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Heparin binding confers prion stability and impairs its aggregation Tuane C. R. G. Vieira,*,1 Yraima Cordeiro,† Byron Caughey,‡ and Jerson L. Silva*,1 *Centro Nacional de Ressonância Magnética Nuclear Jiri Jonas, Instituto de Bioquímica Médica Leopoldo De Meis, Instituto Nacional de Ciência e Tecnologia de Biologia Estrutural e Bioimagem, and †Faculdade de Farmácia, Universidade Federal do Rio de Janeiro, Rio de Janeiro, Brazil; and ‡Laboratory of Persistent Viral Diseases, Rocky Mountain Laboratories, National Institute of Allergy and Infectious Diseases, U.S. National Institutes of Health, Hamilton, Montana, USA The conversion of the prion protein (PrP) into scrapie PrP (PrPSc) is a central event in prion diseases. Several molecules work as cofactors in the conversion process, including glycosaminoglycans (GAGs). GAGs exhibit a paradoxical effect, as they convert PrP into protease-resistant PrP (PrP-res) but also exert protective activity. We compared the stability and aggregation propensity of PrP and the heparin-PrP complex through the application of different in vitro aggregation approaches, including real-time quaking-induced conversion (RT-QuIC). Transmissible spongiform encephalopathy–associated forms from mouse and hamster brain homogenates were used to seed RT-QuIC-induced fibrillization. In our study, interaction between heparin and cellular PrP (PrPC) increased thermal PrP stability, leading to an 8-fold decrease in temperature-induced aggregation. The interaction of low-molecular-weight heparin (LMWHep) with the PrP N- or C-terminal domain affected not only the extent of PrP fibrillization but also its kinetics, lowering the reaction rate constant from 1.04 to 0.29 sⴚ1 and increasing the lag phase from 12 to 19 h in RT-QuIC experiments. Our findings explain the protective effect of heparin in different models of prion and prion-like neurodegenerative diseases and establish the groundwork for the development of therapeutic strategies based on GAGs.—Vieira, T. C. R. G., Cordeiro, Y., Caughey, B., Silva, J. L. Heparin binding confers prion stability and impairs its aggregation. FASEB J. 28, 2667–2676 (2014). www.fasebj.org ABSTRACT

Key Words: glycosaminoglycan 䡠 neurodegeneration

Abbreviations: BBB, blood– brain barrier; BH, brain tissue homogenate; CD, circular dichroism; CJD, Creutzfeldt-Jakob disease; DLS dynamic light scattering; GAG, glycosaminoglycan; H1–3, helix 1–3; HS, heparan sulfate; LMWHep, lowmolecular-weight heparin; LS, light scattering; NP, nucleation-dependent polymerization; PCMA, protein misfolding cyclic amplification; PrP, prion protein; PrPC, cellular prion protein; PrPSc, scrapie prion protein; PrP-res, protease-resistant prion protein; RT-QuIC, real-time quaking-induced conversion; S1–3, sheet 1–3; ShaPrP, Syrian hamster prion protein; ThT, thioflavin T 0892-6638/14/0028-2667 © FASEB

Prions cause transmissible and genetic neurodegenerative diseases, such as bovine spongiform encephalopathy and Creutzfeldt-Jakob disease (CJD). The formation of scrapie prion protein (PrPSc) from cellular prion protein (PrPC) is the central event of prion diseases. PrPC has an unstructured N-terminal domain, a structured globular domain formed by 3 helices (H1, H2, and H3) and a short-stranded ␤ sheet (S1 and S2). The spontaneous conversion of PrPC into PrPSc is impaired by a high-energy barrier between the 2 isoforms of the prion protein. To generate PrPSc, PrPC must partially unfold and oligomerize (1, 2). PrPSc forms toxic aggregates that lead to neuronal cell death (3) and is partially resistant to proteinase K digestion; therefore, this isoform is also called proteaseresistant PrP (PrP-res; ref. 4). These features result from changes in the secondary structure of PrPC, which loses its ␣-helical-rich secondary structure and gains a ␤-sheet-rich conformation during its conversion (4, 5). Misfolded forms of PrP can assemble into amyloid fibrils both in vivo (6) and in vitro (7, 8). The method by which PrP polymerizes and the regions involved in this oligomerization are still unknown. The amino acid residues comprising the S1H1S2 region are highly conserved in various species and are thought to be involved in fibril formation (9, 10). Peptides encompassing this region were shown to aggregate into neurotoxic species (11, 12). However, the S2S3 region has also been suggested to participate in amyloid formation (13, 14). The initial steps of prion in vitro fibrillization have been suggested to involve PrP dimerization and the swapping of the globular domains while conserving the native secondary structure (15). However, the analysis of brain-derived PrPSc samples has suggested that the entire C terminus of the protein is refolded into ␤ strands with no native helices (16). 1 Correspondence: Universidade Federal do Rio de Janeiro, Instituto de Bioquímica, Bloco E Sala 10, 21941-590, Rio de Janeiro, RJ, Brasil. E-mail: J.L.S., [email protected]; T.C.R.G.V., [email protected] doi: 10.1096/fj.13-246777 This article includes supplemental data. Please visit http:// www.fasebj.org to obtain this information.

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In the past 15 yr, many researchers have hypothesized that an as yet unknown adjuvant cofactor plays a role in the process of structural conversion (17, 18). A series of molecules have been tested as adjuvants, including cellular adhesion molecules (19), nucleic acids (11, 20, 21), and glycosaminoglycans (GAGs; refs. 22, 23). GAGs are found inside and outside of cells (on the cellular surface and in the extracellular matrix) and play a pivotal role in cell– cell and cell–substratum interactions (24). GAGs have been implicated in several degenerative diseases, including prion disorders (25– 27). GAGs, especially heparan sulfate (HS) and heparin, are considered crucial molecules for prion conversion and infection (22, 26, 28). Some researchers have suggested that these molecules directly convert the protein into its protease-resistant form (22, 29), whereas others have proposed that these molecules block the interaction of PrP with endogenous GAGs and/or decrease the PrP content at the cell surface (18, 30), resulting in protective action (31–33). These conflicting effects in different systems have not yet been explained. These polysaccharides are structurally heterogeneous, displaying different patterns of sulfation, epimerization, and size, which may account for their different properties (34). The involvement of HS proteoglycans (HSPGs) in the cellular uptake of proteins involved in other neurodegenerative diseases, such as tau (Alzheimer’s disease and frontotemporal dementia) and ␣-synuclein (Parkinson’s disease), was recently demonstrated (35). Notably, a mimetic molecule of heparin blocked neuronal uptake (35). Our group reported that low-molecular-weight heparin (LMWHep) does not induce the conversion of the recombinant murine protein rPrP23–231 (36). The interaction between these 2 molecules induced local conformational changes in the protein, resulting in decreased solvent accessibility, which led to oligomerization and aggregation (36). However, this aggregation was transient and was followed by a stabilization process: after reaching equilibrium, the soluble forms of the protein exhibited a tertiary structure very similar to that of the free protein (36). We also showed that the PrP octarepeat region, a highly conserved region in the PrP N-terminal domain, is important for the PrP– LMWHep interaction at neutral pHs and suggested the existence of a second binding site in the C-terminal domain, near H2 and H3, at acidic pHs (36). In addition, we showed that LMWHep protects rPrP23–231 from aggregation induced by RNA molecules (36). This effect may be related to competition between LMWHep and RNA, because both molecules share the same binding region (20, 36, 37), and this protection may be specific for RNA-induced aggregation. However, this effect may also be related to changes in protein stability or interactions with specific PrP regions that are related to conversion and aggregation. Thus, we investigated whether LMWHep would increase murine PrP stability and, if so, whether these changes in PrP stability would affect its ability to aggregate. The answer to this ques2668

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tion may explain the protective effects that have been attributed to sulfated polysaccharides in different studies. In this study, we found that LMWHep interactions with the N- and C- terminal domains of rPrP23–231 led to kinetic and thermodynamic stabilization, which in turn prevented rPrP23–231 aggregation. MATERIALS AND METHODS Ethics statement Animal tissues used for these experiments were obtained in accordance with the U.S. National Institutes of Health (NIH) Animal Welfare Guidelines per NIH Policy Manual 3040-2 and the Guide for the Care and Use of Laboratory Animals, 8th edition, National Research Council of the National Academies, under animal study protocol 2010-045, as approved by the Animal Care and Use Committee of Rocky Mountain Laboratories, National Institute of Allergy and Infectious Diseases (NIAID), NIH. PrP and LMWHep samples Recombinant full-length PrP23–231 and PrP90 –231 from mice and hamsters, respectively, were expressed in Escherichia coli and were purified by high-affinity column refolding (38). LMWHep, with an average molecular mass of 3000 Da, was obtained from Sigma-Aldrich (cat. no. H3400; St. Louis, MO, USA). The PrP23–231–LMWHep complex was analyzed after sample disaggregation, as observed by Vieira et al. (36). Aggregation was performed in 1.5 ml test tubes (protein LoBind; Eppendorf, Hamburg, Germany), and the samples were further assayed by gel electrophoresis, to confirm that no material was lost during interaction with the test tubes (Supplemental Fig. S1). Brain tissue homogenate (BH) preparation Wild-type C57BL/10 (Prnp⫹/⫹) mice were infected with the RML (Chandler) strain and euthanized by deep isoflurane anesthesia when they reached the clinical stage of disease. Brain tissues were collected, and 10% (w/v) BHs were prepared (39) and stored at ⫺80°C. Far-UV circular dichroism (CD) CD measurements were performed with a J-715 spectropolarimeter (Jasco Corp., Tokyo, Japan) with a cuvette with a 1.0 cm path length. For the thermal denaturation experiments, ellipticity values were monitored at 222 nm as a function of temperature. The samples were heated at a rate of 2°C/min from 20 to 85°C. The ellipticity values from the buffer or the heparin solution were subtracted. Tm values were obtained by adjusting the Hill equation to the signal. The experiments were performed in 10 mM tris(hydroxymethyl)aminomethane (Tris) buffer containing 100 mM NaCl at pH 7.4. Dynamic light scattering (DLS) DLS measurements were performed with a Dynapro Nano Star instrument (Wyatt Technology, Goleta, CA, USA) with a 630 nm laser at 25°C, on 10 ␮M protein and 10 ␮M LMWHep in 10 mM Tris buffer containing 100 mM NaCl (pH 7.4). The protein solution was filtered through a 100 kDa filter before aggregation, and the buffer was filtered through a 0.02 mm

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filter. Temperature-aggregated samples were kept at 70°C for 20 min and then cooled to 25°C. Size distribution by percentage of mass was used for interpretation of the results. Data were collected and processed with Dynamics 7.1.5.6 software (Wyatt Technology). Each measurement was an average of 10 runs of 5 s each. The values shown are the mean of the results of 3 experiments. Syrian hamster PrP109 –149 (ShaPrP109 –149) peptide synthesis and sample preparation ShaPrP109 –149 was acquired from Genemed Synthesis, Inc. (San Antonio, TX, USA). The peptide was synthesized in the solid phase and then purified by using RP-HPLC to ⬎95% purity. ShaPrP109 –149 was solubilized in 50 mM 2-(N-morpholino)ethanesulfonic acid (MES) buffer (pH 5.0), containing 1% SDS and 6 M urea. The concentration of the peptide in solution was determined with a molar extinction coefficient of 8250 M⫺1 · cm⫺1 at 280 nm.

(BMG Labtech, Cary NC, USA) at 42°C for the designated period, with alternating periods of shaking for 1 min (700 rpm double orbital) and rest for 1 min throughout the incubation. The ThT fluorescence measurements (450⫾10 nm excitation and 480⫾10 nm emission; bottom read) were taken every 45 min. LMWHep (final concentration, 25 ␮M) was added to the RT-QuIC buffer in each well, or previously to the rPrP solution (stored in 10 mM phosphate buffer and 130 mM NaCl, pH 5.8 or 7.0) before rPrP was added to the RT-QuIC buffer (reaching a final concentration of 10 ␮M). The averaged sigmoidal curves were analyzed. The first part of the curve, where there were no changes in ThT fluorescence, was considered the lag phase, and the second part, where there was increasing and eventually plateaued ThT fluorescence (indicating the end of fibril formation), was considered the growth phase. The measurements were fitted, as shown by Nielsen et al. (41), to a sigmoidal curve described by the following equation using Sigma Plot (Systat, San Jose, CA, USA): Y ⫽ y i ⫹ m ix ⫹

Light scattering (LS) measurement LS was measured in a PC-1 spectrofluorometer (ISS, Champaign, IL, USA). The samples were illuminated at 320 nm, and the scattering was collected from 300 to 340 nm. The area corresponding to the intensity values was determined and compared with the initial LS value (LS0). The LS:LS0 ratio was used to evaluate the extent of sample aggregation. The ShaPrP109 –149 aggregation kinetics were measured as a function of time (excitation and emission set at 320 nm). The ShaPrP109 –149 stock solution was diluted in 10 mM Tris buffer containing 100 mM NaCl at pH 7.4, and LS was monitored until peptide aggregation was complete. Then, increasing concentrations of rPrP23–231, LMWHep, or rPrP23–231–LMWHep were added to the sample, and the LS spectra were collected. The samples were homogenized before each measurement. We also performed rPrP23–231 LS measurements at temperatures between 20 and 84°C. When the temperature returned to 20°C, the LS values were collected. In vitro conversion assay The in vitro conversion of rPrP23–231 into amyloid fibrils was performed according to the method of Baskakov and Bocharova (7), with some modifications. Briefly, rPrP23–231 samples were diluted to a final concentration of 22 ␮M in 20 mM sodium acetate buffer containing 150 mM NaCl and 6 M urea, and the samples were incubated at 37°C with continuous agitation, with or without LMWHep, for 48 h. After incubation, the samples were evaluated for thioflavin T (ThT) binding. The samples containing this probe were excited at 450 nm, and fluorescence emission was collected from 465 to 520 nm.

y f ⫾ m fx 1 ⫹ e⫺关共x⫺x0兲⁄␶兴

(1)

where Y is the fluorescence intensity, x is time, x0 is the time to 50% of the maximal fluorescence, and y and m are the fluorescence intensity and angular coefficient, respectively, from the lag phase (i) and at the plateau (f). The apparent rate constant, kapp, for the growth of the fibrils was represented by 1/␶, and the lag time was given as x0 ⫺ 2␶. The data were analyzed according to Scheme1, proposed by Jarrett and Lansbury (42).

RESULTS LMWHep binding leads to rPrP stabilization LMWHep binding to rPrP23–231 results in subtle protein conformational changes (36). However, whether this interaction affects the stability of PrP remains unknown. Therefore, we investigated the thermal stability of rPrP23–231 in the presence of LMWHep, by using CD (Fig. 1A). The CD signal variation at 222 nm was higher for the free protein, which had a Tm of 67.55 ⫾ 0.25°C (mean⫾sd), whereas the rPrP23–231–LMWHep complex yielded a Tm of 70.60 ⫾ 0.31°C (mean⫾sd). This result suggested that the LMWHep interaction increases the thermal stability of PrP. We also used LS to evaluate the temperature-induced aggregation of these 2 samples (Fig. 1B). We found that free murine rPrP23–231 underwent significant aggregation at temperatures

Real-time quaking-induced conversion (RT-QuIC) RT-QuIC was performed as described elsewhere (40). Briefly, 98 ␮l of fresh RT-QuIC buffer (10 mM phosphate buffer, pH 7.4; 130 mM NaCl; 0.1 mg/ml rPrP; 10 mM ThT; and 10 mM EDTA) was loaded into the wells of a black 96-well plate with a clear bottom (Nalge Nunc International, Penfield, NY, USA). The reactions were seeded with 2 ␮l of BH (10⫺7 dilution) in a final volume of 100 ␮l. All reactions contained a final concentration of 0.002% SDS. Recombinant fulllength PrP23–231 and PrP90 –231 from mice and hamsters were seeded with their corresponding BHs. The plates were sealed (Nalge Nunc sealer) and incubated in a Fluostar plate reader HEPARIN MODULATES PRION STABILITY AND SEEDING

Scheme 1. Data analysis scheme proposed by Jarrett and Lansbury (42), where C is the cellular form of PrP, S is the scrapie form of the protein, n* represents the critical nucleus size, and Sx is a fibril. 2669

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Figure 1. Effect of temperature on rPrP23–231 and rPrP23–231–LMWHep aggregation. A) Changes in CD ellipticity at 222 nm of PrP23–231 (solid trace) and rPrP23–231–Hep (dashed trace) with increasing temperature (40 – 85°C). Inset shows the fraction unfolded/aggregated (␣). B) Changes in the LS:LS0 ratio of rPrP23–231 (solid trace) and rPrP23–231–LMWHep (dashed trace) with increasing temperature (25– 85°C). After reaching 85°C, the temperature was reduced to 20°C, and the LS values were obtained for rPrP23–231 (solid circles) and rPrP23–231–LMWHep (open circles). The experiments were performed in 10 mM Tris buffer containing 100 mM NaCl at pH 7.4.

Effect of LMWHep on rPrP fibril assembly The ability to propagate in an autocatalytic manner is the key to the infectivity of prions (4). To analyze the effect of LMWHep on murine rPrP23–231 fibril formation, we performed an in vitro conversion assay, as described by Baskakov and Bocharova (7). In this assay, recombinant PrP protein is converted to an amyloid conformation in the presence of low concentrations of urea and guanidine hydrochloride. We observed that TABLE 1. Summary of DLS data obtained for PrP23–231 and PrP23–231–LMWHep, before and after high-temperature treatment Sample

PrP WT PrP23–231–LMWHep PrP WT, 70°C PrP23–231–LMWHep, 70°C

R (nm)

Mass (%)

2.7 ⫾ 0.2 67.3 ⫾ 7.9 1.4 ⫾ 0.6 11.1 ⫾ 4.5 119.0 ⫾ 66.3 9.1 ⫾ 1.6 180.0 ⫾ 43.3 1071.2 ⫾ 109.6 2.7 ⫾ 1.2 188.9 ⫾ 61.7 613.4 ⫾ 233.2

99.8 ⫾ 0.1 0.1 ⫾ 0.1 92.5 ⫾ 11.0 0.6 ⫾ 0.3 6.8 ⫾ 11.3 25.6 ⫾ 0.7 10.0 ⫾ 9.4 64.0 ⫾ 10.0 84.4 ⫾ 4.0 1.8 ⫾ 1.6 12.0 ⫾ 6.2

R, hydrodynamic radius; WT, wild type. Data are means ⫾ se.

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LMWHep decreased the polymerization of rPrP into amyloid structures, as measured by the binding to ThT (Fig. 2). LMWHep delays fibril formation in the RT-QuIC assay Using the RT-QuIC protocol, we also tested the effect of LMWHep on the fibrillization of recombinant PrP seeded with the BHs from animals infected with PrPSc. The addition of LMWHep to each well caused a delay in rPrP fibril formation when the reaction was seeded with infected mouse BH (Fig. 3A). In addition, when the reaction was seeded with infected hamster BH, both 3x105

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⬎65°C, whereas the complex did not aggregate at temperatures within this range, as demonstrated by only a slight increase in LS values (Fig. 1B). This effect was also observed when we used DLS to analyze samples aggregated at 70°C for 20 min (Table 1). Free protein is populated by oligomers and aggregates at ⬃1 ␮m in size, whereas the PrP23–231–LMWHep sample is populated by species with the predicted monomer size (⬃84%; Table 1).

2x105

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Figure 2. Interaction with LMWHep influences rPrP23–231 fibril formation under denaturing conditions. The rPrP23–231 samples were diluted to a concentration of 22 ␮M in 20 mM sodium acetate buffer containing 150 mM NaCl and 6 M urea and were incubated at 37°C with continuous agitation in the presence or absence of 22 ␮M LMWHep for 48 h. After incubation, the samples were diluted in the same buffer, without urea and in the presence of 20 ␮M ThT.

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Figure 3. LMWHep delays and decreases rPrP fibril formation when seeded with infected BHs. For this assay, 98 ␮l of fresh RT-QuIC buffer [10 mM phosphate buffer, pH 7.4; 130 mM NaCl; 0.1 mg/ml rPrP 23–231 (A, B) or rPrP 90 –231 (C); 10 mM ThT; and 10 mM EDTA] was added to the wells of a black 96-well plate with a clear bottom. LMWHep was added to each well at a final concentration of 25 ␮M. The reactions were seeded with 2 ␮l of a 10⫺7 dilution of mouse BH (MoBH; A) or hamster BH (HaBH; B, C). NBH, normal BH. The average ThT fluorescence from a set of quadruplicate wells is reported on the vertical axis.

a delay and a decrease in fibril formation were observed (Fig. 3B). LMWHep altered the lag time and the kapp, as shown in Table 2. This effect was dependent on the salt concentration, as a 300 mM NaCl concentration reduced the LMWHep inhibitory effect (Supplemental Fig. S2). In addition, this effect was significant when LMWHep was added during the lag phase of fibril formation (Supplemental Fig. S3). No effect was detected in the final fibril formation when the compound was added at the end of the exponential phase (Supplemental Fig. S3). The LMWHep protective effect seemed to be dependent on its interaction with the rPrP N-terminal domain, as this effect in the RT-QuIC reaction was not observed when rPrP90 –231 was used as a substrate (Fig. 3C). In the absence of this binding region, a decreased lag time was observed. We also performed the RT-QuIC reactions by adding LMWHep to rPrP at pH 7.0 or 5.8 before it was added to the RT-QuIC cocktail. In these experiments, the LMWHep–rPrP complex was added to the reaction in either its aggregated (after a 1 min incubation) or disaggregated (after a 12 h incubation) form. When rPrP was incubated with LMWHep at pH 7.0, rPrP was converted into fibrils independent of the original sample condition (aggregated or disaggregated; Fig. 4A). In contrast, when the binding reaction occurred at pH 5.8, the aggregated sample was efficiently converted into fibrils, but the disaggregated sample exhibited dramatically reduced fibril formation (Fig. 4B). Electron micrographs of these samples showed the presence of fibrils in aggregated rPrP23–231–LMWHep samples, but very few fibrils in disaggregated rPrP23–231– LMWHep samples (Supplemental Fig. S4). TABLE 2. Observed apparent rate constants and lag times Sample

rPrP23–231 rPrP23–231:LMWHep

kapp (s⫺1)

Lag time (h)

1.04 ⫾ 0.16 0.29 ⫾ 0.07

11.9 ⫾ 0.6 18.9 ⫾ 1.5

A sigmoidal equation was used in the kinetic experiments, to fit the data and obtain the apparent rate constants (kapp) and lag times. Data are means ⫾ se of results of 3 experiments.

HEPARIN MODULATES PRION STABILITY AND SEEDING

Binding of LMWHep affects rPrP23–231 incorporation into ShaPrP109 –149 aggregates The ShaPrP109 –149 peptide contains a loop, a small ␤ strand, and part of the first ␣ helix of the native ShaPrP (43). This peptide promptly aggregated under nondenaturing conditions, and the aggregation reached a plateau after only a few seconds (Fig. 5A). ShaPrP109–141 peptide has been shown to bind to PrPC, thereby seeding its aggregation (44, 45). To evaluate this seeding activity and the effect of LMWHep on this reaction, we added increasing concentrations of mouse rPrP23–231 to aggregated ShaPrP109 –149 peptide. The addition of free protein to the aggregated peptide increased the LS values. No changes in LS were observed when LMWHep or rPrP23–231–LMWHep was added to the aggregated peptide solution. These data indicate that free rPrP was seeded by the ShaPrP109 –149 peptide aggregates but that the rPrP23–231–LMWHep complex impaired this effect (Fig. 5B).

DISCUSSION In this study, the interaction between LMWHep and rPrP23–231 increased PrP protein stability and changed the ability of the protein to aggregate. This finding is supported by several studies showing that sulfated polysaccharides act as protective factors against prion conversion (31–33, 46). However, the role of the sulfated polysaccharides is still controversial, as some studies have shown contrary results (22, 29, 47). In 2011, our group showed that LMWHep interacts with rPrP23–231, causing subtle changes in protein structure; however, these changes were unrelated to protein conversion (36). In the present study, we tested whether the binding of LMWHep to rPrP would affect the thermal stability of the protein. High temperatures are known to induce rPrP conversion into a ␤-sheet-rich conformation that subsequently aggregates (48). Our results showed that LMWHep binding increased the thermal stability of rPrP23–231, leading to an 8-fold 2671

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the inhibitory effect that we observed. However, we should also consider that the structure of the protein may affect the likelihood that the protein will form fibrils. Indeed, Hafner-Bratkoviˇc et al. (15) observed that a higher thermal stability did not directly correlate with a lower conversion efficiency and that the protein structure was more relevant to its conversion efficiency. According to the model proposed by HafnerBratkoviˇc et al. (15), the rPrP fibrils produced under denaturing conditions start to self-polymerize by forming a dimer through a domain-swapping mechanism, and adjuvant molecules could stabilize these dimers. We observed that the rPrP–Hep-soluble complex failed to form amyloid fibrils under the same conditions, suggesting that stabilization of the rPrP monomer prevents dimer formation and, thus, fibrillization. The mechanisms of PrP aggregation and fibrillization still have to be investigated. Many in vitro approaches have been used to propagate PrPSc in attempts to reproduce PrP conversion and fibrillization

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Figure 4. The rPrP–LMWHep complex formed at pH 5.8 is resistant to conversion. For this assay, 98 ␮l of fresh RT-QuIC buffer (10 mM phosphate buffer, pH 7.4; 130 mM NaCl; 0.1 mg/ml rPrP 23–231–LMWHep; 10 mM ThT; and 10 mM EDTA) was loaded into the wells of a black 96-well plate with a clear bottom. LMWHep, at a final concentration of 10 ␮M, was added to rPrP in 10 mM phosphate buffer containing 130 mM NaCl at pH 7.0 (A) or 5.8 (B). The reactions were seeded with 2 ␮l of a 10⫺7 dilution of mouse BH. NBH, normal BH. The average ThT fluorescence from a set of quadruplicate wells is reported on the vertical axis.

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decrease in the temperature-induced aggregation. Changes in stability have been associated with the hereditary forms of prion diseases (49). Protein solvation (50) and electrostatic interactions (51) modulate PrP stability and can affect its conversion tendency. Post-translational modifications, such as N-glycosylation, can also exert these effects (52). The negative charge of LMWHep may also affect these properties and help protect PrP from the effects of high temperatures. Although polysaccharides were shown to induce the fibril formation of a PrP fragment (PrP185–208; ref. 53), these molecules were also shown to inhibit the fibril formation of PrP106–126 (32). These apparently controversial results may be explained by differences in the peptide sequences. The controversial results may also be explained by differences in heparin/HS structure and/or chain length. In our study, we used the full-length prion protein, which may also account for

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Figure 5. rPrP23–231 and LMWHep modulate ShaPrP109 –149 aggregation. A) ShaPrP109 –149 LS (320 nm) after dilution in 10 mM Tris buffer containing 100 mM NaCl (pH 7.4). After the LS values reached a plateau, the samples were increasingly added to the aggregated peptide solution (arrow). B) ShaPrP109 –149 aggregates were incubated with increasing concentrations of rPrP23–231, rPrP23–231–LMWHep, or LMWHep.

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in vivo. Among these methodologies, synthetic peptides that correspond to the PrP regions involved in the secondary structure changes involved in the conversion from PrPC to PrPSc have been widely used (11, 44, 54). We also tested the effects of LMWHep while using this approach with the ShaPrP109 –149 peptide. When diluted in a buffer without denaturing agents, the ShaPrP109 –149 peptide immediately aggregates and forms ␤-sheet-rich structures (11). The addition of peptides encompassing this region of PrPC has been shown to seed PrP aggregation, leading to cellular neurotoxicity (44, 45). We found that the ShaPrP109 –149 peptide seeded rPrP23–231 aggregation, but that the addition of the rPrP23–231–LMWHep complex did not. When the rPrP– LMWHep complex was added to aggregated ShaPrP109–149, we observed that rPrP was not incorporated into the preformed aggregates, suggesting that LMWHep binding modulates prion self-propagation. More recently, protein misfolding cyclic amplification (PMCA) and RT-QuIC reactions have been extensively used to amplify PrPSc (55, 56). PMCA assays have been performed in the presence of different adjuvant cofactors, such as nucleic acids (57), lipids (57, 58), and sulfated polysaccharides (47, 59, 60), to evaluate their influence on PrP conversion. LMWHep enhanced the amplification of variant CJD seeds in PMCA with a narrow range of concentrations, but the effect was negligible with sporadic CJD and 263K seeds, exhibiting the dependence of these reactions on the seed types (47). In the current study, we used the RT-QuIC reaction to further investigate LMWHep’s modulation of PrP fibrillization and saw that LMWHep modulated PrP fibrillization by affecting nucleation and the growth kinetics. Moreover, LMWHep exerted a thermodynamic effect on ShaPrP, as determined by the reduced fibril mass at the end of the reaction. This effect seemed to be triggered by electrostatic interactions with the N-terminal region of rPrP when the LMWHep concentration exceeded a 1:6 molar ratio (PrP: LMWHep). The absence of the N terminus resulted in the opposite effect, with LMWHep causing aggregation to accelerate. The N-terminal region has been shown to modulate the efficiency of PrP conversion and may be important in the initial step of protein conversion, as it may provide a binding surface for PrPSc, thereby leading to an increased conversion efficiency (61, 62). Our results suggest that the interaction of LMWHep with the N-terminal rPrP region modulates its accessibility to the PrPSc seeds, thereby decreasing the conversion’s efficiency. In our previous work (36), we showed that LMWHep leads to transient rPrP aggregation, resulting in a soluble PrP–LMWHep complex after 12 h, with no detected conversion into a PrPSc-like species. This interaction presented a stoichiometry ratio of 1:1 (rPrP: LMWHep) at pH 7.0, and the binding was mediated by the N-terminal octarepeat region. In contrast, at pH 5.8, in addition to binding to the octarepeat domain, we observed heparin interacting with the C-terminal domain near H2 and H3, yielding a molar ratio of 1:2 (rPrP:LMWHep). When the aggregated HEPARIN MODULATES PRION STABILITY AND SEEDING

rPrP–LMWHep (1:2) complex was used as a substrate, we did not observe any inhibition and instead observed an acceleration of the fibrillization. Notably, the disaggregated (soluble) rPrP–LMWHep (1:2) complex that formed at pH 7.0 did not show any aggregation inhibitory effect, whereas the soluble complex that formed at pH 5.8 was dramatically resistant to fibrillization. The H2H3 region appears to be important for PrP conversion and consequent aggregation, as many mutations related to the familial forms of prion diseases have been identified in this region (49, 63). Compounds that interact with this region were shown to increase the protein’s thermal stability, resulting in a stabilized PrPC conformation, acting as a molecular chaperone (63). Our results indicate that the interaction of the PrP N or C terminus with LMWHep at low pH, most likely at the octarepeat or the H2H3 region (Fig. 6A), respectively, impairs conversion and the consequent fibril formation. The molecular details of prion protein conversion and aggregation have been the focus of many investigations (64). Prusiner (65) postulated that the infectious species was a PrP with an altered conformation (S) and that this form would catalyze the conversion of a PrPC monomer (C), suggesting propagation by autocatalysis. Jarrett and Lansbury (42) proposed a nucleation-dependent polymerization (NP) mechanism to explain prion transmission and propagation; in this mechanism, C and S are in equilibrium, and S slowly aggregates to form a critical nucleus (nucleation phase). Once the nucleus is formed, further aggregation becomes thermodynamically favorable, leading to fibril formation and growth (growth phase). PrP aggregation followed by RT-QuIC experiments showed an NP profile, with an initial lag phase followed by rapid growth and a stationary phase. Altogether, our results indicate that the interaction of LMWHep with the N-terminal region of rPrP modulates fibril formation by changing the equilibrium between C and S through an increase in the stability of C (Fig. 6B). Because the availability of S is necessary during both the nucleation and growth phases, LMWHep interferes with the kinetics of both phases. These results also suggest that conversion occurs mainly during the nucleation phase, as this effect was not observed when LMWHep was added during the exponential growth phase. We also observed that the interaction of LMWHep with the C-terminal domain impaired fibrillization, reinforcing the ideas that this domain is the main region that undergoes structural conversion and that interactions at this site affect its conversion (Fig. 6B). Although GAGs have been shown to exert antiprion effects in cell culture (31, 33) and animal models (66), free GAGs may stimulate prion conversion (22). Our studies shed light on the paradoxical effects of sulfated polysaccharides on the cellular PrP structure. Recently, our group showed that LMWHep leads to rPrP23–231 aggregation in vitro but that this effect is transitory and does not directly act on conversion (36). However, cellular GAGs are essential receptor molecules for PrPSc infec2673

Figure 6. Three-dimensional structure and a schematic view of rPrP 23–231 nucleation-dependent polymerization. A) Three-dimensional structure of mouse PrP [Protein Data Bank (PDB) entry 1XYX]. The chain is colored according to amino acid sequence; ␣ helix and ␤ strands are indicated. B) PrPSc catalyzes the conversion of PrPC in an autopropagative manner. This reaction is in equilibrium, and PrPSc slowly aggregates to form a critical nucleus. n, nucleation phase. Once the nucleus is formed, further aggregation becomes thermodynamically favorable, leading to fibril formation and growth (growth phase). The interaction of LMWHep with the N-terminal region modulates fibril formation by increasing PrPC stability and changing the equilibrium between PrPC and PrPSc. Because the conversion of PrPC to PrPSc is necessary at both the nucleation and growth phases, LMWHep interferes with the kinetics of both. The interaction of LMWHep with the rPrP N- and C-terminal domains impairs fibrillization and dramatically reduces fibril formation.

tion (28) and the uptake of other aggregated proteins, such as tau and ␣-synuclein (35). Our results show that the PrP–LMWHep interaction increases PrP stability. Similarly, GAGs may also stabilize the scrapie isoform, contributing to conversion. However, GAGs display protective effects that can be explained by a possible competition with endogenous GAGs. Our data indicate that the rPrP–LMWHep complex is less susceptible to aggregation than the free protein is, through the modulation of fibrillization thermodynamics and kinetics instead of through competition; in this case, LMWHep acts as a true inhibitor (67). HS has been found to aggregate with a variety of amyloid proteins (25–27), and therefore, many therapeutic strategies have been developed to modulate this interaction to affect disease establishment and progression. HS derivatives and analogues have been shown to inhibit PrPSc accumulation in cell culture and prolong the survival of scrapie-infected hamsters (68, 69) and to reduce amyloid A (AA) deposition and AA progression (70, 71). These molecules have been used in clinical trials showing promising results in patients with AA amyloidosis (72). Many therapies have been proposed for prion diseases, but there is still no effective treatment. The intraventricular administration of pentosan polysulfate, an analogue of HS, prolonged the survival of a patient with variant CJD but did not change the progression of the disease (73) and showed no clinical benefit for another patient (74). This treatment also decreased the levels of resistant PrP in the brain of a patient with sporadic CJD (75). Sulfated polysaccharides have been used in many experimental models of prion disease, but their inability to cross the blood– brain barrier (BBB) due to their molecular weight and/or charge hinders their use as therapeutic drugs (76). However, it has been shown that LMWHep fragments can cross the BBB and inhibit a ␤-amyloid amyloidogenic pathway in neuroblastoma cells (77). The administration of LMWHep–Neuroparin (average molec2674

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ular mass of 2100 Da) in Alzheimer’s disease animal models showed its neuroprotective role (78), and Neuroparin can be detected in the brain and cerebrospinal fluid after its intravenous and subcutaneous administration, showing that it may pass through the BBB (79). Mutations in the PrP globular domain that change the packing of H1 and H3 increase the thermodynamic stability of PrP, thereby changing its fibrillization kinetics and consequently decreasing its propensity to aggregate, as well as inhibiting the propagation of transmissible spongiform encephalopathy in animals (80). In this study, we showed that LMWHep, which has a molecular mass of 2100 Da, modulates the fibrillization kinetics and thermodynamics of PrP by increasing PrP protein stability. These results suggest that LMWHep can be used as a therapeutic drug for prion diseases and may also be effective for other neurodegenerative diseases, in that GAGs contribute to the prion-like cellular uptake and propagation of the misfolded proteins involved in these pathologies (35). The authors thank A. Hughson for purifying the recombinant PrP and for preparing the brain homogenate samples and G. and L. Raymond for technical assistance. T.C.R.G.V. received a postdoctoral fellowship from Conselho Nacional de Desenvolvimento Científico e Tecnológico (CNPq) through the Science Without Borders Internship Program and from Fundação Carlos Chagas Filho de Amparo a` Pesquisa do Estado do Rio de Janeiro (FAPERJ). This work was supported by grants from CNPq, Instituto Nacional de Ciência e Tecnologia de Biologia Estrutural e Bioimagem (CNPq INCT Program) and FAPERJ.

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Received for publication November 22, 2013. Accepted for publication March 4, 2014.

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