA). Rosiglitazone was kindly provided by Glaxo Smith Kline (Greenville, NC). Ciglitazone was purchased by Alexis Biochemicals (Lausen, Switzerland). The firefly luciferase vector driven by the PPAR-sensitive peroxisome proliferator response element (PPRE)-3 sequences, mouse mammary tumor virus, was kindly provided by R. Evans (Salk Institute, La Jolla, CA).
Cells
Human ATC cells (FF-1, ARO, HTh74, C643, SW1736, 8305c) and human papillary cancer cells (TPC-1 and NPA) were studied. FF-1 cell line was established in our laboratory as follows. A surgical specimen from an anaplastic thyroid cancer was initially fragmented with a scalpel and subsequently digested for 1 h in DMEM low glucose (Sigma) containing 1 mg/ml type V collagenase (Sigma) in a 37 C water bath with agitation. The cell suspension was then centrifuged and the pellet resuspended in complete RPMI 1640. Cells were then subsequently cultured with RPMI 1640 containing increasing doses (from 1 to 10%) of stripped FCS containing 100 µg/ml penicillin and streptomycin and 2 mM glutamine. Cultures were analyzed by immunofluorescence using an antibody against cytokeratin (Novocastra Laboratories, Newcastle Upon Tyne, UK) that stains exclusively epithelial cells.
ARO and TPC-1 were obtained from Drs. A. Fusco and M. Santoro (Università Federico II, Naples, Italy) and grown in RPMI 1640 (Sigma). HTh74, C643, and SW1736 cells were provided by Dr. N. E. Heldin (Uppsala University Hospital, Uppsala, Sweden), and 8305c cells were purchased from ECACC (Salisbury, UK): these cells were grown in MEM. All media were supplemented with 2 mM glutamine, 10% fetal bovine serum, and 100 µg/ml penicillin and streptomycin.
Viability assay
Cell viability was measured by the dimethylthiazoldiphenyltetra-zoliumbromide (MTT) test: cells (2 x 103/well) were seeded in 96-well plates in 10% FCS-containing medium. After 24 h the medium was removed and replaced with FCS-free medium containing BSA 0.1%. After an additional 24 h, either ciglitazone or rosiglitazone was added at the indicated doses. Parallel experiments were carried out in cells maintained in medium containing 10% FCS. To evaluate whether rosiglitazone affected cell viability regulation by IGF-I, serum-starved cells, incubated in the presence or absence of 10 µM rosiglitazone, were stimulated with 10 nM IGF-I. Cell viability was measured 5 d later by measuring the rate of tetrazolium salts reduction to formazan (MTT, Amersham Life Science, Buckinghamshire, UK), which is proportional to the number of living cells (22). At the end of incubation, the absorbance was read at 540 nm.
Cell growth studies
Cell growth was measured by [3H]thymidine incorporation: cells (3 x 104/well) were plated in 24-well tissue culture plates and grown in their regular medium for 24 h. The medium was then replaced with fresh medium containing 0.1% BSA. Twenty-four hours later, either ciglitazone or rosiglitazone at the indicated doses was added. In parallel experiments the growth effect of 10 nM IGF-I was assessed in the presence or absence of each PPAR agonist. After 24 h, 18.5 kBq/well of [3H]thymidine were added for 4 h. Cell monolayers were then washed twice with cold buffer, incubated with cold 10% trichloroacetic acid solution for 30 min, solubilized with 0.1 N NaOH, and counted by liquid scintillation in a ß-counter.
Migration assay
Cell migration was measured with the Boyden’s chamber technique. Cells (105/well), resuspended in 200 µl medium, were placed on 6.5-mm-diameter polycarbonate filters (8 µm pore size; Corning Costar Corp., Cambridge, MA) containing polycarbonate membranes coated at the lower side with 250 µg/ml collagen IV. Cells were incubated in the presence or absence of 10 µM rosiglitazone and allowed to migrate to the underside of the top chamber for 18 h. IGF-I (10 nM) was added to the lower chamber as chemoattractant.
At the end of incubation, the cells at the upper side of the filter were removed with a cotton swab. Cells that had migrated to the lower side of the filter were fixed with 11% gluteraldeyde for 15 min at room temperature and stained with 0.1% crystal violet in 20% methanol for 20'. After three washes with water and complete drying, the crystal violet was solubilized by immersion of the filters in 10% acetic acid. The concentration of the solubilized crystal violet was evaluated as absorbance at 590 nm. Cell migration was expressed as percent of migrated cells over untreated controls.
Cell cycle and apoptosis evaluation
Cells were cultured in the absence or presence of rosiglitazone and synchronized by culturing them for 24 h in leucine-deprived medium, which induces a block at the G1 restriction point. Cells were then released from the cell cycle block by the addition of regular medium and harvested at 12 h from release. Permeabilized cells were centrifuged and resuspended in PBS containing 20 mg/ml propidium iodide (PI) plus 40 mg/ml RNase (Sigma) for 30 min in the dark. Cells were then subjected to fluorescence-activated cell sorter analysis (Coulter Elite flow cytometer, Beckman Coulter, Milan, Italy) and gated for PI. Sub-G1 cells were scored as apoptotic.
For annexin staining, cells were washed twice with cold PBS and then resuspended in 1 x binding buffer at a concentration of 1 x 106 cells/ml. Cells (1 x 105 cells) were then transferred to a 5-ml tube and incubated with 5 µl annexin V-fluorescein isothiocyanate and 10 µl PI for 15 min at room temperature (20–25 C) in the dark. After the addition of 400 µl of 1 x binding buffer to each tube, annexin-positive cells were scored under a fluorescence microscope. Values obtained were expressed as percent of annexin-positive cells over the total scored cell population.
Apoptosis was also determined with the cell death detection ELISA PLUS kit (Roche Diagnostics), a photometric enzyme immunoassay for the qualitative and quantitative in vitro determination of cytoplasmic histone-associated DNA fragments after induced cell death. Briefly, cells (2 x 104/well) were plated in 12-well tissue culture plates and grown in their regular medium for 24 h. The medium was then replaced with fresh medium containing either 20 µM ciglitazone or 10 µM rosiglitazone for 48 h. At the end of incubation, the absorbance was read at 405 nm.
Assay for caspase-3/7 activity
Caspase-3/7 activity was measured by Apo-ONE homogeneous caspase-3/7 assay (Promega), according to the manufacturer’s instructions. Briefly, 9 x 104 cells were seeded in 35-mm dishes and incubated in the presence or absence of 10 µM rosiglitazone for 48 h. For the assay, 30 µl of solubilized 2 x 105 cells/ml were added to a 96-well plate and incubated with caspase-3/7 Z-DEVD-R110 substrate. The increase of fluorescence was measured every hour in a 8-h period using a spectrofluorimeter Wallac 1420 VICTOR (PerkinElmer, Boston, MA), with an excitation wavelength of 485 nm and an emission wavelength of 535 nm. The assay was carried out in the presence or absence of the caspase inhibitor N-benzyloxycarbonyl-Val-Ala-Asp-fluoromethylketone (Z-VAD-fmk) at the concentration of 30 µM.
Western blot analysis
Cells were grown to 70% confluency and then incubated in the presence or absence of 10 µM rosiglitazone for 48 h. For phospho-Akt analysis, cells were stimulated with 10 nM IGF-I for 10 min. At the end of the incubation, cells were washed with ice-cold PBS and lysed in Laemli buffer containing 62.5 mM Tris-HCl (pH 6.8), 2% sodium dodecyl sulfate, and 9.9% glycerol. Protein concentration was determined by BCA protein assay kit (Pierce, Rockford, IL). For analysis of other proteins, cells were lysed in cold radioimmunoprecipitation assay buffer containing 50 mM Tris (pH 7.4), 150 mM NaCl, 1% Triton X-100, 0.25% sodium deoxycolate, 10 mM sodium pyrophosphate, 1 mM NaF, 1 mM sodium orthovanadate, 2 mM phenylmethylsulfonyl fluoride, 10 µg/ml aprotinin, 10 µg/ml pepstatin, and 10 µg/ml leupeptin. After collection, samples were rotated for 15 min at 4 C. Insoluble material was separated from the soluble extract by centrifugation at 10,000 x g for 10 min at 4 C. Protein concentration was determined by the Bradford assay. Cell lysates were subjected to SDS-PAGE. The resolved proteins were transferred to nitrocellulose membranes and immunoblotted using 1 µg/ml of the indicated antibodies. All immunoblots were analyzed by enhanced chemiluminescence (Amersham, Little Chalfont, UK), autoradiographed, and subjected to densitometric analysis. When appropriate, the nitrocellulose membranes were stripped with buffer Restore (Pierce) for 15 min at room temperature and subsequently reprobed with an anti-ß-actin mouse monoclonal antibody. Densitometric results were corrected for protein loading by normalization for ß-actin expression.
Transient transfection and luciferase assays
To evaluate whether PPAR expressed by ATC cells was functional, cells were transiently transfected with 0.3 µg of a firefly luciferase vector driven by the PPAR-sensitive PPRE3 sequences (PPRE3-tk-luc) (23). Briefly, 2.5 x 104 cells were seeded in 12-well plates and grown for 24 h in complete medium. Transfection was carried out by adding a mixture containing 1 µg DNA and 4 µl Fugene6 in 40 µl of serum-free medium to each well. To normalize values for transfection efficiency, cells were cotransfected with 0.5 µg of a vector coding for the Renilla luciferase. Immediately after transfection, cells were treated with rosiglitazone (10 µM) and recovered 48 h later. Cell extracts and luciferase reagents were prepared with the dual-luciferase reporter assay system (Promega). Both firefly and renilla luciferase activities were then evaluated by a luminometer (Turner 20/20; Turner Industries, Sunnyvale, CA). Normalized reporter activity was calculated as the ratio of firefly luciferase values divided by the renilla luciferase values. Relative fold activation was calculated by dividing normalized values of treated cells by normalized values of control cells.
Gene silencing by small interfering RNA (siRNA)
Cells were plated onto 6-well plates (105/well), maintained in antibiotic-free medium for 24 h, and transfected with a mixture containing Opti-MEM, 8 µl/well LipofectAMINE 2000 (Invitrogen, San Diego, CA), and either 0.5 µg/well scramble siRNA or a mixture of four PPAR siRNA (smart pool; Dharmacon Research, Inc., Lafayette, CO) for 5 h. The sequence of these siRNAs is available from the manufacturer. Cells were then incubated with fresh medium for 48 h until processing.
Real-time PCR
Oligo 6.3 primer analysis software (Molecular Biology Insights, Inc., West Cascade, CO) was used to design appropriate following primer pairs: 5'-TCA GGA AGG CAC TGC TTA TGG-3' (forward) and 5'-GCC CTC TCT GGG CTG ATA ATT-3' (reverse) specific for the human thyroglobulin (Tg); 5'-ACC TGC TCC TCA TCG CCT CT-3' (forward) and 5'-CAA AGG AAG CAG GGC GAC AA-3' (reverse) specific for the human TSH-R; 5'-GGC GTC GCT CCT GTC CAC-3' (forward) and 5'-CGC CCA CAA GCA TGA CAC-3' (reverse) specific for the human NIS; 5'-CTG CGG GAC GGT GAC T-3' (forward) and 5'-CGC GCC TCC CAG ACT-3' (reverse) specific for the human thyroperoxidase (TPO); primer pairs were synthesized by MWG-Biotech (Ebersberg, Germany). Total RNA (5 µg) was reverse transcribed by ThermoScript RT (Invitrogen) and Oligo dT primers. Synthesized cDNA (0.15 µl) was then combined in a PCR using 0.2 µM of specific primers.
ELE-1 (housekeeping gene) amplification was performed using the following primers: 5'-ATT-GAA-GAA-ATT-GCA-GGC-TC-3' (forward) and 5'-TGG-AGA-AGA-GGA-GCT-GTA-TCT-3' (reverse) (fragment size, 280 bp). Quantitative real-time PCR was performed on an ABI Prism 7700 (PE Applied Biosystems, Foster City, CA) using Sybr Green PCR master mix (PE Applied Biosystems) following the manufacturer’s instructions. Amplification reactions were checked for the presence of nonspecific products by agarose gel electrophoresis. Relative quantitative determination (PE Applied Biosystems user bulletin no. 2) of target gene levels was performed by 
Ct values, as previously described (24).
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Results |
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protein protein expression was measured by Western blot in six ATC cell lines and two papillary cancer cell lines. PPAR protein was identified at variable levels at the expected molecular mass of 54 kDa in all cell lines examined except NPA papillary thyroid cancer cells (Fig. 1A
). The highest PPAR levels were observed in FF-1 and ARO ATC cells. To evaluate whether PPAR expressed by ATC cells was functional, each ATC cell line was transfected with a PPRE3-tk-luciferase reporter and then exposed to the PPAR agonist rosiglitazone (10 µM) for 24 h. Rosiglitazone treatment determined a significant luciferase increase (ranging from 150 to 450%) in all cell lines except NPA, demonstrating that PPAR protein was functional in all ATC cell lines (Fig. 1B).
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agonists reduce cell viability of ATC cells agonists, ciglitazone and rosiglitazone, on cell viability was assessed by MTT assay. Dose-response experiments with both compounds were carried out in FF-1 and ARO cell lines. Cells were cultured in monolayers in serum containing medium and MTT staining was evaluated after a 5-d exposure to increasing doses of each PPAR agonist. As shown in Fig. 2A, both PPAR agonists markedly reduced cell viability, with rosiglitazone being more potent than ciglitazone. Both compounds were more effective in serum-free conditions than serum-containing medium (not shown). In FF-1 cells, viability was reduced to 50% at 16 µM rosiglitazone in serum-free medium and at 34 µM in the presence of serum. Corresponding values for ciglitazone were 44 and 64 µM (not shown). Dose-response experiments were then carried out with rosiglitazone in the remaining ATC cell lines (Fig. 2A). In all cell lines cell viability was effectively reduced by incubation with rosiglitazone, starting at 0.1–1.0 µM. In contrast, cell viability of PPAR-negative NPA cells was unaffected by rosiglitazone at doses up to 100 µM (not shown). The effect of rosiglitazone was markedly inhibited when PPAR was knocked down by transfection with specific siRNA cells (Fig. 2B).
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agonists was then evaluated in all ATC cell lines using 10 µM rosiglitazone and 20 µM ciglitazone. Both PPAR agonists reduced ATC cell viability to 44–72% of untreated cells in serum-free conditions (Table 1). Also in these conditions, PPAR agonists were ineffective on NPA papillary cell line. To further evaluate whether this reduction of cell viability was due to PPAR agonist binding to PPAR, we used GW9662, a selective and high-affinity PPAR antagonist, which covalently modifies a cysteine residue in the ligand binding site of PPAR. Preincubation (6 h) with 1 µM GW9662 fully inhibited the effect of 10 µM rosiglitazone or 20 µg ciglitazone on cell viability (Table 1). Taken together, these data indicate that both rosiglitazone and ciglitazone affect cell viability through a PPAR-dependent mechanism.
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agonist effects in a variety of tumor cells, we evaluated whether 9-cis-retinoic acid, at a dose of 1 µM, affected ATC cell viability. Under the conditions used, cis-retinoic acid slightly reduced cell viability only in ARO and FF-1 cells, whereas it was ineffective in the remaining four cell lines (Table 1
). 9-cis-retinoic acid did not increase the effect of PPAR agonists when used in combination (data not shown).
We next investigated whether rosiglitazone was able to potentiate the antitumor effect of doxorubicin, the chemotherapeutic agent most frequently used in ATC treatment. Cells were cultured in serum containing medium and incubated with graded doses of doxorubicin for 5 d. Under these conditions, dose-response experiments showed that a dose of doxorubicin of 0.0250 µM was able to reduce cell survival by 15–60% in all cell lines except HTh74 cells, in which the same effect was achieved with 0.0125 µM doxorubicin (data not shown). A dose of doxorubicin of 0.0125 µM. was therefore used for HTh74 cells and a dose of 0.0250 µM for all the other cell lines in combination experiments with rosiglitazone. The simultaneous exposure to 10 µM rosiglitazone markedly increased the effect of doxorubicin in all ATC cells but not in PPAR-negative NPA cells (Table 2).
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agonists reduce cell proliferation and cell cycle progression in ATC cells agonists. In a variety of cancer cells, PPAR inhibits proliferation and cell cycle arrest at G0/G1 restriction point. We then evaluated in ATC cells whether rosiglitazone induced these alterations in cell cycle progression. Cells were cultured in the absence or presence of rosiglitazone and synchronized by culturing them for 24 h in leucine-deprived medium, which induces a block at the G1 restriction point. Cells were then released from the cell cycle block by the addition of regular medium and harvested at 12 h from release. Cell cycle was analyzed by fluorescence-activated cell sorter. Exposure to rosiglitazone differently increased the proportion of cells in G0/G1 in the cell lines studied (Table 3). Five cell lines also showed a reduction of the proportion of cells in the S phase. One cell line (C643), however, after exposure to rosiglitazone showed an increased proportion of cells in both G0/G1 and S phase with a reduction of the proportion of cells in G2/M phase (Table 3).
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To evaluate whether overall DNA synthesis was slowed, all six ATC cell lines were exposed to either 10 µM rosiglitazone or 20 µM ciglitazone for 48 h and then subjected to a 4 h 3H-thymidine pulse. 3H-thymidine incorporation was significantly reduced by rosiglitazone in five of six cell lines and by ciglitazone in four of six, confirming that the rate of DNA synthesis is slowed by PPAR agonists in most ATC cells (Table 4). By contrast, cis-retinoic acid reduced 3H-thymidine incorporation in only two cell lines and increased it in one (Table 4).
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). The p21cip1 increase ranged from a minimum of approximately 2-fold in FF-1 and 8305c cells to a maximum of approximately 6-fold in C643 cells. Rosiglitazone also induced up-regulation of p27kip1 expression in four of six cell lines (Fig. 3A); the increase was maximal in FF-1 and ARO cells (
2.8-fold in both and 1.8-fold in C643 and SW1736 cells).
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We then evaluated in ATC cells the phosphorylation status of Rb by using an antibody recognizing the hypophosphorylated (inactive) form of the Rb protein. Exposure to rosiglitazone for 48 h increased the hypophosphorylated form of Rb protein by approximately 2-fold in all cells, except HTh74, in which the hypophosphorylated Rb, expressed at a very low level, was increased by approximately 10-fold after rosiglitazone (Fig. 3A). These data suggest therefore that rosiglitazone inhibits CDK-dependent Rb phosphorylation and subsequent cell cycle progression by increasing p21cip1 and/or p27kip1 and decreasing cyclin D1 expression levels.
To investigate the mechanisms underlying the potentiation of doxorubicin effect by rosiglitazone, we evaluated the combined effect of doxorubicin and rosiglitazone on cell cycle molecules. Cells were incubated with doxorubicin at the indicated doses in the presence or absence of 10 µM rosiglitazone for 48 h. Doxorubicin was able to increase the expression of p21cip1 and p27kip1and also rosiglitazone. The greatest increase of both molecules was observed when rosiglitazone and doxorubicin were used in combination (Fig. 3B). Doxorubicin, however, also increased cyclin D1 expression. This increase was partially inhibited by the concomitant exposure to rosiglitazone (Fig. 3B).
PPAR agonists induce an increased rate of cell apoptosis in ATC cells
We next evaluated whether an increased apoptosis rate may contribute to the reduction of cell viability induced by exposure to PPAR agonists. Cells were exposed to either rosiglitazone (0.1–10 µM) or ciglitazone (20 µM) for 48 h, and apoptosis was then evaluated by the measure of cytoplasmic histone-associated DNA fragments. PPAR agonists markedly increased the apoptosis rate in FF-1, HTh74, and C643 cells, whereas they were less effective in SW1736, ARO, and 8305c cells. Rosiglitazone was always more effective than ciglitazone, and its effect was already evident at low doses (0.1–1.0 µM) (Fig. 4). Rosiglitazone effect on ATC cell apoptosis was confirmed by annexin and PI staining in three cell lines (FF-1, ARO, 8305c), in which the effect of rosiglitazone was compared with the effect of staurosporin, a well-known proapoptotic agent (Table 5).
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agonists, we studied the effect of rosiglitazone on some apoptosis regulators, such as Bcl-XL, B-cell leukemia-2 (Bcl-2), Bcl-2-associated X protein (Bax), and second mitochondrion-derived activator (Smac) (also known as Diablo).
The Bcl-2 family includes important apoptosis regulators that can have either proapoptotic (Bax) or antiapoptotic effects (Bcl-2, Bcl-XL). Exposure to rosiglitazone (10 µM) caused a decrease of Bcl-XL expression levels in four cell lines, more marked in C643 and 8305c cells (17 and 15% of untreated, respectively) and less marked in FF-1 and HTh74 (60 and 40%, respectively) (Fig. 5A). Rosiglitazone did not affect Bcl-2 and Bax expression levels (Fig. 5A).
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). To evaluate the role of caspase activation in rosiglitazone-induced apoptosis, ATC cells were exposed to 10 µM rosiglitazone in the presence or absence of 30 µM of the caspase inhibitor Z-VAD-fmk. Exposure to staurosporin (50 nM), a general tyrosine kinase inhibitor that induces apoptosis and activates caspases (27), was used as positive control. Cell apoptosis was measured by determining cytoplasmic histone-associated DNA fragments, as described in Materials and Methods. In all cell lines, both rosiglitazone- and staurosporin-induced apoptosis was almost completely blocked in the presence of Z-VAD-fmk (data not shown).
Activation of caspase-3/7 was then measured in rosiglitazone-exposed cells. As shown in Fig. 5B, rosiglitazone significantly (P < 0.001) induced caspase-3/7 activation in all cell lines at a level comparable with that induced by 50 nM staurosporin. Caspase-3/7 activation by both rosiglitazone and staurosporin was completely inhibited by 30 µM of Z-VAD-fmk (not shown).
PPAR agonists inhibit anchorage-independent growth
Anchorage-independent growth is an important feature of the malignant phenotype. We therefore evaluated the effect of PPAR agonists on anchorage-independent growth by cloning cells in soft agar. Only two cell lines, ARO and SW1736, were able to efficiently form colonies in semisolid agar. At d 5, colonies developed in both the untreated and PPAR agonist-treated cultures. Colonies were then stained and photographed. Both ciglitazone (20 µM, not shown) and rosiglitazone (10 µM) markedly reduced colony formation in these two cell lines. Figure 6A shows the effect of 10 µM rosiglitazone in ARO cells. Dose-response experiments showed that this inhibiting effect was already evident at doses as low as 0.1 µM PPAR agonists (not shown).
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agonists antagonize biological effects of IGF-I in ATC cells by inhibiting Akt phosphorylation and up-regulating PTEN agonists affected IGF-I effects in ATC cells. When cell monolayers were stimulated with 10 nM IGF-I, cell density increased from 131 to 211%, as evaluated by MTT assay; 10 µM rosiglitazone completely blocked this effect (data not shown).
When ARO cells were seeded in semisolid agar in the presence of a reduced amount of serum (2% charcoal-stripped serum), IGF-I was able to stimulate both the number and size of colonies. This IGF-I effect was also completely blocked by the presence of rosiglitazone (10 µM) (Fig. 6B). Most biological effects of IGF-I are responsive to the activation of the insulin receptor substrate (IRS)-1/PI3K/Akt pathway. We therefore evaluated whether LY294002, an inhibitor of PI3K, affected IGF-I-stimulated colony formation. In fact, IGF-I effect was abolished by LY294002 (5 µM) (Fig. 6B). In these culture conditions, the effect of both LY294002 and rosiglitazone was minimal in the absence of IGF-I (not shown).
We also evaluated the effect of rosiglitazone on cell migration in Boyden chambers in response to IGF-I used as chemoattractant. Both the basal and IGF-I stimulated cell migration were significantly inhibited by rosiglitazone in all ATC cell lines (Fig. 7). Incubation with LY294002 (5 µM) also blocked the effect of IGF-I on cell migration (Fig. 7). Taken together these data indicate that IGF-I-induced biological effects such as colony formation and cell migration are PI3K dependent and are inhibited by rosiglitazone.
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). IRS-1 was clearly up-regulated in three cell lines (FF-1, HTh74, and SW1736), whereas IRS-2 content was unaffected (Fig. 8A).
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80% of untreated in both) and markedly decreased in HTh74, C643, and 8305c cells (50% of untreated in all three of them) (Fig. 8B). Very similar results were obtained with an antibody to phosphorylated Thr308 of Akt (not shown). In contrast to Akt, IRS-1 phosphorylation reflected IRS-1 expression levels and was increased by rosiglitazone in FF-1, HTh74, and SW1736, indicating that IRS-1 was functional and not responsible for the inhibition of Akt phosphorylation by rosiglitazone (Fig. 8B). The ratio phospho-IRS-1/phospho-Akt was therefore increased in rosiglitazone-treated cells, compared with untreated cells (Fig. 8B).
Time-course experiments revealed that Akt phosphorylation (both Ser473 and Thr308) was reduced, especially as far as peak values are concerned. In untreated FF-1 cells, Akt phosphorylation at Ser473 remained clearly detectable at 180 min, whereas it was almost switched off after 80 min in rosiglitazone-treated cells (Fig. 8C). Similar results were obtained in HTh74 and C643 cells (data not shown).
The Akt pathway is negatively regulated by the lipid phosphatase PTEN, which is an important antioncogene. In cells exposed to rosiglitazone, PTEN was clearly up-regulated in all cell lines, ranging from 148 ± 12 to 414 ± 27% of untreated cells (Fig. 9). These data indicate that PTEN up-regulation and subsequent inhibition of Akt phosphorylation could be involved in the inhibition by rosiglitazone of the ATC cell biological responses to IGF-I.
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agonists agonists have been reported to promote differentiation in certain cancer cells, we evaluated whether rosiglitazone increased the expression of thyroid-specific differentiation markers, such as Tg, TSH-R, NIS and TPO, in ATC cells. After incubation with 10 µM rosiglitazone for 48 h, mRNA for Tg, TSH-R, NIS, and TPO was measured by quantitative real-time PCR. As shown in Fig. 10, exposure to rosiglitazone increased the expression of these thyroid-specific molecules in ATC cell lines. No significant effect was observed in NPA papillary cancer cells, whereas PPAR was undetectable and nonfunctional.
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protein and that, in these cells, PPAR agonists induce pleiotropic effects consistent with a partial reversion of the EMT, highly characteristic of this aggressive carcinoma.
Biological effects elicited by PPAR agonists in ATC cells included: 1) inhibition of anchorage-dependent and -independent growth and inhibition of cell cycle progression at G0/G1 phase; 2) potentiation of doxorubicin effects; 3) inhibition of cell migration; 4) increased apoptosis rate; 5) down-regulation of the IGF system; and 6) increase of expression of thyroid-specific differentiation markers. These effects will be discussed separately.
PPAR agonists reduced ATC cell viability by inducing both growth arrest and an increased apoptosis rate. They also dramatically reduced anchorage-independent growth by reducing both colony number and size. In addition, our study indicates that PPAR agonists inhibit cell cycle progression in ATC cells by multiple mechanisms, resulting in ATC cell accumulation in G0/G1 phase. The progression in the cell cycle requires cyclin/CDK complexes that result in phosphorylation of the Rb protein. Upon phosphorylation, Rb releases the transcription factor E2F that activates transcription of genes involved in cell cycle progression (26, 27). Rosiglitazone increased the hypophosphorylated (inactive) form of Rb protein, a likely consequence of the effects of up-regulation of CDK inhibitors and down-regulation of cyclin D1. Cyclin/CDK complexes are negatively regulated by CDK inhibitors, in particular p21Cip1 and p27Kip1, that have a key role in mediating G1 arrest by inhibiting cyclin D1-CDK4 and other cyclin/CDK complexes (26, 27, 28, 29). Rosiglitazone up-regulated both p21Cip1/Waf1 and p27Kip1. p21Cip1/Waf1 is normally induced by p53 and is a key mediator of p53-mediated G1 arrest (30). However, p21Cip1/Waf1 may also be induced by p53-independent mechanisms. The ATC cell lines examined in the present study express either a mutated p53 (31) or do not express p53 (SW1736 cells); therefore, the rosiglitazone-mediated p21Cip1/Waf1 up-regulation found in ATC cells appears to be p53 independent. This observation is relevant in view to the fact that p53 is frequently mutated in human ATCs and that p53 mutations are believed to play a role in thyroid cancer dedifferentiation (34).
TZDs had already been reported to influence CDK inhibitors: troglitazone may induce up-regulation of both p21Cip1/Waf1 and p27Kip1 in thyroid cancer cells and pancreatic cells (14, 33), whereas ciglitazone may up-regulate p27Kip1, but not p21Cip1/Waf1, in thyroid cancer cells (34) and p27Kip1 and p18INK4c in hepatocellular carcinoma (35, 36). However, this p53-independent effect of TZDs on CDK inhibitors is a novel observation to our knowledge.
We also demonstrate that rosiglitazone down-regulated cyclin D1 protein in ATC cells. Cyclin D1 is a rate-limiting factor in cell proliferation induced by growth factors in various models and is often overexpressed in cancer cells in which it is induced by several oncogenes and functions as a collaborative oncogene (37, 38, 39). Previous studies carried out in other cell models have reported that PPAR agonists may down-regulate cyclin D1 expression by both inducing proteasome-dependent degradation of the protein (40) and inhibiting its transcription (41).
In addition to cell growth inhibition, PPAR agonists increased ATC cells apoptosis rate. These findings are in agreement with the results of other authors, who showed a predominant apoptotic effect of PPAR agonists in different anaplastic thyroid cancer cells (42). We found that rosiglitazone-induced apoptosis was associated in most ATC cells with the down-regulation of BclxL an antiapoptotic protein of the Bcl-2 family. Bcl2 and Bax were unaffected. Rosiglitazone also caused caspase-3/7 activation, and incubation with the caspase inhibitor Z-VAD-fmk reduced rosiglitazone-induced apoptosis. Also, these are novel findings, although it has been previously reported that Bcl2 may be down-regulated by PPAR agonists in breast and colon cancer and glioblastoma (43, 44, 45).
Interestingly, rosiglitazone potentiated the effect of low doses of doxorubicin, a chemotherapeutic drug frequently used in the therapy of ATC, on cell growth and survival. Interestingly, both rosiglitazone and doxorubicin up-regulated p21Cip1/Waf1 and p27Kip1, and the combination of the two compounds had an additive effect. These data suggest that this drug combination may improve the outcome of ATC patients. These data are in agreement with a very recent report showing that a novel PPAR agonist (RS5444) alone and in combination with paclitaxel inhibits in vitro and in vivo growth of two different ATC cell lines by up-regulating p21Cip1/Waf1 (46).
The ability of cancer cells to migrate and invade the endothelial basal membrane is a prerequisite to develop local invasion and metastatic spreading. ATC is characterized by massive invasion of nearby structures and early development of locoregional and distant metastases. The markedly reduced ATC cell migration caused by rosiglitazone is an intriguing finding and is reminiscent of chemotaxis inhibition of vascular smooth muscle cells and monocytes by PPAR agonists (47, 48). One possible mechanism is Ets-1 transrepression (49), which is known to regulate matrix metalloproteinase-9 and other matrix metalloproteinases that allow digestion of extracellular matrix (50).
An additional antineoplastic mechanism of TZDs in ATC cells is the antagonist effect on IGF-I. Many features of malignant cells, such as unregulated growth and apoptosis and enhanced cell invasion, are modulated by growth factors and cytokines produced in an autocrine or paracrine manner. Among growth factors produced by ATC, IGF-I, and IGF-II have a crucial role in thyroid cancer progression through the activation of their cognate receptor IGF-IR and insulin receptor isoform A (6), which have a mitogenic, antiapoptotic, and transforming potential (51). Rosiglitazone antagonized major biological effects of IGF-I, such as cell migration, survival, and anchorage-independent growth. A possible mechanism for this antagonist effect is the inhibition of the PI3K/Akt pathway by PTEN up-regulation. The PI3K/Akt pathway is a major pathway activated by IGF-I and plays a key role in mediating its biological effects (52). This pathway is negatively regulated by the lipid phosphatase PTEN that dephosphorylates the PI3K product phosphatidylinositol-(3,4,5,)-triphosphate, which, in turn, activates Akt and other downstream kinases (53). The concomitant Akt inhibition and PTEN up-regulation in response to PPAR agonists have already been reported in pancreatic cancer cells (54).
Finally, rosiglitazone increased the expression of thyroid-specific differentiation markers, such as Tg, TSH-R, NIS and TPO, indicating a partial redifferentiation effect. Our results are in agreement with those of others showing that troglitazone increases iodine uptake by cultured follicular cancer cell lines in vitro (55, 56) and that rosiglitazone administration in patients with metastatic thyroid cancer may increase Tg production and radioiodine uptake, albeit at a low degree, by the metastatic tissue (57).
These multiple molecular effects of TZDs affected ATC cell phenotype, inducing morphological (more polygonal, epithelial-like cell morphology; data not shown) changes that are typical of thyroid cell epithelial differentiation and EMT reversal.
It has been reported that some antitumor effects of TZDs are PPAR independent, at least in certain cell models (58, 59). In our system, however, different lines of evidence indicate that the biological effects of TZDs described here require a functional PPAR. First, these effects occurred at doses of rosiglitazone as low as 0.1 µM and only in cells expressing a functional PPAR, whereas they were not observed in PPAR-negative NPA papillary cancer cells. Second, they were inhibited by specific siRNA to PPAR and pretreatment with GW9662, a potent and selective antagonist of PPAR that causes specific and irreversible loss of binding. However, there was no strict correlation between PPAR expression and functional level and the molecular and biological response to TZDs in the different ATC cells lines. This is not surprising because the selective transactivation of the different genes by PPAR activation may vary in the different cells and tissues according to the nature of PPRE in the upstream regions of target genes and the relative abundance of coactivators and corepressors and other nuclear transcription factors that may either compete or cross-talk with PPAR and its heterodimerization partner retinoid X receptor (60). Moreover, the relationship between gene regulation by PPAR and biological effects is obviously affected by the numerous abnormalities in growth and apoptosis regulatory genes present in the different ATC cell lines. However, in all cell lines studied, PPAR agonists had similar antiproliferative, proapoptotic, and differentiating effects.
In conclusion, our data indicate that PPAR agonists induce a partial reversion of the epithelial mesenchymal transition in ATC cells by multiple mechanisms. These effects occur independently of a functional p53, often mutated in ATC, and include partial antagonism with IGF-I effects. Furthermore, because PPAR agonists potentiate the effect of doxorubicin, the most frequently used chemotherapeutic drug in patients with ATC, they may have a clinical role in the multimodal therapy currently used to slow down ATC growth and dissemination.
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First Published Online June 15, 2006
Abbreviations: ATC, Anaplastic thyroid cancer; Bax, Bcl-2-associated X protein; Bcl2, B-cell leukemia-2; CDK, cyclin-dependent kinase; EMT, epithelial mesenchymal transition; FCS, fetal calf serum; IGF-IR, IGF-I receptor; IRS, insulin receptor substrate; MTT, dimethylthiazoldiphenyltetrazoliumbromide; NIS, sodium-iodine symporter; PI, propidium iodide; PI3K, phosphatidylinositol 3-kinase; PPAR, peroxisomal proliferator-activated receptor; PPRE, peroxisome proliferator response element; PTEN, phosphatase and tensin homolog deleted from chromosome 10; Rb, retinoblastoma protein; siRNA, small interfering RNA; Smac, second mitochondrion-derived activator; Tg, thyroglobulin; TPO, thyroperoxidase; TSH-R, TSH receptor; TZD, thiazolidinedione; Z-VAD-fmk, N-benzyloxycarbonyl-Val-Ala-Asp-fluoromethylketone.
Received December 19, 2005.
Accepted for publication June 6, 2006.
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) or rosiglitazone (
) for 4 d in 10% FCS containing medium, and cell viability was then measured by MTT staining. Only rosiglitazone (
) in the lower chamber for 18 h. The migrated cells were stained as described in Materials and Methods. Bars represent the mean ± SD of triplicate experiments.
