Zn Superoxide Dismutase Forms Amyloid Fibrils under Near

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Cu/Zn Superoxide Dismutase Forms Amyloid Fibrils Under Near-Physiological Quiescent Conditions: the Roles of Disulfide Bonds and Effects of Denaturant M. Ashhar I. Khan, Michal Respondek, Sven Kjellstrom, Shashank Deep, Sara Linse, and Mikael Akke ACS Chem. Neurosci., Just Accepted Manuscript • Publication Date (Web): 06 Jun 2017 Downloaded from http://pubs.acs.org on June 7, 2017

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ACS Chemical Neuroscience

Cu/Zn Superoxide Dismutase Forms Amyloid Fibrils Under Near-Physiological Quiescent Conditions: the Roles of Disulfide Bonds and Effects of Denaturant M. Ashhar I. Khan1,3, Michal Respondek1, Sven Kjellström2, Shashank Deep3, Sara Linse2, Mikael Akke1,* 1

Biophysical Chemistry, Center for Molecular Protein Science, Department of Chemistry, Lund University, Lund, Sweden

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Biochemistry and Structural Biology, Center for Molecular Protein Science, Department of Chemistry, Lund University, Lund, Sweden 3

Department of Chemistry, Indian Institute of Technology Delhi, New Delhi, India

* Correspondence to: [email protected]

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Abstract Cu/Zn superoxide dismutase (SOD1) forms intracellular aggregates that are pathological indicators of amyotrophic lateral sclerosis. A large body of research indicates that the entry point to aggregate formation is a monomeric, metal-ion free (apo), and disulfidereduced species. Fibril formation by SOD1 in vitro has typically been reported only for harsh solvent conditions or mechanical agitation. Here we show that monomeric apoSOD1 in the disulfide-reduced state forms fibrillar aggregates under near-physiological quiescent conditions. Monomeric apo-SOD1 with an intact intramolecular disulfide bond is highly resistant to aggregation under the same conditions. A cysteine-free variant of SOD1 exhibits fibrillization behavior and fibril morphology identical to those of disulfide-reduced SOD1, firmly establishing that intermolecular disulfide bonds or intramolecular disulfide shuffling are not required for aggregation and fibril formation. The decreased lag time for fibril formation resulting from reduction of the intramolecular disulfide bond thus primarily reflects the decreased stability of the folded state relative to partially unfolded states, rather than an active role of free sulfhydryl groups in mediating aggregation. Addition of urea to increase the amount of fully unfolded SOD1 increases the lag time for fibril formation, indicating that the population of this species does not dominate over other factors in determining the onset of aggregation. Our results contrast with previous results obtained for agitated samples, in which case amyloid formation was accelerated by denaturant. We reconcile these observations by suggesting that denaturants destabilize monomeric and aggregated species to different extents and thus affect nucleation and growth.

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Keywords: amyotrophic lateral sclerosis, disulfide reduction, ThT fluorescence, protein aggregation, protein unfolding, transmission electron microscopy

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Introduction Amyotrophic lateral sclerosis (ALS) is a fatal neurodegenerative disease selectively affecting motor neurons.1 More than 140 point mutations in Cu,Zn-superoxide dismutase (SOD1) have been associated with the familial form of ALS. In addition, a growing body of data implicate also wildtype SOD1 in sporadic ALS.2-4 The normal, physiological function of SOD1 is to protect the cellular components from the damaging effects of metabolic byproducts, such as superoxide radical anions This function is retained in ALS, where cell toxicity arises from gain of noxious function that appears to be linked to aberrant oligomers or fibrillar aggregates of SOD1.1,

5, 6

Still, despite

extensive studies of ALS-associated variants of SOD1 the defining mechanisms of the disease remain elusive. A number of reports suggest that SOD1-containing pathological inclusions in ALS have similarities with amyloid fibrils present in vitro.7-9 SOD1 fibril formation in vitro is catalyzed by extracts of inclusions from ALS transgenic mouse tissue.10 Conversely, amyloid-like SOD1 fibrils formed in vitro seed intracellular aggregation of endogenously expressed SOD1.11 Furthermore, SOD1 fibrils administered to cell cultures trigger effects similar to those caused by ALS inclusions in vivo, in that they induce elevated expression levels of cytokines and inflammation12 and activate microglia.13 This strong correspondence between in vivo ALS inclusions and in vitro formed SOD1 fibrils motivate further biophysical studies of SOD1 fibril formation to understand the underlying physicochemical factors governing SOD1 aggregation and fibrillar inclusions in ALS.

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Native human SOD1 is a very stable homodimeric protein, whose 153-residue protomers fold into an eight-stranded β-sandwich that has three long loops (loops IV, VI, and VII) extending on one side of the structure. Each protomer coordinates one copper and one zinc ion and maintains a highly conserved intramolecular disulfide bond in the reducing environment of the cytosol. Loop IV contains the Zn2+ binding site and contributes with one side-chain ligand to the Cu2+ binding site, which is otherwise defined by residues located in β-strands 4 and 7. The disulfide bond is formed by residues C57 in loop IV and C146 in β-strand 8 (Figure 1). The metal binding sites and the disulfide bond thus anchor loop IV to the β-sandwich core of SOD1. Loss of metal ions and reduction of the intramolecular disulfide bond destabilize the native dimer and generates an apo monomer, which has been shown to be the entry point to aggregation.1417

Several studies indicate that the gateway to aggregation involves partially unfolded

species,18-20 whereas other reports suggest that aggregation emanates from the fully unfolded protein.21 Many proteins linked to neurodegenerative diseases form amyloid fibrils through a nucleation-dependent polymerization mechanism.22, 23 In the case of SOD1, which is a relatively stable protein, in vitro studies of fibril formation have typically involved extreme experimental conditions, such as elevated temperature, low pH, the presence of denaturant or organic solvents,24,

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or oxidative cross-linking of cysteines to form

disulfide bridges.26 It has also been shown that forceful agitation of the sample under otherwise physiological conditions triggers apo, disulfide-reduced SOD1 to form ordered fibrillar structures similar to those found in other amyloidogenic diseases.21, 26-31 Thus, it has been established that SOD1 forms fibrils under a range of different conditions that are

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all rather harsh and non-physiological, but the question remains whether SOD1 can also form fibrils under quiescent and otherwise physiological conditions.

Figure 1. Structure of monomeric SOD1. The two β-sheets are colored cyan and pink, while loop IV is green, loop VII is purple, and other turns and loops are gray. The C57–C147 disulfide bond is highlighted in yellow. The figure was made using the PyMOL molecular graphics system (Schrödinger, LLC) and PDB entry 2xjk,32 which represents the Cu- and Zn-bound state of the monomeric, pseudo-wildtype variant C6A/F50E/G51E/ C111A employed in the present study.

Considerable attention has been paid to the role of disulfide cross-linking in the aggregation of SOD1. Clinical as well as in vitro studies have shown that intermolecular disulfide bridges are prevalent in SOD1 aggregates, leading to the hypothesis that such interactions are necessary for oligomerization and aggregation. A number of in vivo33-38 and in vitro26, 27, 39 studies have shown the direct involvement of intermolecular disulfide cross-linking in insoluble aggregates, as well as in soluble oligomers. However, this aggregation mechanism has been contradicted by reports suggesting that while the free cysteines may modulate SOD1 aggregation, disulfide cross-linking might not be a critical event.30,

40-43

Other studies have indicated that free thiolates are required to trigger 6


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nucleation of SOD1 fibrils,28 while seeded fibril formation has been shown to be independent of intermolecular disulfide linkages.44 Recent studies further described intramolecular scrambling of disulfide bonds involving all four cysteines (C6, C57, C111, and C146) in apo-SOD1 as a critical step preceding disulfide-mediated oligomerization.45, 46

Thus, there still remains a great deal of uncertainty regarding the role of disulfides in

the aggregation mechanism of SOD1. Here we demonstrate unequivocally that SOD1 forms fibrillar aggregates under quiescent conditions at near-physiological pH, ionic strength, and temperature over a time frame of weeks. Our results further show that intermolecular disulfide bonds are not required for the protein to form aggregates, even in the absence of fibril seeds. Similarly, scrambling of intramolecular disulfide bonds is not required for aggregation. Instead, disulfide reduction affects the lag time of SOD1 fibrillation primarily by removing the bond anchoring loop IV to the protein core and thus promoting local unfolding of the structure, which triggers aggregation. Our results further show that urea denaturation increases the lag time for aggregation, suggesting that different species present along the aggregation pathway have different stabilities towards denaturation, while the population of the globally unfolded state cannot be considered alone as a dominant factor driving aggregation under these conditions.

Results and Discussion We studied aggregation and fibril formation of monomeric apo-SOD1, using the pseudowildtype construct C6A/F50E/G51E/C111A18, 47 denoted pwt-SOD1, under quiescent and near-physiological conditions. The C6A and C111A mutations in pwt-SOD1 prevent aberrant disulfide linkages involving these residues, and the F50E and G51E mutations 7



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introduce charged groups at the native dimer interface, so that charge repulsion makes the protein monomeric.18, 47 To investigate whether intermolecular disulfide cross-linking or exchange of intramolecular disulfides are involved in the aggregation of SOD1, we used a cysteine-free mutant (C57S/C146S pwt-SOD1, denoted SOD1∆C), which eliminates disulfide cross-linking, and compared the behavior of this variant with that of pwt-SOD1 in the disulfide oxidized and reduced states. We carried out a series of fibril formation experiments over several hundred hours, using established methods.48 We monitored fibril formation by thioflavin-T (ThT) fluorescence in PEG-ylated 96-well plates under a variety of conditions that tune the relative populations of reduced/oxidized C57–C146 disulfide (in pwt-SOD1) and folded/unfolded protein (in both pwt-SOD1 and SOD1∆C). We verified that intrinsic ThT fluorescence does not degrade, but is maintained throughout the length of the experiments (Figure S1). We verified the formation of amyloid fibers at the end of the reactions by transmission electron microscopy (TEM). In all experiments we used monomeric protein freshly prepared by analytical gel filtration to make sure that samples did not contain any oligomeric or aggregated species at the start of each experiment (Figure S2). This treatment is critical in order to obtain a high level of reproducibility in fibrillization experiments.49-51 NMR spectra acquired on freshly prepared monomeric samples show that the protein is folded (Figure S3). Given the absence of any peaks from the unfolded state above the baseplane noise and the average signal-to-noise ratio of approximately 250, we estimate the population of unfolded species to be very low under these conditions (Figure S3). Thus, all experiments conducted in the absence of denaturants were initiated using folded, monomeric protein.

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Reduced SOD1 forms fibrils under near-physiological conditions without mechanical agitation. Mechanical agitation of protein solutions substantially accelerates the fibrillation process by catalyzing secondary as well as primary nucleation events due to increased rate of fibril fragmentation and increase of the air-water interface.52 In the case of SOD1, agitation has commonly been employed to accelerate fibril formation, whereas non-fibrillar aggregates have previously been observed in physiological buffer without agitation.27,

45, 53

The experimental conditions that we

employed throughout our study resemble the physiological environment in terms of the pH (7.0), temperature (37°C), and absence of mechanical stimulus in the form of stirring or shaking.

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Figure 2. The lag time for fibril formation by monomeric apo-SOD1 decreases when the native disulfide bond is reduced. (A) pwt-SOD1 and (B) SOD1∆C. ThT fluorescence was measured as a function of time with different concentrations of reducing agent (TCEP) added: blue, 10 equivalents of TCEP; red, 1 equivalent; green, 0.1 equivalent; black, no TCEP. The SOD1 concentration was 200 µM and the temperature was 37 °C. The graphs represent the average of three experiments. The discontinuity in the blue line in panel A is likely caused by instrumental instabilities affecting individual time course experiments. See Figure S5 for plots including all individual data sets.

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Under conditions where the C57–C146 disulfide bond is intact, pwt-SOD1 did not form amyloid structures within a period of four weeks (Figure 2A, the black curve extends to one week; Figure S4 shows data extending to four weeks). However, upon reduction of the disulfide bond by addition of tris(2-carboxyethyl)phosphine (TCEP), pwt-SOD1 formed amyloid structures within a week, as detected by ThT fluorescence (Figure 2A; green, red, blue curves) and verified by direct imaging of fibrils using electron microscopy (Figure 3A). Due to the very long lag time, we did not aim to follow the reaction to completeness, but settled for a semi-quantitative interpretation, which suffices for the purpose of the present investigation. Despite the long lag time (up to several hundred hours), the scatter among replicate time course experiments is relatively limited (see Figure S5) and corresponds to a relative error of roughly 5–10% in estimated lag times, which is on par with that observed for more rapid kinetics, e.g. fibril formation of the Aβ peptide.54 The long lag time is in agreement with results from Furukawa and coworkers,30 who did not detect any aggregation under quiescent conditions for a time period up to 3 days. Increasing the relative population of the disulfide-reduced state by increasing the TCEP:protein ratio resulted in a progressively decreased lag time for aggregation from approximately 280 h in the absence of TCEP to ca 50 h at 10:1 TCEP:SOD1 (Figure 2A), further substantiating that fibril formation originates from the disulfide-reduced state. Also, we find that higher protein concentration leads to a reduction in lag time (Figure S6), as expected for nucleation-dependent amyloid formation.55

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Figure 3. Transmission electron microscopy images of SOD1 amyloid fibrils. (A) Fibrils of pwtSOD1 formed upon reduction of the native C57–C146 disulfide bond with TCEP. (B) Fibrils of the cysteine-free variant SOD1∆C formed without any additives. The scale bar indicates 200 nm.

Intermolecular

disulfide

cross-linking

or

intramolecular

disulfide

scrambling is not required for aggregation of SOD1. A large body of work suggests that intermolecular disulfide bonds are required for oligomerization and aggregation of SOD1,26, 28, 34, 36, 39, 45, 46 while other reports contradict this hypothesis.40-43 Even though the addition of TCEP reduces the equilibrium population of disulfideoxidized species to very low levels, it is in principle possible that pwt-SOD1 could aggregate via transient intermolecular disulfide cross-linking. To stringently investigate the requirement of intermolecular disulfides in the aggregation process of SOD1, we carried out fibril formation experiments with the cysteine-free variant SOD1∆C. The results show that SOD1∆C perfectly mimics the behavior of fully reduced pwt-SOD1 within the margin of error (cf. blue curve in Figures 2A and B), directly demonstrating that free sulfhydryl groups are not required for fibril formation and thereby disproving any obligatory role of disulfide linkage in the aggregation of SOD1. We ascertained that

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the addition of TCEP did not affect the aggregation process of SOD1∆C by performing fibril formation assays with varying amounts of reducing agent present. These experiments showed that there is no significant effect of TCEP on the fibrillation kinetics of the cysteine-free variant (Figure 2B). Furthermore, the fibril morphologies of pwtSOD1 and SOD1∆C do not differ significantly (Figure 3), indicating that the fibrillation processes are highly similar, if not identical, for these two variants. Thus, we conclude that neither intermolecular disulfide cross-linking nor intramolecular disulfide scrambling is a critical element of fibril formation by SOD1. However, the native-like intramolecular disulfide bond clearly retards aggregation, in agreement with earlier reports, suggesting that it is incompatible with the SOD1 fibril structure. Addition of denaturant increases the lag time for fibril formation. As described above, we found that both SOD1∆C and reduced pwt-SOD1 form fibrils under physiologically relevant conditions (quiescent, pH 7, 37 °C) without the addition of denaturing agents (e.g. urea or guanidinium chloride), i.e., under conditions where the population of the fully unfolded state is low (less than 1% in the case of SOD1∆C and even lower for pwt-SOD1; Figure S3). We next investigated the coupling between SOD1 unfolding and fibril formation by adding increasing amounts of urea. In the case of oxidized pwt-SOD1 (with intact intramolecular disulfide bridge), no fibril formation was observed at any population of unfolded SOD1 monomer, pU = 0–99% (Figure S7). Clearly, unfolding alone does not lead to aggregation of pwt-SOD1 under these conditions, in sharp contrast to the effect of disulfide reduction. This further strengthens the conclusion that the native-like intramolecular disulfide-bond is incompatible with the SOD1 fibrillar structure.

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Figure 4. Retardation of SOD1∆C fibril formation with increasing concentration of denaturant. Fibrillation time course experiments on SOD1∆C with varying concentrations of urea yielding populations of the unfolded, monomeric state of: pU < 5% (green), no urea added; pU ≈ 50% (red), 0.22 ± 0.01 M urea; and pU > 99% (blue), 2.58 ± 0.03 M urea. The SOD1 concentration was 200 µM and the temperature was 37 °C. The graphs represent the average of three experiments. The gap in the graphs corresponds to a period between 165 h and 307 h during which the 96-well plate was removed from the plate reader and stored at 37 °C. See Figure S8 for a plot including all individual data sets.

Moreover, the corresponding experiments on SOD1∆C reveal that fibril formation is in fact dramatically slowed down in the presence of urea, even at the lowest concentration of urea, 0.22 M (Figure 4). Apparently, the denaturant effectively suppresses fibril formation. This suggests that urea shifts the folding/unfolding equilibrium and kinetics of the monomer and aggregated forms to different extents and further that urea affects the height of the transition barriers for the steps underlying the aggregation process. This observation contrasts with previous results from experiments

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on agitated SOD1 samples, where the lag time and rate of fibril formation decreased as the concentration of urea increased.21 This may indicate that fibrils are destabilized by urea and therefore break more easily under mechanical agitation compared to fibrils in the absence of urea. Fibril breakage is a secondary process that provides new free ends for elongation and consequently leads to rapid increase in fibrils mass. We conclude that the pathways of aggregation and fibril formation are markedly dependent on the experimental conditions.

Conclusions Here we have shown that SOD1 in the monomeric, metal-free, and disulfide-reduced state forms fibrillar aggregates under near-physiological and quiescent conditions, while the monomeric, metal-free, and disulfide-oxidized state does not (at least not within a timespan of one week). Comparisons of fibril formation experiments conducted with disulfide-reduced pwt-SOD1 and the cysteine-free variant SOD1∆C show that the increased propensity of the disulfide-reduced state, compared to the disulfide-oxidized state, to form amyloids arises from local unfolding of the structure induced by breaking the disulfide bond between loop IV and the β-barrel, rather than from a direct involvement of aberrant disulfide bonds. Thus, we conclude that disulfide bonds are not required for any steps of the aggregation processes. Our results further show that the addition of urea leads to progressively increased lag times for fibril formation under quiescent conditions, in sharp contrast to previous results for agitated samples.21 Apparently urea interferes with fibril nucleation and growth by affecting to different extents the stabilities of monomeric and aggregated or fibrillar species, the outcome of which depends on whether the sample is agitated or

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quiescent. Thus, it is clear that the detailed experimental conditions have a strong bearing on the mechanism of fibril formation. The present sets of data thus provide an important complement to previous results obtained on agitated samples in vitro. Based on experiments conducted under near-physiological and quiescent conditions, our work provides results that fill the gap between previous data from in vitro and cell-based studies.

Methods Materials. All chemicals were of analytical grade and purchased from Sigma-Aldrich (USA). Tris(2-carboxyethyl)phosphine (TCEP) solution neutralized with NaOH was used as reducing agent. The concentrations of ultrapure urea (Bioxtra, U0631) stock solutions were determined by measuring their refractive index. All solutions were prepared in ultrapure water (< 18.2 MΩ cm at 25 °C, MilliQ grade). All reagents and buffers were filtered through a 0.22 µm cutoff filter and degassed before use. The pH of all reagents and buffers was maintained at 7.0 unless stated otherwise. Plasmid design. The plasmids were designed as described18 and obtained from GenScript USA, Inc. All constructs used herein are based on the well-established pseudowild type variant of SOD1 (pwt-SOD1), which is a quadruple point mutant of human wild-type SOD1: C6A/F50E/G51E/C111A; the C to A mutations ensure that aberrant disulfide linkages cannot form and the F50E and G51E mutations introduce charged groups at the dimer interface, thereby making the protein monomeric.47 We also created a cysteine-free mutant (SOD1∆C) that includes the C57S and C146S mutations in addition to the pwt-SOD1 template. The structure of pwt-SOD1 shows no significant deviations from wild-type SOD1, and the structure of the apo form of the cysteine-free mutant

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differs only in a minor rearrangement of loop IV, resulting in looser packing against the core.56 Intact mass determination and sequence composition of all constructs were confirmed by MALDI mass spectrometry (Figures S9 and S10). Protein expression and purification. pwt-SOD1 and SOD1∆C were expressed in Escherichia coli strain BL21 (DE3) pLysS* and purified as described previously,18, 57 except that the heat denaturation step was not included for SOD1∆C, since our protocol does not involve co-expression of the copper chaperone CCS in SOD1∆C. Both pwtSOD1 and SOD1∆C were initially purified using size-exclusion gel chromatography, followed by purification on Source 15Q anion exchange column. Apo pwt-SOD1 was prepared by extensive dialysis against 100 mM sodium acetate buffer at pH 3.8 in the presence of 10 mM EDTA followed by thorough dialysis against 20 mM MOPS buffer, pH 7.0, for storage. Protein sample preparation. All experiments were performed in 20 mM 3-(Nmorpholino)-propanesulfonic acid (MOPS) buffer at pH 7.0, unless otherwise specified. To ensure reproducibility, every experiment was started with the isolation of pure monomeric SOD1 using fast protein liquid chromatography (FPLC) employing an analytical size-exclusion column (Superdex 75 10/300 GL) equilibrated with 20 mM MOPS buffer pH 7.0. Only the central portion of the monomer peak (Figure S2) was collected to minimize any contamination from bacterial proteins or preexisting aggregates. The concentration of the collected protein fraction was estimated by measuring the absorbance at 280 nm on a NanoDrop 2000/2000c spectrophotometer (Thermo Scientific), using an extinction coefficient of 5500 M–1cm–1.

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Fibrillization assay. All samples were prepared in low-binding Eppendorf tubes (Genuine Axygen Quality, Micro tubes MCT-200-L-C) and kept on ice before distributing them in 100 µl aliquots onto a 96-well half-area plate of polyethylene glycolcoated black polystyrene with clear bottom (Corning 3881). Samples were prepared with 50 µM, 100 µM, or 200 µM protein dissolved in 20 mM MOPS buffer pH 7.0 and 40 µM ThT. Different sample conditions were screened by adding TCEP and urea in different combinations and concentrations, see S1 and S2 Tables for a complete listing. We estimated the population of unfolded monomer (pU) in urea-containing solutions based on published kinetic parameters of unfolding.21, 58 For pwt-SOD1, pU = 10% (1.19 ± 0.01 M urea), pU = 50% (1.81 ± 0.03 M urea), and pU = 99% (3.75 ± 0.05 M urea).58 However, the published parameters yield a free energy of unfolding in pure water that corresponds to pU = 20–30% for SOD1∆C,21 in stark disagreement with our results from NMR, which show that pU is less than 1% (Figure S3); this discrepancy could be due to inherent extrapolation errors in the kinetic study, but could also reflect differences in sample age (we used freshly prepared monomeric samples) and sample conditions (buffer, pH, temperature). For this reason, we report conservative ranges of pU for SOD1∆C at three concentrations of urea: pU < 5% (no urea added), pU = 50–60% (0.22 ± 0.01 M urea), and pU > 99% (2.58 ± 0.03 M urea); the use of approximate ranges of pU does not impact on the results presented herein. See Table S1 and S2 for an overview of sample conditions. Each sample was run in triplicate, distributed on the plate in a random manner to mitigate the effects of potential temperature variations across the plate during measurement. Plates were sealed with a plastic film (Corning 3095) and placed either in a Fluostar Omega or Polarstar Optima plate reader (BMG Labtech, Offenburg, Germany) and incubated at 37

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°C without agitation. ThT fluorescence was monitored every 3 min through the bottom of the plate, with the excitation and emission wavelengths set to 440 and 480 nm, respectively. Transmission electron microscopy. Aggregates were visualized by transmission electron microscopy (TEM). Approximately 8 µl of fresh SOD1 aggregates taken from the 96-well plate were applied on 300-mesh carbon-coated grid with Formvar carbon film (EM Sciences). Samples were allowed to adsorb for 2 minutes. After washing with MilliQ water, negative staining with 1.5% uranyl acetate (UA) was performed for 30 seconds. Excessive UA solution was removed and the grid was blotted dry. Images were obtained on a Philips CM120 BioTWINCryo equipped with a postcolumn energy filter (Gatan GIF100), operating at an acceleration voltage of 120 kV. The images were recorded digitally with a CCD camera under low electron dose conditions at a magnification of 1:31,000. NMR

spectroscopy.

Two-dimensional

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N–1H

heteronuclear

single-quantum

coherence (HSQC) spectra were acquired on 2 mM uniformly 15N-labelled apo SOD1∆C dissolved in 20 mM MOPS buffer, pH 7.0, 10% D2O at a temperature of 25 °C, using an Agilent DirectDrive 500 MHz spectrometer. The sample was freshly prepared in monomeric form immediately before each NMR experiment, as described above for the fibrillation assays. NMR data were processed using NMRPipe59 and analyzed using ccpNMR.60 MS analysis. Intact mass analysis and peptide mass fingerprinting were carried out on a MALDI-TOF-TOF-MS instrument, Applied Biosystems Proteomics Analyser 4700 (Applied Biosystems, Framingham, MA). The matrix solution consisted of 5 mg/ml α-

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cyano-4-cinnamic acid (α-CHCA) dissolved in acetonitrile phosphoric acid (50:0.1 v/v). 10 fmol/µl of des-Arg-Bradykinin, m/z 904.468, and 20 fmol/µl of ACTH 18-39, m/z 2465.199, were added to the matrix solution as internal standards. For the analysis, 0.2 µl of sample was mixed with 0.2 µl matrix solution on the MALDI target plate. A dilution series of the protein sample was applied to the MALDI plate and intact weight determination was made in linear mode. For the peptide mass fingerprint analysis, a rapid digestion (60 minutes) using sequence-grade trypsin (Promega) at 1:20 and 1:50 trypsin:protein ratio (w/w) was performed. 50% acetonitrile in 25 mM NH4HCO3 was used as the digestion buffer. The peptide mass fingerprint analysis was performed using the reflector mode of the instrument and subsequent analysis of peptides containing internal sulfur bridges was performed in MS/MS mode. Detailed information about digestion pattern and analysis is given in Supplementary data (Figures S9 and S10).

ASSOCIATED CONTENT Supporting Information ThT fluorescence microplate control experiments without SOD1; analytical gel filtration chromatogram;

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N-1H HSQC spectra; ThT fluorescence microplate fibrillization assays

on disulfide-oxidized pwt-SOD1 without any additives; replicates of ThT fluorescence microplate fibrillization assays on pwt-SOD1 and SOD1∆C at different concentrations of reducing agent (TCEP), protein, or urea; control experiments of ThT fluorescence over time; MS spectra of pwt-SOD1 and SOD1∆C; Tables summarizing experimental conditions and results (PDF).

AUTHOR INFORMATION

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Corresponding author e-mail: [email protected] Author Contributions SL and MA designed the research. MAIK, MR, and SK performed experiments. MAIK, ME, SK, and MA analyzed the data. All authors contributed to writing the manuscript. Funding This research was supported by The Swedish Research Council (621-2010-2478) and the Göran Gustafsson Foundation for Research in Natural Sciences and Medicine (MA). MR was supported by a postdoctoral research fellowship awarded by VR (623-2009-800). MAIK was supported by the European Commission via the Erasmus Mundus Europe Asia scholarship program.

Acknowledgments We thank Risto Cukalevski for assistance with microplate experiments and Kristine Steen Jensen for helpful comments on the manuscript.

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Zn Superoxide Dismutase Forms Amyloid Fibrils under Near

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