Tetradecylthioacetic acid inhibits growth of rat glioma cells ex vivo and in vivo via PPAR-dependent and PPAR-independent pathways
Kjetil Berge,3,4,
Karl J. Tronstad,3,
Esben N. Flindt1,3,
Thomas H. Rasmussen2,
Lise Madsen,
Karsten Kristiansen1 and
Rolf K. Berge
Department of Clinical Biochemistry, Haukeland Hospital, University of Bergen, N-5021 Bergen, Norway,
1 Department of Biochemistry and Molecular Biology, University of Southern Denmark, DK-5230 Odense M and
2 Department of Environmental Medicine, Institute of Community Health, University of Southern Denmark, DK-5230 Odense M, Denmark
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Abstract
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The peroxisome proliferator-activated receptors (PPARs) are transcription factors involved in fatty acid metabolism and energy homeostasis. The PPARs also play crucial roles in the control of cellular growth and differentiation. Especially, the recently emerged concept of ligand-dependent PPAR
-mediated inhibition of cancer cell proliferation through induction of G1-phase arrest and differentiation is of clinical interest to cancer therapy. Tetradecylthioacetic acid (TTA) is a sulphur-substituted saturated fatty acid analog with unique biochemical properties. In this study, we investigated the effects of TTA-administration on cell proliferation in glioma cancer models. The rat glioma cell line BT4Cn, whether grown in culture or implanted in rats, expressed significant levels of PPAR
and PPAR
, with PPAR
being the predominant PPAR subtype. In BT4Cn cells, TTA activated all PPAR subtypes in a dose-dependent manner. In cell culture experiments, the PPAR
-selective ligand BRL49653 moderately inhibited growth of BT4Cn cells, whereas administration of TTA resulted in a marked growth inhibition. Administration of the PPAR
-selective antagonist GW9662 abolished BRL49653-induced growth inhibition, but only marginally reduced the effect of TTA. TTA reduced tumor growth and increased the survival time of rats with implanted BT4Cn tumor. TTA-induced apoptosis in BT4Cn cells, and the administration of TTA led to cytochrome c release from mitochondria and increased the glutathione content in glioma cells. In conclusion, our results indicate that TTA inhibits proliferation of glioma cancer cells through both PPAR
-dependent and PPAR
-independent pathways, of which the latter appears to predominate.
Abbreviations: ANT, adenine nucleotide translocase; 
, mitochondrial membrane potential; NHR, nuclear hormone receptor; PPAR, peroxisome proliferator-activated receptor; PPRE, peroxisome proliferator response element; PTP, permeability transition pore; RXR, retinoid-x receptor; TTA, tetradecylthioacetic acid
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Introduction
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Development of cancer is associated with dysregulation of cell-cycle control, apoptosis and/or differentiation. Several members of the family of nuclear hormone receptors (NHR) play crucial roles in the control of cellular homeostasis, and administration of their cognate ligands has successfully been used in cancer treatment. Thus, retinoids have been used in the management of leukemia (1), head and neck cancer (2) and skin cancer (3). Recently, members of the peroxisome proliferator-activated receptor (PPAR) subfamily of the NHRs, have been demonstrated to be intimately involved in the control of cellular proliferation and differentiation, and accordingly, PPAR agonists have been employed in the treatment of certain cancers (4,5).
The PPAR family comprises PPAR
, PPAR
and PPAR
(also called PPARß, FAAR or NUC1). The PPARs bind as heterodimers with retinoic-x acid receptor (RXR) to a subset of DR-1 elements, peroxisome proliferator response elements (PPREs), and have been shown to regulate expression of genes involved in the transport, metabolism and storage of fatty acids. Transcriptional activation of the PPARRXR heterodimers is enhanced upon binding of a large variety of ligands including saturated and unsaturated fatty acids, arachidonic acid derivatives and a wide range of synthetic drugs with different subtype specificities [reviewed by (69)].
For the last 10 years, administration of peroxisome proliferators, PPAR
agonists, has been known to induce hepatocarcinogenesis in rodents (10). However, cancer development is probably induced by mechanisms secondary to PPAR
transcriptional activation (9). A role in growth regulation for the two remaining PPAR subtypes has also been suggested. Whereas up-regulation of PPAR
expression has been associated with colon cancer (11) and activation of PPAR
stimulates post-confluent proliferation of pre-adipocytes (12,13), opposite effects on cell proliferation are mediated by activation of PPAR
. This PPAR subtype is well known for its role in adipocyte differentiation (14). Furthermore, studies of different cancer cell lines have demonstrated that administration of PPAR
ligands causes G1-phase cell cycle arrest (1518) and induces expression of differentiation markers (1922). Furthermore, PPAR
agonists have been reported to induce apoptosis in certain settings (2325), supporting the idea that activation of PPAR
could be of therapeutic value in cancer treatment.
For the last decade, our group has focused on the hypolipidemic properties of the synthetic fatty acid analog tetradecylthioacetic acid (TTA) (26). The hypolipidemic effect of TTA is probably associated with the ability of TTA to activate PPAR
in the liver (27,28), as several genes containing PPREs are activated upon TTA treatment of rats and hepatocytes in culture. Recently, it has been shown that TTA is a ligand for all PPAR subtypes (2830) (L.Madsen, et al., submitted for publication). It is well documented that the PPAR
subtype plays a central role in hepatic lipid metabolism, but the tissue-specific expression patterns of the PPARs and the fact that PPAR
and PPAR
probably play important roles in fatty acid metabolism in most of these tissues, makes it probable that TTA could also act through PPARs outside the liver. The new data concerning PPAR
- and PPAR
-mediated regulation of proliferation make an analysis of the effects of TTA even more interesting.
Glioma cancers are among the most aggressive cancers and are difficult to treat with chemotherapeutic agents. The BT4Cn cell line is a permanent rat glioma cell line, originally induced in fetal rat brain cells after transplacental exposure to ethylnitrosourea (31). The BT4Cn cell line has been used to study the potentials of anticancer drugs and gene therapy in the treatment of glioma cancer (32,33). Whereas the stages of glioma growth with the concomitant changes in morphological structure are well characterized, relatively little is known about the underlying molecular events or the potential of intervention through transcription factors involved in growth regulation. In this study, we investigated the ability of TTA to affect glioma cell proliferation ex vivo and in vivo. Our results indicate that TTA inhibits BT4Cn growth by PPAR
-dependent and PPAR
-independent pathways, and that TTA induces apoptosis, possibly via release of cytochrome c from the mitochondria.
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Materials and methods
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Chemicals
Chemicals used were: TTA (prepared at the Department of Chemistry, University of Bergen, Norway), as described previously (34), Wy14,643 (Calbiochem, San Diego, CA), BRL49653 (kindly provided by Novo-Nordisk A/S, Bagsvaerd, Denmark), L-165041 (kindly provided by Merck Research Laboratories, Darmstadt, Germany) (35) and GW9662 (kindly provided by Glaxo Wellcome Research and Development, Uxbridge, UK) (36). Chemicals were dissolved in dimethyl sulfoxide (DMSO) and added to a final concentration of 0.1% DMSO in the culture medium. All other chemicals and solvents were of reagent grade from common commercial sources.
Cell culture
The continuous neoplastic cell line BT4Cn were obtained from fetal rat brain cells (31). The cells were grown at 37°C, 5% CO2 and 95% air in Dulbecco's modified Eagle medium [DMEM, high glucose (4.5 g/l)] (Gibco) supplemented with 10% fetal calf serum (Sigma, St. Louis, MO), streptomycin (100 µg/ml) and penicillin (62.5 µg/ml). In apoptosis and cytochrome c release experiments, cells were grown in DMEM supplemented with 10% new born calf serum, streptomycin (100 µg/ml), penicillin (62.5 µg/ml) and three times the prescribed concentration of non-essential amino acids (all from Sigma, St. Louis, MO). Media was changed routinely every 2 days and cells were passaged by trypsinization before confluence. When carrying out growth experiments, cells were plated at low density (~5% confluence) and allowed to settle for 4 h before the medium was changed to medium containing relevant chemicals.
Animal studies
The experiments described have been carried out according to the rules of the Norwegian Animal Research Authority (NARA). Male Norwegian brown rats, BD-IX, were obtained from the Gades Institute, Haukeland Hospital, Bergen, Norway. They were housed in cages in pairs, and maintained on a 12 h cycle of light and dark at a temperature of 20 ± 3°C. During the experiments they weighed 250400 g. They were fed a commercial standard pelleted food and provided tap water ad libitum. Fatty acids were administered by oro-gastric intubation. TTA and palmitic acid were dissolved in 0.5% (w/v) sodium carboxymethyl cellulose at a final stock solution of 75 mg/ml. The animals were administered once a day at a dose of 300 mg/kg body wt, and the feeding was started the day after tumor implantation.
The tumors were implanted subcutaneously or intracranially using stereotactical techniques (37). Before surgery the rats were anaesthetized with 0.2 ml Hypnorm-Dormicum/100 g body wt. For subcutaneous implantation, a tumor was established in vivo by subcutaneous injection of 5x106 tumor cells in the rat's neck. After ~2 weeks, the tumor was removed and cut into 2x2 mm tissue pieces and subcutaneously implanted in the leg. The tissue piece was entered and established 1 cm below the skin incision. The tumor sizes were measured daily and tumor tissue was collected and frozen in liquid nitrogen after ~2 weeks. The intracranially implanted rats were killed when neurological signs were evident.
Growth assays
DNA amounts were determined by fluorometry. Cell samples were harvested in triplicate by trypsinization and dissolved in 1 ml high salt harvest buffer (10 mM Tris, 10 mM EDTA, 2 M NaCl, pH 7.4). Samples were sonicated and properly diluted in high salt buffer (2 mM Tris, 2 mM EDTA, 2 M NaCl, pH 7.4) and the DNA amount was measured on a Hoefer DyNA Quant 200 in high salt buffer containing 0.1 µg/ml Hoechst 33258.
Plasmids
The PPREx3-tk-luc reporter construct containing three copies of the PPRE from the acyl-CoA promoter (38) was kindly provided by R.Evans. Expression vectors encoding murine PPAR
(pSG5PPAR
) (kindly provided by J.D.Tugwood) (6), murine PPAR
(pSG5PPAR
) (kindly provided by by P.Grimaldi) (39), murine PPAR
2 (pSPORTPPAR
2) (kindly provided B.Spiegelman) (40), and murine RXR
(pCMXRXR
) (kindly provided by R.Evans) (41) have been described previously.
Transient transfection
BT4Cn cells were plated in 6-well plates 24 h before transfection. One hour prior to the transfection the medium was changed. Each well was transfected with 3 µg reporter plasmid DNA, 1 µg PPAR expression vector or pSG5 and 1 µg of ß-galactosidase control plasmid by the calciumphosphate method. For transfections with PPAR
, 1 µg RXR
-vector was included. A total amount of 6 mg plasmid DNA/well was used. Five hours later the cells were shocked with 12.5% glycerol in phosphate-buffered saline (PBS) pH 7.3 at room temperature, and fresh media containing 10% (v/v) resin-charcoal-stripped calf serum (AG-1X-8 resin, Bio-Rad, Richmond, CA; Activated Powder Charcoal, Bie & Berntsen, Roedovre, Denmark) was added together with the indicated compound or vehicle (DMSO) alone. After 24 h, cells were harvested and the luciferase and ß-galactosidase activities measured in a Berthold MicroLumat LB96P luminometer using the commercial kit (Galacto-Light, Applied Biosystems, Foster City, CA). Luciferase values were normalized to the ß-galactosidase values.
RTPCR
Total RNA was purified, reverse transcribed (M-MLV Reverse Transcriptase kit, Life Technologies, Carlsbad, CA) and selected cDNA amplified by 25 or 28 cycles of PCR as described previously by Hansen et al. (42). TBP served as internal standard. The following primers were used: TBP: 5'-ACCCTTCACCAATGACTCCTATG-3' and 5'-ATGATGACTGCAGCAAATCGC3'; PPAR
: 5'-CCCTGCCTTCCCTGTGAACTGAC-3' and 5'-GGGACTCATCTGTACTGGTGGGGAC-3'; PPAR
: 5'-GCTATGACCAGGCCTGCAGG-3' and 5'-CCCTTGCACCCCTCACACGCGTC-3'; PPAR
: 5'-GAGCTGACCCAATGGTTGCTG-3' and 5'-GCTTCAATCGGATGGTTCTTC-3'. Fragments were separated on 6% polyacrylamide gels and bands were quantified by phosphoimager (Molecular Dynamics, Sunnyvale, CA) analysis.
Western blotting
One hundred micrograms of whole cell protein in SDS sample buffer (50 mM TrisHCl pH 6.8, 10% glycerol, 2.5% SDS, 10 mM DTE, 10 mM ß-glycerophosphate, 10 mM NaF, 0.1 mM Na orthovanadate, 1 mM PMSF, 1x CompleteTM) were separated by SDSPAGE (10% gels) and blotted onto a PVDF membrane (MSI). Equal loading and transfer were confirmed by Amido black staining. PPAR
was detected by enhanced chemiluminescence using mouse anti-PPAR
antibody (E-8, Santa Cruz Biotechnology, Santa Cruz, CA) and goat horseradish peroxidase-conjugated antimouse antibody (DAKO).
Glutathione assay
Cells suspended in distilled H2O were extracted with equal volumes of cold 5% sulfosalicylic acid with 50 µM dithioerythriol before they were placed at 80°C. Sample preparation and HPLC-analysis was performed according to the method described by Svardal et al. (43).
Determination of cytochrome c
The cells were harvested by scraping and washed in PBS before they were homogenized in H-buffer (0.25 M sucrose, 2 mM HEPES, 0.2 mM EGTA, pH 7.40) using a ball-bearing homogenizer (44) (EMBL, Heidelberg, Germany). Cell debris was removed by centrifugation at 1930 g for 15 min. The supernatant (total homogenate) was centrifuged at 80 000 g for 90 min for preparation of cytosolic (supernatant) fraction. Cytochrome c was determined using a rat/mouse cytochrome c immunoassay provided by R&D Systems, Minneapolis, MN.
Fluorescence microscopy
For microscopic analysis of apoptosis, cells were grown in 8-well chamber slides for 6 days. The cells were fixed in 4% formaldehyde over night and incubated for 30 min with 3 µg/ml Hoechst 33342 in the dark. The cells were analysed in a Leica Orthoplan fluorescence microscope.
Statistics
All fluorometry measurements were performed at least in triplicate and repeated a minimum of two times. Data were then normalized to the control of each experiment. To obtain a measure of dispersion for the overall mean of the control group, the weighed average of the sample standard deviation (SD) from all control experiments were used. A Dunnet two-tailed multiple comparison test were applied to test for equality of means of each sample compared with the control at the significance level
= 0.05 and
= 0.01.
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Results
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Inhibition of glioma cell growth
TTA has chemical and physical properties almost identical to normal saturated fatty acids, but TTA cannot undergo ß-oxidation (45). Figure 1
shows that the presence of 100 µm TTA reduced the accumulation of DNA by ~45% in rat BT4Cn glioma cells after 8 days of treatment, whereas the control (palmitic acid) had no effect on DNA accumulation. These data are in agreement with previous findings showing that TTA reduced [3H]thymidine incorporation in and reduced proliferation of both rat and human glioma cells (46).

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Figure 1. Effect of TTA (A) and palmitic acid (B) on DNA accumulation in BT4Cn cells. Cells were exposed to fatty acids (10, 30 or 100 µM) or vehicle (DMSO) for 8 days. DNA was determined by fluorometry, and the results represent measurement on 10 (TTA) or 2 (palmitic acid) independent experiments. DNA [means ± SD (n = 210)] is presented as percent of DMSO-control; *P < 0.01.
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Expression of PPARs in rat glioma BT4Cn cells
We have recently shown that TTA activates human PPAR
and PPAR
in a concentration-dependent manner (28). Activation of PPAR
has been associated with growth inhibition in several experimental systems and hence, the decreased BT4Cn cell proliferation could be mediated via PPAR activation. Using semi-quantitative RTPCR analysis, RNA expression of the different PPARs in BT4Cn cells was assayed. Expression of PPAR
mRNA was at the detection limit, whereas a significant expression of PPAR
and especially PPAR
mRNA was observed (Figure 2A
). We observed a marked 3-fold increase in the level of PPAR
mRNA at confluence. By 4 days of post-confluent growth, the level of PPAR
mRNA had declined to a level ~2-fold above that observed in exponentially growing cells. Expression of PPAR
followed a similar pattern. However, the magnitude of alterations varied from experiment to experiment, and the differences in expression levels were not statistically significant. Expression of PPAR
and PPAR
mRNA in confluent cells was approximately one-third to one-fourth of that observed in white adipose tissue, a tissue with known high expression of PPAR
, and a lower but still significant level of PPAR
expression (47). Western blotting was used to examine PPAR
expression at the protein level. As shown in Figure 2C
, the PPAR
protein level paralleled the PPAR
mRNA expression profile (Figure 2A and B
).
Transactivation of PPARs in rat glioma cells
Several recent reports have focused on the involvement of PPAR
in the inhibition of cancer cell growth (20,48,49). In contrast, overexpression of PPAR
has been linked to the development of colorectal cancers (11), and in addition, activation of PPAR
induces clonal expansion of pre-adipocytes (12). It is known that ligand-dependent PPAR-mediated transactivation exhibits cell type specificity. To examine how PPAR subtype-selective ligands and TTA-induced PPAR-dependent transactivation, BT4Cn cells were cotransfected with the PPAR responsive PPREx3-TK-luc reporter construct and PPAR
-, PPAR
- or PPAR
-expression vectors and then treated with PPAR-selective agonists or TTA. Figure 3A
shows that cotransfection with vectors expressing either PPAR subtype led to a significant transactivation of the reporter even in the absence of added ligand. Addition of Wy14,643 only very modestly enhanced PPAR
-mediated transactivation. More pronounced and dose-dependent induction of PPAR
- and PPAR
-mediated transactivation was observed with the PPAR
-selective ligand L165041 and the PPAR
-selective ligand BRL 49653, respectively. Interestingly, TTA activated all PPAR subtypes in a dose-dependent manner. We have shown previously that TTA most potently activates human PPAR
- and human PPAR
-mediated transactivation, and that higher concentrations of TTA were required to achieve significant activation of human PPAR
(28,30). In the present study using rodent (mouse) PPARs, a similar trend is apparent in that 100 µM TTA was needed to enhance PPAR
-mediated transactivation, whereas 30 µM TTA slightly enhanced PPAR
- and PPAR
-dependent transactivation suggesting that TTA activates rodent PPARs in BT4Cn glioma cells in the ranking order PPAR
PPAR
> PPAR
(Figure 3A
).

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Figure 3. The effect of TTA and PPAR-selective ligands on PPAR-mediated transactivation in BT4Cn cells. (A) BT4Cn cells were transiently transfected with the PPREx3-TK-luc reporter plasmid and the indicated PPAR expression vector and grown in the presence of PPAR-selective ligands, TTA or vehicle (DMSO) for 24 h. Ligands used were: Wy14,643 (10, 30 and 100 µM), BRL49653 (0.1, 0.5 and 1 µM), L-165041 (0.1, 0.5 and 1 µM) and TTA (10, 30 and 100 µM). Background activity from cells transfected with PPREx3-TK-luc reporter only and treated with vehicle is indicated by `-'. (B) BT4Cn cells were transiently transfected with the PPREx3-TK-luc reporter plasmid, and incubated in the presence or absence of PPAR-selective ligands and TTA for 24 h. Ligands used were: Wy (Wy14,643) (100 µM), BRL (BRL49653) (1 µM), L (L-165041) (1 µM) and TTA (100 µM). Results are presented as ß-galactosidase-normalized luciferase activity ± SD. Similar results were obtained in at least three separate experiments, each performed in duplicate.
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To examine the transactivation potential of endogenously expressed PPAR subtypes, BT4Cn cells were transiently transfected with the PPAR responsive PPREx3-TK-luc reporter plasmid, and treated with Wy14,643, L-165041, BRL 49653 or TTA using the concentrations shown to significantly enhance PPAR-mediated transactivation. In all cases, addition of ligands only slightly enhanced transactivation possibly reflecting the low levels of endogenous PPARs present in the BT4Cn glioma cells (Figure 3B
). Of the PPAR subtype-selective ligands, we found that BRL49653 most efficiently enhanced transactivation in keeping with PPAR
being the predominant PPAR subtype in BT4Cn cells.
Effect of PPAR ligands on BT4Cn growth
To investigate how PPAR activation affected BT4Cn proliferation, we carried out a series of experiments to determine the effect of PPAR-selective ligands. As a measure of cell proliferation we determined the accumulation of DNA in the cultures. Figure 4A and B
show that Wy14,643 and L-165041, at concentrations sufficient to activate PPAR
and PPAR
, respectively, did not affect the accumulation of DNA after 8 days of treatment. However, BRL49653 (Figure 4C
) reduced the DNA accumulation to ~85% of the control. Low concentrations of L-165041 were found to increase growth, supporting the notion that activation of PPAR
induces proliferation. The diminished effect observed by 1 µM L-165041 may relate to the ability of L-165041 to activate PPAR
(35). In conclusion, of the PPAR-selective ligands only BRL49653 was able to inhibit the growth of rat BT4Cn cells, but notably TTA was a much more potent inhibitor of growth than BRL49653.

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Figure 4. The effect of PPAR ligands on BT4Cn cell growth. Cells were treated with the PPAR ligands Wy14,643 (10, 30 and 100 µM) (A), L-165041 (0.1, 0.5, 1 µM) (B), BRL49653 (0.1, 0.5, 1 µM) (C), BRL49653 (1 µM), Wy14,643 (100 µM), L-165041 (1 µM) (D), Wy14,643 (100 µM), L-165041 (1 µM), BRL49653 (1 µM) (E) or TTA (100 µM), BRL49653 (1.0 µM) and/or the PPAR -selective antagonist GW9662 (1 and 10 µM) (F), as indicated for 8 days. DNA [means ± SD (n = 3)] was determined by fluorometry and is presented as percent of DMSO-control. *P < 0.01.
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The transfection analysis showed that 100 µM TTA activated all three PPAR subtypes (Figure 3
). Consequently, we analysed whether combinations of PPAR-selective ligands led to a more pronounced inhibition of proliferation than either ligand alone. None of the employed combinations of PPAR-selective ligands reduced growth beyond that observed with BRL49653 alone (Figure 4D
). Similarly, simultaneous addition of TTA and the PPAR-selective ligands did not result in an inhibition of growth exceeding that observed with TTA alone (Figure 4E
).
The results described above showed that BRL49653 and TTA both reduced growth with TTA being by far the most potent inhibitor. As TTA at 100 µM induced PPAR
-mediated transactivation, we decided to investigate to what extent PPAR
-mediated processes contributed to TTA-induced growth arrest. To this end, we took advantage of the PPAR
antagonist GW9662. In separate experiments we have shown that transactivation by 1 µM BRL49653 is completely abolished by the addition of 10 µM GW9662 (data not shown). Administration of 1 mM or 10 µM GW9662 did not affect the growth of BT4Cn cells. As described above, 1 µM BRL49653 modestly inhibited growth, whereas the BRL49653-mediated inhibition was neutralized by the addition of 10 µM GW9662. Administration of TTA reduced DNA accumulation to 55% of the control, and the simultaneous addition of the PPAR
antagonist only restored the DNA accumulation to 65% of the control (Figure 4F
). Thus, PPAR
-dependent processes contribute to TTA-mediated growth inhibition. However, the contribution of PPAR
- dependent pathways is clearly of minor importance in TTA-mediated arrest of BT4Cn cell proliferation.
TTA-induced apoptosis in BT4Cn cells
We then investigated whether apoptosis contributed to the reduction in DNA accumulation observed upon treatment of rat BT4Cn cells with TTA. Figure 5
shows that 200 µM TTA induced massive apoptosis in BT4Cn cells, as determined by chromatin condensation. When exponentially growing BT4Cn cells were treated with increasing concentrations of TTA we observed a dose-dependent increase in cells exhibiting morphological changes, including chromatin condensation, associated with apoptosis. Thus, following treatment for 24 h with 50, 100 and 200 µM TTA, 11%, 14% and 83% of the cells, respectively, appeared apoptotic. Less than 5% of the control cells showed morphological apoptotic characteristics.

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Figure 5. Apoptotic effect of TTA on BT4Cn cells. Cells were treated with vehicle (medium) (A) and 200 µM TTA (B) for 3 days. The cells were incubated with Hoechst 33342 and condensed chromatin was detected by fluorescence microscopy.
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Cytochrome c release from mitochondria, and subsequent activation of caspases are closely associated with induction and execution of the apoptotic process. Interestingly, TTA-induced apoptosis and increased considerably the release of cytochrome c from mitochondria in BT4Cn cells (Figure 6
).

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Figure 6. Cytochrome c release from mitochondria. Cells were grown in the presence or absence of TTA (200 µM) for 18 h. Cytochrome c [means ± SD (n = 2)] in the cytosolic fraction was measured as described in `Materials and methods', and is presented as ng/mg protein.
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Activation of apoptosis is associated with generation of reactive oxygen species (ROS), and TTA may activate mitochondrial ROS production as it affects the mitochondrial function (50). Furthermore, mitochondrial loss of cytochrome c stimulates mitochondrial superoxide production (51). Thus, ROS and the resulting cellular redox change can be a part of the signal transduction pathway leading to apoptosis. Interestingly, treatment with 100 µM TTA increased the cellular content of glutathione (Table I
). However, when compared with 100 µM TTA, treatment with 200 µM TTA gave a higher degree of apoptosis and resulted in a reduced cellular content of glutathione. These data indicate that TTA treatment may elicit compensatory adjustments in order to counteract changes in the cellular redox state associated with cytochrome c release and apoptosis.
The effect of TTA on tumor growth in vivo
To test whether TTA would also affect growth of glioma cells in a setting resembling the conditions for an in vivo glioma, rats were intracranially implanted with BT4Cn tumors. First, we examined the expression of PPARs in implanted tumors. Figure 8
shows that PPAR subtype expression in implanted tumors paralleled that observed for BT4Cn cells grown in culture (Figure 2
). PPAR
expression was at the detection limit and PPAR
was clearly the predominant PPAR subtype expressed in the tumors. In rats with intracranially implanted BT4Cn tumors, TTA increased the survival time compared with an equal dose of palmitic acid (Figure 7A
). Average survival time was 16.9 ± 1.8 and 14.6 ± 1.2 days for TTA and palmitic acid fed animals, respectively. In another set of in vivo experiments, TTA was shown to inhibit the growth of subcutaneously implanted BT4Cn tumors (Figure 7B
).

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Figure 8. Expression of PPAR mRNA in intracranial BT4Cn implants. RTPCR profiles of PPAR subtype mRNA using TBP as internal standard (25/28 cycles). PA, palmitic acid treatment; TTA, TTA treatment (n = 2).
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Discussion
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In the present study we show that TTA is able to inhibit rat glioma cell growth ex vivo and in vivo. PPAR
is the predominant PPAR subtype expressed in cultured BT4Cn cells and in intracranially implanted BT4Cn tumor cells (Figures 2A and 8
). Of the PPAR subtype-selective ligands, only BRL49653 inhibited the growth of BT4Cn cells ex vivo, an effect that was abolished by the PPAR
antagonist GW9662 (Figure 4F
). These data indicate that only PPAR
mediated ligand-induced inhibition of growth in BT4Cn cells. TTA was a far more potent inhibitor of growth than BRL49653 (Figure 4D and E
). The potential of TTA as a PPAR
ligand was confirmed in transfection experiments (Figure 3A and B
). Treatment with TTA in combination with BRL49653 did not lead to additional inhibition of growth (Figure 4E
), indicating that TTA could fully exploit the potential of PPAR
activation in growth inhibition. However, the irreversible PPAR
antagonist GW9662 did only partly reduce the TTA-mediated growth arrest, clearly demonstrating the existence of PPAR
-independent pathways in the antiproliferative activity of TTA. Moreover, our data show that PPAR
-independent pathways predominate in TTA-mediated growth inhibition.
BRL49653 reduced the accumulation of DNA in glioma cells and modestly induced apoptosis (data not shown). TTA increased the level of apoptosis in confluent BT4Cn cultures (Figure 5
). Thus, our results indicate that ligand-mediated activation of PPAR
elicited an apoptotic response, which contributed to the observed reduction in DNA accumulation in treated BT4Cn cells. The PPAR
-mediated induction of apoptosis may relate to the well-established negative cross talk between PPAR
and NF-kB. Administration of PPAR
ligands have been shown to inhibit NF-
B activity thus quenching anti-apoptotic signals (5255). However, our results also suggest that administration of TTA activates additional PPAR
- independent apoptotic pathways.
The induction of apoptosis observed in response to administration of TTA was paralleled by a substantial mitochondrial release of cytochrome c into the cytosol (Figure 6
). Cytosolic cytochrome c is a pro-apoptotic factor that induces caspase-driven apoptotic cascades (56,57). The exact mechanism behind the TTA-mediated release of cytochrome c is unknown. However, millimolar concentrations of long-chain 3-thia fatty acids have been shown to uncouple oxidative phosphorylation by a fatty acid cycle mechanism resulting in dissipation of the mitochondrial membrane potential (
) (50). Furthermore, it has been shown that long-chain fatty acids interact directly with and induces the opening of the mitochondrial permeability transition pore (PTP) in a Ca2+-dependent manner (58,59). The mitochondrial adenine nucleotide translocase (ANT) seems to be central to the protonophoric activity of long-chain fatty acids (60). ANT is assumed to be part of the PTP (61,62), and it has been reported that ANT is inhibited by long-chain 3-thia fatty acids (50). Mitochondrial permeability transition is considered to be a critical and rate-limiting event in apoptosis and can be triggered by oxidative stress, ADP deficiency and dissipation of 
. It is conceivable that interactions with ANT, dissipation of 
and triggered generation of reactive oxygen species are associated with increased release of cytochrome c, and thus play important roles in TTA-induced apoptosis.
Inhibition of mitochondrial ANT and release of cytochrome c into cytosol is thought to stimulate production of reactive oxygen species (51,63). Therefore, the increased level of glutathione detected in BT4Cn cells treated with low doses of TTA (Table I
) may represent an adaptation to the altered redox conditions in order to escape from cell death. In this notion, increasing the TTA dose to 200 µM apparently led to changes that were beyond the cellular adaptive capacity as a higher level of apoptosis was observed in these cultures. Furthermore, the concomitant reduction in glutathione content in these cells is in accordance with previous reports showing loss in glutathione during apoptosis (51,64). We suggest that the mitochondrial release of cytochrome c and altered cellular redox conditions following TTA treatment may well be a PPAR independent signal transduction pathway during apoptosis.
The limited therapeutic possibilities for the treatment of glioma cancer provide a strong stimulus for the identification of new and selective drugs and the search for molecular targets of relevance to cancer therapy. Studies in our laboratory have shown that 1-14C-labeled TTA is found in rat brain after intravenous administration, and thus is able to cross the bloodbrain barrier. As TTA may provide non-toxic therapy against human glioma cancers, TTA becomes a candidate for a new class of potent drugs in the treatment of glioma cancers.
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Notes
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3 These authors contributed equally to this work 
4 To whom correspondence should be addressed Email: kjetil.berge{at}ikb.uib.no 
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Acknowledgments
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We thank Bjorn Netteland for technical help. This work was supported by the University of Bergen, the Norwegian Research Council, the Norwegian Cancer Society, the Danish Biotechnology Program, the Danish Natural Science Research Council, the Danish Cancer Society and the Novo Nordisk Foundation. We are grateful to Novo Nordisk, Merck Research laboratories and Glaxo Welcome for providing ligands.
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References
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Received February 1, 2001;
revised May 22, 2001;
accepted June 22, 2001.