Prion disease tempo determined by host-dependent substrate reduction

The symptoms of prion infection can take years or decades to manifest following the initial exposure. Molecular markers of prion disease include accumulation of the misfolded prion protein (PrP^Sc), which is derived from its cellular precursor (PrP^C), as well as downregulation of the PrP-like Shadoo (Sho) glycoprotein. Given the overlapping cellular environments for PrP^C and Sho, we inferred that PrP^C levels might also be altered as part of a host response during prion infection. Using rodent models, we found that, in addition to changes in PrP^C glycosylation and proteolytic processing, net reductions in PrP^C occur in a wide range of prion diseases, including sheep scrapie, human Creutzfeldt-Jakob disease, and cervid chronic wasting disease. The reduction in PrP^C results in decreased prion replication, as measured by the protein misfolding cyclic amplification technique for generating PrP^Sc in vitro. While PrP^C downregulation is not discernible in animals with unusually short incubation periods and high PrP^C expression, slowly evolving prion infections exhibit downregulation of the PrP^C substrate required for new PrP^Sc synthesis and as a receptor for pathogenic signaling. Our data reveal PrP^C downregulation as a previously unappreciated element of disease pathogenesis that defines the extensive, presymptomatic period for many prion strains.

Prions are transmissible agents responsible for incurable neurodegenerative diseases in humans and animals. Infection with prions is unique in that they may manifest after a lag phase that can exceed 5 decades from the time of exposure (1). The generation of infectivity from recombinant prion protein (PrP) (2) supports the “protein-only” hypothesis, whereby prion replication corresponds to a conformational transition from the host-encoded, cellular prion protein (PrP^C) substrate into a misfolded infectivity-associated form called PrP^Sc. Initiation of prion disease has an obligatory dependence upon the introduction of a PrP^Sc “seed,” and susceptibility to disease is blocked by PrP gene null alleles (3). Prion disease pathogenesis is associated with a progressive accumulation of PrP^Sc levels, with the qualification that PrP^Sc may encompass subforms that are biophysically and perhaps toxicologically distinct (4, 5).

Besides PrP^C, the Shadoo (Sho) glycoprotein is also expressed in the central nervous system. Sho has a similarity to the N-terminal region of PrP (6, 7) and interacts with several proteins that also interact with PrP^C (8). While the function of Sho remains to be determined, a notable discovery was that levels of the Sho protein are reduced (downregulated) in prion disease (7, 9). Since this effect is apparent in the preclinical phase of disease (10, 11), we speculated that this phenomenon might be of broader importance for pathogenesis. In particular, the status of PrP^C protein levels during prion infection has been investigated little, because of the unaltered steady-state levels of the corresponding mRNA (12, 13) and a co-occurrence of PrP^Sc that may be present in molar excess. Here, to investigate PrP^C, while avoiding the confounding presence of PrP^Sc, velocity gradient centrifugation was used to separate monomeric PrP^C from oligomeric and high molecular assemblies containing PrP^Sc (14). We then applied the conformation-dependent immunoassay (CDI), which measures a “hidden” epitope of PrP^Sc subsequent to its exposure by use of a chemical denaturant. By means of this 2-step procedure, we were able to analyze levels of different PrP species, including PrP^C, protease-resistant PrP^Sc (rPrP^Sc), and protease-sensitive PrP^Sc (sPrP^Sc) (5). Our studies demonstrated both quantitative and qualitative alterations in PrP^C, with a number of implications for our understanding of prion disease pathogenesis.

PrP^C downregulation at end stage in scrapie, Creutzfeldt-Jakob disease, and chronic wasting disease models. The procedure to couple velocity gradient centrifugation with CDI (as explained in the Introduction) has been validated previously for samples derived from hamsters (5) and humans (15). To extend this approach to the analysis of mouse PrP, we recalibrated conditions for the use of the monoclonal antibody 12B2 (mouse PrP epitope at residues 88–92) (ref. 16 and Figure 1, A and B). We then applied the recalibrated technique to the analysis of mouse brain homogenates; these homogenates were derived from WT mouse brains harvested at clinical end stage after infection with the Rocky Mountain Laboratory (RML) isolate of mouse-adapted scrapie prions. Our analyses demonstrated that 93% of the upper gradient fractions, fractions 9 and 10, are composed of PrP^C, while rPrP^Sc and sPrP^Sc make up the remaining 7% (Figure 1B). Since PrP^C is the predominant PrP isoform, we attempted to use the protein within these gradient fractions as a substrate in an in vitro assay to make PrP^Sc; this assay procedure is called protein misfolding cyclic amplification (PMCA) and is conceptually similar to PCR amplification of nucleic acids. However, proteins within the upper fractions of the velocity gradients were incompatible with our PMCA conditions and showed no detectable amplification, even though the levels of PrP^C and concentrations of PrP^Sc after seeding with infected brain homogenate were comparable to the PMCA control experiment with nonfractionated material (Supplemental Figure 1; see complete unedited blots in the supplemental material; supplemental material available online with this article; doi: 10.1172/JCI72241DS1). Nonetheless, the low infectivity of these fractions, as determined by the standard scrapie cell assay (SSCA; a cell monolayer assay to determine the infectious titer for prions), supports the notion that the major component of these upper fractions is indeed PrP^C (Figure 1D). In comparing RML prion-infected and noninfected animals, we discovered that total PrP^C levels were decreased by 41% (P < 0.001) (Figure 1B). Interestingly, the PrP-like protein Sho was found in the same gradient fractions as PrP^C, where it was also reduced in the RML-infected animals by approximately 48% of control values (P < 0.001) (Figure 1, A and C).

Figure 1

Separation of PrP isoforms and prion infectivity by sedimentation velocity in sucrose gradient. (A) The gradient fractionation profile of PrPs present in uninfected brain homogenates of WT FVB mice compared to that of those infected with the RML scrapie isolate strain after separation by ultracentrifugation in sucrose gradient. Fractions collected from the bottom of the tubes were treated with PK or left untreated and analyzed by (A) Western blot using anti-PrP mAb 12B2 and (B) CDI. Protein marker mass is shown in kDa for Western blots, and the asterisk indicates cross-reactivity between secondary antibody and IgG light chain present in the brain homogenates. The concentration of PrP^C (green), total PrP^Sc (brown), and rPrP^Sc (black) in fractions from triplicate samples was determined with CDI with Eu-labeled mAb 12B2 before or after PK treatment (14). (C) Densitometry analysis of the Western blot signal for Sho was estimated with ImageJ. (D) SSCA performed in L929 cells to estimate infectivity in gradient-fractionated RML prions. Data for uninfected animals appears on the left and that for RML-infected animals appears on the right. Data represent average ± SEM

To determine whether PrP^C downregulation is a broad-spectrum characteristic of prion infection, we applied similar analysis to end-stage brain material derived from WT mice infected with different strains of prions. Prion strains are microbiological isolates with distinct pathogenic properties that “breed true”; in molecular terms, they are thought to correspond to differently folded isoforms of PrP^Sc (17). For this purpose, we examined WT mice challenged with the 22L or 139A strains of scrapie prions, Syrian hamsters inoculated with the Sc237 scrapie prion strain, Tg cervid PrP (TgCe) mice (18) inoculated with chronic wasting disease (CWD) prions, Tg human PrP (TgHu) mice (19) challenged with a strain of human sporadic Creutzfeldt Jakob Disease (sCJD) prions, and bank voles inoculated with bank vole–adapted CWD prions. Our further analyses also included mice with a different genetic makeup. The most common allele of the mouse PrP gene (Prnp) is the Prnp^a allele encoding PrP^C-A. Here, we performed RML bioassays using mice that produce different levels of PrP^C-A, which correspond to Tg mice expressing 0.53 times endogenous PrP^C-A levels of WT mice (TgPrnp^a-AL mice) and TgPrnp^a mice (a widely used Tg, commonly referred to as tga20) overexpressing PrP^C-A by approximately 6-fold (20). Excluding TgPrnp^a mice and bank voles, marked decreases (P < 0.001) were noted in the net PrP^C (primarily present in fractions 9 and 10 in infected brains; Supplemental Figures 1–6), and these decreases ranged from 31% ± 3.9% to 68% ± 8.4% (± SEM) (Figure 2). As described for analyses with the RML prion isolate, Sho protein colocalized in fractions 9 and 10 with PrP^C in 22L- and 139A-infected brain material from WT mice at levels 42%–48% of the control values (22L strain, P < 0.001; 139A strain, P < 0.01) (Supplemental Figure 1C).

Figure 2

PrP^C downregulation as a common feature of prion disease. (A) Concentration of PrP^C (green), total PrP^Sc (brown), and PK-resistant PrP^Sc (black) in brains of end-stage infected animals as follows: WT FVB mice challenged with RML, 22L, or 139A scrapie prions; RML-infected TgPrnp^a-AL mice expressing 0.53 times endogenous PrP^C levels of WT mice; RML-infected TgPrnp^a mice overexpressing PrP^C; Syrian hamsters (SHa) inoculated with Sc237 scrapie prions; TgCe “cervidized” mice inoculated with CWD prions; TgHu “humanized” mice challenged with human sCJD prions; and bank voles (Bv109I Prnp) inoculated with CWD prions. (B) Percentage reduction in PrP^C relative to uninfected animals for data represented in A. Healthy mock-infected controls (0; white columns) were compared to inoculated animals (green columns). The columns and error bars represent average ± SEM measured in 3 brains, each sample tested in duplicate or triplicate by CDI. ***P < 0.001, **P < 0.01, determined by ANOVA.

PrP^C quality is altered by prion infection. For the 3 scrapie strains analyzed in WT mice (Figure 1A and Supplemental Figure 1A), we were surprised to discover that residual PrP^C present in infected animals exhibited an altered glycosylation profile. Generally, diglycosylated forms were underrepresented, and there were increased levels of monoglycosylated and unglycosylated PrP^C bands in fractions 9 and 10 (versus the abundance of diglycosylated, monoglycosylated, and unglycosylated protein found in disease-free animals; Figure 3A). In the next experiment, we used gel analysis to examine the upper velocity gradient fractions (fractions 10, 9, and 8) from the 3 mouse-adapted scrapie prion strains. For this purpose, samples were normalized according to PrP immunoblot signal when electrophoresed alongside equivalent gradient fractions from uninfected control animals. Protein samples were examined with or without PNGaseF deglycosylation (Figure 3, B and C). Although low levels of proteinase K–resistant (PK-resistant) PrP^Sc (∼8% of total rPrP^Sc and 2% of PrP^C) were detected in each strain/host combination in the upper fractions of velocity gradients (Figure 1 and Supplemental Figure 1), unexpectedly high amounts of the PrP C2 carboxyl-terminal fragment colocalized in the upper gradient fractions with protease-sensitive PrP^C (Figure 3, C and D). This is of interest because the C2 cleavage fragment is generally considered to be a rPrP^Sc product, not a PrP^C product (21). Furthermore, comparison with the untreated samples indicates that this C2 fragment is primarily in its unglycosylated form (Figure 3B).

Figure 3

Glycosylation and fragmentation of PrP^C altered by prion infection. (A) PrP^C glycoform ratios for protein in gradient fractions 10, 9, and 8 for WT mouse brain and brains of mice with end-stage disease caused by RML, 22L, or 139A scrapie prions were determined for Western blots represented in Figure 1A and Supplemental Figure 1A. Diglycosylated, monoglycosylated, and unglycosylated forms are shown by circles, squares, and diamonds, respectively. The columns and error bars represent average ± SEM measured in 3 brains, each sample tested in duplicate or triplicate by CDI. The same gradient fractions for the 3 mouse-adapted scrapie samples were normalized by immunoblot signal and compared to uninfected controls (“healthy”) in blot analyses (B) before and (C) after PNGaseF deglycosylation. (D) Densitometry was performed to evaluate the net levels of full-length PrP and its C-terminal fragments (PrP C1 and PrP C2), as shown in C.

Preclinical PrP^C downregulation in infected WT mice. As Sho downregulation occurs preclinically (10, 11), we next assessed a time course of WT mice challenged with RML prions or negative control inoculum. Levels of Sho and the different PrP species were analyzed, showing a coincident appearance of the disease-associated PrP C2 fragment and Sho reduction by 120 days after inoculation (dpi), a finding in line with previous reports (10) (Figure 4, A and B). Using velocity gradient centrifugation plus CDI analysis on the time course series, rPrP^Sc began to increase by day 30 and continued to accumulate throughout the disease course (Figure 5A), a trend that paralleled a determination of infectious titer by scrapie cell assay in L929 cells (Figure 4C). However, these data illustrated a downturn for the rate of PrP^Sc replication at 60 dpi (Figure 5A). A biphasic profile was seen for PrP^C in the infected mice, with a marked decrease beginning 60 dpi (Figure 5B), suggesting an inverse relationship to the aforementioned rPrP^Sc levels. This effect was even more noticeable, as it diverged from the slow rise in PrP^C measured in aging mock-inoculated controls (Figure 5B).

Figure 4

Time course analysis of control and pathogenesis-related proteins in infected mouse brain (RML isolate). (A) Samples were analyzed by Western blot of homogenates from sagittally bisected brain. “H” and “S” designate healthy and sick (prion-infected) animals. Antibody probes include the following: untreated samples for total PrP, PK-digested samples for PrP^Sc, fragmentation products (C2, C1) for PrP shown subsequent to PNGase F digestion, and Sho. (B) Percentage change for PrP C2 fragment, total PrP, and Sho (RML Sho) is presented for the time course in infected animals. Baseline Sho levels from mock-infected animals (mock Sho) are also shown. Values were determined from densitometric analysis of blots in A. (C) SSCA performed in L929 cells to estimate time course of infectivity; the dotted line represents assay threshold. The columns and error bars represent average ± SEM measured in 3 brains, each sample tested in duplicate or triplicate by CDI.

Figure 5

Preclinical drop in PrP^C levels, Sho levels, and PrP^Sc replication rate. Time courses were performed for (A and B) WT FVB mice and (C–E) hemizygous Prnp^0/+ mice infected with RML prions and compared to mock-inoculated controls. (A and C) CDI analysis of brain material was used to evaluate rPrP^Sc levels (black diamonds) and to calculate the rPrP^Sc replication rate (red triangles). (B and D) Corresponding CDI data for brain PrP^C for mice in A and C: yellow squares represent control mice and green circles represent RML-infected mice. (E) Western blot analysis was used to compare Sho levels over time in hemizygous Prnp^0/+ mice either inoculated with RML prions (“S” for sick) or mock-inoculated (“H” for healthy) (see analogous Sho data for WT in Figure 4A). The data points and error bars represent average ± SEM measured in 3 brains, each tested in duplicate or triplicate by CDI. ***P < 0.001, **P < 0.01, *P < 0.05, determined by ANOVA.

In the interval of 60 to 140 dpi, PrP^C levels in infected mice fell by over 40%, from 1,200 to 700 ng/ml. To ascertain whether the preclinical drop in PrP^C levels was merely attributable to depletion of PrP^C substrate to make new PrP^Sc, we fit the data deriving from CDI analyses of RML prion-infected animals with data obtained from our previous work on RML prion-infected bigenic mice (22). Specifically, the expression rate of PrP^C (r_E) was derived using the equation r_E = ([PrP^C]/2)/t_1/2), where the half-lives (t_1/2) for net brain PrP^Sc (i.e., sPrP^Sc plus rPrP^Sc) and brain PrP^C were determined to be 36 hours and 18 hours, respectively (see Table 1 and ref. 22). The accumulation rates (r_A) of PrP^Sc and rPrP^Sc were then obtained by linear fit of the data at 90 to 140 dpi with RML isolate: PrP^Sc r_A = –672 + (8.12 × time in days) (R = 0.99); rPrP^Sc r_A = –62.4 + (0.77 × time in days) (R = 0.99). Clearance rates (r_CL) of PrP^Sc and rPrP^Sc were calculated from the half-life of mouse PrP^Sc determined previously (22) in RML prion-infected doxycycline-regulated bigenics and accumulation rates of PrP^Sc and rPrP^Sc obtained in this study: r_CL = (r_A/2)/t_1/2. Finally, the conversion rate (r_C) of PrP^C to PrP^Sc in the brain was determined from the accumulation rate of PrP^Sc corrected for clearance: r_C = r_A – r_CL. According to these calculations, the expression rate of PrP^C was estimated to be approximately 65-fold higher than the conversion rate to total PrP^Sc and approximately 700-fold higher than the conversion rate to rPrP^Sc, thus implying that depletion of PrP^C for the synthesis of PrP^Sc is a negligible factor in determining the drop in steady-state levels of PrP^C (i.e., downregulation). Also, speaking to degenerative changes, PrP^C downregulation preceded alterations in the neuron marker NeuN, and we observed no changes in the GPI-linked Thy1.2 glycoprotein associated with caveolae (ref. 23 and Supplemental Figure 7).

Table 1

Relationship between expression rate of mouse PrPC and conversion rate to PrPSc after infection with RML prions

PrP^C downregulation in models with different levels of starting PrP^C. Although prior studies have defined a profound effect of PrP^C levels in healthy mice on disease incubation times subsequent to inoculation (24, 25), a downregulation effect was never considered in the interpretation of these studies. Consequently, we investigated prion infections in mice underexpressing or overexpressing PrP^C.

To evaluate underexpression, we conducted a time course for RML prion-infected FVB/N hemizygous knockout (Prnp^0/+) mice and controls inoculated with normal brain homogenate. Hemizygous Prnp^0/+ mice are established as generating high levels of infectivity and PrP^Sc months before exhibiting symptoms of prion disease (26). To this effect, we observed that PrP^Sc began to accumulate in the brains of these animals between 60 and 100 dpi (Figure 5C). As in the case of WT animals, a parallel preclinical decrease in PrP^C levels and PrP^Sc replication rate was observed in infected hemizygotes beginning at 100 dpi (nearly 300 days before the expected time in which they succumb to disease; ref. 26), and both parameters continued dropping throughout the time course, reaching levels approximately 40% lower than those of the controls (P < 0.001) (Figure 5, C and D). This was accompanied by a gradual decrease in Sho throughout the duration of the time course (Figure 5E). While this cohort of animals has yet to reach disease end point, the effects of underexpression later in the disease course were explored using the TgPrnp^a-AL mouse line made on a Prnp^0/0 background and expressing 53% of WT levels of PrP^C-A (Supplemental Figure 8). When measured in RML prion-infected mice at disease end point (400–430 dpi), the absolute PrP^C levels in ng/ml lysate were downregulated to the lowest in our survey (Figure 2).

Uninfected adult TgPrnp^a mice overexpress 6 to 7 times PrP^C levels of WT mice and have incubation periods of 60 to 80 dpi (20, 27). By CDI, the highest PrP^C levels in uninfected TgPrnp^a mice (C57BL6 background) were approximately 3 times those seen in WT FVB/N mice (Figure 2A). Infected TgPrnp^a animals and normal brain homogenate inoculated controls were investigated in a time course series (Supplemental Figure 9, A–C). TgPrnp^a animals were also discovered to exhibit a biphasic PrP^C profile, albeit with a smaller effect size (11%, measured between 45 and 60 dpi), that was not significant due to the interindividual variability (±12%) (Supplemental Figure 9B). Nonetheless, a downturn in rPrP^Sc replication rate was noted when comparing the same 2 time points (Supplemental Figure 9A), offering an analogous trend to the effect noted in WT and hemizygous Prnp^0/+ mice (Figure 5, A and C). Like PrP^C, Sho levels were affected little during the disease process in these mice (Supplemental Figure 9C).

These data suggest that notable effect sizes for PrP^C downregulation are associated with long prion incubation times and vice versa. By plotting the incubation times of RML prions in these 3 hosts (all with the same Prnp^a genotype but expressing different levels of PrP^C) versus terminal concentrations of PrP^C and rPrP^Sc, we found that prion infection caused a highly significant shift in the slope of the linear regression between incubation time of the disease and end point levels of PrP^C (Figure 6A and ref. 4). This shift in slope reflects a correlation for PrP^C downregulation in the infected animals, with the largest difference in the data with or without infection occurring in the TgPrnp^a-AL animals. A correlation was also apparent after plotting incubation time and the ratio between end point PrP^C and rPrP^Sc (Figure 6B). Cumulatively, downregulation of PrP^C was linked to the low expression levels of PrP^C and high levels of the accumulated rPrP^Sc at disease end stage and is associated with prolonged incubation time (R = 0.99, P < 0.001). We next applied a similar plotting scheme to the spectrum of prion strains and disease models inoculated in our experiments. We found that this effect is broadly applicable despite independent variables that included 4 different Prnp genotypes, different starting PrP^C levels, and 6 different strains of prions (Figure 6C; R = 0.77, P < 0.05).

Figure 6

Relationships between end point PrP^C/PrP^Sc ratios and prion incubation period. (A) Residual PrP^C plotted against incubation time for the RML prion isolate in 3 mouse genotypes expressing different amounts of the PrP-A protein. Data points for mock-inoculated controls (yellow line) and end-stage RML prion-infected animals (green line) represent, from left to right, TgPrnp^a-AL, WT FVB, and TgPrnp^a mice all infected with the RML prion isolate. (B) Ratios of residual PrP^C/residual PrP^Sc at disease end point plotted against incubation time for mouse genotypes presented in A. (C) Cumulative data for residual PrP^C/residual PrP^Sc at disease end point for 8 prion strain/host genotype combinations.

Assessing PrP^C downregulation in bank voles. Bank vole–adapted CWD prions produce the fastest prion disease in a naturally occurring animal host and hence are important in assessing the interplay between prion replication and disease pathogenesis (28). A time course series of samples was again investigated (Supplemental Figure 9, D–F). Unlike the other prion-infected specimens examined, PrP^Sc generated in the bank vole was primarily composed of oligomers sensitive to PK digestion (Supplemental Figure 6), a form that has been described as the most infectious PrP species (29). In this disease model, neither PrP^C nor Sho levels differed between CWD- and mock-inoculated animals despite the low but continuous accumulation of rPrP^Sc observed in the brain 20 dpi (Supplemental Figure 9, E and F). However, a biphasic replication profile was again noted for rPrP^Sc, with a maximum increase from 20 to 25 dpi and steady decline during the final 5 days of disease (Supplemental Figure 9D). The short incubation period in CWD-infected bank voles may be related to the ability to rapidly generate PK-sensitive oligomers in the absence of Sho downregulation (Supplemental Figure 6 and Supplemental Figure 9F) (and PrP^C downregulation) that may be typically triggered by rPrP^Sc (10, 11).

PrP^C downregulation in cultured cells. We wanted to investigate whether preclinical PrP^C downregulation observed in vivo is associated with PrP^Sc accumulation (as previously demonstrated for Sho; refs. 10, 11) or is a side effect of neurodegenerative change. We explored this possibility using monocultures of cells that lack pathological signs upon prion infection (30). First, we compared N2a neuroblastoma expressing mouse PrP^C-A (N2a) (31) and rabbit RK13-Gag cells expressing elk PrP^C (RK13 Elk21) (32) chronically infected with RML prions or elk CWD prions, respectively. In both cases, corresponding PrP^C reductions were approximately 35% and 55% compared with noninfected controls (Figure 7, A and B). Next, to parallel in vivo time course experiments in the brain, we carried out de novo infections, monitoring RK13 cells stably expressing either mouse Prnp^a or Prnp^b for 5 weeks after treatment with RML-infected brain homogenate (Figure 7 and Supplemental Figure 10). Similar infections with mouse-adapted scrapie prions in vivo have demonstrated drastically different susceptibilities for mice with these genotypes, where mice with the Prnp^a genotype exhibit significantly shorter prion incubation periods than those with the Prnp^b genotype (33). In parallel, expression of PrP^C-A made RK13 cells highly susceptible to RML prion infection (Figure 7, C and D; Supplemental Figure 10A; and refs. 25, 34). In comparison to mock-inoculated cells, propagation of PrP^Sc resulted in a decrease in PrP^C until 21 dpi, with levels then recovering to control levels by 35 dpi (Figure 7D). The converted rPrP^Sc represented 0.3%–0.5% of homologous PrP^C expressed in these cells. In contrast, RK13 cells expressing PrP^C-B were resistant to infection (Figure 7C and Supplemental Figure 10B). Since downregulation did not occur in cells expressing PrP^C-B (Supplemental Figure 10C), the effect seen in Prnp^a-expressing cells is not a response against administration of the brain homogenate but is instead a consequence of a productive infection. Cumulatively, the cell data provide evidence that neither neuronal loss accompanying prion disease nor conversion of PrP^C to PrP^Sc can account, per se, for the PrP^C downregulatory effect.

Figure 7

PrP^C downregulation examined in cell culture models and by PMCA. (A) Western blot analysis was used to show PK sensitivity of cell lysates derived from uninfected N2a, ScN2a (chronically infected with RML scrapie prions), uninfected RK13 Elk21^–, and RKE21⁺ (chronically infected with elk CWD prions). (B) CDI was used to evaluate the relative downregulation of PrP^C in these cell lines. (C) Western blot analysis was used to show PK sensitivity of RK13 cells stably expressing either PrP^C-A or PrP^C-B following exposure to RML prions. (D) CDI was applied to quantify the PrP^C levels in RK13 cells stably expressing PrP^C-A following treatment with healthy (square) or RML-infected brain material (circle). Normal RK13 cells treated with RML-infected brain material (box with circle) were used as a control. (E) PrP^Sc replication rate was evaluated using double PMCA titrations to assess the effect of decreasing PrP^C input and titrated amounts of PrP^Sc “seed” from mouse-adapted RML prions. The columns and data points and error bars represent average ± SEM measured in triplicate. **P < 0.01, ***P < 0.001, determined by ANOVA.

Impact of PrP^C downregulation on prion replication rate. Since prion replication rate decreased in tandem with PrP^C downregulation in our time course studies, we investigated the relationship between these 2 events by simulating rPrP^Sc propagation during the prion infection process in vitro with PMCA (14, 35). First, we examined PMCA products for experiments seeded with increasing amounts of 139A prions in a constant amount of PrP^C; here, PrP^Sc amplification was found to be a saturable component at PrP^Sc seed concentrations of ≥1 ng/ml (Supplemental Figure 11A). This may be analogous to the decreased PrP^Sc replication rate at end point prion disease in the absence of PrP^C downregulation, as described above for TgPrnp^a mice and bank voles. In experiments holding the RML PrP^Sc “seed” constant, dropping PrP^C input in the range of 1,100 to 1,400 ng/ml (similar to the change in PrP^C in early versus end stage disease in WT mice; Figure 5B) had a marked impact upon replication rate (Supplemental Figure 11B), thus indicating that PrP^C is a limiting factor, as previously reported (36). To simulate conditions more precisely over the course of prion infection, RML PrP^Sc seeds and PrP^C substrate were inversely titrated to test the impact on amplification. At PrP^C concentrations below approximately 350 ng/ml and PrP^Sc seed concentrations of ≥1 ng/ml PrP^Sc, the replication rate was held below an amplification index of 20-fold, apparently due to saturation with an abundance of PrP^Sc (Figure 7E). However, at higher ng/ml values of PrP^C, these two traces diverge and hence are associated with a concomitant increase in replication rate, up to a factor of 110-fold (Figure 7E). These inverse titration experiments were extended to include 139A mouse-adapted prions (Supplemental Figure 11C), human sCJD prions (Supplemental Figure 11D), and Syrian hamster Sc237 prions (Supplemental Figure 11E). In every case, a uniform biphasic response was observed for each set of PMCA reactions, in which the most efficient amplification occurred in conditions paralleling an early stage of infection, with a low concentration of PrP^Sc and a high concentration of PrP^C. Therefore, our PMCA data indicate that rising PrP^Sc levels and falling PrP^C levels will converge to suppress prion replication. Furthermore, these results point to an affect of PrP^C downregulation beginning halfway through the incubation time interval in WT mice (Figure 5B).

Two unaddressed factors for prion infection are their long incubation periods and observed discrepancies between PrP^Sc levels and clinical disease (4, 37). However, using capture and conformational antibodies to inventory the different PrP species in the brain (i.e., rPrP^Sc, sPrP^Sc, and PrP^C), we have defined a new variable in disease pathogenesis, namely preclinical downregulation of PrP^C. This downregulation does not emanate from changes in PrP mRNA abundance (12, 13, 38).

PrP^C is indisputably required for the creation of PrP^Sc (2, 3, 24), but depletion by the law of mass action (i.e., a finite pool of PrP^C “substrate” being consumed by a thermodynamically favorable conversion to PrP^Sc) is implausible as the rate of PrP^C synthesis in the brain outstrips the rate of formation of total PrP^Sc and rPrP^Sc by about 1 and 2 orders of magnitude, respectively (Table 1). It is also known from metabolic labeling of chronically infected cell cultures that only approximately 1% to 5% of PrP^C converts into PrP^Sc (39, 40). Declining levels of PrP^C in the course of protracted prion infections were in most instances shadowed by a drop in endogenous mouse Sho (refs. 10, 11; Figures 1, 4, and 5; and Supplemental Figures 1 and 9), a cousin of PrP^C that does not modulate creation of PrP^Sc and is not converted to a protease-resistant isoform (41). Depletion of PrP^C as a simple consequence of synaptic damage or neuronal cell death is unlikely, because PrP^C downregulation occurred nearly 300 days before clinical disease in hemizygous Prnp^0/+ mice and was absent in certain rapid models with an abundance of vacuolation at end stage disease (i.e., TgPrnp^a mice and bank voles; Figure 5D and Supplemental Figure 9, B and E). Moreover, PrP^C downregulation was detected in cell culture models that are undergoing mitosis and have no synapses (Figure 7, A–D). The recovery phase noted 2 weeks later in these de novo infections may reflect selection pressures operating within the cultures of infected but mitotically active cells. From these observations, we suggest first that a host protective response is operating at the level of a cellular biological mechanism, presumably a form of proteostasis. Second, we suggest this response is triggered by the presence of rPrP^Sc and reduces net levels of PrP^C and Sho by a “bystander” effect (Figure 8 and refs. 8, 10, 11). The reduction in net PrP^C levels defined by our experiments stands in apparent contrast to a redistribution effect in persistently infected cells, in which cell surface PrP is reduced and immunohistochemical signal in a trans-Golgi network subcompartment is increased (42).

Figure 8

Summary of PrP^C and Sho downregulation in a prion-infected cell by an unidentified degradation pathway. (A) In a healthy cell, the balance of synthesis rate and degradation rate establish the steady-state level of PrP^C and Sho. (B) We assert that prion-infected cells are characterized by two processes. The first is the well-known biosynthetic conversion of PrP^C to PrP^Sc (top part of cell). We hypothesize a second process (bottom part of the cell) in which the accumulation of PrP^Sc triggers (open arrow) a proteostatic degradative response (Pac-Man shape). This in turn reduces the steady-state concentrations of PrP^C and Sho (indicated by smaller text) by a “bystander effect”. Reduced PrP^C will have an impact on PrP^Sc synthesis (dotted green line). An additional effect of PrP^C reduction may be to attenuate toxic signaling (see the Discussion), but this concept is omitted from the diagram for the sake of simplicity.

Thus far, we have not identified the tell-tale signs of a single canonical proteostatic pathway, such as eIF-2αP (translation axis of the unfolded protein response) (43), LC3-II (autophagy), or ubiquitin (proteasome system) (Supplemental Figure 7). The largely parallel downregulation of PrP^C and Sho indicates remodeling of a membrane microdomain that includes these proteins but does not include Thy-1.2 (Supplemental Figure 7A). However, the changes in the quality of PrP^C (less diglycosylated, preponderance of C2 fragment) contrast with an across-the-board change that might be expected for a hypotheses of mass action depletion or cell death and were unexpected. The C2 aspect is especially notable as Watts and colleagues recently argued that Sho is degraded via an endocytic pathway that requires it to be physically trafficked with PrP^Sc (10). A component of this argument was the inverse correlation between Sho levels and the PrP C2 proteolytic fragment (10), with the latter hypothesized as a stable intermediate in the lysosomal degradation of PrP^Sc (44, 45). However, we have shown that models for other neurodegenerative diseases with activated lysosomes lack downregulation of Sho (11). Although our analyses of total cell lysates reaffirm a correlation between the reduction in Sho levels and the appearance of the protease-resistant PrP C2 proteolytic fragment toward disease end point (Figure 4A), our analyses of fractionated cell lysates demonstrated a surprising abundance of C2 colocalized in the upper gradient fractions. This suggests that protease-resistant C2 found toward disease end point derives from a protease-sensitive, and thus partly PrP^C-like, precursor molecule. While the identity of an endoproteolytic “C2 PrPase” (like C1 PrPase; ref. 46) is unknown, progress in this area will help us to understand early events in PrP^Sc synthesis and early host responses. Another area of interest will be whether other parameters, such as a limited concentration of a cofactor (e.g., phosphatidylethanolamine; refs. 47, 48), can also contribute to a curtailment in the exponential phase for prion replication.

Prior studies have invoked a hypothetical toxic subform of PrP to explain the beneficial effects of a genetic reduction in PrP^C levels that cause a delay between plateauing infectivity levels and the onset of clinical symptoms (4, 49). Here, an alternative explanation emerges from the observed changes with time in PrP isoforms. We show that the ratio between residual PrP^C seen in end-stage disease and the amount of PrP^Sc is clearly correlated with the time lag to disease onset, and present experimental manipulations strongly suggest that this ratio in fact determines time lag. Within this formula, when PrP^Sc and infectivity reach a plateau (4), the extent of PrP^C downregulation becomes the key variable. The difference between quickly and slowly evolving disease can be seen clearly in the different slopes shown in Figure 6A, and we therefore suggest that the slowly evolving infection of the central nervous system (most typical of natural prion disease) can be seen to arise from downregulation of the very same substrate required both for de novo PrP^Sc synthesis and for pathogenic signaling, as defined by grafting experiments (50) and by use of regulated PrP^C expression (49). Interestingly, while the need to hypothesize a toxic subform of PrP^Sc may be less urgent, PrP^C downregulation can be reconciled with the measurements that led some to posit a 2-phase model for prion disease (4) or a form of substrate depletion predicted from analysis of prion disease kinetics (51). Cumulatively, our data show that slowly evolving pathogenesis of preclinical disease is caused by depletion of the PrP^C substrate that is required for PrP^Sc synthesis and toxic signaling. We infer that the natural phenomenon of PrP^C downregulation is partially effective in holding prion disease in check in WT animals. Drugs that enhance this protective response to further reduce PrP^C may have a potent beneficial effect in treating prion disease.

View Supplemental data

We thank Witold Surewicz for recombinant PrP used for calibration in CDI, Earl Poptic from Cleveland Clinic Hybridoma Core Facility for production of 8H4 mAb, and R. Kascsak, H. Rezai, and S. Kar for providing antibodies. We thank F. Jirik (University of Calgary, Calgary, Alberta, Canada) and V. Sim (University of Alberta) for breeding pairs of TgPrnp^a mice. This work was supported by grants from NINDS (NS074317), CDC (UR8/CCU515004), the Charles S. Britton Fund, the Canada Foundation for Innovation, the Canadian Institutes of Health Research (MOP36377), the Alberta Prion Research Institute, and Alberta Innovates — Health Solutions for graduate and postgraduate funding for A. Lau and C.E. Mays and a Scientist award to D. Westaway.

Address correspondence to: David Westaway, University of Alberta, Centre for Prions and Protein Folding Diseases, 204 Brain and Aging Research Building, Edmonton, Alberta, Canada. Phone: 780.492.9024; Fax: 780.492.9352; E-mail: david.westaway@ualberta.ca. Or to: Jiri G. Safar, Case Western Reserve University, 11100 Euclid Ave., Cleveland, Ohio 44106, USA. Phone: 216.368.4609; Fax: 216.368.4090; E-mail: jiri.safar@case.edu.

Conflict of interest: The authors have declared that no conflict of interest exists.

Reference information: J Clin Invest. 2014;124(2):847–858. doi:10.1172/JCI72241.

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