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* This work was supported by the Intramural Research Program of the NIAID, National Institutes of Health. 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. The on-line version of this article (available at http://www.jbc.org) contains supplemental Figs. S1 and S2.
Hemin (iron protoporphyrin IX) is a crucial component of many physiological processes acting either as a prosthetic group or as an intracellular messenger. Some unnatural, synthetic porphyrins have potent anti-scrapie activity and can interact with normal prion protein (PrPC). These observations raised the possibility that hemin, as a natural porphyrin, is a physiological ligand for PrPC. Accordingly, we evaluated PrPC interactions with hemin. When hemin (3–10 μm) was added to the medium of cultured cells, clusters of PrPC formed on the cell surface, and the detergent solubility of PrPC decreased. The addition of hemin also induced PrPC internalization and turnover. The ability of hemin to bind directly to PrPC was demonstrated by hemin-agarose affinity chromatography and UV-visible spectroscopy. Multiple hemin molecules bound primarily to the N-terminal third of PrPC, with reduced binding to PrPC lacking residues 34–94. These hemin-PrPC interactions suggest that PrPC may participate in hemin homeostasis, sensing, and/or uptake and that hemin might affect PrPC functions.
Iron protoporphyrin IX, a natural cyclic tetrapyrrole (cTP),
is vital to cellular homeostasis in either the Fe3+ (hemin) or Fe2+ (heme) oxidation state (supplemental Fig. 1). In hemoglobin and myoglobin the reversible binding of oxygen to the reduced iron of the heme permits oxygen transport and storage (
). This suggests that the modulation of endocytosis of PrPC through its N-terminal domain is important in a conserved physiological function of PrPC.
Based on these observations, we evaluated the possibility of hemin being a physiological ligand of PrPC. Here we show that hemin promotes PrPC clustering, internalization, and degradation in cultured cells. In cell-free reactions, the binding of PrPC to hemin alters the aggregation state and inherent peroxidase activity of the latter.
Preparation of Recombinant PrP (rPrPC)—Cell pellets of Escherichia coli expressing hamster PrP corresponding to residues 23–231 or 90–231 in the pET41 vector (EMD Biosciences) were lysed with BugBuster™ and lysonase (EMD Biosciences) in the presence of EDTA-free protease inhibitors (Roche Applied Science). Inclusion bodies were washed twice with 0.1× BugBuster™ in water and pelleted by centrifugation. The enriched rPrPC was further purified by minor modifications to the method of Zahn et al. (
). The protein was eluted with 10 mm sodium phosphate (pH 5.8), 500 mm imidazole, and 10 mm Tris. Pooled fractions were dialyzed against 10 mm sodium acetate or PBS. The construct containing residues 23–106 was purified in the same manner except the protein was eluted from the nickel column using 10 mm sodium acetate at pH 3.5, and fractions that contain the protein were further purified on an SP-Sepharose column using a salt gradient in sodium acetate at pH 5.0. The protein concentration of rPrPC was determined by absorbance at 280 nm. Purity of the final protein preparations was estimated at ≥99% when analyzed by SDS-PAGE, Western blot, and matrix-assisted laser desorption ionization-mass spectrometry (data not shown).
Preparation of Solutions—Hemin (Mann Research Laboratories Inc.), biliverdin (Frontier Scientific), and bilirubin (Frontier Scientific) were dissolved in Me2SO at 10 mm. Further dilutions were carried out in PBS or serum-free Opti-MEM (Invitrogen). Hemin stock solutions were also prepared in 0.5 m NaOH at 10 mm to investigate the effect of the μ-oxo-dimer of hemin, which is known to form at basic pH.
UV-visible Absorption Spectroscopy—For spectroscopic analysis, rPrPC and hemin were mixed in PBS (pH 7.4) containing 1 mm EDTA prior to measurement of absorbance. Measurements were made on a SpectraMAX 190 plate reader (Molecular Devices). The spectra were acquired between 300 and 800 nm.
Cell Culture—N2a5E4E is a mouse neuroblastoma (N2a) cell line that overexpresses mouse PrPC as described previously (
). Human neuroblastoma cells (NB1) express endogenous levels of PrPC. All cell lines were maintained at 37 °C in a humidified atmosphere of 5% CO2 in Opti-MEM supplemented with 10% fetal bovine serum (Invitrogen) and penicillin/streptomycin (100 units/ml, 100 μg/ml; Invitrogen).
PrPC Binding to Hemin-Agarose—A confluent tissue culture flask (25 cm2) containing cells described above was rinsed three times with PBS and lysed with 600 μl of PBS containing 0.5% Triton X-100 and 0.5% sodium deoxycholate. Cell debris and nuclei were removed by centrifugation at 2,700 × g for 5 min, and 600 μl of postnuclear supernatant was recovered. To the postnuclear supernatant, a protease inhibitor mixture (Complete, Roche Applied Science) was added according to the manufacturer's instructions. The final concentration of NaCl was adjusted to 0.5 m. Additional Triton X-100 and sodium deoxycholate were added to final concentrations of 1%. Sarkosyl (1%) was also added to improve the dissolution of the cell membranes. The final volume was adjusted to 1200 μl after the addition of all reagents and then incubated for 10 min at room temperature to allow the dissolution of membranes. Hemin-agarose beads (Sigma) were washed three times with PBS prior to use. Washed beads containing the equivalent of 0.1 μmol of hemin were added to 200 μl of cell lysate and incubated for 10 min at room temperature. After incubation, the unbound fraction was collected and precipitated with 800 μl of methanol. The beads were washed three times with the same buffer used for the binding step. The methanol precipitate and the beads were resuspended in 100 μl of SDS-PAGE sample buffer and boiled, and 5 μl was subjected to SDS-PAGE with staining for proteins with GelCode Blue (Pierce) or to immunoblot analysis for PrPC using antibody D13 (InPro) for mouse and hamster PrPC and 3F4 for human PrPC.
Immunodetection of Cell Surface PrPC—N2a5E4E cells were plated at low density in a 96-well plate and grown to confluence. At confluence, cells were washed once with serum-free Opti-MEM and treated with hemin, biliverdin, or bilirubin at 0, 1, 3, and 10 μm for 1 h at 37 °C. After treatment, the cells were fixed with 4% paraformaldehyde in PBS for 10 min followed by two washes with PBS. Then the cells were incubated with antibody D13 (InPro) diluted in PBS at 1:1000 for 1 h. After three washes of 5 min each, cells were incubated with secondary antibody conjugated with alkaline phosphatase diluted in PBS at 1:2000 for 1 h. Cells were washed three times and incubated with Attophos substrate (Promega) for 7–15 min until a yellow color was visible. Fluorescence intensity was measured in a SpectaMAX Gemini EM plate reader (Molecular Devices) using 450 nm excitation filter and 520 nm emission filter. The relative fluorescence intensity was calculated based on the signal obtained from untreated cells.
Immunofluorescence—N2a5E4E cells were washed twice with serum-free Opti-MEM and treated with 3 μm hemin for 1 h. After treatment, cells were fixed with 4% paraformaldehyde in PBS for 10 min followed by two washes with PBS. Cells were permeabilized with 0.1% Triton X-100 in PBS for 5 min. To block nonspecific antibody binding, cells were incubated with 10% normal goat serum and 0.1% Triton X-100 in PBS (blocking solution) for 10 min. An antibody against PrPC, SAF-32 (Cayman Chemicals), was diluted in blocking solution (1:200) and added to the cells. After 1 h of incubation, the cells were washed three times with PBS and incubated with secondary antibody anti-mouse IgG conjugated with Alexa 488 fluorescent dye (1:1000) for 1 h. Cells were washed three times and observed by confocal microscopy. All images were acquired with the same confocal parameters.
Biotinylation and Isolation of Cell Surface Proteins—N2aGFP-GPI cells were plated in 24-well plates and cultured for 3 days. At confluence, cells were washed three times with serum-free Opti-MEM and treated with hemin at 0, 1, 3, and 10 μm for 1 h at 37 °C. The cells were washed three times with PBS containing 1 mm CaCl2 and 1.2 mm MgSO4 (PBS Ca2+/Mg2+) on ice. Then 150 μl of 1 mg/ml NHS sulfo-LC biotin was added per well and incubated for 5 min at room temperature. NHS sulfo-LC biotin reacts predominantly with primary amino groups. After biotinylation, the cells were washed three times with PBS Ca2+/Mg2+ containing 100 mm glycine. The cells were then lysed with 200 μl of PBS containing 0.5% Triton X-100, 0.5% sodium deoxycholate, and a protease inhibitor mixture (Complete, Roche Applied Science) (lysis buffer). Cell lysates were incubated with 20 μl of streptavidin Dynabeads (Invitrogen) for 30 min at room temperature and then washed three times with the lysis buffer. The beads were resuspended in 40 μl of SDS-PAGE sample buffer. The samples were subjected to SDS-PAGE and Western blot analysis using the designated antibody for PrPC (D13, InPro), GFP (monoclonal anti-GFP, Roche Applied Science), or NCAM (anti-NCAM, Chemicon). Biotinylated proteins in general were stained with Neutravidin conjugated with alkaline phosphatase (Pierce). To check the effect of hemin on PrPC turnover, biotinylation of N2a5E4E cells was performed as described above but prior to the hemin treatment.
Detergent Insolubility Assay—All cells described were cultured and treated with hemin as described in the section above. The cells were then lysed with 200 μl of lysis buffer. Nuclei and cell debris were removed by centrifugation at 2,700 × g for 5 min at 4 °C, and then Sarkosyl was added to a final concentration of 0.5–1% (
). After 10 min of incubation on ice or at 37 °C, the detergent-insoluble material was recovered by ultracentrifugation at 360,000 × g for 30 min at 4 °C. Supernatant proteins were subjected to methanol precipitation. Pellets that were generated from ultracentrifugation or methanol precipitation were dissolved in SDS-PAGE sample buffer and subjected to immunoblot analyses using antibodies described in the section above.
Metabolic Labeling—Tissue culture flasks (25-cm2) were seeded with equal numbers of human neuroblastoma NB1 cells and grown until they were 80–90% confluent. The cells were preincubated for 1 h with 5 ml of 10 μm hemin in serum-free Opti-MEM followed by a 30-min incubation in 2 ml of methionine-free MEM containing hemin. Then 500 μCi of [35S]methionine was added to each flask and incubated for 30 min. Cells were rinsed twice with PBS and incubated in serum-free Opti-MEM containing hemin for the designated chase time.
Peroxidase Activity—The peroxidase activity was measured by oxidation of 3,3′,5,5′-tetramethylbenzidine (TMB) (Pierce) or 2,2′-azinobis [3-ethylbenzothiazoline-6-sulfonic acid]-diammonium salt (Pierce) by H2O2. Hemin (8 μm) was mixed with various concentrations of rPrPC, prior to the addition of substrate. After substrate addition the reaction was monitored for absorbance at 650 nm on a SpectraMAX 190 plate reader (Molecular Devices).
Hemin-induced PrPC Clustering and Internalization—Several different inhibitors of PrPSc formation, e.g. pentosan polysulfate (
), affect the intracellular localization of PrPC. Because various synthetic cTPs inhibit PrPSc formation, we wondered if hemin, as a natural cTP and potential physiological ligand for PrPC, can also affect PrPC localization. This was tested initially by using an immunofluorescence assay for PrPC detection in fixed and permeabilized cells. To enhance PrPC detection, a neuroblastoma cell line that expresses a high level of PrPC, N2a5E4E, was used. Without hemin treatment, both cell surface and intracellular perinuclear staining was observed. Hemin (3 μm) treatment of N2a5E4E cells for 1 h decreased the immunofluorescence of PrPC on the cell surface and caused some residual surface staining to appear more punctate than in the control cells (Fig. 1). No staining was observed when the primary antibody SAF-32 was omitted from the staining protocol. Furthermore, SAF-32 did not stain CF10 cells generated from PrPC null mice (data not shown) confirming the specificity of the antibody against PrPC. Finally, similar results were obtained with antibody D13 (data not shown). These hemin effects on PrPC localization were not likely because of cytotoxicity because the treatment at ≤10 μm for at least 4 h did not induce any signs of toxicity as judged by morphology or a cytotoxicity assay using 3-(4,5-dimethylthiazol-2-yl)-5-(3-carboxymethoxyphenyl)-2-(4-sulfophenyl)-2H-tetrazolium) (data not shown). Altogether, these results suggested that hemin induced both the aggregation and internalization of PrPC.
Selective Effects of Hemin on PrPC Aggregation—The formation of PrPC aggregates was also evaluated using a detergent insolubility assay (
). Upon hemin treatment, PrPC solubility decreased in a dose-dependent manner in both mouse and human cell lines (Fig. 2, a, b, e, and f). To eliminate the possibility of PrPC aggregation being due to overexpression, the solubility of endogenous PrPC expressed in N2aGFP-GPI cells was also analyzed, and similar results were observed (supplemental Fig. 2). Because hemin can interact with many proteins, we investigated the selectivity of the effects of hemin on the aggregation of PrPC compared with other cell surface proteins such as NCAM (neural cell adhesion molecule), which interacts with PrPC (
). No hemin-induced alteration of NCAM or GFP-GPI solubility was observed, indicating a degree of specificity for the effects of hemin on PrPC solubility (Fig. 2c and supplemental Fig. 2).
Quantifying Hemin-induced PrPC Internalization—To estimate the extent of internalization of PrPC after hemin treatments, relative amounts of cell surface PrPC were quantified using a fluorogenic immunoassay described under “Experimental Procedures.” To increase the PrPC detection in this assay, N2a5E4E cells were used. The PrPC specificity of the assay was indicated by the lack of fluorescence signal when the primary antibody (D13) was omitted and when the assay was applied to primary neuronal cells derived from PrPC null mice. The treatment of N2a5E4E cells with hemin for 1 h caused a dose-dependent reduction of cell surface PrPC (Fig. 3) reaching ∼10% of control levels at 10 μm hemin. It is known that basic solutions (e.g. NaOH) favor the formation of μ-oxo-dimers of hemin. NaOH-treated hemin was slightly more potent than Me2SO-solubilized hemin at causing PrPC internalization at 3 μm, but both solutions were effective overall. In contrast to hemin, its linear tetrapyrrole metabolites biliverdin and bilirubin did not significantly reduce cell surface PrPC (Fig. 3).
Selectivity of Hemin Effects on PrPC Internalization—To further test the selectivity of hemin effects on PrPC internalization, N2aGFP-GPI cells that express endogenous levels of PrPC and a recombinant GFP-GPI were treated with hemin, and cell surface proteins were then biotinylated, captured with streptavidin-coated magnetic beads, and subjected to SDS-PAGE. The gels were either immunoblotted for the detection of individual proteins or stained with a Neutravidin-alkaline phosphatase conjugate to reveal the overall profile of biotinylated cell surface proteins. Consistent with previous assays, the cell surface PrPC signal decreased in a dose-dependent manner (Fig. 4, a and b), but the extent of internalization was lower than the previous assay using the N2a5E4E cells. This discrepancy could be related to different expression levels of PrPC in the two cell types because the N2aGFP-GPI cells express a lower, endogenous level of PrPC, whereas the N2a5E4E cells overexpress PrPC. In contrast, the banding patterns and intensity of many other cell surface proteins were not visibly altered with hemin treatment (Fig. 4a). We also examined hemin effects on NCAM and GFP-GPI. As shown in Fig. 4, a and b, no alteration of NCAM was observed, whereas GFP bands increased with hemin treatment. GFP-GPI is expressed under a cytomegalovirus promoter whose activity can be enhanced by histone acetylation or demethylation (
), the increase of GFP-GPI on the cell surface could be due to increased expression, which was confirmed by a Western blot assay on total cell lysates (data not shown). Collectively, these data provide evidence that the effect of hemin on PrPC internalization was relatively selective.
Hemin-induced Degradation of PrPC—In the immunofluorescence studies of PrPC using N2a5E4E cells, we observed that in some cells, intracellular fluorescence increased as the cell surface staining decreased. However, in other cells, the overall fluorescence intensity decreased (Fig. 1) suggesting that the internalized PrPC might have been degraded. To directly analyze whether degradation of PrPC is induced by hemin treatment, cell surface proteins of N2a5E4E cells were pulse-labeled with biotin for 5 min, incubated with or without hemin (10 μm) for 1 h to allow for turnover, and isolated on streptavidin beads for immunoblot analysis. With hemin treatment, the biotinylated PrPC bands decreased compared with those in untreated cells, whereas the overall banding patterns and intensity of other biotinylated proteins was not noticeably affected (Fig. 5, a and b). These results clarify that the decrease of PrPC signal in response to hemin treatment is not because of reduced expression but to enhanced degradation.
Hemin effects on the biosynthesis and turnover of PrPC were also evaluated by pulse-chase [35S]methionine labeling and radioimmunoprecipitation of PrPC in human NB1 cells that express endogenous levels of PrPC. At time 0, immature glycosylated and unglycosylated forms of PrPC were seen as described previously (Fig. 5c, arrows) (
). After a 1-h chase, mature glycosylated forms predominated (Fig. 5c, asterisk). In hemin (10 μm)-treated cells, these PrPC bands disappeared more rapidly with increasing chase periods, showing increased PrPC turnover relative to that seen in control cells. The quantification of all glycosylated and unglycosylated PrPC bands from two experiments revealed that the loss of pulse-labeled PrPC was accelerated in the presence of 10 μm hemin (Fig. 5d). Altogether, these data show that hemin selectively alters the subcellular localization and turnover of the PrPC.
PrPC Binding to Hemin-Agarose—To evaluate whether hemin can directly interact with PrPC, hemin-agarose affinity chromatography was performed using N2a5E4E and NB1 cell lysates. Amounts of PrPC in bound and unbound fractions were analyzed by Western blotting. A single aliquot of hemin-agarose beads was able to fractionate ∼50% of PrPC from the total cell lysate (Fig. 6, a and b, lane 2). Additional PrPC (∼25%) could be extracted from the lysate with a fresh aliquot of hemin-agarose beads (data not shown). The absence of PrPC binding to agarose beads without hemin confirmed the specificity of the interaction between hemin and PrPC (Fig. 6, a and b, lane 4). To assess the selectivity of PrPC binding, the other proteins of each fraction were stained nonspecifically with GelCode Blue. A number of other proteins from the cell lysates also bound to hemin-agarose, as expected, but most proteins were much more abundant in the unbound fraction (Fig. 6, a and b, lanes 5 and 6). Thus, the hemin-agarose showed some selectivity for binding PrPC. As an additional indication of specificity and to examine whether the octapeptide repeats in PrPC might be involved in hemin binding, we evaluated the hemin-agarose binding of hamster PrPC lacking the octapeptide repeats and flanking sequences (HaPrP Δ34–94). Although the binding of wild-type hamster PrPC was as efficient as the binding of the wild-type mouse and human PrPC, only ∼10% of HaPrP Δ34–94 bound to hemin-agarose (Fig. 6c). These results showed that hemin interacts directly or indirectly with PrPC of multiple species and that PrPC residues 34–94 strongly influence that interaction.
UV-visible Spectroscopy—To obtain additional evidence of direct interactions between hemin and PrPC, we used UV-visible spectroscopy. Hemin is sparingly soluble in aqueous media and, when not bound to proteins, tends to form oligomers that absorb strongly at ∼390 nm (the Soret band). This absorbance maximum (Amax) can shift to different wavelengths upon interaction with other molecules. Using this spectral property of hemin, we evaluated the binding of hemin to purified rPrPC. Spectra of hemin alone at various concentrations showed that the Amax was slightly blue-shifted with increasing concentrations (Fig. 7, a–c, blue lines). However, when hemin was incubated with rPrPC at a 1:1 molar ratio, the Amax red-shifted to 411 ± 3 nm independent of the concentration of the complex (Fig. 7a, dotted green line). These results suggested that PrPC reorganized hemin molecules into distinct oligomeric states. Similar spectral changes were produced with C-terminally truncated rPrP (residues 23–106), which contains the octapeptide repeats (Fig. 7a, dotted pink line). However, the N-terminally truncated rPrP (residues 90–231) did not alter the hemin spectrum indicating that the C-terminal residues 90–231 were not required for hemin binding (Fig. 7a, dotted orange line).
Bovine serum albumin (BSA) is a well known hemin-binding protein that has nanomolar affinity for hemin (
). When BSA interacted with hemin, only a small red shift of Amax (396 ± 2 nm) occurred (Fig. 7b, dotted pink line). These different effects of rPrPC and BSA on the hemin spectrum indicated that PrPC and BSA interact in distinct ways with hemin.
To determine the stoichiometry of the observed hemin-PrPC interactions, increasing concentrations of hemin were added to a fixed rPrPC concentration (Fig. 7c) and vice versa (not shown). At molar excesses of hemin up to ∼10:1, the full Amax red shift was maintained, indicating that each PrP molecule could influence the spectrum of multiple hemin molecules. However, with further increases in the hemin:PrPC ratio, the Amax gradually shifted back toward the Amax of free hemin, suggesting that saturation of the binding to rPrPC had occurred. Taken together, these data indicate that multiple hemin molecules can bind directly to PrPC, primarily via the N-terminal half of the molecule.
Enhancement of the Peroxidase Activity of Hemin by Interactions with rPrPC—Given the observed interactions between hemin and PrPC, we sought clues as to whether such interactions might have additional physiological significance. It has been reported that an excess of free hemin can have cytolytic activity because of its inherent peroxidase activity (
). To see if binding to PrPC might alter such activities of hemin, we compared the peroxidase activity of free hemin and its rPrPC complex. In an assay using TMB as a substrate, the hemin-rPrPC complex showed increased peroxidase activity by up to 3-fold compared with hemin alone (Fig. 8). Similar results were also obtained using 2,2′-azinobis [3-ethylbenzothiazoline-6-sulfonic acid]-diammonium salt as a substrate (data not shown). Consistent with previous studies (
), the binding of hemin to BSA also increased its peroxidase activity (Fig. 8). In contrast, no superoxide dismutase or catalase activities of hemin itself or hemin-rPrPC complexes were observed (data not shown). The fact that the peroxidase activity of hemin is altered by binding to PrPC indicates that the interaction affects the inherent redox properties of this porphyrin.
Hemin interacts with a number of proteins stably or reversibly and orchestrates various vital biological activities. Here we have demonstrated that PrPC is a hemin-binding protein that undergoes aggregation, internalization, and degradation upon exposure to hemin.
Potential Relevance of Hemin Binding and Cellular Trafficking in Biological Activities of PrPC—PrPC constitutively cycles between the plasma membrane and endocytic compartments, and its endocytosis can take place via a clathrin-dependent mechanism (
). These observations suggest that cellular trafficking of PrPC is closely related to its physiological activities. Thus, the fact that hemin binding alters the PrPC trafficking suggests that PrPC may participate in hemin-dependent biological events and/or that hemin binding is relevant in PrPC functions.
The endocytosis of PrPC through clathrin-coated pits requires a transmembrane receptor. Recently, the low density lipoprotein receptor-related protein was identified as the transmembrane receptor that mediates copper-induced PrPC internalization (
). At the moment, it is not clear whether PrPC acts as a receptor for free hemin or interacts with hemin-hemopexin complexes and acts as a co-receptor in compartments such as blood and liver when these complexes are formed. However, given that both copper and hemin are small ligands that have redox properties and appear to bind a similar region of PrPC, it is possible that both of these ligands induce the internalization of PrPC by a related mechanism. Further studies will be required to evaluate this possibility.
PrPC trafficking also influences its conversion into PrPSc and disruption of the normal trafficking of PrPC seems to be a common mechanism for several classes of PrPSc inhibitors (
). Notably, the hemin concentration that effectively altered PrPC trafficking (∼3 μm) also reduced the formation of PrPSc in scrapie-infected cell cultures (data not shown). Therefore, the anti-scrapie activity of hemin in cell culture models might relate to its ability to stimulate the internalization of PrPC.
Roles of Peroxidase Activity of the Hemin-PrPC Complex—Hemin-containing peroxidases react with H2O2 and promote oxidative modifications of proteins, lipids, and halides (
). It is well known that hemin forms oligomers in aqueous media. The fact that co-incubation of hemin with rPrPC caused a red shift in the Soret region of the UV-visible spectrum indicates that hemin oligomers reorganize in the presence of PrPC. As a result, they become more reactive with H2O2 as indicated by the PrPC-induced enhancement of the peroxidase activity of hemin (Fig. 8).
The activity of antioxidant enzymes, such as glutathione peroxidase and superoxide dismutase, is altered in PrPC null mice and in scrapie-infected brains (
). These findings suggest that PrPC participates in the modulation of the activity of these enzymes and/or PrPC itself functions as an antioxidant molecule. Moreover, it has been reported that the flexible N-terminal domain of PrPC, which we have shown contains hemin-binding sites, is influential in cellular responses to oxidative stress (
) are important partners that assist in this function of PrPC. Interestingly, these ligands have distinct binding sites on the PrPC molecule and interact with PrPC on the cell surface suggesting that a formation of the macromolecular complex may occur on the plasma membrane in PrPC-dependent neuritogenesis. On the other hand, these interactions may be disrupted or prevented upon hemin binding to PrPC because the effect of hemin was selective for PrPC trafficking, without affecting the aggregation state or internalization of NCAM (Figs. 2 and 4), which is also endocytosed by a clathrin-dependent mechanism in normal circumstances (
). Thus, PrPC may be a multifunctional protein that requires a mechanism for switching between these functions. Conceivably, the binding of redox-active ligands such as hemin and metal ions can isolate PrPC from a macromolecular complex involved in neuronal plasticity and help organize PrPC into supramolecular assemblies that modulate cellular redox activities and/or sense reactive oxygen species. Further studies will be required to fully elucidate the physiological implications of hemin-PrPC interactions.
We thank Drs. Sonja Best, Valerie Sim, and Kristin McNally for critical reading of this manuscript. We thank Anita Mora and Gary Hettrick for graphics assistance.