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Originally published online as doi:10.1189/jlb.1102521 on May 22, 2003

Published online before print May 22, 2003
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(Journal of Leukocyte Biology. 2003;74:118-125.)
© 2003 by Society for Leukocyte Biology

Activation by prion peptide PrP106–126 induces a NF-{kappa}B-driven proinflammatory response in human monocyte-derived dendritic cells

Silvia M. Bacot*, Petra Lenz{dagger}, Michelle R. Frazier-Jessen* and Gerald M. Feldman*

* Division of Monoclonal Antibodies, Center for Biologics Evaluation and Research, Food and Drug Administration, Bethesda, Maryland; and
{dagger} Laboratory of Cellular Oncology, National Cancer Institute, National Institutes of Health, Bethesda, Maryland

Correspondence: Gerald M. Feldman, Ph.D., Food and Drug Administration, HFB-564, Bldg. 29A, Rm. 3C24, 29 Lincoln Drive, Bethesda, MD 20892. E-mail: feldman{at}cber.fda.gov


   ABSTRACT
TOP
 ABSTRACT
INTRODUCTION
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 REFERENCES
 
Specific prion peptides have been shown to mimic the pathologic isoform of the prion protein (PrP) and to induce a neurotoxic effect in vitro and in vivo. As monocytic cells are thought to play a role in the transmission and pathogenesis of prion disease, the use of these peptides in regulating monocytic cell function is under intense investigation. In the current study, we characterize the ability of prion peptide PrP106–126 to activate specific signaling pathways in human monocyte-derived dendritic cells (DCs). Electrophoretic mobility shift assays establish the activation of transcription factor nuclear factor-{kappa}B within 15 min of exposure, with as little as 25 µM peptide. This signaling cascade results in the up-regulation of inflammatory cytokines interleukin (IL)-1ß, IL-6, and tumor necrosis factor {alpha} (TNF-{alpha}) at the mRNA and protein levels. Phenotypic activation of DCs exposed to PrP106–126 is partly a result of an autocrine TNF-{alpha} response and results in an increased ability of these cells to induce lymphocyte proliferation. The effects of PrP106–126 on DCs were elicited through a receptor complex distinct from that used by human monocytes, demonstrating the ability of this peptide to interact with a multiplicity of receptors on various cell types. Together, these data suggest an involvement of DCs in prion disease pathogenesis.

Key Words: TSE • cell signaling • inflammation


   INTRODUCTION
TOP
 ABSTRACT
 INTRODUCTION
MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 REFERENCES
 
Prion diseases, or transmissible spongiform encephalopathies (TSEs), are uniformly fatal, neurodegenerative disorders found in animals and humans. In these diseases, the normal cell-surface prion glycoprotein (PrPC) is post-translationally modified into a protease-resistant isoform known as scrapie prion protein (PrPsc) [1 ]. In late stages of TSE infection, abnormal PrP (PrPsc) accumulates in the brain, where it is thought to play a direct and indirect role in disease pathogenesis. It has been proposed that the naturally occurring isoform of PrP may play an important role in normal cellular function, including lymphocyte activation [2 ], and synaptic function [3 ]. The presence of PrPC is crucial for the generation of PrPsc, serving as a template for conformational change [4 ]. Although PrPC is predominantly expressed in the brain, it is also found in a wide variety of peripheral tissues, including mononuclear blood cells [2 ]. The prevalence of monocytic cells in tissues affected by prions suggests a possible role in the pathogenesis of prion diseases. In this regard, dendritic cells (DCs) are thought to play a critical role in prion infectivity and translocation [5 , 6 ]. The presence of CD11c+DCs in brains of affected animals is thought to be a result of a combination of the maturation and differentiation of resident microglial cells [7 , 8 ] as well as the recruitment of monocyte-derived DCs to the lesion [9 10 11 ].

Analysis of the human TSEs has revealed the presence of amyloid fibril deposits consisting of insoluble cleavage products from the protease-resistant PrPsc. These peptide fragments, which can reach high concentration levels at their surface interface, have been sequenced and have been shown to consist of a mixture of fragments with amino acid residues mostly between 100 and 140 [12 ]. A synthetic peptide based on this sequence (human PrP90–144) caused neuronal dysfunction and neuropathological changes in transgenic mouse hosts upon intracerebral inoculation [13 ], thereby mimicking the pathological effects of PrPsc. A related peptide consisting of amino acids 106–126 has also been shown to form amyloid-like fibrils in vitro and to display neurotoxic activity toward primary cultures of rat hippocampal neurons [14 ]. PrP106–126 has also been shown to induce nitric oxide synthase (NOS) expression in microglial cells [15 ] and to stimulate a chemotactic response, induction of calcium mobilization, and production of reactive oxygen intermediates in human monocytes [16 , 17 ]. However, the direct effects of prion peptides on monocyte-derived DCs have not been explored, and an analysis of this response is crucial to fully understand the pathogenesis of prion disorders. The current study examines the response of monocyte-derived DCs to PrP106–126 stimulation. We describe a dose-dependent activation of myeloid DCs as determined by a rapid and specific activation of the nuclear factor (NF)-{kappa}B signaling cascade, concomitant with phenotypic activation and an induction of inflammatory cytokines, suggesting the involvement of DCs in the prion disease process.


   MATERIALS AND METHODS
TOP
 ABSTRACT
 INTRODUCTION
 MATERIALS AND METHODS
RESULTS
 DISCUSSION
 REFERENCES
 
Cell culture
Human peripheral blood monocytes were obtained from healthy volunteers by leukapheresis. Monocytes were purified from mononuclear cells by ficoll-hypaque sedimentation followed by countercurrent centrifugal elutriation [18 , 19 ]. Monocytes (>95% CD14+) were resuspended at a density of 1.5–2.0 x 106 cells per ml in Dulbecco’s modified Eagle’s medium (DMEM; Life Technologies, Rockville, MD) containing 10% fetal bovine serum (BioWhittaker, Walkersville, MD), 100 ng/ml recombinant human (rh) granulocyte macrophage-colony stimulating factor (generously provided by Immunex, Seattle, WA), and 100 ng/ml rh interleukin (IL)-4 (generously provided by Schering Plough, Kenilworth, NJ) and were cultured for a minimum of 6 days to allow for differentiation into DCs. A synthetic peptide corresponding to amino acid residues 106–126 of the PrP (PrP106–126), known to contain biological activity [14 , 20 ], was synthesized on an ABI model 433A synergy peptide synthesizer at a 0.1-mmole scale (Applied Biosystems, Foster City, CA) and was used in the stimulation of DCs. A scrambled prion peptide (scrPrP) was used as a negative control. Peptides were dissolved in DMEM and used at concentrations ranging between 10 and 250 µM. Endotoxin levels in the dissolved peptides were undetectable. Lipopolysaccharide (LPS; Sigma Chemical Co., St. Louis, MO) was used at 100 ng/ml.

Preparation of nuclear extracts and electrophoretic mobility shift assays (EMSAs)
After stimulation, cells were washed in cold phosphate-buffered saline, and further reactions were performed at 4°C. Cells were incubated in a hypotonic buffer [10 mM HEPES, pH 7.9, 10 mM KCl, 0.1 mM EDTA, 0.1 mM EGTA, 1 mM sodium orthovanadate, 10 mM ß-glycerophosphate, 1 mM dithiothreitol (DTT), 1 mM 4-(2-aminoethyl) benzenesulfonyl fluoride, and a cocktail of protease inhibitors (Boehringer Mannheim, Mannheim, Germany) followed by lysis with Triton-X 100 (Pierce, Rockford, IL). After removal of cytoplasmic components by centrifugation, nuclear extracts were harvested from the pellet by incubation at 4°C in a high salt buffer (20 mM HEPES, pH 7.9, and 400 mM NaCl in above buffer). Protein concentrations for each extract were determined, and equal amounts of protein were assayed for DNA-binding proteins by EMSA as described previously [21 , 22 ]. Briefly, 5 µg protein was incubated with the 32P-labeled oligonucleotide probe consisting of a double-stranded sequence for the specific signal transducer and activator of transcription (STAT) or NF-{kappa}B consensus site (Promega, Madison, WI) in binding buffer [10 mM Tris, pH 7.5, 100 mM NaCl, 1 mM DTT, 5 mM EDTA, 4% (v/v) glycerol, and 80 µg/ml sonicated salmon sperm DNA]. The sample was then applied to a 6% nondissociating polyacrylamide gel to separate free probe from probe bound to protein. In some experiments, 25 µM of the serine protease inhibitor tosyl-Phe-chloromethylketone (TPCK; Boehringer Mannheim) was used during cell culture to block the degradation of I{kappa}B.

RNase protection assay (RPA)
Total RNA was extracted from PrP106–126-treated DCs using RNAqueous (Ambion, Austin, TX), and 10 µg RNA was used for the RPA. 32P-labeled RNA multitemplate probes (hCK-2 and hCK-3 kits, PharMingen, San Diego, CA) were synthesized per the manufacturer’s directions and allowed to hybridize overnight with the isolated RNA. After RNase digestion with the RNases A and T1 (Ambion), protected fragments were denatured and electrophoresed on a 7-M urea 8% polyacrylamide sequencing gel. The gels were then exposed to Kodak XAR film at -70° overnight.

Fluorescein-activated cell sorter (FACS) analysis
On day 7 of culture, DCs were harvested and resuspended in ice-cold Hanks’ balanced salt solution (HBSS; Life Technologies) containing 0.1% bovine serum albumin (BSA; Life Technologies) and 0.1% sodium azide (Aldrich Chemical Co., Milwaukee, WI). To avoid nonspecific binding of labeled antibody, FcBlock (PharMingen) was added at 1 µg per 1 million cells. Cells were double-stained with monoclonal antibodies (mAb) raised against the following human surface antigens (all antibodies obtained from PharMingen): phycoerythrein (PE)-conjugated CD14 [M5E2, mouse immunoglobulin G (IgG)2a], PE-CD11c (B-ly6, mouse IgG1), PE-CD80 (L307.4, mouse IgG1), fluorescein isothyocyanate (FITC)-conjugated human leukocyte antigen (HLA)-DR (G46-6, mouse IgG2a), FITC-CD40 (5C3, mouse IgG1), FITC-CD83 (HB15e, mouse IgG1), and appropriate isotype-control antibodies. Cells (105 per assay) were incubated with the respective antibody (2.5 µg per sample) for 30–45 min at 4°C before washing. To detect intracellular cytokine production, cells were exposed to various stimuli as indicated, and 1 h later, Brefeldin A (GolgiPlug, PharMingen) was added for an additional 5 h. Cells were then surface-stained with PE-CD11c. Subsequently, cells were submitted to fixation and permeabilization using a Cytofix/Cytoperm kit (PharMingen) according to the manufacturer’s instructions and were stained with FITC-labeled mouse anti-human mAb against IL-1ß (IgG1), tumor necrosis factor {alpha} (TNF-{alpha}; IgG1), IL-6 (IgG2b), and respective isotype controls (all R&D Systems, Minneapolis, MN). Cellular fluorescence was monitored in a FACSCalibur® cell sorter (Becton Dickinson, Mansfield, MA) and analyzed using the CellQuest software provided by the manufacturer. DCs, gated by forward- and side-scatter, expressed high levels of CD11c but were negative for CD14 (data not shown). For some studies, DCs were pretreated with or without 50 ng/ml of EnbrelTM, a soluble TNF receptor used to inhibit the function of TNF (Immunex) [23 24 25 26 27 ].

Mixed lymphocyte reaction (MLR)
DCs were treated with PrP (25 µM), scrPrP (25 µM), or LPS (100 ng/ml) overnight. Cells were then irradiated at 3000 rad, and increasing numbers of {gamma}-irradiated DCs were added in triplicate to 105 monocyte-depleted, allogeneic peripheral blood lymphocytes in flat-bottomed, 96-well plates, as described previously [28 ]. Lymphocyte proliferation was measured 5 days later according to the spectrophotometric method of Mosmann [29 ], using the 3-(4,5-dimethylthiazol-2-yl)-2,5-diphenyl tetrazolium bromide (MTT)-based CellTiter 96® kit obtained from Promega. Absorbance was read at 490 nm, per the manufacturer’s instructions.

Calcium mobilization analysis
For calcium mobilization studies, day 7 DCs were harvested and resuspended in ice-cold HBSS (Life Technologies) containing 0.1% BSA (Life Technologies) at a concentration of 1–2 x 106 cells/ml. Cells were loaded with dye by incubating in 0.1% HBSS containing 2.5 µM Fura-2/AM at 37°C for 30 min in the dark, followed by a series of washes, and resuspended in 2 ml 0.1% HBSS. Cells were monitored for intracellular Ca2+ every 200 ms at 510 nm in a continuously stirred cuvette at 37°C in a model MS-III fluorimeter (Photon Technology, South Brunswick, NJ) after stimulation with formyl-Met-Leu-Phe (fMLP; Sigma Chemical Co.) or PrP106–126. Data are expressed as the relative ratio of fluorescence emitted at 510 nm following sequential excitation at 340 and 380 nm [30 , 31 ].


   RESULTS
TOP
 ABSTRACT
 INTRODUCTION
 MATERIALS AND METHODS
 RESULTS
DISCUSSION
 REFERENCES
 
NF-{kappa}B activation by PrP106–126
To determine if PrP106–126 activates DCs and to ascertain a possible signaling mechanism whereby this prion peptide exerts its effects, nuclear extracts were prepared from PrP106–126-treated DCs at different times and subsequently analyzed for the activation of potential transcriptional activators including STATs 1–6 and NF-{kappa}B via EMSAs. As demonstrated in Figure 1 , stimulation with 25 µM PrP106–126 resulted in the activation of the NF-{kappa}B signaling pathway. This activation occurred within 15-min post-stimulus, peaked around 45 min, and slightly decreased in signal intensity thereafter up to 180 min. In contrast, no activation of other signaling pathways was observed in response to PrP106–126 (data not shown). DCs treated with PrP106–126 for 30 min displayed a dose-dependent increase in NF-{kappa}B activation through 100 µM, and a maximum signal occurred at 50 µM (Fig. 2 ). In contrast, scrPrP had no effect on NF-{kappa}B nuclear translocation (Fig. 2) .



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  		Figure 1. Time course of PrP106–126-induced activation of human monocyte-derived DCs via the NF-{kappa}B pathway. Nuclear extracts were isolated from DCs following exposure to a 25-µM concentration of PrP106–126 for the indicated time. Equal amounts of nuclear protein were allowed to hybridize with labeled probe, and EMSAs performed to analyze the time course of NF-{kappa}B induction. A 15-min stimulus with LPS (100 ng/ml) was used as a positive control. After separation of the bound from free probe by electrophoresis, the dried gel was exposed to Kodak XAR film at -70°C using intensifying screens. Arrows denote DNA–protein complexes specific for the p50/p65 NF-{kappa}B homo- and heterodimers and unbound, excess free probe.

 


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  		Figure 2. PrP106–126 activates human monocyte-derived DCs via the NF-{kappa}B pathway in a dose-dependent manner. Nuclear extracts were isolated from DCs following a 30-min exposure to the indicated concentrations of PrP106–126. Equal amounts of nuclear protein were allowed to hybridize with labeled probe, and EMSAs performed to analyze the ability of PrP106–126 to induce NF-{kappa}B binding in a dose-dependent manner. Arrows denote DNA–protein complexes specific for the p50/p65 NF-{kappa}B homo- and heterodimers and unbound, excess free probe. scrPrP was used as a negative control.

 
For NF-{kappa}B to elicit its signal, it must translocate to the nucleus, and to do so, an inhibitory protein, I{kappa}B, must first be phosphorylated and subsequently degraded. To determine that PrP106–126 activation of the NF-{kappa}B signaling pathway was a result of the degradation of I{kappa}B, DCs were pretreated with the serine protease inhibitor, TPCK, for 1 h before exposure to PrP106–126. Pretreatment with TPCK prevented the induction of the NF-{kappa}B signaling pathway by PrP106–126 (Fig. 3 ), suggesting that the pathway elicited by this peptide causes the degradation of I{kappa}B.



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  		Figure 3. PrP106–126-induced activation of human monocyte-derived DCs is dependent on I{kappa}B degradation. DCs were treated with or without the serine protease inhibitor TPCK (50 µM) for 1 h before a 30-min stimulation with 25 µM PrP106–126. Nuclear extracts were isolated, and equal amounts of nuclear protein were allowed to hybridize with labeled probe before separation by nondenaturing gel electrophoresis. Arrows denote DNA–protein complexes specific for the p50/p65 NF-{kappa}B homo- and heterodimers and unbound, excess free probe.

 
Induction of inflammatory cytokine production by PrP106–126
To investigate the role of NF-{kappa}B activation in relation to gene transcription, inflammatory cytokine mRNA levels were measured by RPA analysis of DCs treated with 25 µM PrP106–126 for up to 16 h. Analysis of the protected fragments demonstrates the ability of the PrP peptide to up-regulate the gene expression of inflammatory cytokines such as IL-1ß, IL-6, and TNF-{alpha} in a time-dependent manner (Fig. 4 ). As before, the scrambled version of the peptide had no stimulatory effect on DC mRNA expression (data not shown). Analysis of these data by phosphor imaging and controlling for the level of RNA demonstrate that PrP106–126 induced a threefold increase in steady-state mRNA levels of the IL-1ß gene within 4 h of stimulation, remaining elevated through 8 h but decaying by 50% by 16 h. Similar kinetics were observed for the genes for TNF-{alpha} and IL-6. Increases in inflammatory cytokines were also observed at the protein level, as determined by FACS staining for intracellular cytokines (Fig. 5 ). Cells treated with PrP106–126 expressed intracellular IL-1ß at a 55-fold higher level compared with untreated cells (P<0.01), tenfold increases were observed for IL-6 and TNF-{alpha} (P<0.01), and a 15-fold increase was observed for IL-12 (P<0.01). The scrambled version of the peptide had no such stimulatory effect on intracellular cytokine expression (Fig. 5) .



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  		Figure 4. RNase protection analysis of cytokine mRNA induction in DCs. Total RNA was extracted from 1.5–2.0 x 107 DCs that were treated with or without PrP106–126 for the indicated periods of time. RNA (5 µg) was allowed to hybridize with the labeled antisense probes before digestion as described in Materials and Methods and was analyzed by RPA. Arrows denote the protected mRNA fragments for TNF-{alpha}, IL-1ß, IL-6, and glyceraldehyde 3-phosphate dehydrogenase (GAPDH).

 


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  		Figure 5. PrP106–126 induces production of inflammatory cytokines in monocyte-derived DCs. Day 7 monocyte-derived DCs were exposed to PrP106–126 (25 µM), scrPrP (25 µM), LPS (100 ng/ml), or were left untreated. Five hours after addition of Brefeldin A, intracellular cytokines IL-1ß, TNF-{alpha}, IL-6, and IL-12 were quantified by flow cytometry as described in Materials and Methods. Dot plots show CD11c expression in FL2 and the respective cytokines in FL1. Numbers represent the percentage of cytokine-producing DCs.

 
Phenotypic activation of DCs by PrP106–126
DCs derived from human monocytes were cultured for 24 h in the presence or absence of PrP106–126 or its scrambled control (scrPrP) and were analyzed for maturation indicators by FACS analysis. PrP106–126 (25 µM) exposure resulted in an up-regulation of major histocompatibility complex (MHC) class II (HLA-DR), CD40, CD80, and CD83 surface expression (Fig. 6 ). In contrast, addition of an equivalent amount of scrPrP did not lead to changes in the surface expression of any of the costimulatory and activation markers tested (Fig. 6) . This phenotypic activation was dose-dependent, although considerable decreases in cell viability were observed at PrP106–126 concentrations approaching 100 µM. To determine if the phenotyic activation of DCs is a result of the direct effects of PrP106–126 or is a result of an indirect activation via the induction of TNF-{alpha}, DCs were treated with or without a soluble TNF receptor (EnbrelTM), which binds TNF-{alpha} with high-affinity [23 24 25 26 27 ]. EnbrelTM pretreatment exhibited a partial reduction in DC maturation in response to PrP106–126 (data not shown), suggesting that TNF-{alpha} induction is partly, although not entirely, responsible for the resulting phenotypic activation in these cells. LPS-induced DC activation was also only partially inhibited with EnbrelTM, whereas TNF-{alpha}-induced DC activation was completely inhibited by EnbrelTM, demonstrating its specificity (data not shown).



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  		Figure 6. Surface expression of differentiation markers on monocyte-derived DCs. Day 6 monocyte-derived DCs were exposed to PrP106–126 (final concentration, 25 µM), and 24 h later, surface expression of HLA-DR, CD40, CD80, CD86, and CD83 was determined by flow cytometry. Untreated cells were used to determine background expression, and scrPrP (25 µM) and LPS (100 ng/ml) served as negative and positive controls, respectively. Numbers in the upper left represent mean fluorescence intensity of the histograms; the numbers in the lower right show the percentage of CD83+ cells. Exposure to PrP106–126 induced expression of MHC class II (HLA-DR), costimulatory molecules (CD40/CD80), and activation marker CD83. Shown are the results of one representative experiment out of a minimum of four experiments.

 
Induction of MLR
PrP106–126-induced changes in DC maturation and activation translated to a functional differentiation, as demonstrated by an increase in allogeneic lymphocyte proliferation (Fig. 7 ). This increase in lymphocyte proliferation closely resembled that observed when DCs are activated with LPS. The addition of scrPrP had no effect on DC-stimulated lymphocyte proliferation (data not shown).



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  		Figure 7. Allogeneic lymphocyte proliferation induced by matured DCs, which were allowed to mature overnight in the absence (Co) or presence of scrPrP106–126 (25 µM), PrP (25 µM), or LPS (100 ng/ml). DCs were then irradiated and co-plated with 1 x 105 lymphocytes per well at the indicated DC:lymphocyte ratio and were incubated at 37°C in 96-well plates for an additional 5 days. Lymphocyte proliferation, as assessed by MTT absorbance, is provided as a function of the DC:lymphocyte ratio. The data represent the mean ± SE of four separate experiments performed in triplicate.

 
PrP106–126 receptor analysis in DCs
In other studies involving monocytes, a specific G protein-coupled receptor (GPCR) formyl peptide receptor-like 1 (FPRL1) was identified as being necessary and sufficient for PrP106–126-induced cell activation [17 ]. To determine if this or related receptor complexes play a role in the PrP106–126–induced activation of DCs, Ca2+ flux studies were performed with day 7 DCs and were compared with freshly elutriated monocytes. Whereas monocytes give a strong, intracellular Ca2+ mobilization when stimulated with fMLP or PrP106–126 (ref. [17 ], data not shown), DCs stimulated with PrP106–126 did not undergo an influx of Ca2+ (Fig. 8 ). In contrast, stimulation of DCs with fMLP induced an immediate and strong intracellular Ca2+ mobilization (Fig. 8) . Similar to what has been observed in monocytes, this response was completely inhibitable with pertussis toxin (PTX; data not shown).



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  		Figure 8. Calcium mobilization induced by PrP106–126 in DCs, which were loaded with Fura-2 and then monitored for changes in cell fluorescence as a reporter for intracellular [Ca2+]. PrP106–126 was used at a final concentration of 25 µM, and fMLP was used at a final concentration of 100 ng/ml. The arrows indicate the times of addition of each stimulus. Data are expressed as the relative ratio of fluorescence as a function of time and are representative of four independent experiments.

 

   DISCUSSION
TOP
 ABSTRACT
 INTRODUCTION
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
REFERENCES
 
The activation of monocyte-derived cells is thought to play a key role in the inflammatory process leading to the pathogenesis of many neurodegenerative diseases, including TSEs [32 33 34 35 36 ]. DCs have been shown to mature and become highly activated in response to several different mediators, including inflammatory cytokines [37 ], oligo CpG motifs [38 ], LPS and other bacterial products [39 ], and ds RNA [40 ]. In this study, we demonstrate the ability of PrP106–126 to activate the NF-{kappa}B signaling pathway in monocyte-derived DCs and through this pathway, to induce the production of inflammatory cytokines and the further differentiation and activation of DCs.

As a major transcriptional activator of inflammatory cytokines, NF-{kappa}B and the cytokines it is capable of activating have been previously linked to the presence and progression of prion disease [41 , 42 ]. Other in vitro studies have demonstrated the ability of PrP106–126 to activate the NF-{kappa}B signaling pathway in microglial cells and monocytes [15 ]. The latter study identifies the receptor with which PrP106–126 interacts as the GPCR FPRL1 [17 ]. In monocyte-derived DCs, however, this receptor has been shown to be down-regulated [43 44 45 ]. Indeed, our data indicate that a GPCR is specifically not involved, as PrP106–126 does not induce a Ca2+ flux in DCs. Furthermore, the effects of PrP106–126 on DCs could not be inhibited with PTX, a specific G protein antagonist (data not shown). Studies are currently underway to identify the receptor complex used by PrP106–126 in DCs.

The induction of cytokine gene and protein expression by PrP106–126-activated DCs is consistent with our current understanding of disease progression in neurological disorders including TSE, Parkinson’s disease, and Alzheimer’s disease [36 , 41 , 42 , 46 ]. Thus, up-regulation of IL-6, IL-1ß, and TNF-{alpha} mRNA expression has been detected in the brains of mice experimentally infected with Creutzfeldt-Jakob disease (CJD) or scrapie [41 ]. In addition, TNF-{alpha} mRNA expression was found to be elevated in human astrocytes and microglial cells when exposed in vitro to PrP106–126 [15 ]. Furthermore, PrP106–126 has been shown to induce the overexpression of the inflammatory cytokines IL-1ß and IL-6 [14 , 47 ] and reactive oxygen intermediates [48 ] in murine microglial cells and astrocytes. Similarly, PrP106–126 stimulated the induction of TNF-{alpha} and NOS through the mitogen-activated protein kinase signaling pathway in human fetal microglial cells [15 ].

We also demonstrate the ability of PrP106–126 to mature and activate DCs as determined by the increased expression of MHC class II (HLA-DR), costimulatory molecules CD40 and CD80, and the maturation marker CD83. These effects are partly a result of the upstream production and availability of TNF-{alpha} by PrP106–126 and are paralleled by an increase in the functional response of PrP106–126-stimulated DCs in terms of their enhanced ability to induce allogeneic lymphocyte proliferation. In this regard, the cellular expression of HLA-DR has been previously demonstrated to be up-regulated in mononuclear cells at lesion sites of some neurodegenerative disorders including Alzheimer’s disease and CJD [29 ].

Although follicular DCs (cells of stromal origin and unrelated to myeloid DCs) appear to support PrPsc replication [49 50 51 52 53 ], the route of transmission for prions from the periphery to the central nervous system (CNS) still remains obscure. Available data suggest that monocyte-derived DCs play a critical role in this process. Prion infectivity titer is found to be very high in CD11c+ DCs, much more so than in any other type of blood mononuclear cells [5 ]. Furthermore, these types of cells have been shown to acquire prion infectivity through the intestinal wall and to aid in transferring this infectivity (directly or indirectly) from the periphery to the CNS [5 , 6 ]. Given our limited knowledge on this subject, the characterization of the intracellular signaling mechanisms used by PrP106–126 and the specific cell types involved could lead to a better understanding of prion pathogenesis, peripheral transmission, and potentially, the development of inhibitory agents to prevent the spread of prions and the pathogenesis of TSE.


   ACKNOWLEDGEMENTS
 
S. M. B. was supported by an appointment in the Postgraduate Research Participation program from the Oak Ridge Institute for Science and Education (TN). The authors thank Drs. David Frucht and Steven Kozlowski (Center for Biologics Evaluation and Research, FDA, Bethesda, MD) for critical review of this manuscript and Dr. Masato Moriguchi (National Institute of Arthritis and Musculoskeletal and Skin Diseases, NIH, Bethesda, MD) for expert technical advice.

Received November 1, 2002; revised March 13, 2003; accepted April 1, 2003.


   REFERENCES
TOP
 ABSTRACT
 INTRODUCTION
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 REFERENCES
 

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C. Julius, M. Heikenwalder, P. Schwarz, A. Marcel, M. Karin, M. Prinz, M. Pasparakis, and A. Aguzzi
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