A seven-residue deletion in PrP leads to generation of a spontaneous prion formed from C-terminal C1 fragment of PrP

Abstract

Prions result from a drastic conformational change of the host-encoded cellular prion protein (PrP), leading to the formation of β-sheet–rich, insoluble, and protease-resistant self-replicating assemblies (PrP^Sc). The cellular and molecular mechanisms involved in spontaneous prion formation in sporadic and inherited human prion diseases or equivalent animal diseases are poorly understood, in part because cell models of spontaneously forming prions are currently lacking. Here, extending studies on the role of the H2 α-helix C terminus of PrP, we found that deletion of the highly conserved ¹⁹⁰HTVTTTT¹⁹⁶ segment of ovine PrP led to spontaneous prion formation in the RK13 rabbit kidney cell model. On long-term passage, the mutant cells stably produced proteinase K (PK)–resistant, insoluble, and aggregated assemblies that were infectious for naïve cells expressing either the mutant protein or other PrPs with slightly different deletions in the same area. The electrophoretic pattern of the PK-resistant core of the spontaneous prion (Δ^Spont) contained mainly C-terminal polypeptides akin to C1, the cell-surface anchored C-terminal moiety of PrP generated by natural cellular processing. RK13 cells expressing solely the Δ190–196 C1 PrP construct, in the absence of the full-length protein, were susceptible to Δ^Spont prions. Δ^Spont infection induced the conversion of the mutated C1 into a PK-resistant and infectious form perpetuating the biochemical characteristics of Δ^Spont prion. In conclusion, this work provides a unique cell-derived system generating spontaneous prions and provides evidence that the 113 C-terminal residues of PrP are sufficient for a self-propagating prion entity.

Mammalian prions are responsible for transmissible spongiform encephalopathies in both humans and animals. Prions result from the misfolding of the host-encoded prion protein (PrP). Under its normal conformation, the cell-surface GPI-anchored PrP (PrP^C) presents a globular domain containing three α-helices and two short antiparallel β strands, preceded by an unstructured N-terminal part (1, 2). In contrast, prions are made from assemblies of β-sheet–rich, insoluble, aggregative, and mostly partially protease-resistant PrP conformers called PrP^Sc in reference to their original identification in scrapie-infected sheep (3). Prion replication appears to proceed by conversion of the normal protein through templated polymerization (4), which explains not only their propagation in tissues but also their intra- or interspecific infectivity. The high-resolution structure of PrP^Sc is not yet resolved, because of inherent difficulties in producing large amounts of purified insoluble assemblies that may nonetheless have some intrinsic heterogeneity with respect to size. Several amyloid models were proposed, and coexistence of different candidate structures has even been suggested (5–7).

The two-third C-terminal part of PrP forming the protease-resistant core of PrP^Sc, the C2 fragment, constitutes the domain necessary and sufficient for prion replication (8, 9). Prion strains are identified by their specific biochemical and/or neuropathological features in the same infected host species (10, 11). Strains result from structural differences in three-dimensional or quaternary structure of PrP^Sc. The N-terminal border of the C2 fragment is strain-dependent and can vary around amino acid positions 80–100.

A C-terminal fragment called C1 results from the natural cleavage of PrP^C by a cellular protease at the α cleavage site, which is between residues 110 and 111 of human PrP (8, 12). C1 is thus smaller than C2 and considered so far too short to be converted into prions, although it encompasses the structured globular domain of the full-length PrPC and is also present at the cell surface (13, 14).

Prions can emerge spontaneously as in sporadic cases of human Creutzfeldt–Jakob disease (CJD), that is, without evidence of infection or contamination. In this context, prion generation requires at first the formation of nuclei stable enough to initiate the polymerization process, which is expected to be a slow and rate-limiting step (15). Indeed, spontaneous prion disease is a rare event, the prevalence of sporadic CJD being of ∼1.5 cases per million per year worldwide. Mutations in PrP can favor PrP spontaneous conversion into prions. Indeed, more than 30 mutations responsible for inherited human prion diseases, including genetic CJD, Gerstmann–Sträussler–Scheinker syndrome (GSS), or fatal familial insomnia, were identified, and these dominant allelic mutations usually show a high penetrance (16). Disease-causing mutations might favor partial unfolding or transient denaturation of PrP^C, which are required for refolding into PrP^Sc, and might also increase stability of initial PrP^Sc seeds.

The cellular and molecular processes underpinning or preventing spontaneous prion generation remain poorly understood. Transgenic mouse models of spontaneous prion formation have proven difficult to obtain. This was achieved for human or mouse PrP bearing some mutations (17, 18). Hallmarks of the disease were not always reproduced in mice, and intriguingly in several instances, prions showed a rather low resistance to protease digestion (19). Prions spontaneously formed in mice overexpressing either anchorless mouse PrP or I109 allele of bank vole PrP (20, 21). Currently, no cellular model for spontaneous prion formation has been reported. Toward this goal we focused here on an intriguing highly conserved threonine-rich region of the α-helix H2 associated with several disease-causing mutations in human PrP (22). In a recent work, we demonstrated that deletion of the cluster of four threonines in the α-helix H2 C terminus has no or marginal effect on ovine prions replication in RK13 cells expressing ovine PrP (23, 24). We now show that specific deletion of the larger H2 C-terminal segment HTVTTTT, which removes three additional residues, causes the spontaneous conversion of the mutant ovine PrP into a new type of prion. This prion exhibits a main protease-resistant core shorter than usual, of C1 size, which was able to infect naïve RK13 cells expressing the mutant C1 segment alone. The potential importance of H2 C terminus for maintenance of normal PrP^C conformation in the cell, the specificities of the new mutant prion, and the surprising conversion of the homologous mutant C1 fragment into a prion entity are discussed.

Results

Δ190–196 deletion does not alter the overall structure of PrP but reduces its stability

We focused here on Δ190–196 ovine PrP (VRQ allelic variant), a mutant PrP with a specific deletion of seven amino acids at the end of helix H2 (Fig. 1). We previously reported that a larger deletion of the H2 C terminus (Δ190–197) did not have a major impact on the structure of the protein, leaving intact the spatial organization of the three α-helices in the globular domain of PrP (23). As with Δ190–197 PrP, structural analysis of recombinant Δ190–196 PrP by CD indicated a conservation of the overall α-helical content compared with WT PrP (Fig. 2A). This is in agreement with NMR analysis of the segment 113–214 of Δ190–196 PrP (C1₁₁₃), which contains the entire sequence of the structured domain and is an equivalent to the natural C1 fragment studied hereinafter. The large dispersion of amide chemical shifts observed in the ¹H-¹⁵N HSQC spectrum of ¹⁵N¹³C-labeled mutant C1₁₁₃ indicated that it maintained a globular core, in addition to its unstructured N-terminal region (Fig. 2B). Moreover, comparison with the spectra of Δ193–196 and Δ190–197 mutant PrPs previously obtained (23, 24) versus WT PrP showed that chemical shift perturbations followed a similar trend, confirming that the structure of the core domain of Δ190–196 C1₁₁₃ is structurally close to those of other mutants (Fig. 2C). Last, analysis of Δ190–196 C1₁₁₃ ¹³Cα chemical shifts yielded the position of the three α-helices within residues 147–159, 175–189, and 203–230, showing that the topology is conserved with respect to WT PrP (Fig. 2C).

Figure 1.

Δ190–196 PrP sequence. On the top is the schematic representation of mature ovine PrP^C (positions 23–234). Secondary structures building the globular part of the protein, the two short beta strands forming a beta sheet, and the three α-helices are indicated. Post-translational modifications such as N-glycan chains (black dots), the disulfide bridge (S–S), and the GPI anchor are also shown. On the bottom, the amino acid sequence of α-helix H2 is highlighted in lavender, and the first residues of H3 are in purple. Residues 190–196 that were removed from the WT PrP are colored in red and replaced by a dotted line for the deletion mutant.

Figure 2.

Structure and stability of recombinant Δ190–196 PrP. A, comparative analysis of the secondary structures of WT PrP (blue) and Δ190–196 PrP (red) by CD. Far UV spectra indicate that the secondary structure of full-length WT PrP is essentially maintained in the mutant protein. B, NMR spectroscopy analysis of the C1-like segment (positions 113–234) of Δ190–196 mutant PrP. 2D ¹H-¹⁵N HSQC spectrum of 250 μm recombinant ¹⁵N¹³C-labeled Δ190–196 C1₁₁₃, acquired at a magnetic field of 18.8 T and a temperature of 298 K. C, combined amide chemical perturbations (obtained with weighing factors of 1 for ¹H and 1/10 for ¹⁵N), measured for each nonproline residue with respect to WT PrP, are represented as superimposed bar diagrams. Perturbations are shown for Δ190–196 C1₁₁₃ (red) and compared with those previously obtained for PrP Δ193–196 (cyan) and Δ190–197 (blue) (23, 24). The positions of the three α-helices, H1, H2, and H3, obtained by analysis of Δ190–196 C1₁₁₃ ¹³Cα chemical shifts by TALOS-N (71) are shown at the top. D, comparison of stability between WT and Δ190–196 mutant PrP. Means melting temperatures (T_m) and standard deviations from five experiments were determined for full-length WT PrP (blue, 57.1 °C ± 0.9 °C) and Δ190–196 PrP (red, 49.9 °C ± 0.5 °C), for proteins resuspended in the same buffer conditions as for CD and NMR analysis (10 mm sodium acetate, pH 5).

The HTVTTTT deletion removed a histidine at position 190, which is the equivalent of His¹⁸⁷ in human PrP (Fig. S1). The pH-dependent protonation of this histidine is thought to play an important role in the electrostatic network and the stability of the globular part of PrP (25, 26). However, the deletion of the other residues might also impact the thermodynamic stability of the protein. We thus tested whether the deletion affected the stability using a thermal shift assay to determine the melting temperature of WT and mutant PrP. The melting temperature of Δ190–196 PrP (49.9 °C) was reduced by 7 °C compared with the WT PrP in sodium acetate buffer (10 mm, pH 5.0) (Fig. 2D). A marked reduction of 8 °C was also observed in these assays, using a different condition, sodium phosphate buffer (250 mm, pH 5.1), that increases the thermal stability of both WT and Δ190–196 recombinant PrPs (Fig. S2). Altogether these observations indicate that Δ190–196 mutant PrP conserves the overall structure of WT PrP but loses some stability.

Expression of the mutant PrP^C in RK13 cells

Ovine Δ190–196 PrP was stably transfected in RK13 cells to generate sublines and clones referred to as Δ190–196 Rov. Mutant PrP was efficiently expressed as a glycoprotein in Rov cells. Western blotting showed that unglycosylated forms of Δ190–196 PrP^C were underrepresented compared with WT or Δ193–197 PrPs generated previously (23) (Fig. 3A). Deglycosylation by PNGase F treatment corroborated the high glycosylation level of Δ190–196 PrP and indicated that the mutant protein was smaller than WT PrP^C by ∼1 kDa, as expected (Fig. 3B). Using an antibody with an epitope in the C-terminal part of PrP rather than in the N-terminal region allowed us to identify both the full-length protein and its natural C-terminal C1 fragment. PNGase treatment was required for accurate identification of PrP^C and C1, because they are both highly glycosylated. The relative proportion of the full-length PrP versus the C1 fragment was roughly similar for the WT and the mutant protein (Fig. 3C). Immunofluorescence showed co-localization of Δ190–196 PrP with WGA, a lectin marker of plasma membrane glycoconjugates, indicating that the mutant protein was correctly addressed to the Δ190–196 Rov cell surface (Fig. 3D) as with the WT protein (27, 28). These observations indicate that Δ190–196 PrP has correct post-translational modifications and cell trafficking.

Figure 3.

Expression of WT and Δ190–196 PrP^C in RK13 cells. A, comparison of the electrophoretic profile of WT PrP, Δ190–196 mutant PrP, and the previously established closely related Δ193–197 mutant (23) by immunoblotting. Molecular mass markers are indicated on the right. On the left, un-, mono-, and biglycosylated species are indicated. B and C, PNGase F treatment was used to resolve the PrP pattern as a single aglycosylated polypeptide. D, confocal microscopy analysis of immunostained cells showed co-localization (merge; in yellow) of Δ190–196 PrP (green) with WGA (red) at the cell surface. Bar, 50 μm. The 4F2 anti-PrP mAb was used in A, B, and D, and Sha31 mAb was used for C.

Spontaneous generation of a self-sustained protease-resistant form of PrP in Δ190–196 rov

The expression of Δ190–196 PrP was turned on by addition of doxycycline, and we followed the fate of the protein over cell passaging by Western blotting, checking for the appearance of PK-resistant forms. Although Δ190–196 PrP was sensitive to PK digestion during the first passages, a PK-resistant form systematically appeared, usually after the fourth or fifth passage (Fig. 4A). This protease-resistant form termed Δ190–196 PrP^res persisted for >1 year of continuous culture (Fig. 5A). The electrophoretic profile of Δ190–196 PrP^res was characterized by a large smear of glycosylated species and the presence of a well-individualized faint band migrating at 14 kDa (Fig. 4A). A second weaker band migrating at 15.5–16 kDa was detected upon overexposure of the blots or when enough material was loaded on the gel (Fig. 5A). Treatment with PNGase F allowed resolving the whole emerging PK-resistant species in two major bands: a main 14-kDa species that had the same size than the C1 fragment and a larger less represented peptide above the 15-kDa molecular mass marker that will be further referred to as 16-kDa PrP^res (Fig. 4B). This indicated that the 14- and 16-kDa bands identified without PNGase treatment (Figs. 4A and 5A) were nonglycosylated native forms of Δ190–196 PrP^res.

Figure 4.

Spontaneous formation of PK-resistant PrP in Δ190–196 Rov cells. Immunoblots of samples from Δ190–196 Rov cell cultures over passaging, passages 2–8 (P2–P8), as indicated at the top of each panel. A, the equivalent of 10 µg of total protein from the cellular lysates were treated or not with PK as indicated at the top of lanes and were loaded on the gel. B, the same samples that are in the upper blot were treated with PNGase F before loading. Full-length mutant PrPs and the C1 fragments are indicated by arrows. The immunoblots were done with Sha31 mAb.

Figure 5.

Persistence of the PK-resistant form in cultures, insolubility of PrP^res, and reproducibility of spontaneous conversion. Immunoblots of PK-treated samples are shown. A, Δ190–196 PrP^Sc was produced persistently up to 1 year of continuous cell culture passage (1 per week). PrP^res at 20, 30, and 52 weeks of culture are shown: 10 µg of total protein in 10 µl of cell lysate were digested and loaded (lanes 2–4). In lane 1 (P20+), the equivalent of 100 µg of protein from the passage at 20 weeks was loaded after PK digestion and concentration of insoluble material at 22,000 × g, to improve detection of the 14- and 16-kDa unglycosylated bands (Sha31 mAb). B, spontaneous emergence of Δ190–196 PrP^res in three populations of Δ190–196 Rov cells obtained from independent transfections (lanes 1–3). C, individual clones isolated from two independent transfections. Five clones (lanes 1–5) with roughly similar Δ190–196 PrP expression levels (upper panel, 4F2 mAb) are shown. They produced spontaneously the PK-resistant form after eight passages of culture (lanes 1, 3, 4, and 5) with the exception of clone 12 (lane 2) (lower panel, Sha31 mAb).

The spontaneous emergence of PK-resistant Δ190–196 PrP was reproducible and occurred systematically from bulk cultures of Δ190–196 Rov obtained from three independent transfections (Fig. 5B). Individual Δ190–196 Rov clones obtained by limiting dilution spontaneously produced Δ190–196 PrP^res, except for one clone, clone 12, despite expression of the mutant PrP to levels similar to those of other clones (Fig. 5C). This “resistant” clone was useful for the infection studies described in the following section, and its existence suggested that currently unrecognized cellular factors are key for the spontaneous generation of Δ190–196 PrP^res. Δ190–196 Rov cells could be frozen and thawed, without affecting the generation of PK-resistant PrP species. This characteristic together with persistence in cell culture recalled that of prion-infected cells.

We next examined whether the biochemical properties of Δ190–196 PrP^Sc resembled those of prions passaged in WT Rov cells. Δ190–196 Rov lysates were treated with increasing PK concentrations and analyzed by Western blotting. Δ190–196 PrP^Sc resisted to higher concentration of PK (Fig. 6, A and B) than 127S prions propagated in WT Rov (Fig. 6C). Δ190–196 PrP^res was recovered by centrifugation at 20,000 × g after PK digestion, indicating that it was insoluble and aggregated (Fig. 5A). The aggregation size of Δ190–196 PrP^Sc was determined by sedimentation velocity. Δ190–196 PrP^Sc formed assemblies with a size in the range of PrP^Sc assemblies formed by subfibrillar prions (Fig. 6D) according to previous reports (29, 30). 127S PrP^Sc assemblies from Rov cells had slightly larger assemblies with respect to size (Fig. 6D). Whether the difference is due to the number of PrP-mers composing the assemblies or to the density of their main core remains to be determined.

Figure 6.

PK resistance and aggregation state of Δ190–196 PrP^res. A, Δ190–196 PrP^res resists to high concentrations of PK. 10 µg of total protein were treated with 2-fold increasing concentrations of PK up to 1 mg/ml. B, samples obtained in A were treated with PNGase F to visualize and determine the molecular mass of polypeptides after PK treatment. C, comparative PK resistance of Δ190–196 PrP^Sc (red squares) and 127S PrP^Sc (black circles). D, sedimentation velocity profile of Δ190–196 PrP^Sc (red line) and 127S PrP^Sc (black line). Fraction number 1 corresponds to the top of the gradient. The results are the means ± S.D. of three independent experiments. In C and D, both proteins were extracted from uninfected Δ190–196 Rov cells and 127S-infected Rov cells, respectively.

To summarize, introduction of the 190–196 deletion in RK13 cells favored the spontaneous and persistent production of PK-resistant PrP^Sc species with an atypical electrophoretic pattern. Δ190–196 PrP spontaneously adopted a conformation that showed hallmarks of a prion: insolubility, aggregation, protease resistance and cell perpetuation. We thus called this entity Δ^Spont prion.

Δ^Spont prion is infectious for cells expressing homologous or closely related mutant PrP

The infectious potential of Δ^Spont prions was primarily tested by cell assay using Δ190–196 or WT Rov. As control, these cells were infected by 127S prions propagated in WT Rov. Naïve Δ190–196 Rov were susceptible to Δ190–196 lysates containing Δ^Spont because they produced PrP^res in large amounts as soon as the second passage postinfection (Fig. 7), whereas mock-infected cells did not. In contrast, WT Rov were not infectable with Δ^Spont prions (Fig. 7). Conversely, 127S could propagate in WT Rov but not in Δ190–196 Rov, at least for the first passages, the spontaneous emergence of Δ^Spont prions on long-term passage obscuring the fate of 127S infection.

Figure 7.

Infectious potential of Δ^Spont prions. Rov cells expressing either WT PrP or Δ190–196 PrP were challenged with cell homogenates from Δ190–196 Rov cells (Δ^Spont inoculum), 127S-infected WT Rov cells (WT-127S), or noninfected RK13 cells (none). The cells were analyzed for the presence of PK-resistant PrP from passages 2–6 postinfection, as indicated. Western blotting was done with Sha31 mAb.

We next challenged the resistant Δ190–196 cell clone 12 and found it readily susceptible to Δ^Spont prion infection. PrP^res was detected as soon as the second passage and up to passage 8 postinfection (Fig. 8A). Δ^Spont prions were thus de novo infectious for cells expressing the homologous mutant protein. The atypical profile of PrP^res, with the characteristic presence of 14- and 16-kDa bands, was faithfully conserved upon infection.

Figure 8.

Conversion of homologous PrP or closely related mutant PrPs by Δ^Spont prions. A, Δ190–196 Rov clone 12 that did not produce spontaneously Δ^Spont PrP^res was infected by Δ^Spont prions (left panel). Rov cells expressing other closely related deletion mutants were also susceptible to the infection by Δ^Spont prions (middle and right panels). Immunoblots show PK-treated samples, eight passages postinfection. B and C, cell homogenates of Δ193–197 (B) or Δ192–197 (C) Rov cells inoculated with Δ^Spont prions (eighth passage, Δ^Spont → Δ193–197 and Δ^Spont → Δ192–197) were used as inocula to infect Rov cells expressing homologous PrP, WT PrP, or the other mutant. The analysis of PK-treated samples harvested after eight passages postinfection is shown.

Previously, we established a set of Rov cells expressing PrP with deletions of different sizes in the C terminus of helix H2. None of these cells produced spontaneously PK-resistant forms of mutant PrP (23). In particular, the Δ193–197 and Δ192–197 cells were susceptible to several ovine prions, including 127S. In contrast, the Δ190–197 cells were resistant to all of them (23). These three mutant cell lines were challenged with Δ^spont prions to determine whether Δ^Spont prion replication was strictly dependent on Δ190–196 PrP. Cells expressing PrP with 190–197, 192–197, or 193–197 deletions were all susceptible to Δ^Spont infection (Fig. 8A). The Δ^Spont PrP^res pattern was mostly maintained in the infected cells.

We knew from our previous work that 127S prions propagated on Δ193–197 Rov cells were still infectious for WT Rov cells, indicating a structural compatibility between this mutant and WT PrP for 127S prion conversion. It was thus appealing to determine whether Δ^Spont propagated on Δ193–197 Rov cells could similarly become infectious for WT cells. The results showed that this was not the case, although ΔSpont prions propagated on Δ193–197 Rov cells were de novo infectious for naïve Δ193–197 cells (Fig. 8B). Similar results were obtained for Δ^Spont propagated on Δ192–197 Rov cells: it was de novo infectious for Δ192–197 Rov cells and infectious for Δ193–197 Rov cells but not for WT Rov cells (Fig. 8C).

Altogether, these cell assays demonstrate the infectivity of Δ^Spont prions and their ability to propagate on cells expressing homologous Δ190–196 PrP or closely related mutants but not WT PrP. This suggests that a certain degree of compatibility in the H2 C-terminal sequence of PrP is required for conversion by Δ^Spont prions.

Molecular typing of Δ^spont prions

Most prion strains can be grouped into two broad categories with respect to their molecular pattern, depending on the N-terminal end point of the C2 PrP^res fragment. Type 1 strains produce PrP^res species starting around position 85 of ovine PrP, whereas type 2 strains PrP^res species are shorter, beginning at position 100. The size of unglycosylated C2 PrP^res is thus a generic way to discriminate between strains. Therefore, we compared the electrophoretic profile of Δ^Spont PrP^res with 127S type 1 prion strain propagated on WT Rov cells and with type 1 (T1^Ov) and type 2 (T2^Ov) strains propagated on Δ193–196 Rov cells, which express a mutant PrP with a size closer to Δ190–196 PrP (Fig. 9). Based on the aglycosylated lower bands, we could ascertain that the main 14-kDa band of Δ^spont prion was shorter than any type 1 or type 2 C2 fragment and that the 16-kDa PK-resistant band was just slightly shorter than the C2 fragment of the type 2 strain. The PrP^res electrophoretic profile of Δ^spont prions was thus clearly atypical with simultaneous presence of two major bands shorter than the usual PK-resistant C2 fragment from “classical” prion strains.

Figure 9.

Molecular typing of Δ^Spont PrP^res. The electrophoretic profiles of prions propagated on Rov cells were compared. The profile of Δ^Spont PrP^res (lanes 2 and 4) with its 14- and 16-kDa bands, as indicated by arrows, was different from those of type 1 strains (127S and T1^ov, lanes 1 and 3, respectively) and type 2 strain (T2^Ov, lane 5) propagated in WT Rov (lane 1) or in Δ193–196 Rov cells (lanes 3 and 5).

To characterize in more detail the nature of the 14- and 16-kDa fragments from Δ^Spont PrP^res, we performed an epitope mapping with anti-PrP antibodies spanning the entire PrP C terminus, after deglycosylation of PrP^res species. Signals produced by Sha31 and 8F9 mAbs were similar, indicating that Δ^Spont PrP^res species are C-terminal fragments of PrP (Fig. 10).

Figure 10.

Epitope mapping of Δ^Spont PrP^res. A, scheme representing mature ovine PrP (positions 25–234, in green) with name and position of mAbs epitopes (open square). The α cleavage site generating C1 fragment is indicated by an arrow. The area covering N-terminal amino acid profile (N-TAAP) variations of C2 PrP^res fragments among prion strains (73) is indicated by the horizontal double-headed dotted arrow. PrP^res fragments resulting from PK digestion of Δ^Spont prions are represented below the scheme, the 14-kDa C1-like PrP^res is in red, the 16-kDa is in orange, and the faint 17-kDa C2-like species is in pink. B, representative Western blots of PNGase F-treated Δ^Spont PrP^res and WT PrP^res revealed with the different anti-PrP mAbs. For the comparative analysis, the same couple of PrP^res samples was loaded several times on a same gel, separated by stained molecular mass markers, and then transferred on a membrane that was split into six parts, each being incubated with a different primary antibody, as indicated.

Regarding the major 14-kDa species, the band was not detected by 12B2 and 8G8 mAbs. Part of the 14-kDa band of higher molecular mass was recognized by 6C2 mAb and thus contained the full epitope of this antibody (¹¹⁴HVAAAGA). The other part of lower molecular mass was 6C2-negative, indicating the loss of at least His¹¹⁴ (Figs. 10 and Fig. S3). Therefore the 14-kDa band contained PrP^res polypeptides with different N-terminal endpoints at the vicinity of the main cleavage site reported for the C1 fragment (8, 31). In noninfected WT Rov cells and in Δ^Spont-free Δ190–196 Rov cells (early passage), part of the C1-PrP^C fragment is 6C2-positive (Fig. S3), suggesting a certain variability in N-terminal endpoints. The PK-resistant 14-kDa band from Δ^Spont PrP^res could thus result from protease digestion of the misfolded full-length Δ190–196 PrP, from a conformational change of the C1 fragment, or from both.

Regarding the 16-kDa band, the epitope mapping indicated a recognition by the 6C2 but not the 8G8 mAb. Therefore the N terminus of these truncated PrP^res polypeptides starts between residues inside the 8G8 epitope (positions 100–105) and residue 113 at the head of 6C2 epitope, most likely close to the 8G8 border based on the molecular mass of these fragments. A faint and fuzzy band was also observed just above the 16-kDa band with all the mAbs used. This indicated the presence of few Δ^Spont PrP^res species of larger size close to the C2 fragments from conventional prions.

Altogether these observations show that Δ190–196 PrP turned spontaneously into a misfolded form producing a complex pattern of PK-resistant species with a main core of ∼14 kDa. These PrP^res fragments of different size reflect the formation of different structures or assemblies by the misfolded protein. The prominence of the 14-kDa band raises the question of whether it could correspond to the minimum and necessary portion of Δ190–196 PrP for Δ^Spont replication as does the C2 fragment for classical prions. As a corollary, this opens the unorthodox question of whether the mutant C1 fragment itself could misfold, either spontaneously or after conversion by Δ190–196 PrP^Sc species produced from the full-length mutant PrP.

Conversion of mutant C1 by Δ^spont prions

To examine the possibility that mutant Δ190–196 C1 PrP^C (ΔC1) alone can misfold into PrP^Sc, either spontaneously or following infection by Δ^Spont prions, we generated Rov cells expressing solely Δ190–196 C1 PrP^C. The C1 fragment is generated by PrP^C cleavage at the α cleavage site, right upstream the N-terminal hydrophobic region. In human brain, the main C1 fragment can start at His¹¹¹ and was referred as to C1-upper, but it can also start at Met¹¹² and was referred to as C1-lower (8). Varying degrees of proteolytically processed C1 fragments at equivalent positions were suggested for ovine PrP (31). We thus built two constructs to express the C1 part of ovine PrP flanked by the N- and C-terminal signal peptides of ovine PrP. The WT and Δ190–196 C1₁₁₃ constructs were designed to start at residues Lys¹¹³, one residue upstream from His¹¹⁴, the equivalent of His¹¹¹ in human PrP, to warrant both a correct and efficient cleavage of the N-terminal peptide signal and the presence of the full 6C2 epitope. The constructs referred as C1₁₁₅ were designed to start at Val¹¹⁵ to produce an equivalent to the C1-lower fragment.

As with WT and Δ190–196 full-length PrP^C, the C1 fragments were highly glycosylated in RK13 cells (Fig. 11A), but the unglycosylated forms were expressed in lower proportion in the two Δ190–196 mutants than in the WT counterparts. The C1 proteins had the expected size (Fig. 11B) and were present at the cell surface (Fig. 11, C and D), indicating that signal peptides were functional and processed by the cells.

Figure 11.

Expression of WT and Δ190–196 C1 polypeptides in RK13 cells. A, immunoblot analysis of stably transfected C1 constructs, before or after PK treatment. The expression of C1-like polypeptides (residues 113–234 and 115–234 of ovine PrP; C1₁₁₃ and C1₁₁₅, respectively) and equivalent fragments with the HTVTTTT Δ190–196 deletion (ΔC1₁₁₃ and ΔC1₁₁₅) was analyzed. B, immunoblot analysis of the same C1 constructs before and after PNGase F treatment. 25 µg of cellular protein was loaded. C, C1 PrP^C expression pattern by immunofluorescence. PrP^C (green) and nuclear marker DAPI (blue) staining of nonpermeabilized fixed cells. Scale bars, 50 μm. D, confocal microscopy imaging of ΔC1₁₁₃ Rov cells co-stained for WGA (red) and PrP (green). The nuclei are stained with DAPI (blue). Merged confocal images or individual channels are shown. Scale bar, 10 μm.

In contrast to full-length Δ190–196 PrP^C, mutant C1 PrP^C did not spontaneously convert into a PK-resistant form over passages, even after several months of cell culture (Fig. 12). However, upon exposure to Δ^Spont prions, these forms were converted into self-replicating PK-resistant PrP^Sc (Fig. 12). These PrP^Sc species were termed ΔC1^Sc. C1 conversion process was specific to Δ^Spont prions and needed the Δ190–196 deletion. 127S prions were not able to convert WT or Δ190–196 C1 PrP^C. Δ^Spont prions were unable to convert WT C1 (Fig. 12), as it was previously unable to do for full-length WT PrP (Fig. 7). As with WT PrP, a certain degree of sequence compatibility between the cellular substrate and the infecting prion is required for C1 conversion by Δ^Spont prion.

Figure 12.

Infection of WT C1 and ΔC1 Rov cells by 127S or Δ^Spont prions. Immunoblots of PK-treated samples at passage 4 (left panel) or passage 8 (right panel) postinfections are shown. Sha31 mAb defines both C1₁₁₃ and C1₁₁₅.

ΔC1^Sc are de novo infectious and preserve Δ^spont PrP^res signature

We finally investigated the infectious potential of ΔC1₁₁₃^Sc and ΔC1₁₁₅^Sc. We used, as inocula, cell lysates of Δ^Spont-infected Δ190–196 C1 at passage 8 (see above). Controls were made to exclude any remnant infectivity of Δ^Spont prions or spontaneous infectivity of the homogenates from cells expressing mutant C1 PrP^C. As shown in Fig. 13, both ΔC1₁₁₃^Sc and ΔC1₁₁₅^Sc were de novo infectious for cells producing the homologous mutant ΔC1 proteins. In contrast cells expressing their WT counterparts were not susceptible to the infection. Remarkably, ΔC1^Sc exhibited the same activity as Δ190–196 PrP^Sc with respect to the panel of susceptible cells. PrP^C from resistant Δ190–196 clone 12 and from the two closely related mutants Δ193–197 and Δ192–197 were converted into PK-resistant PrP^Sc. The Western blotting signature of PrP^res in these cells recalled that produced following Δ^Spont prion infection (compare Figs. 13 and 8A). Not only the characteristic 14-kDa band, but also the 16-kDa band was present, indicating additional conversion of the full-length proteins by ΔC1^Sc. The main strain-specific determinant of Δ^Spont prions was thus enciphered in ΔC1₁₁₃^Sc or ΔC1₁₁₅^Sc.

Figure 13.

Infectivity of ΔC1^Sc prions. Rov cells expressing different forms of mutated, full-length PrP and WT/mutated C1 PrP were left uninfected (none) or exposed to ΔC1^Sc (left panel, ΔC1₁₁₃^Sc; right panel, ΔC1₁₁₅^Sc) obtained from cell homogenates at passage 8 (see Fig. 11). As controls, noninfected (ni) cells and Δ^Spont-infected RK13 cells (control, ctrl) were used. The challenged cells were Δ190–196 Rov cell clone 12 that did not produce spontaneously Δ^Spont prions, Rov cells expressing closely related full-length deletion mutants (Δ193–197, Δ192–196), or populations of cells expressing either WT or a Δ version of C1₁₁₃ (ΔC1₁₁₃) and of C1₁₁₅ (ΔC1₁₁₅). Immunoblots of PK-treated cell lysates at passage 8 postinfection are shown (Sha31 mAb).

Discussion

This work focuses on the spontaneous, in-cell conversion of a deletion mutant PrP into a novel form of prion. Δ^Spont prions showed three main original features: an internal deletion of seven residues in the protease-resistant core, an unusual PrP^res signature, and a remarkable capacity to propagate on the homologous C-terminal C1 segment of PrP, thus generating mutant C1 prions. In addition, this work provides a unique cell model to get insights in cell processes and factors associated with spontaneous prion emergence.

Spontaneous misfolding of ovine PrP with deletion in α-helix H2

Although spontaneous conversion of PrP into prion is a rare event, modifications in PrP can considerably increase this occurrence because not less than ∼40 disease-causing mutations are identified in inherited cases of human PrP (16). Spontaneous prion conversion also occurred in transgenic mice expressing some of these mutations (17–19, 32, 33). Conversion of equivalent mutant PrP in cell culture has so far been disappointing, maybe in part because there is still no easy cell model for efficient human prion replication. We thus considered another approach that was to use a well-characterized reverse genetic model of prion replication and to remove from PrP highly conserved residues that might be important for PrP stability while minimally affecting the overall structure. We previously identified the C-terminal region of the H2 α-helix as an area that meets these prerequisites. We found that the four contiguous threonine residues at the end of the helix were not necessary for replication of several ovine prion strains in RK13 cells (23). Deletion of the threonine cluster (Δ193–196) even favored the replication of strains difficult to propagate in cells expressing WT ovine PrP (23). Thus, we considered this deletion as a good starting point to facilitate the emergence and propagation of a spontaneous prion. Because Δ193–196 deletion was insufficient to cause spontaneous conversion of mutant PrP, we extended it to the three upstream residues His¹⁹⁰, Thr¹⁹¹, and Val¹⁹². Similar residues in human PrP are associated with disease-causing mutations H187R, T188K, and V189I responsible for either GSS or CJD (34–37), highlighting their potential importance for preservation of the normal form of the protein. We found that the simultaneous deletion of residues HTVTTTT maintained the overall PrP structure but strongly reduced the stability of the recombinant ovine PrP. Reduced stability is expected to facilitate partial or complete unfolding of the protein and its spontaneous misfolding (38). Thus the Δ190–196 deletion might have introduced perturbations in the charge equilibrium, salt bridges, and/or hydrophobic interactions. His¹⁸⁷ in human PrP, equivalent to His¹⁹⁰ in ovine PrP, is thought to be a key residue of the electrostatic network stabilizing the globular helical domain of PrP (26, 39). Protonation of histidine is pH-dependent, and the positive charge acquired at acidic pH is thought to be involved in the conformational shift of recombinant PrP in vitro (25, 40, 41). The disease-causing mutation H187R is one of the rare human mutations that markedly reduces PrP stability, and this is attributed to replacement of histidine by a permanently positively charged residue (42). The deletion Δ190–196 not only removed His¹⁹⁰ residue but also brought Lys-197 in position 190, mimicking the replacement of His¹⁹⁰ by another permanently positively charged residue, recalling H187R mutation. This might have contributed to destabilization of Δ190–196 PrP and likely explains why the deletion Δ190–197, which eliminates the lysine residue, did not lead to spontaneous prion generation, whereas the mutant protein remained convertible into PrP^res by Δ^Spont prions. In addition, replacement of each or all the four contiguous threonines (at positions 193–196) by different residues had also affected PrP stability (43, 44). Any of these perturbations alone or in combination may have contributed to the spontaneous conversion of Δ190–196 PrP into Δ^Spont prions. The conservation of HTVTTTT sequence in mammalian PrPs might well reflect preservation of an important region for the dynamics and the maintenance of the helical folding of the protein in a cellular context.

Transmissibility of Δ^spont prions

Δ190–196 PrP was sensitive to protease digestion. However, after several cell passages, a PK-resistant form systematically emerged, indicating spontaneous conformational change of the mutant PrP. The spontaneous formation of PK-resistant PrP was verified in one occasion by an experiment of cell transfection and culture entirely carried out in prion free laboratory, excluding inadvertent contamination. The Δ190–196, or the Δ^Spont PrP^Sc entity as we called it, was shown to be a self-propagating, protease-resistant, insoluble, and aggregated form that was transmissible to other cells. Δ^Spont thus showed all the usual hallmarks of prion replication in cell culture. A basic characteristic of prions is their ability to transmit their own conformational state to homologous PrP^C and in some instance to heterologous PrP^C. Persistence of Δ190–196 PrP^Sc in cell culture indicated some replication, but as we faced a spontaneous conversion that might be reiterated at each passage, we demonstrated de novo infectivity of ΔSpont prion in several cell assays: (i) earlier accumulation of Δ190–196 PrP^Sc in exposed cells; (ii) infection of the Δ190–196 Rov clone 12 that did not produce spontaneously a PK-resistant form of the mutant protein; (iii) infection of Rov cells expressing the Δ190–196 C1 PrP^C; and (iv) infection of Rov cells expressing three other mutant PrPs with close deletions in the H2 C terminus, notably the Δ190–197 mutant that was refractory to infection by conventional ovine prion strains (23). Sequence proximity and/or structural adaptability between Δ^Spont and Δ190–197 PrP might explain it. In contrast, conversion of WT PrP by Δ^Spont prions failed despite many attempts and the use of different cell populations including the historic Rov clone 9 (27) or more susceptible clones (45). In a previous work, we found that the 127S ovine prion strain propagated on WT PrP was easily propagated on Δ193–197 Rov cells and reciprocally (23), suggesting a good structural compatibility for conversion between the Δ193–197 mutant and WT PrP where this strain is concerned. Here, Δ^Spont prion propagated on Δ193–197 PrP was still not able to convert WT PrP, indicating that the Δ193–197 PrP^Sc structure transmitted by Δ^Spont prion rather than only the sequence of the mutant protein causes the transmission barrier.

Biochemical specificities of Δ^spont PrP^Sc

The PrP^res pattern of Δ^Spont was faithfully maintained in the different cell assays. The pattern was complex and showed differentially represented bands corresponding to N-terminal truncations at different end points. It did not fit with any known prion strains regarding the 14- and 16-kDa bands. The minor 16-kDa band was slightly more N-terminally truncated than the classic C2 fragment from type 2 prions. The major 14-kDa band had a size close or similar to that of the C1 fragment of Δ190–196 PrP. These two bands were found in individual clones of Δ190–196 Rov cells with unsimilar relative proportion as in populations of transfected cells. A third, minor, 8G8/12B2-positive band was identified that might be close or similar to that of more classical type1 prions. Thus Δ^Spont PrP^res included fragments close to C2 of type 1 and type 2 plus prominent C1-like species. C2-like protease-resistant fragments are most probably truncated forms of full-length mutant Δ190–196 PrP^Sc, whereas C1-like PrP^res species might result from conversion of the full-length mutant PrP, its C1 fragment, or both. The unconventional PrP^res profile of Δ^Spont prion strongly suggests an original structural organization but might be complicated by conversion of two substrates, the full-length protein and its C1 fragment, rather than only full-length PrP for more classical prions.

Generation of Δ190–196 C1 prion

PrP^C is naturally cleaved into a C-terminal C1 fragment and its complementary N-terminal N1 at the α cleavage site. This cleavage, the efficacy of which is cell-dependent, was attributed to metalloproteases of the ADAM family, but the exact nature of the enzyme remains a subject of controversy and may depend on tissues or cell lines considered (46–50). Moreover, the cleavage is not dependent on a specific sequence, and whether it occurs at the cell surface or inside the cell between the Golgi and the plasma membrane is not clearly established (51, 52). In Rov cells, the C1 fragment appears as a relatively large N-terminally truncated band, beginning around position His¹¹⁴ or Val¹¹⁵ equivalent to that determined in human brain (8, 31). The consensus is that the C1 fragment of PrP^C is shorter than the PrP^res domain and thus cannot be converted into prion. Furthermore, an apparent inverse correlation between C1 levels and cell susceptibility to prions was reported in cell lines, as well as a dominant-negative effect on prion replication in transgenic mice overexpressing C1 (14, 53). Thus, the C1 fragment is often considered as a competitive inhibitor of PrP^C conversion. We found here different outcomes. The main PrP^res domain of Δ^Spont prions had a size close or similar to that of C1, and Δ^Spont prions were able to convert the mutant form of C1. Protease-resistant ΔC1^Sc showed hallmarks of prion replication in cell culture and was transmissible de novo to cells expressing the homologous C1 protein, the homologous full-length mutant PrP, or closely related mutants. To the best of our knowledge, there is no other report of confirmed conversion of C1 or C1-like polypeptides. Two presumptive bovine spongiform encephalopathy cases with C1-like PrP^res signature were reported, but they were finally found to lack prion infectivity after inoculation of brain material to cattle and to bovine PrP transgenic mice (54, 55). The exceptional conversion of ΔC1 into prion after infection by the unconventional Δ^Spont prion is likely due to the conjunction of the sequence modification introduced in the polypeptide and adoption of a misfolded structure significantly different from that of the other prions.

ΔC1^Sc as a driving force for the propagation of the Δ^spont prion

ΔC1 prions had templating activity on the full-length mutant proteins and preserved the PrP^res signature of Δ^Spont prion. Indeed, infection by ΔC1 prions produced not only C1-like PrP^res but also C2-like 16-kDa species that were larger than C1 and thus resulted from conversion of the full-length protein. ΔC1^Sc therefore behaves like a prion strain that maintains the structural information associated with the specific biochemical profile of Δ^Spont prion. This strongly suggests that the 14-kDa ΔC1-like PrP^res core produced by spontaneous conversion of Δ190–196 PrP has at least contributed to Δ^Spont prion propagation and might even be the most important template for the normally folded mutant protein.

Because ΔC1 did not spontaneously form a prion, the primary event in Δ^Spont prion formation may be the spontaneous conversion of full-length PrP. Then the reciprocal capacity of Δ190–196 PrP^Sc to induce conversion of ΔC1 fragment and of ΔC1^Sc to induce full-length mutant PrP conversion would lead to Δ^Spont complex pattern.

It was unexpected that the short ΔC1₁₁₅ or ΔC1₁₁₃ fragments, with lengths of 113 and 115 residues, respectively, could be sufficient to adopt a specific transmissible prion structure, because the lack of WT C1 convertibility into prion was explained by its shortened size compared with C2 PrP^res fragments. Whether ΔC1 prion adopts a structure close to C2 but without the 15 or 30 N-terminal residues or a different structural organization remains an unresolved question. Δ^Spont and even more ΔC1 prions recall somehow PrP106, a mouse PrP with a double deletion (Δ23–88, Δ141–176), that produced a miniprion following infection by RML strain, but not spontaneously (56). The sequence of this mutant protein was very different from that of Δ190–196 ovine PrP or C1. The sequence of PrP106 contained ∼20 residues upstream of the mouse C1 segment and a large internal deletion that removed H1, the β2 strand, and five N-terminal residues of H2 (56), together with the loops between H1 and H2 that are considered to be highly important for prion conversion (57–59). Although structurally different from ΔC1 prion, the miniprion is another example of prion entity constituted from elements of the PrP sequence. A third more distant example of short PrP sequence associated with prion formation is the human PrP with Tyr¹⁴⁵-Stop mutation associated with GSS (60). In contrast to the two precedent mutants, PrP Tyr¹⁴⁵-Stop conserves only the N-terminal moiety of PrP and lacks the whole globular helical domain. Δ^Spont and Δ190–196 C1 prions generated in this work are therefore original and attractive prion entities that deserve to be studied further with transgenic mouse models in the future.

Conclusion

We report the generation of a novel spontaneous mutant prion propagating in cell culture and conversion into prion of both the full-length and the C-terminal C1 fragment of Δ190–196 mutant PrP. We demonstrated that only 113 or 115 residues of the PrP C terminus are sufficient to constitute a self-replicating and transmissible prion entity. Our results also suggest also that the HTVTTTT conserved sequence in the H2 C terminus of PrP is important for prion protein stability and that removal of H2 C-terminal residues is required for productive infection in cells challenged by the spontaneous prion. This work also provides a unique cell culture model for spontaneous prion formation to further study the cell factors and molecular processes involved.

Experimental procedures

Plasmid constructs

Sheep Prnp ORF encoding PrP allotype VRQ (Val¹³⁶, Arg¹⁵⁴, and Gln¹⁷¹) was cloned into pTRE plasmid of the pTeT-on expression system (Clontech) (27). Deletions of residues HTVTTTT (positions 190–196) and N-terminal deletions Δ25–112 and Δ25–114 to generate WT and mutant versions of C1₁₁₃ and C1₁₁₅ were performed by site-directed mutagenesis (QuikChange II mutagenesis kit; Stratagene). Each mutant construct was verified by sequencing.

Antibodies

The anti-PrP mAbs used were as follows: 4F2 directed to the octa-repeat domain (residues 62–94 according to sheep PrP numbering) (61, 62), 12B2 (residues 93–97) (63), 8G8 (residues 100–105) (61, 62), 6C2 (residues 114–121) (64) (Central Veterinary Institute, Wageningen University & Research, Wageningen, The Netherlands) Sha31 (residues 148–155) (62) (Bertin Pharma), and 8F9 (residues 224–234) (65). Sha31and 8G8 were biotinylated and further detected with horseradish peroxidase–conjugated streptavidin. For other mAbs, secondary antibodies were peroxidase-conjugated goat anti-mouse IgG (Abliance) used at 1:5000.

Cell culture and isolation of rov cells

Rov cells are epithelial RK13 cells that stably express either WT or mutant ovine PrP in an inducible manner, by using a tetracycline-inducible system (27). They were obtained by transfection of cells by lipofectamine and puromycin selection. We used cell populations produced by a pool of puromycin-resistant cells, unless indicated otherwise. In some occasions, we used individual clones that were obtained by serial dilution of transfected cells in presence of the selecting agent. The cells were grown in Opti-MEM medium (Invitrogen) supplemented with 10% fetal calf serum and antibiotics and split at 1:4 after trypsin dissociation once a week. To express full-length PrP^C or C-terminal C1 polypeptides, the cells were cultivated in the continuous presence of 1 µg/ml of doxycycline (Sigma).

Prions and prion strains

The spontaneous mutant prion Δ^Spont was propagated on Rov cells expressing the Δ190–196 PrP mutant unless stated otherwise. The 127S ovine prion strain was isolated through serial transmission and cloning by limiting dilution of PG127 field scrapie isolate to tg338 transgenic mice expressing the VRQ allele of ovine PrP (66, 67). 127S was propagated on Rov cells expressing the WT PrP for comparative infection test or determination of PrP^res profile. T1^Ov and T2^Ov prions were originally isolated from serial transmission of a human sporadic CJD case (MM2 type) to tg338 mice (45). For comparison of PrP^res profiles with Δ^Spont, T1^Ov and T2^Ov strains were previously propagated on Rov cells expressing Δ193–196 mutant PrP (23).

Prion infection of cell cultures

To test the infectivity of cell cultures, cells were pelleted, frozen, and thawed three times and sonicated three times for 30 s, and the resulting homogenates were used as inocula to infect naive cell cultures as previously described (23, 68). Homogenates were left for 3 days on the challenged cells. They were then washed with PBS, trypsinized, and seeded at 1/10 dilution in fresh culture medium. The cells were then split at 1:4 dilution after 1 week of culture as for each other successive passage. For infection by Δ^Spont prion, homogenates of Δ190–196 Rov cells harvested at least 9 passages after the addition of doxycycline were used. To test for infectivity of Δ^Spont propagated on Rov cells expressing other deletion mutant PrP or C-terminal C1 polypeptides, inocula were made from cells harvested eight passages postexposure.

Cell lysis, protease digestion, and PNGase F treatment

The cells were washed twice with cold PBS, and whole-cell lysates were prepared in TL1 buffer (50 mm Tris-HCl, pH 7.4, 0.5% sodium deoxycholate, 0.5% Triton X-100). The lysates were clarified by centrifugation for 2 min at 800 × g, and protein concentrations were determined by microBCA assay (Pierce). For PrP^res, the lysates were incubated with 8 µg of PK per 1 mg of protein for 2 h at 37 °C and then centrifuged for 30 min at 22,000 × g. The pellets were dissolved in Laemmli sample buffer and boiled for 15 min at 100 °C. When needed, 500 units of PNGase F (New England BioLabs) and 1% Nonidet P-40 were added to denatured proteins that were further incubated at 37 °C overnight.

Sedimentation velocity fractionation

The experiments were performed as previously described (30). Briefly, the cells were solubilized by addition of a buffer containing 4% (w/v) dodecyl-β-d maltoside and benzonase (0.4 unit/µl). After incubation for 30 min at 37 °C, sarkosyl (N-lauryl sarcosine) was added to give a final concentration of 2% (w/v) in the samples. The incubation was pursued for 30 min at 37 °C. 150 μl of solubilized samples were carefully loaded on a 4.8-ml continuous 10–25% iodixanol gradient (Optiprep, Sigma–Aldrich), with final concentrations of 25 mm HEPES, pH 7.4, 150 mm NaCl, 2 mm EDTA, 0.5% sarkosyl. The gradients were centrifuged at 285,000 × g at 4 °C for 45 min in a swinging-bucket SW-55 rotor. 30 fractions of 160 μl were collected and PK-treated at a final concentration of 50 μg/ml for 1 h at 37 °C. PrP^res contents were analyzed by Western blotting using a biotinylated anti-PrP Sha31 mAb. Signal intensities were quantified using ImageLab software (Bio-Rad) and converted into arbitrary units after normalization. A fixed quantity of human recombinant PrP was employed to calibrate the PrP signals in different gels.

Immunoblotting and detection of PrP^C and PrP^res

Either 4–12% NuPAGE Bis-Tris precast polyacrylamide gels (Invitrogen) or 12% Criterion XT Bis-Tris gels (Bio-Rad) were used for SDS-PAGE. For PrP^C analysis, 50 µg of protein per sample was loaded on the gel. For PrP^res, unless otherwise indicated, the samples corresponding to PK-resistant PrP contained in 25 or 50 μg of cell lysate protein were loaded onto the gel. The transfer of proteins, their detection, and their revelation were described previously (28, 69).

Immunofluorescence, image acquisition, and treatment

The cells were grown on plastic dishes or on glass coverslips in regular medium and washed twice with PBS before fixation for 10 min with 4% paraformaldehyde. The cells were then washed and incubated with the required monoclonal primary antibody (4F2 or Sha31, at 1:5000) in a blocking reagent buffer containing 0.5% crystallin (Roche Diagnostic) and 0.1% Tween 20 in PBS. After washing, the cells were incubated with secondary Alexa Fluor 488–conjugated anti-mouse IgG antibodies (Molecular Probes, Invitrogen) used at a 1:500 dilution, as previously reported (28, 70), and the nuclei were stained with 4′,6-diamidino-2-phenylindole (DAPI). To specifically label the cell surface, rhodamine-conjugated wheat germ agglutinin (WGA, Invitrogen) was incubated for 5 min with living cells and washed once with PBS, and cells were further fixed and processed as described above. Images were acquired either with an Axio observer Z1 microscope (Zeiss) equipped with a CoolSnap HQ2 camera (Photometrics) and driven by the Axio-vision imaging system software. For some experiments, confocal microscopy was performed with a Zeiss LSM 700 microscope (MIMA2 Platform, Institut National de la Recherche Agronomique, Jouy-en-Josas, France) using a Plan-Neofluar 40× (NA 1.3) or Plan-Apochromat 63× (NA 1.4) oil-immersion objective. The images were analyzed using ImageJ version 1.49 software (Wayne Rasband, National Institutes of Health, RRID:SCR_003070).

Expression and purification of recombinant PrP

Recombinant proteins were produced and purified from Escherichia coli as published previously (23). Briefly, by site-directed mutagenesis the deletion Δ190–196 was introduced inside the sequence of the full-length ovine PrP (residues 25–234, VRQ allele) cloned into a pET28 expression vector. WT and mutant proteins produced by E. coli were purified by immobilized metal affinity chromatography on Nickel columns.

CD

The secondary structure of recombinant PrP produced by E. coli was analyzed by CD. Measurements were carried out on Jasco-810 spectropolarimeter. Far-UV CD spectra of full-length ovine PrP and the deletion mutant 190–196 were recorded from 260 to 180 nm at 25ºC in 1-μm-path-length quartz cuvette at a protein concentration of 50 μm in 10 mm sodium acetate buffer at pH 5.0. Each CD spectrum was obtained by averaging six scans collected at a scan rate of 200 nm/min. Baseline spectra obtained with buffer were subtracted for all spectra.

NMR

2D ¹H-¹⁵N HSQC and 3D HNCA spectra of 250 μm recombinant ¹⁵N¹³C-labeled Δ190–196 C1₁₁₃ in 10 mm sodium acetate, pH 5 buffer were acquired on a Bruker NMR AVANCE III spectrometer equipped with a cryoprobe at a magnetic field of 18.8 T and a temperature of 298 K. ¹³Cα chemical shifts were analyzed by TALOS-N software by excluding similar sequences (71)

Fluorescence-based thermal shift assay

Reaction mixtures containing 10 or 20 μm of recombinant PrP in 10 mm sodium acetate, pH 5.0, and SYPRO orange (diluted 500-fold from a 5000-fold stock solution; Invitrogen) were made in duplicate in a 96-well fast PCR plate at a final volume of 20 μl. The experiments were also reproduced in a buffer of 250 mm sodium phosphate, pH 5.1 (Fig. S2). The temperature gradient was carried out in the range of 10 °C to 95 °C, at 3 °C/min with a StepOnePlus real-time PCR system (Applied Biosystems) as previously described (72, 74, 75). Fluorescence was recorded as a function of temperature in real time (excitation with a blue LED source and emission filtered through a 5-carboxyl-X-rhodamine emission filter). The melting temperature (T_m) was calculated with the StepOne software v1.3 (Applied Biosystems) as the maximum of the derivative of the resulting SYPRO Orange fluorescence curves.

Data availability

All of the data in this study are contained within the article.