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The Journal of Immunology, 2006, 177: 6137-6142.
Copyright © 2006 by The American Association of Immunologists, Inc.

Prion Protein Expression by Mouse Dendritic Cells Is Restricted to the Nonplasmacytoid Subsets and Correlates with the Maturation State1

Gloria Martínez del Hoyo*, María López-Bravo*, Patraporn Metharom2,{dagger}, Carlos Ardavín3,4,* and Pierre Aucouturier3,4,{dagger}

* Department of Immunology and Oncology, Centro Nacional de Biotecnología/Consejo Superior de Investigaciones Cientificas, Universidad Autónoma, Madrid, Spain; and {dagger} Université Pierre et Marie Curie, Unité Mixte de Recherche S Institut National de la Santé et de la Recherche Médicale Unité 712, Paris, France


   Abstract
Top
 Abstract
Introduction
 Materials and Methods
 Results
 Discussion
 Disclosures
 References
 
Expression of the physiological cellular prion protein (PrPC) is remarkably regulated during differentiation and activation of cells of the immune system. Among these, dendritic cells (DCs) display particularly high levels of membrane PrPC, which increase upon maturation, in parallel with that of molecules involved in Ag presentation to T cells. Freshly isolated mouse Langerhans cells, dermal DCs, and DCs from thymus, spleen, and mesenteric lymph nodes expressed low to intermediate levels of PrPC. Highest levels of both PrPC and MHC class II molecules were displayed by lymph node CD8{alpha}int DCs, which represent fully mature cells having migrated from peripheral tissues. Maturation induced by overnight culture resulted in increased levels of surface PrPC, as did in vivo DC activation by bacterial LPS. Studies on Fms-like tyrosine kinase 3 ligand bone marrow-differentiated B220– DCs confirmed that PrPC expression followed that of MHC class II and costimulatory molecules, and correlated with IL-12 production in response to TLR-9 engagement by CpG. However, at variance with conventional DCs, B220+ plasmacytoid DCs isolated from the spleen, or in vitro differentiated, did not significantly express PrPC, both before and after activation by TLR-9 engagement. PrP knockout mice displayed higher numbers of spleen CD8{alpha}+ DCs, but no significant differences in their maturation response to stimulation through TLR-4 and TLR-9 were noticed. Results are discussed in relation to the functional relevance of PrPC expression by DCs in the induction of T cell responses, and to the pathophysiology of prion diseases.


   Introduction
Top
 Abstract
 Introduction
Materials and Methods
 Results
 Discussion
 Disclosures
 References
 
The prion protein (PrP)5 was initially described as an essential component of the infectious agents responsible for transmissible spongiform encephalopathies (TSE) (1). TSE are a group of neurodegenerative disorders that include Creutzfeldt-Jakob disease and kuru in humans, bovine spongiform encephalopathy, sheep scrapie, and chronic wasting disease in deer and elk. Although the pathophysiology of TSE remains poorly understood (reviewed in Refs. 2 and 3), an almost invariable feature is the accumulation of an abnormal isoform of PrP (scrapie PrP, designated PrPSc) in infected tissues of affected individuals. PrP was found to be encoded by a unique gene of the host (4), Prnp (PRNP in humans), the structure of which is remarkably conserved between species (5). Its physiological product is expressed as a GPI-anchored membrane protein termed cellular PrP (PrPC), in many tissues at variable levels (6, 7). Because neurons generally display strong PrPC expression, and because the CNS is specifically involved during the clinical stages of TSEs, most studies were devoted to the potential functions of PrPC in the brain (8, 9, 10, 11). Although PrP knockout (Prnp–/–) mice display no gross functional anomaly, the structure of PrP and its regulated expression during development and cell differentiation suggest that its physiological role(s) might be more important than one would expect from observations in experimental animals (12).

Outside the nervous system, PrPC is particularly expressed on the membrane of cells of hemopoietic lineages, including platelets, lymphocytes, and mononuclear phagocytes (7, 13, 14, 15). Published studies have shown that PrPC expression is finely tuned during the differentiation, maturation, and activation processes of T cells (16, 17) and dendritic cells (DC) (18). These observations are of particular interest because of the following: 1) lymphoid tissues are specifically involved in the early stages of TSEs (19, 20); 2) the expression of normal PrPC is required for cells to support prion replication (21, 22, 23); and 3) PrPc might function as a receptor for scrapie PrP (24). Although sites of prion accumulation during the incubation period are relatively well defined in different TSE models (reviewed in Refs. 19 and 25), the mechanisms responsible for prion penetration and transport remain unclear. Because after exposure to pathogens and/or inflammatory compounds DCs migrate from Ag capture areas to the T cell zones of organized lymphoid tissues, these cells represent potential candidates as prion carriers. Indeed, DCs were shown to transport prions to the mesenteric lymph after oral inoculation (26), and to transfer prion infection to the brain (27). Other studies suggest that DCs can degrade TSE agents (28, 29), as do macrophages (30).

Thus, although DCs are clearly involved in TSE pathogenesis, their precise role in the etiology of this disease remains unclear. In this study, we have specifically analyzed the expression of PrPC by DC subpopulations from the skin, thymus, spleen, and lymph nodes (LNs) of the mouse, in an attempt to determine whether distinct DC subsets would be more prone to be differentially targeted by prions. In light of another recent study (31), our results on PrPC expression by DCs should also help in understanding its physiological role in the immune system.


   Materials and Methods
Top
 Abstract
 Introduction
 Materials and Methods
Results
 Discussion
 Disclosures
 References
 
Mice

Five- to 6-wk-old C57BL/6 mice were purchased from IFFA Credo. PrP knockout (Prnp–/–) mice (21) were provided by C. Weissmann (Imperial College, London, U.K.), and backcrossed 10 times on the C57BL/6 background. Animals were bred under strict specific pathogen-free conditions following European recommendations on animal ethics.

Preparation of epidermal Langerhans cells (LCs) and dermal DCs (DDCs)

Ears were split into dorsal and ventral halves and incubated with 0.5% trypsin (Sigma-Aldrich) for 45 min at 37°C, to allow the separation of the epidermis from the dermis. The epidermal or dermal sheets were incubated at 37°C for 12 h in RPMI 1640, with 100 ng/ml murine rGM-CSF (PeproTech). After this incubation period, LCs or DDCs were harvested from the culture medium.

DC-enriched cell fractions

Spleens and thymuses from both control and Prnp–/– mice were cut into small fragments and digested with collagenase A (0.5 mg/ml; Boehringer Mannheim) and DNase I (40 µg/ml; Boehringer Mannheim) in RPMI 1640 supplemented with 5% FCS, for 10 min at 37°C with continuous agitation. Digested fragments were filtered through a stainless-steel sieve, and cell suspensions were washed twice in PBS solution, supplemented with 5% FCS and 5 mM EDTA, containing 5 µg/ml DNase I. Mesenteric LNs (MS-LNs) were mechanically disgregated and filtered through a stainless-steel sieve, and cell suspensions were washed twice in PBS solution, supplemented with 5% FCS and 5 mM EDTA, containing 5 µg/ml DNase I. The cell suspensions were then resuspended in cold isosmotic Optiprep solution (Nyegaard; pH 7.2), density 1.061 g/cm3, and a low-density fraction, accounting for <1% of the starting cell population, was obtained by centrifugation at 1700 x g for 10 min at 4°C. B cells were then removed magnetically after incubation for 30 min at 4°C with sheep anti-mouse Ig-coated magnetic beads (Dynabeads; Dynal Biotech) at a 6:1 bead:cell ratio.

In vivo DC maturation assays

Analysis of in vivo maturation of splenic DCs was performed 3 h after i.v. injection of 25 µg of the TLR-4 ligand LPS from Escherichia coli (Sigma-Aldrich).

In vitro differentiation of DCs with Fms-like tyrosine kinase 3 ligand (Flt3L; Flt3L-DCs)

Flt3L-DCs were generated from lysis buffer-treated bone marrow cells cultured at 1 x 106 cells/ml in 24-well plates, in RPMI 1640 supplemented with 10% FCS, 50 µM 2-ME, 100 U/ml penicillin-streptomycin, 1 mM NaPyr, and 100 ng/ml human Flt3L (PeproTech), at 37°C and 5% CO2.

Maturation of Flt3L-DCs

Day 8 Flt3L-DCs were cultured for additional 24 h in control conditions or in the presence of 6 µg/ml TLR-9 ligand CpG oligodeoxynucleotide-1826: TCCATGACGTTCCTGACGTT (CpG) or 1 mg/ml LPS from E. coli. Inhibition of p-38 MAPK kinase and ERK kinase was achieved by pretreatment with the inhibitors SB203580 (Sigma-Aldrich) and UO126 (Sigma-Aldrich), respectively, for 30 min before addition of CpG.

Flow cytometry

Analysis of DCs from the spleen, thymus, and MS-LNs was performed on DC-enriched fractions freshly isolated or overnight cultured in RPMI 1640 supplemented with 10% FCS, 50 µM 2-ME, 100 U/ml penicillin-streptomycin, 1 mM Na Pyr, and 15% of a J558 cell line-conditioned medium, as a source of GM-CSF. Before staining the samples, all conjugated Abs were carefully titrated to ensure correct quantitative comparisons. After FcR blocking with mAb 2.4G2, DCs were analyzed by gating on CD11c-positive cells, after triple staining with FITC-conjugated anti-CD11c (clone N418); PE-conjugated anti-CD8{alpha} (clone CT-CD8{alpha}); and biotin-conjugated anti-MHC class II (MHC II; clone FD11-54.3), anti-CD40 (clone FGK45), anti-CD86 (B7-2, clone GL1), or anti-PrP (clone SAF83; provided by J. Grassi, Commissariat à l’Energie Atomique, Saclay, France), followed by streptavidin-tricolor (TC; Caltag Laboratories). All analyses, including before and after overnight culture, were performed in comparison with the same cells incubated with the corresponding isotype control Ig.

Analysis of LCs and DDCs was performed by gating on CD11c-positive cells, after double staining with FITC-conjugated anti-CD11c (clone N418) and biotin-conjugated anti-MHC class II (clone FD11-54.3) or anti-PrP (clone SAF83), followed by streptavidin-PE (Caltag Laboratories).

Analysis of plasmacytoid DCs (pDCs) was performed by gating on CD11c-positive cells, after triple staining with FITC-conjugated anti-CD11c (clone N418), PE-conjugated anti-CD45R (B220, clone RA-6B2), and biotin-conjugated anti-MHC class II (clone FD11-54.3) or anti-PrP (clone SAF83), followed by streptavidin-TC.

Phenotypic analysis of Flt3L-DCs was performed after triple staining with FITC-conjugated anti-CD45R (B220, clone RA3-6B2); PE-conjugated anti-CD11b (Mac1, clone MI/70); and biotin-conjugated anti-MHC class II (clone FD11-54.3), anti-CD40 (clone FGK45), anti-CD86 (B7-2, clone GL1), or anti-PrP (clone SAF83), followed by streptavidin-TC.

Analyses were performed on a FACSort flow cytometer (BD Biosciences).

ELISA for IL-12

Cryopreserved supernatants from day 9 Flt3L-DCs cultured at 1 x 106 cells/ml in 24-well plates, in control conditions or in the presence of CpG for the last 24 h, were tested for the presence of IL-12, using a mouse IL-12 p70 ELISA kit (BD Pharmingen).


   Results
Top
 Abstract
 Introduction
 Materials and Methods
 Results
Discussion
 Disclosures
 References
 
PrP expression by DC subpopulations isolated ex vivo

Expression of PrP by the main DC subpopulations present in mouse lymphoid organs was analyzed in DC-enriched cell preparations, after gating for CD11c-positive cells, as described in Materials and Methods.

These included skin LCs and DDCs, CD8+ and CD8– DCs (found in most lymphoid organs, including the thymus, spleen, and LNs), CD8int DCs (which constitute a LN-specific DC subset corresponding to epithelium-related DCs that have migrated to LNs via afferent lymphatics), and B220+ pDCs. These DC subpopulations are endowed with specific functions and differ in their maturation state, which correlates with their level of expression of MHC II and costimulatory molecules. Therefore, as shown in Fig. 1, the analysis of PrP expression has been performed in parallel with that of MHC II expression on conventional DC subpopulations and pDCs, from the skin, thymus, spleen, and MS-LNs, after ex vivo isolation and after an additional overnight culture period that triggers DC maturation.


Figure 1
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  		FIGURE 1. PrP expression by ex vivo isolated DC subpopulations. Histograms show the expression of MHC II and PrP by mouse conventional and pDC subpopulations from the skin, MS-LN, spleen, and thymus, directly after ex vivo isolation (gray profiles), and after 12-h culture in the absence of exogenous cytokines (bold line profiles). Open profiles represent biotin-conjugated isotype control Ab staining. Percentages of cells above background staining are indicated for ex vivo isolated DCs (upper figures) and 12-h cultured DCs (lower bold figures), except for the histograms corresponding to MHC II expression by LCs and DDCs. In these cases, the mean fluorescence intensity values are indicated, to better evaluate the variations in MHC II expression. Expression profiles are represented on a logarithmic scale in the x-axis. Results are representative of five experiments for cells analyzed directly ex vivo, and of three experiments for cells analyzed after 12-h culture.

 
Regarding skin DCs, both freshly isolated LCs and DDCs expressed low to intermediate levels of PrP, which were clearly up-regulated after overnight culture, in parallel with a significant increase of MHC class II expression reflecting the induction of a maturation process. However, because the isolation procedure used to obtain skin DCs involved a trypsin-mediated digestion step, and some surface molecules are known to be trypsin sensitive (this is not the case of MHC II molecules; see Ref. 32), the increased PrP level observed for skin DCs after overnight culture could also reflect partial or total PrP re-expression after proteolysis caused by the isolation procedure.

CD8+ DCs from the thymus, spleen, and LNs expressed low to intermediate levels of PrP. In the spleen and MS-LNs, CD8– DCs expressed similar PrP levels as their CD8+ counterparts. PrP expression by both CD8+ and CD8– DC subsets slightly increased upon overnight culture, this up-regulation being somewhat higher for CD8– DCs. This moderate PrP up-regulation undergone by CD8+ and CD8– DCs after overnight culture was accompanied by a limited increase in MHC II expression.

Interestingly, the LN-associated CD8int DC subset expressed higher levels of PrP than CD8+ and CD8– DCs, as well as higher MHC II levels. In contrast to other DC subsets found in lymphoid organs, CD8int DCs have been demonstrated to represent fully mature DCs, with high levels of membrane MHC II and costimulatory molecules (33). As a result of their maturation status, overnight culture did not promote a significant up-regulation of PrP or MHC II by CD8int DCs.

Finally, B220+ pDCs did not express significant levels of PrP irrespective of their MHC II expression and location, and interestingly, overnight culture proved unable to induce PrP expression on pDCs.

In conclusion, PrP expression appears to be restricted to conventional, nonplasmacytoid, DC subpopulations, and among these, the LN-associated CD8int DC subset displayed the highest level of expression of PrP, in correlation with its maturation state. In addition, overnight culture, which induced a marked increase in the expression of MHC II, determined a strong up-regulation of PrP by skin DCs and, to a lesser extent, by CD8– DCs. Globally, these data suggest that PrP expression level correlates with the maturation status of conventional DC subpopulations.

To confirm this hypothesis, PrP expression was further analyzed on splenic non-pDCs after in vivo maturation induced by LPS, a TLR-4 ligand known to induce conventional DC activation and maturation (Fig. 2). Six hours after i.v. injection of LPS, both CD8– and CD8+ DCs displayed a mature phenotype, as indicated by the up-regulation of MHC II, CD40, and CD86 molecules. After LPS treatment, PrP expression was only slightly increased on CD8+ DCs, but was strongly up-regulated on the CD8– DC subset, as observed after overnight culture of these splenic DC subsets (see Fig. 1). These data further support the view that PrP expression is regulated during conventional DC maturation, this process being differentially regulated in different DC subsets.


Figure 2
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  		FIGURE 2. Effect of in vivo LPS-induced DC maturation on PrP expression. Histograms illustrate the expression of MHC II, costimulatory molecules CD40 and CD86, and PrP by splenic CD8+ and CD8– DCs 6 h before and 6 h after i.v. injection of LPS. Open profiles in upper histograms represent biotin-conjugated isotype control Ab staining. Percentages of cells above background staining are indicated. Expression profiles are represented on a logarithmic scale in the x-axis. Results are representative of two experiments with similar results.

 
PrP protein expression by DCs differentiated in vitro

To obtain further insights into the regulation of PrP expression by mouse DCs, DCs derived from bone marrow precursors driven by the cytokine Flt3L were analyzed for the expression of MHC II, CD40, CD86, and PrP. Interestingly, in contrast to DC cultures generated from bone marrow precursors in the presence of GM-CSF that generate non-pDCs, Flt3L-driven bone marrow cultures have been shown to generate both conventional B220– DCs (equivalent to splenic conventional DCs) and B220+ pDCs (34). Therefore, Flt3L-driven DC cultures represent a more physiological approach for the in vitro analysis of DC subsets present in lymphoid organs, considering that monocyte-derived DCs have been claimed to correspond to inflammatory DCs.

Analysis of DCs from day 10 Flt3L-driven cultures confirmed that, concurring with our data on ex vivo splenic B220+ pDCs, in vitro generated B220+ pDCs did not express PrP, nor did CpG treatment (that caused B220+ pDCs maturation) induce the expression of PrP on these cells (Fig. 3). With regard to in vitro generated B220– DCs, ~30% of them expressed PrP, and in agreement with the data from ex vivo isolated conventional DCs, treatment with the maturation stimuli LPS or CpG promoted a strong PrP up-regulation, paralleled by a significant increase in the expression of MHC II, CD40, and CD86. Cultures in strictly similar conditions, but without TLR ligands, ruled out the possibility that other factors would increase PrP levels.


Figure 3
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  		FIGURE 3. PrP expression by DCs generated in vitro from bone marrow precursors. Histograms show the expression of MHC II, costimulatory molecules, and PrP by conventional B220– DCs and B220+ pDCs derived from Flt3L-driven bone marrow cultures, before and after LPS- or CpG-induced maturation. Percentages of cells above background staining are indicated. Expression profiles are represented on a logarithmic scale in the x-axis. Results are representative of four experiments with similar results.

 
These data further support the concept that PrP expression by conventional, non-pDCs correlates with the maturation process. To reinforce this concept, PrP expression was analyzed on conventional B220– DCs from Flt3L-driven DC cultures after maturation induced by the TLR-9 ligand CpG, in the presence of an inhibitor of p38 MAPK (SB203508), which is involved in the TLR-9 signaling pathway leading to the production of IL-12 by DCs. As shown in Fig. 4, treatment with SB203508 blocked CpG-induced IL-12 production by in vitro generated B220– DCs, and inhibited the up-regulation of PrP, strongly suggesting that the latter was related to TLR-9 triggering. In contrast, treatment with UO126, an inhibitor of ERK kinase, acted synergistically with CpG on the induction of PrP expression and IL-12 production, as expected, because ERK has been proposed to exert an inhibitory effect on the p38 signaling pathway initiated after TLR engagement.


Figure 4
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  		FIGURE 4. Effect of p38 MAPK inhibition on CpG-induced PrP up-regulation. PrP expression and IL-12 production after CpG-induced maturation by conventional B220– DCs, derived from Flt3L-driven bone marrow cultures, in the presence of the p38 MAPK kinase inhibitor SB203508, or the ERK kinase inhibitor UO126. Results are representative of three experiments with similar results.

 
Globally, B220– DC activation and maturation induced by TLR-4 and TLR-9 ligands resulted in the up-regulation of PrP.

Analysis of MHC II and costimulatory molecule expression in PrP-deficient mice

The finding that PrP expression by conventional B220– DCs was correlated with their maturation status prompted us to investigate whether PrP could in turn be involved in the regulation of the expression of MHC II and costimulatory molecules. For this purpose, ex vivo isolated, as well as in vitro differentiated DCs from PrP-deficient (Prnp–/–) mice were analyzed and compared with those obtained from wild-type C57BL/6 control mice. No significant difference was found between survivals of cultured Prnp–/– and wild-type DCs.

As shown in Fig. 5, a slightly higher proportion of CD11c+ spleen DCs from Prnp–/– mice was CD8{alpha} positive as compared with wild-type C57BL/6 mice. A further comparison of six individual mice of each group confirmed that the proportions and absolute numbers of splenic CD8+ DCs were higher in Prnp–/– mice. The mean percentage of CD8+ among CD11c+ B220– spleen cells was 29.9 (SD = 8.8) in control C57BL/6, vs 44.0 (SD = 13.3) in Prnp–/– mice (p < 0.05, Mann-Whitney U test). No significant differences were observed in the expression of MHC II, CD40, and CD86 by ex vivo isolated CD8– or CD8+ splenic DCs, before or after overnight culture (Fig. 5).


Figure 5
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  		FIGURE 5. MHC II and costimulatory molecule expression by ex vivo isolated DCs from PrP-deficient mice. Histograms show the proportion of CD8+ DCs among spleen DCs in control C57BL/6 and PrP-deficient mice (Prnp–/–) in one representative experiment of six, and the expression level of MHC II and costimulatory molecules CD40 and CD86 by CD8+ and CD8– splenic DCs directly after ex vivo isolation, or after overnight culture. Open profiles represent biotin-conjugated isotype control Ab staining. The percentage of cells above background staining is indicated. Expression profiles are represented on a logarithmic scale in the x-axis. Results are representative of three experiments with similar results.

 
Similarly, when conventional B220– DCs generated in Flt3L-driven bone marrow cultures from Prnp–/– mice were compared with those derived from C57BL/6 control mice, no significant differences in the expression of MHC II, CD40, and CD86 were observed, neither before nor after incubation with the maturation stimuli LPS or CpG (Fig. 6).


Figure 6
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  		FIGURE 6. LPS- or CpG-induced maturation of DCs differentiated in vitro from wild-type or PrP-deficient mice. Histograms show the expression of MHC II and costimulatory molecules CD40 and CD86 by conventional B220– DCs, derived from Flt3L-driven bone marrow from C57BL/6 or PrP-deficient mice, before and after LPS- or CpG-induced maturation. The percentage of cells above background staining is indicated. Expression profiles are represented on a logarithmic scale in the x-axis. Results are representative of three experiments with similar results.

 
Therefore, PrP did not appear to play a role in the regulation of the expression of MHC II and costimulatory molecules during DC maturation, but may influence the development of CD8– DCs.


   Discussion
Top
 Abstract
 Introduction
 Materials and Methods
 Results
 Discussion
Disclosures
 References
 
DCs are essential elements of the immune system due to their capacity to induce and regulate Ag-specific immune responses, allowing them to control infections caused by microbial pathogens; to block tumor growth; to exert a complex regulation of T, B, and NK cell responses; and, in addition, to play a pivotal role in the induction and maintenance of T cell tolerance. The remarkable functional diversity of the DC system relies on the plasticity and complexity of the DC differentiation process, which leads to the generation of a large collection of DC subpopulations, endowed with specific functions. DCs can be classified in two major categories: conventional DCs, which in turn include different DC subsets, and pDCs, characterized by their capacity to produce large amounts of type I IFNs during viral infections.

In the present study, we have investigated the expression of PrP by different DC subpopulations endowed with specific phenotypic and functional characteristics. In particular, we have analyzed DCs isolated from different tissues (skin, thymus, spleen, and LNs), before and after overnight culture, or after in vivo induction of DC maturation by LPS treatment. PrP expression was also analyzed on DCs differentiated in vitro from bone marrow precursors in the presence of the cytokine Flt3L, to extend our study to in vitro generated DCs, extensively used both as research tools and for immunotherapeutical purposes.

Previous studies have shown that, outside the nervous system, DCs display the highest expression levels of PrPC, in both humans and mice (7, 17, 18). PrPC was found to be present on murine epidermis LCs (35), on DCs in extrafollicular areas, including T cell zones, of the gut mucosa (7, 36), and DCs of the splenic white pulp (18). PrP was found on the surface of bone marrow-derived human and mouse DCs generated in vitro in the presence of GM-CSF, at levels that increased with LPS stimulation and correlated with that of MHC II and costimulatory molecule CD86 (18, 31). The present study confirms and extends these results, by demonstrating important differences between different DC subsets either analyzed after ex vivo isolation or differentiated in vitro.

A first striking observation is the absence of PrP expression on pDCs, even after maturation, in contrast to conventional DCs. pDCs are featured by their ability to secrete type I IFNs in response to viral infections. Neither ex vivo isolated, nor in vitro differentiated B220+ pDCs expressed detectable PrP, even after overnight culture or stimulation with CpG. This difference with conventional DCs could relate to their developmental origin because pDCs have been proposed to derive, under physiological conditions, from bone marrow lymphoid precursors, based on data obtained both in humans and mice, demonstrating that pDCs express molecules related to the lymphoid lineage, such as pT{alpha}, spi-B, IL-7R, and PIII CIITA, and have IgH gene rearrangements (reviewed in Ref. 37). Whether pDCs can express PrP during in vivo viral infections remain to be determined.

On conventional DCs, PrPC expression was strongly up-regulated after maturation by TLR ligands, such as bacterial LPS and CpG-oligodeoxynucleotides, in parallel with molecules involved in Ag presentation and T cell activation. Interestingly, a recent study showed that membrane PrP on Ag-presenting DCs enhances the stimulation of specific naive T cells, both in vitro and in vivo (31).

On the membrane of T cells, PrPC molecules were found to associate with the TCR/CD3 complex upon engagement with MHC II/Ag peptide or anti-CD3 Abs, as shown by confocal microscopy and coimmunoprecipitation (31, 38). Similar colocalization of PrPC and CD3{epsilon} was demonstrated on Jurkat T cells after a nonspecific stimulation through hypothermal treatment, mimicking an immunological synapse-like structure (39). In contrast, activation of T cells by either mitogens or anti-CD3 Ab enhances their PrPC expression (16, 17), and human T cells with an activated phenotype, based on their CD56 expression, display higher PrPC levels than their resting counterparts (15). Altogether, these data suggest that T cell activation may implicate PrPC on the surface of both partners, i.e., the APC and the T cell.

Higher PrPC levels were found on the surface of spleen DCs that are also CD8{alpha}+. Expression of this marker has been correlated with the secretion of IL-12 and IFN-{alpha} (40, 41). In LNs, the highest PrPC levels are displayed by the CD8int subset that was shown to originate from skin DCs, and which are strong stimulators of Ag-dependent delayed-type hypersensitivity (42). Thus, our data point to a possible involvement of PrPC in T cell activation leading to Th1 responses. The mechanisms involved in T cell activation related to PrPC expressed on DC remain to be explored.

Besides the contribution of our results in clarifying PrPC functions in the immune system, they may help in understanding certain aspects of TSE pathogenesis. Indeed, expression of PrPC by cells of the lymphoreticular system is required for prions to propagate and invade the CNS after peripheral inoculation (23). This may relate to a possible role of membrane PrPC as a receptor for prions, and to the ability of replicating prions at the periphery through PrP neosynthesis. Immunohistochemical studies revealed expression of PrPC by several cell types of the skin, including LCs and DCs, suggesting that they could be initial targets for prion infection (35). Upon activation and maturation, peripheral DCs increase PrP expression, which could help prion replication. Indeed, we found that highest PrPC levels in LN DCs are displayed by a CD8int subset that originates from the skin. In contrast, recent studies suggest that, although DCs can capture prions shortly after oral inoculation and migrate to draining LNs (26), their role in early pathogenesis might not be essential (43) (our unpublished personal data). However, the increased PrP expression by mature DCs present in secondary lymphoid organs might facilitate their involvement in the transfer of prions to sites of neuroinvasion.


   Acknowledgments
 
We thank Jacques Grassi for the gift of SAF83 Ab, Charles Weissmann for Prnp–/– breeders, and Claude Carnaud and Martine Bruley Rosset for helpful discussions and critical reading of the manuscript.


   Disclosures
Top
 Abstract
 Introduction
 Materials and Methods
 Results
 Discussion
 Disclosures
References
 
The authors have no financial conflict of interest.


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

1 This work was supported in part by grants from the Ministerio de Educación y Ciencia of Spain SAF-2003-07291, Groupement d’Intéret Scientifique Maladies à Prions, and European Union Contract QLK5-CT-2002-01044. P.M. was a recipient of an Institut National de la Santé et de la Recherche Médicale Poste Vert fellowship. Back

2 Current address: Centre for Research in Vascular Biology, BioSciences Institute, University College, Cork, Ireland. Back

3 C.A. and P.A. contributed equally to this work. Back

4 Address correspondence and reprint requests to Dr. Pierre Aucouturier, Université Pierre et Marie Curie, Unité Mixte de Recherche S Institut National de la Santé et de la Recherche Médicale Unité 712, 184 rue du Faubourg Saint-Antoine, 75571, Paris Cedex 12, France; E-mail address: aucoutur{at}st-antoine.inserm.fr or Dr. Carlos Ardavín, Department of Immunology and Oncology, Centro Nacional de Biotecnología/Consejo Superior de Investigaciones Cientificas, Universidad Autónoma, 28049 Madrid, Spain; E-mail address: ardavin{at}cnb.uam.es Back

5 Abbreviations used in this paper: PrP, prion protein; DC, dendritic cell; DDC, dermal DC; int, intermediate; LC, Langerhans cell; LN, lymph node; MS-LN, mesenteric LN; pDC, plasmacytoid DC; PrPC, cellular PrP; Flt3L, Fms-like tyrosine kinase 3 ligand; TC, tricolor; TSE, transmissible spongiform encephalopathy. Back

Received for publication May 16, 2006. Accepted for publication August 2, 2006.


   References
Top
 Abstract
 Introduction
 Materials and Methods
 Results
 Discussion
 Disclosures
 References
 

  1. Bolton, D. C., M. P. McKinley, S. B. Prusiner. 1982. Identification of a protein that purifies with the scrapie prion. Science 218: 1309-1311. [Abstract/Free Full Text]
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