PPARδ Is a Type 1 IFN Target Gene and Inhibits Apoptosis in T Cells

ORIGINAL ARTICLE

PPARd Is a Type 1 IFN Target Gene and Inhibits Apoptosis in T Cells Nadya al Yacoub1, Malgorzata Romanowska1, Sybille Krauss2, Susann Schweiger2,3,4 and John Foerster3,4 Peroxisome proliferator-activated receptor b/d (PPARd) is a nuclear hormone receptor regulating diverse biological processes, including b-oxidation of fatty acid and epithelial cell differentiation. To date, the role of PPARd in the immune system has not been thoroughly studied. Here, we show that PPARd is expressed in activated human T cells purified from peripheral blood as well as in T cells isolated from affected psoriasis skin lesions. PPARd is induced in T cells on stimulation with type 1 IFN. Functionally, PPARd enhances proliferation of primary T cells and blocks apoptosis induced by type 1 IFN and by serum deprivation. We show that these cellular functions are mediated by the activation of extracellular signal-regulated kinase1/2 signaling. Our results (1) establish a direct molecular link between type 1 IFN signaling and PPARd, (2) define a functional role for PPARd in human T cells, and (3) suggest that the induction of PPARd by type 1 IFN contributes to the persistence of activated T cells in psoriasis skin lesions. Journal of Investigative Dermatology (2008) 128, 1940–1949; doi:10.1038/jid.2008.32; published online 28 February 2008

INTRODUCTION The transcription factor peroxisome proliferator-activated receptor b/d (PPARd) regulates diverse biological processes, including adipogenesis, glucose metabolism, myogenesis, and macrophage function, as well as proliferative responses involved in colon carcinoma (reviewed in Maisch et al., 2005; Barish et al., 2006; Burdick et al., 2006). As such, it is expressed in various cell types, including adipocytes, endothelial cells, muscle cells, and colon. In the skin, PPARd is expressed in keratinocytes and contributes to the woundhealing response (Icre et al., 2006). In keratinocytes, PPARd is induced by IFN-g, stress-activated kinase, or tumor necrosis factor (TNF)-a by activator protein-1-mediated transcription (Tan et al., 2001; Di-Poi et al., 2003). Psoriasis is a very common chronic inflammatory skin disease, with an estimated prevalence of 2–4% in white populations, exhibiting a non-Mendelian pattern of inheritance (Elder, 2005; Rahman and Elder, 2005). Irrespective of the genetic susceptibility pattern in a given patient, clinical 1

Department of Biology, Free University of Berlin, Berlin, Germany; Department of Human Genetics, Max Planck Institute for Molecular Genetics, Berlin, Germany; 3Department of Pathology and Neuroscience, University of Dundee, Dundee, UK and 4Department of Dermatology, University of Dundee, Dundee, UK

2

Correspondence: Dr John Foerster, Department of Pathology and Neuroscience, University of Dundee, Ninewells Hospital, Level 6, Dundee DD1 9SY, UK. E-mail: [email protected] Abbreviations: ERK, extracellular signal-regulated kinase; GFP, green fluorescent protein; ILK, integrin-linked kinase; PBMC, peripheral blood mononuclear cell; PBS, phosphate-buffered saline; PDK1, pyruvate dehydrogenase kinase 1; RNAi, RNA interference; RT, room temperature; STAT, signal transducer and activator of transcription; TNF-a, tumor necrosis factor-a Received 10 July 2007; revised 17 December 2007; accepted 19 December 2007; published online 28 February 2008

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disease flares are initiated by triggers such as stress and inflammation, culminating in a common yet complex set of aberrations involving disturbed keratinocyte proliferation and differentiation (Gandarillas et al., 1999; Hobbs et al., 2004), activation of type I IFN signaling, endothelial cell activation, neutrophil influx, as well as persistence of activated T cells (Boyman et al., 2004; Kohlmann et al., 2004) and dendritic cells (Lowes et al., 2005) within the skin. Several lines of evidence support a role for PPARd in psoriasis pathogenesis: it is upregulated in lesional psoriatic skin (Westergaard et al., 2003); is induced by TNF-a, a key cytokine in psoriasis (Tan et al., 2001; Di-Poi et al., 2003); regulates keratinocyte differentiation; blocks apoptosis (see below) (Icre et al., 2006); and induces angiogenesis (Piqueras et al., 2006). Recently, we showed that the expression pattern of a large group of transcripts dysregulated in psoriasis is consistent with the activation of PPARd (Romanowska et al., 2007). We also showed that activation of PPARd enhances proliferation of epidermal keratinocytes and that it induces transcription of heparin-binding epidermal growth factor-like growth factor, which in turn induces epidermal hyperplasia in vivo (Zheng et al., 2003; Kimura et al., 2005; Shirakata et al., 2005), suggesting that activation of PPARd contributes to the hyperproliferative phenotype in psoriasis, at least in part, by inducing heparin-binding epidermal growth factor-like growth factor expression. On the basis of available evidence, the role of PPARd in apoptosis appears to be complex and cell type-dependent. Conflicting data exist in particular with respect to keratinocytes. Thus, Di-Poi et al. (2002) reported an inhibitory effect of PPARd on apoptosis by transcriptional upregulation of integrin-linked kinase (ILK) and pyruvate dehydrogenase kinase 1 (PDK1), causing increased phosphorylation of AKT in murine keratinocytes. The same group also reported & 2008 The Society for Investigative Dermatology

N al Yacoub et al. PPARd in T Cells

activated T cells, and other PPAR isoforms are expressed in T cells. As shown in Figure 1a, we detected expression of PPARd both at the mRNA and the protein levels in CD3 þ T cells isolated from peripheral blood, the Jurkat T cell line, as well as CD3 þ T cells isolated from active psoriasis plaques. In each case, PPARd protein was localized almost exclusively to the nucleus. As the expression level was highest in the T cells from psoriasis, which are activated, we next tested whether PPARd expression can be induced by TCR stimulation. To this end, we purified peripheral T cells from buffy coat of healthy probands by magnetic-activated cell sorting-mediated negative depletion of CD19, CD14, CD16, and CD56 lineages and exposed them to stimulatory anti-CD3 antibody either added to the medium, or precoated to the bottom of the culture dishes for 48 hours. We did not observe induction of PPARd in cells from three donors using this protocol (data not shown). Moreover, co-incubation with concanavalin A also only led to a marginal induction of PPARd (Figure 2a). Therefore, TCR stimulation does not cause induction of PPARd in peripheral human T cells.

AKT-dependent protection from ischemia- and H2O2induced apoptosis by PPARd in kidney, in vivo and in vitro (Letavernier et al., 2005). By contrast, none of these effects could be reproduced by other workers in keratinocytes (Burdick et al., 2007) or endothelial cells (Liou et al., 2006). Rather, the latter group identified transcriptional induction of 14-3-3e and, by implication, inhibited mitochondrial translocation of BAD as mechanisms underlying protection from H2O2-induced apoptosis by PPARd in endothelial cells. The inhibition of oxidative stress-induced apoptosis by PPARd blocking may also involve induction of catalase (Pesant et al., 2006). A protective effect by PPARd on celecoxib-induced apoptosis in colon carcinoma cells has also been shown, although, without delineation of the mechanism (Shureiqi et al., 2003). To add another layer of complexity, use of less specific agonists, such as prostacycline, has even led to the suggestion of proapoptotic activity by PPARd in 293T and endothelial cells, although the mechanism underlying this effect remain obscure. In light of diverse activities of PPAR isoforms in various inflammatory processes, their expression and functional role in T cells are not surprising. Thus, PPARg acts proapoptotically in T cells (Harris and Phipps, 2001, 2002; Cippitelli et al., 2003; Soller et al., 2006). PPARa, in turn, appears to counter proinflammatory signaling in T cells, including the activation of IL-2, c-jun, and NFkB (Jones et al., 2002, 2003; Dunn et al., 2007). Surprisingly, very little is known about the expression and function of PPARd in T cells. Here, we report that PPARd is expressed in primary human T cells as well as in T cells derived from active psoriasis and that PPARd is a direct target gene of type 1 IFN. PPARd blocks IFN-induced apoptosis in T cells. Our results establish a direct link between IFN and PPARd signaling and identify an important function of PPARd in a key cell type involved in psoriasis.

PPARd is a transcriptional target of type I IFN

We next used CD3 þ human T cells isolated from peripheral blood to screen for induction of PPARd expression by various agents. As shown in Figure 2a, robust nuclear accumulation of PPARd protein was induced, as expected, after stimulation with TNF-a, presumably acting through a previously characterized cascade involving activator protein-1 signaling (Tan et al., 2001). In contrast, IL-2, or concanavalin A, had no significant effect on PPARd expression. Two proapoptotic stimuli, anti-Fas and hydrogen peroxide, also strongly induced PPARd. As IFNs also regulate apoptosis, we stimulated T cells with IFN-a (type 1) and IFN-g (type 2). Only IFN-a, but not IFN-g, strongly induced PPAR expression in primary T cells. Given the pivotal role of type 1 IFN in psoriasis, and the strong upregulation of PPARd in this disease in vivo, we characterized the induction of PPARd by type 1 IFN in further detail. As shown in Figure 2b, induction of PPARd by type 1 IFN at the mRNA level could not be

RESULTS PPARd is expressed in activated T cells

We analyzed expression of PPARd in human T-cell populations from peripheral blood and from psoriasis skin lesions, as PPARd plays a role in psoriasis, a disease involving PCR :

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Ponceau S Figure 1. Expression of PPARd in human T cells. Detection of PPARd by reverse-transcription PCR (RT-PCR) (top) and western blot (bottom) in T cells expanded from psoriasis lesions, CD3 þ T cells isolated from peripheral blood, and Jurkat T cells. For analysis of protein expression, nuclear (n) and cytosolic (c) fractions were prepared as specified in Materials and Methods and subjected to SDS-PAGE and western blot using 20 mg per lane of cytosolic (c) or nuclear (n) fraction. A Ponceau S stain of the blots is shown at the bottom for control of protein loading.

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prevented by blocking de novo protein translation using cycloheximide, suggesting a direct transcriptional effect. The stimulatory effect observed on incubation with cycloheximide alone is presumably mediated through the induction of IFN-b (Maroteaux et al., 1983; Ringold et al., 1984). Direct transcriptional induction of PPARd by type 1 IFN was also supported by a time-course study, showing notable transcriptional induction at 6 hours and maximal induction at 12 hours after stimulation (Figure 2c). Interestingly, we observed pronounced differences in the magnitude of PPARd induction by IFN-a in T cells from various individuals. These differences were closely correlated with the level of PPARd expression before stimulation, such that cells harboring high endogenous PPARd levels showed less upregulation by IFN-a (Figure 2d; r ¼ 0.92). Taken together, PPARd is regulated in primary human T cells, as in other cell types, by TNF-a and apoptotic and stress signals, but in addition also by type 1 IFN. PPARd stimulates T-cell proliferation

We next sought to address the functional role of PPARd in T cells. As proliferation and apoptosis are two cellular processes regulated by PPARd in other cell types, we studied these in human T cells. As shown in Figure 3a, activation of PPARd by the specific synthetic ligand L-165041 enhanced the proliferation of Jurkat T cells. Conversely, cellular proliferation was inhibited in cells on RNA interference (RNAi)-mediated knockdown of PPARd (Figure 3b). Similarly, the fraction of T cells harboring a PPARd-specific RNAi

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construct decreased by threefold after 4 days in culture, but not in cells harboring control lentivirus (Figure 3c), demonstrating a proliferative disadvantage in PPARd-deficient T cells. Thus, PPARd enhances proliferation in human T cells. PPARd protects from IFN-a-induced apoptosis

Having identified PPARd as a type 1 IFN target in human T cells, we examined whether it regulates the effect of type 1 IFN on T-cell apoptosis. As shown in Figure 4a, knockdown of PPARd in human Jurkat T cells increased the rate of spontaneous apoptosis by approximately twofold from 18 to 33%, as measured by the frequency of annexin V þ cells. Culturing of Jurkat cells in the presence of IFN-a after infection with empty control lentivirus modestly increased the frequency of annexin V þ cells. By contrast, apoptosis dramatically increased from 33 to 65% on IFN-a stimulation after lentivirus-mediated knockdown of PPARd. Conversely, activation of endogenous PPARd in Jurkat cells by the PPARdspecific synthetic ligand L-165041 was able to completely inhibit the (B2-fold) increase in apoptotic cells on IFN-a stimulation (Figure 4b). To verify this effect in primary human cells, we infected phytohemagglutinin-activated CD3 þ T cells from peripheral blood either with PPARd RNAi lentivirus or empty control virus. Weak annexin V staining was evident in approximately 35% and strong annexin V staining in approximately 10% of infected cells, indicative of cellular stress caused by the infection with lentivirus (Figure 4c, upper middle panel). Stimulation with IFN-a caused a significant

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Figure 2. Induction of PPARd expression in naı¨ve primary human T cells. (a) CD3 þ T cells were purified from PBMCs using magnetic-activated cell sorting-mediated negative depletion as detailed in Materials and Methods. Subsequently, cells were incubated for 48 hours with either IL-2 (40 ng ml1), INF-a (200 ng ml1), anti-Fas (20 ng ml1), concanavalin A, H2O2 (1 mM), INF-g (20 ng ml1), or TNF-a (20 ng ml1), as indicated. Nuclear (N) and cytosolic (C) fractions were then subjected to SDS-PAGE and western blot. Per lane, 20 mg of extract was loaded. (b) Naı¨ve primary CD3 þ T cells were stimulated with INF-a cultured for 48 hours in the presence or absence of cycloheximide (5 mM). PPARd mRNA expression was analyzed by RT-PCR. Glyceraldehyde3-phosphate dehydrogenase was used as the internal control to confirm equal amounts of cDNA loading. (c) Naı¨ve primary CD3 þ T cells were treated with 200 ng ml1 INF-a for the time points indicated. PPARd expression was analyzed by RT-PCR as outlined in (b). (d) Naive primary CD3 þ T cells from six healthy donors were stimulated with INF-a for 12 hours, followed by analysis of PPARd RNA expression by PCR. Expression was quantitated densitometrically and normalized to glyceraldehyde-3-phosphate dehydrogenase expression. Data shown represent results from five separate donors.

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Figure 3. PPARd enhances T-cell proliferation. (a) Jurkat T cells were grown in the presence of 1 mM PPARd-specific ligand L-165041 (shaded columns) or vehicle (black). Cell number was determined by counting at the indicated time points. Data shown represent mean±SD of three independent experiments, each performed in duplicate. *Po0.001 in a two-sided paired t-test. (b) Jurkat T cells were infected with PPARd-RNAi lentivirus (black symbols) or empty control virus (gray). Cellular proliferation was assessed by MTS-colorimetric conversion, as described in Materials and Methods, at the indicated time points. Data shown represent mean and SD of the quintuplicate determinations. Two independent experiments showed comparable results. *Po0.001 in a two-sided paired t-test. (c) Jurkat T cells were infected with control lentivirus (pLL3.7) or lentivirus harboring a PPARd-specific siRNA sequence (PPARdRNAi). The frequency of infected cells was determined by FACS analysis of the GFP reporter gene. The percentage of infected cells is shown in the figure. (d) Western blot of Jurkat T cells 48 hours after infection with lentivirus, as specified in (b). Ponceau S staining of the blot is shown for verification of even protein loading. cyt, cytosolic fraction; nuc, nuclear fraction.

increase in annexin Vbright staining cells (from 11 to 18%) only after PPARd knockdown, but not in control virusinfected cells, suggesting that PPARd inhibits type 1 IFNinduced apoptosis in primary human T cells. PPARd protects from serum withdrawal- but not Fas-induced apoptosis

Apoptosis mediated by serum withdrawal proceeds through bypassing the activation of the death-inducing signaling complex triggered by Fas. To elucidate which of these proapoptotic pathways are regulated by PPARd in T cells, we infected Jurkat T cells with PPARd knockdown or control lentivirus and subjected them subsequently to either serum deprivation or stimulation with anti-Fas antibody. Detection of early apoptotic cells was then performed by annexin V staining, whereas cells with reduced DNA content (sub-G1) or advanced DNA fragmentation were visualized by

propidium iodide DNA staining in fixed cells. As shown in Figure 5 (panels labeled ‘‘normal medium’’), knockdown of PPARd alone, in the absence of further treatments, resulted in an approximately twofold increase in early apoptotic annexin V þ (11.1 vs 7.6%) and sub-G1 (61 vs 29%) cells, confirming the results described above. Serum withdrawal increased the percentage of annexin V þ cells by twofold after PPARd knockdown, but not in cells harboring control lentivirus. Similarly, the frequency of cells exhibiting advanced DNA loss was twofold higher after PPARd knockdown than in controls cells (66 vs 39%; Figure 5b). Remarkably, G1- and G2/S-phase cells all but disappeared after PPARd knockdown, suggesting a potent protective effect of PPARd on apoptosis mediated by serum deprivation. Upon Fas activation, Jurkat T cells stained uniformly with annexin Vbright (Figure 5a, panels on right), corresponding to a marked accumulation of sub-G1, but not advanced fragmented cells (Figure 5b), regardless of the www.jidonline.org 1943

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Figure 4. PPARd protects from INF-a-induced apoptosis. (a) Jurkat T cells were infected with empty control lentivirus (pLL3.7) or lentivirus harboring a PPARd-specific siRNA sequence, as indicated in the figure. Infection was confirmed by FACS of the enhanced GFP reporter, as shown on the left. Forty-eight hours after infection, the cells were cultured in the presence or absence of INF-a for an additional 48 hours. Cells were then stained with annexin V-Cy5, followed by FACS analysis of GFP-positive cells. Annexin V-positive cells are indicated by the horizontal bar. (b) Jurkat T cells were stimulated with INF-a for 48 hours in the presence of 1 mM of the PPARd-specific ligand L-165041 or vehicle (DMSO), followed by quantification of annexin V þ cells as outlined in (a). (c) Primary T cells were isolated from peripheral blood using a stimulatory CD3 antibody, as detailed in Materials and Methods, processed as in (a). For quantification of annexin V þ cells, lentivirus-infected cells were gated by FACS of the GFP reporter, as shown on the left. Comparable results were obtained in CD3 þ cells from three independent healthy donors.

presence of PPARd. Thus, PPARd blocks apoptosis induced by serum deprivation but not by Fas activation. Mechanism of antiapoptotic PPARd activity in T cells

The antiapoptotic role of PPARd in keratinocytes has been suggested to proceed by transcriptional activation of PDK and ILK1 (Di-Poi et al., 2002), although this could not be 1944 Journal of Investigative Dermatology (2008), Volume 128

reproduced by other investigators (Burdick et al., 2007). In T cells, we were unable to detect any transcriptional up- or downregulation of either kinase upon PPARd activation by synthetic ligand, or on PPARd knockdown (Figure 6a and b). The results shown above suggest that apoptosis induced by growth factor deprivation through serum withdrawal is inhibited by PPARd. As the antiapoptotic action of growth

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Figure 5. PPARd protects from apoptosis induced by serum withdrawal. (a) Jurkat cells were infected either with empty control lentivirus (pLL3.7) or PPARd-knockdown virus (PPARRNAi), followed by serum withdrawal or treatment with anti-Fas antibody, as detailed in Materials and Methods. Apoptosis was quantified by FACS-mediated detection of annexin V þ cells on gated GFP-positive cells. (b) Determination of cell-cycle distribution by propidium iodide (PI)-mediated DNA staining of GFP-positive cells. Treatment of Jurkat cells was identical to that described in (a). Marker bounds employed to quantify populations with different DNA contents are shown at the bottom of the figure. Comparable results were obtained in cells from two independent donors.

factors contained in serum is mediated through the activation of extracellular signal-regulated kinase (ERK)1/2 signaling, we studied ERK1/2 phosphorylation in Jurkat T cells infected with PPARdRNAi or control lentivirus. As shown in Figure 6c, serum withdrawal led to a strong increase in ERK1/2 phosphorylation, an antiapoptotic response that has been noted previously (Erhardt, Mol Cell Biol; 1999 and Narita, J Biol Chem). By contrast, PPARd knockdown completely blocked ERK1/2 phosphorylation. Furthermore, stimulation of Jurkat cells with IFN-a led to notable phosphorylation of nuclear ERK1/2 only after concurrent activation of PPARd with the synthetic ligand, but not in the absence of PPARd activation. Thus, the protective effect of PPARd on IFN-a and serum withdrawal-induced apoptosis in T cells is mediated, at least in part, by enhanced activation of ERK1/2. DISCUSSION This study identifies PPARd as a target gene of type 1 IFN in T cells. Available evidence suggests that transcriptional regulation of PPARd is complex and cell type-dependent. In murine keratinocytes, PPARd is induced by TNF-a and IFN-g by activation of an activator protein-1-binding site, and repressed by CCAAT/enhancer-binding protein isoforms and Smad3 (Tan et al., 2001, 2004; Di-Poi et al., 2005). This pathway has not been confirmed in primary human

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Figure 6. Phosphorylation of ERK1/2, but not expression of ILK and PDK1 kinases, is regulated by PPARd in T cells. (a) Jurkat T cells were stimulated with 1 mM L-165041 or vehicle (DMSO) for 48 hours. The expression of PDK1 and ILK1 was analyzed by RT-PCR. (b) Jurkat T cells were infected with control lentivirus (pLL3.7) or lentivirus harboring a PPARd siRNA sequence (PPARdRNAi). Forty-eight hours after infection, the expression of PPARd, ILK1, and PDK1 was analyzed by RT-PCR. (c) Jurkat cells were infected as in (b), followed by serum withdrawal (SF) or incubation in normal medium (RPMI) for 24 hours. Western-blot analysis was performed using whole-protein extracts. (d) Jurkat T cells were treated with 200 ng ml1 INF-a in the presence or absence of PPARd-specific ligand GW-501516 for 24 hours. Western blots were performed using nuclear protein extracts.

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keratinocytes, but there may exist species differences, as PPARd is expressed in human, but not murine, adult interfollicular epidermis. In human colorectal cancer cell lines, it is regulated by b-catenin through a T-cell factor/lymphoid enhancer factor site (He et al., 1999), whereas it is induced by vitamin D/retinoid X receptor heterodimers (Dunlop et al., 2005) in several human epithelial cell lines. Although the induction through TNF-a in T cells reported here presumably proceeds through the previously characterized mechanism, the cis-acting elements that are operative downstream of type 1 IFN and TCR activation remain to be determined. In silico analysis of the human PPARd promoter predicts several candidate IFN-stimulated response elements and nuclear factor of activated T cell elements. Clearly, more studies are needed to define the transcriptional regulation of PPARd in T cells in detail. The data presented here show that PPARd blocks apoptosis induced by serum withdrawal. Growth-factor deprivation by serum withdrawal was chosen to further characterize the role of PPARd in apoptosis, as this pathway has been reported to be regulated by PPARd in keratinocytes (Di-Poi et al., 2002). Our results show that PPARd is indeed required for protection from growth-factor deprivation-induced apoptosis, but not by Fas-induced apoptosis in T cells. This activity is, at least in part, mediated by ERK1/2 phosphorylation. It is likely that the activation of ERK1/2 signaling also contributes to the stimulation of T-cell proliferation, which we observed. Possibly, activation of ERK1/2 signaling by PPARd is cell type-dependent, as it has not been observed in keratinocytes (Burdick et al., 2007). As ERK1/2 phosphorylation occurs also in response to oxidative stress (Hill et al., 2006), the induction of PPARd by hydrogen peroxide observed here (Figure 2) may not only protect from oxidative stress-induced apoptosis by induction of catalase (Pesant et al., 2006), but also by activation of ERK1/2. Taken together, PPARd appears to act as an activator of ERK1/2 signaling in response to extracellular stress. Concerning the mechanism of action, although the present data show that the induction of PPARd by IFN-a causes activation of ERK1/2 (Figure 5d), this may not be the only mechanism underlying the modulation of type 1 IFN signaling. Thus, the inhibition of apoptosis in T cells may involve preferential signaling of type 1 IFN through signal transducer and activator of transcription (STAT)3 as opposed to STAT1 or STAT2 (Tanabe et al., 2005). In human T-cell leukemia cells, STAT3 phosphorylation blocks apoptosis by transcriptional induction of the bcl-2 member Mcl-1 (Epling-Burnette et al., 2001). We did not observe transcriptional effects of PPARd on the levels of any STAT isoform either in primary T cells or Jurkat cells (data not shown). However, an indirect regulation of STAT3 function by PPARd is conceivable. Thus, activation of PPARg, which acts antagonistically to PPARd in many biological processes, enhances apoptosis and causes dephosphorylation of STAT3 (Kim et al., 2005). Taken together, several cellular actions may be involved in the antiapoptotic activity of PPARd. 1946 Journal of Investigative Dermatology (2008), Volume 128

The inhibition of IFN-a-induced apoptosis in T cells by PPARd raises a number of intriguing questions about the role of PPARd in the regulation of T-cell survival in vivo (Marrack and Kappler, 2004). Thus, it is conceivable that the induction of PPARd contributes to the antiapoptotic effect of type 1 IFN on activated T cells previously observed in mice (Marrack et al., 1999). Apart from modulating IFN signaling, PPARd may affect activation-induced cell death (Hildeman et al., 2002), possibly by linking T-cell activation to metabolic state (Chinetti et al., 2000; Kostadinova et al., 2005). Therefore, a more detailed analysis of chronic PPARd activation on the persistence of activated T cells in vivo is warranted. Currently, activation of PPARd in vivo by synthetic ligands is being evaluated in clinical trials as a potential long-term treatment for metabolic syndrome (Chang et al., 2007; Sprecher et al., 2007). An increased steady-state level of activated T cells observed after prolonged drug-induced PPARd activation would necessitate a re-appraisal of this treatment approach. Finally, these data represent previously unknown evidence for a contribution of PPARd to the pathogenesis of psoriasis. Recently, we showed that PPARd is highly upregulated in psoriasis lesions and stimulates the proliferation of epidermal keratinocytes (Romanowska et al., 2007). Apart from keratinocyte proliferation, the activation of type 1 IFN signaling and the persistence of activated CD8 þ T cells are additional key features of this complex disease. Strikingly, increased phosphorylation of ERK1/2 has recently been discovered in active psoriasis lesions (Johansen et al., 2005). Therefore, the current results indicate that induction of PPARd by type 1 IFN contributes to the prolonged survival of activated T cells in psoriatic skin by activation of ERK. PPARd-mediated ERK activation also antagonizes PPARg, as this isoform has a proapoptotic effect in T cells (Harris and Phipps, 2001, 2002) and is directly inhibited by ERKmediated phosphorylation (Hu et al., 1996). Consistent with this notion, activation of PPARg has an inhibitory effect on psoriasis, whereas this is not the case with PPARd activation (Kuenzli and Saurat, 2003; Malhotra et al., 2005). Finally, PPARd expression has been detected in dendritic cells (Jakobsen et al., 2006) and its activation stimulates angiogenesis in endothelial cells (Piqueras et al., 2006). Thus, PPARd affects several cell populations critical for psoriasis pathogenesis and its inhibition may represent a promising approach for drug development. MATERIALS AND METHODS Patients and skin samples All experiments with samples obtained from patients were carried out in accordance with the Declaration of Helsinki Principles. Patients undergoing biopsy gave a written statement of consent subject to prior approval by the Charite´ Institutional Review Board and were undergoing in-patient treatment for active disease at the time of sampling. Full-thickness excisional biopsies approximately 3 cm2 in diameter were processed as described (Kohlmann et al., 2004). Briefly, samples were trimmed off fatty tissue, rinsed in prewarmed phosphate-buffered saline (PBS), cut into approximately 2 mm pieces, and placed in AIM-V medium including

N al Yacoub et al. PPARd in T Cells

penicillin/streptomycin and 5% human serum. Cells emigrating from the tissue scaffolds were first re-seeded in 96-well round-bottomed plates and, on outgrowth of T-cell lines, expanded on flat-bottomed plates in the presence of 40 ng ml1 IL-2 (Falcon, Heidelberg, Germany, no. 353077).

and ILK: sense, 50 -atcacacactggatgccgta-30 and anti, 50 -cttccaatgcca ccttcatt-30 (500 bp). Amplification conditions were as follows: for glyceraldehyde-3-phosphate dehydrogenase, annealing at 551C and 25 cycles; for PPARd, 541C and 28 cycles; for PDK and ILK, 551C and 28 cycles.

Reagents

Western blotting

Human INF-a and INF-g were obtained from PBL Biomedical Laboratories, Reutlingen, Germany (no. 11100-1). Recombinant human TNF-a was from PeproTech, Hamburg, Germany (no. C-300001ASS). Anti-CD123-FITC, CD19-MicroBeads, CD14-MicroBeads, a-CD16-FITC, CD3-MicroBeads, and anti-FITC-MicroBeads were purchased from Miltenyi Biotec (Bergisch Gladbach, Germany). Annexin V-Cy5 and anti-CD56-FITC were obtained from BD Biosciences (Heidelberg, Germany). Anti-p44/42 (ERK1/2) and anti-phospho-p44/42 (no. 877-616) were obtained from Cell Signalling (Frankfurt, Germany). Phytohemagglutinin-A and H2O2 were from Sigma-Aldrich (Munich, Germany). The selective PPARd agonist L-165041 (Calbiochem, no. 422175) was diluted and aliquoted in DMSO under N2 atmosphere. Cells were stimulated with 1 mM L-165041 for the time span specified in the Results section. Medium with fresh L-165041 was added every 24 hours for longer stimulations. Control cells received identical volume of vehicle (0.05% vol/vol final DMSO).

Cells were seeded in sixwell plates at a density of 106 ml1 and treated for 48 hours with the reagents specified under Results. Cells were harvested, washed with PBS, and the pellet was frozen in liquid N2. The NE-PER Extraction Kit (Pierce, Waldbronn, Germany, no. 78833) was used to prepare the nuclear and cytoplasmic fractions. Protein concentration was determined using the BCA kit (Pierce, no. 23225). Samples were diluted with tricine sample buffer (Bio-Rad, no. 161-0739) and boiled at 951C for 10 minutes. Per lane, 20 mg protein was loaded on 7.5% SDS-PAGE. After transferring to nitrocellulose membranes (Schleicher & Schell, no. 23225), blots were stained with Ponceau red and blocked with 4% non-fat milk in TBST (Tris-buffered saline with 0,05% Tween 20) for 30 minutes at room temperature (RT). Subsequently, blots were incubated with anti-human PPARd (Cayman Chemical, no. 101720) diluted at 1:300 in 4% milk/TBST overnight at 41C. After washing 3 times with TBST, blots were incubated with anti-rabbit IgG-horseradish peroxidase (dilution 1:2,000; Amersham, Dreieich, Germany, NA934) at RT. Detection was performed using the ECL Plus western blotting detection system (Amersham, no. RPN2132) on a CCD system. Whole-cell protein extracts were prepared by washing cells twice with ice-cold PBS followed by lysis in ice-cold lysis buffer (0.01 M Tris-HCl, pH 7.5; 0.144 mol l1 NaCl; 0.5% Nonidet P-40; 0.5% SDS; 0.1% aprotinin; 10 mg ml1 leupeptin; 2 mM phenylmethanesulfonyl fluoride; and 5 mg ml1 pepstatin). Insoluble aggregates were removed by centrifugation.

Isolation and maintenance of T cells from peripheral blood mononuclear cells Peripheral blood mononuclear cells (PBMCs) were isolated using density-gradient centrifugation with Ficoll-Paque from venous blood collected from healthy donors. Activated CD3 þ T cells were isolated from PBMCs by positive selection using CD3-stimulatory MACS MicroBeads (Miltenyi Biotec, no. 130-050-101) according to standard procedures. Naive, unstimulated CD3 þ T cells were isolated by magnetic microbead-mediated depletion of CD14-, CD19-, CD123-, CD56-, and CD16-positive cells from PBMCs. For each depletion, two consecutive adsorptions to LS þ columns (Miltenyi Biotec) were used. Purity of the resulting CD3 þ cell fraction was quantified by FACS detection of CD3 and was greater than 93% in all cases. Purified primary T cells were cultured in AIMV medium (Gibco, Mannheim, Germany, no. 12055) supplemented with 5% human serum (PAN Systems, no. P30-2401), 1% penicillin/ streptomycin (Gibco, no. 15140-122), and 40 U ml1 IL-2 (PromoKine, Heidelberg, Germany, no. C-61240) at 371C and 5% CO2 in sixwell plates. For activation, negatively sorted T cells were cultured for 16 hours with phytohemagglutinin (5 mg ml1). The human T-cell line Jurkat (obtained from DSMZ, Heidelberg, Germany, no. ACC282) was cultured in RPMI-1640 medium with 10% fetal calf serum and 1% penicillin/streptomycin at 371C and 5% CO2.

Reverse-transcription PCR Total RNA was isolated using the NucleoSpin II extraction kit (Machery-Nagel, no. 740955) and cDNA, which was synthesized using reagents purchased from Invitrogen (Mannheim, Germany). The primers used for PCR were glyceraldehyde-3-phosphate dehydrogenase: sense, 50 -gtcagtggtggacctgacct-30 and anti 50 -aggggt ctacatggcaactg-30 (product: 420 bp); PPARd: sense, 50 -aactgcagatggg ctgtaac-30 and anti, 50 -gtctcgatgtcgtggatcac-30 (483 bp), PDK: sense, 50 -ggttgggaaccactctttca-30 , anti, 50 -ttgtgcaataggccatgtgt-30 (532 bp);

Generation of lentivirus and infection of cells Production of VSV-G pseudotyped replication-deficient lentivirus was carried out as described in Yacoub et al. (2007). For infection of primary T cells, CD3 þ purified cells were activated with phytohemagglutinin for 16 hours and then propagated in AIM-V medium with IL-2 for 7 days. Subsequently, cells were washed, harvested, and resuspended at 3  106 ml1. A volume of 100 ml of cell suspension was mixed with a maximum of 2 ml virus-containing supernatant to obtain a multiplicity of infection of 10. After adding polybrene to a final concentration of 8 mg ml1, the virus–cell mix was spun for 1 hour at 2,000 r.p.m. at RT. After incubating cells at 371C for 3 hours, spin infection was repeated once. Cells were plated in the virus-containing medium, which was replaced 12 hours later. The efficiency of infection was verified in each case using FACS detection of the enhanced green fluorescent protein (GFP) reporter, as shown in the respective figures in the Results section.

Proliferation assay Cells were plated at 2  104 per well in 96-well plates in 100 ml RPMI medium containing 10% fetal calf serum. At the time points indicated under Results, cells were incubated with the MTS reagent (Promega, Heidelberg, Germany) for 1 hour at 371C, as instructed by the manufacturer, followed by detection of absorbance at 490 nm. www.jidonline.org 1947

N al Yacoub et al. PPARd in T Cells

Analysis of sub-G1 cells and annexin V staining Cells to be analyzed (8  105 per sample) were washed twice with PBS and fixed in 5 ml cold 70% ethanol at 41C for 1 hour. After washing with PBS, cells were treated with 0.1 mg RNaseA (SigmaAldrich, no. R 4875) at 371C for 30 minutes in 400 ml PBS. After addition of propidium iodide (20 mg ml1), samples were incubated for an additional 30 minutes at RT. After washing, the samples were resuspended and fluorescence was determined by FACS. For enhanced differentiation between sub-G1 cells and cells exhibiting advanced DNA fragmentation, fluorescence detection was acquired using log scale rather than linear settings. As S- and G2-phase cells were only partially resolved using these settings, they were quantified in a common region, as indicated in the Results section. For detecting early apoptotic cells, staining with annexin V-Cy5 was performed. Cells were collected, washed twice with PBS, and 1  105 cells were resuspended in 100 ml of 1  binding buffer (10  binding buffer: 0.1 M HEPES, pH 7.4, 1.4 M NaCl, 25 mM CaCl2). Next, 5 ml annexin V-Cy5 was added, and cells were incubated for 15 minutes at RT in the dark. After addition of 300 ml of 1  binding buffer, cells were analyzed by FACS on the FL-2 channel in a FACSCalibur machine.

Induction of apoptosis Jurkat cells or peripheral blood-derived CD3 þ T cells were infected with lentivirus and cultured as specified above. Forty-eight hours after infection, cells were either serum-deprived or exposed to antiFas (Upstate Biotech, Schwalbach, Germany, no. 05-201) at a final concentration of 20 ng ml1, or both, as specified in the Results section. CONFLICT OF INTEREST The authors state no conflict of interest.

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