Thermodynamic versus Conformational Metastability in Fibril-Forming

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Thermodynamic Versus Conformational Metastability in Fibril-Forming Lysozyme Solutions Samuele Raccosta, Vincenzo Martorana, and Mauro Manno J. Phys. Chem. B, Just Accepted Manuscript • Publication Date (Web): 17 Sep 2012 Downloaded from http://pubs.acs.org on September 24, 2012

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The Journal of Physical Chemistry

Thermodynamic versus Conformational Metastability in Fibril-forming Lysozyme Solutions Samuele Raccosta ‡§†, Vincenzo Martorana ‡, Mauro Manno ‡* ‡ Institute of Biophysics, National Research Council of Italy, via U. La Malfa 153, I-90146 Palermo, Italy. § Dipartimento di Fisica, Università di Palermo, via Archirafi 36, I-90123 Palermo, Italy.

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ABSTRACT. The role of intermolecular interaction in fibril-forming protein solutions and its relation with molecular conformation is a crucial aspect for the control and inhibition of amyloid structures. Here, we study the fibril formation and the protein-protein interactions of lysozyme at acidic pH and low ionic strength. The amyloid formation occurs after a long lag-time and is preceded by the formation of oligomers, which seems to be off-pathway with respect to fibrillation. By measuring the osmotic isothermal compressibility and the collective diffusion coefficient of lysozyme in solution, we observe that the monomeric solution is kept in a thermodynamically metastable state by strong electrostatic repulsion, even in denaturing conditions. The measured repulsive interaction between monomers is satisfactorily accounted for by classical polyelectrolyte theory. Further, we observe a slow conformational change involving both secondary and tertiary structure, which drives the proteins towards a more hydrophobic conformation. Denatured proteins are driven out of metastability through conformational substates, which are kinetically populated and experience a lower activation energy for fibril formation. Thus, our results highlight the role of electrostatic repulsion, which hinders the aggregation of partially denatured proteins and operates as a gatekeeper favoring the association of those monomers whose conformation is capable of forming amyloid structure.

KEYWORDS. Protein stability, Amyloid fibrils, Electrostatics, Protein interaction, Light Scattering, Virial coefficent.


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INTRODUCTION Amyloid fibrils are linearly elongated protein aggregates,1 characterized by a cross-β sheet quaternary structure running along the main fiber axis.2,3 Amyloid fibrils and deposits have a high clinical relevance, since they are related to several diseases, including Alzheimer’s and prion disease.4 In addition, they have a general biological importance as underlined by the existence of functional amyloids with a positive physiological activity.5 In both perspectives, it is of extreme interest to determine the mechanisms and the factors which are useful to inhibit, promote and control the formation (or the disruption) of amyloid fibrils. A well-established relation exists between the capability of a protein to form amyloid fibrils and its molecular conformation and sequence.6 The propensity towards fibril formation has been related to the amount of β-sheet structure, and several computational or theoretical methods have been developed to rationalize and predict fibrillation rates.7–9 This link has been strengthened by several observations of native proteins capable of forming amyloid fibrils when their conformation is altered by different thermodynamic or environmental conditions, such as e.g. high temperature, low pH, addition of denaturants.10 In this respect, amyloid seems to be a generic structural motif, in which polypeptide chains can organize, and this explains why the molecular bases of amyloid formation are diffusely studied even with model proteins and in nonphysiological conditions.6,10 If the role of protein conformation in amyloid formation has been extensively studied, the details of the intermolecular interaction potentials, which determine protein attraction and cause solution instability, are still not completely explored, and a few studies explicitly focused quantitatively on these aspects.11–17 The physics of intermolecular interaction has been studied for another important type of ordered aggregation, that is crystallization. The formation of


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crystals of native proteins is prompted by the onset of mild protein attractive interactions,18–20 joined to conditions stabilizing the native conformation.21,22 Amyloid aggregation is often correlated to the stabilization of partially unfolded states.23 This is consistent with the fact that hydrophobic attraction plays a main role in the assembly of amyloid structures. In addition, several proteins may form fibrils not only at high temperature, when they undergo partial thermal unfolding, but also at pH far from the isoelectric point, when they are electrically charged.24–27 Indeed, thermal unfolding causes the exposure of hydrophobic residues thus enhancing the hydrophobic attraction, which overwhelms electrostatic repulsion and triggers protein aggregation. However, this scheme typically determines the formation of amorphous aggregates: therefore we may argue that a more subtle mechanism should drive the assembly into ordered fibrillar amyloid-like structures. Sciortino and co-workers have shown that the competition between strong attraction and charge repulsion may determine the formation of elongated colloidal structures, since the elongated geometry reduces the energy cost for the association of molecules with like charges.28,29 Very recently, a remarkable work by Mushol and coworkers has addressed the issue of electrostatic charges in the formation of lysozyme fibrils.17 They found that repulsive charge interactions are a prerequisite for amyloid formation, while net attraction would cause precipitation. Also, they found that a different screening of electrostatic potential can regulate the assembly pathway, by altering the spatial extent of repulsion. In the present work, we are extending their studies on lysozyme fibrils to different conditions, that is at very low ionic strength (11 mM) and at high temperature (60 – 70 "C). In such conditions, lysozyme forms fibrils with a lag time of a few days.30 Our experiments show that at the onset of kinetics the solution is made of partially unfolded lysozyme monomers. Afterwards, small oligomers form with a hydrodynamic radius of about 10 nm. By light scattering (LS) experiments we measured


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the osmotic isothermal compressibility and the collective diffusion coefficient, related to the thermodynamic and hydrodynamic properties of the solution. Our results show quantitatively that the electrostatic repulsion among molecules determines the kinetic stability of the monomeric solutions. Circular dichroism (CD) and intrinsic tryptophan photoluminescence (PL) experiments evidence a slow kinetic shift towards unfolded conformations. This conversion precedes fibrillation and may explain the pathway from the “metastable” state of partially unfolded lysozyme monomers to the stable state of amyloid fibrils, in keeping with a recent characterization of the energy landscape of monomeric human lysozyme solutions.31, 32

EXPERIMENTAL METHODS Sample preparation. Hen egg-white lysozyme (three times crystallized, dialyzed and freezedried) was purchased from Sigma Chemical Co. and used without further purification. Protein powder was dissolved into Millipore Super-Q water and pH 2 was achieved by adding small amounts of 1 M hydrochloric acid. Samples were filtered into a cell through a 0.2 µm Millex-LG syringe filter. For the compressibility measurements, in order to be sure that there were not small pre-existing clusters of proteins, a stock solution was filtered by 30 kDa AMICON ultracentrifuge filters and then diluted by acidic water to obtain different protein concentrations in the range 0.8–12 mg ml−1. These solutions were further filtered directly in cuvette by Millex LG syringe filters. In all experiments the lack of both dust and residual aggregates was checked by preliminary dynamic LS measurements. Protein concentration was determined by UV absorption spectroscopy measurements (Shimadzu UV-2401 PC). The extinction coefficient for lysozyme at 280 nm was taken as 2.46 cm2 mg−1. The ionic species in solution are derived from the dissociation of hydrochloric acid, added to adjust the pH, and to a minor extent from the


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dissociation of sodium acetate, present in the crystallized powder. The ionic strength I of solutions is estimated by taking into account all the ionic species: I = ½ ∑ Zj2nj, where Zj and nj are the electric charge number and the molar concentration of the j-th ionic species, respectively. At increasing protein concentration the ionic strength slightly increases due to the high charge number of protein molecules. However, for diluted lysozyme samples (below 0.3 mM) the calculated ionic strength was estimated to be below 60 mM, while the solvent ionic strength, calculated neglecting the protein contribution, is below 11 mM. The electrical conductivity of lysozyme solution was measured by Multi 3410 digital conductivity meter (WTW GmbH). In the studied concentration range, the conductivity value did not depend upon lysozyme concentration and its value was slightly higher (between 10 and 20%) than the value measured for a 10 mM HCl water solution, consistent with a solvent contribution to the ionic strength between 11 and 12 mM. Differential scanning calorimetry (DSC). DSC data were obtained by using a Hart Scientific (mod. 3705/06) power compensation calorimeter, coupled with a transputer driven control unit,33 and calibrated with naphthalene. 270 µl of a 2.6 mM lysozyme solution and buffer solution were introduced into 1 ml steel sample and reference cells, respectively, and kept under nitrogen laminar flow to avoid air water condensation. The excess specific heat cpex is calculated from the calorimetric profile by subtraction of a quadratic baseline, which is obtained by fitting the calorimetric curve without the peak. Circular dichroism (CD). Far-UV CD spectra were measured on 0.2 mg ml−1 lysozyme samples by using an OLIS (DSM 10 CD) spectrometer (0.1 cm quartz cell, bandwidth 2 nm, step-mode, response time automatically adjusted to maximize signal to noise ratio). The temperature was set by a computer-controlled Peltier device. Mean Residue Ellipticity [θ] for each spectrum was obtained by subtracting the corresponding solvent spectrum. CD spectra were


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analyzed (in the range 178-260 nm) by using the CONTINLL algorithm, and a reference set of 48 proteins (42 soluble and 6 unfolded).34 CONTINLL fits the CD signal of unknown proteins by a linear combination of the spectra of a large basis of reference proteins with known conformations. The analysis yields an estimate of the content of α-helix, α3−10-helix, β-turn, proline-turn and random coil. Photoluminescence (PL). Photoluminescence measurements were performed on 0.18 mg ml−1 lysozyme samples by using a Jasco FP-6500 spectrofluorimeter (1 cm length quartz cell, excitation wavelength 300 nm, excitation and emission bandwidth 3 nm, scan speed 100 nm min−1, response time 2 s). The excitation wavelength was greater than 280 nm to minimize the excitation of tyrosine residues in lysozyme and to avoid bleaching after long exposure. The temperature of the cell holder was regulated by an external recirculating bath. From the spectrum profile the first momentum M1 was calculated as a meaningful parameter that takes into account both centre of band and width. It is defined as M1 = ∫ λP(λ)dλ where P(λ) is the normalized band profile. Atomic force microscopy (AFM). After incubation at high temperature (7 days at 60 and 65 °C, and 3 days at 70 °C), lysozyme samples (18.5 mg ml−1) were diluted 10000 times in Millipore Super-Q water mixed with small amounts of hydrochloric acid. 20 µl drop of each sample was deposited on a mica substrate and dried by a gentle nitrogen flux to remove most of the surface water. Images of protein aggregates were recorded by a Multimode Nanoscope V Atomic Force Microscope (Veeco Instruments, Santa Barbara, CA, USA), operating in air tapping mode (resolution 512x512, scan rate 0.5 Hz). We used rigid cantilevers Nanosensor PPPNCHR-50 (resonance frequency 330 kHz), equipped with silicon tips with a nominal radius of curvature of 7 nm. The analysis of AFM images was performed by the open source software Gwyddion.


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Light scattering (LS). Static and dynamic LS measurements were performed by using a Brookhaven BI200-SM goniometer equipped with either a He-Ne laser (λ0 = 632.8 nm, used for kinetics measurements) or a solid-state laser (λ0 = 532 nm, used for compressibility measurements). The temperature of the thermostated cell compartment was controlled within 0.05 °C by using a thermostated recirculating bath. The scattered light intensity and its time autocorrelation function g2(t) were simultaneously measured at 90° by using a Brookhaven BI9000 correlator. A photodiode was used to collect forward scattered light and to measure the sample turbidity. Absolute values of scattered intensity (Rayleigh ratio) were obtained by normalization with respect to toluene, whose Rayleigh ratio at 632.8 nm and 532 nm was taken as 14.0 10-6 cm−1 and 28.0 10-6 cm−1, respectively.35 The apparent diffusion coefficient Dapp, was obtained

from

the intensity autocorrelation function by fitting to the expression

g2(t) = 1+β2exp{−Dappq2t}2, where β is an instrumental factor, and q is the scattering vector q = 4πñλ0−1sin(θ/2), which depends upon the scattering angle θ, the incident wavelength λ0, and the refractive index of the medium ñ.36 The correlation function during the kinetics of aggregation were fit by a regularization method to obtain the distribution function P(Rh) of hydrodynamic radii Rh.37 This is related to the correlation function of the electric field g1(t) = β-1[g2(t)-1] 0.5 by the following expression: g1(t) = ∫ P(Rh)exp{-D(Rh)q2t} dRh, where q is the scattering vector and D(Rh) = kBT(6ηRh)−1 is the diffusion coefficient of an object with a hydrodynamic radius Rh, with kB being the Boltzmann constant, T the temperature, and η the solvent viscosity. The distribution function P(Rh) is used to partition the total scattered intensity R90 (Rayleigh ratio) into different contributions related to specific species. Namely, the Rayleigh ratio contributed by molecular species with a hydrodynamic radius in the range ∆R = [R1,R2] is R90(∆R) = R90 ∫∆R P(Rh)dRh.


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RESULTS Lysozyme at acidic pH. In the present work, we study the thermodynamic and conformational instability of lysozyme forming amyloid fibrils in acidic solution and at low ionic strength. In such conditions, lysozyme molecule has a high electric charge number (Z=17), and its thermal stability is considerably reduced with respect to physiological pH, as known from calorimetric studies at low pH for the case of human lysozyme.31, 38 Here, we performed differential scanning calorimetry (DSC) experiments at our experimental conditions (pH 2 and ionic strength 11 mM). Fig. 1 shows a comparison between the calorimetric thermograms of the excess heat capacity cpex at pH 7 (100 mM phosphate buffer) and pH 2. Both the onset and the maximum of the thermogram peak at pH 2 occur at lower temperature than at pH 7. Also, the transition is definitively broader at pH 2, suggesting a greater flexibility of the protein and the presence of more conformational substates, in keeping with recent findings.31 The subsequent kinetic experiments are performed in the range 60-70 °C, where the denatured state is largely populated. Kinetics of lysozyme fibrillation. Lysozyme solutions at pH 2 and low ionic strength (11 mM) were incubated at 60, 65 and 70 °C for several days. The long incubation time is required by the long lag-time before the formation of lysozyme fibrils, as known from previous studies on lysozyme in the same conditions30 or at higher ionic strength.17, 39 During incubation, simultaneous static and dynamic LS experiments were performed to measure the scattered light intensity at 90° (given in terms of Rayleigh ratio R90), and the electric field correlation function g1(t). The intensity autocorrelation functions measured in the course of the kinetics were fit by a regularization method to derive the distribution function P(Rh) of hydrodynamic radii Rh. We may distinguish different peaks in P(Rh), related to different species: monomers, oligomers,


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fibrils; a fourth peak related to large objects (as fibrillar or amorphous bundles) is sometimes observed at very long incubation times. The distribution functions were used to partition the total scattered intensity into different contributions related to specific species. The following ranges of hydrodynamic radii were used: 0 – 3 nm for monomers, 3 – 30 nm for oligomers, 30 – 3000 nm for fibrils. Fig. 2 shows the growth of the signal related to monomers (grey circles), oligomers (cyan circle) and fibrils (red circles) for the three temperatures 60, 65 and 70 °C. The Rayleigh ratio of fibrils exhibits a long lag-time as previously reported.17, 30, 39 At 70 °C the lag-phase is extremely reduced, if not suppressed. Also, large micrometer-size objects start to be observable since the very beginning of the kinetics (data not shown). The Rayleigh ratio of fibrillar species does not reach a stationary value within the monitored time window. The Rayleigh ratio of oligomers starts increasing at the onset of the kinetics and reaches a plateau within the first day, accompanied by a decrease of the monomer signal. The latter implies a reduction of the mass concentration of the monomeric species and an increase of the mass concentration of the oligomeric species, which has a constant average hydrodynamic radius of about 8 – 10 nm (see supporting information). In a typical polydisperse solution, the formation of a species with a size of one order of magnitude larger than the monomeric species and a mass concentration of the same order of magnitude of the monomer would cause an increase of the scattered intensity of one or two order of magnitude, since the Rayleigh ratio is proportional to both mass and concentration of the scattering species. In the present experiments, the scattered intensity remains almost constant indicating that the oligomeric species has the same compressibility of the monomeric species. This is a confirmation of the high repulsive interactions, which decrease the concentration fluctuations and hence the scattered intensity. In this context, the oligomeric species may either be a real cluster of bound monomers or a marker of concentration fluctuations,


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analogous to the so called “cluster phase” observed by Stradner et al. in lysozyme solutions at neutral pH.40 The nature of this oligomeric phase is beyond the scope of the present work. Morphological properties of lysozyme fibrils. We studied the morphological properties of samples obtained after incubation of acidic lysozyme solutions for one week at different temperatures by atomic force microscopy (AFM). AFM images show lysozyme fibrils with lengths from a few hundreds of nanometers to a few microns (Fig. 3a). The height of fibrils in Fig. 3a was estimated by measuring the profile perpendicular to the main axis at different places of various fibrils. The fibril height distribution is shown in Fig. 3b, with an average height h = 2.3 nm, which points to the existence of fibrils formed by monomer units along the elongation axis. The observed height of fibrils is quite distributed and we can distinguish simple fibrils and some thicker fibrils. The apparent average width is of about 30 – 40 nm, much larger than fibril height, which is a known effect of tip finite size. By analyzing the height profile along the elongation axis we observed an axial periodicity of 20 nm or higher (Figs. 3c and 3d). A periodic length along the axis of the fibrils (typically related to their thickness) confirms a twisted or coiled structure.41 The twisting of two fibril filaments is also evident in Figs. 3e and 3f. AFM images of fibrils formed by incubation at 60, 65 and 70 °C show analogous morphologies. At 70 °C a larger quantity of amorphous globular aggregates can be observed (see Supplementary Information). Analogously, AFM images on fibrils formed at 65 °C with slightly higher salt concentration (20 mM NaCl) show rare fibrils and a large quantity of globular objects. Osmotic compressibility and collective diffusion. Static LS experiments were performed at different temperatures (from 20 to 70 °C) and at different lysozyme concentrations to measure the Rayleigh ratio at 90°, R90 (Fig. 4a). Freshly prepared and accurately filtered monomeric solutions were quenched in the thermostated cell of the LS device, and measurements were performed shortly after the complete thermalization, which requires a few minutes. In these


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experimental conditions the samples remain monomeric. In diluted monomeric lysozyme solutions, we may assume that the Rayleigh ratio does not depend upon the scattering angle θ. Indeed, both the protein form factor and the structure factor can be taken as equal to 1, since the protein size is much less than the reciprocal of the scattering vector and the protein concentration is considerably low.36 In such conditions, the Rayleigh ratio is proportional to the osmotic isothermal compressibility κT: R90 = KcMw ρkBTκT, where c is the mass concentration, kB is the Boltzmann constant, T is the temperature, Mw is the weight average mass, ρ = NAM0-1c is the protein number concentration, dependent upon the Avogadro number NA and the molecular mass M0, and K is a constant, which depends upon the incident wavelength λ0, the refractive index of the solution ñ and its increment with concentration (whose measured value is [dñ/dc] = 0.172 g−1 cm3, see Supplementary Information): K = (2πñ[dñ/dc] λ0−2)2NA-1. Therefore, the dependence of R90 on protein mass concentration c yields the weight averaged molecular mass of lysozyme, Mw, and the osmotic second virial coefficient, B2, according to the expression:36

[1]

where O(ρ2) stands for any term of the order of ρ2 or higher. In Fig. 4a, the experimental data and the fitting functions are shown according to the expression [1], (and using the nominal molecular mass M0 = 14.3 kDa). Both a linear fit at concentrations below 5 mg ml−1 (that is neglecting all the terms O(ρ2) in [1]) and a quadratic fit at all concentrations (including a term proportional to

ρ2) were performed and gave consistent results. The average Mw in the measured range of temperatures is = 14.3 ± 0.5 kDa, which perfectly agrees with the nominal molecular mass of lysozyme (Fig. 5a). The osmotic second virial coefficient B2 is related to the two-bodies mean-


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field potential U(r),42 accounting for the interaction between two proteins at a center-to-center distance r:

[2]

where β = (kBT)−1. Simultaneously with static LS, dynamic LS experiments were performed to measure the intensity autocorrelation function and to extract the collective diffusion coefficient Dc (Fig. 4b). The dependence of Dc on protein concentration c yields the z-averaged hydrodynamic radius of lysozyme, Rh, and the hydrodynamic coefficient, h2, according to the expression:36, 43

[3]

where O(ρ2) stands for terms of the order of ρ2 or higher and η is the medium viscosity. In Fig. 4b, the experimental data and the fitting functions are shown according to the expression [3]. As for the static LS data, both a linear fit at concentrations below 6 mg ml−1 and a quadratic fit at all concentrations were performed and gave consistent results (the data at concentrations below 1 mg ml−1 were excluded from the fit, due to their large experimental error). The hydrodynamic radius Rh at lower temperatures (below 50 °C) is = 1.65 ± 0.05 nm, while at higher temperatures (above 50 °C) it slightly increases to the value = 1.76 ± 0.08 nm (Fig. 5c). Conformational changes accompanying aggregation. The secondary structure of lysozyme at pH 2 and low ionic strength was studied by far-UV CD. CD spectra do not change up to 50 °C.17, 30 At higher temperature the spectra change as a consequence of thermal unfolding. In Fig.


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6a, we note the conspicuous difference between the spectrum at 25 °C (black dashed line) and the spectrum at 60 °C after thermalization (black solid line). The mean residue ellipticity [θ] between 208 and 230 nm is less negative by increasing the temperature and the minimum at 208 nm shifts towards lower wavelengths, up to 202 nm, as typically observed in a α-to-β transition. Indeed, the deconvolution of spectra by CONTINLL shows a decrease of the content of α-helical structure from about 40% to about 20% from 25 to 60 °C, and a corresponding relative increase of the content of β-turn structure and random coil (see Supplementary Information). Beyond such a fast conformational change due to the temperature increase, one observes a continuous slow change of CD spectra upon incubation at 60 °C, which accompanies and precedes fibrillation (Fig. 6a). In particular, one observes a decrease of the ellipticity [θ] at 209 and 222 nm, which are the minima of a spectrum typical of α-helical structures. A detailed analysis by CONTINLL elicited that the amount of α-helical structure is further decreased (inset of Fig. 6a), and converted mainly into random coil (see Supplementary Information). A similar behavior was observed at 65 °C and 70 °C. Intrinsic tryptophan PL spectra of lysozyme at 25 and 60 °C are displayed in Fig. 6b. These measurements show the effect of temperature increase and time of incubation on the environment of the tryptophan residues.44 By changing temperature from 25 °C (black dashed line) to 60 °C (black solid line), the emission spectrum exhibits a clear red-shift. Although lysozyme has six tryptophans, the main contribution to the intrinsic tryptophan PL is attributed to residue 62, which is completely solvent exposed, and residue 108, which is buried in the protein near the active cleft.45, 46 Therefore, the red-shift can be attributed to a partial unfolding, which consists in changes of the protein tertiary structure and a more consistent solvent exposure of tryptophan 108.47


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Fig. 6b also shows the PL spectra upon incubation at 60 °C. As time goes by, the spectra are further red-shifted with a slow kinetics. This red-shift is displayed in the inset of Fig. 6b by using the first moment of the emission spectra M1 as a significant (and robust) parameter. A similar behavior is observed in samples incubated at 65 and 70 °C.

DISCUSSION Lysozyme is known to form amyloid fibrils in different conditions, such as the addition of alcohols,24, 48 reducing agent,49 and denaturants,50 or at acidic pH and high temperature.17, 30– 32, 39, 51

In the present work we study lysozyme fibrillation at pH 2 and low ionic strength to

determine the kinetics of lysozyme fibrillation and conformational changes, as well as the thermodynamic properties of fibrillation-prone monomeric lysozyme solutions. We performed time lapsed LS experiments at 60, 65 and 70 °C, above the unfolding temperature at pH 2. Fibrils form after a very long lag-time (Fig. 2), as already known at slightly lower temperatures and at higher ionic strength.17, 30, 39 AFM images show long fibrils characterized by thin twisted filaments with a size of a single lysozyme monomer (Fig. 3). These results are consistent with the findings of Hill et al.,17 who observed “monomeric” filaments at moderately low ionic strength and thicker “oligomeric” fibrils at high ionic strength. During the lag-time of fibril appearance, small oligomers with a hydrodynamic radius of about 10 nm form since the very beginning and increase in number within the first day, up to a plateau. The reached stationary value indicates that oligomers may be in equilibrium with monomers.40, 52–54 Furthemore, (i) their size distribution does not overlap with that of the larger fibrillar species (see supporting information); (ii) they do not display any enhanced fluorescent emission due to Thioflavin T binding (see supporting information); (iii) fresh lysozyme solutions seeded with


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15% w/w of oligomeric lysozyme do not exhibit any increased fibrillation rate upon incubation at 65 °C (see supporting information). All these observations suggest that such oligomers may be off-pathway to the fibrillar state. The size of the oligomers is consistent with the observations of spherical oligomers of human lysozyme formed at pH 3, with diameter 8–17 nm and a significant amount of β-sheet structure.51 However, the comparison among the present observation of oligomeric aggregates and analogous findings would require the knowledge of structural details. Such a comparison along with the investigation of the nature of the oligomeric species and the proof that they are off-pathway to fibrillation goes beyond the scope of this work and is left for further studies. The unfolded, “amyloidogenic” state of lysozyme at pH 2. At pH 2, all the polar residues are protonated and lysozyme reaches an electric charge number of Z = 17.55 Moreover, the present experiments were performed at low ionic strength (11 mM) and thus electrostatic interactions are not dramatically screened: in the studied range of temperature (20–70 °C) the Debye length λD has value of about 2.9 nm, which is comparable with the size of the molecule. At high temperature, upon thermal unfolding, the charge could also increase due to the protonation of terminal carboxyl group.56 The unfolding transition occurs between 45 and 70 °C, as detected by different techniques.17, 56 In comparison to higher pH, the transition occurs at lower temperature and is more spread, as confirmed by our DSC measurements (Fig. 1). From CD and PL measurements, we observe a clear difference between 25 and 60 °C in both the secondary and tertiary structure. In particular, the α-helical content is considerably reduced in favour of β-turn and random coil structural motifs (Fig. 6a, see also Supplementary Information), and the tryptophan 108 located in the active cleft is more exposed to solvent (Fig. 6b). In a recent study, the unfolding transition of human lysozyme at acidic pH has been described with a pseudo


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two- state model, in which a native state is converted into an ensemble of denatured states, with a low activation free-energy.31, 32 The measured hydrodynamic radius of lysozyme monomers has an average value of 1.65 nm up to 50 °C, and it exhibits a moderate increase of less than 10% at higher temperatures. The small variation observed upon denaturation is largely expected for two reasons. On one hand, lysozyme partial denaturation does not change significantly the protein volume, as confirmed by the very small variation observed in the radius of gyration measured by Xray scattering.57 On the other hand, hydrodynamic radius is not only related to molecular size and shape but to the overall diffusional properties, which are largely affected by the protein charge and the counterion hydration layers.36 As a matter of fact, a reduction of the ionic strength was known to reduce the experimentally observable gap between the hydrodynamic radius of the denatured state and that of the native state, as well as the actual value of the hydrodynamic radius of lysozyme at low temperature.56 Electrostatic repulsion determines solution stability. The two thermodynamic parameters measured by LS experiments, B2 and h2, shown in Figs. 5b and d, are determined by the interaction potential among solute molecules (expression 2).42 The sign of B2 is related to protein stability and solubility, depending upon the prevalence of repulsive or attractive interactions.18, 20, 58–60 In the measured range of temperature B2 is positive and therefore the solution of monomeric lysozyme is thermodynamically metastable. B2 is essentially constant, except for a moderate increase at the high temperatures (Fig. 5b). These results are consistent with those of Hill et al.,17 who found increasing values of B2 by lowering the ionic strength at pH 2, and with those of Velev et al.,61 who measured B2 of lysozyme solution at very low ionic strength (5 mM) down to pH 3.61, 62 The temperature dependence of B2 is remarkable. Indeed, at high temperature one could expect a decrease of B2, if not a sign inversion. Thermal denaturation


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typically causes the exposure of apolar residues and the enhancement of hydrophobic attraction. In the present case, this effect seems negligible with respect to that of electric charges. The hydrodynamic coefficient accounts for the hydrodynamic effects, that is for the propagation on the motion of a particle of the disturbances due to the motion of the other particles.36 One may expect h2 to be non zero for charged macromolecules (Fig. 5d):43 the considerably high values of h2 may be ascribed to the strong electrostatic repulsion.36, 63, 64 In analogy with the studies on colloidal solutions,20, 22, 65 one may assume as a first approximation a pairwise isotropic potential of “mean force” U(r) between two proteins with a center-to-center distance r.42 In dilute aqueous electrolyte solutions, the protein interaction has been described in the framework of the Derjaguin–Landau–Verwey–Overbeek (DLVO) theory.66 In this context, the protein molecules are modeled as charged, polarizable, hard spheres. The interaction consists of a hard sphere potential Uhs, a repulsive screened Coulomb electric potential Uel, and an attractive dispersion potential Uat. A first contribution to repulsive interaction is due to the steric repulsion, which can be modeled by the hard-sphere potential between two protein molecules with an “equivalent” hard-sphere diameter σ:

[4]

In the case of an ideal system with a repulsive hard sphere potential, the second virial coefficient is B2hs = 4VI (calculated from eqs. 2 and 4 by setting U(r) = Uhs(r)), where VI = πσ3 is an effective interaction volume. The hydrodynamic coefficient is also related to the interaction volume h2hs = 6.55 VI,67 while other treatments yield slightly different values.68


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The interaction volume VI and the equivalent hard-sphere diameter σ depend upon different parameters; VI is related to protein shape and solvation:69 VI = fs3 (v0 + δvw) M0NA−1 where v0 is the dry protein specific volume (v0 = 0.703 cm3 g−1 for lysozyme), vw is the solvent specific volume, δ is the weight of the hydration solvent per weight of protein,69 and fs is a shape correction factor included to take into account a non spherical shape.70 For ellipsoids with an axial ratio between 0.25 and 4, it is reasonable to calculate the interaction volume by using the hydrodynamic radius as an effective hard-sphere equivalent radius (VI = 4/3πRh3).71 Indeed, the calculated shape factors for the hydrodynamic radius Rh and the radius of the interaction volume σ/2 are approximatively equal for moderately asymmetric ellipsoids.71 The screened Coulomb potential is due to an ionic double-layer and can be expressed by the classic Debye-Hückel potential:

[5]

where σ is the hard-sphere diameter, λD = (8πlBNAI)−0.5 is the Debye screening length, lB = e2 (4πεkBT)−1 is the Bjerrum length, e is the electron charge, Z is the number of electronic charges of the protein, ε is the dielectric constant, and I is the ionic strength of the solution, which in the present experiments is 11 mM. At the low ionic strength and high protein charge of the present experiments, Debye-Hückel theory fails due to the large electrostatic energy at the protein surface (Wel > kBT). In such a context, polyelectrolyte theory foresees the collapse of counterions on protein surface (Manning condensation).72 This determines a reduced effective electric charge on the protein molecules. In the case of monovalent counterions and spherical macromolecules with a diameter σ = 3.3 nm)


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comparable with the Debye length (λD = 2.9 nm), one may determine the electrostatic potential by using

the

Debye-Hückel

expression

[5]

and

an

effective

charge

Zeff = σ/lB (1 + σ/2/λD)/ ln(λD/lB).73 By using the hard-sphere potential (eq. 4) plus the electrostatic Debye-Hückel potential (eq. 5) and an equivalent hard-sphere radius equal to the measured hydrodynamic radius, we calculated the second virial coefficient. The results are displayed in Fig. 5b (diamonds). No other contribution to the potential of mean force U(r), neither repulsive nor attractive, was necessary to get values so close to the measured B2. Although this calculation is not able to catch every single details, the agreement with the experimental data is remarkable if one thinks to the considerable amount of approximations underlying the theory. Kinetic instability of lysozyme conformation. Our results show that the monomeric lysozyme solution is thermodynamically metastable. In other words, the proteins undergo a first partial unfolding upon quenching at high temperatures. However, this initial unfolding is not sufficient to enhance the hydrophobic attraction between two proteins, so that it overturns the electrostatic interaction: the interaction remains on average strongly repulsive. The addition of salt is able to screen electrostatic forces and favour aggregation and fibrillation.17, 39 However, a high ionic strength determines a strong damping of repulsive contribution, and may induce amorphous precipitation.17 Increasing temperature has an analogous effect, since the hydrophobic attraction is enhanced with respect to repulsion. Large micron sized aggregates are present in our fibrillation kinetics at 70 °C, and globular objects were found in AFM images at 70 °C, along with fibrillar aggregates (see Supplementary Information). However, our experiments performed at very low ionic strength highlight the important role of strong repulsion to allow fibrillation. Which mechanisms or interactions do allow the protein to associate and move from the “metastable” state of monomeric solution to the stable state of fibrillar aggregates? One may


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argue, as in Hill et al.,17 that electrostatic repulsion prevents amorphous association and selects those pathways that allow the formation of energetically favorable intermolecular contacts.74 This scenario is enriched by the present experiments on the kinetics of conformational changes (Fig. 6). Our observations show a slow continuous change of lysozyme conformation, which starts well before the appearance of fibrils or other aggregates, and continues beyond. These results might be explained as a progressive conformational change of one molecular species that is conformationally unstable due to the disruptive effect of the internal electrostatic repulsion of like charged residues and the cleavage of surface salt bridges.75 Such a progressive conformational change would increase the strength of attractive (hydrophobic) interactions, and therefore the onset of fibrillation would be determined by the prevalence of the attractive term over the repulsive one. In this context, the analysis of the measured thermodynamic parameters holds only at the onset of incubation. Alternatively, one may argue that the spectroscopic changes are due to a shift of the equilibrium among different conformations. This is consistent with a recent work, which characterizes the energy landscape of amyloidogenic human lysozyme and finds an ensemble of partially denatured, almost isoenergetic, states, with different fibrillation propensity.31, 32 Therefore, electrostatic repulsion operates as a gatekeeper for the conformations capable of forming amyloids. When a few proteins with such a conformation associate the equilibrium is slightly shifted towards them, until a significant amount of fibrillated protein is reached and can be detected.

CONCLUSION The conclusions derived from the present experiments are sketched as a cartoon in Fig. 7 in terms of protein energy landscape.23, 32, 76 The present experimental results on isothermal


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compressibility (Figs. 4,5) clearly indicate that at low pH and ionic strength monomeric lysozyme solutions are thermodynamically stable when lysozyme is both in the native state (low temperature) and in the denatured state (high temperature). Our analysis in terms of classical polyelectrolyte theory was able to adequately and quantitatively describe the solution stability (Fig. 5). At high temperature, the monomeric lysozyme solution is metastable, since the most stable state is given by lysozyme aggregates, as we know from both the present experiments (Figs. 2,3) and the recent literature.30, 39 The rationale for such metastability is given by the strong electrostatic repulsion, which provides high energies of activation for the aggregation. This energy landscape is pictorially shown as a thick red line in Fig. 7. The system of denatured proteins is driven out of metastability through specific conformational substates, which are kinetically populated (Fig. 6) and experience lower activation energy for fibril formation, reasonably due to a more conspicuous amount of β-turn and random structures (as revealed by our CD kinetics experiments: Fig. 6a). As discussed above, the fibril-prone substates may be either the most stable state reached through a slow interconversion (solid red arrow between DN and DF in Fig. 7), or a state in equilibrium with the other partially denatured conformation states, which is progressively sequestered by the fibrillation process (solid and dashed red arrows in Fig. 7). While both occurrences are admitted by the present experiments, the latter one is in keeping with other recent works.31, 32 The propensity for fibril formation is reduced by increasing the ionic strength,17 which implies a reduction of Debye screening length and hence a decrease of the activation energy for amorphous aggregation. A temperature increase trivially implies the capability to explore the entire energy landscape and thus results in an effective reduction of fibrillation propensity, as observed above 65 °C in the present work (see Electronic Supporting Information).


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In summary, the present work exploits a novel scientific approach (shared with the recent work of Hill et al.17) by studying quantitatively the intermolecular interaction involved in protein fibrillation. Our experiments elicit a novel interconnection between strong electrostatic repulsion and different partially denatured substates in fibril-forming lysozyme solutions. These results foster the question whether the co-existence of these two aspects is peculiar of lysozyme acidic solutions or it is a more ubiquitous feature of amyloid formation.

ASSOCIATED CONTENT Supporting Information. Analysis of secondary structure changes, AFM images of aggregates at 70 °C or 20 mM ionic strength, measurement of the refractive index increment for lysozyme at pH 2, DLS kinetics of lysozyme fibrillation, ThT fluorescence kinetics, seeding experiments. This information is available free of charge via the Internet at http://pubs.acs.org.

AUTHOR INFORMATION Corresponding Author *Author to whom correspondence should be addressed. Tel.: +39(091)680-9305, Fax: +39(091)680-9349, E-mail: [email protected]

Present Addresses † Present address: Biophysics & Nanoscience Centre CNISM, Facoltà di Scienze, Università della Tuscia, 01100 Viterbo, Italy


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ACKNOWLEDGMENT

We thank M. D’Amico, M. Leone, R. Noto, P. L. San Biagio, V. Vetri for discussions and collaborations. We thank A. Cupane for his suggestions regarding polyelectrolyte theories. One of the authors (S.R.) also thanks D. Bulone for help and discussions. The access to up-to-date AFM instrumentation was possible thanks to the kind support of M. Leone. This work was partially supported by the National Research Council of Italy through the project Intermolecular interaction in protein metastable solution.


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Figure 1. Excess heat capacity cpex of 2.6 mM lysozyme solutions at pH 7, 100 mM phosphate buffer (dashed line), and at pH 2 (solid line).


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Figure 2. Kinetics of fibril formation of 18.4 mg ml−1 lysozyme solutions at pH 2, incubated at 60 (panel a), 65 (panel b) and 70 °C (panel c). The data represent the measured Rayleigh ratio R90 (black circles) and the contributions associated with monomers (grey circles), oligomers (cyan circle), fibrils (red circles).


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Figure 3. AFM images of 18.4 mg ml−1 lysozyme solutions incubated at high temperature. (a),(c): sample incubated at 65 °C for 7 days. (b): distribution of fibril height obtained by sampling the profiles perpendicular to the axis of fibrils in panel a. (d): periodicity on the elongation axis of the fibrils taken along the white segment in panel c. (e),(f): high resolution images of a sample incubated at 70 °C for 3 days.


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Figure 4. (a) Normalized reciprocal Rayleigh ratio R90 , and (b) normalized collective diffusion coefficient Dc as a function of protein concentration (black circles) at pH 2 and at different temperatures, as indicated in each inset. Red solid lines and blue dashed lines are a linear and a quadratic fit to data, respectively.


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Figure 5. Physical parameters obtained from the analysis of LS data. (a): Weight averaged mass Mw. Dashed line indicates the nominal Mw = 14.3 kDa. (b): Second virial coefficient B2 derived from the analysis of experimental data (red circles), and calculated from theory (grey diamonds). (c): Hydrodynamic radius Rh. Dashed line represents the average at temperature below 50 °C ( = 1.65 nm). (d): Hydrodynamic coefficient h2 derived from the analysis of experimental data.


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Figure 6. (a) Kinetics of far-UV CD spectra of 0.21 mg ml−1 lysozyme solutions at pH 2 incubated at 60 °C (black and colored solid lines). The dashed black line is the CD spectrum at 25 °C, before incubation. Inset: behavior of the α-helix content, obtained from CONTINLL analysis, as a function of the incubation time; the continuous line is a guide to the eye. (b) Kinetics of intrinsic tryptophan PL spectrum of 0.18 mg ml−1 lysozyme solutions at pH 2 incubated at 60 °C (black and colored solid lines). The dashed black line is the PL spectrum at 25 °C, before incubation. Inset: behavior of the spectrum first moment M1 as a function of the incubation time; the continuous line is a guide to the eye.


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Figure 7. Energy landscape of metastable lysozyme solution at pH 2 and low ionic strength. The electrostatic potential has a Debye length λD higher than the protein hydrodynamic radius Rh and prevents protein-protein association. Multiple denatured states exhibit different molecular conformations, which thus encounter different energies of activation to undergo either amorphous or amyloid aggregation.


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REFERENCES (1) J. D. Sipe and A. S. Cohen, J. Struct. Biol., 2000, 130, 88–98. (2) M. Sunde and C. Blake, Adv. Protein Chem., 1997, 50, 123–159. (3) J. Greenwald and R. Riek, Structure, 2010, 18, 1244–1260. (4) F. Chiti and C. M. Dobson, Annu. Rev. Biochem., 2006, 75, 333–366. (5) D. M. Fowler, A. V. Koulov, W. E. Balch and J. W. Kelly, Trends Biochem. Sci., 2007, 32, 217–224. (6) L. Goldschmidt, P. K. Teng, R. Riek and D. Eisenberg, Proc. Natl. Acad. Sci. USA, 201, 107, 3487–3492. (7) A.-M. Fernandez-Escamilla, F. Rousseau, J. Schymkowitz and L. Serrano, Nature Biotechnology, 2004, 10, 1302–1306. (8) A. Trovato, F. Chiti, A. Maritan and F. Seno, PLoS Comput. Biol., 2006, 2, e170(1608– 1618). (9) G. Tartaglia, A. P. Pawar, S. Campioni, C. M. Dobson, F. Chiti and M. Vendruscolo, J. Mol. Biol., 2008, 380, 425–436. (10) F. Chiti and C. M. Dobson, Nature Chem. Biol., 2009, 5, 15–22. (11) J. P. Schmittschmitt and J. M. Scholtz, Protein Sci., 2003, 12, 2374–2378. (12) W. Liu, T. Cellmer, D. Keerl, J. M. Prausnitz and H. W. Blanch, Biotechnology and Bioengineering, 2004, 90, 483–490.


32

Page 33 of 40

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


(13) A. M. Corrigan, C. Mller and M. R. H. Krebs, J. Am. Chem. Soc., 2006, 128, 14740– 14741. (14) N. Javid, K. Vogtt, C. Krywka, M. Tolan and R. Winter, Phys. Rev. Lett., 2007, 99, 028101–1–028101–4. (15) K. E. Routledge, G. G. Tartaglia, G. W. Platt, M. Vendruscolo and S. E. Radford, J. Mol. Biol., 2009, 389, 776–786. (16) R. Mezzenga, J.-M. Jung and J. Adamcik, Langmuir, 2010, 26, 10401–10405. (17) S. E. Hill, T. Miti, T. Richmond and M. Muschol, PLoS ONE, 2011, 6, e18171(1)– e18171(12). (18) A. George and W. Wilson, Acta Crystallogr. D, 1994, 50, 361–365. (19) M. Muschol and F. Rosenberger, J. Chem. Phys., 1997, 107, 1953–1962. (20) D. Rosenbaum, P. C. Zamora and C. F. Zukoski, Phys. Rev. Lett., 1996, 76, 150–153. (21) P. M. Tessier, H. R. Johnson, R. Pazhianur, B. W. Berger, J. L. Prentice, B. J. Bahnson, S. I. Sandler and A. M. Lenhoff, Proteins, 2003, 50, 303–311. (22) W. Liu, D. Bratko, J. M. Prausnitz and H. W. Blanch, Biophys. Chem., 2004, 107, 289– 298. (23) C. M. Dobson, Nature, 2003, 426, 884–890. (24) M. R. H. Krebs, D. K. Wilkins, E. W. Chung, M. C. Pitkeathly, A. K. Chamberlain, J. Zurdo, C. V. Robinson and C. M. Dobson, J. Mol. Biol., 2000, 300, 541–549.


33


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

Page 34 of 40

(25) W. Dzwolak, R. Ravindra, J. Lendermann and R. Winter, Biochemistry, 2003, 42, 11347– 11355. (26) M. Manno, E. F. Craparo, A. Podestà, D. Bulone, R. Carrotta, V. Martorana, G. Tiana and P. L. San Biagio, J. Mol. Biol., 2007, 366, 258–274. (27) C. B. Andersen, H. Yagi, M. Manno, V. Martorana, T. Ban, G. Christiansen, D. E. Otzen, Y. Goto and C. Rischel, Biophys. J., 2009, 96, 1529–1536. (28) F. Sciortino, S. Mossa, E. Zaccarelli and P. Tartaglia, Phys. Rev. Lett., 2004, 93, 055701– 1–055701–4. (29) S. Mossa, F. Sciortino, P. Tartaglia and E. Zaccarelli, Langmuir, 2005, 20, 10756–10763. (30) L. Arnaudov and R. de Vries, Biophys. J., 2005, 88, 515–526. (31) A. Dhulesia, N. Cremades, J. R. Kumita, S.-T. D. Hsu, M. Mossuto, M. Dumoulin, D. Nietlispach, M. Akke, X. Salvatella and C. M. Dobson, J. Am. Chem. Soc., 2010, 132, 15580– 15588. (32) A. K. Buell, A. Dhulesia, M. Mossuto, N. Cremades, J. R. Kumita, M. Dumoulin, M. E. Welland, T. P. J. Knowles, X. Salvatella and C. M. Dobson, J. Am. Chem. Soc., 2011, 133, 7737–7743. (33) D. Bulone, L. G. Fornili, S. L. Fornili, M. Lapis and P. L. San Biagio, Meas. Sci. Technol., 1994, 5, 1443–1447. (34) S. Provencher and J. Gloeckner, Biochemistry, 1981, 20, 33–37. (35) M. B. Huglin, S. J. O’Donohue and M. A. Radwan, Eur. Polym. J., 1989, 25, 543–547.


34

Page 35 of 40

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


(36) K. S. Schmitz, An Introduction to Dynamic Light Scattering by Macromolecules, Academic Press, Inc., San Diego, 1990 (37) S. Provencher, Computer Phys. Communicat., 1982, 27, 229–242. (38) P. Claudy, J. Létoffé, A. Bayol, M. Bonnet and J. Maurizot, Thermochimica Acta, 1992, 207, 227–237. (39) S. Hill, J. Robinson, G. Matthews and M. Muschol, Biophys. J., 2009, 96, 3781–3790. (40) A. Stradner, H. Sedwick, F. Cardinaux, W. C. K. Poon, S. U. Egelhaaf and P. Schurtenberger, Nature, 2004, 432, 492–495. (41) A. K. Chamberlain, C. E. MacPhee, J. Zurdo, L. A. Morozova-Roche, H. A. Hill, C. M. Dobson and J. J. Davis, Biophys. J., 2000, 79, 3282–93. (42) J. P. Hansen and I. R. McDonald, Theory of simple liquids, Academic Press, London, New York, San Diego, 1986. (43) P. N. Pusey, In: Liquid Freezing and Glass Transition, (eds. J. P. Hansen, D. Levesque, and J. Zinn-Justine), North Holland Elsevier Science Publishers B. V., Amsterdam, 1992, pp. 763–942. (44) J. R. Lakowicz, Principles of Fluorescence Spectroscopy, Kluwer Academic/Plenum Publishers, New York, 1999. (45) T. Imoto, L. S. Forster, J. A. Rupley and F. Tanaka, Proc. Natl. Acad. Sci. USA, 1971, 69, 1151–1155.


35


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

Page 36 of 40

(46) E. Nishimoto, S. Yamashita, A. G. Szabo and T. Imoto, Biochemistry, 1998, 37, 5599– 5607. (47) M. D’Amico, S. Raccosta, M. Cannas, V. Martorana and M. Manno, J. Phys. Chem. B, 2011, 115, 4078–4087. (48) S. Goda, K. Takano, Y. Yamagata, R. Nagata, H. Akutsu, S. Maki, K. Namba and K. Yutani, Protein Sci., 2000, 9, 369–375. (49) H. Yan, A. Saiani, J. Gough and A. Miller, Biomacromolecules, 2006, 7, 2776–2782. (50) B. Vernaglia, J. Huang and E. Clark, Biomacromolecules, 2004, 5, 1362–1370. (51) E. Frare, M. Mossuto, P. Polverino de Laureto, S. Tolin, L. Menzer, M. Dumoulin, C. Dobson and F. A., J. Mol. Biol., 2009, 387, 17–27. (52) E. T. Powers and D. L. Powers, Biophys. J., 2008, 94, 379–391. (53) R. Carrotta, M. Manno, D. Bulone, V. Martorana and P. L. San Biagio, J. Biol. Chem., 2005, 280, 30001–30008. (54) A. Lomakin, D. S. Chung, G. B. Benedek and D. A. Kirschner, Proc. Natl. Acad. Sci. USA, 1996, 93, 377–382. (55) T. Rosch and E. J.R., J. Phys. Chem. B, 2007, 111, 12591–12598. (56) D. Nicoli and G. Benedek, Biopolymers, 1976, 15, 2421–2437. (57) S. Arai and M. Hirai, Biophys. J., 1999, 76, 2192–2197.


36

Page 37 of 40

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


(58) E. Y. Chi, S. Krishnan, B. S. Kendrik, B. S. Chang, J. F. Carpenter and T. W. Randolph, Protein Sci., 2003, 12, 903–913. (59) M. Manno, C. Xiao, D. Bulone, V. Martorana and P. L. San Biagio, Phys. Rev. E, 2003, 68, 011904–1–011904–11. (60) M. Manno, D. Bulone, V. Martorana and P. L. San Biagio, J. Phys.: Condens. Matter, 2004, 16, S5023–S5033. (61) O. D. Velev, E. W. Kaler and A. M. Lenhoff, Biophys. J., 1998, 75, 2682–2697. (62) E. R. A. Lima, E. C. Biscaia, M. Bostrøm, F. W. Tavares and J. M. Prausnitz, J. Phys. Chem., 2007, 111, 16055–16059. (63) P. Doherty and G. B. Benedek, J. Chem. Phys., 1974, 61, 5426–5434. (64) A. Parmar and M. Muschol, J. Coll. Int. Sci., 2009, 339, 243–248. (65) R. A. Curtis, C. Steinbrecher, M. Heinemann, H. W. Blanch and J. M. Prausnitz, Biophys. Chem., 2002, 98, 249–265. (66) E. J. W. Verwey and J. T. G. Overbeck, Theory fo the Stability of Lyophobic Colloids, Elsevier, New York, 1948. (67) G. K. J. Batchelor, J. Fluid. Mech., 1976, 74, 1–29. (68) B. U. J. Felderhof, J. Phys. A, 1978, 11, 929–937. (69) C. Tanford, Physical Chemistry of Macromolecules, Wiley, New York, 1961. (70) A. Isihara, J. Chem. Phys., 1950, 18, 1446–1449.


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(71) R. Carrotta, M. Manno, F. M. Giordano, A. Longo, G. Portale, V. Martorana and P. L. San Biagio, Phys. Chem. Chem. Phys., 2009, 11, 4007–4018. (72) G. Manning, J. Chem. Phys., 1969, 51, 924–933. (73) G. Manning, J. Phys. Chem. B, 2007, 111, 8554–8559. (74) A. K. Buell, G. G. Tartaglia, N. R. Birkett, C. A. Waudby, M. Vendruscolo, X. Salvatella, M. E. Welland, C. M. Dobson and T. P. J. Knowles, Chembiochem, 2009, 10, 1309–1312. (75) K. Takano, K. Tsuchimori, Y. Yamagata and K. Yutani, Biochemistry, 2000, 39, 12375– 12381. (76) S. M. Vaiana, M. Manno, A. Emanuele, M. B. Palma-Vittorelli and M. U. Palma, J. Biol. Phys., 2001, 27, 133–145.


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Thermodynamic versus Conformational Metastability in Fibril-Forming

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