https://orcid.org/0000-0002-2604-2925
Figures
Abstract
In prion diseases, the species barrier limits the transmission of prions from one species to another. However, cross-species prion transmission is remarkably efficient in bank voles, and this phenomenon is mediated by the bank vole prion protein (BVPrP). The molecular determinants of BVPrP’s ability to function as a universal prion acceptor remain incompletely defined. Building on our finding that cultured cells expressing BVPrP can replicate both mouse and hamster prion strains, we systematically identified key residues in BVPrP that permit cross-species prion replication. We found that residues N155 and N170 of BVPrP, which are absent in mouse PrP but present in hamster PrP, are critical for cross-species prion replication. Additionally, BVPrP residues V112, I139, and M205, which are absent in hamster PrP but present in mouse PrP, are also required to enable replication of both mouse and hamster prions. Unexpectedly, we found that residues E227 and S230 near the C-terminus of BVPrP severely restrict prion accumulation following cross-species prion challenge, suggesting that they may have evolved to counteract the inherent propensity of BVPrP to misfold. PrP variants with an enhanced ability to replicate both mouse and hamster prions displayed accelerated spontaneous aggregation kinetics in vitro. These findings suggest that BVPrP’s unusual properties are governed by a key set of amino acids and that the enhanced misfolding propensity of BVPrP may enable cross-species prion replication.
Author summary
The peculiar behavior of bank voles during prion transmission has attracted significant research interest. Whereas the species barrier restricts the transmission of prions between different species, bank voles are thought to be a universal prion acceptor. However, the molecular determinants of this phenomenon have remained elusive. In this study, we demonstrate that five specific residues in the bank vole prion protein are critical for facilitating cross-species prion transmission. Surprisingly, we found that two residues in the bank vole prion protein may have evolved to counteract its intrinsic misfolding propensity, as they severely restrict spontaneous aggregation as well as prion accumulation following cross-species prion challenge. These results provide new insight into the molecular basis of the species barrier for prion disease.
Citation: Arshad H, Patel Z, Al-Azzawi ZAM, Amano G, Li L, Mehra S, et al. (2024) The molecular determinants of a universal prion acceptor. PLoS Pathog 20(9): e1012538. https://doi.org/10.1371/journal.ppat.1012538
Editor: Amanda L. Woerman, Colorado State University College of Veterinary Medicine and Biomedical Sciences, UNITED STATES OF AMERICA
Received: May 9, 2024; Accepted: August 28, 2024; Published: September 10, 2024
Copyright: © 2024 Arshad et al. This is an open access article distributed under the terms of the Creative Commons Attribution License, which permits unrestricted use, distribution, and reproduction in any medium, provided the original author and source are credited.
Data Availability: All data generated or analyzed during this study are included in this manuscript or its supplemental information.
Funding: This work was funded by grants from the Canadian Institutes of Health Research (PJT-169048 to JCW) and the Natural Sciences and Engineering Research Council (NSERC) of Canada (#RGPIN-2015-05112 to JCW). The funders had no role in study design, data collection and analysis, decision to publish, or preparation of the manuscript.
Competing interests: The authors have declared that no competing interests exist.
Introduction
Prions are infectious protein aggregates that give rise to animal and human neurodegenerative disorders collectively referred to as prion diseases [1–3]. Examples of prion diseases include scrapie in sheep, chronic wasting disease (CWD) in deer and elk, and Creutzfeldt-Jakob disease (CJD) in humans. Prions are composed of a glycosylphosphatidylinositol (GPI)-anchored protein, the prion protein (PrP), which is encoded by the Prnp gene and is expressed within cells of the central nervous system [4]. PrP can exist in two conformational states: cellular PrP (PrPC), which possesses an intrinsically disordered N-terminal domain and an α-helical C-terminal domain, and a disease-causing conformer termed PrPSc, which forms aggregates and is composed almost entirely of β-sheets [5–7]. Distinct PrPSc structures give rise to different prion strains that have unique pathogenic properties [8,9]. PrPSc can template the conversion of PrPC into additional copies of PrPSc, thus resulting in the accumulation and spread of prions within the brain [10,11]. While prion diseases can occur sporadically due to spontaneous generation of PrPSc, the ability of PrPSc to self-propagate means they can also occur via an infectious aetiology. However, a species or transmission barrier limits the ability of prions to pass from one species to another. These barriers are dictated by amino acid sequence differences between PrPC and PrPSc as well as the structural compatibility of PrPC for a given prion strain [12,13]. When PrPC and PrPSc are derived from the same species, and therefore have identical amino acid sequences, efficient templating occurs, leading to robust disease transmission. During cross-species prion transmission, differences in amino acid sequence between PrPC expressed by the host and PrPSc can prevent or hamper disease transmission and can lead to prion strain mutation [14–16].
Bank voles (Myodes glareolus) present an exception to the species barrier, as they can be efficiently infected with prion strains from a wide range of animals with at least partial maintenance of prion strain properties [17–24]. Transgenic mice that express bank vole (BV) PrP are similarly susceptible to cross-species prion infection, arguing that the sequence of BVPrP is structurally compatible with PrPSc from many different species [25,26]. Moreover, BVPrP functions as a permissive substrate for cross-species prion replication in several in vitro prion replication paradigms [27–29], expression of wild-type or mutant BVPrP containing isoleucine at polymorphic codon 109 in mice can lead to spontaneous prion formation [30–33], and recombinant BVPrP can be readily polymerized into infectious aggregates [34–36]. Collectively, this suggests that BVPrP is inherently prone to adopting misfolded conformations, potentially allowing it to serve as a universal or near-universal substrate for prion replication with relaxed sequence identity requirements.
The molecular determinants that govern the anomalous behavior of BVPrP in prion transmission experiments remain incompletely defined. As neither mouse (Mo) nor hamster (Ha) PrP enable cross-species prion transmission, the atypical properties of BVPrP must be enciphered within the 7–8 amino acid sequence differences that exist between these three proteins. Previous studies have implicated subsets of BVPrP-specific residues in facilitating cross-species prion replication. For instance, the presence of specific asparagine residues in BVPrP may facilitate prion replication via formation of steric zippers in the final or intermediary structure of PrPSc [37]. Indeed, when PrP sequences between various species of mice and voles were compared, the presence of asparagine at residues 155 and 170 were inferred to be critical for cross-species prion conversion [19]. More recently, the importance of residues E227 and S230 in BVPrP, which are in the C-terminal region near the GPI anchor attachment site, has been highlighted. These two residues have been suggested to function as a “linchpin” domain within BVPrP, in that they gatekeep the conversion of BVPrPC into PrPSc by several different prion strains [38]. Additionally, transgenic mice that overexpress MoPrP containing these two BVPrP C-terminal residues develop a spontaneous illness that displays certain neuropathological signs of prion disease [39]. However, a comprehensive analysis of the key amino acids that enable BVPrP to function as a universal prion acceptor remains lacking.
Genetically modified mice and in vitro prion replication systems have been widely employed to discern the PrP residues that govern cross-species prion replication. However, both paradigms suffer from important drawbacks. For instance, the generation of transgenic or knock-in mice requires significant financial and time commitments, whereas in vitro systems risk being unphysiological as prion replication occurs in a cell-free environment. Cultured cell models, on the other hand, represent a more tractable system for studying prion replication as they provide the benefit of using intact cells that express membrane anchored PrPC with its correct post-translational modifications [40]. CRISPR/Cas9 gene editing has expanded the range of prion strains that can be studied in cultured cells [41]. For example, CAD5 cells, which can be infected with several different strains of mouse prions [42,43], can be ablated for endogenous MoPrP (CAD5-PrP-/- cells) and then used as a blank canvass to express PrPs from different species [44,45]. Recently, we demonstrated that CAD5-PrP-/- cells expressing BVPrP enable cross-species prion replication following challenge with several different strains of mouse or hamster prions [46]. In this study, we utilized CAD5-PrP-/- cells expressing mutant or chimeric PrPs to systematically identify the amino acid determinants that permit BVPrP to enable cross-species prion replication.
Results
The role of the BVPrP N-terminal domain and N-glycosylation in cross-species prion infection
We have previously demonstrated that monoclonal CAD5-PrP-/- cell lines stably expressing BVPrP are susceptible to infection with several different strains of mouse and hamster prions [46]. Indeed, following three passages post-infection, a monoclonal line of CAD5-PrP-/-(BVPrP) cells exhibits robust levels of proteinase K (PK)-resistant PrP (PrPres), indicative of PrPSc production (S1 Fig). Monoclonal lines are ideal for cellular prion infection experiments as they obviate the need for including a selection antibiotic in the culture medium. While selection antibiotics such as G418 help to maintain PrPC expression levels, they can also interfere with de novo prion infection [47]. Polyclonal cell lines stably expressing BVPrP and cultured in the presence of a low concentration of G418 (0.2 mg/mL) can also become infected following challenge with mouse and hamster prion strains, although they display comparatively lower levels of PrPres at passage three (S1 Fig). To facilitate studying the susceptibility of many PrP variants to cross-species prion infection, we chose to use polyclonal lines of stably transfected CAD5-PrP-/- cells for our experiments and analyzed them following six passages post-infection to allow sufficient time for PrPres accumulation. PrPC expression levels in the stably transfected cell lines were analyzed using the antibodies HuM-D13 (epitope: residues 96–106 of BVPrP), which detects only full-length PrP species, or POM1 (epitope: residues 139–148 of BVPrP), which detects full-length PrP as well as the C1 and C2 endoproteolytic fragments. For detection of PrPres in cell lysates, the antibodies HuM-D13 and HuM-P (epitope: residues 97–106 of BVPrP) were used interchangeably, as both are capable of detecting a wide range of bank vole prion strains following infection of cultured cells or transgenic mice [25,46]. BVPrP is polymorphic at codon 109, where either a methionine (M109) or isoleucine (I109) residue can be present [48]. Although both the M109 and I109 variants of BVPrP are susceptible to cross-species prion infection when expressed in cultured cells [46], we chose to exclusively use the M109 variant for this study since it is less prone to forming prions spontaneously [30].
While not essential for prion replication to occur, post-translational modification of PrP by N-glycans and the disordered PrP N-terminal domain can modulate the conversion of PrPC into PrPSc and the resultant prion strain properties [49–54]. To test the roles of the N-terminal region and N-glycosylation in the ability of BVPrP to enable cross-species prion replication, we generated BVPrP mutants lacking portions of the N-terminal domain as well as mutants with asparagine to glutamine substitutions that abolish the N-glycan attachment sites (S2A Fig). All mutants could be successfully expressed in CAD5-PrP-/- cells (S2B Fig). The mutant BVPrP-expressing cell lines were tested for their ability to replicate two strains of mouse prions (RML and 22L) and two strains of hamster prions (263K and HY), in comparison to CAD5-PrP-/- cells expressing wild-type BVPrP. Mutants lacking the complete N-terminal domain (Δ23–89) as well a mutant lacking the second N-glycan attachment site (N197Q) formed PrPres following challenge with either mouse or hamster prions (S2C–S2F Fig). In contrast, cells expressing mutant BVPrP lacking the first N-glycan site (N181Q), either alone or in combination with the N197Q mutation, did not accumulate PrPres following prion infection. Following prion infection, the patterns of PrPres glycosylation differed in cells expressing wild-type BVPrP compared to cells expressing the Δ23–89 or N197Q mutants, potentially due to strain evolution. These results indicate that the N-terminal domain of BVPrP as well as N-glycosylation at residue N197 are dispensable for cross-species prion replication.
Identification of key residues in BVPrP that enable cross-species prion replication using chimeric mouse/bank vole PrPs
The sequence of mature BVPrP lacking the N- and C-terminal signal sequences differs from the sequence of MoPrP at eight positions (Fig 1A). With respect to the sequence of MoPrP, these changes are insertion of glycine at position 54 as well as the S71G, S79G, L108M, Y154N, S169N, D226E, and R229S substitutions. Because one of the changes is the insertion of a glycine between residues 53 and 54 of MoPrP, the amino acid numbering of BVPrP residues C-terminal to residue 53 is offset by one compared to MoPrP (i.e., residue 108 of MoPrP corresponds to residue 109 of BVPrP). The BVPrP-specific amino acid residues are present throughout the sequence of BVPrP, with three residues found in the disordered N-terminal domain (G54, G72, and G80), three residues in the central globular domain (M109, N155, and N170), and two residues near the C-terminal GPI anchor attachment site (E227 and S230). To investigate which of these residues are critical for enabling cross-species prion replication, three Mo/BVPrP chimeras were initially generated. Mouse chimera 1 (MoC1) consists of MoPrP with the three BVPrP-specific residues from the N-terminal domain, MoC2 contains the three BVPrP residues from the central globular domain, and MoC3 contains the two BVPrP residues near the GPI anchor attachment site (Fig 1A). As CAD5-PrP-/- cells expressing wild-type MoPrP are resistant to hamster prions [44,46], our goal was to identify combinations of BVPrP residues that when added to MoPrP enable replication of hamster prions without losing the ability to become infected by mouse prions. The three chimeras were stably expressed in CAD5-PrP-/- cells and exhibited similar levels of PrP expression to wild-type BVPrP (Fig 1B). Cells expressing the chimeras were challenged with three different hamster prion strains: 263K, HY, and 139H. We note that although 263K and HY prions have different origins (scrapie and transmissible mink encephalopathy, respectively), they exhibit similar strain phenotypes and therefore may not be distinct strains [55]. Cells expressing wild-type MoPrP or BVPrP were used as negative and positive controls, respectively. Of the three chimeras, only cells expressing MoC2 were able to be infected by the three hamster prion strains, as indicated by the presence of PrPres in cell lysates (Fig 1C–1E). Thus, residues M109, N155, and N170 of BVPrP enable cross-species prion replication when inserted into the sequence of MoPrP.
(A) Amino acid sequence alignment of the sequences of BVPrP and MoPrP following removal of N- and C-terminal signal sequences. The sequences differ at eight positions (depicted by red lettering in the BVPrP sequence). The location of residue substitutions in the MoC1, MoC2, and MoC3 chimeras is indicated. (B) Immunoblot for PrPC in undigested lysates from CAD5-PrP-/- cells stably expressing the indicated wild-type or chimeric PrP molecules. The blot was reprobed with an antibody against actin. (C-E) Immunoblots of PrPres levels in lysates from CAD5-PrP-/- cells stably expressing the indicated PrP molecules challenged with either 263K (C), HY (D), or 139H (E) hamster prions. (F) Immunoblot for PrPC in undigested lysates from CAD5-PrP-/- cells stably expressing the indicated wild-type or mutant PrP molecules. The blot was reprobed with an antibody against actin. (G-I) Immunoblots of PrPres levels in lysates from CAD5-PrP-/- cells stably expressing the indicated PrP molecules challenged with either 263K (G), HY (H), or 139H (I) hamster prions. (J) Immunoblot of PrPres levels in lysates from CAD5-PrP-/- cells stably expressing wild-type MoPrP, the MoC1 chimera, or the MoC2 chimera challenged with RML mouse prions. (K-L) Immunoblots of PrPres levels in lysates from CAD5-PrP-/- cells stably expressing wild-type MoPrP, the MoC2 chimera, or Y154N/S169N-mutant MoPrP challenged with RML (K) or 22L (L) mouse prions. PrPC and PrPres were detected using the antibodies HuM-D13 or HuM-P. In all panels, the molecular weight markers indicate kDa.
To determine which residues within MoC2 were sufficient for permitting cross-species prion replication, each possible combination of the three residues was tested. The resultant six new cell lines exhibited roughly similar levels of PrP expression (Fig 1F). We found that the addition of BVPrP residues N155 and N170 (Y154N and S169N mutations in MoPrP) produced levels of PrPres similar to those observed in cells expressing MoC2 for all three hamster prion strains tested (Fig 1G–1I). Of the two residues, N155 appears to be more important as it permitted infection with 263K and 139H prions in isolation, albeit with lower levels of PrPres produced, whereas cells expressing a mutant MoPrP containing just the N170 substitution were resistant to all three hamster prion strains. Importantly, cells expressing either the MoC2 or N155/N170 mutants retained their ability to be infected with two different strains of mouse prions: RML and 22L (Fig 1J–1L). Therefore, the addition of BVPrP residues N155 and N170 enable MoPrP to function as a permissive substrate for cross-species prion replication, akin to wild-type BVPrP.
To further investigate the importance of these two residues, a mutant was generated in which the corresponding mouse residues were substituted into the sequence of BVPrP (N155Y and N170S substitutions). In CAD5-PrP-/- cells, N155Y/N170S-mutant BVPrP expressed at similar levels to wild-type BVPrP (Fig 2A). When N155Y/N170S-mutant cells were infected with mouse (RML, 22L) or hamster (263K, HY) prions, little to no PrPres was observed, suggesting that the cells were resistant to prion infection (Fig 2B and 2C). In contrast, abundant PrPres was observed in cells expressing wild-type BVPrP following challenge with each of the four prion strains. To further test the resistance of this BVPrP mutant to prion infection, RML and 263K prions adapted to the BVPrP sequence by passaging through cells expressing wild-type BVPrP (BV.RML and BV.263K prions) were used as the prion inoculum. Despite the use of BVPrP-adapted prion strains, cells expressing N155Y/N170S-mutant BVPrP were still resistant to prion infection (Fig 2D). Thus, asparagine residues 155 and 170 of BVPrP are necessary for cross-species prion replication. Consistent with our findings, the importance of these asparagine residues for the unusual properties of BVPrP has previously been inferred from prion infection studies in phylogenetically related rodent species as well as from in vitro prion conversion experiments [19,37].
(A) Immunoblot for PrPC in undigested lysates from CAD5-PrP-/- cells stably expressing the indicated BVPrP molecules. Lysate from untransfected CAD5-PrP-/- cells is included as a negative control. The blot was reprobed with an antibody against actin. (B-D) Immunoblots of PrPres levels in lysates from CAD5-PrP-/- cells stably expressing either wild-type or N155Y/N170Q-mutant BVPrP challenged with mouse (B), hamster (C), or bank vole-adapted (D) prion strains. PrPC and PrPres were detected using the antibody HuM-D13. In all panels, the molecular weight markers indicate kDa.
Identification of additional residues in BVPrP that enable cross-species prion replication using chimeric hamster/bank vole PrPs
Our results demonstrate that asparagine residues at positions 155 and 170 of BVPrP are critical for cross-species prion replication. However, the sequence of HaPrP also contains these two residues, yet HaPrP does not enable cross-species prion replication like BVPrP [46]. Thus, BVPrP must contain additional residues that are absent in HaPrP that allow it to act as a permissive substrate for cross-species prion replication. The mature sequence of HaPrP differs from BVPrP at only seven residues (Fig 3A). With respect to the sequence of HaPrP, these changes are M112V, M139I, I203V, I205M, T215V, D227E, and R230S. To investigate which BVPrP residues are critical, we generated two Ha/BVPrP chimeras. Hamster chimera 1 (HaC1) consists of HaPrP with five BVPrP-specific residues from the central globular domain (M112V, M139I, I203V, I205M, and T215V) whereas HaC2 consists of HaPrP with the BVPrP-specific substitutions D227E and R230S near the GPI anchor attachment site (Fig 3A). The two chimeras, along with HaPrP as a negative control and BVPrP as a positive control, were stably expressed in CAD5-PrP-/- cells (Fig 3B). To determine which combination of BVPrP residues enable HaPrP to replicate mouse prions, cells were treated with the RML and 22L strains. Cells expressing either HaC1 or wild-type BVPrP could be successfully infected with both prion strains, with HaC1-expressing cells displaying higher levels of PrPres than cells expressing wild-type BVPrP (Fig 3C and 3D).
(A) Amino acid sequence alignment of the sequences of BVPrP and HaPrP following removal of N- and C-terminal signal sequences. The sequences differ at seven positions (depicted by red lettering in the BVPrP sequence). The location of residue substitutions in the HaC1 and HaC2 chimeras is indicated. (B) Immunoblot for PrPC in undigested lysates from CAD5-PrP-/- cells stably expressing the indicated wild-type or chimeric PrP molecules. The blot was reprobed with an antibody against actin. (C-D) Immunoblots of PrPres levels in lysates from CAD5-PrP-/- cells stably expressing the indicated PrP molecules challenged with either RML (C) or 22L (D) mouse prions. (E) Immunoblot for PrPC in undigested lysates from CAD5-PrP-/- cells stably expressing the indicated wild-type or mutant PrP molecules. The blot was reprobed with an antibody against actin. (F-H) Immunoblots of PrPres levels in lysates from CAD5-PrP-/- cells stably expressing the indicated PrP molecules challenged with either RML (F), 22L (G), or 263K (H) prions. PrPC and PrPres were detected using the antibody HuM-D13. In all panels, the molecular weight markers indicate kDa.
To determine which of the BVPrP residues in HaC1 were critical for enabling cross-species prion replication, each individual BVPrP residue substitution was generated independently in HaPrP and stably expressed in CAD5-PrP-/- cells (S3A Fig). The cell lines were then challenged with RML and 22L prions. Only cells expressing wild-type BVPrP or the HaC1 chimera showed any signs of infection, indicating that none of five BVPrP residues are individually sufficient to enable HaPrP to function as a permissive substrate for cross-species prion replication (S3B and S3C Fig). Therefore, an alternate approach was taken. We hypothesized that individual residue reversions within the HaC1 chimera to the original HaPrP residue would reveal the importance of each of the five distinct residues. Lines of stably transfected CAD5-PrP-/- cells were generated, with each PrP variant expressing at roughly similar levels (Fig 3E). The lines were then challenged with RML and 22L prions. The V203I substitution did not affect the ability of HaC1 to become infected with mouse prions, indicating that BVPrP residue 203 is dispensable for cross-species prion infection (Fig 3F and 3G). Importantly, cells expressing either HaC1 or HaC1 with the V203I substitution retained their susceptibility to hamster 263K prions, indicating that the identified BVPrP residues enable cross-species prion replication and that observed effects are not simply due to sequence similarity between PrPC and PrPSc (Fig 3H). Lower amounts of PrPres were also observed following infection of cells expressing HaC1 with the V215T or I139M substitutions with mouse prions (Fig 3F and 3G). Cells expressing HaC1 with the V215T substitution, but not the I139M substitution, were also susceptible to hamster prions, indicating that, like residue 203, residue 215 of BVPrP is not critical for cross-species prion replication (Fig 3H).
A Ha/BVPrP chimera consisting of HaPrP with BVPrP-specific residues at positions 112, 139, 205, and 215, termed HaC3, was used as a starting point to more precisely map the BVPrP residues that enable HaPrP to serve as a permissive substrate for cross-species prion replication (Fig 4A). All possible two- and three-residue combinations of these four positions were generated and stably expressed in CAD5-PrP-/- cells (Fig 4B and 4C). When challenged with RML prions, only the HaPrP chimera containing the M112V, M139I, and I205M substitutions became infected with prions (Fig 4D). The same chimera was also susceptible to both mouse 22L and hamster 263K prions (Fig 4E and 4F). While no two-residue combination was sufficient to enable replication of RML prions, the addition of BVPrP residues V112 and I139 to HaPrP did permit robust replication of the 22L strain (Fig 4E). Thus, we conclude that three BVPrP residues (V112, I139, and M205) are sufficient for permitting HaPrP to replicate both mouse and hamster prions, although certain strains may require only a subset of these residues.
(A) Amino acid sequence alignment of the sequences of BVPrP and HaPrP with the location of residue substitutions in the HaC3 chimera indicated. (B) Chimeras A-J contain various combinations of BVPrP residues 112, 139, 205, and 215 inserted into the sequence of HaPrP. (C) Immunoblot for PrPC in undigested lysates from CAD5-PrP-/- cells stably expressing the indicated wild-type or chimeric PrP molecules. The blot was reprobed with an antibody against actin. (D-F) Immunoblots of PrPres levels in lysates from CAD5-PrP-/- cells stably expressing the indicated PrP molecules challenged with either RML (D), 22L (E), or 263K (F) prions. For each strain, all samples were transferred to the same membrane. PrPC and PrPres were detected using the antibody HuM-D13. In all panels, the molecular weight markers indicate kDa.
Identification of a C-terminal motif in BVPrP that hampers cross-species prion replication
BVPrP only contains two unique residues that are not found in the sequences of either MoPrP or HaPrP: residues E227 and S230 located near the GPI anchor attachment site. These two residues have been implicated in BVPrP’s propensity to misfold spontaneously and its ability to function as a permissive substrate for prion replication [38,39]. Unexpectedly, we found that chimeric PrPs that lack BVPrP residues E227 and S230 function as better substrates for cross-species prion replication than wild-type BVPrP, based on the higher levels of PrPres observed following infection with hamster or mouse prions (Figs 1C–1E, 1G-1I, 3C, 3D, 3F, 3G and 4D–4F). To further investigate the role of these two residues in cross-species prion replication, we re-tested the chimeric mouse and hamster PrPs containing BVPrP residues E227 and S230, previously referred to as MoC3 and HaC2. New lines of stably transfected CAD5-PrP-/- cells were generated, along with cells expressing wild-type MoPrP or HaPrP as controls (Fig 5A and 5B). Cells expressing MoPrP containing the D226E and R229S substitutions were resistant to RML and 22L prions, indicated by a lack of PrPres following prion infection, whereas cells expressing wild-type MoPrP were readily infected (Fig 5C). Similarly, cells expressing HaPrP containing the D227E and R230S substitutions were resistant to 263K and HY hamster prions (Fig 5D). Both wild-type and mutant versions of MoPrP and HaPrP were expressed at the cell surface, indicating that the C-terminal substitutions do not affect PrPC trafficking (S4A Fig). CAD5-PrP-/- cells expressing HaPrP containing the D227E and R230S substitutions accumulated less PrPres following infection with BV.263K prions (Fig 5E), suggesting that decreased sequence identity between the C-terminal chimeric PrPC substrates and PrPSc in the inoculum is not sufficient to explain the observed effects, as BV.263K prions are more closely sequence matched to the D227E/R230S mutant than wild-type HaPrP. We could not test whether the same is true when using BV.RML prions, since cells expressing wild-type MoPrP are resistant to infection by BVPrP-adapted mouse prion strains [46].
(A) Immunoblot for PrPC in undigested lysates from CAD5-PrP-/- cells stably expressing wild-type or D226E/R229S-mutant MoPrP. (B) Immunoblot for PrPC in undigested lysates from CAD5-PrP-/- cells stably expressing wild-type or D227E/R230S-mutant HaPrP. (C) Immunoblot of PrPres levels in lysates from CAD5-PrP-/- cells stably expressing either wild-type or D226E/R229S-mutant MoPrP challenged with 22L or RML mouse prions. (D-E) Immunoblots of PrPres levels in lysates from CAD5-PrP-/- cells stably expressing either wild-type or D227E/R230S-mutant HaPrP challenged with 263K, HY (D), or BV.263K (E) prions. (F) Immunoblot for PrPC in undigested lysates from CAD5-PrP-/- cells stably expressing wild-type or E227D/S230R-mutant BVPrP. (G-I) Immunoblots of PrPres levels in lysates from CAD5-PrP-/- cells stably expressing either wild-type or E227D/S230R-mutant BVPrP challenged with mouse (G), hamster (H), or bank vole-adapted (I) prion strains. PrPC was detected using the antibodies POM1 or HuM-D13 whereas PrPres was detected using HuM-P or HuM-D13. The blots in panels A, B, and F were reprobed with an antibody against actin. In all panels, the molecular weight markers indicate kDa.
We also conducted the opposite experiment in which the two C-terminal residues in BVPrP were substituted with the equivalent residues that are common to the sequences of both MoPrP and HaPrP. A BVPrP variant with the E227D and S230R substitutions was generated and stably expressed in CAD5-PrP-/- cells (Fig 5F). When these cells were challenged with mouse and hamster prion strains, they exhibited much higher levels of PrPres than cells expressing wild-type BVPrP (Fig 5G and 5H). Once again, sequence similarity between PrPC and PrPSc was not the driving force for this observation since higher levels of PrPres were observed in cells expressing E227D/S230R-mutant BVPrP following infection with BVPrP-adapted RML and 22L prions compared to wild-type BVPrP-expressing cells (Fig 5I). Therefore, the C-terminal region of BVPrP contains amino residues that interfere with prion replication.
To further assess the inhibitory properties associated with BVPrP residues E227 and S230 and how they relate to sequence similarity, we compared four BVPrP substrates harboring various amino acids at positions 227 and 230. These include BVPrP with the wild-type residues (E,S), mouse/hamster equivalent residues (D,R), human PrP residues (Q,S) or residues found in the sequence of elk/ovine/bovine/rabbit/dog PrP (Q,A) (Fig 6A). The BVPrP variants were stably expressed in CAD5-PrP-/- cells, with roughly equal levels of PrPC expression between the four lines (Fig 6B). Moreover, cell surface PrPC expression was observed in all four lines (S4B Fig). When these cells were challenged with mouse (RML, 22L) or hamster (263K, HY, DY) prions, cells expressing wild-type BVPrP accumulated the lowest amounts of PrPres (Figs 6C, 6D and S5). Simply substituting BVPrP residue E227 with the Q227 residue found in human PrP drastically improved the ability of BVPrP to permit cross-species prion replication. Notably, Q227 is not present in any of the mouse or hamster PrPSc inocula, confirming that the differences in PrPres accumulation are not mediated by sequence similarity between PrPC and PrPSc. An independent repeat of these experiments produced similar results (S6 Fig). Moreover, identical results were obtained with four different strains of BVPrP-adapted prions (Fig 6E). In summary, the presence of residues E227 and S230 in wild-type BVPrP substantially reduces susceptibility to cross-species prion infection and the effect is independent of sequence similarity.
(A) Amino acid sequence alignment of the sequences of the C-terminal regions of bank vole, mouse, hamster, human, and elk PrP. A description of the C-terminal BVPrP chimeras is also given. (B) Immunoblot for PrPC in undigested lysates from CAD5-PrP-/- cells stably expressing the indicated wild-type or chimeric BVPrP molecules. The blot was reprobed with an antibody against actin. (C-E) Immunoblots of PrPres levels in lysates from CAD5-PrP-/- cells stably expressing the indicated PrP molecules challenged with either mouse (C), hamster (D), or bank vole-adapted (E) prion strains. Longer-exposure immunoblots are shown in S5 Fig. PrPC and PrPres were detected using the antibody HuM-D13. In all panels, the molecular weight markers indicate kDa.
PrP aggregation kinetics mimic the relative permissiveness of different PrPs to cross-species prion replication
To investigate the mechanism by which BVPrP may permit cross-species prion replication, we generated recombinant (r) full-length untagged PrPs from several different species including bank vole, mouse, hamster, sheep, and elk (Fig 7A). Circular dichroism spectroscopy confirmed that each rPrP was successfully refolded to an α-helical state, consistent with the structure of PrPC (Fig 7B). We then compared the relative propensities of the rPrPs to spontaneously polymerize into aggregates by performing a Thioflavin T (ThT) aggregation assay at 37°C with continuous shaking. The ThT assays were performed in a physiological buffer containing 135 mM salt and a pH of 7.3. rBVPrP formed spontaneous aggregates much more rapidly than the rPrPs from the other four species (Fig 7C). Indeed, the lag phase for rBVPrP aggregation was significantly shorter than the lag phases for aggregation of hamster, ovine, and elk PrP (Fig 7D). rMoPrP displayed intermediate aggregation kinetics, but consistently polymerized more slowly than rBVPrP (Fig 7C and 7D). We also found that the aggregates formed by rBVPrP exhibited higher ThT fluorescence than those composed of the other rPrPs (Fig 7C and 7E). The shapes of the aggregation curves also differed, with rBVPrP initially reaching a higher maximum ThT fluorescence before plateauing at a lower value, whereas the other rPrPs plateaued at ThT fluorescence values similar to the maximum ThT signal attained (Fig 7C and 7F). The underlying explanation for the differences in curve shape and plateau fluorescence remains to be determined but could be related to structural differences among the aggregates and/or distinct aggregation pathways. These results demonstrate that the ability to permit cross-species prion replication is correlated with enhanced spontaneous aggregation kinetics of rPrP.
(A) SDS-PAGE followed by Coomassie blue staining of purified recombinant BV, Mo, Ha, elk, and sheep (Ov) PrP. (B) Circular dichroism spectra of recombinant PrPs. (C) Aggregation curves for rBVPrP (black), rMoPrP (red), rHaPrP (green), rElkPrP (brown), and rOvPrP (blue) as determined using a ThT fluorescence assay. (D) Quantification of the lag phases for the ThT aggregation assays. (E) Quantification of plateau ThT fluorescence values for the aggregation assays. (F) Quantification of max ThT fluorescence:plateau ThT fluorescence ratios for the aggregation assays. For panels C-F, data are mean ± SEM from 15 independent replicates. For panels D-F, statistical significance was assessed using one-way ANOVA with Dunnett’s multiple comparisons test. P-values for pairwise comparisons relative to rBVPrP are shown.
Next, we decided to test the effects of the key residues identified in the cultured cell prion infection experiments on the aggregation kinetics of rBVPrP. We generated rBVPrP with the MoPrP-equivalent residues at positions 155 and 170 as well as MoPrP with the BVPrP-equivalent residues at positions 154 and 169 (Fig 8A). Both mutants were correctly refolded into an α-helical state (Fig 8B). Remarkably, we found that rMoPrP with asparagine at residues 154 and 169 behaved identically to rBVPrP in the ThT aggregation assay, with a similarly reduced lag phase, enhanced maximum ThT fluorescence, and increased ratio of maximum:plateau ThT fluorescence (Fig 8C–8F). Conversely, rBVPrP lacking the two asparagine residues behaved similarly to rMoPrP. Finally, to examine the influence of C-terminal BVPrP residues on the aggregation kinetics of rPrP, we generated rBVPrP with four different residue combinations at positions 227 and 230 (Fig 9A). All four rPrPs were successfully refolded into an α-helical state (Fig 9B). As expected, rBVPrP aggregated rapidly in the ThT assay (Fig 9C). However, rBVPrP containing the C-terminal residues at positions 227 and 230 from either mouse/hamster, human, or elk/ovine/dog/rabbit/bovine PrP aggregated even more rapidly with significantly shorter lag phases than wild-type rBVPrP (Fig 9C and 9D). Therefore, there is a strong correlation between how rapidly a given rPrP spontaneously polymerizes into aggregates in vitro and the ability of the corresponding PrPC to permit cross-species prion replication when expressed in cultured cells.
(A) SDS-PAGE followed by Coomassie blue staining of the indicated purified recombinant PrPs. (B) Circular dichroism spectra of recombinant PrPs. (C) Aggregation curves for rBVPrP (black, n = 29), rMoPrP (red, n = 36), rBVPrP with the N155Y/N170S substitutions (purple, n = 18), and rMoPrP with the Y154N/S169N substitutions (grey, n = 30) as determined using a ThT fluorescence assay. (D) Quantification of the lag phases for the ThT aggregation assays. (E) Quantification of plateau ThT fluorescence values for the aggregation assays. (F) Quantification of max ThT fluorescence:plateau ThT fluorescence ratios for the aggregation assays. For panels C-F, data are mean ± SEM For panels D-F, statistical significance was assessed using one-way ANOVA with Tukey’s multiple comparisons test. P-values for selected pairwise comparisons are shown.
(A) SDS-PAGE followed by Coomassie blue staining of the indicated purified recombinant PrPs. (B) Circular dichroism spectra of recombinant PrPs. (C) Aggregation curves for rBVPrP (black), rBVPrP with the E227D/S230R substitutions (red), rBVPrP with the E227Q substitution (green), and rBVPrP with the E227Q/S230A substitutions (blue) as determined using a ThT fluorescence assay. (D) Quantification of the lag phases for the ThT aggregation assays. For panels C and D, data are mean ± SEM from 12 independent replicates. For panel D, statistical significance was assessed using one-way ANOVA with Dunnett’s multiple comparisons test. P-values for pairwise comparisons relative to rBVPrP are shown.
Discussion
The species barrier in prion disease is predominantly dictated by structural compatibility, which can be conferred by identical or highly similar amino acid sequences between PrPSc seed and PrPC substrate [13]. However, BVPrP demonstrates that a single PrPC substrate can possess the appropriate residues to support the replication of most, if not all prion strains, despite the presence of sequence mismatches between PrPC and PrPSc. In this study, we leveraged the ability of BVPrP to permit cross-species prion replication when expressed in CAD5-PrP-/- cells to identity the key regions in BVPrP responsible for the universal prion acceptor phenotype. Our findings suggest that BVPrP’s anomalous behavior is governed by five amino acid residues that are differentially present in the sequences of MoPrP, HaPrP, and BVPrP. The presence of asparagine residues at positions 155 and 170 of BVPrP explains why BVPrP can permit cross-species prion replication whereas MoPrP cannot. Although HaPrP contains these critical asparagine residues, it lacks essential BVPrP-specific residues at positions 112, 139, and 205, explaining why HaPrP is not a universal prion acceptor. Thus, in some ways, BVPrP can be considered a natural chimera between MoPrP and HaPrP that possesses the right combination of residues to enable cross-species prion replication. However, we unexpectedly found that C-terminal BVPrP-specific residues at positions 227 and 230 hamper cross-species prion transmission, suggesting that the sequence of BVPrP may have been fine-tuned through evolution to counteract its intrinsic propensity to misfold. Furthermore, a striking correlation was observed between the relative susceptibility of a given PrP variant to cross-species replication and the speed at which it forms aggregates spontaneously in vitro. Thus, the ability of PrP to adopt aggregation-competent structures may facilitate crossing of the species barrier.
Previous work has suggested that sequence homology at residues 139 and 155 of PrP is important for efficient templating of PrPC by PrPSc in vitro [56,57]. However, the molecular details of how the five identified BVPrP residues, which include residues 139 and 155, enable cross-species prion replication in cultured cells remain unclear. Conceivably, the residues could act at the PrPC level, potentially by rendering PrPC more prone to aggregation due to localized structural changes that destabilize the protein or promote formation of β-sheet structures. Indeed, rBVPrP aggregated into β-sheet-rich structures much more rapidly than rPrPs from other species, indicative of a higher misfolding propensity and that rPrP, and therefore BVPrPC, may be intrinsically less stable than PrPs from other species. The structure of BVPrPC is very similar to that of PrPC from other mammalian species, except for a more “rigid loop” prior to the second α-helix that is partially mediated by residue N170, which is one of the key BVPrP residues we identified [58]. The presence of a rigid loop has been shown to promote PrP misfolding and spontaneous prion formation as well as modulate interspecies prion transmission, although this appears to be driven by sequence rather than structural homology [59–64]. When coupled with the observation that both elk and horse PrPC also display a rigid loop but do not function as permissive substrates for cross-species prion replication [65,66], this suggests that the rigid loop is not the major determinant of BVPrP’s atypical properties.
Alternatively, the key BVPrP-specific residues could act at the level of PrPSc or hypothetical misfolded intermediate PrP species that occur during the aggregation process. It has been speculated that the relative asparagine content in PrP is a good predictor of susceptibility to prion infection [37], and indeed we discovered that two asparagine residues in BVPrP (N155 and N170) are critical for cross-species prion replication. Due to their polar nature, asparagine side chains can stabilize β-sheets and promote the formation of highly stable zippers or polar clasps [67], potentially lowering energy barriers that would normally dissuade interspecies prion conversion. We found that residue N155 of BVPrP is potentially more important than N170 at facilitating cross-species infection, and this is consistent with structural evidence showing that residue N155 in hamster 263K PrPSc is likely the major determinant that limits transmission of mouse prion strains to hamsters [6]. In the structure of mouse RML PrPSc, residues V111 and I138 (equivalent to V112 and I139 in BVPrP) exist in close spatial proximity within the N-terminal lobe and are found within a hydrophobic pocket that is more tightly packed than in the corresponding region in hamster 263K PrPSc [6,7]. Thus, these residues may help to stabilize the structure of PrPSc, enhancing the efficiency of cross-species prion conversion. In the structure of 263K PrPSc, residue 205 is in proximity to residues near the C-terminal GPI anchor attachment site whereas in the structures of several mouse PrPSc strains it is distal to this region due to a distinct orientation of the PrPSc C-terminus [6–9,68]. Therefore, residue 205 could conceivably influence cross-species prion replication via structural modulation of the PrP C-terminal region. Interestingly, three of the BVPrP residues we identified (V112, I139, and N155) are within or near regions of PrPC thought to play a role in the interaction with PrPSc [69]. It is possible that the five identified BVPrP residues act via diverse and synergistic mechanisms involving PrPC destabilization, PrPSc stabilization, and enhanced PrPC-PrPSc binding to engender BVPrP with the ability to replicate many different prion strains.
A key finding of this study is the discovery of residues in the C-terminal region of BVPrP that hamper cross-species prion replication. This was not simply due to a requirement for sequence identity at residues 227/230 for efficient prion replication, as insertion of a single glutamine residue at position 227 of BVPrP, which is found in the sequence of human PrP but absent in both MoPrP and HaPrP, rendered cells more susceptible to both mouse and hamster prion strains. Thus, despite its ability to function as a universal acceptor of prions, BVPrP can conceivably be rendered even more permissive to prion replication by modifying the amino acids located proximal to the GPI anchor attachment site. Residue E227 is the only truly unique residue in BVPrP as S230 is also found in the sequence of human PrP. It is tempting to speculate that the glutamic acid residue at position 227 arose due to evolutionary pressure to counteract the inherent misfolding propensity of BVPrP, potentially reducing the occurrence of spontaneous or transmitted prion disease in bank voles.
Our findings are opposite to those of previous reports that found that BVPrP residues E227 and S230 are pivotal to the universal prion acceptor phenotype, collectively functioning as a linchpin domain to enable templated or spontaneous structural conversion [38,39]. The reasons for this discrepancy are unclear but could be related to the experimental paradigms employed. We utilized an in vivo cellular environment in which BVPrPC is correctly attached to the outer leaflet of the plasma membrane and would be subject to normal cellular proteostasis mechanisms, whereas in vitro prion amplification assays employing a purified, non-membrane-anchored PrPC substrate were used elsewhere [38]. When the C-terminal region of PrPC is not constrained via attachment to the cell membrane and/or sonication is used to fragment growing PrPSc aggregates, it is possible that the inhibition conferred by BVPrP residues 227/230 can be overridden. Nonetheless, our results further support a model in which the right combination of amino acids at positions 227/230 are important modulators of prion replication efficiency. Consistent with this notion, a mutation in the C-terminal region of PrP can modulate the kinetics of human prion replication [70]. Exactly how residues 227/230 affect prion conversion remains to be determined but could be related to localized structural changes that alter PrPC stability or affect interactions with components of the plasma membrane. One study found that the GPI anchor in human PrP is attached to residue G229, which means that residue S230 would not be present in the mature processed protein [71]. Although the precise attachment site for the GPI anchor in BVPrP has not been experimentally confirmed to the best of our knowledge, this suggests that residue 227 may be a more critical determinant of cross-species prion replication, which is in line with our results.
The striking correlation between the relative permissiveness of a given PrPC to replicate both hamster and mouse prions in cultured cells and the aggregation kinetics of the corresponding rPrP implies that there may be mechanistic links between cross-species prion replication and spontaneous prion generation. Indeed, in transgenic mice, expression of the I109 variant of BVPrP promotes both spontaneous formation of an atypical PrPres species and cross-species prion replication [25,30,31]. We speculate that BVPrP is more conformationally malleable than PrPs from other species, potentially allowing it to form misfolded intermediate species that can either further assemble to initiate spontaneous prion formation or be templated by pre-existing PrPSc in a manner that does not require precise sequence identity between PrPC and PrPSc. It should be noted that we have never observed spontaneous production of PrPres in CAD5-PrP-/-cells stably expressing either the M109 or I109 variants of BVPrP. Whether C-terminal residues in BVPrP attenuate spontaneous prion generation remains to be determined, but we note that changing a single residue in BVPrP to its human equivalent (E227Q) significantly accelerated the spontaneous aggregation of rPrP. Although human PrP contains a misfolding-promoting residue at its C-terminus, it is unlikely to be prone to spontaneous prion conversion since it only possesses two of the five BVPrP residues we have shown to be critical for cross-species prion conversion (I139 and M205). This could explain why spontaneous prion diseases are very rare in humans.
There are several limitations to our study. First, we only used mouse and hamster prion strains to investigate BVPrP-mediated cross-species prion transmission. It is therefore unknown whether the same amino acid determinants in BVPrP govern cross-species transmission involving prion strains from sheep, cervids, cattle, or humans, all of which can be transmitted to bank voles and transgenic mice expressing BVPrP [18–20,25]. While both mouse and hamster prions result in robust PrPres generation following infection of BVPrP-expressing CAD5-PrP-/- cells, detection of infection in cells treated with CWD prions is more challenging and requires amplification using RT-QuIC, which implies low levels of PrPSc accumulation [45,46]. Interestingly, substitution of four elk PrP residues into the sequence of human PrP, one of which is N170, renders mice susceptible to both CWD and CJD prions [64]. Transgenic mice expressing a MoPrP variant containing N170 are also susceptible to CWD prions [60]. Thus, residue N170, which is one of the key BVPrP residues we identified, may also be an important determinant of cross-species CWD prion transmission. Second, we only utilized a single readout of prion infection, relative levels of PrPres detected using antibodies against a single PrP epitope, to assess the ability of a given PrP to permit cross-species prion replication. Thus, we cannot rule out the presence of atypical PrPres species following cross-species prion infection that are only detectable by antibodies that recognize epitopes towards the C-terminus of PrP [72]. Finally, although we have documented a strong parallel between susceptibility of PrP to cross-species prion replication and enhanced spontaneous aggregation kinetics of the corresponding rPrP, it must be noted that rPrP aggregates adopt fibrillar structures distinct from authentic PrPSc and lack the levels of infectivity displayed by brain-derived prions [73,74]. Furthermore, rPrP lacks the N-glycans that are present in cell-expressed PrP, and we found that a BVPrP mutant unable to undergo N-glycosylation was not susceptible to cross-species prion replication. Thus, further studies are required to fully understand any potential links between rPrP aggregation and cross-species prion replication.
Materials and methods
Cell culture
CAD5-PrP-/- cells were generated by CRISPR/Cas9 gene editing of the mouse catecholaminergic cell line CAD5, which is a subclone of the original Cath.a-differentiated (CAD) line [42,44,75]. All experiments utilized CAD5-PrP-/- clone D6 [44]. Cells were cultured in growth medium, which consists of Opti-MEM media (ThermoFisher Scientific #31985088) supplemented with 6.5% (v/v) fetal bovine serum (ThermoFisher Scientific #12483020) and 1X GlutaMAX (ThermoFisher Scientific #35050061). Cells were cultured at 37°C in a 5% CO2 humidified incubator and were maintained by routine passaging at a dilution of 1:5 or 1:3 every 3–4 days. To passage the cells, they were first washed with PBS (ThermoFisher Scientific #14190144) and then treated with enzyme-free cell dissociation reagent (Millipore Sigma #S-014-B). The cells were incubated for 1–3 minutes in a 37°C incubator during the dissociation step and then plated in fresh growth medium. Stably transfected polyclonal CAD5-PrP-/- lines were cultured in growth medium containing 0.2 mg/mL G418 (BioShop #GEN418.5). Monoclonal lines of stably transfected CAD5-PrP-/- cells were cultured in growth medium lacking G418 as previously described [46].
Generation of polyclonal stable cell lines
Expression plasmids encoding wild-type bank vole PrP (M109 variant), mouse PrP, or hamster PrP were generated by cloning the respective Prnp open reading frames between the BamHI and Xbal sites of the vector pcDNA3 [44,46]. The MoC1, MoC2, and HaC1 chimeras were generated by gene synthesis (BioBasic Canada). Other chimeric- and mutant PrP-expressing plasmids were generated by site-directed mutagenesis. For generation of stably transfected CAD5-PrP-/- lines, cells were plated at a density of 7 x 105 cells/well in 6-well plates, and then transfected with a plasmid encoding the desired gene. For transfection, 2 μg of plasmid DNA was mixed with 4 μL of Lipofectamine 2000 (ThermoFisher Scientific #11668019) in 300 μL of Opti-MEM without any additives. This mixture was added to 1.5 mL of Opti-MEM and then applied to CAD5-PrP-/- cells for 24 h. Next, the cells were washed with PBS and detached using enzyme-free dissociation reagent. The entirety of the transfected well was then transferred into a 10-cm dish and cultured in medium containing 1 mg/mL G418. The cells were selected over a period of 12–14 days and then expanded. No clonal selection was performed.
Cell Lysis and immunoblotting
Cells were washed with cold PBS, and then incubated with lysis buffer (50 mM Tris-HCl pH 8.0, 150 mM NaCl, 0.5% (w/v) sodium deoxycholate, and 0.5% (v/v) NP-40) for 30–60 s, swirled, and then lysates collected. A cell scraper was occasionally used when lysing with smaller volumes. Lysates were incubated on ice for 15 min with intermittent vortexing (30 s), and then centrifuged at 5,000x g for 5 min to pellet debris. Protein concentrations in cell lysates were determined using the bicinchoninic acid (BCA) assay (ThermoFisher Scientific #23227). For analysis of PrPC levels, protein was prepared for immunoblotting in 1X Bolt LDS sample buffer (ThermoFisher Scientific #B0007) and then heated for 10 min at 70°C. The samples were run on 10% Bolt Bis-Tris Plus Mini gels (ThermoFisher Scientific #NW00100BOX for 10-well gels) for 35 min at 165 V. Transfer to Immobilon-P PVDF membranes (Millipore Sigma #IPVH00010) was performed for 1 h at 25 V in Tris-glycine transfer buffer (100 mM Tris-HCl pH 8, 137 mM glycine). Membranes were blocked for 1 h at 22°C in blocking buffer (5% (w/v) skim milk in TBS containing 0.05% (v/v) Tween-20 (TBST)), and then incubated overnight at 4°C with antibodies diluted in blocking buffer. The following anti-PrP antibodies were used: recombinant humanized Fab HuM-D13 (1:10,000 dilution), recombinant humanized Fab HuM-P (1:10,000 dilution), and mouse monoclonal POM1 (1:5,000 dilution; Millipore Sigma #MABN2285) [76–78]. The HuM-D13 antibody was provided by Stanley Prusiner (University of California San Francisco). The following day, membranes were washed 3X with TBST (5–10 min per wash), and then incubated with horseradish peroxidase-conjugated secondary antibodies (Bio-Rad #172–1011 or 172–1019, or ThermoFisher Scientific #31414) diluted in blocking buffer for 1 h at 22°C. The membranes were again washed 3X with TBST (10 min per wash). The blots were developed using Western Lightning ECL Pro (Revvity #NEL122001EA) and then chemiluminescent signal was captured by exposure to HyBlot CL x-ray film (Thomas Scientific #1141J52). PrPC blots were reprobed with the rabbit polyclonal anti-actin antibody 20–33 (1:10,000 dilution; Millipore Sigma #A5060).
Prion strains
Hamster prion strains 263K, 139H, Hyper (HY), and Drowsy (DY) were originally obtained as 10% (w/v) brain homogenates in PBS from Jason Bartz (Creighton University). To generate a renewable source of the 263K, HY, and 139H strains, CAD5-PrP-/- cells expressing HaPrP were used [44]. The mouse prion strains RML and 22L were obtained from terminally ill non-transgenic C57BL/6 mice. Brains of prion-infected mice were homogenized in PBS to generate a 10% (w/v) brain homogenate stock, which was aliquoted and stored at -80°C. The homogenization step was conducted using a Minilys homogenizer (Bertin Technologies) with CK14 soft tissue homogenization tubes containing 1.4 mm zirconium oxide beads (Bertin Technologies #P000912-LYSK0-A). Brains were homogenized using 3 cycles of 60 s at maximum speed with each cycle followed by 10 min of incubation on ice. For cellular homogenates, prion-infected CAD5-PrP-/- cells expressing HaPrP were cultured in 10-cm tissue culture plates. After reaching confluency, the cells were washed in cold PBS and then scraped into a small volume of PBS and collected into CK14 soft tissue homogenization tubes supplemented with additional 0.5 mm zirconia beads (BioSpec #11079105Z). The cells were homogenized using 3 cycles of 60 s at maximum speed, with a 10 min incubation on ice between each cycle. Total protein concentrations in cell homogenates were quantified using the BCA assay.
Cellular prion infections
Stably transfected CAD5-PrP-/- cells were plated in either 12- or 24-well dishes at a density of 1.5 x 105 or 1 x 105 cells/well respectively. The following day, cells were challenged with 0.1% (w/v) prion-infected brain homogenate or 100 μg of prion-infected cellular homogenate. After 48–72 h of incubation (depending on cell confluency), the cells were washed 2X with PBS and then passaged at 1:3 dilution into 12-well plates. The cells were then passaged six times in 12-well plates to allow for de novo prion accumulation. At the sixth passage, the cells were scaled up to 6-cm dishes for lysis and analysis of successful prion infection.
Proteinase K digestions
To determine the infection status of cell lines challenged with prions, cells were lysed using lysis buffer, as described above. After determining protein concentration using the BCA assay, 0.5 to 1 mg of cell lysate was digested with PK (ThermoFisher Scientific #EO0491) at a concentration of 50 μg/mL and a PK:protein ratio of 1:50. The digestion was carried out at 37°C for 1 h with shaking at 600 rpm. The digestion was halted by the addition of PMSF to a final concentration of 2 mM, and then sarkosyl (Millipore Sigma #61747) was added to a final concentration of 2% (v/v). Digested samples were ultracentrifuged at 100,000x g for 1 h at 4°C using a TLA-55 rotor (Beckman Coulter) to concentrate the insoluble, PK-resistant fraction. The supernatant was discarded, and the pellet was resuspended in 25 μL of 1X Bolt LDS sample buffer, boiled at 95°C, and then stored at -80°C for subsequent analysis by immunoblotting.
Immunofluorescence microscopy
CAD5 PrP-/- cells stably expressing MoPrP, HaPrP, BVPrP or PrP chimeras were plated in 24-well dishes with a #1.5 glass-like polymer coverslip bottom (Cellvis # P24-1.5P) coated with Poly-D-Lysine (Thermo Fisher #A3890401). Prior to the addition of cells, the coated wells were thoroughly washed and then air dried. Cells were plated at density of 1 x 105 cells/well and then cultured for 48 h until they reached 80–90% confluency. Cells were then fixed with 4% (v/v) paraformaldehyde for 15 min and then washed twice with PBS. Cells were not permeabilized. Following the washes, the cells were blocked in 3% bovine serum albumin (diluted in PBS) for 1 h at room temperature. Cells were then incubated with the anti-PrP antibody POM1 (1:500 dilution) overnight at 4°C in PBS containing 3% bovine serum albumin. The following day, the cells were washed twice with PBS, and then incubated with an AlexaFluor 488-conjugated secondary antibody (Thermo Fisher #A-11001; 1:500 dilution in PBS containing 3% bovine serum albumin) for 2 h at room temperature. Subsequently, cells were washed 2 additional times with PBS. After the final wash, the cells were incubated with DAPI (1:1,000 dilution in PBS) for 10 min, and then washed with PBS. The cells were stored in PBS and then imaged using a Zeiss LSM880 confocal microscope.
Purification of recombinant PrP
Recombinant untagged full-length PrPs (residues 23–230 for mouse PrP, 23–231 for hamster PrP and BVPrP, residues 25–233 for elk PrP and ovine PrP) were expressed and purified as previously described [44,46,47]. Briefly, sequences were inserted into the pET-41 expression vector and then proteins were expressed in E.coli Rosetta2(DE3) cells using an autoinduction system. For BVPrP, the M109 variant was utilized and for sheep PrP, the ARQ allele was used. Cells were lysed using BugBuster Master Mix solution (Millipore Sigma #71456) and then inclusion bodies were isolated. Inclusion bodies were solubilized in 8 M guanidine hydrochloride (GdnHCl) and then PrP was captured using Ni-NTA Superflow beads (Qiagen #30410). Bead-bound PrP was refolded on-column using a 4 h gradient from 6 to 0 M GdnHCl and then eluted using a gradient from 0 to 500 mM imidazole. PrP-containing fractions were dialyzed into 10 mM sodium phosphate pH 5.8, filtered through a 0.22-micron filter, and then stored at -80°C. The concentration of purified recombinant PrP was determined through absorbance at 280 nm using a NanoDrop spectrophotometer. The purity of recombinant PrP was assessed through SDS-PAGE followed by staining with Coomassie brilliant blue. To confirm the successful refolding of recombinant PrP to an α-helical state, circular dichroism experiments on protein dialyzed in 10 mM sodium phosphate pH 7.3 buffer were performed at 22°C using a JASCO J-715 spectropolarimeter.
Thioflavin T aggregation assays
Recombinant PrP was dialyzed in 10 mM sodium phosphate pH 7.3 buffer overnight or for 4 h at 4°C with two buffer changes. To remove any pre-existing aggregates, the dialyzed protein was ultracentrifuged (100,000x g) for 1 h at 4°C. The reaction mixtures were prepared as follows: 0.1 mg/mL recombinant PrP, 10 μM ThT, and 135 mM NaCl in a total volume of 100 μL of 10 mM sodium phosphate pH 7.3. The mixtures were plated in a black, clear-bottom 96-well plate (ThermoFisher Scientific #265301) and then sealed with a clear film (VWR #89134–428). The plate was then placed in a BMG CLARIOstar plate reader and subjected to cycles of 4 min shaking (double orbital, 700 rpm) and 1 min rest/read at 37°C. The ThT readings (excitation 444 ± 5 nm; emission 485 ± 5 nm) were taken every 5 min during the rest period using a gain setting of 1600. For the experiments involving the C-terminal PrP chimeras, the following settings were used to prevent the ThT signal from approaching the detection limit of the instrument: excitation 444 ± 4 nm; emission 485 ± 4 nm; gain = 1500. The lag phases for the aggregation reactions were quantified as previously described [46]. To determine the maximum:plateau fluorescence ratio for each replicate, the maximum ThT fluorescence achieved during the experimental timeframe was divided by the ThT fluorescence at the experimental endpoint (83.25 h). Samples that did not aggregate within the experimental timeframe were excluded from the lag phase, maximum ThT fluorescence, and maximum:plateau ThT fluorescence comparisons.
Statistical analysis
All statistical analysis was performed using GraphPad Prism (version 10.1.1) with a statistical threshold of P < 0.05. Lag phases, maximum ThT fluorescence values, and maximum:plateau ThT fluorescence ratios were compared using one-way ANOVA followed by either Dunnett’s (when comparing all samples to recombinant wild-type BVPrP) or Tukey’s (when performing all possible pairwise comparisons) multiple comparisons tests.
Supporting information
S1 Fig. Infection of monoclonal and polyclonal CAD5-PrP-/- cell lines expressing BVPrP with mouse and hamster prions.
Immunoblot of PrPres levels in monoclonal and polyclonal lines of stably transfected CAD5-PrP-/-(BVPrP) cells infected with either mouse (RML) or hamster (263K, 139H, or HY) prion strains. Cells were analyzed at passage 3 post-infection. PrPres was detected using the antibody HuM-P. Molecular weight markers indicate kDa.
https://doi.org/10.1371/journal.ppat.1012538.s001
(TIF)
S2 Fig. The role of N-glycosylation and the BVPrP N-terminal domain in cross-species prion transmission.
(A) Domain structures of the BVPrP constructs used. For simplicity, the N- and C-terminal signal sequences are not depicted in the diagram. All constructs contain methionine at codon 109. (B) Immunoblot for PrPC in undigested lysates from CAD5-PrP-/- cells stably expressing the indicated BVPrP constructs. The blot was reprobed with an antibody against actin. (C-F) Immunoblots of PrPres levels in CAD5-PrP-/- cells stably expressing the indicated BVPrP constructs challenged with either mouse RML (C), mouse 22L (D), hamster HY (E), or hamster 263K (F) prions. PrPC was detected using the antibody POM1 whereas PrPres was detected using HuM-D13. In all panels, the molecular weight markers indicate kDa.
https://doi.org/10.1371/journal.ppat.1012538.s002
(TIF)
S3 Fig. Single BVPrP residue substitutions in HaPrP are insufficient for enabling cross-species prion replication.
(A) Immunoblot for PrPC in undigested lysates from CAD5-PrP-/- cells stably expressing wild-type HaPrP or BVPrP, the HaC1 chimera, or HaPrP with the indicated BVPrP residue substitutions. The blot was reprobed with an antibody against actin. (B-C) Immunoblots of PrPres levels in lysates from CAD5-PrP-/- cells stably expressing the indicated PrPs challenged with either RML (B) or 22L (C) mouse prions. PrP was detected using the antibody HuM-D13. In all panels, the molecular weight markers indicate kDa.
https://doi.org/10.1371/journal.ppat.1012538.s003
(TIF)
S4 Fig. Chimeric PrPs with C-terminal substitutions are expressed at the cell surface.
(A) Immunofluorescence images of non-permeabilized stably transfected polyclonal CAD5-PrP-/- lines expressing either wild-type MoPrP, wild-type HaPrP, or mutants of MoPrP and HaPrP in which the indicated C-terminal residues were substituted for their BVPrP equivalents. (B) Immunofluorescence images of non-permeabilized stably transfected polyclonal CAD5-PrP-/- lines expressing either wild-type BVPrP or the indicated BVPrP C-terminal chimeras. In both panels, cell surface PrP expression (green) was revealed using the antibody POM1 and nuclei were stained with DAPI (blue). Scale bars = 25 μm.
https://doi.org/10.1371/journal.ppat.1012538.s004
(TIF)
S5 Fig. Longer exposures of immunoblots from cellular prion infection experiments displayed in Fig 6.
Immunoblots of PrPres levels in lysates from CAD5-PrP-/- cells stably expressing the indicated PrP molecules challenged with either mouse, hamster, or bank vole-adapted prion strains. PrPres was detected using the antibody HuM-D13. The molecular weight markers indicate kDa.
https://doi.org/10.1371/journal.ppat.1012538.s005
(TIF)
S6 Fig. Independent repeat of cellular prion infection experiments displayed in Fig 6.
Immunoblots of PrPres levels in lysates from CAD5-PrP-/- cells stably expressing the indicated PrP molecules challenged with either mouse or hamster prion strains. PrPres was detected using the antibody HuM-D13. The molecular weight markers indicate kDa.
https://doi.org/10.1371/journal.ppat.1012538.s006
(TIF)
S1 File. Spreadsheet containing all numerical data from the experiments.
https://doi.org/10.1371/journal.ppat.1012538.s007
(XLSX)
Acknowledgments
We are grateful to Stanley Prusiner for providing the HuM-D13 antibody and Jason Bartz for providing the hamster prion strains. ZP and ZAMA were supported by NSERC undergraduate student research awards.
References
- 1. Colby DW, Prusiner SB. Prions. Cold Spring Harb Perspect Biol. 2011;3(1):a006833. pmid:21421910
- 2. Collinge J. Mammalian prions and their wider relevance in neurodegenerative diseases. Nature. 2016;539(7628):217–26. pmid:27830781
- 3. Prusiner SB. Novel proteinaceous infectious particles cause scrapie. Science. 1982;216(4542):136–44. pmid:6801762
- 4. Watts JC, Bourkas MEC, Arshad H. The function of the cellular prion protein in health and disease. Acta Neuropathol. 2018;135(2):159–78. pmid:29151170
- 5. Riek R, Hornemann S, Wider G, Billeter M, Glockshuber R, Wuthrich K. NMR structure of the mouse prion protein domain PrP(121–231). Nature. 1996;382(6587):180–2. pmid:8700211
- 6. Kraus A, Hoyt F, Schwartz CL, Hansen B, Artikis E, Hughson AG, et al. High-resolution structure and strain comparison of infectious mammalian prions. Mol Cell. 2021;81(21):4540–51 e6. pmid:34433091
- 7. Manka SW, Zhang W, Wenborn A, Betts J, Joiner S, Saibil HR, et al. 2.7 A cryo-EM structure of ex vivo RML prion fibrils. Nature communications. 2022;13(1):4004. pmid:35831275
- 8. Manka SW, Wenborn A, Betts J, Joiner S, Saibil HR, Collinge J, et al. A structural basis for prion strain diversity. Nat Chem Biol. 2023;19(5):607–13. pmid:36646960
- 9. Hoyt F, Alam P, Artikis E, Schwartz CL, Hughson AG, Race B, et al. Cryo-EM of prion strains from the same genotype of host identifies conformational determinants. PLoS Pathog. 2022;18(11):e1010947. pmid:36342968
- 10. Bessen RA, Kocisko DA, Raymond GJ, Nandan S, Lansbury PT, Caughey B. Non-genetic propagation of strain-specific properties of scrapie prion protein. Nature. 1995;375(6533):698–700. pmid:7791905
- 11. Griffith JS. Self-replication and scrapie. Nature. 1967;215(5105):1043–4. pmid:4964084
- 12. Prusiner SB, Scott M, Foster D, Pan KM, Groth D, Mirenda C, et al. Transgenetic studies implicate interactions between homologous PrP isoforms in scrapie prion replication. Cell. 1990;63(4):673–86. pmid:1977523
- 13. Collinge J, Clarke AR. A general model of prion strains and their pathogenicity. Science. 2007;318(5852):930–6. pmid:17991853
- 14. Peretz D, Williamson RA, Legname G, Matsunaga Y, Vergara J, Burton DR, et al. A change in the conformation of prions accompanies the emergence of a new prion strain. Neuron. 2002;34(6):921–32. pmid:12086640
- 15. Castilla J, Gonzalez-Romero D, Saa P, Morales R, De Castro J, Soto C. Crossing the species barrier by PrP(Sc) replication in vitro generates unique infectious prions. Cell. 2008;134(5):757–68. pmid:18775309
- 16. Barria MA, Telling GC, Gambetti P, Mastrianni JA, Soto C. Generation of a new form of human PrP(Sc) in vitro by interspecies transmission from cervid prions. J Biol Chem. 2011;286(9):7490–5. pmid:21209079
- 17. Arshad H, Bourkas MEC, Watts JC. The utility of bank voles for studying prion disease. Prog Mol Biol Transl Sci. 2020;175:179–211. pmid:32958232
- 18. Nonno R, Di Bari MA, Cardone F, Vaccari G, Fazzi P, Dell’Omo G, et al. Efficient transmission and characterization of Creutzfeldt-Jakob disease strains in bank voles. PLoS Pathog. 2006;2(2):e12. pmid:16518470
- 19. Agrimi U, Nonno R, Dell’Omo G, Di Bari MA, Conte M, Chiappini B, et al. Prion protein amino acid determinants of differential susceptibility and molecular feature of prion strains in mice and voles. PLoS Pathog. 2008;4(7):e1000113. pmid:18654630
- 20. Di Bari MA, Nonno R, Castilla J, D’Agostino C, Pirisinu L, Riccardi G, et al. Chronic wasting disease in bank voles: characterisation of the shortest incubation time model for prion diseases. PLoS Pathog. 2013;9(3):e1003219. pmid:23505374
- 21. Pirisinu L, Di Bari MA, D’Agostino C, Marcon S, Riccardi G, Poleggi A, et al. Gerstmann-Straussler-Scheinker disease subtypes efficiently transmit in bank voles as genuine prion diseases. Sci Rep. 2016;6:20443. pmid:26841849
- 22. Nonno R, Notari S, Di Bari MA, Cali I, Pirisinu L, d’Agostino C, et al. Variable Protease-Sensitive Prionopathy Transmission to Bank Voles. Emerg Infect Dis. 2019;25(1):73–81. pmid:30561322
- 23. Nonno R, Di Bari MA, Pirisinu L, D’Agostino C, Vanni I, Chiappini B, et al. Studies in bank voles reveal strain differences between chronic wasting disease prions from Norway and North America. Proc Natl Acad Sci U S A. 2020;117(49):31417–26. pmid:33229531
- 24. Pirisinu L, Di Bari MA, D’Agostino C, Vanni I, Riccardi G, Marcon S, et al. A single amino acid residue in bank vole prion protein drives permissiveness to Nor98/atypical scrapie and the emergence of multiple strain variants. PLoS Pathog. 2022;18(6):e1010646. pmid:35731839
- 25. Watts JC, Giles K, Patel S, Oehler A, Dearmond SJ, Prusiner SB. Evidence that bank vole PrP is a universal acceptor for prions. PLoS Pathog. 2014;10(4):e1003990. pmid:24699458
- 26. Espinosa JC, Nonno R, Di Bari M, Aguilar-Calvo P, Pirisinu L, Fernandez-Borges N, et al. PrPC Governs Susceptibility to Prion Strains in Bank Vole, While Other Host Factors Modulate Strain Features. J Virol. 2016;90(23):10660–9. pmid:27654300
- 27. Orru CD, Groveman BR, Raymond LD, Hughson AG, Nonno R, Zou W, et al. Bank Vole Prion Protein As an Apparently Universal Substrate for RT-QuIC-Based Detection and Discrimination of Prion Strains. PLoS Pathog. 2015;11(6):e1004983. pmid:26086786
- 28. Cosseddu GM, Nonno R, Vaccari G, Bucalossi C, Fernandez-Borges N, Di Bari MA, et al. Ultra-efficient PrP(Sc) amplification highlights potentialities and pitfalls of PMCA technology. PLoS Pathog. 2011;7(11):e1002370. pmid:22114554
- 29. Burke CM, Walsh DJ, Steele AD, Agrimi U, Di Bari MA, Watts JC, et al. Full restoration of specific infectivity and strain properties from pure mammalian prion protein. PLoS Pathog. 2019;15(3):e1007662. pmid:30908557
- 30. Watts JC, Giles K, Stohr J, Oehler A, Bhardwaj S, Grillo SK, et al. Spontaneous generation of rapidly transmissible prions in transgenic mice expressing wild-type bank vole prion protein. Proc Natl Acad Sci U S A. 2012;109(9):3498–503. pmid:22331873
- 31. Watts JC, Giles K, Bourkas ME, Patel S, Oehler A, Gavidia M, et al. Towards authentic transgenic mouse models of heritable PrP prion diseases. Acta Neuropathol. 2016;132(4):593–610. pmid:27350609
- 32. Otero A, Hedman C, Fernandez-Borges N, Erana H, Marin B, Monzon M, et al. A Single Amino Acid Substitution, Found in Mammals with Low Susceptibility to Prion Diseases, Delays Propagation of Two Prion Strains in Highly Susceptible Transgenic Mouse Models. Mol Neurobiol. 2019;56(9):6501–11. pmid:30847740
- 33. Mehra S, Bourkas MEC, Kaczmarczyk L, Stuart E, Arshad H, Griffin JK, et al. Convergent generation of atypical prions in knock-in mouse models of genetic prion disease. J Clin Invest. 2024;134(15):e176344. pmid:39087478
- 34. Fernandez-Borges N, Di Bari MA, Erana H, Sanchez-Martin M, Pirisinu L, Parra B, et al. Cofactors influence the biological properties of infectious recombinant prions. Acta Neuropathol. 2018;135(2):179–99. pmid:29094186
- 35. Erana H, Charco JM, Di Bari MA, Diaz-Dominguez CM, Lopez-Moreno R, Vidal E, et al. Development of a new largely scalable in vitro prion propagation method for the production of infectious recombinant prions for high resolution structural studies. PLoS Pathog. 2019;15(10):e1008117. pmid:31644574
- 36. Erana H, Diaz-Dominguez CM, Charco JM, Vidal E, Gonzalez-Miranda E, Perez-Castro MA, et al. Understanding the key features of the spontaneous formation of bona fide prions through a novel methodology that enables their swift and consistent generation. Acta neuropathologica communications. 2023;11(1):145. pmid:37679832
- 37. Kurt TD, Aguilar-Calvo P, Jiang L, Rodriguez JA, Alderson N, Eisenberg DS, et al. Asparagine and glutamine ladders promote cross-species prion conversion. J Biol Chem. 2017;292(46):19076–86. pmid:28931606
- 38. Burke CM, Mark KMK, Walsh DJ, Noble GP, Steele AD, Diack AB, et al. Identification of a homology-independent linchpin domain controlling mouse and bank vole prion protein conversion. PLoS Pathog. 2020;16(9):e1008875. pmid:32898162
- 39. Kobayashi A, Matsuura Y, Takeuchi A, Yamada M, Miyoshi I, Mohri S, et al. A domain responsible for spontaneous conversion of bank vole prion protein. Brain Pathol. 2019;29(2):155–63. pmid:30051525
- 40. Krance SH, Luke R, Shenouda M, Israwi AR, Colpitts SJ, Darwish L, et al. Cellular models for discovering prion disease therapeutics: Progress and challenges. J Neurochem. 2020;153(2):150–72. pmid:31943194
- 41. Arshad H, Watts JC. Genetically engineered cellular models of prion propagation. Cell Tissue Res. 2023;392(1):63–80. pmid:35581386
- 42. Mahal SP, Baker CA, Demczyk CA, Smith EW, Julius C, Weissmann C. Prion strain discrimination in cell culture: the cell panel assay. Proc Natl Acad Sci U S A. 2007;104(52):20908–13. pmid:18077360
- 43. Oelschlegel AM, Fallahi M, Ortiz-Umpierre S, Weissmann C. The extended cell panel assay characterizes the relationship of prion strains RML, 79A, and 139A and reveals conversion of 139A to 79A-like prions in cell culture. J Virol. 2012;86(9):5297–303. pmid:22379091
- 44. Bourkas MEC, Arshad H, Al-Azzawi ZAM, Halgas O, Shikiya RA, Mehrabian M, et al. Engineering a murine cell line for the stable propagation of hamster prions. J Biol Chem. 2019;294(13):4911–23. pmid:30705093
- 45. Walia R, Ho CC, Lee C, Gilch S, Schatzl HM. Gene-edited murine cell lines for propagation of chronic wasting disease prions. Sci Rep. 2019;9(1):11151. pmid:31371793
- 46. Arshad H, Patel Z, Amano G, Li LY, Al-Azzawi ZAM, Supattapone S, et al. A single protective polymorphism in the prion protein blocks cross-species prion replication in cultured cells. J Neurochem. 2023;165(2):230–45. pmid:36511154
- 47. Arshad H, Patel Z, Mehrabian M, Bourkas MEC, Al-Azzawi ZAM, Schmitt-Ulms G, et al. The aminoglycoside G418 hinders de novo prion infection in cultured cells. J Biol Chem. 2021;297(3):101073. pmid:34390689
- 48. Cartoni C, Schinina ME, Maras B, Nonno R, Vaccari G, Di Baria MA, et al. Identification of the pathological prion protein allotypes in scrapie-infected heterozygous bank voles (Clethrionomys glareolus) by high-performance liquid chromatography-mass spectrometry. J Chromatogr A. 2005;1081(1):122–6. pmid:16013608
- 49. Tuzi NL, Cancellotti E, Baybutt H, Blackford L, Bradford B, Plinston C, et al. Host PrP glycosylation: a major factor determining the outcome of prion infection. PLoS Biol. 2008;6(4):e100. pmid:18416605
- 50. Cancellotti E, Mahal SP, Somerville R, Diack A, Brown D, Piccardo P, et al. Post-translational changes to PrP alter transmissible spongiform encephalopathy strain properties. EMBO J. 2013;32(5):756–69. pmid:23395905
- 51. Nishina KA, Deleault NR, Mahal SP, Baskakov I, Luhrs T, Riek R, et al. The stoichiometry of host PrPC glycoforms modulates the efficiency of PrPSc formation in vitro. Biochemistry. 2006;45(47):14129–39. pmid:17115708
- 52. Katorcha E, Makarava N, Savtchenko R, D’Azzo A, Baskakov IV. Sialylation of prion protein controls the rate of prion amplification, the cross-species barrier, the ratio of PrPSc glycoform and prion infectivity. PLoS Pathog. 2014;10(9):e1004366. pmid:25211026
- 53. Supattapone S, Muramoto T, Legname G, Mehlhorn I, Cohen FE, DeArmond SJ, et al. Identification of two prion protein regions that modify scrapie incubation time. J Virol. 2001;75(3):1408–13. pmid:11152514
- 54. Sevillano AM, Aguilar-Calvo P, Kurt TD, Lawrence JA, Soldau K, Nam TH, et al. Prion protein glycans reduce intracerebral fibril formation and spongiosis in prion disease. J Clin Invest. 2020;130(3):1350–62. pmid:31985492
- 55. Ayers JI, Schutt CR, Shikiya RA, Aguzzi A, Kincaid AE, Bartz JC. The strain-encoded relationship between PrP replication, stability and processing in neurons is predictive of the incubation period of disease. PLoS Pathog. 2011;7(3):e1001317. pmid:21437239
- 56. Priola SA, Chesebro B. A single hamster PrP amino acid blocks conversion to protease-resistant PrP in scrapie-infected mouse neuroblastoma cells. J Virol. 1995;69(12):7754–8. pmid:7494285
- 57. Priola SA, Chabry J, Chan KM. Efficient conversion of normal prion protein (PrP) by abnormal hamster PrP is determined by homology at amino acid residue 155. Journal of Virology. 2001;75(10):4673–80. pmid:11312338
- 58. Christen B, Perez DR, Hornemann S, Wuthrich K. NMR Structure of the Bank Vole Prion Protein at 20 degrees C Contains a Structured Loop of Residues 165–171. Journal of Molecular Biology. 2008;383(2):306–12. pmid:18773909
- 59. Sigurdson CJ, Nilsson KP, Hornemann S, Heikenwalder M, Manco G, Schwarz P, et al. De novo generation of a transmissible spongiform encephalopathy by mouse transgenesis. Proc Natl Acad Sci U S A. 2009;106(1):304–9. pmid:19073920
- 60. Sigurdson CJ, Nilsson KP, Hornemann S, Manco G, Fernandez-Borges N, Schwarz P, et al. A molecular switch controls interspecies prion disease transmission in mice. J Clin Invest. 2010;120(7):2590–9. pmid:20551516
- 61. Sigurdson CJ, Joshi-Barr S, Bett C, Winson O, Manco G, Schwarz P, et al. Spongiform encephalopathy in transgenic mice expressing a point mutation in the beta2-alpha2 loop of the prion protein. J Neurosci. 2011;31(39):13840–7. pmid:21957246
- 62. Bett C, Fernandez-Borges N, Kurt TD, Lucero M, Nilsson KP, Castilla J, et al. Structure of the beta2-alpha2 loop and interspecies prion transmission. FASEB J. 2012;26(7):2868–76. pmid:22490928
- 63. Kyle LM, John TR, Schatzl HM, Lewis RV. Introducing a rigid loop structure from deer into mouse prion protein increases its propensity for misfolding in vitro. PLoS One. 2013;8(6):e66715. pmid:23825561
- 64. Kurt TD, Jiang L, Fernandez-Borges N, Bett C, Liu J, Yang T, et al. Human prion protein sequence elements impede cross-species chronic wasting disease transmission. J Clin Invest. 2015;125(6):2548. pmid:25961458
- 65. Gossert AD, Bonjour S, Lysek DA, Fiorito F, Wuthrich K. Prion protein NMR structures of elk and of mouse/elk hybrids. Proc Natl Acad Sci U S A. 2005;102(3):646–50. pmid:15647363
- 66. Perez DR, Damberger FF, Wuthrich K. Horse prion protein NMR structure and comparisons with related variants of the mouse prion protein. J Mol Biol. 2010;400(2):121–8. pmid:20460128
- 67. Gallagher-Jones M, Glynn C, Boyer DR, Martynowycz MW, Hernandez E, Miao J, et al. Sub-angstrom cryo-EM structure of a prion protofibril reveals a polar clasp. Nat Struct Mol Biol. 2018;25(2):131–4. pmid:29335561
- 68. Hoyt F, Standke HG, Artikis E, Schwartz CL, Hansen B, Li K, et al. Cryo-EM structure of anchorless RML prion reveals variations in shared motifs between distinct strains. Nature communications. 2022;13(1):4005. pmid:35831291
- 69. Solforosi L, Bellon A, Schaller M, Cruite JT, Abalos GC, Williamson RA. Toward molecular dissection of PrPC-PrPSc interactions. J Biol Chem. 2007;282(10):7465–71. pmid:17218310
- 70. Watts JC, Giles K, Serban A, Patel S, Oehler A, Bhardwaj S, et al. Modulation of Creutzfeldt-Jakob disease prion propagation by the A224V mutation. Ann Neurol. 2015;78(4):540–53. pmid:26094969
- 71. Kobayashi A, Hirata T, Nishikaze T, Ninomiya A, Maki Y, Takada Y, et al. alpha2,3 linkage of sialic acid to a GPI anchor and an unpredicted GPI attachment site in human prion protein. J Biol Chem. 2020;295(22):7789–98. pmid:32321762
- 72. Makarava N, Kovacs GG, Savtchenko R, Alexeeva I, Ostapchenko VG, Budka H, et al. A new mechanism for transmissible prion diseases. J Neurosci. 2012;32(21):7345–55. pmid:22623680
- 73. Wille H, Bian W, McDonald M, Kendall A, Colby DW, Bloch L, et al. Natural and synthetic prion structure from X-ray fiber diffraction. Proc Natl Acad Sci U S A. 2009;106(40):16990–5. pmid:19805070
- 74. Wang LQ, Zhao K, Yuan HY, Wang Q, Guan Z, Tao J, et al. Cryo-EM structure of an amyloid fibril formed by full-length human prion protein. Nat Struct Mol Biol. 2020;27(6):598–602. pmid:32514176
- 75. Qi YP, Wang JKT, McMillian M, Chikaraishi DM. Characterization of a CNS cell line, CAD, in which morphological differentiation is initiated by serum deprivation. Journal of Neuroscience. 1997;17(4):1217–25. pmid:9006967
- 76. Williamson RA, Peretz D, Pinilla C, Ball H, Bastidas RB, Rozenshteyn R, et al. Mapping the prion protein using recombinant antibodies. J Virol. 1998;72(11):9413–8. pmid:9765500
- 77. Safar JG, Scott M, Monaghan J, Deering C, Didorenko S, Vergara J, et al. Measuring prions causing bovine spongiform encephalopathy or chronic wasting disease by immunoassays and transgenic mice. Nat Biotechnol. 2002;20(11):1147–50. pmid:12389035
- 78. Polymenidou M, Moos R, Scott M, Sigurdson C, Shi YZ, Yajima B, et al. The POM monoclonals: a comprehensive set of antibodies to non-overlapping prion protein epitopes. PLoS One. 2008;3(12):e3872. pmid:19060956