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1 Medical Biotechnology Center, University of Maryland Biotechnology Institute, Baltimore, Maryland 21201, USA2 Department of Biochemistry and Molecular Biology, University of Maryland School of Medicine, Baltimore, Maryland 21201, USA3 Institute for Animal Health, Compton, Newbury, Berkshire, RG20 7NN, United Kingdom
Reprint requests to: Ilia V. Baskakov, 725 W. Lombard Street, Baltimore, MD 21201, USA; e-mail: Baskakov{at}umbi.umd.edu; fax: (410) 706-8184.
(RECEIVED October 20, 2004; FINAL REVISION January 17, 2005; ACCEPTED January 23, 2005)
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Abstract |
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-sheet-rich fibrillar structure. Full-length recombinant PrP 23-230, however, generates two subpopulations of amyloid in vitro: One is similar to the minor subpopulation of PrPSc, and the other to classical PrPSc. Since no cellular factors or templates were used for generation of the amyloid fibrils in vitro, we speculate that formation of the subpopulation of PrPSc with a short PK-resistant C-terminal region reflects an intrinsic property of PrP rather than the influence of cellular environments and/or cofactors. Our work significantly increases our understanding of the biochemical nature of prion infectious agents and provides a fundamental insight into the mechanisms of prions biogenesis. Keywords: prion protein; amyloid fibrils; conformational transition; proteinase K; Creutzfeldt-Jakob disease
Abbreviations: CJD, Creutzfeldt-Jakob disease • spCJD, sporadic CJD • PrP, prion protein • PrPC, the normal, cellular isoform of PrP • PrPSc, the abnormal, infections isoform of PrP • rPrP, recombinant PrP • rPrP 89-230, recombinant PrP encompassing residues 89–230 • rPrP 23-230, full-length recombinant PrP • 
-rPrP 89-230, -helical isoform of rPrP 89-231 • -rPrP 23-230, -helical isoform of rPrP 23-231 • PK, proteinase K • PrP 27-30, PK-resistant core of classical PrPSc • ThT, Thioflavin T • GdnHCl, guanidinium hydrochloride • FTIR; Fourier transform infrared spectroscopy
Article and publication are at http://www.proteinscience.org/cgi/doi/10.1110/ps.041186605.
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Introduction |
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TOPAbstractIntroductionResults and DiscussionMaterials and methodsReferences |
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PrPC and PrPSc differ substantially in their conformations. Unlike PrPC, PrPSc is a multimeric assembly characterized by enhanced resistance to proteinase K (PK) digestion and by an increase in the amount of -sheet structure (Caughey et al. 1991; Pan et al. 1993). The detailed three-dimensional structure of the PrPC isoform has been determined (Riek et al. 1996, 1997; Donne et al. 1997; Haire et al. 2004), whereas structural information about PrPSc is more limited due to its insolubility, heterogeneity, and highly aggregated state (Gabizon and Prusiner 1990; Wille et al. 2002).
In contrast to PrPC, PrPSc exists in multiple conformations, which are believed to result from infection of animals with different strains of TSE (transmissible spongiform encephalopathy) agent. The existence of different prion strains has been recognized for many years (Dickinson et al. 1968; Fraser and Dickinson 1973), but has only recently been linked to the conformational diversity of PrPSc. According to the template-assisted model, each conformation of PrPSc can act as a unique template, providing conformational constraints for the formation of nascent PrPSc and, therefore, determining the conformation of nascent PrPSc (Cohen et al. 1994). Biochemical assays have been described that distinguish conformers of PrPSc by the extent of PK resistance, the size of the PK-resistant core, thermodynamic stability, epitope presentation, and relative amount of -sheet-rich structure (Bessen and Marsh 1994; Telling et al. 1996; Caughey et al. 1998; Safar et al. 1998; Peretz et al. 2001). Conformationally different subtypes of PrPSc have also been found in patients with sporadic CJD (spCJD) (Parchi et al. 1999; Hill et al. 2003). These PrPSc subtypes have variations in the molecular mass of the PK-resistant core and exhibit different ratios of di-, mono-, and unglycosylated PrPSc. Conformational diversity of PrPSc subtypes is linked to the methionine/valine polymorphism at codon 129 and may also be affected by unidentified host factors (Telling et al. 1995; Parchi et al. 2000).
To distinguish different conformers and to classify subtypes of PrPSc, limited digestion with PK has been used widely since it offers a rapid and accurate assay of altered PrPSc conformation (McKinley et al. 1983; Bessen and Marsh 1994; Safar et al. 1998; Parchi et al. 1999; Hill et al. 2003). Normally, treatment of PrPSc with PK generates a C-terminal PK-resistant core, referred to as PrP 27-30 (McKinley et al. 1983; Oesch et al. 1985). Depending on the subtype of PrPSc, PrP 27-30 exhibits minor variations in gel mobility due to differences in the PK cleavage site between residues 79 and 103 (Parchi et al. 2000). In addition, two novel PK-resistant fragments were identified recently in patients with spCJD (Zou et al. 2003). Deglycosylated forms of these novel PK-resistant fragments migrate at 13 and 12 kDa by SDS-PAGE, and are generated by cleavage at residues 154/156 and 162/167, respectively, retaining the C-terminal region intact (Zou et al. 2003). Multiple factors including gene polymorphism, glycosylation, and as yet unidentified cellular factors may contribute to the broad conformational diversity of PrPSc. It is therefore important to know whether this conformational diversity reflects an intrinsic property of PrP or arises only with assistance from other molecules in a cell. Is unglycosylated recombinant PrP (rPrP) capable of mimicking any known PrPSc subtypes in the absence of the cellular environment or templating?
To address this question we investigated the amyloid form of mouse (Mo) rPrP spanning residues 89–230 (rPrP 89-230) by use of limited proteolysis with PK. In our previous studies, we showed that in vitro conversion of rPrP to the amyloid form in the absence of the cellular environment exhibited peculiar features of the autocatalytic process and was limited by a species barrier, two key aspects of prion propagation (Baskakov 2004). Furthermore, we demonstrated that the amyloid form of Mo rPrP 89-230 generated in vitro induces a transmissible form of prion disease in transgenic mice expressing mouse PrP 89-230 (Baskakov et al. 2002; Legname et al. 2004). Long incubation times observed upon inoculation of the amyloid fibrils suggest that the fibrils generated in vitro have very low infectivity titers. However, shortening of the incubation times observed in the subsequent passages of the synthetic prions in transgenic and wild-type mice allows an alternative explanation. One may speculate that the synthetic prions undergo adaptation upon propagation in vivo, and that the differences in the incubation time are attributed to an apparent transmission barrier. Consistent with this hypothesis are several lines of evidence including biochemical and neuropathological features that indicate that the synthetic prions induced a novel strain of prion disease in experimental animals. Encouraged by these findings, we wished to determine whether the amyloid fibrils, produced in vitro without other cellular components or PrPSc-templating, structurally resembled any known isoforms or subtypes of PrPSc.
Here we demonstrate that that the amyloid fibrils closely resemble the novel subpopulation of PrPSc associated with spCJD. Using PK digestion we identified three major PK-resistant fragments in the amyloid form. These fragments migrated at about 12 kDa, 10 kDa, and 8 kDa by SDS-PAGE and encompassed residues 138/141–230, 152/153–230, and 162–230, respectively, as determined by mass spectrometry. This suggests that the slow propagation rate observed in the first passage and an apparent transmission barrier are due to a proteolytically labile region encompassing residues 89–138. Remarkably, upon digestion with PK, short C-terminal fragments remained assembled in fibrillar structures, maintained high -sheet content, and preserved their seeding activity. Since no cellular factors and templates were used to generate the amyloid form in vitro, we suggest that the formation of the novel subpopulation of PrPSc with this particular PK-resistant core reflects an intrinsic property of PrP.
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Results and Discussion |
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). These results demonstrate that either high concentrations of Gd-nHCl or increased temperatures and intermediate GdnHCl concentrations are required for denaturation of amyloid form. In contrast, -monomeric form of rPrP can be denatured by moderate concentrations of GdnHCl (C1/2 = 2.2 M at room temperature) (Swietnicki et al. 1998). Remarkably, the C1/2 value for denaturation of amyloid fibrils generated in vitro (4.2 M) was identical to the C1/2 value determined for the mouse synthetic prion strain 1 (MoSP1) produced in the brains of Tg9949 mice after inoculation of amyloid fibrils (Legname et al. 2005). Increased resistance toward denaturation is consistent with the formation of a larger solvent-protected core during the transition from monomer into amyloid conformation.
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, left panel). As the PK to protein ratio increased, the size of PK-resistant polypeptides was reduced. Thus, digestion at 1:2000 PK/rPrP ratio produced primarily fragments with apparent molecular masses of 12 kDa and 10 kDa (lane 2), while at a higher ratio (up to 1:20) the major PK-resistant fragments were 10-kDa and 8-kDa peptide (lane 5). The PK/rPrP ratio of 1:20 exceeds the PK-to-protein ratios that are normally used in studies of PrPSc in brain homogenates (Caughey et al. 1998; Safar et al. 1998). Moreover, our PK digestion assay is free from nonprotein components of brain homogenate and from possible interference with PK activity. It is noteworthy that when SDS-sample buffer was not supplemented with urea, we observed stalled bands on the top of the stacking gel instead of 10-kDa and 8-kDa bands. Because treatment under highly denatured conditions, including 3M urea, 2% SDS, and 95°C was necessary for analyses of proteolytic products by SDS-PAGE, it is likely that the PK-resistant core of the amyloid form remained assembled in a highly stable, multimeric form.
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-rPrP 89-230 that may copresent in the preparation of the amyloid fibrils, we treated -rPrP 89-230 with PK using identical digestion conditions. At low concentrations of PK (PK/rPrP ratios 1:2000 and 1:500), -rPrP 89-230 generated two major fragments having apparent molecular masses of 14 kDa and 13 kDa (Fig. 2A
, middle panel, lanes 2,3, respectively). At higher concentrations of PK (PK/rPrP ratios 1:100 and 1:20), these fragments were completely digested (lanes 4,5, respectively). The result of this control experiment is consistent with previous studies that demonstrated that PrPC undergoes proteolytic cleavage at amino acid residues 110/111 to produce a C-terminal fragment referred to as C1 (Chen et al. 1995). Taken together, our studies shows that 8-kDa and 10-kDa fragments are generated from the amyloid fibrils but not from -rPrP 89-230.
To identify the region that is resistant to PK digestion in the amyloid conformation, we used Fabs P and R1, specific to residues 96–105 and 225–230, respectively. The epitope for Fab P was not present in any of the three major PK-resistant products, while the epitope for Fab R1 was present in all three fragments. Using liquid chromatography/mass spectrometry (LC-MS) we showed that the three major PK-resistant bands correspond to polypeptides spanning residues 138/141–230, 152/153–230, and 162–230 (Fig. 2B). Both LC-MS and Western blot analyses demonstrate that the C-terminal region of rPrP 89-230 remains PK-resistant in the amyloid form, while the central region encompassing residues 89–138 is fully susceptible to proteolysis (Fig. 3A).
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;Zou et al. 2003). These fragments were found in 26 of the 29 spCJD brains examined and accounted for up to 24% of all PK-resistant PrP polypeptides in spCJD brains. Chen and coauthors suggested that these short C-terminal fragments were generated from a novel subpopulation of PrPSc that adopts a conformation different from that of "classical" PrPSc (Zou et al. 2003), which is known to produce a core encompassing residues ~90–230 upon PK digestion (for clarity, the minor subpopulation of PrPSc produced in spCJD will be referred to as novel PrPSc). Another group reported that C-terminal PK-resistant fragments of 11–12 kDa were also associated with a certain type of iatrogenic CJD; however, the exact positions of PK cleavage sites were not determined (Satoh et al. 2003). Truncated, PK-resistant C-terminal polypeptides of similar length were found in brain homogenate from mice infected with mouse-passaged hamster PrPSc strain 263K and in cell-free conversion assays under conditions that mimicked the hamster/mouse species barrier (Lawson et al. 2004). The sites of PK cleavage were found to be between residues 130 and 157 as localized by epitope mapping and mobility on a gel (Fig. 3C). Identification of novel C-terminal PK-resistant fragments in patients with sporadic and iatrogenic CJD as well as in mice suggested that their involvement in the pathogenic process may be more common than had previously been recognized (Satoh et al. 2003; Zou et al. 2003; Lawson et al. 2004). On the basis of limited proteolysis, the amyloid form of rPrP 89-230 is similar to the novel rather than classical subpopulations of PrPSc.
Multiple lines of evidence indicate that the central region of PrP, encompassing residues 90–141, is important for prion propagation (Tagliavini et al. 1993; Fischer et al. 1996; Peretz et al. 1997; Chabry et al. 1998). Puzzled by the fact that this region is PK-labile in disease-producing amyloid fibrils and, therefore, expected to undergo rapid digestion by endogenous proteases upon inoculation (Yadavalli et al. 2004), we sought to determine whether the C-terminal PK-resistant core maintains -sheet-rich structure and possesses self-propagating activity. To address these questions we seeded the in vitro conversion reaction with the amyloid form pretreated with PK. For this experiment we used experimental conditions at which spontaneous or nonseeded formation of the amyloid fibrils occurred only after a prolonged lag phase of 60 h (Table 1). Seeding of the in vitro reaction with fibrils not treated with PK substantially reduced the lag phase (Fig. 4A), and the data indicate a linear dependence between the length of the lag phase and the log[amount of seed] (Fig. 4B). Remarkably, the fibrils pre-treated with PK for 1 h at 37°C showed substantial seeding activity. As judged from the length of the lag phase, 2.5% of amyloid fibrils pretreated with PK at 1:20 PK/rPrP ratio exhibited apparent seeding activities that were equivalent to the seeding activities that would be displayed by 0.75% of intact fibrils (Fig. 4A,B; Table 1). At a PK:rPrP ratio of 1:20, only the 10-kDa and 8-kDa fragments, corresponding to residues 152/153–230 and 162–230, respectively, were detectable by Western blotting (Fig. 2A). This experiment illustrates that the C-terminal PK-resistant core of the synthetic prions preserves propagating activity in a cell-free conversion assay.
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). PK-treated fibrils had a higher tendency to coaggregate, probably due to an increase in accessibility of hydrophobic surfaces to the solvent upon PK digestion. Unfortunately, aggregation of PK-treated fibrils into large clumps hampered quantitative analyses of the fibril width.
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-rPrP 89-230 was dominated by strong absorbance at 1651 cm–1 and 1645 cm–1 corresponding to -helices and random coil, respectively (Fig. 5B). The FTIR spectrum of the amyloid form was substantially different from that of the -monomer. The second derivative analysis of the amyloid form revealed a major band at 1622 cm–1, characteristic of -sheet structures with strong intermolecular hydrogen bonds, a smaller band at 1651 cm–1, indicative of -helices and random coil, and a minor band at 1674 cm–1, characteristic of -turns and loops (Fig. 5C). Upon treatment with PK, the relative intensity of bands at 1651 cm–1 and 1674 cm–1 decreased, demonstrating loss of -helical structure and -turns and loops, respectively. The relative intensity of band at 1622 cm–1, a characteristic of -sheet structures, did not change notably. On the other hand, the shift of the band centered at 1622 cm–1 to 1626 cm–1 indicated that -structures with strong intermolecular hydrogen bonds acquired some flexibility upon treatment with PK. Since no cellular factors or templates were used for generation of the amyloid fibrils in vitro, the ability of rPrP 89-230 to adopt a conformation similar to that of novel PrPSc reflects an intrinsic property of PrP that is independent of the influence of cellular environments and/or cofactors. While the N-terminal region of PrP, encompassing residues 23–90, is not critical for transmission of prions, this region is known to influence the conformation of protease-resistant PrP isoforms (Wadsworth et al. 1999; Lawson et al. 2004), and may substantially impact the conformational diversity of fibrils generated in vitro. Therefore, we examined the conformation of the amyloid form generated from murine full-length recombinant PrP (rPrP 23-230).
Conversion of rPrP 23-230 to the amyloid fibrils was carried out using the same solvent conditions as those used for the conversion of rPrP 89-230. FTIR analysis of rPrP 23-230 refolded into the amyloid form produced a spectrum substantially different from that of the corresponding -monomer (Fig. 6A). The spectrum of the amyloid form showed a substantial decrease in intensity of the band at 1654 cm–1 corresponding to -helices and random coil, and an increase in intensity of the band at 1617 cm–1, an indication of -structure with strong intermolecular hydrogen bonds (Fig. 6B).
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, left panel). The major subpopulation of rPrP 23-230 produces PK-resistant fragments of 12 kDa, 10 kDa, and 8 kDa similar to those identified from the PK digest of the amyloid of rPrP 89-230. These fragments appear at low concentrations of PK and show the same pattern of epitope presentation as proteolytic fragments of the amyloid of rPrP 89-230 (Fig. 7B, left, middle, and right panels). The minor subfraction of rPrP 23-230, however, produces several bands with molecular masses between 21 and 16 kDa. These bands account for ~5%–10% of all PK-resistant products, as judged by SDS-PAGE (Fig. 7A, left panel). At PK:rPrP ratios of 1:100 and 1:50, these bands are further digested to generate a single PK-resistant product with molecular mass of 16 kDa (Fig. 7A, left panel, lanes 6,7). The 16-kDa fragment is consistently resistant to PK even upon incubation at the PK:rPrP ratio of 1:10 at 37°C for 1 h. This polypeptide contains epitopes for Fab P (residues 96–105), Fab D18 (residues 133–157), and Fab R1 (residues 225–230) (Fig. 7B). As a control to confirm that 8-kDa, 10-kDa, and 16-kDa PK-resistant fragments are generated from the amyloid fibrils, we used -rPrP 23-230. At PK/rPrP ratios of 1:100 and 1:50, -rPrP 23-230 was completely digested with no PK-resistant fragments being detected (Fig. 7A, left panel, lanes 6,7).
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). The majority of fibrils display relatively weak Thioflavin T (ThT)-fluorescence as seen by fluorescence microscopy (Fig. 8A). A minor subpopulation of fibrils (5%–10%), however, exhibits substantially brighter emission, which remains stable even after harsh PK treatment (Fig. 8A). This result is consistent with the two conformers identified by limited digestion in Figure 6. Two subpopulations of the fibrils were also seen by electron microscopy, where the minor subpopulation accounted for ~5% of all fibrils (Fig. 8B).
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). The major subpopulation is more sensitive to PK digestion and produces short C-terminal PK-resistant fragments similar to those generated from novel PrPSc. The minor subpopulation is characterized by enhanced resistance to PK and possesses a C-terminal resistant core identical to the PK-resistant fragments of classical PrPSc.
It is notable that conformers biochemically identical to both novel and classical PrPSc can be generated in vitro without any cellular cofactors or PrPSc-dependent templating. To date, the relationship between the novel and classical PrPSc remains speculative, but the results of proteolytic digestion of PrPSc carried out under partially denaturing conditions may provide an important link between the two forms (Kocisko et al. 1996). Caughey and coauthors showed that PK treatment of in vivo-derived hamster PrPSc in the presence of 2.5 M GdnHCl resulted in digestion of the N-terminal regions including residues 90–115 and, possibly, 90–143, while the C-terminal region remained intact (Fig. 3D;Kocisko et al. 1996). The C-terminal fragments of PrPSc generated under partially denaturing conditions (Kocisko et al. 1996) were similar to those produced from novel PrPSc under native conditions (Zou et al. 2003). If the classical and novel subpopulations of PrPSc indeed share a similar substructure, the novel subpopulation may represent a metabolic intermediate or byproduct of formation of classical PrPSc. Remarkably, the third site targeted by PK in both the amyloid fibrils and the novel subpopulation of PrPSc (residue 162) coincides with the sequence YYR (residues 162–164) (Paramithiotis et al. 2003). Epitope mapping using antibodies raised to a peptide including this sequence has shown that the epitope containing these residues is buried in the native PrPC form, but becomes solvent-exposed upon conversion to pathological PrPSc form.
Since the relative proportions of two amyloid conformers generated in vitro are opposite to that of novel and classical PrPSc in sporadic CJD, as yet unidentified cellular cofactors may be required for efficient conversion of rPrP 23-230 into a conformer similar to classical PrPSc. Such a cofactor may promote assembly by counteracting the electrostatic repulsion of positively charged N-terminal regions that remain exposed to the solvent upon conversion to the amyloid form. Polyanions such as sulfated glycans or RNA may serve this function. RNA and heparin sulfate have been shown to bind to the N-terminal region of PrPC (Gonzalez-Iglesias et al. 2002; Warner et al. 2002; Gabus et al. 2004) and, furthermore, both RNA and sulfated glycans were found to stimulate cell-free conversion of PrPC into protease-resistant PrPSc-like forms (Wong et al. 2001; Adler et al. 2003; Deleault et al. 2003).
While the amyloid fibrils formed by rPrP 89-230 were shown to cause a transmissible form of prion disease in experimental animals, the incubation times observed upon inoculation of the fibrils was longer than that exhibited by most known PrPSc strains (Legname et al. 2004). It is important to note that natural strains of PrPSc evolved through natural selection (Bartz et al. 2000). Only those strains that show very fast rates of propagation and, correspondingly, shorter incubation times are preferred by many laboratory investigators. Moreover, most strains that are currently used in laboratories were repassaged many times and therefore adapt well to a particular host. One can speculate that the shortening of the incubation times observed in the second passages of synthetic prions may be attributed to the adaptation to the host, suggesting that an apparent transmission barrier precludes efficient propagation of the amyloid fibrils in the first passage. A transmission barrier is typically observed when the sequence of PrPSc in the inoculum does not match that of PrPC in the recipient animals (Pattison and Jones 1968; Prusiner et al. 1990; Scott et al. 1993). Besides the differences in the sequences of PrPs, different strains of prion disease have different transmission barriers, presumably as a result of different conformations of PrPSc (Barron et al. 2003). Since the sequence of the amyloid fibrils was identical to that of endogenously expressed PrPC (mouse PrP 89-230), it is likely that the apparent transmission barrier is due to the unique conformational properties of the amyloid fibrils and, in particular, due to the proteolytic liability of residues 90–138. In addition, the lack of glycosylation in the amyloid fibrils may affect their conformational compatibility to PrPC and, therefore, also contribute to the slow rate of propagation in the first passage and the apparent transmission barrier. Remarkably, after primary passage was accomplished, much shorter incubation times in the second passage of the synthetic prions were observed (Legname et al. 2004).
PK resistance of the central region and, in particular, the 3F4 epitope (residues 108–111) has historically been exploited to distinguish PrPSc from PrPC as a biochemical marker for the presence of prion infection (McKinley et al. 1983; Oesch et al. 1985). Taken together, this study and that by Legname and coauthors demonstrate that the amyloid fibrils generated in vitro are capable of transmitting a prion disease despite having a proteolytically labile region encompassing residues 90–138. These findings may explain reports of prion infections in the absence of detectable PK-resistant PrPSc (Lasmézas et al. 1997; Manuelidis et al. 1997; Barron et al. 2001; Tremblay et al. 2004). These observations have serious implications for the development of effective prion diagnostics.
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Materials and methods |
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In vitro conversion of PrPs into the amyloid fibrils
To form amyloid fibrils we used two different formats, manual and automated. In the manual format, a stock solution of rPrP 89-230 or rPrP 23-230 (4 mg/mL) in 6 M GdnHCl was diluted to the final protein concentration of 0.5 mg/mL and incubated at 37°C in 20 mM Na-acetate buffer, 1 M GdnHCl, 3 M urea, 150 mM NaCl (pH 5.0) with continuous shaking at 600 rpm using a Delfia plate shaker (Wallac) in conical plastic tubes (Eppendorf) in a 0.6-mL reaction volume. The kinetics of fibril formation were monitored using a ThT-binding assay. Aliquots withdrawn during the time course of incubation at 37°C were diluted into 5 mM Na-acetate buffer (pH 5.5) to a final concentration of rPrP of 1 µM, and then ThT (Molecular Probes) was added to a final concentration of 10µM. Six emission spectra (from 460 to 520 nm) were recorded for each sample in 0.4-cm rectangular cuvettes with excitation at 445 nm on a FluoroMax-3 fluorimeter (Jobin Yvon); both excitation and emission slits were 4 nm. Spectra were averaged and the fluorescence intensity at emission maximum (482 nm) was determined. In a typical experiment, we observed a 50-fold increase in ThT-fluorescence per 1 µM of rPrP 89-230 upon conversion into the amyloid form.
To monitor the kinetics of amyloid formation in seeding experiments, we used the automated format. The conversion in the automated format was carried out in PBS buffer, 1 M GdnHCl, 3 M urea (pH 7.0) in a 0.15-mL volume in 96-well plates and in the presence of ThT (10 µM). Our preliminary studies using the manual format demonstrated that the presence of 10 µM ThT in the reaction mixture does not interfere with the kinetics of amyloid formation (data not shown). Three Teflon spheres (2.38-mm diameter, McMaster-Carr) were placed into each well of a 96-well plate; then the reaction mixture containing rPrP and ThT was pipetted into wells, and the plates were covered by ELAS septum sheets (Spike International), and incubated at 37°C upon continuous shaking at 900 rpm in a Fluoroskan Ascent CF microplate reader (ThermoLabsystems). The kinetics were monitored by bottom reading of fluorescence intensity every 5 min using 444 nm excitation and 485 nm emission filters.
The lag phase of amyloid formation was determined by fitting the time-dependent changes in the fluorescence of ThT (F) over time of the reaction (t) to the following equation:
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where A is the base level of ThT fluorescence during the lag phase, B is the difference between final level of ThT fluorescence at plateau and the initial level during the lag phase, k is the rate constant of fibril growth (h–1), and tm is the observed time at midpoint of transition. The lag time (tl) of fibril formation was calculated as: tl = tm–2/k. The apparent seeding activities in the reactions seeded with 2.5% of fibrils pretreated with PK were calculated using the linear dependence between the length of the lag phase and Lg[amount of seed].
GdnHCl-induced denaturation of the amyloid fibrils
Mo rPrP 89-230 in amyloid conformation (7 µL, 0.4 mg/mL) was suspended in PBS (30 µL, pH 7.2) containing GdnHCl (0.5–5.5 M). This solution was incubated for 30 min at a fixed temperature (24°C or 57°C) and then diluted to 300 µL with 8 M GdnHCl and water to adjust the final concentration of GdnHCl to 0.55 M. Fluorescence spectra were recorded in the presence of 10 µM thioflavin T (ThT) in 0.4-cm rectangular cuvettes, with excitation at 445 nm, on a FluoroMax-3 fluorimeter (Jobin Yvon) at 24°C; both excitation and emission slits were 4 nm. ThT bound to the amyloid fibrils showed typical fluorescence spectra with the emission maximum at 482 nm. Background ThT fluorescence was subtracted. C1/2 values were calculated according to the equation C1/2 = 
G/m, where G is apparent free energy of denaturation, and m represents the GdnHCl concentration dependence of the free energy of denaturation. G and m values were calculated using least-squares fit of the data to a two-state model using the linear extrapolation method as described (Santoro and Bolen 1988).
Proteolysis with proteinase K
The amyloid forms or -helical forms of rPrP 89-230 (0.2 mg/mL) and rPrP 23-230 (0.2 mg/mL) were dialyzed and treated with PK at 37°C for 1 h in 100 mM Tris-HCl buffer (pH 7.2). Digestion was stopped by quenching with PMSF (2 mM), followed by addition of 8 M urea, to a final concentration of 3 M, and 4X sample buffer to a final concentration of 1X. Samples were heated at 95°C for 5 min and analyzed by SDS-PAGE by use of precast 12% NuPage SDS gels (Invitrogen). For Western blot experiments, proteins were electroblotted onto Immobilon P PVDF membrane (Millipore), incubated with anti-PrP Fabs (0.2 µg/mL) followed by incubation with goat anti-human IgG F(ab)2 fragment conjugated with HRP, and detected using the ECL system (Pierce).
FTIR spectroscopy
FTIR spectra were measured by means of a Bruker Tensor 27 FTIR instrument (Bruker Optics) equipped with an MCT detector cooled with liquid nitrogen. Both -rPrP and amyloid fibrils were dialyzed against 10 mM Na-acetate buffer (pH 5.5), and 10 µL of each isoform (3 mg/mL of -rPrP, or 0.3 mg/mL of the amyloid fibrils) was loaded into a BioATRcell II. Then, 128 scans at 2-cm–1 resolution were collected for each sample under constant purging with nitrogen, corrected for water vapor; background spectra of buffer were subtracted.
Mass spectrometry
Online capillary LC-MS analyses were performed at the Institute for Animal Health proteomics facility as described (Gill et al. 2000). Briefly, samples were diluted to ~1 pmol/µL in 95:5 H2O:acetonitrile with 0.05% trifluoroacetic acid (TFA) (solvent A), and ~20 pmol was injected onto a preconcentration trap and desalted by washing with the above solvent. Components were separated on a home-packed capillary HPLC column (180-µm i.d., 5-µm beads, 300 Å pore size, Jupiter C18, Phenomenex), previously equilibrated with solvent A, and were eluted by an increasing gradient of solvent B, where solvent B was 5:95 H2O:acetonitrile with 0.05% TFA. The flow rate was ~1 µL/min. The column eluent was passed directly to a Quattro II tandem quadrupole mass spectrometer (Waters UK) operated in positive ion mode. The instrument was equipped with a continuous flow nanospray source and acquired full scan mass spectra (m/z 300–2100) every 5 sec.
Negative staining and electron microscopy
Negative staining was performed on carbon-coated 100-mesh grids coated with 0.01% of poly-L-lysine solution prior to staining. The samples were adsorbed for 30 sec, washed with 0.1 M and 0.01 M Na-acetate for 5 sec each, stained with freshly filtered 2% uranyl acetate for 30 sec, dried, and then viewed in a Zeiss EM 10 CA electron microscope.
Epifluorescence microscopy
Epifluorescence microscopy was carried out on an inverted microscope (Nikon Eclipse TE2000-U) with the illumination system X-Cite 120 (EXFO Photonics Solutions) connected through fiber optics using a 1.3 aperture Plan Fluor 100x NA objective. The emission was isolated from Rayleigh and Raman-shifted light by a combination of filters: an excitation filter 485DF22, a beam splitter 505DRLPO2, and an emission filter 510LP (Omega Optical). Digital images were acquired using a cooled 12-bit CoolSnap HQ CCD camera (Photometrics). Fibrils were prestained with ThT (10 µM) for 3 min prior to imaging.
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References |
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Barron, R.M., Thomson, V., Jameison, E., Melton, D.W., Ironside, J., Will, R., and Manson, J.C. 2001. Changing a single amino acid in the N-terminus of murine PrP alters TSE incubation time across three species barriers. EMBO J. 20: 5070–5078.[CrossRef][Medline]
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) or 57°C (•) in the presence of variable concentrations of GdnHCl. The GdnHCl concentration was then adjusted to 0.55 M, followed by a ThT-binding assay. The experiments were conducted at least in duplicate. The lines represent the results of the fitting to a two-state model (
