Journal of Leukocyte Biology Myeloid cells, immune suppression, tumor immunology
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(Journal of Leukocyte Biology. 2003;73:665-672.)
© 2003 by Society for Leukocyte Biology

Activation-induced PPAR{gamma} expression sensitizes primary human T cells toward apoptosis

Anja Tautenhahn, Bernhard Brüne and Andreas von Knethen

University of Kaiserslautern, Institute of Cell Biology, Erwin-Schroedinger-Strasse, Germany

Correspondence: Andreas von Knethen, University of Kaiserslautern, Institute of Cell Biology, Erwin-Schroedinger-Strasse 13/420A, 67663 Kaiserslautern, Germany. E-mail: aknethen{at}rhrk.uni-kl.de


   ABSTRACT
TOP
 ABSTRACT
INTRODUCTION
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 REFERENCES
 
Phytohemagglutinin (PHA) elicited expression of peroxisome proliferator-activated receptor {gamma} (PPAR{gamma}) in primary human T cells via the PPAR{gamma}3 promoter, as shown by reverse transcription-polymerase chain reaction. Electrophoretic mobility shift assay demonstrated no correlation between PPAR{gamma} expression and its activation. However, addition of specific PPAR{gamma} agonists such as ciglitazone or 15-deoxy-{Delta}12,14-prostaglandin J2 (15d-PGJ2) for 1 h following PHA pretreatment provoked PPAR{gamma} activation verified by supershift analysis. Taking the proapoptotic properties of PPAR{gamma} into consideration, we analyzed induction of apoptosis in activated T cells in response to PPAR{gamma} agonists. Cells exposed to PPAR{gamma} agonists alone revealed minor cell death compared with controls, whereas treatment with 15d-PGJ2 or ciglitazone for 4 h subsequent to PHA stimulation significantly increased cell demise, which was attenuated by the pan-caspase inhibitor zVAD, pointing to apoptosis as the underlying mechanism. These data may be relevant for pathophysiological conditions accompanied with lymphopenia of T cells under conditions such as sepsis.

Key Words: inflammation • lymphocyte • cell death


   INTRODUCTION
TOP
 ABSTRACT
 INTRODUCTION
MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 REFERENCES
 
T cells are important players of the acquired immune system, and diseases linked with T cell apoptosis, such as sepsis, cause severe, pathological effects. Hence, understanding mechanisms leading to T cell apoptosis or necrosis, thereby affecting the immune state of the body, is of great interest. With regard to induction of programmed T cell death, FAS and tumor necrosis factor gained considerable attention [1 ]. More recent studies focused on the new candidate, peroxisome proliferator-activated receptor {gamma} (PPAR{gamma}), leading to T cell apoptosis when appropriately activated [2 , 3 ]. The anti-inflammatory and proapoptotic properties of the nuclear hormone receptor family, known as PPARs, originally identified in association with obesity, diabetes, and arteriosclerosis, were recently established [4 , 5 ]. So far, three subtypes, PPAR{alpha}, PPARß (also known as PPAR{delta}), and PPAR{gamma}, have been elucidated [6 ]. Upon ligand binding, the transcription factor translocates into the nucleus, forms a heterodimer with the retinoic acid receptor, binds to its specific recognition site in the promoter of target genes, and modulates its transcription. Lately, PPAR{gamma} has been described to act in an anti-inflammatory manner in cells of the immune system. In primary murine- and human-resting T cells, PPAR{gamma} is expressed [3 , 7 ], although contradictory reports exist showing that resting T lymphocytes do not express PPAR{gamma} [8 , 9 ]. In transformed T cell lines such as Jurkat and CEM, moderate PPAR{gamma} expression is observed [8 ]. Biological effects of PPAR{gamma} demand its activation. Therefore, in T cells, expression and activation of PPAR{gamma} inhibit T cell activation by scavenging the transcription factor, nuclear factor of activated T cells (NF-AT), known to be responsible for interleukin-2 (IL-2) expression [10 ]. Moreover anti-CD3-induced expression of interferon-{gamma} was suppressed in CD4+T cells in response to PPAR{gamma} activation [7 ]. In addition, it is acknowledged that activated protein-1 and NF-{kappa}B are attenuated by PPAR{gamma} in T cells [11 ]. Besides its anti-inflammatory effect, which lowers proinflammatory cytokine expression, PPAR{gamma} is known to induce T cell apoptosis in the murine system when its physiological agonist 15-deoxy-{Delta}12,14-prostaglandin J2 (15d-PGJ2) or the synthetic compound troglitazone was supplied simultaneously with T cell-activating agents such as phorbol 12-myristate 13-acetate, ionomycin or antigen, and antigen-presenting cells [3 ]. In transformed, human T cell lines, PPAR{gamma} agonists directly induce apoptosis without pretreatment of cells [8 ].

Concerning these properties of PPAR{gamma}, we hypothesized a role of PPAR{gamma} in mediating a feedback loop in T cells. Following T cell activation, PPAR{gamma} expression is induced to orchestrate T cell response. As a test system, we studied PPAR{gamma} expression in Jurkat T cells and in primary human T cells derived from peripheral blood in response to phytohemagglutinin (PHA) treatment. Induction of PPAR{gamma} provoked by PHA showed no DNA-binding capacity but sensitized T cells toward PPAR{gamma} agonist-mediated apoptosis. We conclude that expression of PPAR{gamma} in activated T cells may be one likely mechanism to regulate their immunological response in inflammation, depending at least in part on neighboring cells, such as monocytes, macrophages, or endothelial cells, to provide PPAR{gamma} agonists.


   MATERIALS AND METHODS
TOP
 ABSTRACT
 INTRODUCTION
 MATERIALS AND METHODS
RESULTS
 DISCUSSION
 REFERENCES
 
Materials
Diethylamine (DEA)-NONOate and PHA were purchased from Sigma (Deisenhofen, Germany). The polyclonal anti-PPAR{gamma} antibody was from Alexis (Grünberg, Germany). The polyclonal anti-PPAR{alpha}- and anti-Fas-neutralizing antibodies and the monoclonal anti-Fas antibody were obtained from Santa Cruz Biotechnology (Heidelberg, Germany). Cold shock protein D (CspD) and the alkaline phosphatase-labeled anti-digoxigenin (DIG) antibody were from Roche (Mannheim, Germany). Culture supplements and fetal calf serum (FCS) were ordered from Biochrom (Berlin, Germany). The pan-caspase inhibitor zVAD, 15d-PGJ2, WY14643, and ciglitazone were bought from Biomol (Hamburg, Germany). Oligonucleotides were ordered from Qiagen Operon (Hilden, Germany) and Eurogentec (Seraing, Belgium). All chemicals were of the highest grade of purity and were commercially available.

Cell culture
The human T cell leukemia Jurkat were maintained in RPMI 1640 supplemented with 100 U/ml penicillin, 100 µg/ml streptomycin, and 10% heat-inactivated FCS (complete RPMI). All experiments were performed using complete RPMI. 15d-PGJ2, WY14643, and ciglitazone were dissolved in dimethyl sulfoxide and added as indicated.

T cell isolation and culture
Whole blood (36 ml) obtained from healthy donors was collected by the Vacutainer CPT tube system (Becton Dickenson, Heidelberg, Germany) using the standard technique proposed. The cell layer containing peripheral blood mononuclear cells (PBMC) was received following centrifugation (1500 g, 15 min, 24°C) within 1 h of blood collection. Cells were recovered and washed twice in phosphate-buffered saline (PBS), and T cells were isolated using the magnetic cell-sorting technology (Miltenyi Biotec, Bergisch Gladbach, Germany), as described by the supplier to enrich CD3+ cells. In brief, 107 total PBMC were resuspended in 80 µl PBS supplemented with 2 mM EDTA and 0.5% bovine serum albumin (PBS/E/B). Then 20 µl CD3 MicroBeads per 107 total cells was added. Cells were mixed, incubated for 15 min at 4°C, washed with PBS/E/B, and resuspended in 500 µl PBS/E/B per 108 cells. The LS+ column was placed in the magnetic field of a MidiMacs separator and washed with 3 ml PBS/E/B before addition of the cells; unlabeled cells passed through. The positive cells were flushed out of the column with 5 ml PBS/E/B using the plunger supplied. Flow cytometry confirmed that this population was 95–98% pure (CD3+ vs. CD3–). Cells were grown in 24-well tissue plates (Greiner Labortechnik, Frickenhausen, Germany) in complete RPMI. After 2 h of recovering, cells were used for the experiments.

Measurement of mitochondrial membrane potential ({Delta}{psi})
Following individual incubations, cells were loaded with 40 nM fluorochrome 3,3'-dihexyloxacarbocyanide iodide (DiOC6(3); Molecular Probes, Leiden, The Netherlands) for 30 min, after which the dye is accumulated in mitochondria that contain an intact membrane potential [12 ]. After PHA and agonist treatment, {Delta}{psi} was measured on a FACSCalibur flow cytometer (Becton Dickinson) using CellQuestPro software. At least 10,000 cells were accumulated for analysis. Results are given here in percentage of total cells with intact {Delta}{psi}.

Annexin V-fluorescein isothiocyanate (FITC)/propidium iodide (PI) staining
Following depicted incubations, 2 x 105 cells in 100 µl binding buffer were labeled with 5 µl Annexin V-FITC and 2.5 µl PI for 15 min on ice in the dark to differentiate between apoptotic and necrotic cell death using an Annexin V-FITC/PI-staining kit (Immunotech, Marseille, France). Afterward, 150 µl binding buffer was added, and cell samples were analyzed immediately using a FACSCalibur flow cytometer and CellQuestPro software. A minimum of 10,000 cells was analyzed.

Nuclear protein extraction
Preparation of crude nuclear extract was basically as described [13 ]. Briefly, following cell activation for the times indicated, 4 x 106 cells were washed in 1 ml ice-cold PBS, centrifuged at 1000 g for 5 min, resuspended in 400 µl ice-cold hypotonic buffer [10 mM HEPES/KOH, 2 mM MgCl2, 0.1 mM EDTA, 10 mM KCl, 1 mM dithiothreitol (DTT), 0.5 mM phenylmethylsulfonyl fluoride (PMSF), pH 7.9], left on ice for 10 min, vortexed, and centrifuged at 15,000 g for 30 s. Sedimented nuclei were resuspended in 50 µl ice-cold saline buffer (50 mM HEPES/KOH, 50 mM KCl, 300 mM NaCl, 0.1 mM EDTA, 10% glycerol, 1 mM DTT, 0.5 mM PMSF, pH 7.9), left on ice for 20 min, vortexed, and centrifuged at 15,000 g for 5 min at 4°C. Aliquots of the supernatant containing nuclear proteins were frozen in liquid nitrogen and stored at -70°C. Protein was determined using a Bio-Rad II kit.

Electrophoretic mobility shift assay (EMSA)
An established EMSA method with slight modifications was used [14 ]. Nuclear protein (10 µg) was incubated for 20 min at room temperature with 2 µg poly(dI-dC) from Pharmacia, 2.5 µl buffer D (20 mM HEPES/KOH, 20% glycerol, 100 mM KCl, 0.5 mM EDTA, 0.25% Nonidet P-40, 2 mM DTT, 0.5 mM PMSF, pH 7.9), 5 µl buffer F (20% Ficoll-400, 100 mM HEPES/KOH, 300 mM KCl, 10 mM DTT, 0.5 mM PMSF, pH 7.9), and 200 fmol 5'-DIG-labeled oligonucleotide in a final volume of 25 µl. Supershift antibodies (2 µg) were added as indicated. DNA–protein complexes were resolved at 80 V for 1 h in a taurine-buffered, native 6% polyacrylamide gel (4% for supershift), blotted onto a positively charged nylon membrane (Nytran Supercharge, Schleicher and Schuell, Keene, NH), and cross-linked for 5 min on a UV-transilluminator. DNA was detected using an alkaline phosphatase-labeled anti-DIG antibody and visualized using the substrate CSPD according to the manufacturer’s protocol. Oligonucleotides with the consensus PPRE site (bold letters) were used [15 ]: 5'-GGT AAA GGT CAA AGG TCA AT-3'; 3'-A TTT CCA GTT TCC AGT TAG CCG-5'.

RNA extraction and semiquantitative reverse transcriptase-polymerase chain reaction (RT-PCR)
RNA was extracted using peqGOLD RNAPure (Peqlab, Erlangen, Germany) according to the distributor’s manual. RT reactions and PCR for human PPAR{gamma}, IL-2, and glyceraldehydes 3-phosphate dehydrogenase (GAPDH) were performed using the Advantage RT-for-PCR kit and the Advantage 2 polymerase mix (Clontech, Heidelberg, Germany). Sequences of the primers were as follows: PPAR{gamma} (548–1262) [16 ], TA = 63°C: 5' > 3' ATG GCC ATT GAG TGC CGA GTC TG, 3' > 5' GGC TTT TGA GGA ACT CCC TGG TCA; PPAR{gamma} promoter 1 (PPAR{gamma}1) [17 ], TA = 63°C: 5' > 3' GGT CGG CCT CGA GGA CAC CG; PPAR{gamma} promoter 2 [17 ], TA = 63°C: 5' > 3' GTG AAT TAC AGC AAA CCC CTA TTC CAT GC; PPAR{gamma} promoter 3 [18 ], TA = 63°C: 5' > 3' CAT TTT GTC AAC GAG AGT CAG CCT TTA ACG; IL-2 (241–456) [19 ], TA = 61°C: 5' > 3' TTA AGT TTT ACA TGC CCA AGA AGG CC, 3' > 5' ACC AAC GAC AGA GTA GAC GTA TAA GT; GAPDH (47–1048) [20 ], TA = 63°C: 5' > 3' ATG GTG AAG GTC GGT GTG AAC GG, 3' > 5' TTA CTC CTT GGA GGC CAT GTA GGG C.

Annealing temperatures were calculated using the primer design program Oligo (Molecular Biology Insights, Colorado Springs, CO). The number of amplification cycles (25 for GAPDH; 30 for IL-2 and PPAR{gamma}) was necessary to achieve exponential amplification, where product formation is proportional to starting DNA. PPAR{gamma} promoter use was determined using a specific 5' primer (PPAR{gamma}1, PPAR{gamma}2, PPAR{gamma}3) and the common 3' primer from the PPAR{gamma} primer set. Products were run on a 1% agarose gel and ethidium bromide-stained. Controls of isolated RNA omitting RT were used during PCR to guarantee DNA-free, RNA preparations (data not shown).

Statistical analysis
Each experiment was performed at least three times, and statistical analysis was performed using the two-tailed Student’s t-test. Otherwise, representative data are shown.


   RESULTS
TOP
 ABSTRACT
 INTRODUCTION
 MATERIALS AND METHODS
 RESULTS
DISCUSSION
 REFERENCES
 
PPAR{gamma} is induced in response to T cell activation
In a first set of experiments, we stimulated Jurkat T cells with increasing concentrations (0.5, 1, and 5 µg/ml) of PHA, a known T cell mitogen. Cells were harvested after 6, 15, and 24 h. RT-PCR revealed that in Jurkat T cells treated with 5 µg/ml, PHA expression of PPAR{gamma} was slightly but significantly induced after 6 h, reaching a plateau after 15 h and staying elevated at 24 h (Fig. 1 A ). These results were reproduced in primary T cells derived from peripheral blood from healthy donors. In these cells, PPAR{gamma} was clearly induced by 10 µg/ml PHA after 24 h and was more pronounced at 48 h (Fig. 2 A ). Activation of T cells was verified by IL-2 RT-PCR, a cytokine that is known to be expressed in response to mitogen stimulation (Figs. 1B and 2B) . As expected, IL-2 expression was induced in response to PHA in a time- and concentration-dependent manner. Samples were normalized to a GAPDH standard (Figs. 1C and 2C) .



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  		Figure 1. Expression of PPAR{gamma} in activated Jurkat T cells. Cells were stimulated with 0.5, 1, and 5 µg/ml PHA for 6, 15, and 24 h or remained as controls. Cells were harvested, total RNA was isolated for semiquantitative RT-PCR, and expression of (A) PPAR{gamma} and (B) IL-2 was analyzed. PCR results are normalized for (C) GAPDH expression. Data are representative of three similar experiments.

 


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  		Figure 2. Expression of PPAR{gamma} in activated CD3+ cells. Primary human T cells were stimulated for 24 h or 48 h with 5 and 10 µg/ml PHA or remained as controls. Cells were harvested, total RNA was isolated for semiquantitative RT-PCR, and expression of (A) PPAR{gamma} and (B) IL-2 was analyzed. PCR results are normalized for (C) GAPDH expression. Data are representative of three similar experiments.

 
PPAR{gamma} expression is conferred via its {gamma}3 promoter
PPAR{gamma} is known to be expressed from three different promoters [17 , 18 ]. Of note, contradictory reports exist describing the promoter used for PPAR{gamma} expression in T cells [3 , 21 , 22 ]. To clarify this question, we analyzed the promoter used in our system. As shown in Figure 3 , RT-PCR, using three sets of primers, each with the same 3' primer but a promoter-specific 5' primer, established PPAR{gamma} induction via {gamma}3 in Jurkat (Fig. 3A) as well as primary human T cells (Fig. 3B) .



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  		Figure 3. PPAR{gamma}3 promoter-dependent expression of PPAR{gamma} in activated T cells. (A) Jurkat T cells were stimulated with 5 µg/ml PHA for 15 h. Cells were harvested, and total RNA was isolated for semiquantitative RT-PCR. (B) Primary human T cells were stimulated for 24 h with 10 µg/ml PHA or remained as controls. Cells were harvested, and total RNA was isolated for semiquantitative RT-PCR. Data are representative of three similar experiments.

 
PPAR{gamma} is expressed but not activated
Taking into consideration that PPAR{gamma} expression may be directly linked to its activation, we went on to analyze the DNA-binding capacity of PPAR{gamma} using the EMSA. As shown in Figure 4 , nuclear extracts prepared from cells treated for 15 h and 24 h with increasing concentrations of PHA (0.5, 1, or 5 µg/ml) did not reveal significant DNA binding of PPAR{gamma} compared with untreated controls. These data suggest that in T cells, expression of PPAR{gamma} can be separated from its activation. To prove the assumption that PPAR{gamma} can bind to DNA after providing an exogenous ligand, we evaluated activation of PPAR{gamma} in response to PPAR{gamma} agonists. Experimentally, Jurkat T cells were treated for 15 h with 5 µg/ml PHA. Afterward, the PPAR{gamma}-specific synthetic ligand ciglitazone (3 µM) was added for 1 h. As shown in Figure 5A , 3 µM ciglitazone enhanced binding of PPAR{gamma} to DIG-labeled oligonucleotides compared with the PHA-treated sample (Fig. 5A , lanes 6 vs. 5), which showed only a minor signal in Jurkat T cells. Importantly, in cells treated with 3 µM ciglitazone for 1 h, a PPAR{gamma} DNA-binding activity was missing (Fig. 5A , lane 3). Also, the physiological PPAR{gamma} agonist 15d-PGJ2 (1 µM) caused PPAR{gamma} activation (Fig. 5B , lane 2). Taking inflammatory conditions into consideration, we were also interested in testing the effect of nitric oxide (NO) on PPAR{gamma} activation. It is interesting that NO, supplied from DEA-NONOate (10 µM), provoked PPAR{gamma} DNA binding in activated Jurkat cells (Fig. 5B , lane 3). To provide unequivocal proof for the involvement of PPAR{gamma} in DNA binding, we conducted a supershift analysis (Fig. 5C) . PPAR{gamma} activation achieved with 3 µM ciglitazone in PHA (10 µg/ml)-pretreated, primary T cells was supershifted with a PPAR{gamma} antibody, whereas an unrelated PPAR{alpha} antibody left the response unaltered.



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  		Figure 4. Activation of PPAR{gamma} in response to PHA. Activation of PPAR{gamma} was analyzed by EMSA using a specific PPAR response element (PPRE) oligonucleotide, as described in Materials and Methods. Jurkat T cells were stimulated for (A) 15 h or (B) 24 h with 0.5, 1, and 5 µg/ml PHA. In controls, cell stimulation was omitted. Data are representative of experiments performed at least three times.

 


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  		Figure 5. Activation of PPAR{gamma} in response to agonists. Activation of PPAR{gamma} was analyzed by EMSA using a specific PPRE oligonucleotide, as described in Materials and Methods. (A) Jurkat T cells were stimulated for 15 h with 5 µg/ml PHA. One hour before preparation of nuclear extracts, 3 µM ciglitazone was added. For controls, cell stimulation was omitted. Data are representative of three similar experiments. (B) Jurkat T cells were stimulated for 15 h with 5 µg/ml PHA. One hour before preparation of nuclear extracts, 10 µM DEA-NONOate or 1 µM 15d-PGJ2 was added. For controls, cell stimulation was omitted. Data are representative of three similar experiments. (C) Supershift analysis of the active PPAR complex was performed as described in Materials and Methods. Primary human T cells were stimulated for 24 h with 10 µg/ml PHA and further for 4 h with 3 µM ciglitazone. For supershift analysis, a PPAR{gamma} antibody (lane 2) or a PPAR{alpha} antibody (lane 3) was included. Lane 1, PPAR analysis without antibody addition. Data are representative of three similar experiments.

 
PPAR{gamma} sensitizes transformed T cells toward apoptosis
Taking the anti-inflammatory, proapoptotic properties of PPAR{gamma} into account, we performed a set of experiments to analyze the effect of PPAR{gamma} activation in T cell apoptosis by determining the {Delta}{Psi} using DiOC6(3) (Figs. 6 and 7 ) as a marker of apoptosis. Jurkat T cells treated for 19 h with 5 µg PHA revealed that 14.5 ± 10% of all cells lost their {Delta}{Psi}, thus indicating apoptosis (Fig. 6A) . 15d-PGJ2 (1 µM), being added for the last 4 h of PHA stimulation, enhanced apoptosis in Jurkat cells up to 58.7 ± 14% (Fig. 6B) . Treatment of the cells with 1 µM 15d-PGJ2 for 4 h only did not alter Jurkat T cell apoptosis 3.8 ± 2% (Fig. 6C) compared with the untreated control (data not shown). To gain insight into mechanisms leading to cell death, we conducted a set of experiments applying zVAD. For these experiments, we used DiOC6(3) and Annexin V-FITC/PI staining to support our hypothesis regarding the involvement of apoptosis (Fig. 7) . As shown in Figure 7A , treatment of Jurkat T cells for 19 h with 5 µg/ml PHA and 3 µM ciglitazone being present for the last 4 h revealed 79.5 ± 12% apoptosis. Loss of cell viability after PHA/ciglitazone treatment was markedly reduced (26.8±11%) in the presence of zVAD, as illustrated in Figure 7B . Adding ciglitazone for 4 h only elicited 19.4 ± 7% apoptosis, as depicted in Figure 7C . To verify results obtained by the DiOC6(3) method, we analyzed Annexin V-FITC/PI staining in parallel to asses apoptotic (early apoptosis Annexin V-FITC-positive and late apoptosis Annexin V-FITC/PI-positive) and necrotic (only PI-positive) cells. Results are presented in insets shown in Figure 7 . The PPAR{gamma}-specific, synthetic agonist ciglitazone induced a minor apoptotic response (16.2±9%) using Annexin V-FITC/PI (Fig. 7C , inset). Pretreatment of cells with 5 µg/ml PHA for 15 h and coincubation for an additional 4 h with 3 µM ciglitazone resulted in a dramatic increase of apoptotic cell death (81.7±7%), as given in the inset of Figure 7A . zVAD suppressed ciglitazone-mediated cell death in PHA-activated Jurkat T cells, which is shown in the inset of Figure 7B . Cell death was significantly reduced to 25.0 ± 7%. Therefore, results obtained by {Delta}{Psi} staining [DiOC6(3)] were corroborated by Annexin V-FITC/PI staining.



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  		Figure 6. PPAR{gamma} agonists trigger loss of {Delta}{Psi}. Jurkat T cells (1x106) were incubated with (A) 5 µg/ml PHA for 19 h, (B) 5 µg/ml PHA for 15 h and further for 4 h with 1 µM 15d-PGJ2, or (C) 1 µM 15d-PGJ2 for 4 h. Percentage of viable cells was analyzed using DiOC6(3) by flow cytometry as described in Materials and Methods. Data are representative of three similar experiments.

 


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  		Figure 7. PPAR{gamma} agonists trigger apoptosis. Jurkat T cells (1x106) were incubated with (A) 5 µg/ml PHA for 15 h and further for 4 h with 3 µM ciglitazone, (B) 5 µg/ml PHA for 15 h and subsequently with a combination of 3 µM ciglitazone and 100 µM zVAD for 4 h, or (C) 3 µM ciglitazone for 4 h. Percentage of viable cells was analyzed using DiOC6(3) and Annexin V-FITC/PI staining in parallel by flow cytometry as described in Materials and Methods. Data are representative of three similar experiments.

 
Taking into account that Fas-mediated apoptosis is an important mechanism of programmed cell death in T cells [1 ], we elucidated its role in our system by including an anti-Fas-neutralizing antibody. The system was established with significant reduction of anti-Fas (0.05 µg/ml, 4 h)-induced apoptosis (set as 100%) to 58.4 ± 4%, as expected when cells were preincubated with anti-Fas-neutralizing antibody (0.5 µg/ml, 1 h). In contrast, ciglitazone-elicited apoptosis was only marginally affected. Apoptosis (set as 100%) was reduced by 7.7 ± 3%, which points to Fas-independent cell death. Consequently, PHA-induced PPAR{gamma} expression sensitized Jurkat T cells toward PPAR{gamma} agonist-mediated apoptosis in a Fas-independent manner. To verify the physiological importance of these results, experiments were repeated using primary blood-derived T cells.

PPAR{gamma} sensitizes primary T cells toward apoptosis
As shown in Figure 8 A , untreated, primary T cells showed 19.2 ± 8% apoptosis after 28 h. Addition of 10 µg/ml PHA for 28 h left the amounts of cells with intact mitochondria (86.5±8%) comparable with controls (Fig. 8B) . Treatment with 3 µM ciglitazone only resulted in no apoptotic response (20.0±5%) compared with the PHA-treated or vehicle-exposed controls (Fig. 8C vs. A and B). In contrast, application of 3 µM ciglitazone to activate PPAR{gamma} for the last 4 h of PHA stimulation increased T cell apoptosis to 43 ± 9% after 28 h (Fig. 8D) . Administration of WY14643 for 4 h resulted in 21 ± 11% apoptosis, which was not significantly different from untreated controls (Fig. 8 ,E vs. A). The PPAR{alpha}-specific agonist WY14643 (10 µM) did not alter the level of apoptosis (17.6±9%) in response to PHA stimulation as shown in Figure 8F . Addition of anti-Fas-neutralizing antibody (0.5 µg/ml) to PHA-activated cells 1 h before ciglitazone treatment resulted in 10.5 ± 3% reduction of cell death, which points to receptor-independent apoptosis. Therefore, we propose a Fas-unrelated but PPAR{gamma}-mediated effect leading to apoptosis.



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  		Figure 8. PPAR{gamma} agonists trigger apoptosis in activated, primary human T cells. Primary T cells (1x106) were (A) untreated, (B) stimulated with 10 µg/ml PHA for 28 h, (C) incubated with 3 µM ciglitazone for 4 h, (D) prestimulated with 10 µg/ml PHA for 24 h and further for 4 h with 3 µM ciglitazone, (E) incubated with 10 µM WY14643 for 4 h, and (F) stimulated with 10 µg/ml PHA for 24 h and additionally with 10 µM WY14643 for 4 h. Percentage of viable cells was determined using DiOC6(3) by flow cytometry as described in Materials and Methods. Data are representative of three similar experiments.

 

   DISCUSSION
TOP
 ABSTRACT
 INTRODUCTION
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
REFERENCES
 
Here, we showed induction of PPAR{gamma} via its {gamma}3 promoter by promoter-specific RT-PCR in Jurkat as well as human primary peripheral blood-derived T lymphocytes in response to PHA stimulation. PPAR{gamma} expression in T cells was not correlated with its activation. Thus, addition of specific PPAR{gamma} agonists such as ciglitazone or 15d-PGJ2 following PHA pretreatment provoked PPAR{gamma} activation as defined by EMSAs with the notion that supershift analysis verified the involvement of PPAR{gamma}. By this means, PPAR{gamma} activation likely gave rise to an increase of cell death as shown by DiOC6(3) and AnnexinV staining. Cell death occurred from apoptosis in a caspase-dependent manner, as cell death was attenuated by zVAD. Apoptosis, however, was not suppressed by addition of the anti-Fas-neutralizing antibody before incubation with ciglitazone, thus pointing to Fas-independent apoptosis signaling. Our data support the concept that PPAR{gamma} is one important regulator of human T cell function, not only by inhibiting T cell activation [10 ] but by modulating survival of activated T cells during inflammation.

The transcription factor PPAR{gamma} is involved in regulating immune functions [5 ]. Inhibition of proinflammatory cytokine expression as well as induction of apoptosis have been noticed in response to its activation in various cellular systems [2 , 3 , 8 , 10 , 11 , 23 24 25 26 27 ]. In T cells, PPAR{gamma} was shown to inhibit proliferation and activation when provided simultaneously with the T cell-activating agents by inhibiting NF-AT-dependent gene expression such as IL-2 synthesis [3 , 8 , 10 ]. In this report, we focused our attention on expression of PPAR{gamma} in primary human and Jurkat T cells in response to the mitogen PHA. We found that stimulation of T cells with PHA provoked induction of PPAR{gamma} in a time- and concentration-dependent manner, which is in line with the report of Cunard et al. [28 ] showing a similar phenomena in murine splenocytes. Taking into consideration that PPAR{gamma} can be expressed from three different promoters in the human system [17 , 18 ], we demonstrated the involvement of the PPAR{gamma}3 promoter by promoter-specific RT-PCR, not only in Jurkat but also in primary T cells. EMSAs were performed to evaluate the DNA-binding capacity of PPAR{gamma} in activated T cells, which revealed no binding of the transcription factor to its specific PPRE. Therefore, expression of PPAR{gamma} in activated T cells is not correlated to its activation. Consequently, PPAR{gamma}-specific agonists have to be provided exogenously. We demonstrated that addition of the PPAR{gamma} physiological agonist 15d-PGJ2, the synthetic compound ciglitazone, and NO donors provoke DNA binding of PPAR{gamma} in T cells, which is corroborated for other cell systems such as mesangial cells and macrophages [29 , 30 ]. These results have potential physiological significance, as agonists or activators of PPAR{gamma} can be released by neighboring cells, such as endothelial cells or macrophages known to express inducible NO synthase, cyclooxygenase-2, or 12/15-lipoxygenase in inflammatory tissue as established sources for PPAR{gamma}-activating mediators [21 , 31 32 33 ]. Activation of PPAR{gamma} in prestimulated T cells by the addition of exogenous agonists or activating compounds, using concentrations that provoked DNA binding, led in turn in our experimental model to an increase of cell death as shown by the breakdown of the {Delta}{Psi} (DiOC6(3)) and phosphatidylserine exposure at the outer leaflet of the plasma membrane (Annexin-V staining), thus pointing to apoptosis as the underlying mechanism. Further evidence for apoptotic cell death came from the use of zVAD, which significantly lowered PPAR{gamma}-dependent cell death in activated T cells. In some analogy to our analysis, Wang et al. [9 ] provided evidence that PPAR{gamma}-specific agonists, such as 15d-PGJ2 or troglitazone, provoked transcriptional activity in murine T cells, but in contrast to our work, no apoptosis was observed. The difference between their study and our study might be a result of divergences in the experimental setup. Their mechanistic studies were performed using the early murine hematopoietic F5.12 cell line carrying an expression plasmid for PPAR{gamma}1, whereas our experiments were done with stimulated human T cells. However, recent findings revealed that some effects of thiazolidinediones and cyclopentones could occur independent of PPAR{gamma} activation [34 , 35 ]. To rule out any unspecific impact, the synthetic, PPAR{alpha}-specific agonist WY14643 [36 ] was included in our experiments, showing no impact on apoptosis of PHA-treated T cells, thus indicating a likely PPAR{gamma}-dependent mechanism.

Involvement of death domain receptors in apoptosis upon PPAR{gamma} activation was recently described in A549 cells [37 ]. In this study, apoptosis was monitored 72 h after initial stimulation. The authors provide evidence showing that in epithelial cells, PPAR{gamma}-conferred apoptosis is death domain-dependent, mediated by caspase-8 activation, related to bax induction but unrelated to Bid-cleavage or cytochrome C release. In our hands, apoptosis occurred in T cells no later than 4 h after agonist addition in a Fas-independent manner. Different cell systems as well as different time points might be explanations for these contrasting data. More experiments have to be performed to elucidate the detailed pathway responsible for PPAR{gamma}-mediated apoptosis in T cells. Induction of PPAR{gamma} in response to T cell activation most likely sensitizes cells toward PPAR{gamma}-specific agonists supplied exogenously, e.g., by neighboring cells, which concomitantly provoked PPAR{gamma}-dependent apoptosis. This mechanism might be of physiological importance to guarantee down-regulation of T cell-immune responses during inflammation but may also be of pathophysiological significance in diseases known to be accompanied by lymphopenia such as sepsis. Understanding molecular signaling pathways leading to PPAR{gamma} expression as well as PPAR{gamma} conferred that apoptosis in activated T cells will give new insights for therapeutic interventions.


   ACKNOWLEDGEMENTS
 
We are grateful to Novartis Stiftung für Therapeutische Forschung for financial support.

Received October 11, 2002; revised January 17, 2003; accepted January 21, 2003.


   REFERENCES
TOP
 ABSTRACT
 INTRODUCTION
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 REFERENCES
 

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