The peroxisome proliferator-activated receptor γ (PPAR-γ) belongs to a receptor superfamily of ligand-activated transcription factors involved in the regulation of metabolism and inflammation. Oral administration of PPAR-γ agonists ameliorates the clinical course and histopathological features in experimental autoimmune encephalomyelitis, an animal model for multiple sclerosis (MS), and PPAR-γ agonist treatment of PBMCs from MS patients suppresses PHA-induced cell proliferation and cytokine secretion. These effects are pronounced when cells are preincubated with the PPAR-γ agonists and reexposed at the time of stimulation, indicating a sensitizing effect. To characterize the mechanisms underlying this sensitizing effect, we analyzed PPAR-γ expression in PMBCs of MS patients and healthy controls. Surprisingly, MS patients exhibited decreased PPAR-γ levels compared with controls. PHA stimulation of PBMCs from healthy controls resulted in a significant loss of PPAR-γ, which was prevented by in vitro preincubation of the cells or in vivo by long-term oral medication with the PPAR-γ agonist pioglitazone. Differences in PPAR-γ expression were accompanied by changes in PPAR-γ DNA-binding activity, as preincubation with pioglitazone increased DNA binding of PPAR-γ. Additionally, preincubation decreased NF-κB DNA-binding activity to control levels, whereas the inhibitory protein IκBα was increased. In MS patients, pioglitazone-induced increase in PPAR-γ DNA-binding activity and decrease in NF-κB DNA-binding activity was only observed in the absence of an acute MS relapse. These results suggest that the sensitizing effect observed in the preincubation experiments is mediated by prevention of inflammation-induced suppression of PPAR-γ expression with consecutive increase in PPAR-γ DNA-binding activity.

Multiple sclerosis (MS)2 is an inflammatory demyelinating autoimmune disease of the CNS that affects more than one million people worldwide (1). The peripheral activation of autoreactive CNS-Ag-specific CD4+ T cells seems to be the first step in the disease process, followed by migration of activated lymphocytes across the blood-brain barrier, infiltration of the brain parenchyma and local secretion of proinflammatory cytokines and chemokines leading to subsequent destruction of the myelin sheath and axonal loss (1, 2). Currently used therapeutic agents such as recombinant IFN-β and glatiramer acetate seem to influence this pathogenic process at several stages, such as inhibition of T cell activation (3), reduction of transmigration (4), as well as interference with the secretion of a number of proinflammatory cytokines (5). However, the beneficial effects of these substances are limited, prompting the search for additional therapeutic options (3). Several studies have recently demonstrated that agonists of the peroxisome proliferator-activated receptor γ (PPAR-γ) reduce the clinical and histopathological features of experimental allergic encephalomyelitis (EAE), an animal model of MS induced by repetitive immunization of susceptible animals with myelin derived-peptides (6, 7, 8). Moreover, PPAR-γ-deficient heterozygous mice develop an exacerbated EAE with prolonged clinical symptoms accompanied by an increased expansion of CD4+ and CD8+ T cells and pronounced Th1 response, demonstrating a critical role of PPAR-γ in the regulation of CNS inflammation and demyelination (9).

PPAR-γ is a member of the nuclear hormone receptor superfamily of ligand-activated transcription factors that are related to retinoid and steroid hormone receptors (10). It is expressed in different cell types of the immune system, e.g., macrophages (11), microglia (12), B-lymphocytes (13) and T-lymphocytes (14), as well as in glial and neuronal cells (15). Upon ligand binding, it heterodimerizes with the retinoid X receptor and binds to PPAR-response elements (PPRE) located in the promoter region of target genes, thus influencing gene transcription. Several studies have documented a negative interaction between PPAR-γ and proinflammatory transcription factors like NF-κB, NFAT, AP-1, and STAT both via direct interaction and by sequestration of essential cofactors (16, 17). PPAR-γ agonists include naturally occurring substances such as the prostaglandin D2 metabolite 15-deoxy-Δ (12, 14) prostaglandin J2 (PGJ2) (18), as well as several synthetic ligands such as the antidiabetic thiazolidinediones, e.g., pioglitazone (PIO) and ciglitazone, and several nonsteroidal anti-inflammatory drugs (19). Over the past decade, a number of antiproliferative and anti-inflammatory properties of PPAR-γ agonists have been characterized. In monocytes and macrophages, PPAR-γ activation inhibits the expression of a number of proinflammatory mediators (17, 20) and attenuates the oxidative burst in macrophages (21). In T-lymphocytes, PPAR-γ activation inhibits both Ag-specific and nonspecific T cell activation, T cell proliferation, and the production of several proinflammatory cytokines (14, 22, 23, 24). Interestingly, the anti-inflammatory and antiproliferative effects of PPAR-γ agonists are increased when cells are preincubated with PPAR-γ agonists and reexposed at the time of proinflammatory stimulation, indicating a sensitizing effect of these substances (25). The present study further examines the mechanisms underlying this sensitizing effect.

RPMI 1640, PBS, penicillin, streptomycin, and Trizol were purchased from Invitrogen Life Technologies. FCS was obtained from PAN Biotech and PHA from Sigma-Aldrich. PIO was purchased from Alexis Biochemicals.

Ten MS patients (6 females and 4 males) fulfilling established diagnostic criteria (26) and 10 healthy donors (HDs) (6 females and 4 males) were included in the study. Six MS patients had relapsing-remitting disease and four had secondary progressive disease. Four of the patients with relapsing-remitting disease received treatment with IFN-1β, the remaining two patients received treatment with glatiramer acetate. Patients with secondary progressive disease did not receive any treatment. The mean age of MS patients was 37.9 years and 35.2 years for HDs. The mean expanded disability status scale was 3.2 ± 1.7, and the mean duration of disease was 8.5 ± 5.4 years. Informed consent was obtained from all patients and HDs.

PBMCs derived from four patients suffering from type II diabetes that were on long-term oral medication with PIO (Actos) at a dose of 15 mg/day for at least 6 months were used as additional controls after giving their written consent. At the time of PBMC preparation, these patients did not suffer from any infectious or neoplastic disorder and did not take any immunosuppressive medication. The mean age of these patients was 47.6 years.

PBMCs were isolated by Ficoll standard density gradient centrifugation (Nycomed). Jurkat cells were purchased from Deutsche Sammlung von Mikroorganismen und Zellkulturen and grown in RPMI 1640 containing 100 U/ml penicillin G, 100 μg/ml streptomycin, and 10% FCS at 1 × 105 cells per well. PBMCs were cultured in the presence of PHA (5 μg/ml) in 200 μl of RPMI 1640 medium supplemented with 2 mM glutamine, 100 U/ml penicillin G, 100 μg/ml streptomycin, and 5% autologous serum in 96-well round-bottom microtiter plates (Nunc) at 1 × 105 cells per well.

T cell proliferative responses were determined using a nonradioactive proliferation assay measuring the incorporation of BrdU as described previously (25). The sensitivity of BrdU incorporation is comparable to standard radioactive [3H]thymidine incorporation assays. BrdU was added to the cultures 24 h before harvesting the cells for ELISA. BrdU assays were purchased from Boehringer Mannheim. Absorbance was measured at 450 nm on an Optimax ELISA reader (Molecular Devices). All proliferation assays were conducted in triplicates.

Cell culture supernatants for determination of IFN-γ were collected for all experimental conditions outlined above. Supernatants were centrifuged at 5000 rpm for 5 min, and stored at 20°C until assayed. Cytokine measurements were performed using a commercially available ELISA (R&D Systems) following the manufacturer′s instructions. The detection limit was 8 pg/ml. All measurements were performed in duplicate, and mean values of the two measurements were used for statistical analysis. Absorbances were measured at 450 nm on an Optimax ELISA reader (Molecular Devices).

Supernatants were collected and frozen at −20°C until analysis. Using the cytometric bead array “Inflammation” (BD Biosciences), the levels of IL-6 and TNF-α were measured simultaneously using a FACSCalibur flow cytometer (BD Biosciences) according to the manufacturer’s instructions. Subsequent analysis of the data was performed using the BD CBA software.

For Western blotting, cells were lysed in RIPA buffer (150 mM NaCl, 10 mM Tris-HCl (pH 8.0), 1% Nonidet P-40, 0.5% deoxycholic acid, 0.1% SDS, 5 mM EDTA) containing 0.7% PMSF, 0.2% aprotinin, and 0.2% leupeptin. Protein aliquots (120 μg) for detection of PPAR-γ and 60 μg protein aliquots for detection of IκBα were mixed with an equal amount of 4× SDS sample buffer, boiled at 98°C for 5 min, centrifuged, and separated by 10% SDS-PAGE. The gels were then transferred onto polyvinylidene fluoride membranes (Millipore). Membranes were blocked with 5% low-fat milk powder in PBS and washed three times with PBS/Tween 20 (PBST). Primary anti-PPAR-γ-Ab 210–118 (Alexis Biochemicals) was added at a 1/750 dilution in TBST containing 3% BSA and 0.01% sodium azide overnight at 4°C. Membranes were washed with TBST/Triton 1%, and HRP-conjugated goat anti-rabbit IgG (Sigma-Aldrich) was added at a dilution of 1/2500 in TBST for 1 h at room temperature. Primary anti-IκBα Ab (sc-203; Santa Cruz Biochemicals) was added at a 1/500 dilution in TBST containing 3% BSA and 0.01% sodium azide overnight at 4°C. Membranes were washed with TBST/Triton 1%, and HRP-conjugated goat anti-rabbit IgG (Sigma-Aldrich) was added at a dilution of 1/3000 in TBST for 1 h at room temperature. Bands were visualized by ECL reagent.

PPAR-γ protein was detected at ∼45 kDa. The specificity of the Ab used has been confirmed by Western blotting of lysates of N2A cells transiently transfected with PPAR-γ cDNA and of embryonic mouse PPAR-γ knockout and wild type fibroblasts (15).

For control of equal protein loading, membranes were afterward incubated with stripping solution (62.5 mM Tris-HCl (pH 6.8); 2% SDS, 100 mM 2-ME) for 30 min at 60°C, and after extensive washing with water and PBS again incubated with blocking solution for 1 h. Membranes were then incubated overnight with primary Ab to β-actin (1/5000; diluted in PBS containing 0.05% Tween 20 and 5% skim milk), again washed extensively with PBST, and incubated for 2 h with blocking solution containing HRP-conjugated rabbit-anti-mouse IgG (1/10,000).

After treatment, cells were washed with ice-cold PBS and centrifuged at 2000 × g for 5 min. Cells were washed in 1 ml of buffer A (10 mM HEPES (pH 7.9), 1.5 mM MgCl2, 10 mM KCl, 0.1% Nonidet P-40, and 0.5 mM DTT) and pelleted at 2000 × g for 5 min. The supernatant was discarded, and pellets were resuspended in 80 μl of buffer A containing 0.1% Triton X-100. After incubation at 4°C for 10 min, the homogenate was centrifuged at 2000 × g for 5 min, the nuclear pellet was resuspended in 60 μl of 20 mM HEPES (pH 7.9), 0.42 M NaCl, 25% (v/v) glycerol, 1.5 mM MgCl2, and 0.2 mM EDTA, incubated on ice for 30 min and centrifuged at 15,000 × g for 20 min at 4°C. Extracts were stored at −80°C until use.

For determination of PPAR-γ DNA-binding, a double-stranded oligonucleotide containing the PPRE from the acyl-CoA oxidase gene was used. The sequence containing the consensus PPRE site is as follows: 5′-GGT AAA GGT CAA AGG TCA ATC GGC-3′, and 3′-CCA TTT CCA GTT TCC AGT TAG CCG-5′.

For detection of NF-κB DNA binding, the following sequence was used: 5′-AGT TGA GGG GAC TTT CCC AGG C-3′, and 3′-TCA ACT CCC CTG AAA GGG TCC G-5′.

Complementary oligonucleotides were annealed in 10 mM Tris (pH 8.0), by heating to 65°C for 1 min and slow cooling at room temperature. Double-stranded oligonucleotides (10 pmol) are 5′-end labeled with [γ-32P]ATP (5000 Ci/mmol; Amersham Biosciences) and T4 polynucleotide kinase (New England Biolabs), purified using a G25 quick spin column (Roche Molecular Biochemicals) and diluted to a final volume of 150 μl.

EMSA experiments were performed using the gel shift assay system from Promega. Nuclear extracts (15 μg per reaction) were preincubated in 1 × binding buffer (4% glycerol, 1 mM MgCl2, 0.5 mM EDTA, 0.5 mM DTT, 50 mM NaCl, 10 mM Tris-HCl (pH 7.5), 0.5 mg/ml poly(dI-dC) · poly(dI-dC)) for 1 h on ice. For supershift and competition assays, 2 μl of Ab (polyclonal anti-p50-Ab, sc-8414X, and anti-p65-Ab, sc-372X, both obtained from Santa Cruz Biochemicals, for NF-κB-EMSA; polyclonal anti-PPAR-γ-Ab 210-118 from Alexis for PPAR-γ-EMSA) or an excess of unlabeled double-stranded oligonucleotides were added. After addition of 1 μl of labeled probe to each reaction and a 20-min incubation on ice, DNA-protein complexes were separated on 8% native polyacrylamide gels at 220 V with 0.5 × TBE as running buffer. Gels were vacuum-dried and visualized by autoradiography.

Jurkat cells were transfected with 10 μg of plasmid DNA by electroporation using a Bio-Rad Gene Pulser II with a capacitance extender plus (Bio-Rad). The conditions used were 250 V and 975 μF. The PPAR-γ1 reporter plasmid was provided by Dr. Johan Auwerx, Strasbourg, France. Twenty minutes after transfection, cells were either pretreated for 48 h with 5 μM PIO or left untreated. After 48 h, transfected but untreated cells were either stimulated with PHA alone or cotreated with PHA and 10 μM PIO for 48 h; transfected cells pretreated with 5 μM PIO were then stimulated with PHA and cotreated with 5 μM PIO for another 48 h. Afterward, cells were harvested in 250 μl of 1× reporter lysis buffer (Promega) for luciferase activity. The luciferase reporter assay was performed according to the manufacturer’s protocol, and the relative light intensity was measured with a luminometer. The experiment was performed twice with triplicates for each group.

For Western blots and EMSAs, bands were scanned and intensities were measured using NIH Image program. Graphs show mean ± SEM. Differences between controls and the different groups were assessed by Student’s t test (two tailed, unpaired); a value of p < 0.05 was considered to be significant.

We first assessed the antiproliferative effects of PIO in a concentration not exhibiting significant toxic effects assessed by MTT assay (data not shown). PBMCs from 10 MS patients and 10 healthy controls were either stimulated with PHA alone (5 μg/ml) for 48 h, coincubated with PHA and 10 μM PIO for 48 h, or pretreated with 5 μM PIO for 48 h followed by coadministration of PHA (5 μg/ml) together with 5 μM PIO for another 48 h. Untreated PBMCs served as control (Fig. 1). In healthy controls, coincubation lead to a 29% decrease of cell proliferation compared with PHA stimulation alone, whereas preincubation with PIO lead to a 73% decrease in cell proliferation compared with PHA stimulation alone. In MS patients, these antiproliferative effects of PIO were less pronounced, i.e., 21% decrease of proliferation by coincubation and 64% decrease of proliferation by preincubation, albeit this difference did not reach statistical significance when compared with healthy controls.

FIGURE 1.

Antiproliferative effects of the PPAR-γ agonist PIO on PHA-induced proliferation of PBMCs from MS patients and HDs. PBMCs from 10 MS patients (white bars) and 10 HDs (black bars) were stimulated with PHA alone (5 μg/ml) for 48 h, coincubated with PHA and 10 μM PIO (PHA+PIO) for 48 h, or first pretreated with 5 μM PIO for 48 h followed by coadministration of PHA (5 μg/ml) together with 5 μM PIO for another 48 h (PHA+PIO pre). Untreated PBMCs served as control. Proliferation was assessed by BrdU incorporation and expressed as percentage of the maximal proliferative response. Maximal proliferative response of PBMCs from MS patients and healthy controls was equal. ∗, Proliferative response compared with PHA stimulation (∗∗∗, p < 0.001); †, proliferative response in preincubated vs coincubated PBMCs (†††, p < 0.001). †, p < 0.05); §, proliferative response in MS patients compared with healthy controls (§§, p < 0.01).

FIGURE 1.

Antiproliferative effects of the PPAR-γ agonist PIO on PHA-induced proliferation of PBMCs from MS patients and HDs. PBMCs from 10 MS patients (white bars) and 10 HDs (black bars) were stimulated with PHA alone (5 μg/ml) for 48 h, coincubated with PHA and 10 μM PIO (PHA+PIO) for 48 h, or first pretreated with 5 μM PIO for 48 h followed by coadministration of PHA (5 μg/ml) together with 5 μM PIO for another 48 h (PHA+PIO pre). Untreated PBMCs served as control. Proliferation was assessed by BrdU incorporation and expressed as percentage of the maximal proliferative response. Maximal proliferative response of PBMCs from MS patients and healthy controls was equal. ∗, Proliferative response compared with PHA stimulation (∗∗∗, p < 0.001); †, proliferative response in preincubated vs coincubated PBMCs (†††, p < 0.001). †, p < 0.05); §, proliferative response in MS patients compared with healthy controls (§§, p < 0.01).

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Additionally, analysis of cytokine production by PBMCs from an identical experimental setup using PBMCs from the same donors as shown in Fig. 1 revealed a decrease in the PHA-induced secretion of TNF-α, IL-6, and IFN-γ by PIO treatment both in healthy controls and in MS patients (Fig. 2). Again, preincubation with PIO significantly augmented this decrease in cytokine secretion in MS patients and in controls. However, in MS patients, the effect of PIO cotreatment on cytokine secretion was significantly diminished when compared with healthy controls, whereas pretreatment of cells from MS patients with PIO was nearly as effective in reducing cytokine secretion as in cells from healthy controls (Fig. 2).

FIGURE 2.

Anti-inflammatory effects of PIO on PHA-stimulated PBMCs from MS patients and healthy donors. PBMCs from 10 MS patients and 10 healthy donors were stimulated with PHA alone (5 μg/ml) for 48 h, coincubated with PHA and 10 μM PIO (PHA+PIO) for 48 h, or first pretreated with 5 μM PIO for 48 h followed by coadministration of PHA (5 μg/ml) together with 5 μM PIO for another 48 h (PHA+PIO pre). Untreated PBMCs served as control. Supernatants were collected 48 h after PHA stimulation. Secretion of IFN-γ was measured by ELISA, secretion of TNF-α and IL-6 was assessed by immunobead assay. Cytokine secretion is expressed as percentage of the maximal response, maximal response in MS patients, and in healthy controls was equal. ∗, Cytokine secretion compared with PHA stimulation (∗∗, p < 0.01; ∗∗∗, p < 0.001); †, cytokine secretion by preincubated vs coincubated PBMCs (†, p < 0.05; ††, p < 0.01; †††, p < 0.001); §, cytokine secretion in MS patients compared with healthy controls (§, p < 0.05; §§, p < 0.01).

FIGURE 2.

Anti-inflammatory effects of PIO on PHA-stimulated PBMCs from MS patients and healthy donors. PBMCs from 10 MS patients and 10 healthy donors were stimulated with PHA alone (5 μg/ml) for 48 h, coincubated with PHA and 10 μM PIO (PHA+PIO) for 48 h, or first pretreated with 5 μM PIO for 48 h followed by coadministration of PHA (5 μg/ml) together with 5 μM PIO for another 48 h (PHA+PIO pre). Untreated PBMCs served as control. Supernatants were collected 48 h after PHA stimulation. Secretion of IFN-γ was measured by ELISA, secretion of TNF-α and IL-6 was assessed by immunobead assay. Cytokine secretion is expressed as percentage of the maximal response, maximal response in MS patients, and in healthy controls was equal. ∗, Cytokine secretion compared with PHA stimulation (∗∗, p < 0.01; ∗∗∗, p < 0.001); †, cytokine secretion by preincubated vs coincubated PBMCs (†, p < 0.05; ††, p < 0.01; †††, p < 0.001); §, cytokine secretion in MS patients compared with healthy controls (§, p < 0.05; §§, p < 0.01).

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Next, we investigated PPAR-γ expression levels in untreated PBMCs from 10 MS patients and 10 healthy donors (Fig. 3,a). All MS patients exhibited a profound reduction in PPAR-γ expression levels of ∼65%, independent from disease course or current therapy. Treatment of PBMCs from eight healthy donors with PHA for 48 h lead to a distinct decrease in PPAR-γ expression (Fig. 3,b) that was prevented by preincubation but not by coincubation with PIO. In strong contrast, PBMCs derived from four diabetic patients on long-term oral treatment with PIO did not reveal this PHA-induced loss of PPAR-γ. Additional in vitro treatment of the cells with PIO did not change PPAR-γ expression levels further (Fig. 3,c). To estimate whether the changes in PPAR-γ levels by PHA and by PIO are caused by changes in PPAR-γ gene transcription, transformed human T cells (Jurkat cells) were transfected with a reporter gene construct containing the human PPAR-γ1 promoter and afterward treated with PHA and PIO (Fig. 3 d). These cells have proven to be comparably sensitive to the anti-inflammatory and antiproliferative effects of PPAR-γ agonists as PBMCs (25). PHA stimulation alone led to a 50% decrease in PPAR-γ1 promoter activity compared with baseline activity. Cotreatment of PHA-stimulated cells with PIO prevented the PHA-induced decrease in PPAR-γ1 promoter activity but did not result in an increase in PPAR-γ1 promoter activity known from PIO treatment of Jurkat cells under noninflammatory conditions (data not shown). Preincubation with PIO lead to a significant increase in PPAR-γ1 promoter activity comparable to the one observed after incubation with PIO alone.

FIGURE 3.

PPAR-γ expression in PBMCs from MS patients, healthy controls, and diabetic patients under long-term oral treatment with PIO. Analysis of PPAR-γ protein levels in untreated PBMCs from 10 MS patients and 10 healthy controls by Western blotting (a). Determination of PPAR-γ expression in PBMCs from eight healthy donors after PHA stimulation alone (5 μg/ml) for 48 h, after coincubation with 10 μM PIO (PHA+PIO) for 48 h, and after pretreatment of cells with 5 μM PIO for 48 h followed by coadministration of PHA (5 μg/ml) together with 5 μM PIO for another 48 h (PHA+PIO pre). Untreated PBMCs served as control (b). PPAR-γ expression in PBMCs from four diabetic patients receiving long-term oral PIO treatment after PHA stimulation alone (5 μg/ml) for 48 h, after coincubation with 10 μM PIO for 48 h (PHA+PIO), and without in vitro treatment of cells (c). β-actin Western blot served as control for all Western blots. The intensity of each band was measured by densitometry; graphs demonstrate mean and SEM of each condition. Jurkat cells transfected with the PPAR-γ1 promoter were stimulated with PHA (5 μg/ml) for 48 h, cotreated with PHA and 10 μM PIO for 48 h (PHA+PIO), or pretreated with 5 μM PIO for 48 h followed by coadministration of PHA together with 5 μM PIO for another 48 h (PHA+PIO pre). The graph shows the relative luciferase intensity compared with untreated, but transfected cells (d). Asterisks indicate significant differences between controls and respective groups (∗, p < 0,05; ∗∗, p < 0,01); †, significant differences between pretreated and non-pretreated cells (†, p < 0.05).

FIGURE 3.

PPAR-γ expression in PBMCs from MS patients, healthy controls, and diabetic patients under long-term oral treatment with PIO. Analysis of PPAR-γ protein levels in untreated PBMCs from 10 MS patients and 10 healthy controls by Western blotting (a). Determination of PPAR-γ expression in PBMCs from eight healthy donors after PHA stimulation alone (5 μg/ml) for 48 h, after coincubation with 10 μM PIO (PHA+PIO) for 48 h, and after pretreatment of cells with 5 μM PIO for 48 h followed by coadministration of PHA (5 μg/ml) together with 5 μM PIO for another 48 h (PHA+PIO pre). Untreated PBMCs served as control (b). PPAR-γ expression in PBMCs from four diabetic patients receiving long-term oral PIO treatment after PHA stimulation alone (5 μg/ml) for 48 h, after coincubation with 10 μM PIO for 48 h (PHA+PIO), and without in vitro treatment of cells (c). β-actin Western blot served as control for all Western blots. The intensity of each band was measured by densitometry; graphs demonstrate mean and SEM of each condition. Jurkat cells transfected with the PPAR-γ1 promoter were stimulated with PHA (5 μg/ml) for 48 h, cotreated with PHA and 10 μM PIO for 48 h (PHA+PIO), or pretreated with 5 μM PIO for 48 h followed by coadministration of PHA together with 5 μM PIO for another 48 h (PHA+PIO pre). The graph shows the relative luciferase intensity compared with untreated, but transfected cells (d). Asterisks indicate significant differences between controls and respective groups (∗, p < 0,05; ∗∗, p < 0,01); †, significant differences between pretreated and non-pretreated cells (†, p < 0.05).

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We next investigated whether the observed changes in PPAR-γ expression levels are accompanied by changes of PPAR-γ DNA-binding at the consensus sequence of the human PPRE in PBMCs from six healthy donors. PHA stimulation alone did not result in changes of DNA-binding activity, but did prevent the expected increase in DNA-binding activity after cotreatment with PIO (Fig. 4). In contrast, preincubation of PBMCs with PIO resulted in a profound increase in DNA-binding activity of PPAR-γ comparable to the one observed after PIO treatment under noninflammatory conditions (data not shown).

FIGURE 4.

Changes in PPAR-γ DNA-binding activity by PIO under inflammatory conditions in PBMCs from healthy controls. EMSA determination of PPAR-γ DNA-binding activity in PBMCs from six healthy donors without treatment, after PHA stimulation alone (5 μg/ml) for 48 h, after coincubation with 10 μM PIO (PHA+PIO) for 48 h, and after pretreatment of cells with 5 μM PIO for 48 h followed by coadministration of PHA (5 μg/ml) together with 5 μM PIO for another 48 h (PHA+PIO pre). Human embryonic kidney (HEK) cells transfected with or without PPAR-γ served as control. Preincubation of nuclear extracts from PPAR-γ-transfected HEK cells with an anti-PPAR-γ-Ab led to a profound decrease in PPAR-γ-DNA-binding activity, demonstrating the reaction specificity. The intensity of each band was measured by densitometry; graphs demonstrate mean and SEM. ∗, Significant differences between control and different treatments (∗∗, p < 0.01); †, significant differences between PHA stimulation alone and pretreatment with PIO (†, p < 0.05).

FIGURE 4.

Changes in PPAR-γ DNA-binding activity by PIO under inflammatory conditions in PBMCs from healthy controls. EMSA determination of PPAR-γ DNA-binding activity in PBMCs from six healthy donors without treatment, after PHA stimulation alone (5 μg/ml) for 48 h, after coincubation with 10 μM PIO (PHA+PIO) for 48 h, and after pretreatment of cells with 5 μM PIO for 48 h followed by coadministration of PHA (5 μg/ml) together with 5 μM PIO for another 48 h (PHA+PIO pre). Human embryonic kidney (HEK) cells transfected with or without PPAR-γ served as control. Preincubation of nuclear extracts from PPAR-γ-transfected HEK cells with an anti-PPAR-γ-Ab led to a profound decrease in PPAR-γ-DNA-binding activity, demonstrating the reaction specificity. The intensity of each band was measured by densitometry; graphs demonstrate mean and SEM. ∗, Significant differences between control and different treatments (∗∗, p < 0.01); †, significant differences between PHA stimulation alone and pretreatment with PIO (†, p < 0.05).

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Another important mechanism of PPAR-γ-mediated anti-inflammatory action is the negative interaction with NF-κB, one of the central transcription factors involved in the expression of proinflammatory genes. Therefore, we investigated the influence of PIO treatment on the DNA-binding activity of NF-κB. As depicted in Fig. 5,a, cotreatment with PIO did only marginally decrease NF-κB DNA-binding activity induced by PHA stimulation, whereas pretreatment reduced NF-κB DNA-binding activity back to control levels. As the activity of NF-κB can be influenced by changes in the expression levels of its cytoplasmic inhibitors, especially by IκBα, we next investigated whether PIO influences IκBα expression levels under inflammatory conditions. Cotreatment of PBMCs with PHA and PIO did not influence the expression levels of IκBα when compared with PHA stimulation alone, whereas pretreatment with PIO substantially increased IκBα expression (Fig. 5 b). Exposure to PHA alone mildly enhanced IκBα levels at 48 h. This is in accordance with previous reports showing biphasic expression pattern of IκBα in response to inflammatory stimuli (27).

FIGURE 5.

Changes in NF-κB-DNA-binding activity and IκBα expression levels by PIO treatment under inflammatory conditions in PBMCs from healthy controls. Determination of NF-κB-DNA-binding activity by EMSA in PBMCs from six healthy donors after PHA stimulation alone (5 μg/ml) for 48 h, after coincubation with 10 μM PIO for 48 h, and after pretreatment of cells with 5 μM PIO for 48 h followed by coadministration of PHA (5 μg/ml) together with 5 μM PIO for another 48 h. Untreated PBMCs served as control. Supershift experiments with Abs directed against p50 and p65 and cold competition with excessive unlabeled oligonucleotide demonstrate the specificity of the reaction (a). Western blot analysis of IκBα expression in PBMCs from six healthy donors without treatment, after PHA stimulation alone (5 μg/ml) for 48 h, after coincubation with 10 μM PIO for 48 h, and after pretreatment of cells with 5 μM PIO for 48 h followed by coadministration of PHA (5 μg/ml) together with 5 μM PIO for another 48 h (b). β-actin Western blot served as control. The intensity of each band was measured by densitometry; graphs demonstrate mean and SEM. ∗, Significant differences between control and different treatments (∗, p < 0.05; ∗∗, p < 0.01), †, significant differences between PHA stimulation alone and pretreatment with PIO (†, p < 0.05).

FIGURE 5.

Changes in NF-κB-DNA-binding activity and IκBα expression levels by PIO treatment under inflammatory conditions in PBMCs from healthy controls. Determination of NF-κB-DNA-binding activity by EMSA in PBMCs from six healthy donors after PHA stimulation alone (5 μg/ml) for 48 h, after coincubation with 10 μM PIO for 48 h, and after pretreatment of cells with 5 μM PIO for 48 h followed by coadministration of PHA (5 μg/ml) together with 5 μM PIO for another 48 h. Untreated PBMCs served as control. Supershift experiments with Abs directed against p50 and p65 and cold competition with excessive unlabeled oligonucleotide demonstrate the specificity of the reaction (a). Western blot analysis of IκBα expression in PBMCs from six healthy donors without treatment, after PHA stimulation alone (5 μg/ml) for 48 h, after coincubation with 10 μM PIO for 48 h, and after pretreatment of cells with 5 μM PIO for 48 h followed by coadministration of PHA (5 μg/ml) together with 5 μM PIO for another 48 h (b). β-actin Western blot served as control. The intensity of each band was measured by densitometry; graphs demonstrate mean and SEM. ∗, Significant differences between control and different treatments (∗, p < 0.05; ∗∗, p < 0.01), †, significant differences between PHA stimulation alone and pretreatment with PIO (†, p < 0.05).

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PBMCs taken from one MS patient in the absence of a clinically detectable MS relapse (patient A) exhibit an increase in PPAR-γ DNA-binding activity after pretreatment with PIO 48 h before PHA stimulation, accompanied by a decrease in PHA-induced NF-κB DNA-binding activity. (Fig. 6). These changes resemble those observed in healthy controls. However, in a patient suffering from an acute MS relapse at the time of the investigation (patient B), which means that the cells have already been activated in vivo, pretreatment with PIO in vitro before PHA stimulation did not result in an increase in PPAR-γ DNA-binding activity and lead only to a slight decrease in NF-κB DNA-binding activity (Fig. 6).

FIGURE 6.

Changes in PPAR-γ DNA-binding activity and NF-κB-DNA-binding activity by PIO treatment under inflammatory conditions in PBMCs from MS patients. Determination of PPAR-γ DNA-binding activity and NF-κB-DNA-binding activity by EMSA in PBMCs from one MS patient without acute inflammation (patient A) and one patient with an acute MS relapse at the time of the investigation (patient B). PBMCs were either stimulated with PHA alone (5 μg/ml) for 48 h or pretreated with 5 μM PIO for 48 h followed by coadministration of PHA (5 μg/ml) together with 5 μM PIO for another 48 h. Untreated PBMCs served as control. For PPAR-γ, preincubation of nuclear extracts from PPAR-γ-transfected HEK cells with an anti-PPAR-γ-Ab leading to a profound decrease in PPAR-γ-DNA-binding activity, for NF-κB, supershift of lysates from PHA-treated PBMCs (patient A) with Abs against p50 and p65 demonstrate the specificity of the reaction.

FIGURE 6.

Changes in PPAR-γ DNA-binding activity and NF-κB-DNA-binding activity by PIO treatment under inflammatory conditions in PBMCs from MS patients. Determination of PPAR-γ DNA-binding activity and NF-κB-DNA-binding activity by EMSA in PBMCs from one MS patient without acute inflammation (patient A) and one patient with an acute MS relapse at the time of the investigation (patient B). PBMCs were either stimulated with PHA alone (5 μg/ml) for 48 h or pretreated with 5 μM PIO for 48 h followed by coadministration of PHA (5 μg/ml) together with 5 μM PIO for another 48 h. Untreated PBMCs served as control. For PPAR-γ, preincubation of nuclear extracts from PPAR-γ-transfected HEK cells with an anti-PPAR-γ-Ab leading to a profound decrease in PPAR-γ-DNA-binding activity, for NF-κB, supershift of lysates from PHA-treated PBMCs (patient A) with Abs against p50 and p65 demonstrate the specificity of the reaction.

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It is well known that proinflammatory cytokines play a major role in the pathogenesis of MS as well as in EAE, an established animal model of MS. Several cytokines like TNF-α, IFN-γ, and IL-6 are regularly found in MS brain lesions and in spinal cord infiltrates of EAE mice (6, 28, 29). Moreover, cytokine expression in PBMCs from MS patients correlates well with disease activity and precedes the onset of clinical symptoms by 4 weeks, suggesting an essential role in the development of MS relapses (30). We show that PIO significantly decreases cell proliferation as well as secretion of several proinflammatory cytokines in PHA-stimulated PBMCs from MS patients and healthy controls, i.e., TNF-α, IL-6, and IFN-γ, thus attenuating inflammatory molecules known to be involved in MS pathogenesis. These effects are much more pronounced, when cells are preincubated with PIO before inflammatory stimulation, indicating a sensitizing effect induced by pretreatment of the cells with the PPAR-γ agonist. However, in MS patients, these anti-inflammatory effects of PIO treatment were significantly reduced when compared with healthy controls. Because untreated PBMCs from MS patients exhibited a strong reduction in PPAR-γ expression and inflammatory stimulation of PBMCs from healthy controls equally resulted in loss of PPAR-γ, we propose that inflammatory conditions affect PPAR-γ expression. It has already been demonstrated that proinflammatory stimuli like TNF-α and IFN-γ suppress the expression of PPAR-γ in adipocytes and bone marrow stromal cells, whereas PPAR-γ agonists are able to increase the expression of their own receptor (31, 32, 33). However, these cell types predominantly express PPAR-γ2, whereas PBMCs express PPAR-γ1. In our experiments, coincubation with PIO was not able to prevent the inflammation-induced loss of PPAR-γ, whereas preincubation with the drug stabilized PPAR-γ levels. Additionally, long-term oral PIO treatment prevented the PHA-induced loss of PPAR-γ expression in PBMCs from diabetic patients, demonstrating that the concentrations of PIO achieved by a standard oral treatment in vivo are sufficient to protect from inflammation-induced loss of PPAR-γ. These changes in PPAR-γ expression are probably due to changes in PPAR-γ gene transcription, because reporter gene assays revealed a decrease in PPAR-γ1 promotor activity by inflammatory stimulation. Moreover, stimulation abrogated the agonist-induced increase in promoter activity upon cotreatment with PIO, whereas pretreatment with PIO lead to a pronounced increase in the activity of the PPAR-γ1 promoter even under inflammatory conditions. These results suggest that after inflammatory stimulation in vitro or in vivo in MS patients even in the absence of clinical symptoms, PPAR-γ1 promoter activity is suppressed resulting in decreased PPAR-γ expression levels and that this inflammation-induced decrease in PPAR-γ expression can be prevented either by preincubation with PIO in vitro or by oral treatment with PIO as demonstrated in PBMCs from diabetic patients.

As changes in PPAR-γ expression do not necessarily affect PPAR-γ activity, the influence of PHA and PIO on PPAR-γ binding activity to its response element was investigated in PBMCs from healthy controls. Preincubation with PIO markedly increased the DNA-binding activity of PPAR-γ, whereas coincubation of PIO and PHA did not alter PPAR-γ DNA-binding activity. These data show that inflammatory stimulation impairs PPAR-γ DNA-binding activity in parallel to PPAR-γ protein levels, and this effect can be prevented by preincubation with a receptor agonist. Interestingly, PHA stimulation alone did not result in a decrease in PPAR-γ DNA-binding activity. This is in contrast to the reduced PPAR-γ expression levels observed after inflammatory stimulation. A possible explanation for this discrepancy is that inflammatory conditions may induce compensatory mechanisms like an increase in PPAR-γ DNA-binding activity by endogenous activators. This hypothesis is strengthened by the observation that cotreatment with the receptor agonists was not able to increase PPAR-γ DNA-binding activity further, possibly due to already maximal activity of PPAR-γ DNA binding.

In addition to PPRE-mediated gene regulation, PPAR-γ negatively interferes with a number of transcription factors such as AP-1, NF-κB, and NFAT, all involved in proinflammatory gene expression (16, 17, 24). NF-κB, a key factor of inflammatory signaling, initiates the transcription of various cytokines relevant for MS pathogenesis, including TNF-α, IL-6, and IFN-γ, (34) and inflammatory active white matter lesions exhibit increased NF-κB expression in microglia and invading macrophages (35). Additionally, mice deficient in the p50 subunit of NF-κB are significantly resistant to MOG EAE that was accompanied by a deficiency of Ag-specific T cells to differentiate into effector T cells in vivo (36). Preincubation of PBMCs from healthy controls reduced NF-κB DNA-binding activity back to control levels, whereas coincubation with PHA and PIO did not affect the inflammation-induced increase of NF-κB DNA-binding activity. Because NF-κB activation is tightly regulated by proteolysis of inhibitory proteins (IκB) that sequester NF-κB in the cytoplasm, we investigated whether alterations of NF-κB DNA-binding activity are accompanied by changes of IκBα protein levels. Coincubation with PIO did not alter IκBα levels, but preincubation significantly increased IκBα. These data suggest that pretreatment with PIO leads to an increase in PPAR-γ DNA-binding activity with concomitant increase in the expression of IκBα and subsequent inhibition of NF-κB DNA-binding activity. This hypothesis is strengthened by the observation that heterozygous PPAR-γ mice develop spontaneous NF-κB activation, accompanied by an increase in IκBα phosphorylation with concomitant decrease in IκBα expression levels (37). However, alternative mechanisms such as direct interaction between PPAR-γ and NF-κB may also account for the observed decrease in NF-κB activation.

In MS patients, PIO-induced increase in PPAR-γ DNA-binding activity and concomitant decrease in NF-κB DNA-binding activity was only observed in the absence of an acute inflammatory event, because pretreatment of PBMCs with PIO in vitro failed to increase PPAR-γ activity in a patient with an acute MS relapse at the time of the investigation, whereas in a patient without apparent inflammation, PIO treatment was equally effective in enhancing PPAR-γ DNA-binding activity and lowering NF-κB DNA-binding activity as in healthy controls. This is in contrast to PPAR-γ expression levels, as untreated PBMCs from MS patients exhibited a strong reduction in PPAR-γ expression independently from disease activity. Thus, it seems conceivable that preincubation with PIO in the absence of acute inflammatory activity in vivo, i.e., in nonactivated PBMCs, is still able to increase PPAR-γ activity despite decreased PPAR-γ expression levels but not in PBMCs from patients with an acute MS relapse when cells have already been activated in vivo before PIO treatment in vitro. These results suggest that treatment with PIO can prevent inflammation-induced loss of PPAR-γ activity only when given before an acute inflammatory event. With regard to the therapeutic potential of PPAR-γ agonists for the treatment of MS, these observations underline the significance of an early onset of treatment before an acute relapse.

Taken together, our data demonstrate a protection from inflammation-induced loss of PPAR-γ expression, an increase in PPAR-γ DNA-binding activity, and hence an enhancement of PPAR-γ-mediated anti-inflammatory and antiproliferative effects by PIO pretreatment, thus representing a promising strategy to counterbalance inflammatory activity in MS patients by elevating the threshold for further inflammatory events.

The authors have no financial conflict of interest.

The costs of publication of this article were defrayed in part by the payment of page charges. This article must therefore be hereby marked advertisement in accordance with 18 U.S.C. Section 1734 solely to indicate this fact.

2

Abbreviations used in this paper: MS, multiple sclerosis; PPAR-γ, peroxisome proliferator-activated receptor γ; EAE, experimental allergic encephalomyelitis; PPRE, PPAR-response element; HD, healthy donor; PIO, pioglitazone.

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