Neutralizing Mutations Significantly Inhibit Amyloid Formation by Human Prion Protein and Decrease Its Cytotoxicity

Journal Pre-proof Neutralizing Mutations Significantly Inhibit Amyloid Formation by Human Prion Protein and Decrease Its Cytotoxicity Jun-Jie Huang, Xiang-Ning Li, Wan-Li Liu, Han-Ye Yuan, Yuan Gao, Kan Wang, Bo Tang, Dai-Wen Pang, Jie Chen, Yi Liang PII:

S0022-2836(19)30697-7

DOI:

https://doi.org/10.1016/j.jmb.2019.11.020

Reference:

YJMBI 66338

To appear in:

Journal of Molecular Biology

Received Date: 12 July 2019 Revised Date:

22 October 2019

Accepted Date: 26 November 2019

Please cite this article as: J.-J. Huang, X.-N. Li, W.-L. Liu, H.-Y. Yuan, Y. Gao, K. Wang, B. Tang, D.W. Pang, J. Chen, Y. Liang, Neutralizing Mutations Significantly Inhibit Amyloid Formation by Human Prion Protein and Decrease Its Cytotoxicity, Journal of Molecular Biology, https://doi.org/10.1016/ j.jmb.2019.11.020. This is a PDF file of an article that has undergone enhancements after acceptance, such as the addition of a cover page and metadata, and formatting for readability, but it is not yet the definitive version of record. This version will undergo additional copyediting, typesetting and review before it is published in its final form, but we are providing this version to give early visibility of the article. Please note that, during the production process, errors may be discovered which could affect the content, and all legal disclaimers that apply to the journal pertain. © 2019 Elsevier Ltd. All rights reserved.

Neutralizing Mutations Significantly Inhibit Amyloid Formation by Human Prion Protein and Decrease Its Cytotoxicity Jun-Jie Huang 1, Xiang-Ning Li 1, Wan-Li Liu 1, Han-Ye Yuan 1, Yuan Gao 1, Kan Wang 1, Bo Tang 2, Dai-Wen Pang 2, Jie Chen 1 and Yi Liang 1, ∗ 1 - Hubei Key Laboratory of Cell Homeostasis, College of Life Sciences, Wuhan University, Wuhan 430072, China 2 - Key Laboratory of Analytical Chemistry for Biology and Medicine (Ministry of Education), College of Chemistry and Molecular Sciences, Wuhan 430072, China

Running Title: Protective polymorphism decreases PrP toxicity

∗

Correspondence to Yi Liang: [email protected].

Abbreviations used: ANS, 8-Anilino-1-naphthalene-sulfonic acid; CD, Circular dichroism; DAPI, 4’,6’-diamino-2-phenylindole; DIC, differential interference contrast; FTIR, Fourier transform infrared; GdnHCl, guanidine hydrochloride; PI, propidium iodide; PrP, prion protein; PrPC, the normal cellular prion protein; PrPSc, the abnormal pathologic prion protein; ROS, reactive oxygen species; TAMRA, 5(6)-carboxy-tetramethylrhodamine electron

microscopy;

ThT,

N-succinimidyl

thioflavin

encephalopathy. 1

T;

TSE,

ester;

TEM,

transmissible

transmission spongiform

Abstract Prion diseases, such as Creutzfeldt-Jakob disease and bovine spongiform encephalopathy, are fatal neurodegenerative diseases that affect many mammals including humans, and are caused by the misfolding of prion protein (PrP). A naturally occurring protective polymorphism G127V in human PrP has recently been found to greatly attenuate prion diseases, but the mechanism has remained elusive. We herein report that the hydrophobic chain introduced in G127V significantly inhibits amyloid fibril formation by human PrP, highlighting the protective effect of the G127V polymorphism. We further introduce an amino acid with a different hydrophobic chain (Ile) at the same position and find that G127I has similar protective effects as G127V. Moreover, we show that these two neutralizing mutations, G127V and G127I, significantly decrease the human PrP cytotoxicity resulting from PrP fibril formation, mitochondrial damage, and elevated ROS production enhanced by a strong prion-prone peptide PrP 106-126. These findings elucidate the molecular basis for a natural protective polymorphism in PrP and will enable the development of novel therapeutic strategies against prion diseases.

Keywords: prion protein; neutralizing mutation, protein aggregation, prion diseases, protein phase separation

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Introduction Prions, meaning “protein infectious agents,” were initially described by Stanley B. Prusiner [1]. Prion protein (PrP) misfolding in many mammals, including humans, causes prion disease or transmissible spongiform encephalopathy (TSE) [2-6]. Furthermore, prion-like diseases such as Alzheimer’s disease and amyotrophic lateral sclerosis follow similar principles of prion disease [4,7-10]. Key steps for the transformation of the cellular PrP, PrPC, into the pathological PrP, PrPSc [1,4,12,13], are the formation of the PrPC-PrPSc complex followed by transformation into the PrPSc-PrPSc complex [1,4,11-18]. The N-terminal domain of PrPC is disordered and the C-terminal domain has α-helical structure [19]. The primary structure of PrPSc is the same as that of PrPC, but there are critical differences between the two isoforms with respect to their β-sheet content, solubility in detergents, resistance to protease activity, and resulting mitochondrial dysfunctions [1,6,12,14,20-23]. Previous studies have established that a peptide (PrP 106-126) from wild-type human PrP owns some pathogenic and physicochemical properties of PrPSc, leading to its use as a synthetic surrogate for PrPSc to induce mitochondrial damage [21,24] and PrPSc-like cytotoxicity [25-27]. A naturally occurring protective polymorphism G127V in human PrP has been identified in people living in regions where kuru was prevalent [28], and has been recently reported to greatly attenuate prion disease [29]. The G127V polymorphism, in which G127 and V127 are described as “wild-type” and “mutant”, respectively, is located at the boundary between the N-terminal domain and the C-terminal domain of

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PrPC [19,30,31]. Molecular dynamic simulations and structural features of the mutated protein from nuclear magnetic resonance (NMR) studies [30,31], as well as in vitro aggregation studies [32], have suggested that the mutation hinders fibril formation and elongation by preventing monomer-monomer and monomer-fibril interactions. Liquid-liquid phase separation by low-complexity domains of proteins such as PrPC and various heterogenous nuclear ribonucleoproteins (hnRNPs) is the first step of protein aggregation, followed by a liquid-to-solid phase transition, which generates membrane-less organelles and drives pathological fibril formation, which are critical events functionally linked to prion disease and prion-like diseases [33-36]. Very recently, wild-type PrPC has been reported to be able to undergo liquid-liquid phase separation in phosphate-buffered saline (PBS) buffer [35]. In this paper, we studied the effect of the G127V polymorphism in different cell-free and cell assays to clarify the mechanism underlying the very interesting protective effect of G127V against infection with prion disease observed in humans and animal models. First, we reported that the hydrophobic chain introduced in G127V modulated the in vitro phase separation and significantly inhibited amyloid fibril formation by human PrP, highlighting the protective effect of the G127V polymorphism. Second, we introduced an amino acid with a different hydrophobic chain (Ile) at the same position and found G127I to have protective effects similar to those of G127V. Third, we observed significantly fewer PrP fibrils in G127V PrP-expressing cells or G127I PrP-expressing cells than in wild-type PrP-expressing

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cells. Finally, our analysis in cells expressing wild-type and mutated PrP showed that PrP 106-126 caused much more severe mitochondrial impairment, reactive oxygen species (ROS) production, and cytotoxicity in wild-type PrP-expressing cells than in G127V PrP-expressing cells or G127I PrP-expressing cells. These findings reveal the molecular basis for a natural protective polymorphism in the human PrP, which could be exploited to develop therapeutic strategies against prion disease.

Results Neutralizing mutations modulate the liquid-liquid phase separation of PrPC To understand why a natural protective polymorphism in PrP, G127V, is refractory to prion disease, we first tested the hypothesis that G127V PrPC might be less efficient than wild-type PrPC in liquid-liquid phase separation in PBS buffer. Wild-type PrPC and its single variant G127V were labeled by 5(6)-carboxy-tetramethylrhodamine N-succinimidyl ester (TAMRA, red fluorescence) and liquid droplets of 50 µM PrPC in PBS (red) were observed by differential interference contrast (DIC) confocal microscopy with excitation at 546 nm (Figs. 1 and 2). Unexpectedly, both wild-type PrPC and G127V PrPC formed liquid droplets in PBS buffer (on ice) at an early stage of phase separation (30 min) (Fig. 1A-F). However, compared to the liquid droplets formed by wild-type PrPC in PBS buffer (on ice), which were able to fuse to one another to increase the size from ~5 µm (30 min) to 10-20 µm (72 h) (Figs. 1A-C and 2A-C), droplets formed with G127V PrPC or G127I PrPC did not fuse, and interestingly, the preformed droplets reverted back to a homogenous well-mixed

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solution upon prolonged incubation (71.5 h) (Figs. 1D-I and 2D-I), suggesting that the capability of PrPC to undergo liquid-liquid phase separation is modulated, but not inhibited by G127V and G127I mutations. All protein phase separation experiments were repeated three times, and the results were reproducible (Figs. 2 and S1). We obtained a series of PrP mutants with different substitutions at position 127 and found that G127A PrPC, G127E PrPC, G127K PrPC and G127W PrPC also form liquid droplets in PBS buffer (on ice) (Fig. S2). Furthermore, control experiments showed that wild-type PrPC, G127V PrPC and G127I PrPC did not form liquid droplets in water (on ice) for 30 min and that 50 µM PrPC in water did not exhibit phase separation (Fig. S3). Collectively, these data demonstrate that two neutralizing mutations, G127V and G127I, modulate the liquid-liquid phase separation of PrPC. We then used fluorescence recovery after photobleaching (FRAP) to analyze the dynamics of PrPC molecules within the liquid droplets of wild-type PrPC (Fig. 3A-F), G127V PrPC (Fig. 3G-L) or G127I PrPC (Fig. 3M-R), formed in PBS buffer containing 10% Ficoll 70 (on ice). Ficoll 70, a crowding agent, was used to enhance the stability of PrPC phase separated droplets formed in a short time, in favor of FRAP measurements. Immobile PrPC molecule fraction in the droplets was evaluated using the final relative change in fluorescence intensity of PrPC in liquids after photobleaching. Fluorescence intensity changes in a bleached region over time were normalized against the first time point before photobleaching. After photobleaching of wild-type PrPC droplets, a 27% recovery of the PrPC fluorescence was observed when the fluorescence recovery time was 120 s (Fig. 3A-F), and an about 36% immobile

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wild-type PrPC molecule fraction in the droplets was also observed. After photobleaching of G127V droplets or G127I droplets, however, an 85% recovery or a complete recovery of the PrPC fluorescence was observed when the fluorescence recovery time was 120 s (Fig. 3G-R), and an ~4% immobile G127V molecule fraction or ~0% immobile G127I molecule fraction in the droplets was also observed. The photobleaching percentages of droplets formed with wild-type PrPC, G127V PrPC and G127I PrPC were 51%, 34% and 52%, respectively, suggesting that the difference could be in the recovery rather than the initial impact (Fig. 3). These data demonstrate that two neutralizing mutations, G127V and G127I, modulate the dynamics of PrPC molecules within PrPC phase separated droplets and significantly inhibit the phase transition of PrPC from liquid droplets to solid (potentially fibrillar) material.

Neutralizing mutations significantly inhibit the fibrillization of PrP To further understand why a natural protective polymorphism in PrP, G127V, is refractory to prion disease, we constructed six human PrP variants with mutations at position 127 and investigated their amyloid fibril formation with different cell-free approaches (Fig. 4). We utilized thioflavin T (ThT), a fluorescent dye that specifically binds

to

the

β-sheet

conformation

of

fibrils

[22,23,32,37-39];

8-anilino-1-naphthalene-sulfonic acid (ANS), a fluorescent dye that specifically binds to the solvent-exposed hydrophobic clusters of fibrils [22]; and SDS-PAGE of Sarkosyl-soluble PrP [23] to study the kinetics of fibril formation. For wild-type human PrP and its variants G127V, G127I, G127A, G127E, G127K and G127W, we

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determined time-dependent ThT fluorescence (Fig. 4A), Sarkosyl-soluble fractions and ANS fluorescence (Fig. 4C-H). The lag time of G127V fibrillization (Fig. 4B: 21.35 ± 0.68 h, p = 0.0000080; Fig. 4C: over 22 h; Fig. 4H: 18.99 h) was significantly greater than that of wild-type PrP (Fig. 4B: 6.46 ± 0.55 h; Fig. 4C: over 8 h; Fig. 4H: 7.06 h), confirming that the G127V mutation delays the kinetics of fibril formation. Furthermore, the G127I mutation had protective effects similar to those of G127V (Fig. 4B: 31.57 ± 2.41 h, p = 0.000061; Fig. 4D: over 42 h; Fig. 4H: 38.49 h). Compared with the fibril formation lag time in wild-type PrP, however, the lag time of fibril formation was increased to some extent in G127W PrP (Fig. 4B: 9.72 ± 0.72 h, p = 0.0034; Fig. 4D: over 11 h) and significantly decreased in G127K PrP (Fig. 4B: 2.98 ± 0.14 h, p = 0.00045; Fig. 4D: over 6 h). Furthermore, the fibril formation lag time in G127A (Fig. 4B: 8.69 ± 1.21 h, p = 0.044; Fig. 4D: over 14 h) or G127E (Fig. 4B: 4.10 ± 1.00 h, p = 0.023; Fig. 4D: over 8 h) PrP was not significantly different from that in wild-type PrP. Collectively, these data demonstrate that the hydrophobic chains introduced in G127V and G127I significantly inhibit the fibrillization of PrP. Fibril formation of wild-type, G127V and G127I PrPs was confirmed by our transmission electron microscopy (TEM) experiments (Fig. S4). After fibril formation of wild-type, G127V, G127I, G127A, G127E, G127K and G127W PrPs, the samples were ultracentrifuged. The protein bands in the supernatant after centrifugation were not observed by SDS-PAGE (Fig. S5), indicating that there is no soluble protein left after the formation of fibrils. We then employed far-UV circular dichroism (CD) and Fourier transform infrared

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(FTIR) spectroscopy [23,40,41] to analyze the effect of different mutations on the secondary structure of PrP. Effect of different mutations on the tertiary structure of PrP was analyzed using near-UV CD spectroscopy. Amino acid substitutions at position 127 did not change the secondary structure of PrPC at its native state (Fig. S6A). A negative peak at 208 nm and a negative peak at 222 nm were all observed in wild-type PrPC, G127V PrPC, G127I PrPC, G127A PrPC, G127E PrPC, G127K PrPC and G127W PrPC, indicative of a predominant α-helical structure (Fig. S6A). A negative peak at ~275 nm was all observed in wild-type PrPC and its six mutants, and the shapes of the near-UV CD spectra of these seven proteins indicate that the tertiary structure of PrPC in the native state is not significantly altered by amino acid substitutions at position 127 (Fig. S6B). A negative peak at ~218 nm was all observed in fibrils formed by wild-type PrP and its six variants, and the shapes of the far-UV CD spectra of fibrils formed by these seven proteins are indicative of β-sheet rich structures (Figs. 4I and S7A). The most prominent difference between the far-UV CD spectra of fibrils formed by wild-type PrP and those of G127V and G127I mutants is the signal intensity (Fig. 4I), which could be due to the difference in fibril clamping. Compared to the FTIR spectrum of wild-type PrP fibrils, which had an amide I’ peak at approximately 1630 cm-1 showing a high β-sheet structure, the FTIR spectra of G127V fibrils and G127I fibrils had an amide I’ peak at approximately 1645 cm-1 (Fig. 4J). However, there is no significant difference between the far-UV CD and FTIR spectra of fibrils formed by wild-type PrP and those of G127A, G127E, G127K and G127W mutants (Fig. S7). Collectively, these data demonstrate that two neutralizing

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mutations, G127V and G127I, have effects on the far-UV CD and FTIR spectra of fibrillar forms of PrP. To check whether the conformational stability of human PrPC is affected by mutations, we conducted experiments on urea-induced unfolding of this protein at pH 5.0. Effect of different mutations on the conformational stability of PrPC represented by its unfolded fraction ([θ]222) during urea unfolding was analyzed using far-UV CD spectroscopy. For wild-type human PrPC and its six variants with mutations at position 127, we determined urea-dependent unfolded fraction of PrPC (Fig. S8A). The [Urea]1/2 values of G127V PrPC (3.65 ± 0.31 M, p = 0.081), G127I PrPC (3.70 ± 0.23 M, p = 0.034), G127A PrPC (3.45 ± 0.29 M, p = 0.24), G127E PrPC (3.42 ± 0.25 M, p = 0.25), G127K PrPC (2.97 ± 0.36 M, p = 0.42) and G127W PrPC (3.84 ± 0.14 M, p = 0.0074) were not significantly different from that of wild-type PrPC (3.19 ± 0.18 M), indicating that the G127V, G127I, G127A, G127E, G127K and G127W mutations do not significantly affect the conformational stability of human PrPC (Fig. S8B). Here, [Urea]1/2 is corresponding to the urea concentration required to unfold half of PrPC. Similarly, the G126V mutation does not affect the conformational stability of mouse PrPC [32].

Neutralizing mutations have significantly weaker aggregation ability in cells than wild-type PrP To compare the amount of PrP fibrils in G127V PrP-expressing cells or G127I PrP-expressing cells with those in wild-type PrP-expressing cells, we studied the

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aggregation of wild-type, G127V and G127I PrPs in RK13 cells using Western blotting. The RK13 cell line was chosen because it does not express endogenous PrP, thus avoiding interference by wild-type endogenous protein [42]. RK13 cells transiently expressing wild-type, G127V and G127I PrPs were cultured for 2 days. The normalized amount of insoluble PrP aggregates in those cells (Fig. 5B) was determined as a ratio of the density of insoluble PrP aggregate bands over the total density of all PrP bands in cell lysates (Figs. 5A and S9) after sample ultracentrifugation. The densities of the 35-kDa and 25-kDa bands of wild-type PrP in the pellets were much greater than those of G127V (Figs. 5A and S9), and G127V formed a significantly lower level of Sarkosyl-insoluble aggregates in cells than did wild-type PrP (0.843 ± 0.155 for G127V versus 1.000 ± 0.196 for wild-type PrP; p = 0.023) (Fig. 5B). We suspected that the hydrophobic chain of Val-127 might have blocked the intrinsic ability of PrP to form Sarkosyl-insoluble aggregates in cells. To test this hypothesis, we introduced another amino acid (Ile) with a different hydrophobic chain at the same position and found G127I to have a similar protective effect as G127V (Figs. 5A, 5B and S9). G127I also formed a significantly lower level of Sarkosyl-insoluble aggregates in cells than did wild-type PrP (0.610 ± 0.220 for G127I versus 1.000 ± 0.196 for wild-type PrP; p = 0.040) (Fig. 5B). Therefore, the two neutralizing mutations, G127V and G127I, provide PrP significantly weaker aggregation ability in cells than the wild-type form. To investigate how neutralizing mutations inhibit the aggregation of PrP in cells, we analyzed the effects of such mutants on the subcellular localization of PrP. RK13

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cells transiently expressing wild-type, G127V and G127I PrP, cultured for a half day and then incubated with PBS buffer and 50 µM PrP 106-126 for 1.5 days, were immunostained with the anti-PrP antibody 8H4 and with IgG conjugated to Alexa Fluor 555 (red), stained with DAPI (blue), and observed by confocal microscopy (Figs. S10 and 5C-N). Wild-type, G127V and G127I PrPs in the native state were all mainly attached to the plasma membrane (Fig. S10), indicating that amino acid substitutions at position 127 do not impair the localization of PrP at the cell surface. However, when wild-type, G127V and G127I PrPs were treated with PrP 106-126, the two neutralizing mutations displayed much weaker aggregation ability in cells (Fig. 5G-N) than did wild-type PrP, and PrP 106-126 caused translocation of wild-type PrPC but not G127V PrPC or G127I PrPC from the membrane to the cytoplasm (Fig. 5C-N). Wild-type PrP formed fibrils (red spots in Fig. 5D and 5E) in the cytoplasm of wild-type PrP-expressing cells treated with PrP 106-126. Collectively, these data suggest that the hydrophobic chains introduced in G127V and G127I inhibit the aggregation of PrP in cells by preventing the redistribution of PrPC from the cell surface to the cytoplasm.

Neutralizing mutations protect against mitochondrial damage in cells caused by PrP aggregation To address the question of the relationship between PrP aggregation and mitochondrial damage, we utilized TEM [43,44] to study the influences of transiently expressed wild-type, G127V and G127I PrP on mitochondrial damage in RK13 cells

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incubated with 50 µM PrP 106-126 (Fig. 6). The normal morphology of mitochondria in cells expressing wild-type, G127V and G127I PrP not treated with PrP 106-126 (Fig. 6A and 6B) or in cells transiently expressing G127V or G127I incubated with the peptide (Fig. 6C and 6D), highlighted by white arrows, was tubular or round. PrP 106-126 treatment caused severe mitochondrial impairment in cells expressing wild-type PrP, and nearly all the mitochondria in the cells became either vacuolized with broken mitochondrial cristae, which is highlighted by yellow arrows, or swollen, which is highlighted by blue arrows; these processes were accompanied by the accumulation of lysosomes (highlighted by green arrows) in the cells (Fig. 6D). Autolysosomes, highlighted by red arrows, were also observed in the cells (Fig. 6D). In contrast, PrP 106-126 treatment did not cause severe mitochondrial impairment in G127V PrP-expressing cells or G127I PrP-expressing cells; only a few mitochondria in the cells became vacuolized with broken mitochondrial cristae, and lysosomes and autolysosomes were not observed (Fig. 6C and 6D). As controls, RK13 cells were also incubated with PBS buffer (Fig. S11A and S11B) or treated with 50 µM PrP 106-126 (Fig. S11C and S11D). PrP 106-126 did not cause any mitochondrial impairment in cells not expressing PrP (Fig. S11C and S11D). Collectively, these data demonstrate that PrP 106-126 causes much more severe mitochondrial impairment in cells expressing wild-type PrP than in cells expressing G127V PrP or G127I PrP. Therefore, the two neutralizing mutations, G127V and G127I, protect against mitochondrial damage in cells caused by PrP aggregation and enhanced by PrP 106-126.

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Neutralizing mutations significantly decrease PrP aggregation-induced ROS production and apoptosis in cells To examine the role of PrP aggregation in cellular ROS production, flow cytometry and the ROS probe 2’,7’-dichlorofluorescin diacetate (DCFH-DA) [42] were used to determine ROS levels in cells transiently expressing PrP. SH-SY5Y and RK13 cells expressing wild-type (Figs. 7 and S12, A and D), G127V (Figs. 7 and S12, B and E) and G127I (Figs. 7 and S12, C and F) PrPs were cultured for a half day and then incubated with PBS buffer (Figs. 7 and S12, A-C) or 50 µM PrP 106-126 (Figs. 8 and S12, D-F) for 1.5 days. The ROS levels caused by PrP aggregation with PrP 106-126 in SH-SY5Y cells (Fig. 7D) and RK13 cells (Fig. S12D) expressing wild-type PrP were 4.84% and 3.87%, respectively, which were markedly greater than those in SH-SY5Y and RK13 cells not treated with the peptide and transiently expressing wild-type PrP (Figs. 7A and S12A, 0.29% and 0.59%, respectively). In contrast, PrP 106-126-treated and untreated SH-SY5Y and RK13 cells transiently expressing G127V or G127I showed similar low ROS levels (Figs. 7 and S12, B and E or C and F). As controls, SH-SY5Y cells (Fig. S13A and S13B) and RK13 cells (Fig. S13C and S13D) treated with PrP 106-126 (Fig. S13B and S13D) and those cells not treated with the peptide (Fig. S13A and S13C) displayed similar low ROS levels, indicative of PrP 106-126 having no clear effect on ROS levels in cells not expressing PrP. To examine the role of PrP aggregation in cytotoxicity, flow cytometry with annexin V-FITC and propidium iodide (PI) staining [8,10,42,45-47] was used to study the influences of transiently expressed wild-type, G127V and G127I PrP on toxicity in

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living SH-SY5Y cells incubated with 50 µM PrP 106-126 (Fig. 7G-L). The percentage of early apoptotic cells among living cells treated with the peptide for 36 h and expressing wild-type PrP was 11.01% (Fig. 7J), which was significantly greater than that in cells not treated with the peptide and expressing wild-type PrP (Fig. 7G, 0.49%). In contrast, PrP 106-126-treated and untreated SH-SY5Y cells transiently expressing G127V or G127I showed similar low rates of early apoptosis (Fig. 7K and 7H, 2.00% and 0.68% respectively, or Fig. 7L and 7I, 1.68% and 0.52% respectively). As controls, cultures of SH-SY5Y cells treated with 50 µM PrP 106-126 for 36 h (Fig. S14B, 2.67%) only contained a mildly higher percentage of early apoptotic cells than cultures that were not treated with the peptide (Fig. S14A, 0.65%). Interestingly, when treated with PrP 106-126, wild-type PrP-expressing cells, G127V PrP-expressing cells, G127I PrP-expressing cells and cells not overexpressing PrP all showed similar rates of late apoptosis (Figs. 7J-L and S14B). We can conclude that PrP 106-126 causes much more severe ROS production and early apoptosis in wild-type PrP-expressing cells than in G127V PrP-expressing cells or G127I PrP-expressing cells. Therefore, the two neutralizing mutations, G127V and G127I, significantly decrease ROS production and apoptosis in cells caused by PrP aggregation enhanced by the peptide.

Discussion Prion disease is a type of infectious fatal neurodegenerative disease primarily caused by the misfolding of PrPs in humans, cattle, sheep, and cervid species [2-6].

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The conversion from PrPC into PrPSc is an important step in the initiation of prion disease [1,4,12,13]. A naturally occurring protective polymorphism in human PrP, G127V, has been recently reported to cause positive evolutionary selection by preventing PrP fibril formation and greatly attenuating prion disease [28-32], but the mechanism behind this phenomenon is largely unknown. The roles of the Met-129 or Val-129 polymorphism in human PrP are well documented regarding the susceptibility of prion disease [48]; the Gly-127 or Val-127 polymorphism is always associated with the Met-129 or Val-129 polymorphism, but Val-127 is seen exclusively on Met-129 PRNP alleles [28,29]. In the present study, we focused on a natural protective polymorphism in PrP, G127V, and one novel protective mutant G127I, and reported a biophysical characterization of PrP variants that highlight the protective effect of the G127V polymorphism. We showed that the liquid-liquid phase separation of PrPC occurs in G, V and I derivatives, but is reversed in V and I upon prolonged incubation. Our FRAP data indicated that the droplets of G derivative contain significant amount of non-movable (solid, potentially fibrillar) material, while V and I droplets do not contain significant amount of solid material. We observed a significantly lower amount of insoluble G127V PrP fibrils than wild-type PrP fibrils 2 days after transient expression of the proteins, consistent with the delayed kinetics of G127V PrP fibril formation. We demonstrated that the hydrophobic chain introduced in G127V modulates the in vitro phase separation and significantly inhibits amyloid fibril formation by human PrP. Data are also included showing that the G127I mutation has regulatory and protective effects similar to those of G127V. Overall, our

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data indicated that the liquid-liquid phase separation of PrPC per se is not impaired by V or I residue at position 127. However, the phase separation of PrPC is modulated and the subsequent formation of PrP fibrils is impaired by G127V or G127I. A defect in fibril formation could antagonize the fusion of droplets and result in their eventual dissolution. Thus, it is possible that the impact of G127V and G127I on the phase separation of PrPC could be a consequence of the effect on fibrillization. Different from the previously reported data in which the G127V mutation is able to slow down the fibrillization rate of PrP [30-32], we herein investigate the liquid-liquid phase separation and the subsequent fibril formation of wild-type PrP and its two neutralizing mutations, G127V and G127I, as well as their impacts on mitochondrial damage, ROS production and apoptosis in cells. The following findings of our study have advanced the field of the phase separation and fibril formation of proteins in neurodegenerative diseases [9,33,34,49]. First, we found that two neutralizing mutations, G127V and G127I, modulated the liquid-liquid phase separation of PrPC and significantly inhibited the subsequent fibrillization of PrP (Fig. 8). This finding is supported by our demonstration that a single mutation at position 127 in PrPC (from Gly-127 to Val-127 or Ile-127) resulted in solvent-exposed hydrophobic clusters that modulated the liquid-liquid phase separation of PrPC and significantly inhibited the formation of PrP fibrils. Second, we revealed that PrP with the two neutralizing mutations had significantly weaker aggregation ability in cells than wild-type PrP and protected against mitochondrial damage in cells caused by PrP aggregation (Fig. 8). The hydrophobic chains introduced in G127V and G127I sequestered PrPC at the cell

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surface and thus inhibited the aggregation of PrP in cells by preventing the translocation of PrPC from the membrane to cytoplasm. Third, we observed that the two neutralizing mutations, G127V and G127I, significantly decreased not only PrP aggregation-induced ROS production but also PrP aggregation-induced apoptosis (Fig. 8). PrP aggregation under pathological conditions causes mitochondrial damage in cells. Because mitochondria are the major organelle for producing oxidative signals, mitochondrial dysfunction leads to elevated ROS production, and aggravated oxidative stress further enhances PrP aggregation in cells and causes cytotoxicity (Fig. 8). Our study enhances our understanding of how a natural protective polymorphism in PrP regulates the phase separation, amyloid formation and cytotoxicity of PrP and helps explain the mechanism underlying the very interesting protective effect of this natural polymorphism in PrP against infection with prion disease observed in humans and animal models.

Materials and Methods Materials Two dyes, ThT and DAPI, were purchased from Sigma-Aldrich (St. Louis, MO). The mouse anti-PrP monoclonal antibodies 3F4 and 8H4 were obtained from BioLegend (San Diego, CA) and Abcam (Cambridge, UK), respectively. Alexa 555-conjugated fluorescent secondary antibody and the ROS probe DCFH-DA (2’,7’-dichlorofluorescein diacetate) were purchased from Beyotime (Nantong, China). Guanidine hydrochloride (GdnHCl) and acetonitrile were obtained from Promega

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(Madison, WI) and Fisher Scientific (Fairlawn, NJ), respectively. Sarkosyl was purchased from Amresco (Solon, OH). Ni-Sepharose was purchased from GE Company (Pittsburgh, PA). All other chemicals used in this study were of analytical grade and were produced in China. Protein purification A plasmid encoding wild-type human PrP (23-231) was a kind gift from Dr. Geng-Fu Xiao (Wuhan Institute of Virology, Chinese Academy of Sciences). The gene for PrP 23-231 was constructed in the vector pET-30a (+), and PrP mutants G127V, G127I, G127A, G127E, G127K and G127W were constructed by site-directed mutagenesis with a wild-type PrP template and the primers shown in Table S1. All PrP plasmids were transformed into Escherichia coli. Recombinant wild-type PrP and its mutants were expressed in E. coli BL21 (DE3) cells (Novagen, Merck, Darmstadt, Germany) and purified by high-performance liquid chromatography (HPLC) on a C4 reverse-phase column (Shimadzu, Kyoto, Japan) as described by Baskakov [50] and Liang [23,42]. After purification, human PrPs were dialyzed against 1 × PBS buffer (pH 7.0) for 1 day, concentrated, filtered and stored at -80°C. SDS-PAGE and mass spectrometry were used to confirm that all the purified human PrPs were single species with an intact disulfide bond. We used a NanoDrop OneC Microvolume UV-Vis Spectrophotometer (Thermo Scientific, Waltham, MA) to determine the concentrations of wild-type PrPC and its mutants using their absorbance at 280 nm and molar extinction coefficients calculated from the composition of the protein (http://web.expasy.org/protparam/).

19

Liquid-droplet formation The freshly purified wild-type PrPC, G127V PrPC, G127I PrPC, G127A PrPC, G127E PrPC, G127K PrPC and G127W PrPC were incubated with TAMRA (red fluorescence) at a PrPC:TAMRA molar ratio of 1:3 for 1 h. These proteins labeled by TAMRA were filtered and concentrated to 100 µM and then diluted 1:1 with 2 × PBS buffer to a final concentration of 50 µM on ice to induce liquid-liquid phase separation [35] for 30 min or 72 h. Images of 10 µl samples were captured using a Leica TCS SP8 laser scanning confocal microscope (Wetzlar, Germany), and liquid droplets of 50 µM PrPC in PBS were observed by DIC confocal microscopy with excitation at 546 nm. Wild-type PrPC, G127V PrPC and G127I PrPC diluted with doubly distilled water to a final concentration of 50 µM on ice for 30 min were used as controls. No phase separation for 50 µM PrPC in water was observed by DIC confocal microscopy with excitation at 546 nm. Fluorescence recovery after photobleaching (FRAP) Wild-type PrPC, G127V PrPC and G127I PrPC, labeled by TAMRA, were filtered and concentrated to 100 µM and then diluted 1:0.2:0.8 with 10 × PBS buffer and 25% Ficoll 70 to a final concentration of 50 µM on ice to induce liquid-liquid phase separation in the presence of 10% Ficoll 70 for 30 min. FRAP measurements were performed on a Leica TCS SP8 laser scanning confocal microscope (Wetzlar, Germany) with excitation at 546 nm. Droplets of a size of ~10 µm were selected. For each droplet, a square (~2.5 µm × 2.5 µm) was bleached at 80% transmission for 20 flashes (1.3 s per flash), and postbleach time-lapse images were collected (40 frames,

20

3.8 s per frame). Images were analyzed using Leica LAS AF Lite. Fibril formation To form fibrils from wild-type PrP and its mutants in denaturing conditions, we used the protocols described by us previously [23]. Briefly, wild-type PrP and its variants were dialyzed against 1 × PBS buffer (pH 7.0) containing 2 M guanidine hydrochloride (GdnHCl) and diluted to a final concentration of 10 µM. Samples were then incubated with shaking at 220 rpm at 37°C. ThT binding assays Wild-type PrP and its single variants were incubated in PBS buffer (pH 7.0) containing 2 M GdnHCl with shaking at 220 rpm and 37°C for 15-52 h and analyzed by ThT binding assays. The final concentrations of human PrP and ThT were 1 and 125 µM, respectively. A Cytation 3 Cell Imaging Multi-Mode Reader (BioTek, Winooski, VT) was employed to measure ThT fluorescence produced with excitation at 450 nm and emitted at 480 nm. The lag time of the fibrillization of wild-type PrP and its mutants was determined by a sigmoidal equation [10,38,39] using the ThT fluorescence data. Revised Student’s t-test was used to perform statistical analyses of the lag time. Values of p < 0.005 were considered to indicate statistically significant differences. The following notation is used throughout: ∗, p < 0.005, ∗∗, p < 0.001, and ∗∗∗, p < 0.0001 relative to wild-type PrP. ANS binding assays Wild-type PrP and its variants were incubated in PBS buffer (pH 7.0) containing 2 M GdnHCl with shaking at 220 rpm and 37°C for 14-60 h and analyzed by ANS

21

binding assays. The final concentrations of human PrP and ANS were 1 and 125 µM, respectively. The fluorescence spectra for ANS binding were recorded between 400 and 650 nm with excitation at 380 nm on the Cytation 3 Cell Imaging Multi-Mode Reader. The lag time of the fibrillization of wild-type PrP and its mutants was determined by a sigmoidal equation using the ANS fluorescence data. SDS-PAGE of Sarkosyl-soluble fractions To distinguish human PrP monomers, which are Sarkosyl soluble, from human PrP fibrils, which are Sarkosyl insoluble, Sarkosyl-soluble SDS-PAGE experiments for fibril formation of wild-type PrP and its single variants were carried out as previously described in detail [23]. Briefly, 10 µM PrP samples were taken at multiple time points during aggregation, incubated with 2% Sarkosyl for half an hour at 25°C, then mixed with 2 × loading buffer (without SDS, β-mercaptoethanol, and heating) and separated by 12.5% SDS-PAGE. The soluble PrP monomers were detected by SDS-PAGE with Coomassie Blue R250 staining. CD spectroscopy CD spectra of wild-type PrP and its single variants were measured using a Jasco J-810 spectropolarimeter (Jasco Corp., Tokyo, Japan). We used a quartz cell with a 1-mm light path to measure far-UV CD spectra recorded between 195 and 250 nm at 25°C, and 10 µM PrP samples were dialyzed against 20 mM NaAc buffer (pH 5.0) for 3 h in advance to ensure that GdnHCl had been removed. We also used a quartz cell with a 10-mm light path to measure near-UV CD spectra of 100 µM PrP samples recorded between 250 and 350 nm at 25°C. The spectra of all scans were corrected

22

relative to the buffer blank, and all experiments were repeated five times. Fourier transform infrared spectroscopy Attenuated total reflection FTIR spectra of 10 µM wild-type PrP and its single mutants during fibril formation were measured using a Nicolet 5700 FTIR spectrophotometer (Thermo Electron, Madison, WI). PrP fibril samples were harvested by ultracentrifugation for 150,000 g for half an hour and washed with H2O. Then the pellets were vacuumized overnight. The dried PrP fibril samples were resuspended in D2O and stored at 4°C. FTIR spectra were recorded between 400 and 4000 cm-1 at 4 cm-1 resolution and at 25°C, and each sample was repeated 32 times. The average FTIR intensity was recorded and corrected for D2O. We used OMNIC 8 software to address the FTIR intensity with 5-point smoothing. TEM of PrP fibrils Human PrP (10 µM) was incubated in PBS buffer (pH 7.0) containing 2 M GdnHCl with shaking at 220 rpm and 37°C. Fibril formation of human PrP was confirmed by electron microscopy of negatively stained samples. The incubation time was chosen within a time range of the plateau of each kinetic curve of ThT fluorescence. PrP fibril samples (10 µl) were loaded on copper grids for 60 s and washed with H2O for 10 s. Samples on grids were then stained with 2% (w/v) uranyl acetate for 50 s and dried in the air at 25°C. The stained samples were examined using a JEM-1400 Plus transmission electron microscope (JEOL, Tokyo, Japan) operating at 100 kV. SDS-PAGE of the supernatant after centrifugation of PrP fibril samples Human PrP (10 µM) was incubated in PBS buffer (pH 7.0) containing 2 M GdnHCl

23

with shaking at 220 rpm and 37°C for 15 h for wild-type PrP, 30 h for G127V, 52 h for G127I, 16 h for G127A, 11 h for G127E, 9 h for G127K and 15 h for G127W. PrP fibril samples were then ultracentrifuged at 150,000 g for half an hour. After centrifugation, the supernatant was collected and separated by 15% SDS-PAGE. The protein bands were detected by SDS-PAGE with Coomassie Blue R250 staining. Wild-type PrPC (10 µM) in water was used as a control. Determining conformational stability of PrPC Here, the conformational stability of PrPC is represented by its unfolded fraction ([θ]222) during urea unfolding. For urea-induced unfolding studies performed at pH 5.0 (in 20 mM acetate buffer), samples (10 µM) of wild-type PrPC and its single mutants were incubated in increasing concentrations of urea (0-8 M) for 2 h at 25°C. The change in the far-UV ellipticity at 222 nm was monitored using the Jasco J-810 spectropolarimeter. [Urea]1/2, corresponding to the urea concentration required to unfold half of PrPC, was determined by fitting the urea-induced unfolding data to a sigmoidal equation. Wild-type PrPC was used as a control. The [Urea]1/2 values are expressed as the mean ± S.D. (with error bars) of values obtained in 3 independent experiments. Revised Student’s t-test was used to perform statistical analyses. Values of p < 0.005 were considered to indicate statistically significant differences. The following notation is used throughout: ∗, p < 0.005, ∗∗, p < 0.001, and ∗∗∗, p < 0.0001 relative to wild-type PrPC. Cell culture and transfection RK13 and SH-SY5Y cells were cultured in minimum essential media and in

24

Dulbecco’s modified Eagle’s medium (Gibco, Invitrogen, Mulgrave, VIC, Australia), respectively, supplemented with 10% (v/v) fetal bovine serum (FBS, Gibco), 100 U/ml streptomycin, and 100 U/ml penicillin in 5% CO2 at 37°C. Wild-type, G127V and G127I PrPs that had been constructed in the pCMV-Tag 2B vector were transiently transfected into RK13 and SH-SY5Y cell lines by Lipofectamine® 2000 (Invitrogen, Carlsbad, CA). Sarkosyl-insoluble Western blotting RK13 cells transiently expressing wild-type, G127V or G127I PrP were cultured for 2 days. Wild-type PrP and its single variants in RK13 cells were harvested and resuspended in lysis buffer (pH 7.6) containing 1% Triton X-100, 50 mM Tris, 150 mM NaCl, 1 mM phenylmethanesulfonyl fluoride, and protease inhibitors on ice for half an hour. The cell lysates were centrifuged at 10,000 g for 10 min. Half of the supernatant was incubated with 1% Sarkosyl for half an hour at 37°C. The mixture was then ultracentrifuged at 150,000 g for half an hour and washed with 1 × PBS buffer (pH 7.4). The Sarkosyl-insoluble pellets were boiled in SDS-PAGE loading buffer for 15 min. The other half of the supernatant, which served as the total protein sample, was also boiled in SDS-PAGE loading buffer for 15 min. Samples were separated by 12.5% SDS-PAGE and then Western blotted as described in detail in the above proteinase K digestion assays. The Sarkosyl-insoluble pellets from those cells were probed with 3F4, and the supernatants from those cells were probed using 3F4 and the anti-β-actin antibody. The amount of loaded protein was normalized using a BCA Protein Quantification kit (Beyotime). To calculate the amount of

25

Sarkosyl-insoluble PrP, ImageJ software (NIH, Bethesda, MD) was used to assess the densitometry of PrP bands. The normalized amount of insoluble PrP aggregates in those cells was calculated as the ratio of the density of insoluble PrP aggregate bands to the total density of all PrP bands in cell lysates. RK13 cells transiently expressing wild-type PrP were used as a control. The normalized amounts of insoluble PrP aggregates are expressed as the mean ± S.D. (with error bars) of values obtained in 3 independent experiments. Statistical analyses were performed using Student’s t-test. Values of p < 0.05 were considered to indicate statistically significant differences. The following notation is used throughout: ∗, p < 0.05, ∗∗, p < 0.01, and ∗∗∗, p < 0.001 relative to wild-type PrP. PrP peptide A toxic prion peptide, PrP 106-126, was synthesized by Beijing Genomics Institute (BGI), Shenzhen, China, dissolved in 1 × PBS buffer (pH 7.4) to a concentration of 1 mM,

and

kept

frozen

until

use.

The

sequence

of

PrP

106-126

is

KTNMKHMAGAAAAGAVVGGLG. Laser scanning confocal analysis RK13 cells transiently expressing wild-type, G127V and G127I PrP were cultured for a half day, incubated with 50 µM PrP 106-126 or 1 × PBS buffer for 1.5 days at 37°C, fixed, permeabilized, immunostained with the mouse monoclonal anti-PrP antibody 8H4 and IgG conjugated to Alexa Fluor 555 (red), and stained with DAPI (blue). Images were captured using an Olympus FluoView FV1000 laser scanning confocal microscope (Tokyo, Japan) as described in detail previously [42].

26

Ultrathin TEM RK13 cells transiently expressing wild-type, G127V and G127I PrP were cultured for a half day and then incubated with 50 µM PrP 106-126 or 1 × PBS buffer for 1.5 days. After prefixation with 3% paraformaldehyde and 1.5% glutaraldehyde in 1 × PBS buffer (pH 7.4), the cells were harvested and postfixed in 1% osmium tetroxide for 1 h with an ice bath; the samples were then dehydrated in graded acetone and embedded in 812 resins. Ultrathin sections of cells were prepared using a Leica Ultracut S Microtome (Buffalo Grove, IL) and negatively stained using 2% uranyl acetate and lead citrate. The doubly stained ultrathin sections of cells were examined using a JEM-1400 Plus transmission electron microscope (JEOL) operating at 80 kV. Oxidative stress detection RK13 and SH-SY5Y cells transiently expressing wild-type, G127V and G127I PrP were cultured for a half day, and then incubated with 50 µM PrP 106-126 or 1 × PBS buffer for 1.5 days. The cellular ROS levels were determined using a ROS assay kit (Beyotime, Nantong, China) according to the manufacturer’s protocol. Briefly, the cells were washed twice with 1 × PBS buffer and incubated with 5 µM DCFH-DA for 20 min at 37°C. The percentage of the total number of cells (~10,000) containing ROS was determined on an EPICS XL-MCL flow cytometer (Beckman Coulter) using EXPO32 MultiComp software (Beckman Coulter) as described in detail previously [42]. The RK-13 and SH-SY5Y cell lines transiently expressing wild-type PrP were used as controls. All ROS detection experiments were repeated three times. Annexin V-FITC apoptosis detection assay

27

SH-SY5Y cells transiently expressing wild-type, G127V and G127I PrP were cultured for a half day and then incubated with 50 µM PrP 106-126 or 1 × PBS buffer for 1.5 days. Apoptotic cells were detected by flow cytometry after staining with an annexin V-FITC apoptosis detection kit (Beyotime). Briefly, the cells were harvested after digestion with 2.5 mg/ml trypsin (Promega, Madison, WI), washed with 1 × PBS buffer at 4°C, and resuspended in 185 µl of binding buffer. The samples were then incubated with 5 µl of annexin V-FITC and 10 µl of PI for 10 min at 4°C in the dark. Annexin V binding was analyzed using an EPICS XL-MCL flow cytometer (Beckman Coulter, Fullerton, CA), and the percentage of apoptotic cells was calculated from the total number of cells (~3 × 104 cells) as described in detail previously [42]. All apoptotic blot experiments were repeated three times. Statistical analysis The data shown for each experiment were based on at least three technical replicates, as indicated in individual figure legends. Data are presented as the mean ± S.D., and p-values were determined by Revised Student’s t-test or Student’s t-test. All experiments were further confirmed by biological repeats.

Acknowledgments We sincerely thank Dr. Geng-Fu Xiao (Wuhan Institute of Virology, Chinese Academy of Sciences) for the kind gift of the human PrPC plasmid and Dr. Xiang-Dong Fu (University of California, San Diego) for insightful discussions and critical comments throughout the investigation. We thank Dr. Zhiyin Song for his

28

technical assistance with the TEM of ultrathin sections of cells. This work was supported by the National Natural Science Foundation of China Grants 31370774, 31770833, and 31570779, the National Key Basic Research Foundation of China Grants 2013CB910702 and 2012CB911003, and the Fundamental Research Fund for the Central Universities of China 2015204020201.

Author Contributions: J.-J.H. X.-N.L. and Y.L. designed the experiments. J.-J.H. X.-N.L., W.-L.L., H.-Y.Y., Y.G., K.W., B.T. and Y.L. performed the experiments. D.-W.P. and J.C. contributed the new reagents. All authors were involved in data interpretation and discussion. J.-J.H. and Y.L. wrote the manuscript with contributions from all other authors.

Conflict of Interest Statement: The authors declare no competing financial interests.

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Figure legends

Fig. 1. Wild-type PrPC, G127V PrPC and G127I PrPC all form liquid droplets in PBS buffer at the early stage of phase separation. (A-I) Wild-type PrPC (A-C), G127V PrPC (D-F) and G127I PrPC (G-I) were diluted 1:1 with 2 × PBS buffer to a final concentration of 50 µM on ice to induce liquid-liquid phase separation for 30 min. Wild-type PrPC, G127V PrPC and G127I PrPC were labeled by TAMRA (red fluorescence), and liquid droplets of 50 µM PrPC in PBS (red) were observed by DIC confocal microscopy with excitation at 546 nm. The scale bars represent 25 µm.

Fig. 2. G127V and G127I modulate liquid-liquid phase separation of human PrPC. (A-I) Wild-type PrPC (A-C), G127V PrPC (D-F) and G127I PrPC (G-I) were diluted 1:1 with 2 × PBS buffer to a final concentration of 50 µM on ice to induce liquid-liquid phase separation for 72 h. Wild-type PrPC, G127V PrPC and G127I PrPC were labeled by TAMRA (red fluorescence), and liquid droplets of 50 µM PrPC (red) in PBS were observed by DIC confocal microscopy with excitation at 546 nm. The scale bars represent 25 µm.

Fig. 3. G127V and G127I remarkably accelerate fluorescence recovery after photobleaching of liquid droplets and significantly inhibit the phase transition of human PrPC. (A-R) Time course of FRAP after internal photobleaching of liquid droplets of wild-type PrPC (A-F), G127V PrPC (G-L) and G127I PrPC (M-R). The

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internal photobleaching is marked by a black square. Pre-bleach represents the time before photobleaching (A, G and M), and time 0 indicates that for photobleaching (B, H and N). The fluorescence recovery time is indicated on top of the corresponding recovery image (C-F, I-L, and O-R). Wild-type PrPC (A-C), G127V PrPC (D-F) and G127I PrPC (G-I) were diluted 1:0.2:0.8 with 10 × PBS buffer and 25% Ficoll 70 to a final concentration of 50 µM on ice to induce liquid-liquid phase separation in the presence of 10% Ficoll 70 for 30 min. Wild-type PrPC, G127V PrPC and G127I PrPC were labeled by TAMRA (red fluorescence), and liquid droplets of 50 µM PrPC (red) in PBS and 10% Ficoll 70 were observed by confocal microscopy with excitation at 546 nm. The scale bars represent 2.5 µm.

Fig. 4. G127V and G127I significantly inhibit amyloid formation of human PrP. (A) Samples (10 µM) of wild-type PrP (open square) and its single mutants G127V (open circle), G127I (solid circle), G127A (solid square), G127E (open up triangle), G127K (open down triangle) and G127W (solid up triangle) were incubated in PBS buffer (pH 7.0) containing 2 M GdnHCl with shaking at 220 rpm and 37°C for 15, 30, 52, 16, 11, 9 and 15 h, respectively, and analyzed by ThT binding assays. The solid lines show the best sigmoidal fit for the ThT intensity-time curves. (B) The lag time of fibril formation of wild-type PrP and its six variants with mutations at position 127 was determined by a sigmoidal equation using the data from (A). The ThT fluorescence intensity (A) and lag time (B) are expressed as the mean ± S.D. of the values obtained in 3 independent experiments. G127V, p = 0.0000080; G127I, p =

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0.000061. Revised Student’s t-test was used to perform statistical analyses. Values of p < 0.005 were considered to indicate statistically significant differences. The following notation is used throughout: ∗, p < 0.005, ∗∗, p < 0.001, and ∗∗∗, p < 0.0001 relative to wild-type PrP (a control). (C and D) SDS-PAGE analysis of time-dependent Sarkosyl-soluble PrP, including wild-type PrP and its mutations G127V (C), G127I, G127A, G127K, G127E and G127W (D). Briefly, 10-µM PrP samples were incubated with 2% Sarkosyl and separated by 12.5% SDS-PAGE. The soluble PrP monomers were detected by SDS-PAGE with Coomassie Blue R250 staining. (E-G) ANS fluorescence spectra of wild-type PrP (E) under denaturing conditions for 0 (black), 6 (red), 8 (green), 10 (blue) and 12 (magenta) h are shown, and those of G127V (F) and G127I (G) PrPs under denaturing conditions for 0 (black), 12 (red), 20 (green), 28 (blue) and 36 (magenta) h are also shown. (H) Samples (10 µM) of wild-type PrP (open square) and its single mutants G127V (open circle) and G127I (open triangle) were incubated in PBS buffer (pH 7.0) containing 2 M GdnHCl with shaking at 220 rpm and 37°C for 14, 30 and 60 h and analyzed by ANS binding assays. The solid lines show the best sigmoidal fit for the ANS intensity-time curves. (I and J) The secondary structure of amyloid fibrils produced from 10 µM wild-type (black), G127V (red) and G127I (blue) PrPs under denaturing conditions for 12, 30 and 48 h, respectively, monitored by far-UV CD (I) and FTIR (J).

Fig. 5. G127V and G127I PrP have significantly weaker aggregation ability in cells than wild-type PrP. (A) The Sarkosyl-insoluble pellets from RK13 cells transiently

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expressing wild-type, G127V or G127I PrP were probed with 3F4, and the supernatants from those cells were probed using 3F4 and the anti-β-actin antibody. RK13 cells transiently expressing wild-type, G127V or G127I prion proteins were cultured for 2 days. (B) The normalized amount of insoluble PrP aggregates in RK13 cells transiently expressing wild-type, G127V or G127I PrP was determined as a ratio of the density of insoluble PrP aggregate bands over the total density of all PrP bands in cell lysates. RK13 cells transiently expressing wild-type PrP were used as a control. The normalized amounts of insoluble PrP aggregates are expressed as the mean ± S.D. (with error bars) of values obtained in 3 independent experiments. G127V, p = 0.023; G127I, p = 0.040. Statistical analyses were performed using Student’s t-test. Values of p < 0.05 were considered to indicate statistically significant differences. The following notation is used throughout: ∗, p < 0.05, ∗∗, p < 0.01, and ∗∗∗, p < 0.001 relative to wild-type PrP (a control). (C-N) G127V and G127I, which are mainly attached to the plasma membrane, display much weaker aggregation ability in cells than wild-type PrP, which is mainly located in the cytoplasm. Wild-type, G127V or G127I PrPs were treated with 50 µM PrP 106-126. RK13 cells transiently expressing wild-type (C-F), G127V (G-J) and G127I (K-N) PrP were cultured for a half day, then with 50 µM PrP 106-126 for 1.5 days at 37°C, fixed, permeabilized, immunostained with the anti-PrP antibody 8H4 and IgG conjugated to Alexa Fluor 555 (red), stained with DAPI (blue), and observed by confocal microscopy. The scale bars represent 25 µm.

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Fig. 6. TEM images show that G127V and G127I protect against mitochondrial damage in RK13 cells caused by PrP aggregation and are enhanced by PrP 106-126. (A-D) RK13 cells transiently expressing wild-type, G127V and G127I PrPs were cultured for a half day and then incubated with PBS buffer (A and B) or 50 µM PrP 106-126 (C and D) for 1.5 days. The enlarged regions B and D show twenty-five-fold enlarged images from A and C, respectively, and display the detailed structures of mitochondria in RK13 cells. Nuclei are highlighted using black arrows (A and C). The normal morphology of mitochondria in RK13 cells incubated with PBS buffer (B) or in RK13 cells expressing G127V and G127I PrPs incubated with 50 µM PrP 106-126 (D), which are highlighted by using white arrows, was tubular or round. PrP 106-126 treatment caused severe mitochondrial impairment in RK13 cells expressing wild-type PrP (D). Nearly all the mitochondria in the cells became vacuolized, and the mitochondrial cristae were broken, which is highlighted by yellow arrows, or became swollen, which is highlighted by blue arrows; these processes were accompanied by the accumulation of lysosomes (highlighted by green arrows) in the cells. Autolysosomes, highlighted by red arrows, were also observed in the cells. In contrast, PrP 106-126 treatment did not cause severe mitochondrial impairment in RK13 cells expressing PrPG127V and PrPG127I (D), and only a few mitochondria in the cells became vacuolized with broken mitochondrial cristae (highlighted by yellow arrows). Samples were negatively stained using 2% uranyl acetate and lead citrate. The scale bars represent 5 µm for A and C and 1 µm for B and D.

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Fig. 7. G127V and G127I significantly decrease PrP 106-126-enhanced PrP cytotoxicity resulting from PrP fibril formation and elevated ROS production. (A-L) SH-SY5Y cells transiently expressing wild-type (A, D, G and J), G127V (B, E, H and K) and G127I (C, F, I and L) PrP were cultured for a half day and then incubated with PBS buffer (A-C and G-I) or 50 µM PrP 106-126 (D-F and J-L) for 1.5 days. The percentage of cells with ROS was determined by flow cytometry using the ROS probe DCFH-DA. SH-SY5Y cells transiently expressing wild-type PrP were used as a control. The percentage of apoptotic cells was also determined by flow cytometry. The four quadrants distinguished by annexin V-FITC/PI staining represent viable cells (R4 quadrant), early apoptotic cells (R5 quadrant), late apoptotic cells (R3 quadrant) and operation-damaged cells (R2 quadrant).

Fig. 8. A hypothetical model shows how G127V and G127I inhibit amyloid fibril formation of PrP and protect against mitochondrial damage, elevated ROS production and PrP cytotoxicity. Mitochondrial damage, elevated ROS production and PrP cytotoxicity are key functional consequences of the aggregation of human PrP. G127V and G127I modulate liquid-liquid phase separation of PrPC, significantly inhibit amyloid fibril formation of PrP and decrease its cytotoxicity. PrP aggregation under pathological conditions causes mitochondrial damage in cells, and mitochondrial dysfunction leads to elevated ROS production. The aggravated oxidative stress further enhances PrP aggregation in cells and finally causes PrP cytotoxicity.

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Research Highlights •

Neutralizing mutations modulate the liquid-liquid phase separation of PrPC.

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Neutralizing mutations significantly inhibit the fibrillization of PrP.

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Neutralizing mutations protect against mitochondrial damage in cells.

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Neutralizing mutations decrease ROS production and cytotoxicity of PrP.