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Oncological Endocrinology (M.G.C., N.F., M.P., L.C., R.P., G.B.), Azienda Sanitaria Ospedaliera, San Giovanni Battista, and Department of Clinical Pathophysiology (M.G.C., O.B., G.B.), University of Turin, 10126 Turin, Italy
Address all correspondence and requests for reprints to: Prof. Giuseppe Boccuzzi, Dipartimento di Fisiopatologia Clinica, Via Genova 3, 10126 Torino, Italy. E-mail: giuseppe.boccuzzi{at}unito.it.
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Abstract |
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Introduction |
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5% of thyroid cancers) are resistant to conventional therapy and are almost always fatal within a few months of diagnosis (4). Furthermore, chemotherapy and radiotherapy give disappointing results, and new therapeutic approaches are needed. An increasing body of evidence concerning the importance of epigenetic changes in cancer onset and progression has raised interest in the manipulation of transcription as a mode of cancer therapy; altering gene expression through chromatin modification now seems to be a viable target. Consistent with this, histone deacetylase (HDAC) inhibitors have emerged as a promising new class of anticancer drugs (5, 6, 7), and both natural and synthetic inhibitors have been characterized (8). They are structurally different but share the capacity to enhance cell differentiation (9), induce apoptosis (10, 11, 12), inhibit cancer cell growth (13), and revert oncogene-transformed cell morphology (14). However, a number of limitations hamper their clinical use; some have a short half-life and/or significant toxic side effects in vivo (15). Conversely, valproic acid (VPA), a potent anticonvulsant that acts also as a class I selective HDAC inhibitor (16), produces mild adverse effects in man, even when serum levels exceed the normal therapeutic range while receiving antiepileptic therapy (17). The drug alters the expression of a critical subset of target genes, and this selective modulation probably explains both its therapeutic efficiency and the paucity of side effects. Moreover, it also has useful pharmacokinetic properties, with a significantly longer biological half-life than the other HDAC inhibitors (16). VPA has already been proposed for redifferentiating treatment of hematological malignancies (18) and neuroblastoma (19). Recently, we showed (20) that VPA can promote the redifferentiation of poorly differentiated thyroid cancer cells, restoring their ability to capture and concentrate iodine.
In addition to its redifferentiating property, VPA has been reported to affect the growth of several transformed cells (19, 21, 22, 23, 24) and to induce apoptosis in human leukemia cell lines (25) and in endometrial cancer cells (26).
The study aimed to define the effect of VPA on the growth of poorly differentiated thyroid carcinoma cells. We demonstrate that, at concentrations that induce cell redifferentiation and restore sensitivity to radio-iodine therapy, VPA strongly inhibits cell growth through induction of apoptosis and cell cycle arrest. We suggest that additional mechanisms (apoptosis and cell cycle arrest) might cooperate with VPA’s ability to restore radio-iodine sensitivity in treating poorly differentiated thyroid carcinoma.
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Materials and Methods |
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Papillary thyroid carcinoma (N-PA) cell line was a kind gift from Prof. Mauro Papotti and Dr. Paola Cassoni (Pathology Service, Department of Oncology, University of Turin). The subline of N-PA cells we used does not express any markers of differentiation, as we report elsewhere (20), and may thus be considered a good model of dedifferentiated carcinoma. Poorly differentiated papillary carcinoma (BHT-101) cells were purchased from Deutsche Sammlung von Mikroorganismen und Zellculturen (Braunschweig, Germany). Human thyroid follicular epithelial cell line (Nthy-ori 3-1) was purchased from European Collection of Cell Cultures (Salisbury, UK). N-PA and Nthy-ori 3–1 cells were routinely maintained in 25-cm2 flasks, at 37 C in 5% CO2-95% humidity, in RPMI 1640 (Sigma, St. Louis, MO) with 100 IU/ml penicillin and 100 µg/ml streptomycin added, supplemented with 10% heat-inactivated fetal calf serum (FCS) (Euroclone, Wetherby, West York, UK). The BHT-101 cell line was maintained in DMEM plus 20% heat-inactivated FCS.
Once a week, cells were detached with trypsin/EDTA and reseeded at a dilution of 1:2–1:4.
Cell viability assay
Cells were seeded at 3 x 103 cells/well in 96-multiwell plates (Corning, New York, NY) in culture medium plus 10% FCS. After 24 h, they were treated with medium alone or with medium containing different doses of VPA (0.5, 1, 1.5, or 3 mM) (Sigma) for up to 11 d. At d 1, 4, 7, 9, and 11, viable cells were determined by a 3–4,5-dimethylthiazol-2,5 biphenyl tetrazolium bromide (MTT; Sigma) assay, as described elsewhere (27). Briefly, 10 µl 12-mM MTT was added to each well; after a further 4-h incubation, 100 µl 0.04-N NaCl in isopropanol was added, and absorbance at 495 nm was measured using a plate reader (Model 680 Microplate Reader; Bio-Rad, Hercules, CA). Eight replicates were done to determine each data point.
Cell death detection ELISA
For apoptosis studies, 3 x 105 cells were seeded in 96-multiwell microtiter plates and treated with culture medium with or without VPA (0.5, 1, 1.5, or 3 mM) for 4 d; apoptosis was then evaluated using Cell Death Detection ELISAPLUS (Roche, Basel, Switzerland), following the manufacturer’s instructions. This assay is based on a quantitative sandwich-enzyme-immunoassay principle using monoclonal antibodies directed against DNA and histones, respectively. The assay provides the specific determination of mono- and oligonucleosomes in the cytoplasm fraction of cell lysates. Apoptosis was expressed as an enrichment factor, calculated as a fraction of the absorbance of treated cells vs. untreated controls.
Poly-(ADP-ribose)-polymerase (PARP) Western blot
In 75-cm2 flasks, 1 x 106 cells were cultured, treated with 0.5–3 mM VPA, and incubated for 4 d. After VPA treatment, cell pellets were resuspended in 150 µl extraction buffer [per sample, 100 µl 50-mM glucose, 25 mM Tris-HCl (pH 8), 10 mM EDTA, 1 mM phenylmethylsulfonylfluoride were mixed, just before use, with 50 µl 50-mM Tris-HCl (pH 6.8), 6 M urea, 6% 2-mercaptoethanol, 3% sodium dodecyl sulfate, 0.003% bromphenol-blue]. Cell extracts were sonicated and incubated for 15 min at 65 C. Extracts (20 µl/well) were subjected to SDS-PAGE (T = 8%) and electroblotted on a PVDF membrane. The membrane was probed with a rabbit polyclonal antibody anti-PARP (Roche) as primary antibody. The peroxidase-linked antirabbit antibody (Amersham Biosciences, Little Chalfont, UK) was used as secondary antibody. Chemiluminescence was detected, following the enhanced chemiluminescence manufacturer’s instructions. Bands were photographed and analyzed with Kodak 1D Image Analysis software (Eastman Kodak Co., Rochester, NY).
The membrane was also stained with Ponceau S Solution (Sigma) to check protein loading.
Caspase activity assay
In 75-cm2 flasks, 1 x 106 cells were seeded and exposed to 3 mM VPA for 48 h. After VPA treatment, caspases-3, caspase-8, and caspase-9 activities were determined using colorimetric assay kits (R&D Systems, Inc., Minneapolis, MN), following the manufacturer’s instructions. Briefly, cells were lysed and incubated with colorimetric substrates (DEVD-pNA for caspese-3, IETD-pNA for caspase-8, and LEHD-pNA for caspase-9) for 2 h at 37 C. After incubation, the chromophores were quantified spectrophotometrically at a wavelength of 405 nm.
Cell cycle analysis
Cells were treated with 0.5–3.0 mM VPA for up to 96 h. At 24 h, 48 h, 72 h, and 96 h, respectively, all cells were collected, fixed in 70% ethanol for 30 min on ice, and incubated in propidium iodide solution (20 µg/ml propidium iodide, 0.2 mg/ml ribonuclease A in PBS) for 1 h at room temperature. Analysis of the whole cell population, as well as that of the viable fraction alone, was performed using an EPICS XL flow cytometer (Coulter Corp., Hialeah, FL).
RNA extraction and RT-PCR analysis
Cells were seeded at 2 x 105 cells/well in six-well plates; after 48 h, they were treated with 0.5–3.0 mM VPA for 48 h. Total RNA was extracted from both cell lines using TRIzol Reagent (Invitrogen, Groningen, The Netherlands), following the method developed by Chomczynski and Sacchi (28). Total RNA was reverse-transcribed at 42 C for 40 min using AMV reverse transcriptase (Finnzymes, Espoo, Finland) and oligodT primer (Invitrogen). The PCR system contained 5 µl of 10x PCR buffer, 10 µl of RT product, 0.2 mM deoxynucleotide triphosphate (Finnzymes), 1.25 U Taq DNA polymerase (Finnzymes), 50 ng each of sense and antisense primers in a total vol of 50 µl; primers: 5'-TTG TGA AGG CAG GGG GAA G and 3'-GGA AGG TCG CTG GAC GAT TTG A for p21, 5'-TCA AAC GTG CGA GTG TCT AAC G and 3'-TTG GGG AAC CGT CTG AAA CAT TTT for p27, 5'-ACC CCT TAA GGA TCT TCC TG and 3'-TCC AGG GTA TAT CCA GTC TTT CG for cyclin A, 5'-GGA TGC TGG AGG TCT GCG AGG AAC and 3'-GAG AGG AAG CGT GTG AGG CGG TAG for cyclin D1, 5'-GGA GAA CTT CCA AAA GGT GG and 3'-CTG GCT TGG TCA CAT CCT GG for cdk2, 5'-TCT CGA TAT GAG CCA GTG GCT G and 3'-TCC ACG GGG CAG GGA TAC ATC for cdk4, 5'-CTC ACC CTG AAG TAC CCC ATC G and 3'-CTT GCT GAT CCA CAT CTG CTG G for ß-actin. The expected PCR products were 495 bp for p21, 535 for p27, 936 for cyclin A, 514 for cyclin D1, 875 for cdk2, 732 for cdk4, and 885 for ß-actin. Amplification was carried out as follows: for p21, 1x (95 C, 3 min), 35x (95 C, 1 min; 60 C, 1 min; 72 C, 1 min), and 1x (72 C, 7 min). For p27, cyclin A, and cdk2, 1 x (95 C, 3 min), 40 x (95 C, 1 min; 55 C, 1 min; 60 C, 1 min), and 1 x (72 C, 7 min). For cyclin D1 and cdk4, 1 x (94 C, 2 min), 40 x (94 C, 1 min; 65 C, 1 min; 72 C), and 1 x (72 C, 7 min). For ß-actin, 1 x (94 C, 1 min), 35 x (94 C, 30 sec; 58 C, 30 sec; 72 C, 30 sec), and 1 x (72 C, 7 min). PCR products were electrophoresed on 1.5% agarose gel in the presence of ethidium bromide. Gels were photographed and analyzed with Kodak 1D Image Analysis software. The net intensity of bands in each experiment was normalized to the intensity of the corresponding ß-actin band before comparison between VPA-treated cells and untreated control.
Western blot analysis
In 75-cm2 flasks 1 x 106 cells were seeded and treated with 0.5–3 mM VPA for 3 d. After treatment, cells were lysed in RIPA buffer (PBS, pH 7.4; 1% Nonidet P-40; 0.1% sodium dodecyl sulfate; 0.5% sodium deoxycholate; 100 µg/ml phenylmethylsulfonylfluoride; 30 µl aprotinin; 100 mM NaVO4), extracted at 4 C for 30 min, and centrifuged at 4 C for 20 min at 15,000 x g. Equal amounts of protein (50 µg protein/lane) were subjected to SDS-PAGE (T = 8%) and electroblotted onto a PVDF membrane; the membrane was probed with the following primary antibodies: antihuman p21 (sc-397, 1:200 dilution; Santa Cruz Biotechnology Inc., Santa Cruz, CA); antihuman cyclin A (Clone BF 683, 1:1000 dilution; Upstate, Lake Placid, NY); antiactin (Monoclonal Anti-actin clone AC-40, 1:1000 dilution; Sigma) to check protein loading.
Proteins were detected with enhanced chemiluminescence Western blot reagents, following manufacturer’s instructions. Bands were photographed using Kodak 1D Image Analysis software.
Statistical analysis
Data are expressed throughout as means ± SEM, calculated from at least three different experiments. Statistical comparisons between groups were performed with ANOVA (one-way ANOVA), and the threshold of significance was calculated with the Bonferroni test. Caspase activities were compared with the paired t test. Statistical significance was set at P < 0.05.
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Results |
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Cells were treated with different doses of VPA for up to a maximum of 11 d. As shown in Fig. 1
, VPA significantly reduced the number of viable N-PA and BHT-101 cells (viability ratio < 1.2). The effect appeared on both cell lines after 4 d of treatment at VPA concentrations of 1 mM or more. On the contrary, at the concentrations effective on poorly differentiated cancer cell lines, VPA had no effect on the number of viable Nthy-ori 3-1 cells, the cell viability ratio being more than 3 even after 11 d of treatment (data not shown).
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As shown in Fig. 2, VPA induced apoptosis in both N-PA and BHT-101 cells; using the cell death detection ELISA, the enrichment factor was increased significantly in both cell lines by treatment with VPA at concentrations between 1 and 3 mM.
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, PARP was cleaved in both N-PA and BHT-101 cells treated with 1.5 and with 3.0 mM VPA. The sensitivity of the ELISA kit is higher than that of Western blot used to detect PARP cleavage, which explains why the VPA’s ability to induce apoptosis at a concentration of 1 mM became evident only with the ELISA technique.
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, caspase 3 is significantly activated in both cell lines treated with VPA (paired t test, P < 0.05 in both cell lines). Caspase-dependent apoptosis can be initiated either through the extrinsic (caspase 8) or through the intrinsic (caspase 9) pathway. These initiator caspases subsequently activate the effector caspase 3. Figure 4C shows the activation of caspase 9 in both N-PA and BHT-101 cells (P < 0.05 in both cell lines); on the contrary, no statistically significant increase in caspase 8 activity was found (Fig. 4B).
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Effect of VPA on the cell cycle
Because HDAC inhibitors affect both cell proliferation and cell survival, we assessed the effect of VPA on cell cycle progression. As shown in Table 1, flow cytometry revealed that VPA increased the sub-G1 population of treated N-PA cells vs. controls, in accordance with apoptosis induction, as demonstrated by nucleosome formation and PARP cleavage. The same effect was observed in BHT-101 cells (data not shown). The increase started after 48-h VPA treatment and was dose- and time-dependent up to 96 h. In addition to apoptosis induction, when we analyzed only viable cells, VPA at concentrations between 1 and 3 mM induced a significant growth arrest in G1. The effect appeared after 48 h of treatment, as shown in Fig. 5, in both N-PA and BHT-101 cells. A total of 61.4% of untreated N-PA cells were in G0–G1 compared with 72.2% of cells cultured with 1 mM VPA, 77% of those with 1.5 mM VPA and 78.6% of those with 3 mM VPA. As far as BHT-101 cells are concerned, 60% of untreated cells were in G0–G1, 76% of 1 mM VPA-treated cells, 80% of 1.5 mM, and 84% of 3 mM cells. The effect of VPA on G1 arrest was similar at 72 and 96 h of treatment (data not shown).
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To further clarify the mechanism of VPA-induced G1 cell cycle arrest, we examined the effect of the drug on the expression of different genes involved in cell cycle control; p21 was up-regulated both at the mRNA (Fig. 6) and at the protein level (Fig. 7), and cyclin A was down-regulated both at the mRNA (Fig. 6) and at the protein level (Fig. 7). Conversely, mRNA expression of p27, cyclin D1, cdk2, and cdk4 were unchanged upon treatment of either cell line with VPA; nor did immunoblotting of p27 and cyclin D1 show any difference in protein levels (data not shown).
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Discussion |
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TopAbstractIntroductionMaterials and MethodsResultsDiscussionReferences |
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The mechanisms underlying VPA’s effect on cell growth include both apoptosis induction and cell cycle arrest; after VPA treatment at 1 mM or above, the percentage of cells in sub-G1 increased, and cytoplasmatic nucleosome formation and PARP cleavage, well-known apoptosis markers, occurred in both cell lines. Moreover, selective G1 arrest was induced in cultured cells, 1 mM VPA again being the lowest effective dose, suggesting that cell cycle arrest also collaborates in inhibiting cell growth induced by VPA.
Apoptosis, physiologically programmed cell-death, proceeds through two different pathways: the death receptor, or extrinsic, pathway; and the intrinsic pathway. HDAC inhibitors have been reported to activate the extrinsic pathway: apidicin induces apoptosis through FAS/FAS ligand expression in leukemia cells (29); desipeptide-induced apoptosis selectively involves the TNF receptor, initiating caspase 8 and effector caspase 3 (30). Conversely, we show here that the intrinsic pathway is involved in VPA-induced apoptosis; we observed no activation of caspase-8, whereas caspase-9, which finally induces caspase-3, was clearly activated. These data are in agreement with the observation that, in leukemia cells, overexpression of Bcl-2, which blocks the intrinsic pathway, inhibits apoptosis mediated by superoylanilide hydroxamic acid (SAHA) (31), another HDAC inhibitor. Moreover, SAHA induces cleavage and activation of Bcl-2 interacting domain, resulting in mitochondrial membrane damage and reactive oxygen species production (32), thus activating the intrinsic pathway. It is likely that the specific pathway used by each HDAC inhibitor to induce apoptosis mainly depends on specific HDAC enzymes inhibited and on the cell type.
The G1 arrest induced by VPA is in line with the reported effect of this drug in endometrial carcinoma cell lines (26), and stresses its ability to favor the local remodeling of chromatin. This effect is shared by other HDAC inhibitors, such as sodium butyrate, trichostatin A, and MS-275, in various cancer cell lines (33, 34, 35). Butyrate, trichostatin A, depsipeptide, oxamflatin, MS-275, and SAHA induce expression of the CDKN1A gene, which encodes the cyclin-dependent kinase inhibitor p21 that, in turn, inhibits cell cycle progression and causes cell cycle arrest in G1 (13, 36, 37). Our data show that the induction of p21 plays a role in VPA-induced cell cycle arrest in poorly differentiated thyroid cancer. However, even if the role of p21 in G1 arrest is documented in many cancer cells, it is clear from studies with p21–/– cells that it is not the sole determinant responsible for this event (38, 39); the cell cycle arrest caused by HDAC may also be mediated by an altered expression of cyclins A and D and of p27KIP1, resulting in decreased activity of cdk4 and cdk2 (40). Here, we show that mRNA expression and protein level of cyclin A are clearly reduced in VPA-treated cells, whereas expressions of p27, cyclin D1, cdk2, and cdk 4 were unchanged, in line with the opinion that many regulator genes cooperate in controlling cell cycle progression.
It is generally believed that cells arrested in G1 evolve toward differentiation, a fate mutually exclusive with apoptosis (40, 41). However, what factors determine whether a cancer cell undergoes cell cycle arrest, differentiation, or death remains unresolved. Drug levels and cell type have been considered to be the main factors favoring toxicity or G1 arrest; HDAC inhibitors are usually toxic at higher doses and induce G1 arrest at lower doses. In our model, on the contrary, we show that the HDAC inhibitor VPA arrests the cell cycle, induces apoptosis, and promotes differentiation, all at the same dose (
1 mM VPA), making a dose-dependent effect unlikely. The differentiation effect we have reported elsewhere in N-PA cells (20) also occurred in BHT-101 cells (data not shown), where VPA induces NIS expression and increases iodine uptake to the same extent as it does in N-PA cells. As far as VPA is concerned, it might cause growth arrest in G1 and thus differentiation of some cells, which finally reexpress NIS protein and reacquire the ability to concentrate iodine; but, at the same time, the cells that escape differentiation may progress through the cell cycle and proceed toward apoptosis.
In conclusion, the identification of VPA as an inducer of both cell differentiation (20) and apoptosis argues that this drug might play a dual role in thyroid cancer management; VPA increases the efficacy of radiometabolic therapy, promoting iodine reuptake by tumor cells, and affects tumor growth by acting on the cell cycle and on cell death.
VPA levels reached in patients treated for epilepsy are usually not above 100 µg/ml (0.7 mM). Only limited toxicity occurs when the concentration is below 3.1 mM, and severe side effects develop when the concentration is above 5.9 mM; 1 mM VPA, which is the expected plasma level for use in treating poorly differentiated thyroid cancer, is just above therapeutic levels for epilepsy and thus appears clinically achievable. Translational and clinical studies will ultimately determine the clinical utility and safety of VPA as an option for the treatment of poorly differentiated thyroid cancers, which are known not to respond to conventional therapy.
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Footnotes |
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First Published Online December 7, 2004
Abbreviations: FCS, Fetal calf serum; HDAC, histone deacetylase; MTT, 3–4,5-dimethylthiazol-2,5 biphenyl tetrazolium bromide; NIS, Na+/I– symporter; PARP, poly-(ADP-ribose)-polymerase; SAHA, superoylanilide hydroxamic acid; VPA, valproic acid.
Received July 13, 2004.
Accepted November 29, 2004.
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. Int J Oncol 20:97–106[Medline]
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