19 Feb 2009: Singh A, Mohan ML, Isaac AO, Luo X, Petrak J, et al. (2009) Correction: Prion Protein Modulates Cellular Iron Uptake: A Novel Function with Implications for Prion Disease Pathogenesis. PLoS ONE 4(2): 10.1371/annotation/194f4e44-20f0-48eb-bbe9-14e21d18909b. doi: 10.1371/annotation/194f4e44-20f0-48eb-bbe9-14e21d18909b View correction
Converging evidence leaves little doubt that a change in the conformation of prion protein (PrPC) from a mainly α-helical to a β-sheet rich PrP-scrapie (PrPSc) form is the main event responsible for prion disease associated neurotoxicity. However, neither the mechanism of toxicity by PrPSc, nor the normal function of PrPC is entirely clear. Recent reports suggest that imbalance of iron homeostasis is a common feature of prion infected cells and mouse models, implicating redox-iron in prion disease pathogenesis. In this report, we provide evidence that PrPC mediates cellular iron uptake and transport, and mutant PrP forms alter cellular iron levels differentially. Using human neuroblastoma cells as models, we demonstrate that over-expression of PrPC increases intra-cellular iron relative to non-transfected controls as indicated by an increase in total cellular iron, the cellular labile iron pool (LIP), and iron content of ferritin. As a result, the levels of iron uptake proteins transferrin (Tf) and transferrin receptor (TfR) are decreased, and expression of iron storage protein ferritin is increased. The positive effect of PrPC on ferritin iron content is enhanced by stimulating PrPC endocytosis, and reversed by cross-linking PrPC on the plasma membrane. Expression of mutant PrP forms lacking the octapeptide-repeats, the membrane anchor, or carrying the pathogenic mutation PrP102L decreases ferritin iron content significantly relative to PrPC expressing cells, but the effect on cellular LIP and levels of Tf, TfR, and ferritin is complex, varying with the mutation. Neither PrPC nor the mutant PrP forms influence the rate or amount of iron released into the medium, suggesting a functional role for PrPC in cellular iron uptake and transport to ferritin, and dysfunction of PrPC as a significant contributing factor of brain iron imbalance in prion disorders.
Citation: Singh A, Mohan ML, Isaac AO, Luo X, Petrak J, et al. (2009) Prion Protein Modulates Cellular Iron Uptake: A Novel Function with Implications for Prion Disease Pathogenesis. PLoS ONE 4(2): e4468. doi:10.1371/journal.pone.0004468
Editor: Hilal Lashuel, Swiss Federal Institute of Technology Lausanne, Switzerland
Received: September 17, 2008; Accepted: December 26, 2008; Published: February 12, 2009
Copyright: © 2009 Singh et al. This is an open-access article distributed under the terms of the Creative Commons Attribution License, which permits unrestricted use, distribution, and reproduction in any medium, provided the original author and source are credited.
Funding: Funding source, NIH. Sponsor had no role at all in the study.
Competing interests: The authors have declared that no competing interests exist.
Prion protein (PrPC) is an evolutionarily conserved cell surface glycoprotein expressed abundantly on neuronal cells. Despite its ubiquitous presence, the physiological function of PrPC has remained ambiguous. The best characterized role for this protein remains its involvement in the pathogenesis of familial, infectious, and sporadic prion disorders, where a change in the conformation of PrPC from a mainly α-helical to a β-sheet rich PrP-scrapie (PrPSc) form renders it infectious and pathogenic –. The mechanism by which PrPSc induces neurotoxicity, however, is not clear. Studies over the past decade have clarified several aspects of this process , , . Prominent among these is the resistance of transgenic mice lacking neuronal PrPC expression to PrPSc induced toxicity, implicating PrPC as the principal mediator of the neurotoxic signal , . However, prion infected transgenic mice expressing PrPC only on astrocytes accumulate PrPSc and succumb to disease , leaving the matter unresolved. Adding to the complexity is the development of prion specific neuropathology in mice over-expressing normal or mutant PrP in the wrong cellular compartment in the absence of detectable PrPSc, suggesting the presence of additional pathways of neurotoxicity , . Although brain homogenates from these animals are not infectious in bioassays, these models suggest that a disproportionate change in the physiological function of PrPC is as neurotoxic as the gain of toxic function by PrPSc. Investigations on both fronts are therefore essential to uncover the underlying mechanism(s) of neurotoxicity in these disorders.
Efforts aimed at understanding the physiological function of PrPC and pathological implications thereof have revealed several possibilities, varying with the model, the physiological state, and the extra- and intracellular milieu in a particular tissue. Some of the reported functions include a role in cell adhesion, signal transduction, and as an anti-oxidant and anti-apoptotic protein , , . While the importance of these observations cannot be under-estimated, they fail to provide a direct link between PrPC function and dysfunction to prion disease pathogenesis. In this context, it is interesting to note that PrPC binds iron and copper, and is believed to play a functional role in neuronal iron and copper metabolism , . Since both iron and copper are highly redox-active and neurotoxic if mis-managed, it is conceivable that dysfunction of PrPC due to aggregation to the PrPSc form causes the reported accumulation of redox-active PrPSc complexes in prion infected cell and mouse models, inducing a state of iron imbalance –. A phenotype of iron deficiency in the presence of excess iron is noted in sporadic Cruetzfeldt-Jakob disease (sCJD) affected human and scrapie infected animal brain tissue, lending credence to this assumption .
To explore if PrPC is involved in cellular iron metabolism, we investigated the influence of PrPC and mutant PrP forms on cellular iron levels in human neuroblastoma cells expressing endogenous levels (M17) or transfected to express 6–7 fold higher levels of PrPC or mutant PrP forms. The following parameters were evaluated: 1) total cellular iron, 2) intracellular labile iron pool (LIP), 3) iron content of ferritin, and 4) levels of iron uptake proteins transferrin receptor (TfR) and transferrin (Tf) and iron storage protein ferritin that respond to minor changes in the LIP , . Our data demonstrate that PrPC increases cellular iron levels and the cells demonstrate a state of mild overload, while pathogenic and non-pathogenic mutations of PrP alter cellular iron levels differentially, specific to the mutation.
Normal and mutant PrP forms influence cellular iron levels differentially
The influence of PrP expression on cellular iron status was evaluated in M17 cells expressing endogenous PrPC or stably transfected to express 6–7 fold higher levels of PrPC or the following mutant PrP forms: 1) PrP231stop that lacks the glycosylphosphatidyl inositol (GPI) anchor and is secreted into the medium, 2) PrPΔ51–89 that lacks the copper binding octa-peptide repeat region, 3) PrPΔ23–89 that lacks the N-terminal 90 amino acids, and 4) PrP102L associated with Gerstmann-Straussler-Scheinker disease (GSS), a familial prion disorder (Fig. 1A). Expression of PrP in transfected cell lines was assessed by separating cell lysates on SDS-PAGE and probing transferred proteins with the PrP specific monoclonal antibody 3F4 . As expected, the di-, mono-, and unglycosylated forms of PrPC, PrPΔ51–89, PrPΔ23–89, and PrP102L migrating between 20 and 37 kDa are detected (Fig. 1B, lanes 2–5). Deletion mutations PrPΔ51–89 and PrPΔ23–89 migrate faster than PrPC and PrP102L as expected (Fig. 1B, lanes 3 and 4). M17 lysates show barely detectable levels of PrPC, while transfected cell lines express significantly higher levels of PrPC and mutant PrP forms (Fig. 1B, lanes 1–5).
(A) Diagrammatic representation of PrPC and mutant PrP forms evaluated in this study. (B) Lysates of M17, PrPC, PrPΔ51–89, PrPΔ23–89, and PrP102L were resolved by SDS-PAGE and immunoreacted for PrP and β-actin. All transfected cell lines express 6–7 fold higher levels of PrP relative to non-transfected M17 cells (lanes 1–5). (C) Cell lines in (B) were radiolabeled with 59FeCl3-citrate complex, washed with PBS supplemented with 100 µM DFO to chelate surface bound iron, and lysed. Equal amount of protein from each sample was spotted on a PVDF membrane, air dried, and exposed to an X-ray film.
To evaluate if PrPC or mutant PrP forms influence cellular iron uptake, M17, PrPC, PrPΔ51–89, PrPΔ23–89, and PrP102L cells cultured in serum-free medium for 1 hour were radiolabeled with 59FeCl3-citrate complex for 4 hours in the same medium, washed with PBS supplemented with 100 µM desferrioxamine (DFO) to remove surface bound iron, and lysed in non-denaturing buffer. Equal amount of protein from lysates was spotted on a PVDF membrane, air-dried, and exposed to an X-ray film. Surprisingly, PrPC and PrP102L cells incorporate significantly more 59Fe, while PrPΔ51–89, PrPΔ23–89 cells take up less 59Fe than M17 controls (Fig. 1C).
Major 59Fe labeled proteins in these cells were identified by separating cell lysates prepared in non-denaturing buffer on a 3–20% native gel in duplicate. One part of the gel was dried and subjected to autoradiography (Fig. 2, lanes 1–5), while the other was transferred to a PVDF membrane under native conditions and probed for ferritin and Tf using specific antibodies ,  (Fig. 2, lanes 6–15). Autoradiography shows a prominent iron labeled band consistent with ferritin (Fig. 2, lanes 1–5 and 6–10, black arrow), and a faster migrating band representing Tf (Fig. 2, lanes 1–5 and 11–15, open arrow) (the lower part of the autoradiograph is over-exposed to highlight the Tf band). Compared to M17 lysates, the amount of 59Fe bound to ferritin is higher in PrPC and PrP102L lysates, and lower in PrPΔ51–89 and PrPΔ23–89 lysates (Fig. 2, lanes 1–5). On the other hand, Tf bound iron is higher in M17 compared to PrPC, PrPΔ51–89, and PrPΔ23–89 lysates, and equivalent to PrP102L lysates (Fig. 2, lanes 1–5 and 11–15). The slower migrating iron labeled bands (*) probably represent a complex of Tf and TfR (Fig. 2, lanes 1–5) , . Probing for ferritin shows a major band and minor slower migrating forms probably representing ferritin complexes (Fig. 2, lanes 6–10, black arrow). Probing for Tf shows oligomers or glycosylation variants of Tf that correspond to 59Fe labeled purified transferrin fractionated similarly (Fig. 2, lanes 1–5, 11–15; Fig. S1). The relative levels of ferritin and Tf proteins in the samples correspond to radioactive iron in labeled ferritin and Tf bands in all samples (Fig. 2, lanes 1–15). Similar results were obtained when the cells were labeled with 59FeCl3-citrate complex for 16 hours or with purified 59Fe-Tf for 4 and 16 hours (data not shown), indicating similar uptake of non-transferrin and Tf bound Fe by these cells. Silver staining of re-hydrated autoradiographed gel confirms equal loading of protein for all samples analyzed (Fig. S1). Quantitative comparison of ferritin iron and levels of PrP, ferritin, and Tf between the cell lines is shown below in Fig. 4.
Radiolabeled lysates were fractionated on a 3–20% native gradient gel in duplicate. One set was subjected to autoradiography (lanes 1–5) and the other was transblotted and probed for ferritin and transferrin under native conditions (lanes 6–15).
The identity of iron labeled bands in Fig. 2 was further confirmed by cutting each band from fractionated PrPC lysates and re-fractionating electro-eluted proteins on SDS-PAGE followed by immunoblotting (Fig. S2). Lane 1 represents proteins eluted from the loading well that did not enter the running native gel. Lanes 2, 3 and 5 represent iron labeled bands that resolve adequately on native gels, and lane 4 represents unlabeled section of the gel that serves as a negative control. Sequential immunoreaction with specific antibodies confirms the presence of PrP in band 1, TfR in bands 1 and 2, ferritin in band 3, and Tf in band 5 (Fig. S2). Band 4 does not react with antibodies to known iron binding proteins. Silver staining shows co-migration of a few other un-identified proteins with bands 1–3, and almost none with bands 4 and 5 (Fig. S2).
To determine if PrPC mediates iron uptake directly, a modified non-denaturing gel system with a 3–9% gradient was used to separate 59Fe-labeled PrP effectively. Accordingly, M17 and PrPC cells were radiolabeled with 59FeCl3-citrate complex for 4 hours as above, and lysates were fractionated in duplicate under non-denaturing conditions. One part was dried and exposed to an X-ray film, while the other was transferred to a PVDF membrane and probed for PrP, ferritin, TfR, and Tf. As in Fig. 2, the amount of 59Fe incorporated by ferritin in PrPC cells is significantly higher than M17 cells (Fig. 3A, lanes 1 and 2, black arrow). A slower migrating 59Fe labeled band corresponding to Tf/TfR complex is detected in M17 lysates (Fig. 3A, lanes 1, 7, and 9, open arrow). Unlike Figure 2, PrP is resolved on this less concentrated gel system and is detected by PrP specific antibody 3F4 (Fig. 3A, lane 4, arrow-head). However, a corresponding 59Fe labeled band is not detected in lane 2, though pure 59Fe-labeled recombinant PrP is readily detected by this method as demonstrated previously . Evaluation of iron modulating proteins shows higher levels of ferritin and lower levels of TfR and Tf in PrPC lysates relative to M17 as in Fig. 1 above (Fig. 3A, lanes 5–10, arrow-head). Ferritin and Tf/TfR complex show corresponding iron labeled bands as expected (Fig. 3A, compare lanes 1, 2 with 5, 6, 9, 10). Fractionation of the same samples by SDS-PAGE followed by immunoblotting confirms increased levels of ferritin and decreased levels of Tf and TfR in PrPC lysates compared to M17 controls (Fig. 3B, lanes 1 and 2). Together, these results demonstrate that PrPC increases total cellular iron, ferritin iron, and ferritin levels, and decreases Tf and TfR levels. However, the absence of 59Fe-labeled PrPC indicates that either the association of PrP with 59Fe is transient or relatively weak and disrupted after cell lysis, or alternatively, PrP facilitates the incorporation of 59Fe into ferritin by an indirect mechanism that does not involve the formation of a PrP-iron complex.
(A) 59Fe-labeled M17 and PrPC lysates were fractionated on a 3–9% native gradient gel and auto-radiographed (lanes 1 and 2), or immunoblotted as above with antibodies specific to PrP, ferritin, TfR, and Tf (lanes 3–10). (B) Immunoblotting of the same samples following fractionation by SDS-PAGE shows similar differences in the levels of PrP, ferritin, Tf, and TfR as in (A) after normalization with actin (lanes 1 and 2). (C) 59Fe-labeled M17, PrPC, and PrP231stop lysates were fractionated by native gel electrophoresis and subjected to autoradiography (lanes 1–3). (D) Unlabeled lysates prepared from M17, PrPC, and PrP231stop lysates, and methanol precipitated proteins from the medium sample of PrP231stop cells were fractionated by SDS-PAGE and immunoblotted for PrP using 3F4 (lanes 1–4). (E) Membrane from (D) was re-probed for ferritin, Tf, TfR, and β-actin (lanes 1–3).
To evaluate if expression of PrPC on the cell surface is required for iron uptake, a similar evaluation was carried out in cells expressing PrP231stop that lacks the GPI anchor and is secreted into the medium. Radiolabeling of M17, PrPC, and PrP231stop cells with 59FeCl3-citrate complex for 4 hours shows significantly more 59Fe-ferritin in PrPC cells compared to M17 as above, and minimal change in PrP231stop samples (Fig. 3C, lanes 1–3, black arrow). Western blotting of M17, PrPC, and PrP231stop lysates and medium sample from PrP231stop cells cultured overnight in serum-free medium with 3F4 shows the expected glycoforms of PrP in PrPC lysates, and undetectable reactivity in M17 and PrP231stop lysates as expected (Fig. 3D, lanes 1–3). However, significant reactivity is detected in the medium of PrP231stop cells, demonstrating adequate expression and secretion of PrP231stop in transfected cells (Fig. 3D, lane 4) , . Re-probing of lysate samples for ferritin, Tf, and TfR shows increased levels of ferritin and decreased levels of Tf and TfR in PrPC samples compared to M17 lysates (Fig. 3E, lanes 1 and 2). PrP231stop lysates show minimal change in ferritin levels, and surprisingly, lower levels of Tf and TfR relative to M17 lysates (Fig. 3E, lanes 1 and 3). This observation is surprising since 59Fe-ferritin levels in PrP231stop cells are as low as M17, and yet the cells do not show increased levels of Tf and TfR as in M17-cells. Reaction for β-actin confirms equal loading of protein in all samples (Fig. 3E, lanes 1–3).
Quantitative comparison of ferritin iron and levels of ferritin, Tf, and TfR shows significant differences between cell lines. Thus, relative to M17 cells, PrPC cells show an increase in ferritin iron and ferritin levels to 570 and 565%, and a decrease in Tf and TfR levels to 70 and 75% respectively. A similar comparison of mutant cell lines relative to PrPC cells shows the following: PrPΔ51–89 cells show a decrease in ferritin iron and ferritin to 7.0, 6.9%, and insignificant change in Tf and TfR levels. PrPΔ23–89 cells show a similar decrease in ferritin iron and ferritin levels to 7.5 and 7.2%, an increase in Tf to 120%, and insignificant change in TfR levels. PrP102L-cells show a decrease in ferritin iron and ferritin levels to 89 and 90%, and an increase in Tf and TfR levels to 300 and 142% respectively. PrP231stop cells show a decrease in ferritin iron and ferritin to 27 and 16%, and a decrease in Tf and TfR levels to 89 and 67% respectively. Quantification of PrP expression relative to M17 shows levels of 650, 710, 750, 610, and 5% in PrPC, PrPΔ51–89, PrPΔ23–89, PrP102L, and PrP231stop cells respectively (Fig. 4).
Quantitative evaluation after densitometry of ferritin iron and levels of PrP, ferritin, Tf, and TfR in PrPC, PrPΔ51–89, PrPΔ23–89, PrP102L, PrP231stop-cells relative to non-transfected M17 controls. Values are mean±SEM of 11 independent experiments. The y-scale is linear but has been re-scaled after the break to illustrate the data clearly. For M17 vs. PrPC *p<0.001, **p<0.01, and for PrPC vs. mutant cell lines #p<0.001, ##p<0.01).
Considering the tightly orchestrated and coordinated balance between cellular iron levels and iron uptake and storage proteins , , these results indicate a mild iron overload in PrPC-cells relative to M17-cells, and an indefinable phenotype in mutant cell lines since the iron uptake proteins Tf and TfR do not respond to ferritin iron levels as expected. Since Tf and TfR levels are reflective of the biologically available intracellular labile iron pool (LIP) that is maintained within the physiological range by ferritin, these results indicate a disconnect between ferritin iron and the cellular LIP, or a failure of the iron regulatory loop involving the LIP, iron binding proteins 1 and 2, TfR, and ferritin to induce appropriate response.
Mutant PrP forms influence the uptake of iron by ferritin
The influence of normal and mutant PrP forms on intracellular LIP was evaluated in M17 and transfected cell lines cultured in complete medium under normal culture conditions. All cell lines were loaded with the iron binding dye calcein-AM, and the increase in fluorescence in response to salicylaldehyde isonicotinoyhydrazone (SIH), a cell permeable iron chelator, was measured (Fig. 5A) . Relative to M17 cells, PrPC cells show an increase in LIP to 143%, an expected observation since the ferritin iron levels of these cells are also higher than M17 cells (compare Figs. 5A and 4). A similar evaluation of mutant cell lines relative to PrPC-cells shows a decrease in LIP to 95, 78 and 67% in PrPΔ51–89, PrPΔ23–89, PrP102L-cells, and an increase to 155% in PrP231stop cells respectively (Fig. 5A). These results indicate that Tf and TfR levels in mutant cell lines observed in Fig. 4 above respond to the LIP rather than ferritin iron content as expected. More importantly, these results indicate a block in uptake or increased uptake of iron by ferritin in specific cell lines, accounting for the disproportionate levels of ferritin iron and intracellular LIP, and the unexpected response of Tf and TfR to cellular iron content.
(A) Indicated cell lines were loaded with calcein and intracellular LIP was estimated by quantifying the SIH chelatable iron pool. Values are mean±SEM. n = 12 for M17 and PrPC, and 7 for mutant cell lines. *p<0.001, **p<0.01, #p<0.001, ##p<0.01. (B) The same cell lines were exposed to 0.1 mM FAC for 16 hours and 50 µg of protein from cell homogenates was spotted on a PVDF membrane and reacted with Ferene-S, a dye that forms a blue reaction product with iron .
To evaluate if the difference in ferritin iron content of different cell lines is maintained in the presence of excess extra-cellular iron, M17, PrPC, PrPΔ51–89, PrPΔ23–89, PrP102L, and PrP231stop-cells were cultured overnight in the presence of 0.1 mM ferric ammonium citrate (FAC). (This dose of FAC was found to cause <1% cell death after overnight exposure). After washing the cells with PBS supplemented with 100 µM DFO to remove surface bound iron, cells were disrupted with glacial acetic acid and equal amount of protein from each cell line was spotted on a PVDF membrane. Reaction with Ferene-S, a dye that forms a blue reaction product with iron , shows a marked increase in protein bound iron in all cell lines compared to unexposed controls (Fig. 5B). More importantly, each cell line reflects cell-specific differences in protein bound iron as observed for ferritin iron above (Fig. 5B). Fractionation of lysates by SDS-PAGE followed by immunoblotting for PrP, ferritin, and TfR shows up-regulation of PrP and ferritin, and down-regulation of TfR to undetectable levels in FAC exposed lysates (Fig. S3 A, lanes 1–4) . Up-regulation of PrP in response to FAC appears to be at the mRNA level (Fig. S3 B). These results suggest a dominant role for PrP in the transport of extracellular iron to ferritin both under normal culture conditions and in the presence of excess extra-cellular iron.
Together, the above results demonstrate a state of relative iron overload in PrPC-cells compared to M17 controls as indicated by an increase in intracellular LIP and iron content of ferritin, increase in iron storage protein ferritin, and decrease in iron uptake proteins Tf and TfR. Relative to PrPC-cells, mutant PrP expressing cells show a substantial decrease in ferritin iron in PrPΔ51–89, PrPΔ23–89, and PrP231stop-cells, and relatively less reduction in PrP102L-cells. Intracellular LIP is reduced in PrPΔ23–89 and PrP102L, minimally altered in PrPΔ51–89, and substantially increased in PrP231stop-cells relative to PrPC-cells. Tf and TfR respond to LIP levels in some cell lines, but show an unexpected change in others, reflecting a state of cellular iron imbalance.
Stimulation of PrP endocytosis increases, and cross-linking decreases ferritin iron content
Further support for the role of PrP in mediating cellular iron uptake was obtained by assessing iron incorporation into ferritin following stimulation or disruption of PrPC endocytosis by 3F4, a well characterized monoclonal antibody specific for methionine residues 109 and 112 of human PrP . A similar approach has been used successfully to down-regulate mouse PrP using Fab fragments of PrP specific antibodies . Initial evaluation revealed that 3F4 concentrations of 1 and 12 µg/ml are optimal for stimulating and disrupting endocytosis of PrPC respectively without compromising cell viability.
To evaluate the effect of antibody treatment morphologically, M17 and PrPC-cells exposed to 1 µg/ml of 3F4 for 5 days were fixed, permeabilized, and reacted with anti-mouse-FITC. Both M17 and PrPC-cells show minimal reactivity at the plasma membrane, but significant reactivity in endocytic vesicles that are more prominent in PrPC cells (Fig. 6A, panels 1 and 2, arrow-head). These observations suggest significant endocytosis of PrPC along with 3F4. Untreated PrPC-cells reacted with 3F4-anti-mouse-FITC show punctuate reaction at the plasma membrane and minimal intracellular reaction as expected for normal distribution of PrPC (Fig. 6A, panel 3, arrow). Exposure to 12 µg/ml of 3F4, however, cross-links PrPC at the plasma membrane and reduces its endocytosis significantly (Fig. 6B, panels 1 and 2). As a control, mouse neuroblastoma cells (N2a) expressing mouse PrP that does not react with 3F4 were exposed to 3F4 and reacted with mouse PrP-specific antibody 8H4 followed by anti-mouse-FITC. Examination shows normal distribution of PrPC at the plasma membrane and some reactivity in the Golgi region as expected (Fig. 6B, panel 3) . Exposure of PrPC cells to anti-Thy-1, a monoclonal antibody to an irrelevant GPI-linked protein abundant on neuronal cells shows normal distribution of PrPC when reacted with 8H4-anti-mouse-FITC (Fig. 6B, panel 4), confirming the specificity of 3F4 mediated endocytosis and cross-linking of PrPC.
(A) Immunostaining of M17 and PrPC cells exposed to 1 µg/ml of 3F4 for 5 days shows a prominent reaction in vesicular structures in M17 and PrPC cells (panels 1 and 2). Coalesced vesicles simulating aggregated PrPC are evident near the Golgi region and in the cytosol of PrPC cells (panel 2). Untreated PrPC-cells reacted with 8H4-anti-mouse-FITC show a prominent reaction at the plasma membrane as expected (panel 3). (B) Reaction of M17 and PrPC cells exposed to 12 µg/ml of 3F4 for 4 hours with anti-mouse FITC shows cross-linking of PrP on the plasma membrane of M17 and PrPC cells (panels 1 and 2, arrow) and a slight increase of reactivity in vesicular structures in the latter (panel 2, arrow-head). Similar exposure of N2a-cells to 3F4 and PrPC-cells to anti-Thy1 antibody followed by immunoreaction with 8H4-anti-mouse-FITC shows plasma membrane and Golgi reaction of endogenous PrP in N2a cells (panel 3) and plasma membrane distribution of PrP in Thy-1 exposed cells (panel 4). (Mouse PrP expressed by N2a cells does not react with 3F4).
The effect of increased endocytosis of PrPC on ferritin iron content was evaluated by radiolabeling cells cultured in the presence of 1 µg/ml of 3F4 with 59FeCl3 for the last 4 hours of the incubation, and analyzing radiolabeled lysates as in Figure 1 above. Fractionation by non-denaturing page shows a significant increase in ferritin iron in the 3F4 exposed lysate compared to untreated control (Fig. 7A, lanes 1 and 2, open arrow). Analysis by SDS-PAGE and immunoblotting shows 2–3 fold increase in reactivity for all PrP glycoforms with anti-PrP antibodies 3F4 and 8H4 (Fig. 7A, lanes 3–6). However, the 18 kDa fragment that results from recycling of PrPC from the plasma membrane is not increased in 3F4 exposed lysates, indicating stimulation of PrPC internalization and possible intracellular accumulation by 3F4 binding rather than increased recycling from the plasma membrane (Fig. 7A, lanes 5 and 6) . The 50 kDa band represents internalized 3F4 (Fig. 7A, lanes 4 and 6). Immunobloting for ferritin, Tf, and TfR shows an increase in TfR, and minimal change in ferritin and Tf levels (Fig. 7A, lanes 7 and 8). Quantification by densitometry shows an increase in ferritin iron to 271%, and insignificant change in ferritin and Tf levels by 3F4 treatment. The increase in TfR levels to 175% is probably due to co-endocytosis with PrP-antibody complex (Fig. 7B). Measurement of cellular LIP revealed insignificant difference between 3F4 exposed and untreated controls after 24 hours (data not shown) or 5 days of treatment, indicating efficient transport of iron to ferritin within this time frame (Fig. 7C). PrPC cells treated with anti-Thy-1 antibody, however, demonstrated a significant decrease in LIP after 5 days of incubation with 3F4 (Fig. 7C).
(A) PrPC-cells exposed to 1 µg/ml of 3F4 for 5 days were radiolabeled with 59FeCl3 for 4 hours, and lysates were fractionated on a non-denaturing gel and auto-radiographed (lanes 1 and 2). Equal aliquots of the same samples were boiled in SDS-containing sample buffer and fractionated in duplicate by SDS-PAGE followed by immunoblotting with PrP specific antibodies 3F4 and 8H4 (lanes 3–6). Subsequently, the membranes were re-probed for ferritin, Tf, TfR, and β-actin (lanes 7 and 8). (B) Quantification by densitometry shows an increase in ferritin iron and TfR levels, and insignificant change in Tf levels in 3F4 exposed cells. Values are mean±SEM of three independent experiments. *p<0.001 compared to untreated cells. (C) Estimation of LIP after exposing the cells to 1 µg/ml of 3F4 or anti-Thy-1 antibody for 5 days shows insignificant difference between untreated and 3F4 treated PrPC cells, and a decrease in anti-Thy-1 treated cells. *p<0.001. n = 5.
A similar evaluation of cells exposed to 12 µg/ml of 3F4 for 4 hours shows significantly less increase in ferritin iron compared to untreated controls (Fig. 8A, lanes 1 and 2, open arrow). Separation by SDS-PAGE and immunoblotting shows increase in PrP reactivity (Fig. 8A, lane 4) and an increase in the levels of ferritin, Tf, and TfR (Fig. 8A, lanes 5 and 6). Quantification shows an increase in ferritin iron to 148%, and an increase in the levels of ferritin and TfR to 153 and 146% respectively. Tf levels show insignificant change by this treatment (Fig. 8B). A similar increase in ferritin iron is observed when M17 cells expressing endogenous levels of PrP are exposed to 3F4 (Fig. S4 lanes 1 and 2), ruling out the effect of over-expression of PrPC on these observations. Exposure to equivalent amounts of anti-Thy-1 does not alter ferritin iron content significantly (Fig. S4, lane 3). Measurement of intracellular LIP after 4 hours of exposure to 12 µg/ml of 3F4 shows an increase to 170% in treated cells compared to untreated controls. Exposure to similar concentrations of anti-Thy-1 shows a decrease to 70% (Fig. 8C), an unexpected effect that requires further evaluation.
(A) PrPC-cells exposed to 12 µg/ml of 3F4 for 4 hours were radiolabeled with 59FeCl3 in the last 2 hours, and lysates were fractionated on a native gel followed by autoradiography (lanes 1 and 2). Equal aliquots of lysates were fractionated by SDS-PAGE as above and immunoblotted with 3F4 (lanes 3 and 4). The membrane was re-pobed for ferritin, Tf, TfR, and β-actin (lanes 5 and 6). (B) Quantification by densitometry shows an increase in ferritin iron, ferritin, and TfR levels, and insignificant change in Tf levels by 3F4 treatment. *p<0.001, **p<0.025. n = 3. (C) Estimation of LIP after exposing the cells to 12 µg/ml of 3F4 or anti-Thy-1 antibody for 4 hours shows an increase in 3F4 exposed cells, and a decrease in anti-Thy-1 treated cells. *p<0.001. n = 7.
The above results indicate that stimulation of PrPC endocytosis over a prolonged period increases iron incorporation into ferritin, whereas cross-linking of PrPC that is likely to result in its degradation following endocytosis has relatively less effect on ferritin iron. The increase in intra-cellular LIP by cross-linking PrP without any increase in ferritin iron probably reflects inefficient transport of iron to ferritin in the absence of PrP, as observed for certain mutant forms of PrP. The levels of ferritin, Tf, and TfR probably reflect an artifactual change due to membrane perturbation by antibody treatment rather than a response to intracellular LIP.
PrP does not modulate release of iron from cells
To determine if the difference in cellular iron levels between cell lines is due to differential release into the medium, M17, PrPC, PrPΔ51–89, PrPΔ23–89, and PrP102L cells were cultured in the presence of 3H-thymidine overnight to monitor cell proliferation and radiolabeled with 59FeCl3 for 4 hours as above. Labeled cells were washed with PBS containing 100 µM DFO to remove surface bound 59Fe, and chased in complete medium for 30 minutes to 16 hours. At the indicated times equal aliquots of medium were retrieved and released 59Fe was quantified in a γ-counter. Kinetic analysis shows minimal difference in extracellular iron between cell lines after normalizing with 3H-thymidine (Fig. 9A). Estimation of cell-associated 59Fe after 16 hours of chase shows more 59Fe in PrPC and PrP102L, and significantly less in PrPΔ51–89 and PrPΔ23–89 compared to M17 lysates as observed in Fig. 1 above (Fig. 9B). However, the fold difference in ferritin iron content between M17 and other cell lines is significantly less after 16 hours of chase, and represents steady state levels of iron content in each cell line. Evaluation of possible ferroxidase activity of recombinant PrP using plasma as a positive control yielded negative results (Fig. 9C). Though informative, this result does not rule out possible ferroxidase activity of cell-associated PrP, a technically challenging assay that has yielded inconclusive results (data not shown).
(A) Cells expressing PrPC, PrPΔ51–89, PrPΔ23–89, and PrP102L were radiolabeled with 59FeCl3, washed with PBS supplemented with DFO, and chased in complete medium for 30, 60, 90, 120 min, and 16 hours. At the indicated time points equal aliquots of medium samples were quantified in a γ-counter. Estimation of released 59Fe does not show a significant difference between the indicated cell lines at any time point. n = 6 experiments in triplicate. (B) Cell associated 59Fe after 16 h of chase reflects the ferritin iron content of each cell line noted in Figure 1 above, though the difference between cell lines is significantly less. (C) Possible ferroxidase activity of recombinant PrP was measured using the established colorimetric method  with modifications. Negative controls included water and albumin supplemented with copper, and positive controls included plasma in the absence or presence of copper. Recombinant PrP does not show detectable ferroxidase activity either in the absence or presence of copper, whereas plasma shows a robust reaction under similar conditions.
The results presented in this report demonstrate an unprecedented role of PrP in facilitating iron uptake by cells and its transport to cellular ferritin. Using a combination of neuroblastoma cell lines expressing normal and mutant PrP forms, we demonstrate that over-expression of PrPC increases intracellular LIP and the amount of iron deposited in ferritin. Pathogenic and non-pathogenic mutations of PrP over-expressed to the same extent as PrPC alter cellular LIP and ferritin iron content differentially, specific to the mutation. Certain cell lines, especially cells expressing anchorless PrP231stop, demonstrate increased LIP in the presence of decreased ferritin iron, while PrP102L-cells display low LIP in the presence of adequate ferritin iron. Furthermore, stimulation of endocytosis by PrP specific antibody increases ferritin iron, while cross-linking at the plasma membrane increases LIP but has minimal effect on ferritin iron, indicating that alteration of PrP function or cellular localization disturbs the homeostasis between ferritin iron and cellular LIP. The differential incorporation of iron by mutant cell lines is maintained in the presence of excess extra-cellular iron, demonstrating a dominant role of PrPC in iron uptake and transport. The positive effect of PrPC on cellular iron is mainly due to enhanced uptake since the amount released into the culture medium is not altered in any of the cell lines tested. Together, these observations suggest a role for PrPC in mediating iron uptake and transport to ferritin directly, or by interacting with other iron modulating proteins. Below we discuss these data with reference to possible functions of PrPC in cellular iron metabolism, and the implications thereof in inducing imbalance in iron homeostasis observed in prion disease affected brains , , .
It is surprising that a GPI-linked protein such as PrPC is involved in iron transport to ferritin since PrPC is a membrane protein that undergoes vesicular transport while ferritin is cytosolic . Normally, cellular iron uptake is mediated by the Tf/TfR dependent and independent pathways, the former being most prominent and well characterized especially in neuroblastoma cells. In the Tf/TfR dependent pathway, ferric iron captured by Tf is taken up by the cells through TfR-mediated uptake via clathrin coated pits. Tf-bound ferric iron is released in the acidic environment of the endosomes, reduced to ferrous iron by an endosomal ferric reductase Steap3, and transported across the endosomal membrane by DMT1 to cytosolic ferritin where it is oxidized to the fairly inert ferric form by ferritin H-chain and stored , , . In the Tf-independent pathway, iron is taken up by an unknown transport mechanism, possibly non-specifically by fluid phase of endocytosis, and stored in ferritin. Ferritin regulates the biologically available LIP in the cell, and is itself regulated by iron regulatory proteins (IRPs) 1 and 2 , , , . In neuroblastoma cells, the LIP is a function of total cellular iron, and an increase in cellular iron is accompanied by increased ferritin content to maintain the LIP within safe limits , . Where might PrP intersect with this tightly orchestrated mechanism of iron uptake, transport, and storage? Three potential mechanisms are plausible: 1) modulation of uptake at the plasma membrane independently or by interacting with the Tf/TfR dependent pathway, 2) facilitation of iron transport to cytosolic ferritin across the endosomal membrane by promoting ferric iron release from Tf and/or its reduction for transfer through DMT1 , or 3) assistance in deposition into ferritin by oxidizing ferrous iron to the ferric form. It is unlikely that PrP facilitates export of iron from neuroblastoma cells based on our observations.
At the plasma membrane, PrPC could take up iron directly from the extra-cellular milieu and deliver to an endosomal compartment as suggested for copper . However, this seems unlikely for three reasons; 1) 59Fe-labeled PrPC could not be detected in radiolabeled cells although labeled recombinant PrP is easily detected using the same procedure , 2) 59Fe-labeled recombinant PrP loses its label to Tf when added to cells, indicating lower affinity for iron relative to Tf (unpublished observations), and 3) intra-cellular LIP is high in cells expressing anchorless PrP231stop despite low ferritin iron content, indicating efficient uptake of iron in the absence of cell surface PrPC. It remains plausible, though, that PrPC modulates iron uptake by the Tf/TfR pathway at the plasma membrane or in an endosomal compartment .
It is also possible that extracellular iron induces the movement of PrPC from detergent insoluble membrane domains where it normally resides to the proximity of TfR in a similar manner as in the presence of copper . Here, it may enhance the binding of iron loaded Tf to its receptor, or stimulate the endocytosis of Tf/TfR complex by a direct or an indirect interaction. In this context, it is interesting to note that PrPC undergoes endocytosis through clathrin coated pits after associating with a transmembrane protein through its N-terminal domain , suggesting that the reported co-localization of PrPC with Tf and TfR within endosomes may reflect a functional association rather than co-residence due to a common mode of endocytosis . Assuming this scenario, the increase in TfR levels by stimulation of PrP endocytosis by 3F4 and the differential effect of mutant PrP forms on ferritin iron content may be explained by a change in the rate of endocytosis, or altered interaction of normal and mutant PrP forms with Tf or TfR due to misfolding –. We have previously reported increased endocytosis and defective recycling of mutant PrP102L in neuroblastoma cells , a fact that may account for increased ferritin iron in these cells. Though attractive, this model fails to explain decreased ferritin iron in the presence of significantly high LIP in cells expressing anchorless PrP231stop and by cross-linking PrP at the plasma membrane, indicating a role downstream from the plasma membrane. The up-regulation of PrPC at the transcriptional and translational level when cells are exposed to excess extra-cellular iron (supporting information) perhaps reflects its function as an iron regulatory protein, though a protective response to oxidative stress cannot be ruled out under these experimental conditions . However, since all cell lines display similar differences in 59Fe-ferritin content when labeled with 59FeCl3 or purified 59Fe-Tf (unpublished observations), it is likely that PrPC functions downstream of the iron uptake pathways specific for free and Tf bound iron, perhaps in an endosomal compartment.
Keeping the above facts in mind, it is plausible that PrPC functions as a ferric reductase along with Steap3 to facilitate the transport of ferric iron released from Tf across the endosomal membrane to cytosolic ferritin. This assumption is supported by the fact that PrPC functions as a copper transport protein by reducing copper (II) prior to transfer to copper (I) specific trafficking proteins within cells . Such a function would explain the low ferritin iron content in cells expressing mutant PrP lacking the octapeptide region responsible for reducing copper (II) , the observed up-regulation of PrPC in response to exogenous iron, increase in ferritin iron by increased expression of PrPC and stimulation of PrP endocytosis, and co-localization of PrPC and ferritin in cells exposed to excess iron . However, decreased ferritin iron despite high LIP levels in cells expressing anchor-less PrP and the opposite scenario in PrP102L-cells suggests an additional role in iron transport between the LIP and cellular ferritin, a function that is hard to explain merely by the altered reductase activity of mutant proteins. Although we could not detect measurable ferroxidase activity of recombinant PrP, such a function of cell associated PrPC would explain the facilitative effect of PrPC on iron incorporation into ferritin. Further studies are required to resolve this question.
Despite obvious shortcomings in our data in explaining the mechanistic details of cellular iron modulation by PrP, this report clearly shows the effect of PrP and its mutants on iron uptake and transport. We demonstrate a state of mild iron overload mediated by PrPC, and mild iron deficiency or imbalance by pathogenic and non-pathogenic mutations of PrP. The positive role of PrPC on cellular iron levels is further supported by a recent study where transgenic mice lacking PrPC expression (PrP−/−) recover slowly from experimentally induced hemolytic anemia , indicating a functional role for PrPC in iron uptake by hematopoietic cells. These findings take on a greater significance since prion disease affected human and animal brains show signs of iron imbalance , a potentially neurotoxic state due to the highly redox-active nature of iron. It is conceivable that dysfunction of PrP due to aggregation combined with the formation of redox-active PrPSc aggregates  induces brain iron imbalance, contributing to prion disease associated neurotoxicity. Future studies are required to define the precise biochemical pathway of iron modulation by PrP, and develop therapeutic strategies to prevent iron induced neuronal death in prion disorders.
Materials and Methods
Antibodies and chemicals
Monoclonal anti-PrP antibodies 3F4 and 8H4 were obtained from Signet (Dedham, MA) and Drs. Man-Sun Sy (Case Western Reserve University) and Pierluigi Gambetti (National Prion Surveillance Center, Case Western Reserve University) respectively. Antibody against human ferritin was purchased from Sigma (St. Louis, MO), anti-transferrin from GeneTex (San Antonio, TX), anti transferrin receptor from Zymed Laboratories Inc (Carlsbad, CA), and anti-Thy 1.1 from eBioscience (SanDiego, CA). Secondary antibodies tagged with HRP or fluorophores FITC and TRITC were obtained from Amersham Biosciences (England) and Southern Biotechnology Associates (Birmingham, AL) respectively. Ferrous ammonium sulfate, Ferene S, and all other chemicals were purchased from Sigma. All cell culture supplies were obtained from Invitrogen. 59FeCl3 was from Perkin-Elmer.
Cell lines and culture conditions
Human neuroblastoma cells (M17) were obtained from J. Biedler (Memorial Sloan-Kattering Cancer Center, New York) and purchased from ATCC. M17 cells expressing PrPC, PrP231stop, PrPΔ51–89, PrPΔ23–89, and PrP102L were generated and cultured as described in previous reports , . For this study M17 cells from two different sources were transfected at least three separate times and bulk transfected cells were used to avoid cloning artifacts. Similarly transfected cells from two different investigators and cells cultured in DMEM supplemented with 10% FBS and Opti-MEM supplemented with different lots of FCS were also tried to avoid errors due to culture conditions.
Radiolabeling with 59FeCl3
M17, PrPC, and mutant PrPΔ51–89, PrPΔ23–89, PrP102L, and PrP231stop cells cultured overnight to 80% confluency were serum starved for 1 h and incubated with 59FeCl3-citrate complex (1 mM sodium citrate and 20–25 µCi of 59FeCl3 in serum free Opti-MEM; molar ratio of citrate to iron was maintained at 100:1) for 4 h at 37°C in the incubator. At the end of the incubation cells were washed 3 times with ice cold PBS and lysed with native lysis buffer (0.14 M NaCl, 0.1 M HEPES, pH 7.4, 1.5% Triton X-100 and 1 mM PMSF). Aliquots of lysates were mixed with glycerol (to a final concentration of 5%) and traces of bromophenol blue, and equal amount of protein from each sample was resolved on 3–9% native gradient gel. For fractionation on SDS-PAGE, the same samples were mixed with 4× SDS-sample buffer, boiled for 10 min and resolved on SDS-PAGE followed by immunoblotting.
Native gradient gel electrophoresis, autoradiography, immunoblotting and electroelution
Electrophoresis of lysates was performed using a Hoefer SE 600 vertical apparatus with a cooling system. Linear 3–20% (Fig. 1) or 3–9% (Fig. 3) gradient polyacrylamide gels were prepared as described by Vyoral et al.  with modifications. The gel mixture contained 0.375 M Tris, pH 6.8, 1.5% Triton X-100, and 1.18 mM ammonium persulfate. N,N,N′,N′-Tetramethylethylenediamine (TEMED) was added to a final concentration of 5.38 mM. Radiolabeled lysates mixed with glycerol were subjected to electrophoresis using electrode/running buffer (25 mM Tris, 192 mM glycine pH 8.3, and 1.5% Triton X-100) under constant current (100 mA) for 4 h at 4°C. Gels were either electroblotted or vacuum dried (BioRad) and exposed to X-ray film (Kodak BioMax XAR) fitted with intensifying screens. For Western Blotting, gels were washed thoroughly with electrode buffer without Triton X-100 for 2 h (each wash of 200 ml, 10 min) on a slowly rocking platform to remove Triton. The gel was electroblotted to a PVDF membrane using BioRad semi-dry electroblotting system with anode buffer (25 mM Tris, pH 10.4) and cathode buffer (25 mM Tris, 39 mM glycine, pH 9.2) at 25 V for 90 min. Membranes were further processed for immunodetection as described below. To confirm the identity of iron labeled proteins, iron bands were excised from native gels and proteins were electro-eluted using Biorad electro-eluter at 60 mA for 4 h. Eluted proteins were concentrated by methanol precipitation and analyzed by SDS-PAGE.
SDS-PAGE and Western blotting
Cells cultured under different conditions were fractionated by SDS-PAGE and immunoblotted as described previously , . The following antibody dilutions were used: 8H4 (1:3000), 3F4 (1:5000), ferritin (1:1000), Tf (1:6000), TfR (1:3000), actin (1:7500), secondary antibodies conjugated with horseradish peroxidase (1:6000). Immunoreactive bands were visualized by ECL (Amersham Biosciences Inc.).
Measurement of intracellular calcein-chelatable iron
Cellular labile iron pool (LIP) was assayed as described by Tenopoulou et al.  using the iron sensitive fluorescent dye calcein. When incubated with cells as a lipophilic calcein-AM-ester (molecular probes), it enters the cells and is cleaved by cellular esterases to release calcein that binds iron and is quenched by this reaction. Upon addition of the cell permeable iron chelator salicylaldehyde isonicotinoyhydrazone (SIH), iron is released from calcein that regains its fluorescence (recorded at λex 488 nm and λem 518 nm). Briefly, 5×105 M17 cells or cell lines expressing PrPC and mutant PrP forms plated in 35 mm Petri dishes were washed with PBS containing 1 mg/ml BSA and 20 mM Hepes, pH 7.3 and incubated with 0.25 µM calcein-AM for 20 min at 37°C in same buffer. After calcein loading, cells were trypsinized, washed and re-suspended in 1.0 ml of the above buffer without calcein-AM and placed in a 24 well micro-plate in a thermostatically controlled (37°C) fluorescence plate reader (Microtek). The fluorescence was monitored at λex 488 nm and λem 518 nm. Iron-induced quenching of calcein was reduced by the addition of 20 µM SIH. Cell number and viability was checked by Trypan Blue dye exclusion and results were expressed as ΔF/106 cells.
Detection of iron with Ferene S
Cell lines cultured overnight in complete medium or in the presence of 0.1 mM ferric ammonium citrate (FAC) were washed with PBS supplemented with EDTA to chelate surface bound iron and pelleted. The pellet was dissolved in 50 µl of acetic acid and equal amount of protein (50 µg) was spotted on a PVDF membrane and immersed in a freshly prepared solution of Ferene S (0.75 mM 3-[2-pyridyl]-5, 6-bis(2-[-furyl sulfonic acid]-2, 4-triazine, 2% (v/v) acetic acid, 0.1% thioglycolic acid) (24) for 30 minutes at 37°C. Ferene reacts with iron in the presence of acetic acid and thioglycolic acid to form a dark blue complex. Stained membranes were de-stained with 2% acetic acid and scanned.
Stimulation of endocytosis with 3F4 antibody
M17 and PrPC cells were cultured in DMEM supplemented with 5% FBS and 1% PSF at 37°C in a humidified atmosphere in absence or presence of 1 µg/ml of 3F4 for 5 days , . Medium containing 3F4 was replaced every 2nd day and care was taken to make sure that the cells did not achieve confluency. On the 5th day, cells were washed and incubated with serum free DMEM for 1 h, followed by radiolabeling with 59FeCl3-citrate complex in DMEM for 4 h as above. In a separate experimental paradigm, N2a, M17 and PrPC cells were radiolabeled as above in the presence of 12 µg/ml of 3F4 or Thy-1 4 h. After labeling, cells were washed, lysed in native lysis buffer, and analyzed as above.
Immunostaining and fluorescence microscopy
Cell lines subjected to different experimental conditions were processed for immunostaining as described in a previous report .
Estimation of iron export from cells
Cell lines expressing different PrP forms were radiolabeled with 59FeCl3-citrate complex as above. Cell surface bound iron was chelated with 3 washes of PBS supplemented with DFO (100 µM) and the cells were chased in complete medium for different time periods. A 50 µl aliquot of the medium was retrieved at each time point and counted in a γ-counter. After 16 h, cells were lysed and cell associated iron was measured in a gamma counter.
Estimation of ferroxidase activity of recombinant PrP
Ferroxidase activity of PrP was measured by the published colorimetric method using 3-(2-pyridyl)-5,6-bis(2-[5furylsulfonic acid])-1,2,4-triazine that forms a colored Fe2+ complex with ferrous iron (44) with the following modifications: Reagent A: 0,45 mol/l sodium acetate, pH 5.8, reagent B: 130 mmol/l thiourea, 367 µM/l Fe(NH4)(SO4)2×6H2O, reagent C (chromogen): 18 mmol/l 3-(2-pyridyl)-5,6-bis(2-[5-furylsulfonic acid])-1,2,4-triazine in 0.01 M Tris pH 7.0. Each sample contained either 1 µl of water or 1 µl of 300 µM CuSO4, 6 µL of the sample (undiluted human plasma, human serum albumin 70 g/l (Sigma A1653-5G) in PBS or recombinant prion protein (0.6 µg/ml) and 820 µl of reagent A. Multichannel pipette (Finnpipette) was used for the rapid addition of the reagent B (substrate) to minimize the time difference in sample processing. Sample quadruplicates were incubated at 37°C for 4 min. Unoxidized Fe2+ was reacted with 60 µl of chromogen solution (reagent C) and absorbance was measured at 600 nm with Smart Spec Plus (BioRad) spectrophotometer. Copper was added to provide two copper ions per PrP molecule, and was also added to human albumin and plasma samples. The amount of PrP protein in PrP-containing samples (3.6 µg/sample) roughly corresponds to a known amount of ceruloplasmin in 6 µl of undiluted human plasma. As a control, purified 99% human serum albumin was used (70 g/l in PBS) to mimic the total protein concentration in plasma. As a blank samples were supplemented with 6 µl of de-ionized water instead of albumin solution, plasma or recombinant PrP solution.
RNA Isolation and Northern blotting
M17 and WT cells cultured in the absence or presence of 0.1 mM FAC for 24 h were washed with cold PBS, trypsinized, and collected in 1.5 ml eppendorf tubes. Total RNA was isolated by using SV total RNA isolation kit (Promega, Madison, WI) and quantified. 15 µg of total RNA was fractionated on 0.8% formaldehyde agarose gel followed by blotting to positively charged Nylon membranes (Roche diagnostics). Membranes were hybridized with DIG-labeled PrP or β-actin probes and binding was detected by the CSPD reagent.
Data are presented as the mean±SEM values. Statistical evaluation of the data was performed by using Students t-test (unpaired).
(A) Apotransferrin (Sigma) was radiolabeled with 59FeCl3-citrate complex and resolved on a native gel as in Fig. 2. Tf migrates as three distinct bands representing different conformational forms. (B) Autoradiographed gel from Figure 2 was re-hydrated and stained with silver to ensure equal loading of proteins (Beta-actin does not resolve on this native gel).
(7.48 MB TIF)
Lysates of PrPC cells labeled with 59FeCl3-citrate complex were resolved on native gel as in Fig. 1 and exposed to an X-ray film to visualize iron labeled bands (panel A). Marked areas were excised from the wet gel, proteins were electro-eluted, and resolved by SDS-PAGE followed by sequential immunoblotting with antibodies specific to PrP, TfR, ferritin, and Tf (panel B). Finally, the membrane was stained with silver to visualize all proteins (panel B). Band 1 that includes proteins in the loading well reacts strongly for PrP and TfR. Band 3 reacts specifically for ferritin, while band 5 represents Tf. No detectable proteins are present in band 4. Silver staining shows 4 prominent proteins in bands 1–3, the identity of which is currently unknown.
(10.07 MB TIF)
(A) Lysates of M17 and PrPC-cells treated as in Figure 5 were fractionated by SDS-PAGE and transferred proteins were probed for PrP, ferritin, TfR, and β-actin (lanes 1–4). (B) FAC exposed M17 and PrPC-cells show up-regulation of PrP mRNA compared to untreated controls (lanes 1–4).
(2.03 MB TIF)
M17-cells exposed to buffer, 3F4, and anti-Thy-1 antibody were radiolabeled with 59FeCl3-citrate complex and lysates were resolved on native gel followed by autoradiography (lanes 1–3). Equal aliquots of the same samples were resolved by SDS-PAGE followed by immunoblotting for β-actin to ensure equal loading of protein (lanes 1–3).
(1.00 MB TIF)
Conceived and designed the experiments: NS. Performed the experiments: AS MLM AOI XL JP NS. Analyzed the data: AS MLM AOI JP DV NS. Contributed reagents/materials/analysis tools: DV. Wrote the paper: NS.
- 1. Caughey B, Baron GS (2006) Prions and their partners in crime. Nature 443: 803–810.
- 2. Aguzzi A, Heikenwalder M (2006) Pathogenesis of prion diseases: current status and future outlook. Nat Rev Microbiol 4: 765–775.
- 3. Tatzelt J, Schätzl HM (2007) Molecular basis of cerebral neurodegeneration in prion diseases. FEBS J 274: 606–611.
- 4. Saá P, Castilla J, Soto C (2006) Ultra-efficient replication of infectious prions by automated protein misfolding cyclic amplification. J Biol Chem 281: 35245–35252.
- 5. Deleault NR, Harris BT, Rees JR, Supattapone S (2007) Formation of native prions from minimal components in vitro. Proc Natl Acad Sci USA 104: 9741–9746.
- 6. Aguzzi A, Heikenwalder M, Polymenidou M (2007) Insights into prion strains and neurotoxicity. Nat Rev Mol Cell Biol 8: 552–561.
- 7. Harris DA, True HL (2006) New insights into prion structure and toxicity. Neuron 50: 353–357.
- 8. Mallucci G, Dickinson A, Linehan J, Klohn PC, Brandner S, et al. (2003) Depleting neuronal PrP in prion infection prevents disease and reverses spongiosis. Science 302: 871–874.
- 9. Chesebro B, Trifilo M, Race R, Meade-White K, Teng C, et al. (2005) Anchorless prion protein results in infectious amyloid disease without clinical scrapie. Science 308: 435–439.
- 10. Jeffrey M, Goodsir CM, Race RE, Chesebro B (2004) Scrapie-specific neuronal lesions are independent of neuronal PrP expression. Ann Neurol 55: 781–792.
- 11. Roucou X, LeBlanc AC (2005) Cellular prion protein neuroprotective function: implications in prion diseases. J Mol Med 83: 3–11.
- 12. Westergard L, Christensen HM, Harris DA (2007) The cellular prion protein (PrP(C)): its physiological function and role in disease. Biochim Biophys Acta 1772: 629–644.
- 13. Brown DR, Qin K, Herms JW, Madlung A, Manson J, et al. (1997) The cellular prion protein binds copper in vivo. Nature 390: 684–687.
- 14. Pauly PC, Harris DA (1998) Copper stimulates endocytosis of the prion protein. J Biol Chem 273: 33107–33110.
- 15. Kim NH, Park SJ, Jin JK, Kwon MS, Choi EK, et al. (2000) Increased ferric iron content and iron-induced oxidative stress in the brains of scrapie-infected mice. Brain Res 884: 98–103.
- 16. Hur K, Kim JI, Choi SI, Choi EK, Carp RI, et al. (2002) The pathogenic mechanisms of prion diseases. Mech Ageing Dev 123: 1637–1647.
- 17. Basu S, Mohan ML, Luo X, Kundu B, Kong Q, et al. (2007) Modulation of proteinase K-resistant prion protein in cells and infectious brain homogenate by redox iron: implications for prion replication and disease pathogenesis. Mol Biol Cell 18: 3302–3312.
- 18. Moos T, Morgan EH (2004) The metabolism of neuronal iron and its pathogenic role in neurological disease. Ann N Y Acad Sci 1012: 14–26.
- 19. MacKenzie EL, Iwasaki K, Tsuji Y (2001) Intracellular iron transport and storage: from molecular mechanisms to health implications. Antioxid Redox Signal 10: 997–1030.
- 20. Petrak JV, Vyoral D (2001) Detection of iron-containing proteins contributing to the cellular labile iron pool by a native electrophoresis metal blotting technique. J Inorg Biochem 86: 669–675.
- 21. Vyoral D, Petrák J, Hradilek A (1998) Separation of cellular iron containing compounds by electrophoresis. Biol Trace Elem Res 61: 263–275.
- 22. Rogers M, Yehiely F, Scott M, Prusiner SB (1993) Conversion of truncated and elongated prion proteins into the scrapie isoform in cultured cells. Proc Natl Acad Sci USA 90: 3182–3186.
- 23. Campana V, Caputo A, Sarnataro D, Paladino S, Tivodar S, et al. (2007) Characterization of the properties and trafficking of an anchorless form of the prion protein. J Biol Chem 282: 22747–22756.
- 24. Baker E, Richardson D, Gross S, Ponka P (1992) Evaluation of the iron chelation potential of hydrazones of pyridoxal, salicylaldehyde and 2-hydroxy-1-naphthylaldehyde using the hepatocyte in culture. Hepatology 15: 492–501.
- 25. Chung MC (1985) A Specific iron stain for iron-binding proteins in polyacrylamide gels: application to transferrin and lactoferrin. Anal Biochem 148: 498–502.
- 26. Kascsak RJ, Rubenstein R, Merz PA, Tonna-DeMasi M, Fersko R, et al. (1987) Mouse polyclonal and monoclonal antibody to scrapie-associated fibril proteins. J Virol 61: 3688–3693.
- 27. Peretz D, Williamson RA, Kaneko K, Vergara J, Leclerc E, et al. (2001) Antibodies inhibit prion propagation and clear cell cultures of prion infectivity. Nature 412: 739–743.
- 28. Chen SG, Teplow DB, Parchi P, Teller JK, Gambetti P, et al. (1995) Truncated forms of the human prion protein in normal brain and in prion diseases. J Biol Chem 270: 19173–19180.
- 29. Liu X, Theil EC (2005) Ferritin as an iron concentrator and chelator target. Ann N Y Acad Sci 1054: 136–40.
- 30. Ohgami RS, Campagna DR, Greer EL, Antiochos B, McDonald A, et al. (2005) Identification of a ferrireductase required for efficient transferrin-dependent iron uptake in erythroid cells. Nat Genet 37: 1264–1269.
- 31. Burdo JR, Connor JR (2003) Brain iron uptake and homeostatic mechanisms: an overview. Biometals 16: 63–75.
- 32. Moos T, Rosengren Nielsen T, Skjørringe T, Morgan EH (2007) Iron trafficking inside the brain. J Neurochem 103: 1730–1740.
- 33. Aguirre P, Mena N, Tapia V, Arredondo M, Núñez MT (2005) Iron homeostasis in neuronal cells: a role for IREG1. BMC Neurosci 6: 3.
- 34. Brown LR, Harris DA (2003) Copper and zinc cause delivery of the prion protein from the plasma membrane to a subset of early endosomes and the Golgi. J Neurochem 87: 353–363.
- 35. Miura T, Sasaki S, Toyama A, Takeuchi H (2005) Copper reduction by the octapeptide repeat region of prion protein: pH dependence and implications in cellular copper uptake. Biochemistry 44: 8712–8720.
- 36. Shyng SL, Moulder KL, Lesko A, Harris DA (1995) The N-terminal domain of a glycolipid-anchored prion protein is essential for its endocytosis via clathrin-coated pits. J Biol Chem 270: 14793–14800.
- 37. Peters PJ, Mironov A Jr, Peretz D, van Donselaar E, Leclerc E, et al. (2003) Trafficking of prion proteins through a caveolae-mediated endosomal pathway. J Cell Biol 162: 703–717.
- 38. Mishra RS, Gu Y, Bose S, Verghese S, Kalepu S, et al. (2002) Cell surface accumulation of a truncated transmembrane prion protein in Gerstmann-Straussler-Scheinker disease P102L. J Biol Chem 277: 24554–24561.
- 39. Choi CJ, Anantharam V, Saetveit NJ, Houk RS, Kanthasamy A, et al. (2007) Normal cellular prion protein protects against manganese-induced oxidative stress and apoptotic cell death. Toxicol Sci 98: 495–509.
- 40. Zivny JH, Gelderman MP, Xu F, Piper J, Holada K, et al. (2008) Reduced erythroid cell and erythropoietin production in response to acute anemia in prion protein-deficient (Prnp−/−) mice. Blood Cells Mol Dis 40: 302–307.
- 41. Gu Y, Fujioka H, Mishra RS, Li R, Singh N (2002) Prion peptide 106–126 modulates the aggregation of cellular prion protein and induces the synthesis of potentially neurotoxic transmembrane PrP. J Biol Chem 277: 2275–2286.
- 42. Jin T, Gu Y, Zanusso G, Sy M, Kumar A, et al. (2000) The chaperone protein BiP binds to a mutant prion protein and mediates its degradation by the proteasome. J Biol Chem 275: 38699–38704.
- 43. Tenopoulou M, Kurtz T, Doulias PT, Galaris D, Brunk UT (2007) Does the calcein-AM method assay the total cellular ‘labile iron pool’ or only a fraction of it? Biochem J 403: 261–266.
- 44. Erel O (1998) Automated measurement of serum ferroxidase activity. Clin Chem 44: 2313–2319.
- 45. Singh A, Isaac AO, Luo X, Mohan ML, Cohen ML, et al. (2009) Abnormal brain iron homeostasis in human and animal prion disorders. Plos Pathogens, in press.