Self-Assembly of Ovalbumin Amyloid Pores: Effects on Membrane

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Self-assembly of Ovalbumin Amyloid Pores: Effects on Membrane Permeabilization, Dipole Potential and Bilayer Fluidity

Mily Bhattacharya* and Priyanka Dogra

Department of Chemical Sciences, Indian Institute of Science Education and Research (IISER), Mohali, Knowledge City, Sector 81, S.A.S. Nagar, Mohali-140306, India. * Corresponding author email: [email protected]

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Abstract: Amyloid assembly is inherently a stochastic and a hierarchical process comprising the genesis of heterogeneous, transiently-populated prefibrillar aggregates that are characterized to be non-native oligomeric conformers. These oligomers could be either off-pathway or onpathway species enroute to amyloid fibrils that are associated with a variety of neurodegenerative disorders namely, Alzheimer’s, Parkinson’s, prion diseases as well as in localized and systemic amyloidoses (type II diabetes and dialysis-related, respectively). Morphological characterizations of these prefibrillar aggregates indicated that apparently, the doughnut or annular structure is commonly shared among various prefibrillar species irrespective of the diverse native structures and aggregation mechanisms. In this work, we have elucidated the self-assembly mechanism of amyloid pore formation from ovalbumin using a range of biophysical techniques that shed light into the time-dependent protein structural changes as aggregation progressed. Additionally, based on several evidences suggesting amyloid pore-mediated cytotoxicity, we have investigated the annular amyloidmembrane interaction using a comprehensive biophysical approach. The influences of annular pores on the intramembrane dipole potential and bilayer fluidity, as a consequence of membrane permeabilization, were examined in a protein concentration- and time-dependent manner that provided important insights into the pore-membrane interactions. An instantaneous membrane permeabilization kinetics suggested that plausibly a detergent-like carpet mechanism during membrane disruption was effective. Moreover, it was inferred that a loss in the membrane integrity resulted in the generation of both disordered lipid and disoriented water dipoles that reside at the immediate vicinity of the membrane bilayer. These key findings may have implications in amyloid pore induced deleterious effects during amyloid-membrane interactions. 2


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1. Introduction Protein conformational diseases initiating from an unrestrained self-assembly followed by a deposition of the misfolded protein aggregates and amyloids are implicated in a variety of physiological disorders such as Alzheimer’s, Parkinson’s, prion diseases as well as in systemic amyloidosis.1-8 The initial pivotal step in amyloid assembly is considered to be the formation and accumulation of partially-unfolded intermediate state(s) of a protein, as a result of mutation in the amino acid sequence of the polypeptide chain or changes in solution conditions such as pH, ionic strength and temperature.4, 9-12 The fact that nearly every protein can access the amyloid state, whether they are disease-associated or not, indicates that protein aggregation is a generic phenomenon and is likely to be independent of the native structure.13 However, it has been documented that the amino acid sequence and environmental conditions have a significant impact on the fibrillation kinetics, fibril morphology and the architecture along with the fibril stability.13 Amyloid fibrillation is inherently a hierarchical and a stochastic process during which generation of a multitude of heterogeneous, transient and non-native oligomeric conformers occurs that serve as amyloidogenic precursors.9, 14-18 These oligomeric species or prefibrillar aggregates could be either on-pathway or off-pathway entities during amyloid assembly and are considered to be more cytotoxic owing to their ability to interact with plasma membranes.19-23 For example, recent reports on α-synuclein oligomers suggest that the “small” doughnut-shaped oligomers are off-pathway species and exhibit relatively higher cytotoxicity compared to another type of coexistent, transientlypopulated “large” oligomers that eventually transform into amyloid fibrils.24 Moreover, it is now well-recognized that although the mechanism of polydisperse prefibrillar aggregate formation could be diverse, the annular pore-like morphology is one of the most common features shared by the prefibrillar oligomers as indicated by AFM and TEM images.25-27 3


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In recent years, many studies have proposed that among several divergent modes of amyloidinduced cytotoxicity, annular pore-mediated membrane disruption is also of primary significance.28-33 The emerging hypotheses from these studies state that membrane disruption may occur by either one or a combination of two broadly classified mechanisms.21, 28, 34-36 According to the “barrel-stave” mechanism, the annular protofibrils form discrete ion channel-type pores within the membrane resulting in localized, non-specific ruptures of the bilayer as suggested by marker-selective dye release assays and ion conductance measurements. In contrast to membrane poration, another mode of membrane disruption known as “detergent-like carpet” mechanism could also compromise the membrane integrity. In this scenario, the amyloid aggregates act as surfactants and cover the membrane surface that results in a reduction in the membrane surface tension. This, in turn, plausibly allows the extraction of lipids from either one or both the leaflets leading to membrane fragmentation and bilayer thinning that has been observed in amyloid oligomers derived from islet amyloid polypeptide (IAPP), Amyloid-β, α-synuclein, β2-microglobulin etc.22, 37-39 However, owing to inherent complexity in the elucidation of polydisperse, transiently-populated oligomeric structures coupled with various physico-chemical determinants underlying amyloidmembrane interactions, accurate analysis and assignment of a particular mechanism still remains a challenging task. Here, we have used a model protein-membrane system and a comprehensive biophysical approach to address issues related to alterations in various structural features of model membranes such as bilayer fluidity and dipole potential upon addition of protein amyloid pores that exhibited membrane permeabilization. We have used ovalbumin, a 385-residue, 45 kDa glycoprotein comprising nine α-helices and three β-sheets that harbor three tryptophans (Trp 148, Trp 184, Trp 267), four free cystines (Cys 11, Cys 30, Cys 367, Cys 382), and one 4


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solvent accessible disulfide bond (Cys 73-Cys 120) (Figure 1a). Ovalbumin belongs to the serpin superfamily in which misfolding and aggregation of neuroserpin has been reported to result in neuronal cell death leading to Alzheimer-like dementia.40 Several reports also indicate that co-localization of neuroserpin aggregates along with the Aβ aggregates is quite prevalent in the senile plaques isolated from patients diagnosed with Alzheimer’s disease.40 Additionally, it has been reported that ovalbumin forms amyloid aggregates under certain conditions.41-44 It has also been documented that ovalbumin forms nanoscopic amyloid annular pores in a hierarchical fashion and a wealth of structural insights into these higherorder supramolecular pores were obtained at the molecular level from Raman spectroscopic measurements.42 In the present study, we have carried out a detailed mechanistic investigation on ovalbumin aggregation using a variety of biophysical tools such as steady-state and time-resolved fluorescence spectroscopy, circular dichroism and multi-angle light scattering. The multiple structural probes were used in-tandem to investigate the kinetics of conformational- and size changes during the process of amyloid assembly with an emphasis on delineating the early key steps. A plausible mechanism of ovalbumin aggregation was proposed based on our kinetic analysis. Additionally, interactions between ovalbumin aggregates and synthetic lipid vesicles were examined using a variety of fluorescence assays in both time- and protein concentration-dependent manner. These experiments provided important insights into the changes in the membrane structural parameters such as, lipid orderedness and fluidity of the acyl chains, as a consequence of vesicle permeabilization.

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2. Results An investigation into the nanoscale morphological transitions of ovalbumin aggregates by AFM revealed a time-dependent increment in the population and dimensions of supramolecular, higher-order annular pores (Figure 1b, c) as reported earlier by us.42 The following sections describe monitoring the alterations in protein structure as a function of time using a probe-dependent approach and the subsequent effects of protein aggregates on model membranes. The kinetic parameters of amyloid aggregation were deduced and analyzed by fitting the rates of changes of the spectroscopic read-outs using a singleexponential function.

2.1 Kinetics of conformational- and size changes As mentioned previously,42 an enhancement in the Thioflavin-T (ThT) fluorescence emission (Figure 2a) confirmed that ovalbumin formed β-sheet-rich amyloid aggregates at 65 °C with an average rate constant of ~114 x 10-3 min-1 Additionally, the far-UV CD spectra of ovalbumin showed an evolution of β-sheets at the expense of α-helices and the average rate constant was estimated to be ~80 x 10-3 min-1 (Figure 2b). The intrinsic (tryptophan) fluorophore was utilized to study the ovalbumin conformational- and size-changes as aggregation progressed. The tryptophan fluorescence intensity, monitored at 350 nm, increased slightly during the aggregation with no apparent shift in the emission maxima (Figure S1a; Supporting Information). On the other hand, the tryptophan fluorescence anisotropy (eq. 1; Supporting Information) showed a progressive increase as a function of time (Figure 2c) indicative of an overall size growth of aggregates and the average rate constant was extracted to be ~ 238 x 10-3 min-1. A small but measureable increase in the tryptophan fluorescence anisotropy was observed prior to heating of the sample (0 min), 6


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suggestive of a plausible formation of soluble oligomers under aggregating conditions. This was independently verified by monitoring the tryptophan fluorescence anisotropy as a function of ovalbumin concentration that confirmed the formation of oligomers at concentrations ≥50 µM (Figure S1b). Thus, from the tryptophan steady-state fluorescence kinetics, it was inferred that large-sized aggregates are formed during the aggregation event. Next, 1,8-anilinonaphthalene-sulfonic acid-ammonium salt (ANS) was used as a hydrophobic reporter to monitor the changes in hydrophobicity during the course of aggregation. Free ANS in aqueous solution fluoresces weakly at ~515 nm whereas a significant enhancement in the ANS fluorescence intensity is observed upon binding to hydrophobic pockets with a concomitant blue shift to ~475 nm.45 In our study, we observed a shift of ~8 nm (from ~475 to ~467 nm) in the ANS fluorescence spectra as a function of time (Figure S1c; Supporting Information). A progressive increase in both the ANS fluorescence intensity (at 475 nm; Figure 2d) and anisotropy (Figure 2d inset) were observed and the apparent rate constants were estimated to be ~423 and ~233 x 10-3 min-1, respectively. Also, prior to heating of the sample, the ANS fluorescence anisotropy showed a substantial increase (~0.15) compared to that of the monomer (~0.12) indicating the formation of soluble oligomers and hence, corroborated the tryptophan fluorescence anisotropy results. Taken together, the ANS fluorescence data indicated that the molten-globule state of ovalbumin formed soluble oligomers, mediated by hydrophobic collapse, which upon heating led to the formation of amyloid aggregates with enhanced hydrophobicity and size growth. Comparison between the ANS fluorescence intensity and anisotropy kinetics revealed that the conformational change occurred twice as faster than the overall size growth. An overall comparison of the ThT fluorescence and tryptophan as well as ANS fluorescence kinetics suggests that during ovalbumin aggregation, the cross-β amyloid binding sites evolve at a much slower rate 7


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compared to the protein conformational-and size changes.

2.2 Dynamics and dimensions of ovalbumin aggregates In order to gain further insights into the dynamical nature of the ovalbumin aggregates, picosecond time-resolved fluorescence anisotropy decay measurements were carried out. Time-resolved fluorescence anisotropy decay measurements allow one to separate the local and the global dynamics of a fluorophore that are otherwise unresolved in steady-state fluorescence anisotropy measurements due to time-averaging.45-46 The local and global dynamics can be distinctly identified by two different rotational correlation times φfast and φslow, respectively (obtained from the decay analysis; eq. 2-4; Supporting Information). The slow rotational correlation time, φslow, is attributed to the global dynamics that is commonly associated with the hydrodynamic volume and hence, size of the protein (eqs. 5,6; Supporting Information).46 Prior to the anisotropy decay measurements, the fluorescence lifetime of tryptophans was measured at different time-points and three lifetime components were recovered. The mean fluorescence lifetime (~2.3 ns) of tryptophans was observed to be constant at all stages of ovalbumin aggregation (Figure S2; Supporting Information). Next, the fluorescence lifetime of ANS (added to the aggregates at various time intervals) was measured as aggregation progressed and again, three lifetime components were extracted. Comparison of the mean fluorescence lifetime of ANS revealed an increase from ~11 ns to ~15 ns during the course of aggregation (Table 1 and Figure S3). A closer inspection revealed that the longest lifetime component (τ3) of ANS increased from ~17.1 ns to ~18.2 ns whereas the short lifetime components (τ1, τ2) remained almost constant (see Table 1). The amplitude (α3) associated with τ3 increased significantly at the expense of the amplitudes of the shorter lifetime components. Following these observations, we carried out time-resolved 8


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fluorescence anisotropy decay measurements only on ANS (Figure 3a-c and Table 1) since ANS has a much longer fluorescence lifetime (> 10 ns) compared to tryptophan when bound to hydrophobic pockets.45 The fast and slow rotational correlation times for the initial (preincubated) oligomeric species were analyzed to be ~0.3 and ~43 ns, respectively and the average hydrodynamic radius was estimated to be ~3.6 nm that corroborated our AFM results.42 For the aggregates formed after 15 minutes of heating, the fast and slow rotational correlation times were recovered to be ~0.2 and >200 ns, respectively and the average hydrodynamic radius was determined to be ~6.6 nm thus, clearly indicating an increase in the overall aggregate size. After 1 hour of heating, the recovered fast and slow rotational correlation times were ~0.2 and >300 ns, respectively and the average hydrodynamic radius was estimated to be >6.9 nm. Interestingly, an overall comparison of the amplitudes (βfast; Table 1) associated with the fast rotational correlation time of ANS demonstrated a significant drop from ~0.6 to ~0.2 suggesting that the local tumbling of ANS became more restricted as aggregation progressed. These results indicated that the hydrophobic core becomes more tightly packed upon amyloid assembly. Therefore, time-resolved fluorescence anisotropy measurements provided insights into both structural packing and overall size growth during the amyloid formation. We would like to mention here that during the later stage of aggregation, one would expect an even larger increase in the average hydrodynamic size that cannot be accurately estimated by fluorescence anisotropy decay since the overall rotational correlation time is much longer than the mean fluorescence lifetime. Therefore, a light scattering based technique for size estimation was utilized to shed light into the changes in aggregate size in the later stage of aggregation.

2.3 Insights into the changes in size and molar mass during assembly Next, we carried out multi-angle light scattering (MALS) experiments to determine the 9


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growth kinetics of aggregates. Unlike traditional dynamic light scattering experiments, MALS offers unique capabilities to directly estimate the average molar mass of the aggregates without requiring any pre-assumption of the aggregate shape (globular, rod-like etc.).47 Estimation of the average molar mass reveal an average number of protein monomers that are involved in the aggregate formation. Additionally, the average hydrodynamic radii of the aggregated species could be extracted from the MALS data analysis at every time-point. Figure 3d,e show the changes in the average hydrodynamic radii and average molar masses of the aggregated species, respectively as a function of time. The average hydrodynamic radius of the oligomeric species was ~3 nm (Figure 3d) which increased to ~8 nm after heating the sample for 15 minutes hence, suggestive of an increase (~2.6-fold) in the size due to aggregation and corroborated the time-resolved anisotropy data. Upon heating the sample further, the hydrodynamic radii attained a plateau and largely remained unaltered at a value of ~20 nm. This suggested that the soluble oligomers associated to form large-sized aggregates upon heating. However, prolonged heating of the sample did not show any further change in the hydrodynamic radii. Kinetic analysis of the average hydrodynamic radii data revealed an apparent rate constant of ~27 x 10-3 min-1. Figure 3e depicts the change in the average molar masses of the ovalbumin aggregates as a function of time. After 15 minutes of heating, the average molar mass of the aggregates was observed to be ~900 kDa which is ~20-fold higher than that of the monomeric ovalbumin (~45 kDa). This indicated that on an average, ~20 monomeric units of ovalbumin were present in the aggregates. A progressive increase in the average molar mass was observed as a function of time that revealed the formation of large-sized aggregates with an association of ~90 monomers after 2.5 hours of heating. The apparent rate constant of the average molar mass kinetics was recovered to be ~15 x 10-3 min-1 which is almost half of that observed for 10


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the hydrodynamic radii kinetics. This implies that at longer timescales, the change in average hydrodynamic radii becomes insensitive whereas, the change in average molar mass is observed as long as the protein monomers add onto the aggregates. Thus, the average molar mass kinetics appears to be slower than the kinetics of hydrodynamic radii.

2.4 Comparison of the rates observed from different spectroscopic read-outs An overall comparison of the ovalbumin aggregation kinetics monitored by various structural probes (Figure 3f) revealed that the molten-globule oligomers undergo rapid hydrophobic collapse and conformational changes preceding the overall size growth of the amyloid pores that are in good agreement with the results obtained from our Raman spectroscopy and AFM measurements.42 However, the evolution of cross-β binding sites occurs at a slower rate compared to the size growth. We would also like to point out that though the aggregation depicts an apparent completion within few hours (as demonstrated by steady-state fluorescence and CD), the average molar mass of the aggregates continue to grow (MALS) implying that the growth of aggregates occurs albeit at a much slower rate.

2.5 Ovalbumin-membrane interactions: Insights into membrane permeabilization, intramembrane dipole potential and acyl chain fluidity Several reports in the literature suggest that amyloid pores cause cytotoxicity by virtue of their ability to disrupt lipid membranes.20-21,

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Therefore, in order to discern whether the

ovalbumin annular pores have the potential to disrupt model membranes, several assays were employed to probe amyloid pore mediated liposomal membrane permeabilization and to gain insights into the subsequent effects on the lipid orderedness as well as on the acyl chain fluidity of the constituent lipids. For the following studies, we chose the zwitterionic lipid 111


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palmitoyl-2-oleoyl-sn-glycero-3-phosphatidylcholine (POPC) as a model membrane system since it is a predominant constituent of biological membranes and is chemically inert.48 2.5.1. Membrane permeabilization studies To probe membrane permeabilization, the well-known calcein release assay49-50 was utilized whereby POPC large unilamellar vesicles (LUVs) were prepared encapsulating a very high, self-quenching concentration of calcein (40 mM; see Experimental Section; Supporting Information). Upon vesicle permeabilization, calcein is released and therefore, an enhancement in its fluorescence intensity is observed due to relief in self-quenching. In our study, an instantaneous increase in calcein fluorescence was observed upon addition of ovalbumin aggregates formed at various time intervals. Figure 4a shows one of the representative calcein release kinetic traces although the efficiency of calcein release was less than 100% (even at 1:1 protein:liposome molar ratio and upon prolonged incubation) when compared to that observed upon addition of 1% (v/v) Triton X-100 to the liposomes (Figure S4a; Supporting Information). This observation is consistent with other reports on amyloid aggregates (protofibrils and/or pores) derived from physiologically relevant, disease-related proteins

namely,

β2-microglobulin,51

α-synuclein,52

etc.

although

the

reason

is

incomprehensible. It is presumed that either a fraction of liposomes are more susceptible to disruption or a plausible fusion/coalescence of liposomes occurs, upon addition of protein aggregates, which leads to a reduced liposome disintegration by the aggregates.51 Additionally, the calcein fluorescence attained saturation quickly and remained unchanged even at prolonged timescales. The extent of calcein release enhanced as a function of aggregation states formed at various time points presumably due to an increase in the population of higher-order supramolecular annular pores (as observed by AFM previously)42 which saturated after 4 hours (Figure 4b). Next, in order to discern the minimum 12


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concentration of protein aggregates required for vesicle disruption, the calcein release was monitored as a function of protein concentrations (from 50 nM to 20 µM) at each aggregation state (i.e. at different time-points) while keeping the liposome concentration constant. We observed that the protein concentration of 50-100 nM, at all time-points, did not exhibit any disruptive effect (Figure S4b; Supporting Information) even upon prolonged incubation. For the 15 minute aggregate sample containing a heterogeneous mixture of annular pores and worm-like fibrils,42 the minimum concentration that showed small but significant disruption of calcein encapsulated LUVs was 1 µM (Figure 4c) and the calcein release efficiency (~2.5%) was comparable to that exhibited by both the native ovalbumin and the 0 minute aggregation sample (prior to incubation at 65 ºC) at 10 µM. For aggregate samples withdrawn at later time-points, the optimum concentration for calcein release was 250 nM although the extent of calcein fluorescence varied as a function of time as well as with increasing protein concentrations (Figure 4c, d). It was also observed that the highest concentration of protein (20 µM), formed at later stages of aggregation, exhibited the most effective LUV disruption with a calcein release efficiency of ~43%. However, as expected, the native ovalbumin and the 0 min aggregate sample did not show any variation in calcein efflux efficiencies as a function of concentration. 2.5.2. Insights into bilayer acyl chain fluidity Next, we embarked upon assessing alterations in lipid acyl chain fluidity upon amyloid poremembrane interactions, which is an important structural feature of membranes and plays a significant role in the proper functioning of membrane proteins.53-54 For these experiments, we monitored the changes in steady-state fluorescence anisotropy of a lipophilic probe namely, 1,6- diphenyl-1,3,5-hexatriene (DPH) that was incorporated into the LUVs during liposome preparation (Experimental Section; Supporting Information). DPH anisotropy has 13


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been extensively utilized to monitor the acyl chain dynamics as well as packing order density.55-57 Additionally, insights into the variations in membrane fluidity upon addition of amyloid aggregates derived from physiologically relevant proteins viz. Aβ,58 transthyretin (TTR),59 α-synuclein60 and model proteins namely, lysozyme61 were obtained that reported a few inconsistencies. For example, Aβ peptides were demonstrated to decrease the bilayer fluidity whereas α-synuclein and TTR were shown to increase the fluidity while lysozyme fibrils did not show any changes. In our study, we observed DPH fluorescence anisotropy of ~0.1 prior to the addition of aggregates to DPH-POPC liposomes. After adding ovalbumin amyloids (protein concentration of 10 µM) formed at various time-points, we observed a rise in the anisotropy that attained a plateau (~0.27) plausibly due to an increase in liposomal chain rigidity and packing density. As expected, both the native and the 0 min ovalbumin sample did not demonstrate any constraining effects on the acyl chain dynamics and hence, the DPH anisotropy remained constant at ~0.1 (Figure 5a). Monitoring the changes in DPH anisotropy at varying protein concentrations (250 nM-20 µM) whilst keeping the liposome concentration constant exhibited an interesting trend (Figure 5b, c & Figure S5a-d). Figure 5b shows DPH anisotropy kinetics traces upon adding various concentrations of ovalbumin formed after 4 hours. As evident from the plots, the protein concentrations of 250 nM (data not shown) and 500 nM did not show any increase in the DPH anisotropy. A very small increase in the DPH anisotropy (~0.13) was observed for 1 µM protein concentration which showed a progressive and significant increase (from ~0.13 to ~0.32) at higher protein concentrations. The highest anisotropy value of ~0.32 observed during ovalbumin-POPC interactions is similar to that observed for POPC (r = 0.33) in its gel phase.62 Also, the timecourse of DPH anisotropy measurements indicated that the anisotropy attained a plateau within 10-15 minutes (Figure S5a-d; Supporting Information) and reached similar end-points (Figure S5e) irrespective of protein concentrations at later aggregation stages. However, 14


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based on our results gathered from calcein release experiments, these DPH anisotropy data are quite surprising since one would expect an increase in the chain fluidity as a result of vesicle disruption. In order to probe whether upon vesicle rupture, the displaced hydrophobic DPH translocated into the amyloid pores at different aggregation states, changes in DPH anisotropy as a function of ovalbumin aggregation were monitored. In these experiments, 500 nM of DPH (same concentration as that of in DPH-POPC LUVs) was added into the aggregates (protein concentration: 10 µM) at various time-points. The anisotropy of free DPH was ~0.09. In the presence of aggregates, the DPH anisotropy sharply enhanced to ~0.33 (0 min sample) and increased further to ~0.38 (15 min sample onwards; data not shown) that is closer to its reported limiting anisotropy value.63 Comparison of DPH anisotropies for the 0 min sample in the presence and absence of liposomes at a constant protein concentration revealed that the anisotropy was lower (~0.1) in the presence of vesicles whereas it was higher (~0.33) in the absence of vesicles. Interestingly, such lower DPH anisotropy correlates well with the calcein permeabilization efficiency of the 0 min sample (~2.5%; Figure 4b) which implies that due to very low disruption potency, the plausible displacement of embedded DPH from the POPC LUVs followed by its translocation into oligomers does not occur. Hence, taken together, we speculate that the rise in DPH anisotropy as a consequence of pores-induced liposome disintegration could be attributed to one or both of the following factors. Since the supramolecular amyloid pores are hydrophobic, it is very likely that the vesicle-embedded lipophilic DPH could get displaced from the disintegrated membrane interior and move into the hydrophobic core of ovalbumin aggregates and/or DPH could get simultaneously inserted between the co-aggregates formed by ovalbumin pores and fragmented POPC.

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2.5.3. Insights into intramembrane dipole potential After establishing that the ovalbumin amyloid annular pores compromise membrane integrity, next we probed the ordered orientation of the constituent zwitterionic POPC lipid molecules, upon addition of ovalbumin amyloid pores, by monitoring the changes in dipole potential of the POPC LUVs using a potentiometric/voltage-sensitive fluorescent dye namely, 4-(2-[6-(Dioctylamino)-2-naphthalenyl]ethenyl)-1-(3-sulfopropyl) pyridinium salt (di-8ANEPPS) that was incorporated into the LUVs during vesicle preparation (see Experimental Section; Supporting Information). The dipole potential (ΨD) of a membrane denotes the potential difference inside the membrane bilayer which is an indicator of the orientation of the lipid dipoles (polar head groups and ester linkages of the acyl group to the glycerol backbone) within the membrane and confined water dipoles at the membrane interface (~3 nm thick).64 It affects the translocations of various ions and macromolecules across the membranes as well as influences the structure and functions of membrane proteins. Since the dipole potential is operative at a very small distance of a membrane bilayer, it gives rise to a significantly strong electric field of 108-109 V/m and any change in the dipole potential results in a shift in the absorption and emission spectra of the di-8-ANEPPS dye that is located at the membrane interface (~1.2 nm from the centre of the membrane bilayer).65 Hence, based on the results obtained from calcein leakage assay, we hypothesized that upon addition of ovalbumin amyloid pores, a drop in the dipole potential would be observed owing to aggregate binding-induced vesicle disruption and subsequent random orientation of the constituent dipolar lipids and water molecules. In order to monitor the dipole potential, the dual-wavelength fluorescence ratiometric approach was utilized66-67 (see Experimental Section; Supporting Information) to eliminate the effects of photobleaching and variation in the dye concentration. Recently, it has also been reported that the fluorescence ratio (R) of di16


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8-ANEPPS is only dependent on the membrane dipole potential and is not influenced by “specific molecular interactions”.68 Additionally, care was taken to record the fluorescence intensity at the red-edge (670 nm) of emission spectra to exclude any effects of membrane acyl chain fluidity/viscosity on the measured dipole potential.69 Utilization of di-8-ANEPPS fluorescence to monitor the effects of fibrillar proteins on model membranes has been reported earlier.70 In our study, addition of ovalbumin aggregates (at 10 µM protein concentration) indeed resulted in a reduction in the dipole potential with a sharp drop at the 15 min aggregate sample (Figure 6a). As aggregation proceeded further, a gradual yet significant reduction in the dipole potential was observed that saturated after 4 hours. The latter observation suggested that the vesicle disintegration with a subsequent randomization of the dipolar lipids and water dipoles were influenced by the time-dependent enhancement in population as well as an increment in the pore sizes hence, corroborating the calcein release data. Next, the reduction in the dipole potential was investigated in a protein concentrationdependent manner (250 nM–20 µM) at various stages of aggregation. Figure 6b, c represent comparative changes in the dipole potential as a function of concentration for samples prior to incubation (0 min) at elevated temperature and at different time-points of aggregation. It appears that at nanomolar protein concentration for all time-points, the dipole potential dropped marginally plausibly, due to a very low liposome disruption efficiency. For protein concentrations ≥1 µM, a small but measurable drop in the dipole potential was observed for the 0 min sample whereas quite a significant reduction in ΨD was measured with an advancement of the aggregation stages. However, it is to be noted that for the 0 min sample at concentrations ≥5 µM, changes in the ΨD exhibited a slight protein concentration dependence unlike the calcein release experiments. This could, probably, be attributed to the displacement of water molecules upon binding of the initial soluble oligomers to the liposomal surface, 17


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thus resulting in random orientations of water dipoles. Taken together, our calcein efflux and dipole potential measurements indicated that upon addition of ovalbumin oligomers and annular pores to the liposomal membranes, binding of the aggregates coupled with a vesicle rupture occurred. Consequently, the lipid and/or water dipoles disoriented which was manifested as changes in the di-8-ANEPPS fluorescence located at the membrane interface and the encapsulated calcein was released from the aqueous interior of the ruptured vesicles.

3. Discussion 3.1 Ovalbumin forms amyloid aggregates at low pH The outcomes obtained from all the fluorescence read-outs, CD and MALS in the present study along with the previously reported Raman spectroscopy and AFM measurements42 confirmed that the partially-unfolded, molten-globule conformer of ovalbumin forms cross βsheet-rich amyloid annular pores in the presence of salt at elevated temperature. It was observed that ovalbumin amyloid aggregation proceeds without any lag phase and apparently gets completed within few hours which is in line with a previous report.71 The fluorescence anisotropy measurements along with the MALS data indicated the formation of spherical oligomers which was also demonstrated earlier by AFM.42 This is identified to be one of the early key steps in ovalbumin aggregation that is similar to the initial dimer and/or trimer formed by an aggregation-prone intermediate, as has been demonstrated using single molecule fluorescence techniques during neuroserpin aggregation.72 However, the CD spectra of the oligomers did not show any significant conformational change compared to the monomeric molten-globule73 which is in agreement to that reported earlier.74 Therefore, these oligomers

represent

conformationally

unconverted,

aggregation-competent

soluble

aggregates. As the temperature was raised to trigger the aggregation, a rise in the β-sheet 18


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content at the expense of α-helices was observed by both CD and Raman spectroscopic measurements.42 Also, fluorescence anisotropy, average hydrodynamic radii and molar mass measurements suggested an increase in the aggregate size. Hence, based on our overall kinetics data, we propose that raising the temperature facilitates the formation of disordered regions within these oligomers that undergo conformational rearrangement from a predominantly α-helical structure to a β-sheet-rich amyloid annular pores that precede the overall size growth of aggregates to large pores. Additionally, we found that the increase in average molar mass and hydrodynamic radii of the ovalbumin aggregates are correlated by a power-law with an exponent of 2.2 that indicated the formation of random polymers by “endto-end association” and plausibly, absence of any secondary aggregation mechanisms, similar to that observed for neuroserpin polymerization.75

3.2 Insights into hydrophobicity and hydrogen bonding interactions within amyloid annular pores In many instances of protein aggregation and amyloid assembly, it has been demonstrated that both electrostatic and hydrophobic interactions play a very vital role in promoting aggregation.76-77 Ovalbumin has an isoelectric point of ~4.5 which acquires a net charge of +43 at pH 2.2. At higher protein concentration and upon thermal incubation, addition of salt screens the electrostatic repulsions between the positively charged polypeptide chains and favours hydrophobic association of the thermally-unfolded regions that drives the aggregation forward. In our study, a sharp increase in the ANS fluorescence intensity indicated the formation of ovalbumin aggregates with enhanced hydrophobicity. Additional evidence favouring an increase in hydrophobicity and a possible sequestration of tryptophans into the hydrophobic regions was provided by the Raman spectra of aggregates that showed an 19


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increase in the tryptophan hydrophobicity ratio (I1360/I1340) and a modest weakening of hydrogen bonding strength between the tryptophanyl indole ring and the surrounding water molecules.42

3.3 Implications of ovalbumin pores-membrane interactions on annular pore-induced cytotoxicity According to several reports, the annular pores and protofibrils are considered to be more cytotoxic than the mature amyloid fibres since they permeabilize cell membranes by a host of putative mechanisms.19 Recently, annular protofibrils have been isolated from brain samples diagnosed with Alzheimer’s disease, hence reasserting the cytotoxicity of the doughnutshaped protofibrillar aggregates.38 In the present study, we have investigated the membrane disruption efficiency induced by the ovalbumin supramolecular annular pores using a variety of fluorescence assays that reported the changes in various structural features of the liposomal membranes. It is also known that monomeric ovalbumin does not bind to POPC LUVs.78 Calcein is encapsulated into the aqueous core of the liposomal bilayer, DPH is inserted into the hydrophobic core formed by the lipid acyl chains and di-8-ANEPPS is located at the membrane interface where a chemically heterogeneous environment encompassing water molecules, lipid head groups etc. prevails. Results from the calcein leakage experiments suggested that with an advancement in aggregation, ovalbumin amyloid pores disrupted liposomal membranes with an enhancing potency that could be attributed to the time-dependent evolution of large, increasingly/predominantly populated higher-order annular pores. An instantaneous membrane permeabilization kinetics was observed which implied that probably, the annular pores disrupted the membranes by a detergent-like carpet mechanism similar to that observed in anti-microbial peptides-induced membrane activity though the structural determinants for membrane impairment are quite varied.79-80 The DPH 20


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anisotropy measurements gave rise to two major alternate possibilities in the interpretation of acyl chain fluidity. Though an increase in DPH anisotropy was observed upon adding pores into the vesicles which could be assigned to an increase in the chain rigidity, the probability of translocation of DPH from the bilayer interior into the hydrophobic core of the pores and/or insertion of DPH within protein-and-fragmented lipid co-aggregates cannot be ruled out in light of few recent reports.71-72 Recently, while monitoring the effects of Aβ aggregates on membranes using various spectroscopic techniques, a similar increase in DPH steady-state fluorescence anisotropy was observed that was ascribed to the translocation of non-covalently bound DPH from DPPC liposomes into Aβ aggregates.81 Studies using small angle X-ray scattering (SAXS) and two-photon fluorescence microscopy suggested the presence of soluble

co-aggregates

of

fibrillar

α-synuclein

and

lipid

vesicles

upon

vesicle

permeabilization.82 In other reports, extraction of lipids from the outer membrane leaflet of distorted, blebby liposomes by amyloid fibrils has been hypothesized to be one of the key mechanisms of liposome disintegration.22, 37 Membrane dipole potential (ΨD) is another very crucial structural determinant of changes in the membrane structural integrity which was reported by interfacially-positioned di-8-ANEPPS (located at ~1.2 nm from the bilayer centre)65 in the present study. ΨD denotes the potential difference within the membrane bilayer which arises due to variation in the orientations of interfacial water dipoles and the lipid molecular dipoles. Recently, MD simulations suggested the presence of a “mosaic of water orientation structures” at POPC bilayer/water interface83 that has been experimentally shown to comprise three distinct hydrogen-bonded water structures.84 As discussed earlier, an expected reduction in the dipole potential was observed that was ascribed to the disruptionmediated randomization of the constituent lipid and confined water dipoles. Additionally, penetration of water molecules has been proposed to be responsible for the decline in ΨD.65 Therefore, we propose that binding of annular pores onto the liposomal surface probably 21


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disrupts/weakens the hydrogen-bonded water network at the membrane interface and displaces water molecules that penetrate upon membrane permeabilization. Moreover, loss in the membrane integrity leads to an isotropic orientation of the lipid dipoles and all of these factors result in the observed loss in potential. In summary, we have delineated the cascade of molecular events that eventually lead to the formation of higher-order amyloid pores with a hydrophobic core and a high β-sheet content. Results obtained from the membrane activity studies provided some insightful clues into the pore-mediated membrane disintegration. We believe that a comprehensive biophysical approach, used in this study on model protein-membrane systems, would be useful in deciphering detailed molecular mechanism of protein amyloid-membrane interactions that would provide key insights into amyloid aggregates-induced deleterious effects.

4. Experimental Section For setting up ovalbumin aggregation reaction, we used our previously reported protocol.42 Various structural probes were utilized to monitor the aggregation kinetics and all the experimental as well as data analysis details are provided in the Supporting Information. Also, for all the experimental details of preparation of a variety of liposomes, monitoring ovalbumin-vesicle interactions, see the Supporting Information.

5. Acknowledgements We thank the members of the Mukhopadhyay lab for reading the manuscript critically. M.B. thanks the Department of Science & Technology (DST), New Delhi for the Women Scientist grant and DST-SERB for the Young Scientist grant. P.D. thanks DST for the INSPIRE fellowship. We are also grateful to Ms. M. Kombrabail, Mr. Satyanarayan and Prof. G. Krishnamoorthy (TIFR Mumbai) for their help with time-resolved fluorescence and MALS 22


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measurements, Prof. N. Periasamy (TIFR Mumbai) for providing us with the time-resolved fluorescence data analysis software and Dr. S.K. Pal (IISER Mohali) for allowing us to use the light scattering instrument. We gratefully acknowledge Dr. Samrat Mukhopadhyay (IISER Mohali) for discussion.

Supporting Information The Supporting Information contains the Experimental Section wherein all the experimental and data analysis details have been described. Also, a few additional figures (Figures S1-S5) are provided in the Supporting Information.

23


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Phospholipid/Cholesterol Membranes: A Theoretical Study. J. Phys. Chem. B 2011, 115, 4160-4167. 69.

Clarke, R.J.; Kane, D.J. Optical Detection of Membrane Dipole Potential: Avoidance

of Fluidity and Dye-Induced Effects. Biochim. Biophys. Acta 1997, 1323, 223-239. 70.

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Amyloid Protein and Phospholipid Membranes Is Dependent on Protein Aggregation State. J. Chinese Inst. Chem. Eng. 2008, 39, 321-328. 71.

Azakami, H.; Mukai, A.; Kato, A. Role of Amyloid Type Cross β-Structure in the

Formation of Soluble Aggregate and Gel in Heat-Induced Ovalbumin. J. Agric. Food Chem. 2005, 53, 1254-1257. 72.

Chiou, A.; Hagglof, P.; Orte, A.; Chen, A.Y.; Dunne, P.D.; Belorgey, D.; Karlsson-Li,

S.; Lomas, D.A.; Klenerman, D. Probing Neuroserpin Polymerization and Interaction with Amyloid-β Peptides Using Single Molecule Fluorescence. Biophys. J. 2009, 97, 2306-2315. 73.

Bhattacharya, M.; Mukhopadhyay, S. Structural and Dynamical Insights into the

Molten-Globule Form of Ovalbumin. J. Phys. Chem. B 2012, 116, 520-531. 74.

Hu, H.Y.; Du, H.N. α-to-β Structural Transformation of Ovalbumin: Heat and pH

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M.; Martorana, V.; Cupane, A.; Bolognesi, M.; Manno, M. The Tempered Polymerization of Human Neuroserpin. PLoS One 2012, 7, e32444. 76.

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Serpell, L.C. Hydrophobic, Aromatic, and Electrostatic Interactions Play a Central Role in Amyloid Fibril Formation and Stability. Biochemistry 2011, 50, 2061-2071. 77.

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Understanding Amyloid Aggregation by Statistical Analysis of Atomic Force Microscopy Images. Nat. Nanotechnol. 2010, 5, 423-428. 78.

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Cullin, C.; Lecomte, S. Interaction of Aβ(1-42) Amyloids with Lipids Promotes "OffPathway" Oligomerization and Membrane Damage. Biomacromolecules 2015, 16, 944-950. 80.

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Suzuki, M.; Miura, T. Effect of Amyloid β-Peptide on the Fluidity of

Phosphatidylcholine Membranes: Uses and Limitations of Diphenylhexatriene Fluorescence Anisotropy. Biochim. Biophys. Acta 2015, 1848, 753-759. 82.

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Protein/Lipid Coaggregates Are Formed During α-Synuclein-Induced Disruption of Lipid Bilayers. Biomacromolecules 2014, 15, 3643-3654. 83.

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a Neutral Zwitterionic Lipid/Water Interface Revealed by Molecular Dynamics Simulations. J. Phys. Chem. Lett. 2014, 5, 4343-4348. 84.

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Structures at a Zwitterionic Lipid/Water Interface Revealed by Heterodyne-Detected Vibrational Sum Frequency Generation. J. Am. Chem. Soc. 2012, 134, 7842-7850.

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Figure and Table Legends: Figure 1. (a) Crystal structure of ovalbumin (PDB ID: 1OVA) generated using PyMol (DeLano Scientific, LLC, CA). The tryptophans, free cystines and the single disulfide bond are shown as magenta spheres, yellow ribbons and yellow stick, respectively. (b,c) Formation of annular pores by ovalbumin showing a progressive increase in the population and dimensions of the doughnut-shaped amyloids after (b) 15 minutes and (c) 4 hours of incubation as observed by AFM.

Figure 2. Aggregation kinetics of ovalbumin investigated at pH 2.2, 50 mM NaCl and 65 ºC using a probe-dependent approach. (a) Evolution of ThT fluorescence (indicated by a black upward arrow) as a function of aggregation. The gray line denotes ThT fluorescence prior to incubation (0 min); the inset graph shows changes in the ThT fluorescence intensity (green filled circles) at 480 nm and the solid black line represents the single-exponential fit. (b) FarUV CD spectra representing the secondary structural changes during aggregation. Far-UV CD spectra representing the secondary structural changes during aggregation. For clarity, a few representative spectra at 0 min (blue), 10 min (violet), 1 hour (green), 1.5 hours (dark cyan), 3 hours (orange) and 4 hours (red) are shown. The inset graph shows a plot of ratiometric ellipticity (θ218/θ222, magenta filled circles) depicting the evolution of β-sheets at the expense of α-helices and the black solid line represents the single-exponential fit. (c) Changes in the tryptophan fluorescence anisotropy (filled circles), monitored at 350 nm, which is fitted by a single-exponential function (solid line) and (d) changes in the ANS fluorescence intensity and anisotropy (inset graph; red filled circles) monitored at 475 nm. The solid line in (d) and the corresponding inset graph represents the single-exponential fit. The fitted lines were used to recover the apparent rate constants. All the experiments were 34


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carried out at room temperature and repeated at least thrice.

Figure 3. Time-resolved fluorescence anisotropy decay profiles (gray scatter plots) of ANS bound to ovalbumin aggregates at (a) 0 min, (b) 15 min and (c) 1 hour. The black solid line in each decay profile represents the bi-exponential fit that was used to recover the rotational correlation times. (d, e) Aggregation kinetics of ovalbumin monitored by multi-angle light scattering technique at room temperature. Changes in the (d) average hydrodynamic radii and (e) average molar mass of aggregates as a function of time. The solid line in each graph represents the fit obtained using single-exponential function that was used to recover the apparent rate constants. (f) Comparison of the observed rate constants of ovalbumin aggregation kinetics obtained from multiple structural probes that were used to investigate protein conformational- and size changes during amyloid assembly.

Figure 4. Ovalbumin pore-induced permeabilization of calcein-encapsulated POPC LUVs. (a) Disruption kinetics of liposomes monitored by calcein fluorescence released from the vesicles. The black arrow indicates addition of amyloid pores (protein concentration: 10 µM), formed after 2 hours, to the POPC vesicles. (b) Changes in the calcein release efficiency as a function of native monomeric and aggregated forms of ovalbumin at a protein concentration of 10 µM at room temperature. The release efficiency is expressed as a percentage of the maximum calcein release observed upon addition of Triton X-100 to the vesicles. The error bars denote the standard deviations obtained from at least three separate measurements. (c) Comparison of the extent of calcein release from POPC LUVs as a function of ovalbumin concentration at early (15 min) and late (4 hours) aggregation stages. The error bars denote the standard deviations obtained from at least three separate measurements. (d) A 3dimensional column plot for comparison of calcein efflux efficiencies as a function of 35


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ovalbumin concentration at various aggregation stages (at different time-points).

Figure 5. (a) Alterations in the acyl chain fluidity of POPC LUVs monitored by changes in DPH fluorescence anisotropy as a function of native monomeric and aggregated forms of ovalbumin at a protein concentration of 10 µM. The error bars denote the standard deviations obtained from at least five separate measurements. (b) Kinetics of changes in the acyl chain fluidity, monitored by DPH fluorescence anisotropy, as a function of ovalbumin amyloid pores (4 hours) in a concentration-dependent manner. The black arrow denotes the addition of protein aggregates. (c) Comparison of the changes in the bilayer fluidity of DPH/POPC LUVs as a function of ovalbumin concentration at early oligomeric (0 min) and late (4 hours) aggregation stages. The error bars indicate the standard deviations obtained from at least five separate measurements.

Figure 6. (a) Changes in the POPC membrane dipole potential (ΨD) as a function of native monomeric and aggregated forms of ovalbumin at a protein concentration of 10 µM, monitored by a voltage-sensitive fluorophore di-8-ANEPPS, at room temperature. The reduction in ΨD is expressed as the differences between the dipole potential of di-8ANEPPS/POPC LUVs in the presence of ovalbumin to that observed in the absence of ovalbumin. The black line is an eye-guide and the error bars denote the standard deviations obtained from at least five separate measurements. (b) Comparison of the extent of dipole potential reduction of POPC LUVs as a function of ovalbumin concentration at oligomeric (0 min) and late (4 hours) aggregation stages. The error bars denote the standard deviations obtained from at least five separate measurements. (c) Changes in the dipole potential monitored as a function of ovalbumin concentration at various aggregation stages (at different

36


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time-points). The black lines serve as eye-guides. All the experiments were repeated at least five times at room temperature.

Table 1. Typical recovered parameters associated with picosecond time-resolved fluorescence intensity and anisotropy decays of ANS in ovalbumin aggregates a a

The errors associated with the mean fluorescence lifetimes and rotational correlation times

(fast and slow) of ANS with their respective amplitudes are: (a) For 0 min (oligomer): τm = 11.49 ± 0.30 ns; φfast = 0.35 ± 0.14 ns, βfast = 0.33 ± 0.01 ns; φslow = 42 ± 1 ns, βslow = 0.67 ± 0.01 ns. (b) For 15 min (aggregate): τm = 15.5 ± 0.01 ns; φfast = 0.27 ± 0.11 ns, βfast = 0.24 ± 0.08 ns; φslow = 226 ± 24 ns, βslow = 0.76 ± 0.08 ns. (c) For 60 min (aggregate): τm = 14.08 ± 1.14 ns; φfast = 0.20 ± 0.04 ns, βfast = 0.21 ± 0.04 ns; φslow = 300 ± 17 ns, βslow = 0.79 ± 0.04 ns. b The results on the monomeric protein are taken from Ref. 73.

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Figure 1

(a)

(b)

(c)

W 184 W 267

W 148

250 nm

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(a)

(b)

18 12

2

0.8 0.6

0.6 0.4 0.2 0.0 45

470

90

135 180 Time (min)

225

480 490 500 Wavelength (nm)

510

(c)

0.18

Oligomer Monomer

0.17 0

20

40

60

80

1.00 0.96 0

70

0

140 210 Time (min)

280

-6

520

0.20

0.19

1.04

-12

270

100 120 140 160

195

Normalized ANS Fl. Intensity (A.U.)

0

0.0 460

6

1.08

210 225 Wavelength (nm)

240

1.0

(d)

0.22

0.8 ANS Fl. Anisotropy

0.2

0.8

3

0.4

1.0

1.12 [Θ]218/[Θ]222

-1

[θ ] (X 10 deg.cm .dmol )

1.0

ThT Fl. Intensity (A.U.)

Normalized ThT Fl. Intensity (A.U.)

Figure 2

Trp Fluorescence Anisotropy

1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

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0.6

0.4

0.20 0.18

Oligomer

0.16 0.14

Monomer

0.12 0

30

60 90 120 150 180 Time (min)

0.2 0

30

Time (min)

60

90 120 Time (min)

150

180

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Figure 3

0.20

0.20

(a)

0.20

(b)

0 min

0.12

0.12

0.12

[r(t)] 0.08

0.08

0.08

0.04

0.04

0.04

5

10

15 20 t / ns

25

30

0.00

35

0

5

10

15 20 t / ns

25

30

25 Average Molar mass (x 104) (Da)

(d) 20 15 10 5 0 0

20

40

60 80 100 120 140 160 Time (min)

400

(e)

300

200

100

0 0

20

40

0.00

35

60 80 100 120 140 160 Time (min)

0

Avg. rate constant (k) (X 10-3) (min-1)

0

1h

[r(t)]

0.16

[r(t)]

0.16

0.00

(c)

15 min

0.16

Average Rh (nm)

1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

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5

10

15 20 t / ns

500

25

30

35

Intensity (Fluo., CD) Fluo. Anisotropy, Avg. Rh

(f) 400

Avg. Molar Mass

300 200 100 0

Trp

ANS

ThT

Rh

β/α α

Molar Mass

Read-outs

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(a)

40

(c)

0 h

Time (min)

10

24

8

h

6

4

4

h

2

2

0

m in

4000

10

15

6000

20

m in

8000

(b)

0

Amyloid sample added

10000

30

Na tiv e

12000

Calcein Efflux Efficiency (%)

Calcein Fluorescence (A.U.)

Figure 4

Calcein Efflux Efficiency (%)

1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

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15 min 4h

(d) 30 20 10 0 1 10 [Ovalbumin] (µ µM)

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Figure 5

0.30

(b)

DPH Fl. Anisotropy

0.20

0.15

0.10

0.25

10 µM

0.20

5 µM 1 µM

0.15

500 nM

h 24

h 4

h 2

m in 15

m in

0.10

0

0.3

20 µM

DPH Fl. Anisotropy (final)

(a) 0.25

N at iv e

DPH Fl. Anisotropy (final)

1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

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0

5

10

15 20 Time (min)

25

30

(c)

0 min 4h

0.2

0.1

0.0 1 10 [Ovalbumin] (µ µM)

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-20

-30

h 24

h 4

h 2

m in 15

m in 0

N

at iv e

-40

Changes in Dipole Potential (mV)

(a)

-10


Figure 6


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0 -20

(b)

-40 -60 -80

0 min 4h

1 10 [Ovalbumin] (µ µM)

(c)

0 -20 -40 0 min 15 min 2h 4h 24 h

-60 -80 0

5

10 15 [Ovalbumin] (µ µM)

20

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Table 1

Fluorescence lifetime in ns (amplitude)

Mean lifetime in ns

Rotational correlation times in nsa (amplitude)

Time-points (protein state)

τ1 (α1)

τ2 (α2)

τ3 (α3)

τm

χ2

φfast (βfast)

φslow (βslow)

r0

rss

χ2

Molten-globule (monomer)b

1.1 (0.14)

6.47 (0.38)

15.8 (0.48)

10.2

0.99

0.11 (0.59)

23.7 (0.41)

0.160

0.043

1.16

0 min (oligomer)

0.82 (0.13)

6.50 (0.37)

17.14 (0.50)

11.2

0.98

0.34 (0.34)

43.0 (0.66)

0.177

0.088

1.24

15 min (aggregate)

1.21 (0.06)

6.84 (0.17)

18.25 (0.77)

15.5

1.03

0.20 (0.25)

249.1 (0.75)

0.182

0.129

1.11

1 hour (aggregate)

0.67 (0.11)

6.62 (0.16)

18.20 (0.73)

14.7

1.02

0.21 (0.24)

300 (0.76)

0.203

0.147

1.02

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TOC Graphic

For Table of Contents Only

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Self-Assembly of Ovalbumin Amyloid Pores: Effects on Membrane

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