The Journal of Neuroscience, December 1, 2004, 24(48):10908-10917; doi:10.1523/JNEUROSCI.3987-04.2004
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Neurobiology of Disease
Peroxisome Proliferator-Activated Receptor 
Induces a Clearance Mechanism for the Amyloid-
Peptide
Ira Espuny Camacho,1
Lutgarde Serneels,1
Kurt Spittaels,2
Pascal Merchiers,2
Diana Dominguez,1 and
Bart De Strooper1
1Neuronal Cell Biology and Gene Transfer Laboratory, Center for Human Genetics, Vlaams Interuniversitair Instituut voor Biotechnologie and Katholieke Universiteit Leuven, 3000 Leuven, Belgium, and 2Galapagos Genomics NV, 2800 Mechelen, Belgium
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Abstract
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We investigated whether peroxisome proliferator-activated receptor 
(PPAR) could be involved in the modulation of the amyloid cascade causing Alzheimer's disease. Inducing expression or activating PPAR using synthetic agonists of the thiazolinedione family results in a dramatic decrease in the levels of the amyloid-
(A) peptide in the conditioned medium of neuronal and non-neuronal cells. PPAR does not affect expression or activity of any of the secretases involved in the generation of the A peptide but induces a fast, cell-bound clearing mechanism responsible for the removal of the A peptide from the medium. Although PPAR expression is generally low in the CNS, induction of PPAR expression during inflammation could be beneficial for inducing A clearance. We confirm that the A clearance mechanism can indeed be induced by PPAR activation in primary murine-mixed glia and cortical neuronal cultures. Our results suggest that PPAR-controlled mechanisms should be explored further as potential drug targets for Alzheimer's disease treatment.
Key words: PPAR; TZD; A; APP; NSAIDs; Alzheimer's disease
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Introduction
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Alzheimer's disease (AD), the most common form of dementia in the elderly, is a neurodegenerative disease characterized by the deposition of extracellular amyloid- (A) plaques and the formation of intracellular tangles in the CNS. A plaques are mainly composed of A peptides, protein fragments derived by proteolytic cleavage of the amyloid precursor protein (APP). An inflammatory component appears to contribute also to the pathogenesis of AD with activated microglia, the principal immune cells in the brain, extensively associated with the A plaques and secreting proinflammatory cytokines (Perlmutter et al., 1990
; McGeer et al., 1993; Tan et al., 1999). An important number of retrospective epidemiological studies have provided evidence that long-term treatment with some nonsteroidal anti-inflammatory drugs (NSAIDs) reduce the risk for AD, delay its onset, and slow the progression of the disease (Rich et al., 1995; McGeer and McGeer, 1996; Stewart et al., 1997; Mackenzie and Munoz, 1998; in t' Veld et al., 2001). Initial in vitro experiments showed that the treatment of microglia and monocytes in culture with NSAIDs attenuated the secretion of proinflammatory cytokines in response to fibrilar A. Furthermore, the conditioned medium of these stimulated microglia was neurotoxic, whereas the medium from microglia stimulated with A but treated with NSAIDs displayed a protective effect in neurons (Klegeris et al., 1999; Combs et al., 2000). Interestingly, the thiazolidinedione (TZD) drugs (Willson et al., 2000), agonists of the peroxisome proliferator-activated receptor (PPAR), showed the same anti-inflammatory neuroprotective effect as NSAIDs (Combs et al., 2000). Because some NSAIDs are agonists of PPAR (Lehmann et al., 1995), and given the evidence for a direct role of NSAIDs on amyloid pathology (Lim et al., 2000; Weggen et al., 2001; Jantzen et al., 2002; Q. Yan et al., 2003), the possible role of PPAR in this context deserves additional investigation. PPAR is a nuclear transcription factor belonging to the PPAR family. These transcription factors play important physiological roles in the regulation of lipid metabolism (Mangelsdorf et al., 1995; Lemberger et al., 1996). PPAR is involved in adipocyte differentiation (Chawla et al., 1994; Tontonoz et al., 1994; Hu et al., 1995; Rosen et al., 1999), insulin action (Olefsky, 2000; Steppan et al., 2001), and cell proliferation (Wang et al., 2001; Okano et al., 2002). The recent discovery that PPAR stimulation reduces inflammation in vitro (Lemberger et al., 1996; Colville-Nash et al., 1998; Jiang et al., 1998; Petrova et al., 1999; Ricote et al., 1999; Combs et al., 2000) and in vivo (Heneka et al., 2000; Dehmer et al., 2004), together with the fact that PPAR agonists (Willson et al., 2000) have been used for years in the treatment of diabetes type II, have raised the hope that PPAR could become a drug target for the treatment of neurological diseases with an inflammatory component-like AD. Based on the NSAID studies mentioned above, we speculated that activation of PPAR could have, in addition to its well known immunomodulating role, a more direct effect on the amyloid cascade in AD. We therefore investigated the effects of PPAR activation, either by expressing PPAR or by using TZD agonists, on APP processing and A clearance in different cell lines and primary cultures of brain cells. We demonstrate that the activation of PPAR directly affects the stability of A externally added to the cell, suggesting the activation of a rapid clearance mechanism.
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Materials and Methods
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Generation of adenoviruses. The Ad5/[green fluorescent protein (GFP), human (h) APP695sw, hAPP-C99, hPPAR, hPPAR
, hPPAR
, human retinoic X receptor (hRXR), mNotch 
E-myc, and notch intracellular domain (NICD)] recombinant adenoviruses were generated by Galapagos Genomics NV as described previously (Michiels et al., 2002).
Drugs and peptides used in culture. Rosiglitazone (stock solution prepared in ethanol), pioglitazone (solution in DMSO), and the PPAR antagonist GW9662 (solution in DMSO) were obtained from Alexis Biochemicals. Troglitazone (in DMSO) and retinoic acid (solution in DMSO) were obtained from Biomol Research Laboratories (Plymouth Meeting, PA). A1-40 synthetic peptide was purchased from Sigma (St. Louis, MO).
Antibodies. The antibodies used for A detection were mouse monoclonal WO2 (epitope A4-10; Abeta, Heidelberg, Germany), rabbit polyclonal B7/8 (A peptide), generated as described previously (De Strooper et al., 1995), and mouse monoclonal 4G8 (epitope A17-24; Senetec PLC). For APPs detection, we used the mouse monoclonal 6E10 antibody (A1-16) obtained from Senetec PLC. For APPssw detection, we used the rabbit polyclonal 54 antibody obtained from Mary Savage (Cephalon, West Chester, PA). Full-length APP, -APP-CTF, and -APP-CTF were detected using the rabbit polyclonal B63.1 antibody generated against the last 15 amino acids of APP as described (C. Esselens, B. De Strooper, and W. Annaert, unpublished observations). Mouse monoclonal anti-Myc (9E10) antibody was purchased from Sanver Tech (Turnhout, Belgium). Rabbit polyclonal anti-cleaved Notch (val 1744) was obtained from Cell Signal/Westburg. The rabbit polyclonal antibodies anti-hPPAR (H-100), anti-hPPAR (H-74), and anti-hRXR (D-20) were obtained from Santa Cruz Biotechnology (Santa Cruz, CA). Mouse monoclonal anti--actin antibody (A5441) was obtained from Sigma, and polyclonal anti-SP1 antibody (5C059) was obtained from Santa Cruz Biotechnology.
Cell culture. Primary murine-mixed glial and cortical neuronal cultures were established from brains of embryonic day 14 or 17 fetal mice as described previously (Cai et al., 2001). Briefly, the dissected brain cortices were suspended in HBSS supplemented with 0.25% trypsin and 0.01% DNaseI and incubated at 37°C for 10 min. The tissues were then transferred to DMEM (Invitrogen, San Diego, CA) supplemented with 10% (v/v) fetal bovine serum (FBS) and dissociated by repeated trituration. The dispersed cells were collected by centrifugation and plated at 75 x 104 cells/well on 6-well cell culture plates (coated with poly-D-lysine) in B27/Neurobasal media (Invitrogen). For neuronal cultures, 24 hr after plating, cells were treated with cytosine arabinoside (3 x 10-7 M) to prevent non-neuronal (glia) cell proliferation and used 48 hr later. Glia cells were allowed to grow for 7 d before starting the experiments. Cells were infected with recombinant adenovirus with doses of virus defined as 1000, 2000, and 3000 multiplicities of infection (MOI) for 24 hr maximum and kept in culture for another 40-42 hr. Neuronal cultures were treated after APPsw infection with TZD drugs for 9 hr.
Human embryonic kidney 293 (HEK293) APPsw (Swedish mutation) cells, kindly provided by Christian Haass (Adolf Butenandt Institute, Ludwig-Maximilians University, Munich, Germany), and HEK293 cells were kept in culture using DMEM with 10% FBS. For viral infection, cells were seeded at 500,000 cells/6-well plate or 112,000 cells/24-well plate and cultured at 37°C in the presence of 5% CO2. After 24 hr of incubation, cultures (60% confluence) were infected with recombinant adenovirus with 1, 3, 10, 30, or 100 MOI. After 48 hr, medium was discarded, and a minimum volume of DMEM supplemented with 1% FBS was added to the cells for different time periods. When TZD drugs were used, cells were treated for 16 hr, and then samples of conditioned medium were analyzed for A detection. IMR-32 cells were grown in culture in Eagle's minimum essential medium (Invitrogen) supplemented with 1 mM sodium pyruvate, 0.1 mM nonessential amino acids, 1.5 gm/l sodium bicarbonate, and 10% FBS. Cells were differentiated toward a more neuronal phenotype by incubating the cells for 7 d in differentiation medium [medium as described above with 1 mM Bt2cAMP and 2.5 µM bromodeoxyuridine (Sigma)] before infection for 48 hr using 1000 MOI recombinant adenovirus.
Radioactive labeling. IMR-32 differentiated cells and primary murine cortical cultures were labeled with 100 µCi/ml 35S-methionine for 5-7 hr. After labeling, A was immunoprecipitated from the media with 4G8 or B7/8 antibodies, and APP was immunoprecipitated from cell extracts prepared in double-immune precipitation buffer with B63.1 antibody. Samples were resolved in SDS-PAGE gel and analyzed by phosphor-imaging.
Gel electrophoresis and immunoblotting. For Notch Emyc, NICD, APPsw full-length, -APP-CTF, and -APP-CTF detection, cells were lysed in 1% Triton, and postnuclear fractions were isolated by centrifugation at 10,000 x g at 4°C for 15 min. Protein concentrations were determined by the Bradford assay (1976). Proteins were resolved in 10% Bis-Tris SDS-PAGE gels (Invitrogen) and transferred to polyvinylidene difluoride (PVDF) membranes for Western blot detection with c-Myc, anti-NICD (val 1744), or B63 antibodies. For A intracellular detection, cells were lysed in 200 µl of ice-cold radioimmunoprecipitation buffer (0.1% SDS, 0.5% natrium deoxycholate, 1% NP-40, and 5 mM EDTA in TBS, pH 8.0), and postnuclear fractions were isolated by centrifugation at 10,000 x g at 4°C for 10 min. All material was used for A immunoprecipitation using B7/8 antibody. Immunoprecipitated samples were loaded in 12% Bis-Tris SDS-PAGE gels (Invitrogen) and transferred to PVDF membranes for immunoblotting using WO2 antibody. Nuclear cell extracts were prepared as described previously (Andrews and Faller, 1991), and protein concentrations were determined by the Bradford assay (1976). Samples were loaded on 7% Tris-acetate gels (Invitrogen) and transferred to PVDF membranes for immunoblot detection using specific antibodies. Samples of conditioned medium were resolved in 12% Bis-Tris for secreted A detection, or in 4-12% Bis-Tris for APPs and APPs detection, and transferred to PDVF membranes. Immunoblot analysis was performed using WO2, 6E10, or 54 antibodies, respectively. Detection of signal was performed using the chemiluminescence kit (PerkinElmer Life Sciences, Emeryville, CA).
-Secretase cell-free assay. HEK293 APPsw cells were transduced for 48 hr. Sixteen hours after infection, cells were harvested. Membranes were prepared as described previously (Nyabi et al., 2003). Briefly, cells were homogenized with a ball-bearing cell cracker (10 µM), and postnuclear fractions were isolated by centrifugation (800 x g for 10 min). Microsomal fractions were isolated by ultracentrifugation (100,000 x g for 1 hr) and washed twice with 0.02% saponin. The pellet was solubilized in 0.5% 3-[(3-cholamidopropyl)dimethylammonio]-1-propanesulfonate choline acetyltransferase buffer, pH 7, for 1 hr at 4°C, and extracts were cleared (100,000 x g for 1 hr). Solubilized membranes (5 µg) were incubated without or with substrate (APP C100-flag) for 16 hr at 37°C.
A stability assay. HEK293 cells were infected for 48 hr, followed by 16 hr in DMEM and 1% FBS before adding the A1-40 synthetic peptide for different time points. Conditioned medium was collected and subjected to immunoblotting analysis for A stability.
Statistical analysis. Values were expressed as means ± SEM.
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Results
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hPPAR expression reduces steady-state levels of A peptide in the culture medium of HEK293 cells
HEK293 cells stably transfected with APPsw (Citron et al., 1992; Lannfelt et al., 1993; Rossor et al., 1993) were transduced in duplicate with recombinant adenovirus driving GFP (control), hAPPsw, or hPPAR expression. A levels in the medium were assessed by Western blot after 16 hr (Fig. 1A, lanes 1-6). High levels of hPPAR expression resulted in a remarkable reduction of A in the culture medium. In contrast, additional expression of hAPPsw caused, as expected, an increase in A production (Fig. 1A).
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Figure 1. Analysis of the processing of APP in hPPAR-overexpressing cells. HEK293 APPsw cells were transduced with recombinant adenovirus (MOI 100) driving expression of GFP (lanes 1, 2), hPPAR (lanes 3, 4), hAPP695sw (lanes 5, 6), or APP-C99 together with GFP (lanes 7, 8) or hPPAR (lanes 9, 10). Cells and conditioned medium were collected 16 hr after infection. A, Western blot detection of secreted A with WO2 antibody. B, Western blot detection of APPs with 6E10 antibody. C, Western blot detection of APPssw with 54 antibody. D, E, Full-length APP, -APP-CTF, and -APP-CTF were detected by Western blot using B63 antibody. F, Levels of endogenous -actin are shown as a loading control for the cell extracts. G, Cells were transduced with adenovirus driving expression of either GFP or hPPAR with different MOI (1, 3, 10, 30, and 100). A levels were detected with WO2 antibody.
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Both A40 and A42 levels were decreased to a similar extent by hPPAR expression as assessed by a sandwich ELISA (data not shown). Furthermore, we could demonstrate an inverse correlation between the levels of A present in the medium and the levels of hPPAR expression in the cell (Fig. 1G).
hPPAR expression does not modulate expression nor activity of the - and - secretases
APP is cleaved either by the - or the -secretase, resulting in the release of the APP ectodomain (APPs or APPs, respectively). The respective APP C-terminal fragments remaining in the cell membrane (-APP-CTF and -APP-CTF) become substrates for the -secretase complex (see below) (De Strooper et al., 1998). It is possible that PPAR expression modulates the activity of these secretases because increased -secretase or decreased -secretase could indeed result in decreased A generation. Using specific antibodies, we found comparable levels of APPs and APPs in the medium (Fig. 1B,C, lanes 1-6). In addition, the membrane-bound -APP-CTF and -APP-CTF were not significantly affected by PPAR expression (Fig. 1D, lanes 1-6). To further confirm that the effect of PPAR on A levels was not mediated by an effect on the direct precursor of the peptide, -APP-CTF, we expressed a synthetic APP-C99 construct together with hPPAR or GFP and tested again the levels of A in culture (Fig. 1A-F, lanes 7-10). High levels of PPAR again caused a down-regulation of A despite the high expression of APP-C99 (Fig. 1E, lanes 7-10). We further confirmed by Western blotting that levels of expression of the aspartic protease BACE1 or -secretase (Hussain et al., 1999; Sinha et al., 1999; Vassar et al., 1999; R. Yan et al., 1999), and of the candidate -secretases ADAMs 9, 10, and 17 (Pan and Rubin, 1997; Koike et al., 1999; Lammich et al., 1999) were not affected by hPPAR expression (data not shown). We conclude, therefore, that neither the expression nor the activity of - or -secretase nor the stability of the APP-C99 fragment were affected in hPPAR-expressing cells.
-Secretase activity is not affected by PPAR activation
The -APP-CTF and -APP-CTF, products from the - and the -secretase activities, are further cleaved by presenilin/-secretase to generate the secreted p3 and A peptides, respectively, and the cytoplasmic APP intracellular domain (AICD). The fact that APP-CTFs did not accumulate in the hPPAR-transduced cells already suggests normal activity of the -secretase complex (Fig. 1E). However, given the evidence in the literature that NSAIDs can modulate -secretase (Weggen et al., 2001; Eriksen et al., 2003) and knowing that they are also agonists of PPAR (Lehmann et al., 1995), we checked the possibility of PPAR regulating -secretase activity in more detail.
We first evaluated whether the generation of the AICD was affected by PPAR expression. We used a reporter assay based on a C99-Gal4 BD-VP16 AD construct (Karlstrom et al., 2002) that contains the transmembrane domain of APP fused via its AICD domain to the Gal4-binding domain and to the VP16 activation domain. After cleavage, this domain is released from the cell membrane and translocates to the nucleus, where it binds to the Gal4 upstream activator sequence, activating the expression of a luciferase reporter gene. No differences were observed in cells transduced with either control GFP vector or with the hPPAR vector (data not shown), indicating that AICD production was not altered by hPPAR expression at least within the limits of detection of this assay. Next we used a cell-free assay to measure directly the generation of A (Nyabi et al., 2003) (Fig. 2A). Membranes purified from GFP control (Fig. 2A, lanes 1, 2) and hPPAR-transduced cells (Fig. 2A, lanes 3, 4) were incubated either without (Fig. 2A, lanes 1, 3) or with a flag-tagged substrate for -secretase, representing the C-terminal 100 amino acids from APP (APP C100-flag) (Fig. 2A, lanes 2, 4) for 16 hr at 37°C. A production was not altered by hPPAR (Fig. 2A, lanes 2, 4). The substrate is also incubated alone (Fig. 2A, lane 5), showing no degradation. This result implies that downregulation of the A levels observed in hPPAR-expressing cells is not attributable to a total or partial inactivation of the -secretase complex.
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Figure 2. Analysis of-secretase cleavage of APP and of Notch. A, Membranes from HEK293 APPsw cells expressing GFP (lanes 1, 2) or hPPAR (lanes 3, 4) were purified and incubated without (lanes 1, 3) or with (lanes 2, 4) APP C100-flag substrate at 37°C for 16 hr. A was detected with WO2 antibody. hPPAR does not affect A generation in this assay. B-D, HEK293 APPsw cells were coinfected with mNotchE-myc and either GFP (lanes 2, 3) or hPPAR adenovirus (lanes 4, 5). Cells coinfected with NICD and GFP (lane 6) are shown as a positive control for the detection of NICD. Cells only infected with GFP (lane 1) are used as a negative control. Cells were treated for 4 hr with lactacystin (10 µM) to avoid proteasome degradation of NICD. B, Detection of Notch E-myc and NICD using an anti-myc antibody. Levels of the substrate NotchE and NICD were comparable in GFP- and hPPAR-transduced cells, showing a normal processing by -secretase. C, Detection of NICD with antibody (val 1744) specific for the cleaved form of Notch. D, Conditioned medium was collected, and A levels were detected with WO2 antibody to confirm the activity of PPAR in this experiment.
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Notch processing is not affected by hPPAR expression
An important substrate of the -secretase complex is the Notch receptor. After Notch cleavage, NICD is released and translocates to the nucleus, where it activates transcription of genes involved in cell-fate decisions (Jarriault et al., 1995). To investigate in a more direct way the effect of PPAR on -secretase activity, we transduced HEK293 APPsw cells with mNotchE-myc recombinant adenovirus in combination with either hPPAR (Fig. 2B-D, lanes 4, 5) or GFP (Fig. 2B-D, lanes 2, 3). mNotchE-myc is a truncated Notch protein lacking most of the ectodomain that undergoes proteolytic cleavage by the -secretase complex, releasing NICD (Kopan et al., 1996; De Strooper et al., 1999). HEK293 APPsw cells infected with GFP alone (Fig. 2B-D, lane 1) or together with NICD (Fig. 2B-D, lane 6) were used as controls. Similar levels of mNotchE-myc (the substrate) and NICD (the cleavage product) were detected with the anti-myc antibody (Fig. 2B). Using an antibody against cleaved Notch (Val 1744) that recognizes a neo-epitope in NICD after -secretase cleavage (Fig. 2C), we confirmed that similar levels of NICD were generated in GFP- or hPPAR-expressing cells. We noticed the presence of two NICD-like bands (Fig. 2B,C). Because the two bands were detected by both the anti-NICD antibody specific for the -secretase cleavage site at the N terminus and by anti-myc, recognizing the epitope at the C terminus of NICD, both bands must represent full-length NICD. Therefore, we suggest that they correspond to different posttranslational modifications of NICD in HEK293 cells.
To prove that hPPAR was active in the transduced cells, we also checked the levels of secreted A in the conditioned medium. Confirming the results displayed in Figure 1, cells overexpressing hPPAR showed strongly reduced levels of the A peptide (Fig. 2D). Thus, -secretase cleavage of mNotchE-myc is not affected by hPPAR overexpression.
We finally assessed by Western blotting the expression levels of all the -secretase components described in the literature thus far (i.e., presenilin, nicastrin, Pen 2, and Aph 1a) (De Strooper et al., 1998; Yu et al., 2000; Francis et al., 2002; Goutte et al., 2002; De Strooper, 2003). No changes were observed in hPPAR-transduced compared with GFP-transduced cells (data not shown).
We conclude that neither -secretase activity on APP or Notch nor the expression of the known components constituting -secretase complex are affected by hPPAR expression.
hPPAR does not cause accumulation of intracellular A-peptide
We next investigated whether the decreased levels of secreted A could be explained by a deficit in A secretion. We transduced HEK293 APPsw cells with recombinant GFP or hPPAR adenovirus and analyzed postnuclear extracts measuring intracellular A levels at four different postinfection time points by Western blot (Fig. 3B). A in GFP-transduced cells tends to slightly increase over time, whereas A is downregulated in hPPAR-overexpressing cells at 3 hr after infection, and levels remain low and even tend to slightly decrease over time. Levels of secreted A were also measured in this experiment at the same four postinfection time points (Fig. 3A), confirming our previous results. Thus, analogous with the secreted A, the levels of intracellular A are also decreased after hPPAR overexpression, ruling out the possibility of intracellular A accumulation.
Downregulation of A is not mediated by a secreted A-degrading enzyme
The steady-state levels of A are the result of a dynamic equilibrium between production and degradation of the peptide. Because production of A does not seem to be affected, we further investigated whether the turnover of A was increased in hPPAR-expressing cells. We first tested whether a secreted A-degrading enzyme could be involved in A turnover. In a cell-free in vitro assay, conditioned medium from GFP-transduced HEK293 APPsw cells that contain high levels of A was incubated in a 50:50 ratio with fresh medium (Fig. 4A, lane 1) or with medium from hPPAR-transduced HEK293 APPsw cells (Fig. 4A, lane 2) at 37°C for 2 hr. The A peptide was not degraded faster after incubation with medium from hPPAR-overexpressing cells. Longer incubation up to 16 hr also did not result in any measurable effect on A (data not shown). There was no A detectable in medium from hPPAR-transduced cells (Fig. 4A, lane 3) or in medium alone (Fig. 4A, lane 4), as expected. These results rule out basically that a secreted protease is responsible for the observed A degradation. However, it could be argued, as a limitation of the in vitro assay, that a potential soluble protease may need cell-bound factors to be active. To test this possibility, we added conditioned medium from hPPAR-transduced cells to GFP-transduced HEK293 control cells coinfected with APPsw adenovirus (Fig. 4B, lanes 1-4). As controls, we added medium from GFP-transduced HEK293 cells (Fig. 4B, lanes 5-8) or unconditioned fresh medium (Fig. 4B, lanes 9-12). The level of A in the conditioned medium was not different at any of the four time points investigated (Fig. 4B). Together, these results show that the effect of hPPAR on A levels is not mediated by an A-degrading enzyme secreted to the conditioned medium.
Next we investigated the effect on secreted and intracellular A levels in PPAR-transduced HEK293 APPsw cells by the addition of a proteasome inhibitor (10 µM lactacystin), a proton pump ATPase inhibitor (0.1 µM bafilomycin), or an alkalinization agent (30 mM NH4Cl) (data not shown). None of the mentioned treatments rescued the levels of secreted or intracellular A to control levels (GFP-transduced HEK293 APPsw cells). Therefore, we conclude that PPAR effect on A levels is not mediated by proteasome degradation or by a pH-sensitive protease (such as lysosomal degradation).
Clearance of the A40 peptide is increased after hPPAR expression
We next investigated the possibility that hPPAR expression results in an increased cell-dependent A clearance (and rapid degradation). For that purpose, we added exogenously synthetic A40 peptide to the conditioned medium of HEK293 cells and incubated the cells over a 9 hr period (Fig. 5A). In the hPPAR-expressing cells, a decline in A40 synthetic peptide is already observed at 7 and 20 min, with very little remaining A at 3 hr. In GFP-transduced cells, the levels of A40 peptide remain stable. No increase in A was detected in extracts from hPPAR-expressing cells, implying that the peptide was effectively and rapidly degraded once it was taken up by the cells (data not shown). These results show that the stability of the A peptide in the medium is specifically affected in cells with increased levels of expression of the hPPAR.
A peptide downregulation is specific for the PPAR isoform of the PPAR family
We next checked whether other members of the PPAR family had similar effects on A levels (Willson et al., 2000). In addition, we also questioned whether increased expression of the hRXR, known to form the active heterodimer transcription factors in the PPAR family (Miyata et al., 1994; Blumberg and Evans, 1998), could also affect the levels of A present in culture. For this purpose, HEK293 APPsw cells were transduced with GFP control, hPPAR-, hPPAR-, hPPAR-, or hRXR-expressing recombinant adenoviruses. We controlled the expression of the different constructs by immunoblotting the nuclear fractions of the transduced cells with specific antibodies (Fig. 6B-D). No antibody is available to detect expression of the hPPAR gene in Western blotting. Therefore, we had to assume that hPPAR was expressed to a similar extent as its homologs in this experiment. Regardless, as shown in Figure 6A, levels of A secreted into the medium were not significantly altered by any of the other hPPAR or hRXR transcription factors. Thus, we conclude that the effect of hPPAR on A degradation is specific.
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Figure 6. Downregulation of A levels is a specific effect of the hPPAR isoform of the PPAR family. HEK293 APPsw cells were infected either with GFP, hPPAR, hPPAR, hPPAR, or hRXR recombinant adenoviruses. A, Samples from medium were analyzed by Western blot, and APPs and A were detected with WO2 antibody. B-D, Levels of expression of hRXR, hPPAR, and hPPAR were detected by Western blot from nuclear extracts using specific antibodies. E-G, Levels of expression of the SP1 transcription factor are detected as a loading control. The results show that A is downregulated specifically after hPPAR overexpression.
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Similar levels of A downregulation are obtained through activation of endogenous hPPAR
Obviously, both from a cell biological and a pharmacological point of view, it is important to demonstrate that a similar effect on A could be obtained by activating endogenously expressed PPAR. We therefore treated cells in culture with several synthetic PPAR ligands from the TZD drug family (Berger et al., 1996; Willson et al., 2000) (rosiglitazone, troglitazone, and pioglitazone) or with 9-cis-retinoic acid (9-cis-RA), an agonist of the RXR (Mangelsdorf et al., 1992). TZD drugs are direct agonists of the PPAR transcription factor, whereas 9-cis-RA activates the heterodimeric complex formed by PPAR-RXR (Mukherjee et al., 1997). We incubated HEK293 APPsw cells with increasing concentrations of troglitazone or pioglitazone (1, 3, 10, and 30 µM), rosiglitazone (30 µM), or 9-cis-RA (30 µM). As a control for solvent interactions, we treated the cells also with DMSO or ethanol alone. Levels of A and APPs were assessed by Western blot (Fig. 7A-D). As shown in Figure 7A-D, A is decreased after treatment with troglitazone (1, 3, 10, and 30 µM), pioglitazone (1, 3, 10, and 30 µM), 9-cis-RA (30 µM), and rosiglitazone (30 µM) in a dose-dependent manner. The levels of APPs in the conditioned medium are not significantly affected, demonstrating the absence of toxicity of the treatments and equal loading (Fig. 7A-D). Thus, activation of the endogenous levels of PPAR present in HEK293 APPsw cells, through its specific agonists or through 9-cis-RA, activating the PPAR-RXR heterodimer, resulted in a notable reduction of the A levels, comparable with that seen with high levels of hPPAR expression in the cell. To confirm that the TZD effect on A is mediated by PPAR transcriptional activity, we incubated the TZD PPAR-activated cells with the PPAR antagonist GW9662 to reverse the effects (Huang et al., 1999; Willson et al., 2000). As shown in Figure 8, treatment with this antagonist restored A levels to control ones (DMSO treatment), confirming the role of PPAR activation in A downregulation.
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Figure 7. Downregulation of A is reached through TZD activation of endogenous PPAR. A-D, HEK293 APPsw cells were transduced with GFP adenovirus and incubated for 16 hr with several TZD drugs or only transduced with hPPAR adenovirus. Levels of APPs and A were detected by Western blot (WO2 antibody) from samples of conditioned medium. A, Cells were treated with 1, 3, 10, and 30 µM troglitazone with DMSO (solvent) or transfected with hPPAR without treatment. B, Cells were treated with 1, 3, 10, and 30 µM pioglitazone with DMSO or transfected with hPPAR. C, Cells were treated with 30 µM 9-cis-RA with DMSO or transfected with hPPAR. D, Cells were treated with 30 µM rosiglitazone, ethanol (solvent), or transfected with hPPAR. A-D, A quantification from two to four independent experiments. Values of A were normalized for APPs (A/APPs) and referred to the GFP control value set as 100%.
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A is downregulated in a differentiated human neuroblastoma cell line, in primary cultures of murine glia cells, and in murine cortical neurons through an hPPAR-activated pathway
We next tested whether the effect of PPAR on A levels could be confirmed in cells and cell lines that are more relevant for AD. First, we used the human neuron-related cell line IMR-32 (Donnelly-Roberts et al., 1998). The cells were differentiated and then transduced with either GFP or hPPAR adenovirus and metabolically labeled with 35S-methionine for 5 hr. No secreted A was detected in hPPAR-expressing cells, whereas control cells secreted detectable levels of A (Fig. 9A). It should also be noted that in these experiments APP was expressed at endogenous levels. We next tested whether A could be downregulated in primary cultures of murine-mixed glia cells cotransduced with APPsw together with either GFP or hPPAR. Levels of A present in the culture medium are clearly downregulated after hPPAR expression in glia cells (Fig. 9B). Finally, we found in primary cortical neuronal cultures cotransduced with APPsw and hPPAR and treated with increasing concentrations of troglitazone (3, 10, and 30 µM) a decrease of A levels. Probably reflecting the relative low efficiency of transduction of primary neurons, we obtained an 
30% reduction of A burden when treated with 30 µM troglitazone (Fig. 9C). These data confirm in three additional and relevant cell lines that an A clearance pathway can be activated by hPPAR.
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Discussion
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The work presented here shows a PPAR-specific effect on the regulation of the A steady-state levels in several cell lines and, more importantly, in primary neuronal and mixed glial cell cultures. The effect is conserved in human and in rodent cells. We convincingly showed that high levels of expression of PPAR strongly reduce the levels of secreted and intracellular A in a dose-dependent manner. This effect is (1) specific for the PPAR receptor, because overexpression of any of the other isoforms of the PPAR family did not significantly alter the levels of A; (2) physiologically relevant, because similar downregulation of A is reached viaendogenous PPAR activation by TZD agonists; (3) mediated by the transcriptional activity of PPAR, as demonstrated by the reversion of the effect on A using the GW9662 PPAR antagonist; and (4) independent on levels of expression of APP or on the presence of the Swedish mutation used in some of our assays, as shown by our experiments with differentiated IMR-32 cells expressing endogenous wild-type APP.
Based on this evidence, we conclude that the effect on A is specific, that the effect on A is not an artifact (e.g., caused by the use of adenoviral transduction), that the mechanism involved is present in many cell types (HEK293, IMR-32, primary mixed glia and neuronal cultures), and, finally, that this mechanism can be activated via drug treatment. Our data contrast with two other studies in which either PPAR expression was induced in Chinese hamster ovary cells (Sagi et al., 2003) or endogenous PPAR was activated with TZD in HEK293 cells (Q. Yan et al., 2003). No changes in A levels were observed in these two studies. However, our experiments show that the PPAR effect on A levels is only observed if sufficient time is given to the cells to activate the clearing mechanism and that relatively high levels of PPAR expression are required. We hypothesize that differences in the levels of expression of PPAR or other cellular factors could be possible reasons for the discrepancy between our and previous studies. In any event, we observed in a reproducible and consistent manner that A peptides are reduced in culture media by the action of PPAR. We want to remark that A was detected with at least three different antibodies directed against different A epitopes, indicating that the loss of A signals in our detection system reflects likely complete degradation of the peptide.
Recent data showed that neuroblastoma cells treated with cytokines produced increased levels of A40 and A42 in culture, which could be reversed by the action of TZD or NSAIDs (Sastre et al., 2003). However, our data indicate that the PPAR-mediated effect on A is independent of inflammation and additionally provides some evidence of a cellular A clearance mechanism involved. In vivo data from an AD mouse model treated with either pioglitazone, one of the TZD drugs that crosses the blood-brain barrier, or with the NSAID ibuprofen showed a reduction in the number and size of the A plaques after ibuprofen but not pioglitazone treatment (Q. Yan et al., 2003). However, when total A levels from brain homogenates were analyzed, a reduction in the soluble and insoluble A40 and A42 species was observed after pioglitazone treatment as well, agreeing with our observations. In addition, we noticed that the pioglitazone treatment in this mouse study did not have any effect on microglia activation, in discordance with previous in vitro (Combs et al., 2000) and in vivo (Dehmer et al., 2004) data. We suggest that higher doses and other treatment regimens are required before final conclusions on the effect of TZD drugs on amyloid pathology in vivo can be made.
In any event, our data provide evidence that PPAR can induce an A clearing mechanism in a variety of cells. A previous study proposed PPAR as a regulator of the transcriptional levels of the -secretase enzyme BACE1 because the mechanism involved in A downregulation (Sastre et al., 2003). Neuroblastoma cells, when treated with cytokines, showed higher levels of the mRNA transcript, the protein, and the activity of BACE1. The effect of cytokines was again reverted by the action of TZD or NSAIDs (Sastre et al., 2003). Although this mechanism could indeed operate in situations in which significant inflammation occurs, our findings in noninflammatory conditions show that high levels of PPAR downregulate A without detectable changes in the levels of expression of BACE1 (as measured by Western blot) or in its activity as measured by the generation of the APPs and -APP-CTF. This was further confirmed by the overexpression experiment using an APP-C99 construct, mimicking the cleavage of APP by -secretase. The results showed clearly that neither the generation nor the stability of APP-C99 is affected by hPPAR. In addition, normal expression and activity of the -secretase complex was also detected, suggesting that changes in the generation of A are not involved in the effects observed in our studies. Moreover, we demonstrated that the cleavage of an artificial Notch substrate or the generation of AICD, a possible important -secretase product of APP, is affected by PPAR. Therefore, we conclude that neither - or -secretase cleavage of APP is changed by PPAR expression.
We therefore postulate that the mechanism of action of PPAR is involved with an increase in the rate of A turnover. We confirmed this hypothesis by experiments showing a rapid clearance of A40 synthetic peptide from the culture medium of PPAR-overexpressing cells. The fact that no increase in the signal of intracellular A was detected in the cell extracts strongly suggests that the A peptide is indeed effectively degraded. Interestingly, the degradation of A seems to depend on the presence of cell-bound factors because medium extracted from PPAR-overexpressing cells was not able to degrade A peptide on its own. Besides, we could not find evidence for an involvement of the proteasome or lysosomal systems in this A-degradation mechanism, or at least compounds known to block these systems did not affect A clearance in our hands. We hypothesize that PPAR mediates an activation of a cell-dependent clearance mechanism probably by transcriptional activation of one or several proteins involved in the uptake from the medium and intracellular degradation of the peptide. We want to stress that we believe that this mechanism is physiologically relevant because we could demonstrate its activity not only in HEK293 cells but also in differentiated neuroblastoma IMR-32 cells and in primary murine neuronal and mixed glial cultures (the latter consists mainly of astroglia cells but contains also a small population of microglia and oligodendroglia). Perhaps this clearance mechanism becomes activated in the brain in some pathological situations in which high levels of PPAR are present. Indeed, high levels of PPAR have been detected in brains from AD patients (Kitamura et al., 1999). Importantly, we showed that when we simulated such conditions in culture by expressing PPAR in primary murine cortical neuronal cultures, treatment with TZD compounds indeed resulted in a downregulation of A levels. This not only demonstrates that the PPAR-regulated A clearance mechanism is present in cells of the central nervous system but also strongly suggests that additional work in vivo is needed to fully explore the potential of TZD compounds in the treatment of AD.
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Footnotes
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Received April 24, 2004;
revised October 20, 2004;
accepted October 20, 2004.
This work was supported by a Pioneer award from the Alzheimer's Association (B.D.S.); the Fund for Scientific Research, Flanders; the Katholieke Universiteit Leuven (Geconcerteerde onderzoeksactie); the European Union (Abnormal proteins in the pathogenesis of neurodegenerative disorders); and the Federal Office for Scientific Affairs, Belgium (IUAP P5/19). We thank Christian Haass for the generous gift of the human embryonic kidney cells stably transduced with amyloid precursor protein (Swedish mutation) (APPsw); Mary Savage (Cephalon) for the 54 antibody specifically recognizing-secretase-processed APPsw; Katrien Horre, Kathleen Craessaerts, and Siska Deforce for the preparation of the primary glia and cortical cultures; and Simon Reeve for helpful criticism.
Correspondence should be addressed to Dr. Bart De Strooper, Neuronal Cell Biology and Gene Transfer Laboratory, Center for Human Genetics, Vlaams Interuniversitair Instituut voor Biotechnologie and Katholieke Universiteit Leuven, Herestraat 49, 3000 Leuven, Belgium. E-mail: Bart.destrooper{at}med.kuleuven.Ac.be.
Copyright © 2004 Society for Neuroscience 0270-6474/04/2410908-10$15.00/0
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Abstract
Introduction
