The authors have declared that no competing interests exist.
Conceived and designed the experiments: LB PPDL. Performed the experiments: MP GDF LT EF DK M. Brucale IT M. Bisaglia. Analyzed the data: MP GDF LT EF DK M. Brucale M. Bisaglia LB PPDL. Contributed reagents/materials/analysis tools: BS DA. Wrote the paper: LB PPDL.
Current address: Istituto di Biofisica, Consiglio Nazionale delle Ricerche, Povo (TN), Italy
The aggregation of α-synuclein into amyloid fibrils constitutes a key step in the onset of Parkinson's disease. Amyloid fibrils of α-synuclein are the major component of Lewy bodies, histological hallmarks of the disease. Little is known about the mechanism of aggregation of α-synuclein. During this process, α-synuclein forms transient intermediates that are considered to be toxic species. The dimerization of α-synuclein could represent a rate-limiting step in the aggregation of the protein. Here, we analyzed four covalent dimers of α-synuclein, obtained by covalent link of the N-terms, C-terms, tandem cloning of two sequences and tandem juxtaposition in one protein of the 1–104 and 29–140 sequences. Their biophysical properties in solution were determined by CD, FT-IR and NMR spectroscopies. SDS-induced folding was also studied. The fibrils formation was analyzed by ThT and polarization fluorescence assays. Their morphology was investigated by TEM and AFM-based quantitative morphometric analysis. All dimers were found to be devoid of ordered secondary structure under physiological conditions and undergo α-helical transition upon interaction with SDS. All protein species are able to form amyloid-like fibrils. The reciprocal orientation of the α-synuclein monomers in the dimeric constructs affects the kinetics of the aggregation process and a scale of relative amyloidogenic propensity was determined. Structural investigations by FT IR spectroscopy, and proteolytic mapping of the fibril core did not evidence remarkable difference among the species, whereas morphological analyses showed that fibrils formed by dimers display a lower and diversified level of organization in comparison with α-synuclein fibrils. This study demonstrates that although α-synuclein dimerization does not imply the acquisition of a preferred conformation by the participating monomers, it can strongly affect the aggregation properties of the molecules. The results presented highlight a substantial role of the relative orientation of the individual monomer in the definition of the fibril higher structural levels.
α-Synuclein (aS) is a small (140 amino acids) protein, highly expressed in the central nervous system where it constitutes about 0.5–1.0% of the entire cytosolic protein content. In fibrillar form, it is the major component of intracellular deposits of proteins and lipids called Lewy bodies and Lewy neurites, which are hallmarks of Parkinson's disease (PD) and other related neurological disorders
The amino acidic sequence of aS includes a conserved N-terminal domain containing a series of amphipathic repeats that modulates membrane binding; a central region, between residues 61–95, that comprises the highly aggregation-prone NAC sequence
The aggregation of aS could be described as a process, which follows a nucleated polymerization model. Unfolded aS molecules are initially distributed among several related conformations. At the onset of the fibrillization process, soluble species undergo self-assembly into stable oligomeric intermediates able to grow through monomer accretion, thereby evolving into final amyloid-like fibrils
aS stable covalent dimers were obtained through oxidation or nitration mechanisms or via di-tyrosine binding
To elucidate the role of dimerization in the aS aggregation process, we produced and characterized four dimers of aS, obtained by covalently linking two aS monomers via their terminals. A cysteine residue was added at either the N- or C-terminal of aS, providing head-to-head “NN” or tail-to-tail “CC” dimerization through the formation of a disulfide bond. An “NC” dimer, formed by a tandem repeat of the aS sequence, was obtained as a single polypeptide chain. Another dimer, called “double core” or “DC” dimer, is constituted by two consecutive central, highly amyloidogenic regions, containing aS residues from 1 to 104 and from 29 to 140. This last dimer was conceived to both draw up the hydrophobic and amyloidogenic regions, avoiding the interferences of the terminal parts, and to impose a constrain of proximity between the two amyloidogenic regions of aS.
Our study demonstrates that bringing aS molecules in close proximity is not a sufficient condition for the protein to assume a specific conformation. On the other hand, the relative orientation of each molecule in the dimer plays a critical role in the structural arrangement of aS fibrils.
Proteinase K from
The pET28b (Novagen) plasmid was used for the expression of recombinant human aS in
Native polyacrilamide gel electrophoresis was performed on 12% polyacrylamide gel, with a constant current of 25 mA, using a Mini–PROTEIN II Bio-Rad electrophoresis system (Hercules, CA). 5 µg of protein were loaded into each well, and the protein bands were visualized by Coomassie blue staining.
Reverse phase-high pressures liquid chromatography (RP-HPLC), was conducted on a Jupiter C4 column (4.6×150 mm, Phenomenex, USA). Elution was obtained with a linear gradient of acetonitrile (0.085% TFA) vs water (0.1% TFA), from 10 to 40% in 5 min and from 40 to 45% in 40 min at a flow rate of 1 ml/min. The identity of the protein material was assessed by mass spectrometry using an ESI-QTOF Micro instrument (Waters, Milford, MS, USA). Gel filtration chromatography (GF) was performed with a Superdex 200 10/300GL column (Amersham Biosciences, Uppsala, Sweden), using an ÄKTA FPLC system (Amersham Biosciences, Uppsala, Sweden), eluting at 0.4 ml/min in 20 mM in Tris-HCl, 150 mM NaCl, pH 7.4, and recording the absorbance at 214 nm. 50 µg of each protein samples (aS, NN, CC, NC and DC dimer), dissolved in the eluting buffer, were loaded into the column. The hydrodynamic volume of the analytes was determined on the basis of their distribution coefficients, Kd, calculated as the following formula: Kd = (Ve-V0)/(Vt-V0), where Ve, Vt and V0 are the standard proteins, the total and the void elution volume, respectively. The calibration function (y = 1.15–0.38×) was obtained by plotting Kd against the logarithm of the molecular weight of proteins. A mixture of proteins of known MW was used (bovine α-lactalbumin, 14 kDa; carbonic anhydrase, 29 kDa; ovalbumin, 45 kDa; bovine serum albumin, 66 kDa; β-amylase, 200 kDa; apoferritin, 443 kDa and, thyroglobulin, 669 kDa). Blue dextran and 0.05% dimethyl sulfoxide (DMSO) were loaded to estimate V0 and Vt.
Protein concentrations were determined by absorption measurements at 280 nm using a double-beam Lambda-20 spectrophotometer (Perkin Elmer, Norwalk, CT). The extinction coefficients of the proteins at 280 nm are 5960 (aS), 12045 (NN and CC dimer), 11920 (NC dimer) and 7450 M−1⋅cm−1 (DC dimer), as evaluated from their amino acid composition by the method of Gill and von Hippel
Deuterated aS and dimers were prepared by three cycles of dissolution in D2O, filtration with a 20 nm pore-size filter (Whatman, Maidstone, UK), freezing of the protein solution at −80°C, followed by lyophilisation. The spectra of aS and dimers in solution were registered after dissolving the deuterated proteins in 20 mM Tris·DCl, 150 mM NaCl, pH* 7.2 (uncorrected for isotopic effects) at a concentration of ∼5 mg/ml. Spectra were recorded at 20–22°C using a Perkin Elmer 1720× spectrometer (Norwalk, CT, USA), purged with a continuous flow of N2 gas. Protein samples were placed between a pair of CaF2 windows separated by a 50 µm Mylar spacer. For each protein sample, 50 interferograms were accumulated at a spectral resolution of 2 cm−1. The spectra were analyzed using the Grams 32 program version 4.14 (Galactic Industries Corporation, Salem, NH). Buffer spectra were recorded under identical conditions to those used for protein samples and subtracted from the spectra of the latter. The second derivative of the amide I and II bands was used to identify the different spectral components. Thereafter, curve fitting was performed with Gaussian and Lorentzian line shapes, and with bandwidths varying between 15 and 25 cm−1
FT-IR spectra of dimer ultra-centrifuged fibrils (90000 rpm for 2 hours at 4°C) were also recorded. Fibrils were obtained after 4–6 weeks of incubation of deuterated protein in the buffer previously described, at a protein concentration of 70 µM, at 37°C under shaking at 500 rpm in a thermo-mixer (Compact, Eppendorf, Hamburg, DE).
The expression of aS dimers for NMR studies (15N-labeled proteins) was achieved by growing cells in M9 minimal medium. Heteronuclear Single Quantum Correlation (HSQC) spectra were acquired on a Bruker Avance DMX spectrometer equipped with a gradient triple resonance probe. 15N-labelled aS samples (100–350 µM) were dissolved in PBS buffer containing 10% D2O. The experiments (256 increments of 512 time points each) were acquired at 283 K with 16 transients. The spectral widths were 3 ppm (1H) and 22 ppm (15N) and the frequency offsets were 8 ppm (1H) and 116 ppm (15N). Prior to Fourier transformation, the data were multiplied by a 90° shifted
70 µM aSG141C142 was mixed with 5 molar excess of tris(2-carboxyethyl)phosphine in 20 mM Tris buffer, pH 7.0. After 30 min, 5 fold molar excess Oregon Green 488 maleimide (Molecular Probes, Invitrogen, CA, USA) previously dissolved in 20 mM Tris pH 7.0, was added to aSG141C142 solution. The reaction was held 4 hour at 45°C, then was analyzed by RP-HPLC by using a Jupiter C4 column (Phenomenex, CA, USA). Elution was obtained with a linear gradient of acetonitrile (0.085% TFA) vs water (0.1% TFA), from 39 to 46% in 14 min at a flow rate of 0.6 ml/min. Fluorescence polarization (FP) measurements were performed with a fluorescence plate reader DTX 880 Multimode Detector (Beckman Coulter, IN, USA), equipped with an excitation filter at 485 nm and two emission filters at 535 nm. Excitation and emission filters were equipped with polarizer. Measurements were carried out at 37°C, 1000 rpm shaking. 1 mg/ml of protein species was dissolved in PBS, 0.05% sodium azide, and mixed with 1/100 molar ratio of aSG141C142 labelled with Oregon Green 488. Time points were collected every 6 or 12 hours and FP (y) values were plotted against the incubation time (t). The data were fitted with a sigmoidal curve that allowed to evaluate the slope at the flex point (t50) and the length of the lag-phase.
Protein aggregation was carried out in PBS buffer. The protein solutions were filtered with a 0.22 µm pore-size filter (Millipore, Bedford, MA, USA) and incubated at 37°C for up to 14 days at a protein concentration of 70 µM (1 mg/ml), under shaking at 500 rpm in a thermo-mixer (Compact, Eppendorf, Hamburg, DE). Aliquots of the samples were withdrawn during incubation for further analysis.
The ThT binding assays were performed accordingly to LeVine
Aliquots of the aggregation samples after 20 days of incubation were examined by transmission electron microscopy (TEM). The samples were diluted 3 times with PBS. A drop of the samples solution was placed on a Butvar-coated copper grid (400-square mesh) (TAAB-Laboratories Equipment Ltd, Berks, UK), dried and negatively stained with a drop of uranyl acetate solution (1%, w/v). TEM pictures were taken on a Tecnai G2 12 Twin instrument (FEI Company, Hillsboro, OR, USA), operating at an excitation voltage of 100 kV).
For AFM analysis, aliquots (10 µl) from aggregation mixtures were diluted in 90 µl of PBS buffer, left to equilibrate at room temperature for 10 min then deposited on freshly cleaved mica (RubyRed Mica Sheets, Electron Microscopy Sciences, Fort Washington, USA) and left to adsorb for 2 min. The mica surface was then rinsed with ∼500 µl of ultrapure water and dried under nitrogen. AFM imaging was performed in tapping mode with NSC15 phosphorous-doped silicon probes (MikroMasch, Tallin, Estonia) on a NanoScope IIIa SFM system equipped with a Multimode head and a type E piezoelectric scanner (Veeco, Santa Barbara, CA, USA). Raw SFM images were processed only for background removal (flattening) using Gwyddion v2.26. Diameters of the aggregates were measured from the images via a custom script
Proteolysis experiments were carried out on ultra-centrifuged (90000 rpm for 2 hours at 4°C) aggregation samples obtained after 1 month of incubation. Fibrils were dissolved in PBS, pH 7.4 and treated with proteinase K or porcine trypsin (E/S ratio of 1∶1000 and 1∶50, by weight, respectively). The reactions were quenched after 2 hours by acidification with TFA in water (4% v/v). The proteolysis mixtures were ultra-centrifuged. The soluble fractions were directly analyzed by RP-HPLC, while the pellets were incubated over night with 7.4 M Gnd-HCl before the analysis. RP-HPLC analyses were conducted using a Vydac C18 column (4.6 mm×250 mm; The Separations Group, Hesperia, CA), eluted with a gradient of acetonitrile/0.085% TFA vs. water/0.1% TFA from 5 to 25% in 5 min, from 25 to 28% in 13 min, from 28 to 39% in 3 min, from 39 to 45% in 21 min at a flow rate of 1 ml/min. The sites of cleavage along the polypeptide chains were identified by mass spectrometry analyses of the protein fragments purified by RP-HPLC.
The produced aS dimers were identified by MS and isolated to homogeneity. Their chemical characterization is thoroughly described in the Supplementary Material (
The secondary structure of NN, CC, NC and DC in PBS buffer was evaluated by far-UV CD, FT-IR and NMR spectroscopy. The far UV CD spectra, recorded between 198 and 250 nm (
The spectra were recorded in PBS buffer pH 7.4 at a protein concentration of 5 µM, using a quartz cuvette with 1 mm of pathlength. NN, CC, NC and DC are represented respectively by a long, medium, short dash and dotted line. Inset: Far UV CD of aS (continuous line).
For FT-IR spectra, deuterated proteins were dissolved in a saline buffer (20 mM Tris·DCl, 150 mM NaCl, pH* 7.2, uncorrected for isotopic effects), and the IR absorbance was recorded between 1500 and 1750 cm−1 to evaluate the contribution of both Amide I and II bands (
Curve fitting was performed with Gaussian and Lorentzian lineshapes and with bandwidths varying between 15 and 20 cm−1. The peak position of the amide band components was deduced from the second derivative spectra (
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1530 | β-sheet/turns | 10 | 3 | 17 | 5 | 3 | - |
1560–1568 | Glu (COO−) | 12 | 25 | 25 | 22 | 14 | 8 |
1580–1586 | Asp (COO−) | 9 | 5 | 6 | 7 | 7 | - |
1610–1612 | Tyr/β-sheet | - | 12 | 21 | 12 | 5 | - |
1640 | Random | 55 | 38 | 17 | 42 | 54 | 69 |
1656–1671 | Turns | 14 | 17 | 14 | 12 | 17 | 23 |
Peak position of the amide I band components, as deduced by the second derivative spectra.
Percentage area of the amide I band components, as obtained by integrating the area under each deconvoluted band.
To verify whether dimerization can promote interactions, 1H-15N HSQC spectra of aS dimers were recorded (
aS HSQS spectrum is reported with resonance assignments, where space permits. In red are indicated new peaks of dimers spectra that are not present in aS spectrum. For CC dimer, experiments were performed in the absence and in the presence of 10 mM DTT and the difference maps (CC-dm) was calculated by subtracting the data matrix obtained after 2 hours of incubation in the presence of DTT from the reference.
aS is able to acquire a α-helix conformation in the presence of SDS. In the titration experiments over a range of SDS concentration (0–2 mM), the presence of an isodichroic point at 203 nm suggested for aS a simple two-state conformational transition between random coil and α-helix
NN, CC, NC and DC are represented respectively by black triangle up, black triangle down, empty square, empty circle, respectively. SDS dependence curve of aS is also reported (black circle).
The kinetics of fibril formation of aS dimers was monitored using Thioflavin T (ThT) fluorescence (
The aggregation processes were conducted at a protein concentration of 1 mg/ml. Fluorescence intensity of ThT is reported as percentage of the plateau of emission intensity corresponding to each curve. NN, CC, NC and DC are represented respectively by black triangle up, black triangle down, empty square, empty circle, respectively. aS aggregation curve is indicated by black circles. Error bars were calculated from three independent aggregation experiments. For FP, lag-phase, (white bars) and curve slope at t50 (black bars) for NN, CC and aS were reported. The values of lag phase and slope were deduced from four independent aggregation experiments.
The aggregation properties of NN and CC dimers were tested with a complementary technique based on fluorescence polarization (FP), which intensity is proportional to the diffusion coefficient of the fluorescent probe linked to the protein
FT-IR spectra were recorded to evaluate the type and content of secondary structure in the proteins aggregates (
Protein fibrils obtained after one month of incubation were dissolved in 20 mM Tris·DCl, 150 mM NaCl pH* 7.2. The peak position of the amide band components was deduced from the second derivative spectra (
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1560–1568 | Glu (COO−) | 23 | 33 | 48 | 3 | 8 |
1580–1586 | Asp (COO−) | 5 | - | - | 3 | 4 |
1612–1619 | Aggregated β-sheet | 31 | 18 | 13 | 32 | 46 |
1634–1636 | β-sheet | 18 | - | 18 | - | - |
1640–1650 | Random | - | 33 | 11 | 38 | 16 |
1656–1668 | Turns | 19 | 15 | 9 | 22 | 24 |
1680–1689 | Anti-parallel aggregated β-sheet | 4 | 1 | 1 | 2 | 2 |
Peak position of the amide I band components, as deduced by the second derivative spectra.
Percentage area of the amide I band components, as obtained by integrating the area under each deconvoluted band. The areas corresponding to side chain contributions located at 1700–1710 cm−1 have not been considered.
The morphology of aggregates formed by all the dimers was investigated by transmission electron (TEM) and atomic force (AFM) microscopy (
Right column: distributions of the apparent diameters of the fibrillar aggregates, as measured from AFM images via an automated procedure (see main text). Gaussian fits are shown (solid black lines) where applicable.
Fibrils formed by aS have a measurable periodicity of 82±7 nm as estimated by AFM height profiles, and clearly appear as formed by two left-hand-twisted filaments in TEM images. Fibrils formed by NN and CC dimers instead appear as single filaments even at the maximum AFM and TEM resolutions available to us. NC fibrils rarely show a periodicity of ∼80 nm, suggesting that they are also formed by twisted sub-fibrillar filaments, but the vast majority of NC aggregates does not show such features, thus hindering statistically significant morphological observations on this specific aspect (
To define the region(s) of the proteins involved in the core of the fibrils formed by the different types of dimers, an approach based on proteolytic digestion coupled to mass spectrometry was used. Regions of the protein sequence normally exposed to proteases exhibit limited accessibility when involved in β-sheet structure
Sequences cleaved off are represented in white, protected regions are indicated in black. The numbers refer to N-terminal of each peptide. As DC is constituted by the sequence 1–104 linked to 29–140 of aS, the sequence notation of its proteolytic fragments refers to the aS sequence numbering. In the case of fragments encompassing the two sequences, these were indicated with numbering of the segments deriving from both sequences separated by a slash.
DC dimer sequence is constituted by 1–104 linked to 29–140 of aS. The sequence notation reported for its proteolytic fragments refers to aS sequence numbering, therefore, the fragments encompassing the two sequences were indicated with the numbering of the segments deriving from both sequences separated by a slash.
This study was aimed to address two issues. The first one refers to the possibility to speed up the aggregation process thought an increase of local concentration of aS by generating covalent dimers. The second issue explored is the topology of the individual aS molecules as building blocks of the aggregated fibers. aS dimers formation has been reported in the literature as product of redox chemistry, as described as an example for Tyr125 based dimerization upon nitrative stress
In our study the procedure used to produce aS dimers relies on two strategies: the first was focused on the formation of dimers with a minimal number of constraints on the fibrillation process. aS molecules were linked with different reciprocal orientations through their N-terminals (V3C), the C-terminals (adding 141G and 142C) or directly cloned in tandem. The second strategy led to the production of a fourth dimer, in which the two fibrils core regions (residues 1–104/29–140) were juxtaposed. The dimerization of aS
The kinetic properties of the aggregation process, for the different dimer types, were first analyzed in terms of fibril formation. The ThT assay, that reports the growth of fibrils, showed only for the DC an aggregation behavior characterized by the presence of a shorter lag phase in comparison to length observed for aS and all other dimers, thus suggesting that the all the latter aggregate following a qualitatively similar nucleated polymerization process. Differences emerged in the fibril elongation phase starting immediately after the lag phase, as the CC dimer presented a steeper rise in the ThT fluorescence signal, which could be interpreted as a facilitated accretion of the dimer building blocks on the fibrils as a consequence of the topology of the individual aS molecules in the fibrils. This possibility will be discussed again later within the context of the proposed structural model for the fibrillar assemblies. The DC dimer presents a more aggregation prone behavior, but the end product seems to differ from the mature fibrils formed by aS, an observation that hinders the direct comparison of the kinetic data from these species.
To explore an earlier stage in the aggregation process, we analyzed the formation of oligomers as detected by fluorescence polarization assay. Using a protocol described by Luk et al.
The quantitative morphological characterization of aS fibrillar products based on TEM and AFM images is in accord with the fibril structure model first proposed by Fink and coworkers
The structural characterization of the fibrillation products of the dimers may provide the rationale to contextualize the results presented here in the light of the recent SSNMR studies on aS fibrils. Several laboratories focused on the definition of the aS monomer structure within fibrils
The last evidence that provides further support to the model proposed by Vilar et al.
In conclusion, this study allows to propose both a tentative scheme of the tertiary structure organization of the monomers in the fibrils, in which as previously proposed
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