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. 2009 May 20;29(20):6752–6760. doi: 10.1523/JNEUROSCI.0789-09.2009

p53-Dependent Transcriptional Control of Cellular Prion by Presenilins

Bruno Vincent 1,2,✉, Claire Sunyach 1,2, Hans-Dieter Orzechowski 3, Peter St George-Hyslop 4,5, Frédéric Checler 1,2,✉
PMCID: PMC6665908  PMID: 19458243

Abstract

The presenilin-dependent γ-secretase processing of the β-amyloid precursor protein (βAPP) conditions the length of the amyloid β peptides (Aβ) that accumulate in the senile plaques of Alzheimer's disease-affected brains. This, together with an additional presenilin-mediated ε-secretase cleavage, generates intracellular βAPP-derived fragments named amyloid intracellular domains (AICDs) that regulate the transcription of several genes. We establish that presenilins control the transcription of cellular prion protein (PrPc) by a γ-secretase inhibitor-sensitive and AICD-mediated process. We demonstrate that AICD-dependent control of PrPc involves the tumor suppressor p53. Thus, p53-deficiency abolishes the AICD-mediated control of PrPc transcription. Furthermore, we show that p53 directly binds to the PrPc promoter and increases its transactivation. Overall, our study unravels a transcriptional regulation of PrPc by the oncogene p53 that is directly driven by presenilin-dependent formation of AICD. Furthermore, it adds support to previous reports linking secretase activities involved in βAPP metabolism to the physiology of PrPc.

Introduction

Transmissible spongiform encephalopathies (TSE) result from the unconventional conversion of the normal cellular prion protein (PrPc) into its pathogenic, protease-resistant scrapie isoform PrPs.c. (Aguzzi and Polymenidou, 2004). The propagation mechanism apparently does not require any nucleic acid as was proposed in the “protein only” theory (Prusiner, 1998). Tremendous efforts have been devoted to understand the mechanisms by which PrPc converts into PrPs.c. Although the precise process remains to be established, it appears clearly that “prion infection” requires the crucial presence of endogenous PrPc, since PrPc-deficient mice are fully resistant to infection and toxicity after inoculation with pathogenic scrapie-enriched material (Büeler et al., 1993; Brandner et al., 1996). However, since prion diseases are extremely rare with no more than one case per million of individuals in human beings, it is important to understand the physiological role and regulation of PrPc.

PrPc was reported to participate in lymphocyte activation (Cashman et al., 1990), cellular adhesion processes (Rieger et al., 1997; Gauczynski et al., 2001; Mangé et al., 2002), neurite growth (Santuccione et al., 2005), synaptogenesis (Kanaani et al., 2005), cellular signaling (Mouillet-Richard et al., 2000), and cell viability (for review, see Linden et al., 2008). Concerning transcriptional regulation of PrPc, it has been reported that PrPc mRNA levels could be increased either under stress conditions such as ischemia and hypoglycemia, or by some growth factors (Linden et al., 2008). However, little is known about the molecular pathways underlying these regulatory events. The functional characterization of the human prion promoter (Funke-Kaiser et al., 2001) identified two regulatory regions where sequence analysis revealed consensus sequences for AP-1, Sp1, and Sp2 factors (Mahal et al., 2001; Bellingham et al., 2009). Our in silico analysis of the human PRioN protein (PRNP) gene promoter also revealed a motif partly matching the binding sequence targeted by the oncogene p53. We have examined the ability of p53 to modulate PrPc promoter transactivation and mRNA expression and we show that p53 acts as a transcriptional activator of PrPc by binding directly to the suspected promoter sequence. Furthermore, we previously established that the β-amyloid precursor protein (βAPP-intracellular domain amyloid intracellular domain (AICD) generated after presenilin-dependent γ-secretase-mediated cleavage of βAPP behaves as a transcriptional activator of p53 (Alves da Costa et al., 2006). Here, we show that presenilins also control PrPc transcription in a presenilin-dependent manner by increasing AICD-induced p53 expression. Our study is the first demonstration of the involvement of presenilins in the control of PrPc expression and reveals a new pathway linking γ-secretase and p53 to the direct control of PrPc transcription. Since endogenous PrPc is necessary to allow PrPs.c. infectious process, presenilin-dependent control of PrPc levels could be seen as a means to modulate not only the physiological function of PrPc but also PrPs.c. accumulation. This hypothesis is nicely supported by a recent study showing that a γ-secretase inhibitor significantly impairs the accumulation of pathogenic prions in vivo (Spilman et al., 2008), suggesting that such compounds could represent a new class of therapeutic agents against prion diseases.

Materials and Methods

Cell culture and transfections.

Embryonic mouse fibroblasts [presenilin-deficient (PS−/−), βAPP-deficient (APP−/−), APP−/−/APLP2−/−, p19Arf−/−, and p19Arf−/−/p53−/−, as well as their respective wild-type controls], βAPP-over-expressing HEK293 cells and primary cultured neurons were obtained and maintained in culture as previously described (Vincent et al., 1996; Kamijo et al., 1997; Marambaud et al., 1997; Heber et al., 2000; Herreman et al., 2000; Pardossi-Piquard et al. 2005). Transient transfections in HEK293 cells were performed with Lipofectamine 2000 reagent (Invitrogen), whereas fibroblasts were transiently transfected by means of the mouse embryonic fibroblasts Nucleofector kit (Amaxa Biosystems) as described previously (Sunyach et al., 2007).

Immunoprecipitation, SDS-PAGE, and Western blotting.

N1-containing conditioned media were immunoprecipitated and submitted to 16.5% Tris-tricine gels as previously described (Vincent et al., 2000). βAPP, APLP1, APLP2, Fe65, Tip60, and p53 were separated on 8% Tris-glycine gels, whereas PrPc, tubulin, PS1, and PS2 were analyzed on 12% Tris-glycine gels. C50 and C59 fragments were separated on 16.5% Tris-tricine gels. Proteins were then transferred onto Hybond-C nitrocellulose membranes and incubated with the following antibodies: monoclonal SAF32 (N1 and PrPc; SPIBio), monoclonal anti-β-tubulin or anti-β-actin (Sigma), polyclonal anti-NTF-PS1 or anti-Loop-PS2 (Dr. Gopal Thinakaran, University of Chicago, Chicago, IL), polyclonal anti-Fe65 and monoclonal anti-myc (for myc-tagged C50 and C59; Dr. L. Mercken, Vitry-sur-Seine, France), monoclonal anti-HA (HA-tagged Tip60; Covance), polyclonal anti-APLP1 or APLP2 (EMD Biosciences), monoclonal 22C11 (βAPP; Roche Applied Science), or polyclonal anti-p53 (Dr. Jean-Christophe Bourdon, Dundee, Scotland, UK). Blots were revealed by incubations with goat anti-rabbit or goat anti-mouse secondary antibodies coupled to peroxidase (Beckman Coulter) and chemiluminescence recording using a Luminescence Image Analyzer LAS-3000 (Raytest). Quantification of data was performed with the Aida Image Analyzer software (Raytest).

Chronic γ-secretase inhibitor treatments.

Chronic treatment of 4-d-old primary cultured neurons with the γ-secretase inhibitor DAPT (N-[-(3, 5-difluorophenacetyl)-l-alanyl]-S-phenylglycine t-butylester; 10 μm) was performed by addition of the inhibitor at 0 and 9 h for a total incubation of 24 h then PrPc immunoreactivity was monitored by Western blot as described above.

Semiquantitative and real-time PCR analyses.

Total RNA was extracted and purified from mouse embryonic fibroblasts or HEK293 cells with the NucleoSpin RNA II kit (Machery-Nagel). Two micrograms of total RNA were reverse-transcribed using oligo (dT) priming and avian myeloblastosis virus reverse transcriptase (Promega). For semiquantitative PCR, reactions were performed at 94°C for 30 s, then 55°C for 1 min, and finally 68°C for 2 min using 40 cycles, followed by a final extension of 7 min at 68°C. PCR products were analyzed on a 1% agarose gel stained with ethidium bromide. Real-time PCR were performed in an ABI PRISM 5700 sequence detector system (Applied Biosystems) using the SYBR Green detection protocol as outlined by the manufacturer. Specific primers for semiquantitative or real-time PCR were designed using the Primer Express software (Applied Biosystems) and were as follows: mouse PrPc: forward, 5′-CTG CTG GCC CTC TTT GTG AC-3′ and reverse 5′-CTT TTT GCA GAG GCC GAC AT-3′. Human PrPc: forward, 5′-AAT CAA GCA GCA CAC GGT CA-3′ and reverse 5′-TCG GTG AAG TTC TCC CCC TT-3′.

Expression levels of human p53 and PrPc genes and mouse PrPc gene were normalized by monitoring RNA levels of human GAPDH and mouse γ-actin genes using the following primers: forward, 5′-TGG GCT ACA CTG AGC ACC AG-3′ and reverse 5′-CAG CGT CAA AGG TGG AGG AG-3′ for human GAPDH; and forward 5′-CAC CAT CGG TTG TTA GTT GCC-3′ and reverse 5′-CAG GTG TCG ATG CAA ACG TT-3′ for mouse γ-actin.

Measurements of PrPc promoter transactivation.

The 1543 bp of the 5′ untranslated and promoter region of the human PrPc gene was subcloned into the luciferase reporter vector pGL3basic and used to measure PrPc promoter transactivation as has been extensively described (Funke-Kaiser et al., 2001). Cells grown in 12-well plates were cotransfected with PrPc promoter-luciferase, β-galactosidase (to normalize transfection efficiencies), and the indicated cDNAs with Lipofectamine (HEK293 cells) or with the Amaxa Nucleofector kit (mouse embryonic fibroblasts). Thirty-six hours after transfection, luciferase and β-galactosidase activities were measured with appropriate enzyme assay systems (Promega).

Site-directed mutagenesis of PrPc promoter.

Mutation of the p53 putative binding site located on the human PrPc promoter-luciferase reporter construct (Mahal et al., 2001) was introduced using the Quick change site-directed mutagenesis kit (Stratagene) following manufacturer's specifications. The two following sets of primers were purchased from Eurogentec: forward, 5′-CCTATTTTCCCCAGGGAGCACCTGGTTTACGCCC-3′; reverse, 5′-GGGCGTAAACCA-GGTGCTCCCTGGGGAAAATAGG-3′. The resulting construct (CATG replaced by AGCA) was verified by sequencing.

Chromatin immunoprecipitation assay.

Chromatin immunoprecipitation assay (ChIP) was performed according to the instructions of the Chip-IT kit (Active Motif). Briefly, to prepare chromatin, HEK293 cells were seeded in three 150 mm dishes and allowed to reach 70–80% confluency. Cells were fixed, recovered in PBS, cross linked, and processed for chromatin preparation. DNA obtained was digested with the shearing enzyme provided in the kit as recommended by the supplier, yielding chromatin fragments of 200–500 bp in size. Each immunoprecipitation was performed on 50 μg of chromatin in the ChIP immunoprecipitation buffer supplied, with 5 μg of anti-p53 primary antibody (Active Motif) or irrelevant antibody (IgG and RNApol IgG) as negative controls. Immune complexes were collected with 40 μl of a solution of protein G-Sepharose. After elution, cross links were reversed and RNA digested using RNase (100 μg/ml). To digest proteins, SDS (1%), and proteinase K (100 μg/ml) were added, and the samples were incubated overnight at 37°C. The beads were washed as recommended and DNA was purified on columns provided. PCR amplification was performed using primers specific for the −326/−150 bp region of the PrPc promoter (forward, 5′-CAGGAGCCACACAGTTGAAACAGA-3′; reverse, 5-AGGGTGATTTACGTAAAATAGCAAA-3′).

Transgenic mouse brain tissue preparation.

Pieces of brains from APP−/− and APP−/−/APLP2−/− mice (Heber et al., 2000), and Fe65 transgenic and Fe65/AICD transgenic mice (Ryan and Pimplikar, 2005), together with their respective wild-type controls, were homogenized in lysis buffer (10 mm Tris/HCl, pH 7.5, containing 150 mm NaCl, 0.5% Triton X-100, 0.5% deoxycholate, 5 mm EDTA). Protein expressions were analyzed by Western blot as described above.

Immunofluorescence analysis.

Wild-type, PS−/−, or APP−/−/APLP2−/− fibroblasts were grown on glass coverslips in 35 mm dishes. At 50% of confluence, cells were fixed with 1.5% paraformaldehyde for 20 min at room temperature and washed three times with PBS. Permeabilization was performed by the addition of 0.1% Triton X-100 for 5 min followed by three PBS washes (nonpermeabilized cells were maintained in PBS during this time). After incubation for 1 h with 1% nonfat milk in PBS (to limit nonspecific fixation of antibodies), cells were incubated overnight at 4°C with the indicated primary antibodies in 1% nonfat milk/PBS). After three washes with PBS, cells were incubated for 1 h with adequate secondary antibodies conjugated to Alexa Fluor-594 and Alexa Fluor-488 respectively (Interchim). Coverslips were washed, incubated with DAPI to counterstain the nuclei and mounted in Vectashield-mounting medium (Vector Laboratories). Staining was visualized as previously described (Sunyach et al., 2007).

Statistical analysis.

Statistical analyses were performed with the PRISM software (GraphPad) by using the unpaired t test for pairwise comparisons.

Results

Presenilins modulate cellular prion protein at a transcriptional level

We examined the influence of presenilins on the expression of PrPc. Figure 1A shows that PrPc immunoreactivity was drastically decreased (50 ± 3.7% inhibition compared with wild-type cells, n = 20, p < 0.0001) in homogenates prepared from presenilins 1- and 2-deficient (PS−/−) mouse fibroblasts. As expected, the secretion of the N-terminal metabolite of PrPc referred to as N1 (Chen et al., 1995; Vincent et al., 2000, 2001) was concomitantly reduced by PS deficiency (Fig. 1A). In situ staining of PrPc on nonpermeabilized intact cells revealed a drastic decrease in cell surface-associated immunofluorescence in PS-deficient fibroblasts (Fig. 1B, left panels). PrPc labeling on Triton X-100 permeabilized cells revealed a similar decrease of PrPc expression triggered by PS depletion (Fig. 1B, right panels), indicating that the reduction of membrane-associated PrPc expression was not caused by an intracellular accumulation of the protein that would have been triggered by a defect in PS-dependent trafficking of PrPc to the plasma membrane.

Figure 1.

Figure 1.

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Presenilins deficiency reduces PrPc immunoreactivity. A, Wild-type (WT) or PS−/− mouse fibroblasts were incubated for 8 h in serum-free medium; then the secreted PrPc N-terminal metabolite (N1) was immunoprecipitated and analyzed by 16.5% Tris/Tricine electrophoresis, and Western blot as described in Materials and Methods. PrPc, tubulin, and presenilin 1 and 2 immunoreactivities were monitored in cell homogenates as described in Materials and Methods. B, WT and PS−/− cells were fixed and labeled under nonpermeabilized or Triton X-100-permeabilized conditions with antibodies directed toward PrPc (SAF32), N-terminal (NTF), and C-terminal (CTF) parts of presenilins (α-NTF PS1 and α-CTF PS2, respectively) as described in Materials and Methods. Cells were then counterstained with DAPI to visualize nuclei and analyzed on an Axioplan microscope.

We assessed whether the observed reduction in PrPc immunoreactivity resulted from a decrease in its mRNA transcription. We took advantage of the design of a PrPc promoter construct that harbors 1543 bp of the 5′-untranslated promoter region of the PrPc human gene in-frame with a luciferase reporter gene (Funke-Kaiser et al., 2001) to establish that PrPc promoter transactivation was drastically reduced in PS-deficient fibroblasts (63 ± 2.5% of reduction compared with wild-type cells, n = 15, p < 0.0001) (Fig. 2C). This was accompanied by a lowering of PrPc mRNA levels measured by both semiquantitative RT-PCR (Fig. 2A) and real-time PCR analyses (46.6 ± 8.4% of reduction, n = 9, p < 0.0001) (Fig. 2D) in PS-deficient fibroblasts. Importantly, we were able to substantially rescue PrPc promoter transactivation (1.88 ± 0.13-fold increase, p < 0.0001) (Fig. 2C) and mRNA levels (1.69 ± 0.14-fold increase, p < 0.05) (Fig. 2D) after transient overexpression of both PS1 and PS2 in PS-deficient cells (Fig. 2B). Overall, the above data demonstrate that the PSs control PrPc expression at the transcriptional level.

Figure 2.

Figure 2.

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Presenilins regulate PrPc expression at a transcriptional level. A, Analysis of PrPc and actin mRNA levels by semiquantitative PCR in wild-type (WT) and PS−/− fibroblasts. B–D, PS−/− fibroblasts were transiently transfected with either empty pcDNA3 vector (CT) or with PS1 and PS2 cDNA (PS1/PS2) as described in Materials and Methods. Thirty-six hours after transfection, protein expression (B), PrPc promoter transactivation (C), and mRNA levels (D) were monitored. Bars in C and D express percentage of mock-transfected WT cells and represent the means ± SEM of 10 or 6 independent experiments, respectively. *p < 0.05; **p < 0.0001.

βAPP and APLPs are involved in the regulation of PrPc transcription

PSs are pleiotropic proteins (Checler, 1999) that are thought to harbor an aspartyl protease activity referred to as γ-secretase (Wolfe et al., 1999). To assess whether the PS-dependent control of PrPc expression could be linked to γ-secretase activity, primary cultured neurons were exposed to DAPT, a specific PS-directed inhibitor (Dovey et al., 2001). Figure 3A clearly shows that chronic treatment of neurons with DAPT significantly reduced PrPc expression (37 ± 4% inhibition, n = 3, p < 0.0001). The βAPP is historically the first protein described as a substrate of the PS-dependent γ-secretase activity (De Strooper et al., 1998). Furthermore, PS-dependent cleavage of βAPP yields a C-terminal metabolite called AICD that has been suggested to behave as a transcription factor (Müller et al., 2008). We therefore examined whether βAPP could be involved in the control of PrPc expression and promoter transactivation. We first showed that βAPP overexpression triggers a significant increase of PrPc immunoreactivity (1.6 ± 0.07-fold increase, n = 8, p < 0.0001) (Fig. 3B) in human stably transfected HEK293. To rule out any artifactual effect resulting from overexpression procedure and to confirm that endogenous βAPP indeed controls PrPc expression, we monitored PrPc immunoreactivities in βAPP-deficient fibroblasts, and we observed a reduction of PrPc expression in absence of βAPP (19.3 ± 3.6% inhibition, p < 0.0001) (Fig. 3C), a phenotype reminiscent of that observed in presenilin-deficient cells (Fig. 1A). Interestingly, the lowering of PrPc expression was further amplified in βAPP−/−/APLP2−/− fibroblasts (63 ± 6% inhibition, p < 0.0001) (Fig. 3C) that are lacking APLP2, a member of the βAPP family that also undergoes PS-dependent γ-secretase-like cleavages (Scheinfeld et al., 2002). Interestingly, we show that PrPc immunoreactivity was fully restored by transient transfections of βAPP or APLP2 cDNAs in βAPP−/−/APLP2−/− cells (Fig. 3D). It should be noted that APLP1, another member of the βAPP family that is expressed in the brain but absent in fibroblasts, mimics the potential of rescue harbored by βAPP and APLP2 (Fig. 3D). The above findings were further supported by in vivo data showing a dramatic reduction of PrPc expression in both βAPP- and βAPP/APLP2-deficient mouse brains (Fig. 3E). Finally, immunofluorescence staining of PrPc at the cell surface of permeabilized and nonpermeabilized βAPP−/−/ APLP2−/− cells indicated that the combined absence of βAPP and APLP2 drastically impaired PrPc expression at the plasma membrane (Fig. 4A), as was observed in PS-deficient cells.

Figure 3.

Figure 3.

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PrPc expression is reduced by a γ-secretase inhibitor and is modulated by βAPP and APLP2 in cells and in mice brain. A, Primary cultured neurons were chronically treated with the γ-secretase inhibitor DAPT (10 μm) for a total period of 24 h; then PrPc and tubulin immunoreactivities were analyzed by Western blot as described in Materials and Methods. B, PrPc expression in stably transfected mock (mock)- and βAPP (APP)-expressing HEK293 cells (left). C, PrPc, tubulin, βAPP, and APLP2 immunoreactivities in homogenates of wild-type (WT), βAPP-deficient (APP−/−), and APP−/−/APLP2−/− fibroblasts (left) were monitored as described in Materials and Methods. Bars corresponding to densitometric analyses express percentage of control WT cells and represent the means ± SEM of 20 independent experiments. *p < 0.0001. D, Empty pcDNA3 (CT), βAPP (APP), APLP1, or APLP2 cDNAs were transiently transfected in APP−/−/APLP2−/− double knock-out fibroblasts. Thirty-six hours after transfection, PrPc and tubulin immunoreactivities (left) or transgene expressions (right) were monitored as described in Materials and Methods. E, PrPc, tubulin, βAPP, and APLP2 immunoreactivities were monitored in whole homogenates prepared from WT, APP−/−, and APP−/−/APLP2−/− mice brains as described in Materials and Methods.

Figure 4.

Figure 4.

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βAPP and APLP2 regulate PrPc at a transcriptional level. A, Wild-type (WT) and APP−/−/APLP2−/− cultured fibroblasts were fixed and labeled with the indicated antibodies in nonpermeabilized or Triton X-100 permeabilized conditions as described in Materials and Methods. Cells were then counterstained with DAPI to visualize nuclei (in blue) and processed as described in Figure 1B. B, Transactivation of the PrPc promoter was measured in WT, APP−/−, and APP−/−/APLP2−/− cells as described in Materials and Methods. Bars corresponding to luciferase activities (normalized for β-galactosidase activities) express percentage of control activity recovered in WT cells and represent the means ± SEM of 20 independent experiments performed in triplicates. ****p < 0.0001. C, D, Analysis of PrPc mRNA levels in WT, APP−/−, and APP−/−/APLP2−/− fibroblasts by semiquantitative (C) and real-time (D) PCR. Bars in D express percentage of control mRNA levels measured in WT cells and represent the means ± SEM of three independent experiments realized in triplicates. **p < 0.01; ***p < 0.003. E, APP−/−/APLP2−/− fibroblasts were transiently transfected with either empty pcDNA3 vector (CT) or βAPP cDNA. Thirty-six hours after transfection, βAPP immunoreactivity (left), PrPc promoter transactivation (middle), and mRNA levels (right) were measured as described in Materials and Methods. Bars express percentage of control promoter transactivation or mRNA levels recovered in WT cells and represent the means ± SEM of 8 or 3 independent experiments, respectively. *p < 0.03; ****p < 0.0001.

Four lines of data indicate that βAPP and APLP2 control PrPc expression at a transcriptional level. First, the transactivation of the human PrPc promoter was significantly impaired by βAPP and βAPP/APLP2 depletion (63 ± 5% reduction, p < 0.0001 and 81 ± 3% decrease, p < 0.0001, respectively) (Fig. 4B). Second, βAPP and βAPP/APLP2 deficiency lowered PrPc mRNA levels as shown by semiquantitative (Fig. 4C) and quantitative real-time PCR (70 ± 14% decrease, p < 0.01 and 69 ± 10% inhibition, p < 0.003, respectively) (Fig. 4D). Third, βAPP partially restored both PrPc promoter transactivation (2.07 ± 0.05-fold increase, p < 0.0001) and PrPc mRNA levels (3.02 ± 0.37-fold increase, p < 0.03) (Fig. 4E). Fourth, both luciferase activity (2.75 ± 0.14-fold increase, n = 6, p < 0.0001) and PrPc mRNA levels (1.93 ± 0.11-fold increase, n = 10, p < 0.0001) were significantly augmented after βAPP overexpression in HEK293 cells, ruling out a possible cell specific βAPP-mediated control of PrPc transcription (data not shown).

The γ-secretase-derived βAPP metabolite AICD controls PrPc expression and promoter transactivation

The βAPP-related effects on PrPc expression and transcription were fully reminiscent of those associated with PSs. Since the latter appeared dependent on γ-secretase activity, we reasoned that the production of a βAPP metabolite generated by PS-dependent γ-secretase activity could account for the observed modulation of PrPc mRNA levels and protein expression. Interestingly, AICD59 (C59) and AICD50 (C50) derive from the γ- and ε-secretase attacks of βAPP, respectively, and have been suggested to modulate a series of genes at a transcriptional level (Müller et al., 2008). Therefore, we examined the possibility that AICD could modulate the transcription of PrPc. The adaptor protein Fe65 and the histone acetyltransferase Tip60 have been reported to enhance AICD immunoreactivity and to promote AICD translocation to the nucleus (Cao and Südhof, 2001). Therefore, we transiently transfected C50 or C59 together with Fe65 and Tip60 in βAPP−/−/APLP2−/− fibroblasts (Fig. 5A). Clearly, both C50 and C59 overexpression significantly enhanced PrPc immunoreactivity (1.14 ± 0.05-fold increase, p < 0.005 and 1.55 ± 0.16-fold increase, p < 0.005, respectively) (Fig. 5A), PrPc promoter transactivation (1.13 ± 0.04-fold increase, p < 0.002 and 1.64 ± 0.12-fold increase, p < 0.0001, respectively) (Fig. 5B), and PrPc mRNA levels (2.47 ± 0.12-fold increase, p < 0.05 and 1.73 ± 0.04-fold increase, p < 0.05, respectively) (Fig. 5C). Here again, both C50 and C59 heighten PrPc mRNA levels (2.41 ± 0.26-fold increase, p < 0.01 and 1.68 ± 0.19-fold increase, p < 0.03, respectively) in HEK293 cells (Fig. 5D). Finally, we took advantage of transgenic mice overexpressing both C59 and Fe65 (Ryan and Pimplikar, 2005) to determine whether the AICD-mediated control of PrPc expression described here also takes place in vivo in mouse brain. Indeed, C59/Fe65 transgenic mice show enhanced cerebral expression of PrPc compared with Fe65 single transgenic animals (Fig. 5E). Since PS-dependent γ-secretase activity is able to cleave other substrates than βAPP, thereby generating other ICDs, we examined the effect of notch ICD (NICD), mΔENotch (which engenders NICD after γ-secretase cleavage), or the γ-secretase-derived C-terminal products of E- and N-cadherins (E-Cad/CTF2 and N-Cad/CTF2, respectively), and established that none of these fragments were able to modulate PrPc levels (data not shown).

Figure 5.

Figure 5.

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Presenilin-dependent control of PrPc transcription is mediated by βAPP intracellular domains. A–C, APP−/−/APLP2−/− fibroblasts were transiently transfected with empty pcDNA3 vector (CT) or C50/C59 cDNA together with Fe65 and Tip60-encoding plasmids as described in Materials and Methods. Thirty-six hours after transfection, PrPc expression (A), promoter transactivation (B) and mRNA levels (C) were monitored as described in Materials and Methods. Bars express percentage of corresponding control values obtained with pcDNA3-transfected cells and represent the means ± SEM of 12 (A), 3 (B), or 4 (C) independent determinations. *p < 0.05; ****p < 0.0003. D, HEK293 cells were transiently transfected with empty pcDNA3 vector (CT) or either C50 or C59 in combination with Fe65 and Tip60 cDNA in as described in Materials and Methods. Thirty-six hours after transfection, mRNA levels were monitored by real time PCR. Bars express percentage of control (pcDNA3-transfected cells) and represent the means ± SEM of 3 independent experiments performed in triplicates. **p < 0.03; ***p < 0.01. E, Expressions of PrPc, C59, Fe65, and tubulin were analyzed by Western blot, as described in Materials and Methods, in singly (Fe65) or doubly (Fe65/C59) transgenic mice brain homogenates.

AICD-mediated regulation of PrPc is p53-dependent and occurs through direct binding of p53 to PrPc promoter

The question arises by which mechanisms AICD could control PrPc promoter transactivation, and thereby, mRNA and protein levels. We have previously demonstrated that AICDs can positively control p53 at the transcriptional level (Alves da Costa et al., 2006). In silico examination of the PrPc promoter revealed a putative although incomplete p53-binding site (el-Deiry et al., 1992) (Fig. 6B). It was therefore tempting to speculate on the possibility that p53 could account for the AICD-mediated regulation of PrPc transcription. Six lines of independent data confirm that this was indeed the case. First, PrPc mRNA levels were reduced by p53 depletion (Fig. 6A). Second, C50- and C59-mediated effects on PrPc mRNA levels were fully p53-dependent because the C50/59-induced increase in PrPc mRNA was prevented by p53 depletion (Fig. 6A). Third, PrPc expression (decrease of 34 ± 8%, n = 5, p < 0.01) (Fig. 6C), promoter transactivation (decrease of 49 ± 5%, p < 0.001) (Fig. 6D), and mRNA levels (decrease of 55 ± 15%, p < 0.01) (Fig. 6E) were reduced by p53 depletion. Fourth, p53 cDNA transfection in p53-deficient cells rescued PrPc promoter activation (2.44 ± 0.07-fold increase, p < 0.0001) (Fig. 6D) and mRNA levels (2.6 ± 0.13-fold increase, p < 0.001) (Fig. 6E). Fifth, ChIP, using a set of primers framing the putative p53 binding site described above, showed that p53 physically bound to the human PrPc promoter (Fig. 6F, lane 5). Sixth, mutation of the putative p53 binding site located on the PrPc promoter reporter construct not only resulted in a significant decrease of baseline luciferase activity in p19Arf −/− cells expressing endogenous p53 levels (Fig. 6G, compare WT and Mut), but also abolished the ability of overexpressed p53 to trigger PrPc promoter transactivation (Fig. 6G, compare WT/p53 and Mut/p53). This set of data brings the demonstration that p53 could act as a direct activator of PrPc promoter transactivation and strongly argues in favor of a p53-mediated AICD-dependent regulation of PrPc transcription.

Figure 6.

Figure 6.

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Regulation of PrPc transcription by p53. A, p19Arf−/− (Arf−/−) and p19Arf−/−p53−/− (Arf−/−/p53−/−) fibroblasts were transiently transfected with Fe65 and Tip60 cDNA together with either empty pcDNA3 vector (CT), C50 or C59 cDNA. Thirty-six hours after transfection, PrPc mRNA levels were measured by real-time PCR as described in Materials and Methods. Right panel shows representative Western blot analyses of each of the over-expressed proteins. Bars express percentage of control (Arf−/− cells transfected with pcDNA3, Fe65 and Tip60 cDNA) and represent the means ± SEM of 4–8 experiments performed in duplicates. *p < 0.05; #p < 0.0005; ns, not statistically significant. B, Alignment of typical p53 consensus binding motive (el-Deiry et al., 1992) with the partial p53-binding site identified on human PrPc promoter (Mahal et al., 2001) where involved nucleotides are highlighted in high caps. C, PrPc expression was monitored in Arf−/− and Arf−/−/p53−/− fibroblasts as described in Materials and Methods. D–E, Arf−/− and Arf−/−/p53−/− fibroblasts were transiently transfected with empty pcDNA3 vector (CT) or p53 cDNA. Thirty-six hours after transfection, PrPc promoter transactivation (D) and RT-PCR analysis of mRNA levels (E) were monitored as described in Materials and Methods. Bars express percentage of control pcDNA3-transfected Arf−/− cells and represent the means ± SEM of 7 and 4 independent determinations, respectively. **p < 0.01; ***p < 0.005; ****p < 0.001; #p < 0.0005. F, ChIP analysis of p53 binding on the PrPc promoter sequence was performed as described in Materials and Methods. Lane 1: molecular markers, 2: DNA input, 3: IgG control, 4: RNA pol IgG, 5: anti p53 IgG, 6: water, 7: PrPc promoter reporter construct. G, Arf−/− fibroblasts were transiently transfected with pcDNA3 or p53 cDNA together with wild-type PrPc promoter (WT, white bars) or with PrPc promoter mutated on its partial p53-binding site (Mut, gray bars). Thirty-six hours after transfection, luciferase activity was measured as described in Materials and Methods. Values are expressed as a percentage of control (cells transfected with the wild-type PrPc promoter alone, WT) and are the mean ± SEM of 6 experiments performed in triplicates. *p < 0.05; ****p < 0.001; ns, not significant.

Discussion

Although the role of the PrPc in transmissible spongiform encephalopathies has been clearly established (Aguzzi and Polymenidou, 2004), its physiological function as well as the regulation of its expression are yet poorly understood. The demonstration that PrPc-deficient mice are viable with no obvious deleterious phenotype (Büeler et al., 1992) would suggest either that endogenous PrPc does not fulfill any vital functions, or that such functions could be complemented by yet unknown proteins/mechanisms. Nevertheless, several studies indicated that PrPc could contribute to several distinct processes, including lymphocyte activation, synaptic transmission, cell adhesion, signaling, and apoptosis (for review, see Linden et al., 2008). Other works also suggested that PrPc could be involved in the neuronal differentiation of PC12 cells triggered by either interleukin-6 or nerve growth factor, as shown by increased expression of PrPc mRNA levels (Lazarini et al., 1994). Consistent with a pivotal link between NGF and PrPc, PrPc mRNA expression was also increased after intracerebral injection of this growth factor in cholinergic neurons (Mobley et al., 1988). Accordingly, Satoh et al. (1998) reported on cell-specific modulation of PrPc mRNA levels after treatment with other cytokines (interleukin 1β), growth factors (tumor necrosis α), and tumor promoting drugs (phorbol esters).

Molecular cloning of the human PRNP gene promoter facilitated the understanding of the transcriptional regulation of PrPc (Funke-Kaiser et al., 2001; Mahal et al., 2001). Thus, the 5′-flanking region of PRNP revealed several putative binding sites for transcription factors including Sp1, AP1, AP2, c-REL, and Nkx2–5, suggesting stimulus-dependent and cell-specific mechanisms of transcriptional regulation. Interestingly, Mahal et al. (2001) also reported a CATG sequence located 746 nt upstream to the transcriptional start site of the human PRNP gene. This CATG motif partially mimics the p53 consensus binding site sequence (el-Deiry et al., 1992). Whether this putative p53-binding domain was functional remained to be established. Nevertheless, the concomitant presence of p53 and SP-1 binding sequences on the PRNP promoter was interesting with respect to the fact that these two transcriptional factors were reported to physically interact to form functional hetero-complexes (Borellini and Glazer, 1993; MacLeod, 1993; Gualberto and Baldwin, 1995).

Our study clearly demonstrates that p53 acts as a functional activator of the PRNP gene promoter transcription. First, p53 deficiency drastically lowers PrPc expression, mRNA levels, and promoter transactivation that all could be rescued by transient transfection of p53 cDNA. Second, substitution of the putative p53-binding site CATG by AGCA fully abolishes p53-dependent upregulation of PrPc transcription. Third, ChIP unravels a direct interaction between p53 and PrPc promoter. Therefore, our study establishes a physical and functional link between p53 and PRNP promoter triggering transcriptional activation and increased expression of PrPc. Noteworthy, a very recent study described an ATM (ataxia-telangectasia-mutated)-mediated binding of p53 to the PRNP promoter in response to copper-induced oxidative stress that promotes an elevation of PrPc (Qin et al., 2009).

It should be noted, however, that the PRNP gene did not emerge when a global mapping of p53-binding sites in the human genome was performed recently by ChIP coupled to paired-end ditag sequencing (PET) (Wei et al., 2006). This likely reflects a weak physical PRNP promoter/p53 interaction as could have been expected from the fact that the human PrPc promoter only contains one of the two canonical p53-binding CATG half-sites.

Since we previously established that PSs could modulate p53 expression (Alves da Costa et al., 2006), this prompted us to examine first whether PSs could control PrPc expression. A first set of experiments clearly showed that PS depletion lowered PrPc levels in cell homogenates as well as in intact cultured cells, and reduced PrPc mRNA levels and its promoter transactivation. Interestingly, PrPc immunoreactivity, promoter transactivation, and mRNA levels could be restored by PS1 and PS2 expression in PS-deficient fibroblasts. This clearly established that endogenous PSs could modulate PrPc transcription. That this phenotype was linked to the reported ability of PSs to display γ-secretase activity was demonstrated by the fact that the presenilin-binding γ-secretase inhibitor DAPT indeed reduced PrPc expression in primary cultured neurons.

PS-dependent γ-secretase targets a series of substrates (Wakabayashi and De Strooper, 2008) that could have been theoretically considered as putative mediators of p53-dependent PS-associated modulation of PrPc. We examined the possibility that PS/γ-secretase-mediated cleavages of βAPP and APLP could be responsible for this phenotype, because we previously established that AICD and ALID (βAPP-like intracellular domain), the intracellular metabolites of the βAPP and APLP, respectively, generated by PS-dependent γ-secretase activity, could control p53 levels (Alves da Costa et al., 2006). Five lines of data indicate that this was indeed the case: first, βAPP overexpression increases PrPc expression; second, βAPP and APLP depletion lowered PrPc immunoreactivity, promoter transactivation. and mRNA levels in fibroblasts as well as in mice brains; third, βAPP and APLP cDNA transfection could rescue PrPc promoter transactivation and mRNA levels in βAPP/APLP2 deficient cells; fourth, AICD increases PrPc expression, promoter transactivation, and mRNA levels after overexpression in both βAPP/APLP2-deficient fibroblasts, whereas AICD-transgenic mice display higher PrPc levels than wild-type animals; and fifth, AICD-mediated modulation of PrPc mRNA levels was fully abolished by p53 deficiency. Overall, our study demonstrates for the first time that PSs control PrPc levels via its ability to generate AICD. Thereafter, AICD increases p53 that ultimately acts as a transcriptional activator of PRNP gene promoter (Fig. 7). Moreover, it appears that AICD-mediated cascade leading to the modulation PrPc is rather specific since other γ-secretase-derived fragments such as NICD or E- and N-cadherin C-terminal fragments did not affect PrPc levels.

Figure 7.

Figure 7.

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Schematic representation of the γ-secretase-mediated presenilin-dependent transcriptional activation of PrPc via AICDs and p53. The cleavage of βAPP by the β-secretase BACE1 yields the C99 fragment that undergoes subsequent attack by the presenilin-dependent γ-secretase. This results in the concomitant productions of Aβ and AICD. AICD associates with the adaptor protein Fe65 and the histone acetyltransferase Tip60 to form a stable complex that translocates into the nucleus where it can induce the promoter transactivation of several gene targets including p53 (Alves da Costa et al., 2006) that acts as a direct activator of PrPc promoter transactivation directly (this study). Therefore, PS-dependent γ-secretase-mediated production of AICD ultimately leads to a p53-dependent augmentation of cellular prion protein expression at the plasma membrane.

Interestingly, various neuropathological stigmata have previously been shown to label both Alzheimer's disease (AD)- and prion-affected brains (Powers et al. 1991; Hainfellner et al., 1998), and it has been reported that PrPc gene polymorphism could increase the risk factor of developing early onset AD (Dermaut et al., 2003; Riemenschneider et al., 2004). Furthermore, cell biology approaches also indicated that several molecular effectors apparently bridge Alzheimer's disease and prion pathophysiology. Thus, PrPc undergoes constitutive and PKC-regulated proteolysis by membrane-bound proteases of the disintegrin family of metalloproteases, namely ADAM10 and ADAM17 (Vincent et al., 2000, 2001). This proteolytic attack that takes place at the 111/112 peptidyl bond, i.e., in the middle of the putative neurotoxic 106–126 region of PrPc (Forloni et al., 1993), is reminiscent of the well characterized α-secretase cleavage of βAPP, which occurs inside the Aβ sequence, is performed by the same enzymes, and is similarly upregulated by protein kinase C (Checler and Vincent, 2002). βAPP and PrPc not only undergo common metabolic processes but can apparently cross talk. Thus, the levels of PrPc and its distribution within cholesterol-rich lipid rafts strongly impact on the β-secretase cleavage of βAPP and thereby reduce Aβ production by restricting the access of BACE1 to βAPP (Parkin et al., 2007). Together with the present data, these observations establish that the secretases responsible for the metabolism of βAPP are also able to interfere with PrPc physiology.

Several studies have clearly established that endogenous PrPc was necessary for the PrP-scrapie-associated (PrPs.c.) pathology since mice devoid of PrPc resist infection by pathogenic inoculates (Büeler et al., 1993; Brandner et al., 1996). In this context, strategies aimed at reducing endogenous levels of PrPc could be seen as theoretical methods to protect against prion infections. This could be envisioned by increasing disintegrin-mediated cleavages as we have proposed (Checler and Vincent, 2002). Is γ-secretase inhibition another putative therapeutic approach to interfere with prion propagation? This possibility has been supported by a recent very interesting study showing that an orally administered γ-secretase inhibitor, combined to quinacrine, was able to reduce PrPs.c. in the neocortex and hippocampus of mouse brains (Spilman et al., 2008). The present demonstration of a PS-dependent and γ-secretase-mediated control of PrPc transcription provides the molecular basis of the above empirical observation and suggests that γ-secretase inhibitors could slow down the spread of pathogenic PrPs.c. in prion-infected animals simply by reducing its endogenous PrPc template. Therefore, our work not only describes the tumor suppressor p53 as a transcriptional activator of PRNP gene promoter transcription but also unravels a new PS-dependent function in the control of PrPc and potentially opens new perspectives concerning the treatment of prion-associated pathologies.

Footnotes

This work was supported by the Fondation pour la Recherche Médicale and by the Conseil Général des Alpes-Maritimes. We thank Drs. S. Pimplikar (Cleveland, OH), U. Müller (Heidelberg, Germany), M.F. Roussel (Memphis, TN), B. De Strooper (Leuven, Belgium), and P. Saftig (Göttingen, Germany) for kindly providing cell lines and brain samples. We sincerely thank T. C. Südhof (Dallas, TX), G. Thinakaran (Chicago, IL), J. C. Bourdon (Dundee, Scotland, UK), and L. Mercken (Vitry-sur-Seine, France) for their generous gift of cDNAs and antibodies.

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