Biophysical groundwork as a hinge to unravel the biology of α-synuclein aggregation and toxicity

Biophysical groundwork as a hinge to unravelthe biology of α-synuclein aggregationand toxicity

Nicoletta Plotegher, Elisa Greggio, Marco Bisaglia and Luigi Bubacco*Molecular Physiology and Biophysics Unit, Department of Biology, University of Padova, Padova, Italy

Abstract. Alpha-synuclein (aS) and its aggregation properties are central in the developmentand spread of Parkinson’s disease. Point mutations and multiplications of the SNCA gene encodingaS cause autosomal dominant forms of the disorder. Moreover, protein inclusions found in thesurviving neurons of parkinsonian brains consist mainly of a fibrillar form of aS. Aggregates of aS,which form a transient, complex and heterogeneous ensemble, participate in a wide variety oftoxic mechanisms that may be amplified by aS spreading among neighbouring neurons. Recently,significant effort has been directed into the study of the aS aggregation process and the impact ofaS aggregates on neuron survival. In this review, we present state-of-the-art biophysical studies onthe aS aggregation process in vitro and in cellular models. We comprehensively review the newinsights generated by the recent biophysical investigations, which could provide a solid basis fromwhich to design future biomedical studies. The diverse cellular models of aS toxicity and theirpotential use in the biophysical investigation are also discussed.

1. Introduction 2

2. Aggregation kinetics 52.1. Fluorescent dyes for amyloid fibrils 62.2. Monitoring aggregation through the intrinsic properties of aS 7

2.2.1. Absorbance variation in aggregating aS solution 72.2.2. Dynamic light scattering 82.2.3. Intrinsic tryptophan fluorescence 8

2.3. Fluorophores covalently bound to aS monomers 92.3.1. Fluorescence correlation spectroscopy 92.3.2. Fluorescence polarization 9

2.4. Novel approaches 92.5. Aggregation inhibitors and enhancers 112.6. Kinetic models 12

3. Monomeric aS conformations 143.1. Monomeric aS structural analysis 15

3.1.1. Paramagnetic relaxation enhancement-NMR measurements and molecular dynamics (MD)simulation 15

* Author for Correspondence: Luigi Bubacco, Molecular Physiology and Biophysics Unit, Departmentof Biology, University of Padova, Via U. Bassi 58B, 35121 Padova, Italy.Fax: +390498276300; Email: [email protected]

Quarterly Reviews of Biophysics 47, 1 (2014), pp. 1–48. © Cambridge University Press 2014 1doi:10.1017/S0033583513000097 Printed in the United States of America

3.1.2. Single-molecule force spectroscopy experiments 163.1.3. Fluorescence measurements on aggregation prone monomeric aS conformers 16

3.2. aS conformation and aggregation on membranes 17

4. aS oligomeric species 184.1. Physical methods for obtaining aS oligomers 214.2. Chemical oligomers 22

5. aS fibrils: structure and properties 235.1. Fibril structure 23

5.1.1. Fourier transform infrared and small angle X-ray spectroscopy measurements 235.1.2. NMR and EPR to probe aS fibril structure 24

5.2. Fibril imaging 255.2.1. Fibrils observed with TEM 265.2.2. Fibrils measured by AFM 265.2.3. Super resolution fluorescence imaging of aS fibrils 27

5.3. Fibril modelling 28

6. Aggregation in cell models 296.1. Aggregation kinetics and aggregate properties in cells 29

6.1.1. aS fused with fluorescent proteins (FPs) or tags 306.1.2. TEM and ThS staining on fixed cells 316.1.3. Advanced fluorescence microscopy techniques applied to the study of aS aggregation

in cells 316.2. aS aggregates toxicity 336.3. Misfolded aS transmission in cell models 35

7. Conclusions 37

8. Acknowledgements 38

9. References 38

1. Introduction

Protein misfolding and aggregation are related to many diverse and severe pathologies, severalof which are neurodegenerative disorders (Ross & Poirier, 2004). Considering that age is arisk factor for most neurodegenerative diseases and considering the expected increase in theaverage lifespan, modern biomedical science is intensely committed to studying the molecularmechanisms responsible for neurodegeneration. In particular, biophysicists, molecular and cellbiologists and clinicians are exploiting a multitude of different approaches to tackle this issue.One of the most studied neurodegenerative disorders linked to protein aggregation is

Parkinson’s disease (PD). The relevant misfolding protein is the 140 amino acid α-synuclein(aS). aS aggregation is not only related to PD etiopathogenesis but also to other neurodegenera-tive disorders, i.e., dementia with Lewy Bodies (LBs), multiple system atrophy (MSA) andAlzheimer’s disease (AD). These diseases, because of their shared aS involvement, are collectivelytermed synucleinopathies (Trojanowski & Lee, 2003).The primary structure of aS (Fig. 1) is divided into three main regions: (i) the N-terminus

(N-term), including the first 60 amino acids, (ii) the non-beta amyloid component (NAC) regionfrom residue 61 to residue 95, which is highly hydrophobic and acquires a β-sheet structure when

2 N. Plotegher et al.

aS forms amyloid fibrils (Weinreb et al. 1996), and (iii) the C-terminus (C-term), which includes14 negatively charged acidic residues and does not exhibit a defined secondary structure. TheN-term together with the NAC region can form an amphipathic α-helix that is responsible forprotein–membranes interactions (Bartels et al. 2010) (see Section 4.1). As will be discussedbelow, most studies suggest that aS has a disordered structure in solution under physiologicalconditions. Upon binding to phospholipid vesicles and lipid membranes, the N-term residuesadopt a helical structure; conversely, aS seems to acquire a β-sheet structure in the fibrilsfound in the LBs in parkinsonian brains.aS is widely expressed in the central nervous system and is mainly localised in the presynaptic

terminals, in both cytosolic and membrane-bound forms (Clayton & George, 1999). Human aSbelongs to the synuclein protein family, which includes two other proteins, β-synuclein andγ-synuclein. The other two synucleins share a high sequence identity with aS (52 and 63%,respectively) and exhibit similar expression patterns (Clayton & George, 1998).The physiological functions of aS are still poorly understood, but mounting evidence ascribes

to the protein a role in synaptic vesicle recycling and docking, Soluble NSF Attachment ProteinReceptor (SNARE) complex assembly (Burre et al. 2010) and neurotransmitter release (Liu et al.2004). aS has been shown to involved in synaptic plasticity and learning (Clayton & George,1998), but its activity is likely dispensable, as knock-out mice are viable and present only subtlealterations in dopamine (DA) release and reduced levels of striatal DA (Abeliovich et al. 2000),supporting the hypothesis that the protein is a DA release modulator. Moreover, it has beenshown that aS can interact with microtubules (Esposito et al. 2007) and actin (Alim et al.2004) and can affect actin structure and dynamics (Sousa et al. 2009). Interestingly, aS knock-outmice also showed impairment in working and spatial long-term memory, as evaluated by the

-+

20 40 100 12060

Ser87 Ser129

N C

N-term NAC C-term

1 61 95 140

A30P E46KH50Q

G51D

A53T

80

Fig. 1. Schematic representation of aS sequence. The three regions in which the protein is typically dividedare reported: (i) amino acids 1–60 correspond to the N-term, an amphipathic region responsible for aSmembrane interactions, which acquires an α-helical structure together with part of the central region ofthe protein, in the presence of detergents or when bound to lipid membranes. As shown, all thepathological mutations of aS identified so far are located at the N-term (A30P, E46K, H50Q, G51Dand A53T). (ii) The NAC region (non-beta amyloid component) is the part of the protein that drives thefirst steps of the aggregation process and folds into a β-sheet secondary structure when aS aggregatesand form amyloid fibrils. (iii) The C-term is characterized by the presence of several negative chargesand by the fact that it does not acquire any defined secondary structure features, neither in the fibrilsnor when aS is bound to the membranes. The protein sequence shows seven repeats with anapolipoproteins lipid-binding motif [EGS]-K-T-K-[EQ]-[GQ]-V-XXXX, which are represented in thefigures as red rectangles (www.uniprot.org). The schema reports also aS charges distribution and theposition of the two serines found to be relevant to PD because phosphorylated in parkinsonian brainsor PD models.

Biophysical properties of α-synuclein 3

behavioural tests (Kokhan et al. 2012). However, neuronal dysfunctions, alterations in synapticstructures and decreased survival rate were observed in triple knock-out mice lacking all threesynuclein isoforms, revealing the importance of these three proteins for the functionality ofthe central nervous system (Greten-Harrison et al. 2010).PD pathogenesis is mainly sporadic, but ∼5–10% of PD cases are linked to mutations in single

genes. Autosomal recessive, early-onset forms of this disorder are inherited through mutationsin the PINK1, parkin or DJ-1 genes, which encode proteins involved in the maintenance of mito-chondrial function during oxidative stress (Berman & Hastings, 1999). Autosomal dominant,late-onset PD, on the other hand, is caused by mutations in the Leucine-rich repeat kinase 2(LRRK2) gene (Funayama et al. 2002), the aS-coding SNCA gene (Kasten & Klein, 2013) andVacuolar Sorting Protein 35 (VPS35) (Zimprich et al. 2011).Monogenic forms of PD closely resemble idiopathic cases and represent a rich opportunity for

revealing the nature of PD pathology. Both idiopathic and genetic forms of PD are characterizedby progressive death of dopaminergic neurons in the substantia nigra pars compacta. However, notall dopaminergic neurons in the brain are affected (e.g., dopaminergic neurons of the ventraltegmental area are spared), and other brain regions seem to be affected at later stages(Dickson, 2007). Functional imaging (i.e., positron emission tomography) suggests that at least70% of the dopaminergic neurons must degenerate before manifestation of the first symptomsof parkinsonism (Brooks, 1998).The evidence that linked aS and genetic PD was the discovery of single point mutations, i.e.,

A30P, E46K and A53T (Kruger et al. 1998; Polymeropoulos et al. 1997; Zarranz et al. 2004), inthe gene SNCA and the gene’s duplication (Chartier-Harlin et al. 2004) and triplication (Singletonet al. 2003) leading to autosomal dominant forms of the disease. Recently, two other mutations inthe central region of aS (G51D and H50Q) were associated with familial PD (Kiely et al. 2013;Proukakis et al. 2013). Susceptibility to PD is increased in the presence of risk variants on theSNCA gene, i.e., in the 3′UTR or the promoter region (Fuchs et al. 2008).Protein and lipid aggregates, termed LBs and Lewy Neurites (LNs), are found in the surviving

neurons in brains of patients with a PD diagnosis. The observation that the main constituent ofLBs and LNs is an aggregated fibrillar β-sheet-rich form of aS is additional evidence linking theprotein to PD (Hirsch et al. 1988; Spillantini et al. 1997; Takeda et al. 1998).aS and its propensity to aggregate seem to be crucial in PD etiopathogenesis: in the past years,

several studies have attempted to unravel the mechanisms of aS oligomerization and fibril forma-tion and, more importantly, to define the relationship between these processes and neuronaldeath. Unfortunately, the molecular mechanism of aS aggregation has not been completely elu-cidated in vitro and remains quite obscure in vivo. To further complicate this matter, the processseems to be heavily influenced by experimental conditions (Giehm et al. 2011a), which can lead todifferent aggregation products and kinetic parameters. With these premises, it is clear that study-ing aS aggregation in cell and animal models is important to pin down the (patho)physiologicallyrelevant aS aggregation products and conditions. Several diverse biophysical approaches have re-cently been developed and used to study aS aggregation. Fluorescence spectroscopy, microscopyand scattering techniques have been exploited to evaluate the kinetics of the aggregation processand to describe the properties of both aggregation intermediates and fibrillar end products.Moreover, a large number of possibly toxicity mechanisms related to aS aggregation have beenproposed, but the actual causes of neuronal death are still objects of debate. The gain-of-function,toxic mechanism(s) related to aS mutations or aggregate formation is most likely responsible foraS-induced cell death and should be further elucidated.


In this review, we will try to summarize the key findings on this challenging topic, with the aimof reconstructing the plethora of available biophysical data into a comprehensive biological frame.Moreover, we will suggest important biological question in this field that could be dissected usingbiophysical techniques.In particular, the structural characterization of oligomers remains a tricky but crucial issue,

considering their heterogeneity and transient nature. Structural information of oligomers,which can be effectively studied using biophysical methods, is necessary to define the relationshipbetween aS oligomer-induced toxicity and their properties.

2. Aggregation kinetics

Unfolded monomeric aS, in extreme or pathological conditions, tends to acquire a transientβ-sheet structure prone to aggregation. The first steps of self-interaction of aS monomersoccur rarely, but after an initial lag phase, monomers start to aggregate onto the newly formednucleation centres to form oligomeric species. As soon as the oligomer concentration reachesa critical point, they are rapidly converted into protofilaments, protofibrils and, finally, amyloidfibrils. The kinetic properties that govern aS amyloid formation can be described by a sigmoidaltime dependence: an initial lag phase, suggesting a nucleation mechanism (see Section 1.2.2)where oligomers act as nuclei; a subsequent phase of rapid oligomer assembly into higher mol-ecular weight species, resulting in a decrease in oligomer concentration and an increase in fibrillarspecies concentrations; and finally, the process reaches a plateau when mature fibrils become themajority of the polymers in solution and continue their growth. However, the process describedabove emerges mainly from in vitro experiments and is likely more complex in the cellular contest,which is characterized by different competing ligands and pathways (Kaylor et al. 2005), leading tomultiple forms of aggregation intermediates, oligomers and fibrils.Among these different aS aggregation products, oligomers are the least well-defined mainly

because of their ample heterogeneity: it has been shown (see Section 4 for details) that aS oligo-mers can acquire ensembles of different secondary structures and morphologies depending onthe conditions under which they are formed. Therefore, reaction conditions cannot be used tounambiguously define or classify oligomers. The same can be said for a classification based onoligomers’ structural properties because the structural differences among the ensemble of oligo-mers do not allow discrimination of individual species. One possibility is to define aS oligomersas all the species that are still able to revert to monomers in a chemical equilibrium. However, thisdefinition does not account for those aS oligomers that are off-pathway in the aggregation pro-cess and that neither reverts to aS monomers nor converts to aS fibrils. For these reasons, thedefinition of aS oligomeric species may not be unequivocal and remains an open question thatneeds to be answered. Conversely, protofilaments, protofibrils and fibrils are more homogeneousand experimentally manageable species, as will be discussed in Section 5.The primary forces that drive aS fibrillization are hydrophobic interactions due to a hydro-

phobic stretch in the middle of the aS sequence (from residues 71 to 82), which is composedof 12 amino acids (VTGVTAVAQKTV) (Giasson et al. 2001). In this paper, the authors showedhow the introduction of a charged residue in this stretch of the fibril core region, i.e., A76R orA76E, decreased the fibrillization rate in vitro. Moreover, a synthetic peptide corresponding to thehydrophobic stretch mentioned above can form filaments when aggregated alone. It should bementioned that β-synuclein, which seems not to aggregate in vitro, lacks this hydrophobic stretchof amino acids (Yamin et al. 2005). Interestingly, it has been shown that β-synuclein can be


induced to fibrillize in particular conditions (metal ions, pesticides), further suggesting that evennon-amyloidogenic proteins can be forced to form fibrils under certain circumstances (Suk et al.2010; Yamin et al. 2005). A recent report suggests that β-synuclein behaves similarly to aS: itaggregates and forms amyloid fibrils, and oligomeric aggregation intermediates are able to per-meabilize membranes and to damage mitochondria, causing neuronal death (Taschenbergeret al. 2013).The most commonly used technique to study the kinetics of aS aggregation in vitro is fluores-

cence spectroscopy because measuring the variation in the fluorescence properties of suitableexogenous fluorophores over time allows quantitative analysis of the aS aggregation process.This type of data provides the statistical basis for the comparison of different numericalmodels proposed to describe the kinetic properties of aS aggregation (Morris et al. 2009).Moreover, comparative analysis of the aggregation data obtained under different conditionsled to the identification of the relevant parameters of aS fibrillization in vitro, i.e., protein con-centrations, pH values, buffer conditions, temperature, agitation speed and type and presenceof solution–air interfaces (Giehm & Otzen, 2010).

2.1 Fluorescent dyes for amyloid fibrils

In this section, we will present an overview of the fluorescent dyes used to study the time depen-dence of amyloid fibril formation. The fluorescence emission of the dye Thioflavin T (ThT)increases upon binding to β-sheet structures. Therefore, measuring ThT fluorescence intensityvariations as a function of time in an aggregation assay has allowed the monitoring of β-sheetformation and thereby the fraction of aS fibrils. ThT is typically excited at 450 nm, and the emis-sion spectrum is recorded between 460 and 550 nm, with a maximum at 480 nm. The fluores-cence intensities of ThT and Thioflavin S (ThS) are usually measured in vitro (LeVine, 1993)and in cells or tissue (Luk et al. 2009), respectively. However, ThS can also be used in vitro;there are two main differences between the two dyes, which are both derived from the samechemical precursor: ThT undergoes a change in its emission spectrum and is positively chargedunder binding conditions, whereas ThS has a negative charge under the same conditions and itsemission spectrum only increases in intensity in the presence of β-sheets (LeVine, 1993). ThT ispreferred over ThS for in vitro assays because the shifted ThT emission peak in the presence ofamyloid is more easily distinguished from the background fluorescence of free dye in solution. Toour knowledge, it is still unclear why ThT is not used to stain cells or tissues, but it may be relatedto its reduced binding specificity in a complex environment.The binding mechanism of ThT to amyloids is not fully understood; Krebs et al. (2005)

proposed that the ThT molecules intercalate perpendicularly to the backbone of aS in aβ-sheet conformation and therefore are parallel to the long axes of protofilaments, protofibrilsor fibrils. However, even though the binding between aS fibrils and ThT shows high specificity,no binding constants or evidence for the mode of binding aS fibrils have been reported, althoughthe binding constants were estimated using absorbance spectroscopy data of ThT and fibrilsolution for the amyloids formed by lysozyme (i.e., 7.5×106 and 5.6×104 M−1) (Sulatskayaet al. 2011). A crystallographic structure of ThT bound to β-2 microglobulin amyloid-like aggre-gates allowed the definition of the photophysical properties of ThT when bound to β-sheets(Wolfe et al. 2010). The authors proposed that the fluorophore’s electronic ground state distri-bution is stabilized sterically and electrostatically by β-sheets, that emission due to non-radiativerelaxation is minimized, and that consequentially the quantum yield increases. However, it should


be mentioned that despite its use in several studies to evaluate the kinetic properties of aS aggre-gation, emission measurements of ThT fluorescence often show variable emission efficiency andlow reproducibility. These difficulties can only in part be ascribed to the intrinsic variability in thekinetics of the aS aggregation process. Indeed, it has been shown that ThT micelles can be pres-ent in solution and enhance fluorescence intensity (Khurana et al. 2005), even in the absence ofthe β-sheet structure. It has also been proposed that the surfaces responsible for the interactionwith ThT can be shielded upon conversion of protofilaments to protofibrils and fibrils, causinga reduced ThT emission intensity due to decreased binding (Krebs et al. 2005). These findingssuggest that this method may not provide a precise determination of the kinetic properties ofamyloid fibril formation, not only in the case of aS, in which part of the variability is due toaS, but also for the other amyloidogenic proteins.Furthermore, the presence of other exogenous compounds, such as molecular chaperones or

molecules designed to interfere with aS aggregation, can induce a significant bias in ThT intensitymeasurements (Hudson et al. 2009), reducing the reliability of this method for the evaluation ofaggregation enhancers or inhibitors. Finally, ThT is likely unable to properly bind β-sheets smallerthan a fibrillar structure, preventing the detection of aggregation intermediates, which are, accord-ing to the latest hypothesis, the most toxic species in vivo (Winner et al. 2011) (see Section 5.3).The second most common extrinsic dye used to study amyloid fibrils formation is Congo Red.

This dye, when observed through a polarized filter, shows birefringent emission upon binding toamyloid fibrils and has been used in particular to identify amyloids in post-mortem brain tissues.Moreover, the absorbance spectrum of fibril-bound Congo Red shows a red-shift comparedwith the free form of the dye (Conway et al. 2000). Potential caveats are that Congo Red hasbeen shown to induce oligomerization of native proteins and to bind to partially folded confor-mations of several proteins, not exclusively to amyloid fibrils (Khurana et al. 2001). For these rea-sons, Congo Red uses in the study of the kinetic properties of the aS aggregation process or in thedetection of amyloid fibrils in vitro has to be approached with care. Up-to-date reviews are avail-able for most of the suitable dyes used to detect amyloid fibrils, and these describe in depth thebinding modes and the possible applications of these assays for studying protein aggregation(Groenning, 2010).However, the potential pitfalls in the use of extrinsic dyes to study the subsequent steps in the

aggregation process and the limited reproducibility of data concerning aS fibril formation (Giehmet al. 2011a) propelled the development of alternative methods to evaluate aS oligomerization andfibril formation.

2.2 Monitoring aggregation through the intrinsic properties of aS

2.2.1 Absorbance variation in aggregating aS solution

aS aggregation can be monitored by collecting aliquots from an aS aggregation assay solutionat different time points and, after pelleting the fibrils present in the sample, measuring theamount of soluble aS still present. Even if this method does not provide information aboutthe aS aggregation process, i.e., from the early interactions to oligomerization to fibril formationand elongation, it is still useful because it does not require the use of extrinsic dyes or mutationsin the aS sequence to monitor the process. Absorbance variation measurements were originallyused by Wood et al. (1999) to analyse samples in an aggregation assay in which the fibrillizationwas accelerated by adding fragmented (sonicated) mature fibrils, termed seeds. After demonstrat-ing that aS seeds can function as nuclei, promoting amyloid fibril assembly, the authors


defined aS fibril formation as a nucleation-dependent process, in which seeds or early oligomericintermediates accelerated mature fibril formation. This idea was recently exploited to reproducecellular models for aS aggregation (see Section 6).

2.2.2 Dynamic light scattering

The kinetic properties of aS aggregation were also studied using dynamic light scattering(DLS), which allows the determination of hydrodynamic radii of macromolecules in solution.In a time-course analysis of aggregation, fibrils were pelleted and supernatants collected andassessed at different time points. DLS showed an increase in the amount of species with largerradii (Dusa et al. 2006) and allowed partial characterization of the intermediate aS oligomers.However, this technique has two main limitations: (i) DLS signal intensity depends on the massesof the objects in solution, so the methods lose their ability to detect small oligomers whenlarger objects are present in solution, making DLS unsuitable for studying the whole aS aggre-gation process; and (ii) the radii calculated for each weighted fraction are the results of algorithmsthat are not standardized and are derived from strong approximations about aggregate shapes.

2.2.3 Intrinsic tryptophan fluorescence

Another method for studying aS oligomerization exploits the intensity variation in the intrinsictryptophan fluorescence of aS mutants. Given that the aS sequence lacks tryptophans, the fluor-escent amino acid was introduced at different positions of the wild type (wt) aS sequence bysingle-site mutagenesis. This mutagenesis has both an advantage because the position in whichthe tryptophan is introduced can be freely chosen according to the experimental strategy and apotential disadvantage because it introduces a large and bulky amino acid that is not presentin the native sequence of aS and that likely influences aS aggregation properties.The tryptophan emission maximum is typically at ∼355 nm for unfolded proteins in which the

benzimidazole moiety is fully exposed to the solvent. Upon confinement in a more hydrophobicenvironment, the spectrum displays a blue-shift; this variation can be used to monitor foldingor aggregation processes that perturb the local environment of the benzimidazole. Kaylor et al.generated the Y125W/Y133F/Y136F mutant, which shows the kinetic properties of fibrillizationsimilar to those of the wt aS (as evaluated by ThT emission measurements), as well as the similarassociated morphology (as evaluated by transmission electron microscopy, TEM) (Kaylor et al.2005). The Trp emission maximum wavelength was evaluated as a function of time for the pel-lets, in which the insoluble aggregates were collected. The wavelength value of maximum emis-sion remained 352 nm for the insoluble fibrils, indicating that the Trp125 was still exposed to thesolvent. Conversely, a blue-shift, from 356 to 352 nm, was recorded in the supernatants duringthe latest stages of the aggregation process, suggesting that the larger oligomers present in thesolution at longer incubation time have less solvent-exposed tryptophan.Dusa et al. (2006) prepared the Y39W mutant, and its tryptophan fluorescence was again

measured as a function of aggregation time. This Y39W mutant showed a 2-fold decrease inthe fibrillization rate in comparison with the wt aS. Therefore, the insertion of a Trp in the aSsequence has some potential as a structural probe, but it also has a clear drawback in the pertur-bation that this modification induces, which calls for a careful evaluation of each designedmutant.


2.3 Fluorophores covalently bound to aS monomers

To study the dynamics of aS aggregation by monitoring variations in fluorescent properties,the protein has to be modified. In these paragraphs, we will survey studies in which aS hasbeen covalently bound to specific fluorophores. The binding is usually done through thiol groupsby insertion of a cysteine in a specific position in the aS sequence, exploiting the fact that aS doesnot have any cysteines in its native sequence. In most cases, the cysteine is added at the C-term tolimit the effects that both the amino acid and the fluorophore could have on aS aggregationproperties. Modification in other regions, i.e., the NAC, could be more informative about aS rear-rangements but would also more likely to be invasive because this region is the site of the primaryinteractions that drive aS fibrillization.

2.3.1 Fluorescence correlation spectroscopy

Fluorescence correlation spectroscopy provides information about the distribution of the dif-fusion coefficients of fluorescent molecules in a sample. For this reason, it was used to monitorthe diffusion coefficients of aS conjugated to a fluorophore at different time points in an aggre-gation assay. The A140C aS mutant labelled with Alexa Fluor 488 was used by Nath et al. ( 2010)to investigate the early stages of aS oligomerization. The authors could demonstrate the presenceof progressively larger oligomeric species with time: after 7 h of aggregation, they observed aheterogeneous distribution of oligomers with different diffusion coefficients and confirmedthe concentration dependence of the early stages of the aggregation process. Although theauthors could not characterize any single oligomeric species due to the intrinsic heterogeneityof the sample, it is interesting to note that the FCS technique is sensitive to the early intermediatesand that its use for the study of small oligomers is not hampered by the presence of largeraggregates.

2.3.2 Fluorescence polarization

Fluorescence polarization (or anisotropy) measurements are a useful tool to evaluate aS aggre-gation and can reveal not only larger aggregates or amyloid fibrils but also oligomers.Preliminary anisotropy measurements on aS fibrils were performed by Dusa et al. (2006), but

fluorescence polarization data concerning the kinetics of the whole aS aggregation process werefirst reported by Luk et al. (2007). Fluorescence polarization allows the monitoring of changes inthe relative molecular masses of the objects in solution, revealing not only soluble monomers ormultimers but also larger aggregates and insoluble fibrils. The main advantages are that fluor-escence polarization allows detection of oligomeric species and therefore is the most suitabletool to evaluate the effects of aggregation inhibitors acting at the early stages of aS oligomeriza-tion. To perform an fluorescence polarization experiment, a fraction of aS monomers in theaggregation assembly must be conjugated with a fluorophore. For example, Luk et al. (2007)added a cysteine to the wt aS sequence at position 141 and labelled the protein by reactingsuccinimidyl-ester forms of the Oregon Green 488 and Alexa Fluor 594 with purified aS cysteinemutant.

2.4 Novel approaches

The approaches mentioned above are well established and provided important information aboutthe aS aggregation process. However, most of these studies manifest some limitations: ThTexperiments have limited reproducibility, evaluation of aS fluorescence variations requires


modifications of the aS sequence, and most of the studies, for example, the ThT, DLS and absor-bance variation experiments, do not allow the characterization of the whole aS aggregationprocess.Therefore, investigators embarked on the exploration of alternative techniques to achieve

a more thorough characterization of the kinetic properties of aS fibrillization. The favouredapproaches are those capable of tackling the early stages of the process, which are the lessknown stages, and of providing several types of information during the same measurement.In 2008, a new fluorescence aggregation assay, which provides information on oligomeric

species and on amyloid fibrils, was developed (Thirunavukkuarasu et al. 2008). Pyrene-labelledA19C, A90C and A140C aS mutants were monitored in an aggregation assay, and several fluor-escence parameters were measured. This assay provides a wide range of information, includingthe detection of the early stages during aS aggregation process, the decrease of the monomer frac-tion and the increase of the molecular masses in solution can be easily detected. These parametersare also commonly measured using other probes, but a progressive decrease in the sample’s po-larity, an increase in the steady-state anisotropy and differences in intramolecular binding can becharacterized at the same time, providing a wider set of information concerning the aS aggre-gation process.More recently, Yushchenko et al. (2010) introduced a new method that is particularly sensitive

to the early intermediates of the aggregation process. They measured excited-state intramolecularproton transfer (ESIPT) in the A140C aS mutant conjugated to a thiol-reactive (maleimide) probe(MFC), based on 2-(2-furyl)-3-hydroxychromone FC (aS-MFC). aS-MFC has two emissionbands, the intensities of which were denoted T* and N*; the authors showed that the intensityratio T*/N* increased upon aggregation and the T* band exhibited a red-shift, meaning that theprobe was confined to a less protic and less polar environment. The aggregation reaction can befollowed continuously from the early steps of the process using this technique. Moreover, ESIPTmeasurement of aS-MFC can provide a multiparameter set of data, which is more informative incomparison to other conventional assays.Variations in the fluorescent properties of a dansyl probe were also used to study the aggre-

gation of aS (Yap et al. 2011). The authors prepared four cysteine mutants (G7C, V26C,L100C and Y136C) on which to place the dansyl moiety in the aS sequence. The dansyl reagentis particularly suitable to study proteins in solution because it is sensitive to the local environmentand can act as fluorescence energy transfer acceptor, i.e., it is able to provide a great deal ofinformation by measuring the same fluorescence signal in a single experiment. 1.5% of dansyl-derivatized aS was used to characterize wt aS aggregation kinetic, by monitoring variations indansyl emission intensity and light scattering. These data suggested that the first steps of oligo-merization mainly involve the ends of the protein – the C- and N-term disordered regions.A very reliable method to study aS aggregation kinetic was recently proposed by Cremades et al.

(Cremades et al. 2012). They used single-molecule two-colour coincidence detection (TCCD) andForster resonance energy transfer (FRET) experiments to estimate the change in oligomerfraction during the aS aggregation process. aS was conjugated to two different fluorophores(Alexa Fluor 488 and 633) and a time series TCCD analysis was performed on different aliquotstaken from the aggregation mix. The presence of oligomers was detected in correspondence offluorescence emission bursts for both colours. Characterization of different oligomeric speciesbelonging to different aggregation pathways was performed over time, leading to the identifica-tion of four lag phases and oligomerization rates for these species, before conversion into fibrillarspecies. This method, first demonstrated in 2006 by the same group (Orte et al. 2008), is sensitive


and able to detect oligomeric aggregation intermediates during the aggregation process, althoughit requires an experimental apparatus specifically designed for the collection of this type of data.It was recently shown that aS oligomers and aggregates develop an intrinsic fluorescence emis-

sion that can be used in a quantitative assay to monitor amyloid fibril growth (Pinotsi et al. 2013).The major advantage of this method is that it is label-free: monitoring the variations in theintrinsic emission allows the study of the kinetics of aggregation without the introduction ofextrinsic dyes that can interfere with the process. The authors of the paper report that the effectsof lacmoid on aS fibrillization were properly revealed by measuring the intrinsic fluorescenceemission, whereas the measurement of ThT fluorescence led to the incorrect conclusion thatlacmoid can hinder the aggregation process because lacmoid interacts with ThT and alters itsfluorescence properties. Interestingly, this method could be extended to the characterization ofother amyloidogenic proteins.These different and novel methods could seem less approachable compared with the

traditional ones, but they provide more refined information about the process studied.However, these methods do suffer from the drawback that few of the required experimental set-ups exist, hindering robust replication by multiple-independent laboratories.

2.5 Aggregation inhibitors and enhancers

Early on, aS fibrillization has been suggested as a possible drug target to slow or stop neuro-degeneration, and it has been shown to be strongly affected by various factors and moleculesthat can either enhance or inhibit the process.The fibril elongation rate was shown to be directly proportional to protein concentration

(Wood et al. 1999), which may explain the pathogenicity associated with SNCA gene multipli-cation (Singleton et al. 2003) that leads to an increased dosage of aS.The kinetic parameters of fibrillization for the pathological mutants were compared with those

of the wt: A53T showed an increased rate of aggregation and a shorter lag phase, whereas A30Pdisplayed a longer lag phase and a slower growth rate, leading to a delay in the midpoint of thefibrillization transition (Li et al. 2001). In addition, the third mutant E46K, discovered afterA30P and A53T, showed an increased fibrillization rate, similar to A53T (Fredenburg et al.2007), with a lag phase twice as long (∼8 h) compared with the wt one. Published data concerningthe most recently discovered aS pathological mutants are not yet available, but the emerging pic-ture implies that the aS mutants, more than the aggregation process per se, are the culprits inneurodegeneration.Covalent chemical modifications of aS, such as nitration, oxidation (associated with an inflam-

matory response) and phosphorylation, lead to variations in both the rates and the final productsof the aggregation process. Methionine oxidation reduces aS aggregation propensity in vitro: thepresence of a 4-fold molar excess of oxidized protein (Uversky et al. 2002) completely inhibitsfibril formation. It has also been shown that the presence of a sub-stoichiometric concentrationof nitrated aS in the aggregation assembly interferes with the fibrillization process (Yamin et al.2003). However, aS with tyrosine nitration has been detected in LB, suggesting that interferingwith aggregation leads to the sequestration of the nitrated aS into the aggregates (Duda et al.2000; Giasson et al. 2000).In addition, the fraction of aS phosphorylated in position 87 (pSer87) and 129 (pSer129) is

enhanced in LBs and in the brains of patients affected by synucleinopathies (Fujiwara et al.2002; Paleologou et al. 2010). Of relevance to aggregation, it has been shown that pSer87,


generated in vitro by casein kinase 1 (CK1) phosphorylation, inhibits aS fibrillization (Paleologouet al. 2010); similarly, Paleologou et al. demonstrated that CK1-mediated phosphorylation of aShampers aS aggregation (Paleologou et al. 2008), whreas Fujiwara et al. (2002) reported thatpSer129 aS, phosphorylated using casein kinase 2 (CK2), shows an increased fibrillization pro-pensity. It is noteworthy that Ser129 is not in the core of aS fibrils and can also be phosphorylatedafter fibrillization in a physiological environment (Paleologou et al. 2008). Interestingly, aS is phos-phorylated at Ser129 not only by casein kinases but also by Polo-like kinase 1, 2 and 3, both invitro and in vivo, with different phosphorylation efficiencies (Mbefo et al. 2010). However, the re-lationship between phosphorylation and aggregation is still an open question and needsclarification.Divalent and trivalent metal ions (for example, Al3+, Cu2+, Fe3+ or Pb2+) have been shown to

strongly accelerate the aS aggregation rate in vitro, although their relevance to the process in vivoremains to be proven (Bisaglia et al. 2009). Similarly, polyanions (Cohlberg et al. 2002; Liu et al. 2005)or polycationic species (Goers et al. 2003) accelerate the aS aggregation rate in vitro. The effect ofthese compounds depends on their concentration and, in the case of polymers, also on theirlength, and suggests a critical role for electrostatic interactions in modifications of the aggregationrate. It has been proposed that the masking of negative charges at the C-term could lead to thecollapse of aS to a more amyloidogenic conformation.Other molecules, such as catechins or chaperones (Bieschke et al. 2010; Dedmon et al.

2005a), or properly designed peptides (El-Agnaf et al. 2004; Sievers et al. 2011), can slow or in-hibit aS aggregation by direct interaction with the monomer or with other aggregationintermediates.

2.6 Kinetic models

From the plethora of data available in the literature on the time dependence of aS (but also ofother amyloid proteins) fibrillization in vitro, investigators attempted to develop kinetic modelsto describe and predict aS aggregation behaviour.The proposed models are based on different types of assumptions and mathematical analysis

(for an exhaustive review and the details of the mathematical treatments, see Morris et al. 2009),but none of them, to our knowledge, emerge as more significant than the others. This result ismost likely due to variability in the data, which depend on the chosen experimental conditions, aswell as to the complexity of the described process and the consequent complexity of the pro-posed models. Furthermore, the real test that could validate a kinetic model is its applicationto the data on aS aggregation in a cellular environment, a possibility that is hindered by the sparseexperimental data on the process in live cells (see Section 6).Each kinetic model provides a fitting of the data based on different assumptions about the

mechanism that leads to oligomer and fibril formation in vitro, but the limiting step remainsthe intrinsically low reproducibility of aS aggregation kinetic data. In the sigmoid curve, theslope at t50 and the length of the lag phase change significantly (lag phase length varies bytwo orders of magnitude) by varying the incubation conditions (Pronchik et al. 2010). In particu-lar, agitation (Pronchik et al. 2010) and shear forces due to mechanical influences, such as cen-trifugation (Bhak et al. 2009), have been identified as the most relevant parameters in the intrinsicvariability that affect the aggregation process. Pronchik et al. performed aggregation experimentsto determine a direct correlation between shaking rate and fibrillization rate. In the literature,three hypotheses have been proposed: (i) a faster fibrillization rate upon faster shaking is due


to an increased rate of fibril fragmentation and the consequent generation of more active nu-cleation sites; (ii) vigorous mixing eliminates local concentration gradients and promotes fibrilelongation, but is limited by the mass transfer rate; and (iii) in the presence of hydrophobic–hydrophilic interfaces, aS behaves as a surfactant, i.e., it concentrates, attempting to minimizethe exposure of its hydrophobic residues to water solution by clustering, and acquires anaggregation-prone conformation. By aggregating aS under gentle agitation and mixing withbeads at different concentrations and with different surface hydrophobicity, the authors observedthat only the hydrophobic–hydrophilic interface hypothesis is consistent with their results:they showed how the concentration (the actual active surface area per aS molecule) and thetype (polytetrafluorethylene, polymethylmethacrylate and glass) of interface directly correlatewith aggregation rates. Based on their results, they propose that the irreproducibility oftenobserved in aS kinetic data is due to the variability in heterogeneous air–water interfaces duringagitation. Moreover, they suggested that aS partitioning at these interfaces causes an increasein local protein concentration and drives the equilibrium towards conformations that couldmore easily prompt amyloidogenesis.Relatively high concentrations (more than 1 mg/ml∼70 μM) and/or the presence of aS seeds

can further enhance reproducibility (Giehm & Otzen, 2010; Giehm et al. 2011a), but these para-meters can act only from the second or third stage of the process (Fig. 2), when oligomers(or seeds) form and fibrils are growing in length. However, the main source of variability remainsthe first step of the aggregation when aS undergoes conformational rearrangements whose prob-ability seems to be extremely low. For this reason, we suggest that the reproducibility issue cannotbe completely overcome at this point, partially because of technical limits but also because of theintrinsic properties of aS.Two recent papers explore the possibility of accelerating the aS fibrillization process by prepar-

ing chimaeric aS dimers (Pivato et al. 2012) or by inducing dimerization in a mutated aS throughan inducible dimerizing domain (Roostaee et al. 2013). In this second case, the induction ofdimerization between the C-term of two aS molecules led to an increase in the aggregationrate compared with the wt aS.In Pivato et al. the comparison between different aS dimers is reported: aS cys mutants were

designed to form aS dimers bound through disulphide bonds between the two N- or C-termini(respectively, NN or CC dimers). Moreover, a tandem dimer, consisting of two aS molecules(NC dimer), and a double-core dimer, consisting of two NAC regions (DC dimer), weredesigned. Interestingly, CC and DC dimers showed faster aggregation kinetics when studied byThT compared wyth the wt, whereas NN and NC dimers exhibited a negligible increase inThT fluorescence intensity over time, indicating slower aggregation. These latest results suggestthat it is not only the increase in the local concentration of aS molecules due to dimerization butalso the chosen constraints in terms of the relative orientation of aS molecules in these dimersthat strongly affect the kinetics of the aggregation process.A final comment on the kinetics of aS aggregation is on the stirring issue: most of the literature

regarding aS aggregation in vitro describes experiments performed while shaking. This consider-ation led to a question on the relevance of stirring for the aggregation process in cells and in PDmodels more generally. In our opinion, stirring should be seen as an efficient accelerator of theaggregation process and a price to pay to reduce the aggregation time frame to values experimen-tally approachable. It remains clear that from a physiological point of view, stirring has no rel-evance, but stirring in vitro allows experiments to fast forward through the years that aS seemsto require to aggregate in parkinsonian brains.


3. Monomeric aS conformations

aS, because of its lack of a defined secondary structure when purified from recombinant over-expression systems, has been considered an intrinsically disordered protein (Weinreb et al.1996). In a quite recent and disputed paper (Bartels et al. 2011), aS was described as anα-helical homo-tetramer when purified from mammalian cells overexpressing aS or fromhuman red blood cells. However, these results could not be replicated by other groups(Fauvet et al. 2012b), who found aS as an unfolded monomer in erythrocytes, mammalian celllines and Escherichia coli. Interestingly, aS with a modification of ten amino acids added at theN-term showed an α-helical tetrameric conformation when purified from E. coli and studiedby nuclear magnetic resonance (NMR) (Wang et al. 2011). Nonetheless, Gurry et al. showedthat the very same modified aS in solution was present as a monomer and, in a small fraction,

AGKTKEGVLYVGSKTKEGVVHGVATVAEKTKEQVTNVGGAVVTGVTAVAQKTVEGAGSIAAATGFVKKDQLGKNEEGAPQE

40 50 60 70 80 90 100

(1)

(2)

(3)

(4)

40 50 60 70 80 90 100

1

1,2

0,8

Fig. 2. Core region of aS fibril. Amino acidic sequence and β-strands representation for different models:(1) β-strands distribution in aS fibril as obtained from EPR experiments (Chen et al. 2007). (2) β-strandsdistribution in two different type of aS identified in a solid-state NMR study (Heise et al. 2005). (3) Fiveβ-strands were proposed by Vilar et al. in a model based on solid-state NMR experiments (Vilar et al.2008). (4) Most recent β-strands distribution in the aS fibrils core region as defined by solid-state NMRassignment of 106 out of 140 residues of the aS sequence (Comellas et al. 2011). Below, a schematicrepresentation of the result obtained using Chou–Fasman algorithm to calculate the average β-sheetpropensity of the central part of aS, starting from residue 30 up to residue 110 (Suk et al. 2010). Asexpected, there is a good correspondence between the calculated β-sheet propensity and the β-strandsmeasured using structural techniques described above.


as trimers and tetramers presenting either a β-strand or an α-helical secondary structure(Gurry et al. 2013). Conversely, Binolfi et al. reported that aS directly studied within E. coliwith a method that does not require cell lysis or protein purification showed an NMR signaltypical of monomeric and disordered proteins (Binolfi et al. 2012). An alternative scenario pro-posed by Trexler & Rhoades (2012), is that N-terminal acetylation of aS in combination witha mild purification from bacteria results in an α-helical oligomeric aS form. However, othergroups studied structural effects of acetylation on aS by NMR and reported that aS acetylation,which was shown to be an abundant modification in human brains (Anderson et al. 2006), wasonly associated to a transient variation in N-term aS α-helical propensity and did not lead to theformation of stable tetramers (Fauvet et al. 2012a; Kang et al. 2012). These contradictory resultskeep the issue open: the majority of data points to monomeric aS as the physiologically relevantspecies, whereas only a few reports suggest the presence of functional aS tetramers. It goes with-out saying that more work is required to resolve the controversy.The first stage of aS fibrillization is believed to be the formation of a transient form of mono-

meric aS that is prone to aggregation. This step is a very rare event crucial for the initiation ofoligomerization, and it has been studied using three strategies: (i) ensemble solution techniques,such as NMR, fluorescence measurements and molecular dynamics (MD) simulations; (ii) single-molecule force spectroscopy experiments, which are useful for studying protein folding andunfolding at the single-molecule level and have been used to characterize the conformationalchanges aS undergoes before aggregation begins; and (iii) fluorescence microscopy, whichwas used to study perturbations in aS conformations induced by cellular membrane interfaces,showing that in the presence of lipids, the propensity of aS to aggregate is modified.

3.1 Monomeric aS structural analysis

3.1.1 Paramagnetic relaxation enhancement-NMR measurements andmolecular dynamics (MD) simulation

The intrinsic heterogeneity of the aS ensemble is challenging for the investigators using NMRspectroscopy and MD simulation with regard to obtaining a high-resolution characterization ofaS disordered states. The information thus obtained can only be associated with the averageproperties of the ensemble and not with single aS molecule conformations. Using a spin-labelcovalently bound to the protein through a cysteine strategically placed in its amino acid sequence,investigators acquired 1H–15N correlation (HSQC) spectra of the protein. The combination ofHSQC and paramagnetic resonance enhancement allowed the identification of long-range inter-actions between the central part of aS and the C-term, suggesting that the native aS ensembleconsists of heterogeneous structures. Matching constraints obtained by the data analysis andMD simulations, the authors found that the distribution of aS monomers in solution is composedof conformers with various gyration radii and is considerably different from the distributionexpected for a fully random coil protein (Allison et al. 2009; Bertoncini et al. 2005; Dedmonet al. 2005b). Using the same techniques, other research groups investigated monomeric aS tran-sient interactions, folding and rigidity, by considering the effect of acidic pH, which is an exper-imental condition that promotes aS aggregation by decreasing its negative charge. Theseexperiments allowed the identification of aS conformations that could be prone to aggregationin vitro but went undetected at physiological pH values (Cho et al. 2009; McClendon et al.2009; Wu & Baum, 2010). Another method that allows us to explore the thermodynamicsof the system by perturbing the aS conformation distributions, is to work at supercooled


temperature. Using this setup, Wu et al. (2009) measured aS ensembles that are less averaged byNMR relaxation and provided information on aS conformations. It should be mentioned that inthe context of an extremely slow process, such as aS aggregation, in which a very small fraction ofthe molecules present is likely to be able to drive the process, ensemble solution techniques mayfail to identify the relevant species. Pushing the experimental conditions far from what can bephysiologically expect may lead to the identification of otherwise undetectable forms thatcould be relevant but must be considered with caution.

3.1.2 Single-molecule force spectroscopy experiments

NMR experiments and MD simulation can only provide ensemble-averaged information on aSmonomer structures. Conversely, single-molecule force spectroscopy methods based on atomicforce microscopy (AFM)-driven mechanical unfolding permits the characterization of aS confor-mational variants at the individual molecule level. Sandal et al. first proved the polymorphic natureof aS in solution by measuring its behaviour in response to a mechanical unfolding force (Sandalet al. 2008). Later on, the same group (Brucale et al. 2009), as well as others more recently (Hervaset al. 2012), further characterized the aS conformation ensemble using force spectroscopy andthe different conformations that could be ascribed to aS pathological mutants. The identificationof stable conformers that precede aS oligomerization may be important because they could betargets for a possible therapeutic intervention.

3.1.3 Fluorescence measurements on aggregation prone monomeric aS conformers

A fluorescence technique that was used to study conformational changes in aS monomers issingle-molecule Forster resonance energy transfer (smFRET) (Trexler & Rhoades, 2010). Thismethod allows determination of the distances between two fluorophore-labelled amino acidsby measuring FRET between donor and acceptor. Trexler et al. using different labelling sites,were able to characterize the heterogeneous and dynamic state of aS in solution at differentpH and in the presence of charged molecules, i.e., in conditions that promote aS aggregation.The authors showed that at low pH, a collapse of the C-term onto the NAC region occurs,while the N-term and the central part of the protein are less affected. The construct labelledat the two ends of the protein, i.e., in positions 9 and 130, presented the same type of structuralcollapse observed at pH 3.0 and at pH 7.4, showing the C- and N-terms in close proximity. Thesedata suggest that aS in solution undergoes substantial conformational variations. However, it isstill unclear if these conformational rearrangements are responsible for the first interactions lead-ing to protein oligomerization. Another possibility is that the proximity of the C- and N-termscan protect the NAC region from those interactions that lead to aS aggregation.Yap et al. (2011), measuring dansyl-conjugated aS cysteine mutants, further confirmed that aS

aggregation requires both C- and N-term rearrangements. In particular, conformational modifi-cations that involve both residues 7 and 136, in the unstructured C- and N-term regions, respect-ively, occur early in the aggregation process before aS fibril formation. Conversely, changes atresidues 26 and 100, closer to the fibrillar core, occur later in association with fibril formation.Taken together, these NMR, MD, smFRET and single molecule force spectroscopy data on

aS molecules suggest that the unfolded state of aS in solution in vitro consists of an ensembleof different, transient and dynamic conformations, some of which can be, in specific conditions(low pH, higher temperature, different ionic strength), prone to establish transient or stable bondswithin the same aS monomer and/or with other aS molecules in vitro, driving the first steps


of protein oligomerization. The tested conditions could mimic the environmental changes thatoccur in neurons and that lead to protein conformational modification and aggregation.

3.2 aS conformation and aggregation on membranes

Recently, a new aspect has been proposed to be relevant in the aggregation process: the accumu-lation of aS on membrane surfaces. When monomeric aS interacts with vesicles or membranes,the N-term of the protein acquires an α-helical conformation that can be a bent helix, anextended helix or an antiparallel helix-turn-helix structure, depending on the membrane proper-ties. Conversely, the C-term, as amply demonstrated, remains unstructured (Borbat et al. 2006;Bussell et al. 2005; Davidson et al. 1998; Ferreon et al. 2009; Georgieva et al. 2010; Jao et al.2008; Middleton & Rhoades, 2010; Trexler & Rhoades, 2009; Ulmer et al. 2005). For amyloido-genic proteins, but not for aS, a direct conversion was observed between the α-helical confor-mation that most of the proteins acquire upon membranes binding (Butterfield & Lashuel,2010) and the β-sheet structures typical of amyloid oligomers and fibrils. Furthermore, an in-crease in fibril formation was observed when aS was aggregated in the presence of brain-derivedmembranes (Lee et al. 2002) and anionic detergents or fatty acids micelles (Necula et al. 2003),suggesting that membranes also play an important role in aS aggregation. Interestingly, it hasbeen proposed that the binding of aS to the membrane increases the local concentration, favour-ing the triggering of the aggregation process (Auluck et al. 2010).Reynolds et al. (2011) showed by supercritical angle fluorescence microscopy (SAF) that aS in

a sub-physiological concentration in solution tends to aggregate in the presence of negativelycharged supported lipid bilayers. FRET images of donor-labelled (Dy647-N-hydroxysuccinimide)supported lipid bilayers and acceptor-labelled (Cy7-monofunctional NHS ester dye molecules) aSwere performed by SAF microscopy, with the total protein–lipid ratio going from 1:21.6(200 nM) to 1:1.44 (3 μM). Most of the protein remains in solution, and initially, the interactionbetween aS and the lipids led to the switch of the protein to the α-helical conformation previouslymentioned. Later on, a structural rearrangement occurs, leading to the formation of aggregates onthe lipid bilayer of a smaller part of the aS molecules, while the majority of the protein is still insolution. The aggregates were also observed by cryo scanning electron microscopy, showing a sizevarying from a few micrometres to tens of micrometres and displaying a morphology typical ofamyloid fibrils. It seems that the ability of aS to fulfil its physiological function(s) at the mem-branes depends on the protein–lipid ratio and on the type of lipids. It could be that under patho-logical conditions, aS binding to the membrane is no longer protective and leads to aS aggregationon the membranes, which can damage membranes through different possible mechanisms. TheaS aggregation described here is time and concentration dependent: at nM concentration,aS aggregates led to membrane damage, destabilization, thinning and, more unexpectedly, lipidextraction from the membrane, while at μM concentration, disruption of the bilayer occurred.A solid-state NMR (ss-NMR) study on aS aggregation in the presence or absence of anionicphospholipids showed that fibrils are formed in both cases, but when lipids are present, thefibril structure exhibits major structural perturbations at the N-term (Comellas et al. 2012).Therefore, the authors proposed a model for aS aggregation in the two different cases, accountingfor the revealed structural differences.The increase in the local concentration of aS at the membranes can induce not only increased

aggregation but can also trigger membrane permeabilization induced by monomeric aS thatassembles into oligomers at the membranes upon the application of a voltage (Tosatto et al. 2012).


These oligomers are ordered and show a recursive structure similar to the pores formed by somebacterial toxins. The relationship between these objects and the previously mentioned aggregatesformed at the membranes and observed by other groups remains unclear.

4. aS oligomeric species

As previously mentioned, aS oligomeric species are heterogeneous, transient and poorly defined.In the last 10 years, their relevance in the molecular studies on synucleinopathies has grown con-sistently. Since 2001, several studies have proposed oligomer toxicity, especially for their annularstructures, similar to the pore-forming toxins, and their purported ability to interact with lipidmembranes. In 2004, Pountney et al. (2004) were able to purify aS annular oligomers from inclu-sions in patients affected by MSA. More recently, the E57K aS mutant, the only one able to formoligomeric species that do not convert into amyloid fibrils, was used to study oligomer toxicity inan in vivo mice model (Winner et al. 2011).In an early work, Conway et al. (2000) observed that both A30P and A53T mutants shared

a faster oligomerization rate in vitro but not a faster fibrillization, suggesting that a faster fibrilliza-tion rate is not directly linked to a higher cytotoxicity in PD. Even the E46K mutant, which alsoshowed an increased aggregation rate with respect to wt, can form soluble, non-fibrillar specieswith sizes and chemical properties similar to wt soluble annular protofibrils (Fredenburg et al.2007).Based on these observations and on the fact that aS oligomers are still poorly understood

despite their importance, a major goal of current research on aS aggregation from the perspec-tives of both basic science and new therapeutic strategies is their biophysical characterization. Thelatter is germane to defining the toxicity mechanism(s) of oligomers. A detailed molecular under-standing of aS pathological forms can open a new avenue to the design of specific drugs tailoredagainst these toxic conformations.Two roles were mainly ascribed to aS oligomers, although other toxic mechanisms have been

recently suggested (Section 6.2): (i) membrane permeabilization through a mechanism typical ofpore-forming toxins; and (ii) PD spreading through a mechanism similar to prion protein(Polymenidou & Cleveland, 2012), where misfolded aS passes from one cell to another, triggeringaggregation and subsequent toxic events. To our understanding, there are some main issues to benoted: first, objects spreading from cell to cell should be stable; second, the spreading, as will bediscussed in detail later, may occur through different pathways; third, it remains to be determinedif the transferred objects themselves become toxic for the recipient cells or if they work as a tem-plate to seed a new aggregation process whose products are responsible for the induced toxicity.The occurrence of aS-mediated membrane permeabilization is supported by in vitro and cellular

experiments: some oligomeric species obtained from recombinant aS showed a pore-like struc-ture when observed using microscopy techniques (Fredenburg et al. 2007; Lashuel et al. 2002;Quist et al. 2005; Winner et al. 2011), and they exhibited a clear electrophysiological activity onvesicles (Fredenburg et al. 2007; Volles & Lansbury, 2002) and membrane models (Kim et al.2009; Schmidt et al. 2012; van Rooijen et al. 2010), similar to the one expected for pore-formingtoxins (Lashuel & Lansbury, 2006).The cell-to-cell spreading process of aS was highlighted by clinical data that showed the pres-

ence of LBs in grafted neurons in individuals affected by PD who died 10 years after surgery(Kordower et al. 2008; Li et al. 2008). This observation strongly supports a cell-to-cell propagationmechanism to explain the spread of aS pathology in these brains. Based on numerous


post-mortem analyses, in 2003 Braak formulated a staging classification of PD wherein LBs firstappear in the olfactory bulb and only at later stages are found in the midbrain and neocortex(Braak et al. 2003), suggesting a progressive involvement of different brain regions, possiblydue to the passage of aggregated aS. In support of this evidence, PD cellular models for aS ag-gregation have largely been used to study the mechanism of aS spread: in particular, it has beenshown that either oligomeric species or seeds obtained from recombinant aS can be up-takenby healthy recipient cells in PD cellular models, in primary neurons cultures and in mice models.These aS species are able to trigger the aggregation process (Danzer et al. 2009; Desplatset al. 2009; Hansen et al. 2011; Luk et al. 2009, 2012; Nonaka et al. 2010; Volpicelli-Daley et al.2011).In view of these facts, the study of oligomeric aS aggregation intermediates in relationship to

their putative pathogenic effects in synucleinopathies becomes particularly relevant; however, thisstudy presents some considerable challenges. As previously mentioned, aS oligomeric species areheterogeneous and difficult to define, which becomes an issue when trying to study aS oligomercharacteristics in relation to their noxious effects. We propose a minimal but sound definition foraS oligomers as ‘objects consisting of several aS monomers that exhibit newly acquired functionalproperties in the cell’, which, unfortunately, remains inadequately vague.The first problem encountered when approaching the study of aS oligomers is the fact that

their structural, morphological and functional properties are extremely sensitive to the experimen-tal conditions under which they are formed (i.e., temperature, pH, buffer), to the presence ofreactive molecules, to the presence of other proteins during the aggregation process, and to physi-cal or chemical treatments (as reported in Section 3.1, 3.2 and 3.3). The second problem is thatsome of the oligomers, mainly obtained in vitro using recombinant aS, lack a robust validationin vivo. From the pathological point of view, it is clear that different oligomeric species acquireimportance only when there is evidence of both their presence and functional impact in PDcell or animal models or in post-mortem brain tissue and, consequentially, when they can be linkedto the pathology.Despite their evident relevance, aS oligomeric species often form transient species that are

difficult to prepare, isolate and analyse (Giehm et al. 2011b; Kim et al. 2008; Lashuel et al.2002). Moreover, in some instances, they may be in equilibrium with aS monomers, whichagain depends on experimental conditions, and may decrease their molar fraction when, forexample, an isolation method such as gel filtration (GF) is used (Bhak et al. 2009). This latterconsideration could explain the differences in the estimated amount of oligomeric speciesobserved when aS aggregated in solution was analysed by GF (Lashuel et al. 2002) or by smallangle X-ray scattering (SAXS) (Giehm et al. 2011b). Lashuel et al. showed that a type of solubleaS oligomeric species, termed protofibrils, could be obtained by simply incubating aS wt, A30Pand A53T on ice. The protofibril fraction that can be separated by GF was always <15% ofthe total protein in solution. The estimated apparent molecular mass of the oligomeric speciesis 600 kDa on average, corresponding to more than 42 aS molecules. In the same sample,Lashuel et al. could observe annular and/or tubular oligomers by EM that, in their speculation,consisted of 20–24 aS monomers. A similar result was obtained by an independent techniquemore recently. Single-molecule photobleaching was used by Zijlstra et al. (2012) to determinethe average number of monomers in aS oligomeric species. The A140C aS mutant was labelledwith Alexa Fluor 488, and wt aS was incubated together with different percentages of labelledprotein at a concentration of 1 mM for 18 h at room temperature with mixing at 1250 rpm.aS oligomers were purified by GF, and through analysis of the number of bleaching steps,


the authors were able to quantify the average number of aS monomers that constituted thepurified oligomers, i.e., between 25 and 32, depending on the labelling densities.Conway et al. (2000) generated oligomeric intermediates by incubating aS in solution at 37 °C

without agitation, isolated oligomers with a molecular weight higher than 600 kDa, as determinedby GF, and analysed them using AFM. These oligomers showed a spherical shape and heightsvarying between 2 and 6 nm. Upon incubation for 72 h at 4 °C, the spherical oligomers formedlarger circular or elliptical ring-like structures, with diameters between 35 and 55 nm for thecircular rings and with widths between 35 and 55 nm and lengths between 65 and 130 nm forthe elliptical rings.Giehm et al. performed aggregation of aS by shaking at 37 °C in the presence of glass micro-

spheres to increase reproducibility and, using SAXS, found that 30–40% of the protein formedoligomeric species (SAXS measurement does not affect the monomer–oligomer equilibrium).Moreover, they could reconstruct a low-resolution structure of the oligomers and foundthat aS oligomers in these conditions are shaped as closed wreaths 18 nm long and 9 nmwide, with a diameter of ∼4.5 nm. Comparing the volumes of monomers versus oligomers(460±30 nm3) obtained by modelling and starting from the constraints established by SAXSmeasurements, the authors suggested that the oligomers are constituted by ∼16 aS molecules.Moreover, the model suggested that these oligomers are on-pathway and may be the buildingblocks of aS fibrils (Giehm et al. 2011b).The oligomeric structure proposed by Giehm et al. could also account for the pore-forming

activity thought to be one of the toxic mechanisms of aS aggregates. However, consideringthat oligomers are transient and heterogeneous, it seems quite improbable that ∼30–40% ofthe aS in solution consists of a monodisperse ensemble of oligomers. Our interpretationis that this result is an approximation due to the low resolution of the technique, which, however,remains one of the most effective techniques used in vitro to characterize these aggregationintermediates.In addition, E46K protofibrils, isolated by GF after protein lyophilization and resuspension,

show a recurrent annular shape (d=10–15 nm) (Fredenburg et al. 2007), similar to that observedfor wt, A30P and A53T aS (Conway et al. 2000; Lashuel et al. 2002).Other biophysical techniques have been used in the attempt to classify oligomeric species.

Kaylor et al. (2005) identified two main species by fluorescence spectroscopy, i.e., early andlate oligomers: the first ones are transient and convert rapidly to fibrils at the end of the lagphase, while the late oligomeric species coexist with the fibrils even at the end of the reaction.A Fourier transform infrared spectroscopy (FTIR) analysis of the same two oligomeric speciesin comparison with aS monomers or fibrils showed that the early oligomeric species have a frac-tion of β-sheets comparable with that of aS fibrils (∼43%), while in the late oligomers, the β-sheetstructure is ∼18%. The authors also generated the triple aS mutant Y125W/Y133F/Y136F, suit-able for performing FRET measurements. The differences in FRET efficiency allowed determi-nation of the mean distance between Tyr39 and Trp125 for the different aS species during theaggregation process. The average value decreased from 2.49 nm for the monomers to 2.19 and1.88 nm for the early and late oligomers, respectively; the fibril spectrum showed no energy trans-fer, most likely because the fibril structure quenches the tyrosine emission. To further character-ize these oligomeric species, the same research group performed DLS experiments (Dusa et al.2006): they measured the scattering signal of the supernatants at different time points duringthe aggregation reaction. At the early phases of the fibrillization, only monomers were foundin the sample (mean radii were inferred from the scattered light: Rmonomer=3.1±0.2 nm),


while after a few hours, oligomers with Rearly oligomer=22±2 nm were revealed. At the end ofthe fibrillization, smaller oligomeric species were in the sample supernatants (Rlate oligomers=12–15 nm).Apetri et al. (2006) analysed aS aggregates during an entire fibrillization process at 37 °C

without shaking (initial monomer concentration 300 μM) to study oligomeric intermediates’morphology and secondary structure. As first species, they found spheroidal oligomers thatwere divided into two different classes on the basis of their heights, as detected by AFM(1.4–3.5 or 4.0–7.5 nm); after a few days, chains of spheres were found in the samples, withmean heights of ∼5 nm, which disappeared upon fibril formation. The secondary structure ofthe different oligomeric species was, in this case, evaluated by Raman spectroscopy: the authorsassigned the band at 1650 cm−1 that typically corresponds to α-helical structures mainly to aSmonomers in solution. The same analysis revealed that the two oligomeric species have a de-crease in their α-helical content and an increase in their β-sheet structure content comparedwith the monomers.More recently, experiments using Trp fluorescence emission variations were performed to eva-

luate the residues involved in the formation of aS oligomeric species (van Rooijen et al. 2009b).Oligomers of aS Trp mutants (F4W, Y39W, A69W, A90W, A124W and A140W) were obtainedby aggregating mutated aS at a 1 mM concentration overnight at room temperature and 300rpm, followed by 2 h at 37 °C without shaking. The aggregated aS was then filtered (0.2 μm)and injected into a GF column, and the oligomers were collected in the void volume. While allthe aS Trp mutants in the monomeric form have a Trp emission maximum between 347 and355 nm, the peaks of the F4W, Y39W, A69W, A90W mutants were considerably blue-shifted inthe oligomeric species, suggesting that these residues in the oligomers were less exposed to the sol-vent. A124W and A140W showed a minimal blue-shift, suggesting that the negatively chargedC-term is not buried in the oligomer core, which seems to comprise between residues 4 and 90.A recent step forward is studied by Cremades et al. (2012), which showed that aS oligomeric

species are present as a heterogeneous ensemble during the aS aggregation process using single-molecule fluorescence experiments (smTCCD). The oligomer distribution was divided into threedifferent classes: small (2–5 monomers), medium (5–15 monomers) and large (15–150 mono-mers) oligomers, confirming the variability reported before. The mass fraction of oligomers pres-ent in solution at all the observed time points (<5%) is in good agreement with part of the resultsmentioned above.As previously reported, the oligomeric species obtained in vitro vary considerably according

to the experimental conditions, and this heterogeneity is further enhanced in experiments thatinvolve additional physical or chemical procedures to induce oligomer formation. The use ofelaborate protocols aims to obtain more reproducible results and/or to prove that an excessof some chemical compounds can lead to the formation of specific types of oligomers with par-ticular toxicity. However, it remains to be proven whether these procedures lead to the identifica-tion of aS oligomers that are relevant to the study of the molecular mechanisms ofsynucleinopathies or if they are just another paradigm of aS oligomeric species of unprovenrelevance among all others.

4.1 Physical methods for obtaining aS oligomers

Several physical methods have been applied to aS oligomer ‘production’. In a first case, lyophi-lization of recombinant proteins after purification and their subsequent resuspension led to the


formation of stable aS oligomers that were analysed by different biochemical and biophysicalmethods (Apetri et al. 2006; Lashuel et al. 2002). Fragmentation of aS fibrils by supercoolingwas also used to obtain an enriched preparation of aS oligomers with electrophysiological activityon artificial membranes and whose structural features were extensively characterized in twopapers (Kim et al. 2008, 2009). In particular, oligomer morphologies were characterized byAFM and TEM, whereas their secondary structure, a β-sheet non-fibrillar structure, was studiedby solid-state NMR.The sonication of recombinant aS fibrils results in the formation of so-called ‘seeds’. Although

seeds are heterogeneous objects of variable size, they turned out to be an extremely useful tool inaggregation experiments in which the introduction of seeds as preformed nucleation centres intothe aggregation mix led to more reproducible results (Giehm & Otzen, 2010; Wood et al. 1999).Moreover, aS seeds have been used to trigger aS aggregation in the cellular compartments usingcell lines or primary neuronal cultures (Luk et al. 2009; Nonaka et al. 2010; Volpicelli-Daley et al.2011; Waxman & Giasson, 2010), as will be discussed more in detail in Section 6.1.

4.2 Chemical oligomers

The presence of specific chemical compounds in the aggregation mix was shown to influence theaS aggregation process, leading to the formation of off-pathway oligomerization products throughthe formation of covalent bonds. Among these products, particularly interesting are the oligo-mers formed in the presence of docosahexaenoic acid (DHA) (De Franceschi et al. 2011) or4-hydroxy-2-nonenal (HNE) (Nasstrom et al. 2011; Qin et al. 2007b). DHA is a fatty acid presentin high abundance in the brain regions where LBs are present, whereas HNE is a lipid peroxi-dation product that can modify proteins, most likely by forming adducts or aggregates in parkin-sonian brains (Yoritaka et al. 1996). De Franceschi et al. obtained covalent oligomers afterincubation of aS with DHA at the molar ratio of 1:50; these oligomers are characterized by an-nular and spherical morphologies, as detected by AFM and TEM. Moreover, FTIR and circulardichroism (CD) analysis showed a high content of α-helical and random structures but almostno β-sheet structure, suggesting that these oligomers are not on-pathway intermediate aggregatesof mature amyloid fibrils. DHA can therefore modulate the properties of the covalent aS oligo-mers, which were shown to be toxic to SH-SY5Y neuroblastoma cells (De Franceschi et al. 2011).HNE–aS oligomers also showed an annular morphology, as detected by AFM (inner diameter

30–50 nm, outer diameter 80–100 nm), but also a curved protofibrillar structure 100–200 nmlong and 2–4 nm high. However, in contrast to DHA–aS oligomers, their secondary structureis characterized by a shift in the CD spectra to wavelengths typical of a β-sheet conformation(Nasstrom et al. 2011). HNE–aS oligomers are internalized by SH-SY5Y, which show a reducedviability induced by the oligomers’ toxicity.Another physiologically relevant molecule responsible for the formation of covalent oligomers

of aS is DA. DA is particularly important in PD pathogenesis because the prominent death ofdopaminergic neurons in parkinsonian brains suggests that this neurotransmitter may play akey role in the toxic events that lead to cell death and neurodegeneration. Bisaglia et al. showedthat incubation of aS and DA leads to the formation of aS oligomeric species that are stable whenseparated in a sodium dodecyl sulphate (SDS)–PAGE (Bisaglia et al. 2007). The structure ofaS/DA oligomers was also studied by groups (Cappai et al. 2005; Rekas et al. 2010) usingSAXS and CD; size-exclusion chromatography allowed separation of aS/DA dimers, trimersand higher molecular weight oligomers. Dimers and trimers showed an increased content


of β-sheets and turns, while SAXS data permitted the elaboration of a low-resolution modelfor the aS/DA oligomers shape. It is worth noting that 3,4-dihydroxyphenylacetaldehyde, i.e.,the monoamine oxidase metabolic product of DA, can form SDS insoluble oligomeric adductswith aS (Burke et al. 2008).Interestingly, the presence of metal ions in the aggregation assay also induces the formation of

SDS-resistant aS oligomers, whose main characteristic is again their annular or spherical structure(Kostka et al. 2008; Lowe et al. 2004). The type of metal ion appears to affect the dimensions andshape of the aS oligomers formed: spherical particles (0.4–8 nm) are obtained from aS in thepresence of copper(II), iron(III) and nickel(II); and larger spherical oligomers (5–8 nm) resultfrom the aggregation of aS with magnesium(II), cadmium(II) and zinc(II). Moreover, calcium(II) and cadmium(II) lead to the formation of annular oligomers with diameters, respectively,of 70–90 and 22–30 nm (Lowe et al. 2004). To summarize, aS oligomeric species generatedin vitro have to be taken cum grano salis. The numerous studies reported above suggest thatthe heterogeneity observed can hardly be ascribed only to the intrinsic heterogeneity of the aSaggregation products and is most likely strongly affected by the experimental conditions underwhich aggregation assays are performed. On this ground, it appears difficult to firmly ascribea pathological function to these oligomers without robust proof of their presence and activityin PD cell or animal models or, even better, in parkinsonian brains.The key future challenge for PD researchers is to conceive of an experimental method to

observe these oligomers in action within the cell to determine which among the several aSoligomeric species are truly relevant for the pathology.

5. aS fibrils: structure and properties

5.1 Fibril structure

5.1.1 Fourier transform infrared and small angle X-ray spectroscopy measurements

Secondary structure information of aS fibrils can be obtained by FTIR (Fourier transform infra-red) spectroscopy, which provides a fingerprint of the secondary structure distribution rather thanprecise structural details. FTIR spectra of resuspended aS fibrils (Kaylor et al. 2005; Qin et al.2007a) showed a maximum at 1630–1635 cm−1, characteristic of the vibrational frequenciesof intramolecular parallel β-sheets. The broad shoulder at ∼1650 cm−1 reflects the presence ofdisordered structures, i.e., parts of the N- and C-terms. We are aware that FTIR analysis cannotcompete with NMR in term of resolution; however, it is clear that it is much more affordable andeasier to use FTIR fingerprints to compare the secondary structure characteristics. Furthermore,this technique has potential for the study of aggregation directly in cells, as recently reviewed forother aggregating proteins (Miller et al. 2013).Information on fibril quaternary structure can be obtained by SAXS experiments on full-length

fibrils (Giehm et al. 2011b). As previously reported, Giehm et al. presented SAXS data on aSoligomeric species that they suggested represent the building blocks of mature fibrils. TheseSAXS experiments allowed the authors to obtain a reconstruction of aS fibrils derived fromthe analysis of the data on oligomeric species, whose resolution is very low compared with themodels obtained using the methods previously introduced. However, this reconstruction allowsan establishment of a relationship between aS oligomer structure and aS fibril formation, provid-ing a possible characterization of aS assembly, in which fibrils are built by a class of aS oligomers.


5.1.2 NMR and EPR to probe aS fibril structure

Among the aS aggregation products, amyloid fibrils are the best characterized, being stablein solution and easy to isolate. They share several structural characteristics with amyloidfibrils formed by other aggregating proteins (Dobson, 1999) linked to neurodegenerative dis-orders, such as AD, Huntington’s disease and amyotrophic lateral sclerosis (Ross & Poirier,2004).A first attempt to define the structure of fibrillar aS was accomplished by X-ray diffraction

(Serpell et al. 2000). Serpell et al. measured X-ray diffraction patterns of dried fibrils obtainedfrom wt aS, from a deletion mutant consisting of the first 120 residues, and from the pathologicalmutants A30P and A53T. All samples showed a ring shape at a spacing of ∼0.47 nm, and the wtalso showed a weak diffraction at ∼1.0–1.1 nm; the X-ray data suggest a β-sheet structure for allthese fibrils, with the β-strands perpendicular to the fibril axis.More resolved structural information has been acquired by nuclear and electron magnetic

resonance techniques, which were applied to the characterization of the aS fibril structure.The main goal was to define both the structures of the fibril core and of the N- and C-termsto ultimately define a model for aS amyloid fibrils.The structural features of aS fibrils were first studied by site-specific spin labelling and by

electronic paramagnetic resonance (EPR) (Der-Sarkissian et al. 2003). This work was based ona nitroxide spin label, used as reporter molecule, whose structural environment and proximityto other reporter molecules can be deduced from the modulation of its EPR properties. EPRspectral analysis allowed the determination of loops, buried sites and secondary structureelements in amyloid fibrils. The data suggested that aS fibrils have a core region (including notonly the NAC domain) organized in an ordered, in-register, parallel structure, consisting of68 amino acids from residues 34 to 101. The N-term region showed a more heterogeneousand less ordered structure, while the C-term was described as completely unfolded.In an ss-NMR study (Heise et al. 2005), the authors used correlation experiments, reverse

isotope labelling and novel strategies for the characterization of the aS fibril structure. Theresidue-specific resonance assignments of 48 amino acids in the aS core region (from residues38 to 95) were obtained using two-dimensional (2D) and 3D correlation experiments. Thesedata also allowed determination of the structure of the β-sheet core region, constituted in the pro-posed model by six β-strands. Within this core region, a turn and a loop were identified (at resi-dues 66–68 and 83–86). Moreover, the authors found, in accordance with the previous EPRstudy (Der-Sarkissian et al. 2003), that the N-term starting from residue 22 is quite rigid, whilethe C-term is mobile and has no secondary structure. Through the differences in 13C and 15Nshifts found for the amino acids between E83 and I88 and for the side-chain atoms, Heiseet al. identified two different types of fibrils: a relatively ordered form and a disordered one(Heise et al. 2005). The parallel TEM imaging experiments performed suggested that the differ-ences in the structure detected by ss-NMR correspond to at least two different fibril morpholo-gies (see Section 4.2).An EPR study was performed on fibrils obtained from 83 different point mutation spin-

labelled derivatives to define more precisely the fibril boundaries of the core region and to de-termine its secondary structure elements (Chen et al. 2007). Consistent with previous work,Chen et al. found a core packed region that extended from amino acids 36 to 98 and, withinthis region, some mobile and flexible areas. Moreover, they confirmed the fibrils’ parallel arrange-ment, in which the β-strand planes are perpendicular to the fibril axis.


In 2008 Vilar et al. (2008) presented new NMR experiments on aS fibrils. Hydrogen–deuterium(H/D) exchange measurements suggested, as expected, that most of the residues from 30 to 110were not exposed to the solvent. Moreover, H/D exchange showed biphasic behaviourconsisting of two exchange rates, consistent with the presence of two distinct conformationsin the sample, as previously reported (Heise et al. 2005). Residues 1–27 and 104–140 appearedflexible and showed no secondary structure, as indicated by their fast exchange rates. ss-NMRexperiments on 15N,13C-labelled aS fibrils allowed the assignment of 70% of the residues inthe core region (35–95), but no heterogeneity was detectable with this technique in the samples.Based on ss-NMR and EPR experiments performed by other groups and on their own work,Vilar et al. proposed a new structural model, in which the core of the fibrils (from residues30 to 110) is a β-sandwich consisting of five β-strands with different possible packings andsolvent exposure, while the N- and C- terms are unstructured. This model also accountsfor the data on the proteolysis resistance of the fibrils in solution (Pivato et al. 2012; Qin et al.2007a).In a recent work, Comellas et al. by assigning via ss-NMR 106 residues, versus the 62 residues

presented in a previous ss-NMR study (Heise et al. 2005), demonstrated that the core of the fibrilgoes from residues 38–96 (Comellas et al. 2011). The novelty of their model is due to the iden-tification of a motif repeated twice in the fibrils’ secondary structure, consisting of a long β-strandand two short β-strands, from residues 38 to 66 and from 68 to 96. Moreover, the authors couldidentify structural features in both the N- and C- terms (Comellas et al. 2011), in disagreementwith the previous NMR and EPR works, in which these parts of the protein were describedas completely unfolded. More recently, the same research group published a comparison betweeneach pathological mutant and wt aS fibrils in terms of structural features, based on an ss-NMRstudy. The authors showed that, despite the slower aggregation rate of A30P mutants, the result-ant amyloid fibrils have chemical shifts and secondary structure similar to the wt aS (Lemkau et al.2012), whereas the structures of A53T and E46K fibrils show some small differences aroundthe mutated region and significant variations compared with the wt aS (Lemkau et al. 2013).Given these latest results, the mutants’ pathogenicity may be attributable not only to the proper-ties of the aggregation intermediates but also to the characteristics of the final products of theaggregation process.Over the past decade, structural data has provided a satisfactory description of the aS fibrillar

core and allowed the elaboration of models that describes the aS monomer structure within thefibril (Fig. 3).The latest model clearly suggests that the pathological mutations E46K and A53T are struc-

tured, while the missense mutation A30P is located in an unstructured loop; moreover, theN- and C-terms are not completely unfolded, as hypothesized in the first studies, but presentsome minor structural elements that could be crucial, as we will see in the next section, forthe formation of mature fibrils.

5.2 Fibril imaging

TEM and AFM are the main imaging techniques used to study aS fibrils and to determine theirmorphology, in particular, their mean diameters, mean lengths and periodicity. Optical imaging,even in its more recent developments, such as stimulated emission depletion microscopy(STED), still has not achieved a sufficient resolution compared with the actual aS fibrildimensions.


5.2.1 Fibrils observed with TEM

TEM allows imaging of aS fibrils, but its resolution is limited not only by the electrons’wavelength but also by the staining procedure. Information on fibril length and periodicitycan therefore be estimated, but diameter measurements are strongly biased by the presenceand dimensions of the grains of uranyl acetate salt. Therefore, TEM experiments on fibrils aretypically used to qualitatively evaluate aggregate morphology and, as such, are reported inmost of the publications on aS aggregation (Heise et al. 2005; Vilar et al. 2008).

5.2.2 Fibrils measured by AFM

AFM imaging, which does not require staining and is less invasive, is typically used to measurefibrils’ diameters and periodicity. AFM can be applied to biological studies using different ima-ging modes (Casuso et al. 2011), and in most of them, it only mildly affects fibril integrity andstructure and therefore has been used to develop structural models of fibrils and hypotheseson fibril assembly.This method allowed identification of a strong dependence of aS fibril morphologies, lengths

and heights on the experimental conditions, i.e., solution pH, salt concentration and aS concen-tration (Hoyer et al. 2002; van Raaij et al. 2008), in good agreement with the kinetics and the struc-tural data previously discussed.Van Raaij et al. performed AFM experiments to determine the dependence of fibril length

distribution on protein concentration and found that longer fibrils are obtained by increasingthe initial aS concentration. Moreover, they found a concentration threshold below whichthere is no significant fibrillization. This concentration has been estimated to be below 15 μM.Hoyer et al. (2002) studied aS fibril morphology at different pHs (from pH 4.0 to 7.0) and used

AFM to identify the periodicity in the obtained fibrils. They found only untwisted fibrils when aSwas aggregated at pH 7.0, whereas at pH 6.0, they could find two twisting modes: right-handed

3450

5156

62

66

6879

8085

8896

*

*

30

38

106

*

108

N-term

C-term

Fig. 3. Schematic representation of the arrangement of an individual aS molecule within a fibril, followingthe results obtained by Comellas et al. by solid-state NMR. N-term and C-term do not present a definestructure, but still present some structural features, in disagremeent with part of the previous works.Conversly, the central part of the protein is folded into six β-strands: solid-state NMR allowed also theidentification a repeated motif, constituted by one long and two short β-strands (from amino acids 38 to66 and from 68 to 96). Structural features (turns, loops and β-strands) and the associated number ofresidues are reported in the schema. Fibril axis is perpendicular to the plane that contains the foldedprotein (Comellas et al. 2011).


with an ∼45 nm periodicity and left-handed with a periodicity of ∼95 nm. At lower pH, theyreported a decreased amount of detectable fibrils, while amorphous aggregates were abundant.Conversely, other authors (Qin et al. 2007a; van Raaij et al. 2006) reported twisting in fibrilsobtained at pH higher than 6.0, and, moreover, van Raaij et al. compared the morphologicalcharacteristics of wt aS, A30P and E46K fibrils by measuring their heights and periodicitieswith a robust statistical analysis. They found mean heights of 10.0±1.1, 8.1±1.2 and9.0±0.8 nm for wt aS, A30P and E46K fibrils, respectively, and periodicities of 81±24,103±20 and 76±34 nm. The authors were not able to characterize A53T fibrils because, intheir experimental conditions, they could only obtain large amorphous aggregates(diameter>1 μm) for this mutant.Khurana et al. performed AFM experiments that allowed the characterization of fibril mor-

phology and led them to hypothesize a fibril assembly mechanism at the late stages that leadsto mature fibrils (Khurana et al. 2003). They proposed that aS monomers, in aggregating con-ditions, first assemble into protofilaments, which are 3.8±0.6 nm high; after protofilamentelongation, two of them intertwine and form protofibrils (mean height 6.5±0.6 nm), likely tominimize the exposure of hydrophobic interfaces. For the same reasons, two protofibrils inter-twine into mature fibrils that are 9.8±1.2 nm high. The authors also reported a clear periodicityof 100–150 nm for protofibrils and an amplitude of ∼1 nm.In 2007, Qin et al. reported detailed height measurements for aS fibrils, digested fibrils and

deletion mutants’ fibrils lacking the N-term, C-term or both, to determine the role of the twoterminals in fibril assembly (Qin et al. 2007a). The authors proposed an average height for canoni-cal fibrils of 13.6±0.7 nm, while all the other fibrils had different sizes. In particular, it hasbeen shown that the removal of the C-term (truncated aS from amino acid 1 to 103) leads tothe formation of protofibrils (mean height 5.7±0.3 nm), while in the absence of the N-term(from amino acid 30 to 140), the mean height is 10.7±0.5 nm, suggesting that the fibrils’ struc-ture is conserved but that the mean diameter is smaller because of the N-term removal. Theabsence of both the N- and C-terms leads to the formation of protofilaments whose mean heightis 2.9±0.3 nm, a value comparable with the data on trypsin-digested fibrils (2.8±0.2 nm).The results obtained by Qin et al. highlighted a key role for the C- and N-terms in mature fibril

formation and strongly supported the model previously proposed by Khurana et al. on the latesteps of aS fibril hierarchical assembly (Khurana et al. 2003).Recently, different types of chimaeric aS covalent dimers were generated to investigate the rela-

tive orientation of aS monomers within the amyloid fibrils (Pivato et al. 2012). The dimers wereobtained by disulphide bridge linking of aS molecules in different ways, i.e., through their N-term,through their C-term, or by simply expressing wt monomeric aS in tandem. A fourth dimer, inwhich the two fibril core regions of the two aS molecules were juxtaposed, was also preparedand purified. These chimaera induce different minimal constraints to the possible orientationsof the aS molecules stacking within the fibrils, showing that CC and NN dimers exhibit morphol-ogies and mean diameters very similar to wt aS, whereas NC shows shorter diameters and theDC dimer exhibits a very broad distribution of diameters from 3 to 15 nm, as if the fibrils inthis case were clustering.

5.2.3 Super resolution fluorescence imaging of aS fibrils

Fluorescence microscopy has rarely been used to study aS aggregation in vitro because of its lowresolution. More recently, super resolution methods have been applied to the characterization of


fibril features and their growth dynamics; however, these techniques are not widely used, mainlybecause they are not commonly available. In a very recent paper, Roberti et al. (2012) studied aSfibril structure and growth using photoactivated localization microscopy (PALM). They achieveda resolution of ∼20 nm and were able to identify, using aS conjugated with two different deriva-tives of rhodamine spiroamides, i.e., RSA577 and RSA617, the dynamics of fibril accretion,which, interestingly, occur in their aggregation process only at one single end of the fibril.Recently, Ries et al. (2013) used the super resolution method termed binding-activated localizationmicroscopy (BALM) to image aS fibrils in vitro. This method is based on the use of an amyloid-specific dye (pentamer-formyl thiophene acetic acid) that binds to aS fibrils, fluoresces, and isthen bleached, to determine fibrils’ diameters without being diffraction-limited. The apparentdiameter of aS fibrils measured by BALM is 14 nm, meaning that the achieved resolutionis below 20 nm. In both the cases, PALM and BALM images of aS fibrils resemble the AFMmicrographs very closely but do not add information about fibril features.These methods could be interesting if used in cells to characterize aggregates in a more bio-

logical milieu. The resolution of this method can break the diffraction limit and is therefore verypowerful in fixed cells; however, the acquisition is very slow, which presents a problem for itsapplication to the imaging of live cells (see Section 6.1).

5.3 Fibril modelling

Taking into account most of the structural data on aS fibrils present in the literature, it has beenpossible to recapitulate the results in a structural schema representing the most probablefolding of aS molecules within the fibrils (Fig. 3), which is in good agreement with the putativeaS β-sheet propensity calculated from aS primary sequence (Fig. 2) (Suk et al. 2010; Zibaee et al.2007). Moreover, the assembly of aS molecules in fibrils was proposed to be a general hierarchicalmechanism, from protofilaments to protofibrils to fibrils.However, the recognition of polymorphisms in fibrils samples by NMR, EPR and microscopy

techniques and the differences in the quantitative analysis on fibril diameters, even when per-formed with the very same technique (Sections 4.1 and 4.2), suggest that even small changesin the general aggregation conditions and in the sample preparation can lead to large variationsin the results obtained.In terms of differences between the fibrils obtained in vitro from wt aS and from the pathologi-

cal mutants, it seems that there are some structural differences around the mutation regions, atleast for A53T and E46K, while morphological or dimensional differences can be found for all ofthe mutants.An interesting observation arose from an aggregation assay performed with chimaeric aS

mutants, in which β-sheet-prone regions in the aS sequence were shuffled (Suk et al. 2010).The authors showed that even after scrambling the aS sequence, some of the mutants, butnot all, were still able to form fibrils. This result suggested that the formation of aS fibrils is asso-ciated with the possible spatial rearrangements of the β-sheets composing the structure.Moreover, the formation of the initial nucleation centre is affected by the presence of gapsbetween the different β-sheet regions.Our opinion on this issue is that many of the structural and morphological differences among

fibrils reported in the literature may not only depend on the different aggregation intermediates,including protofilaments, protofibrils and fibrils (see Section 3.2), present in the same samplebut may also be, for the multiple amyloidogenic mechanisms of fibril formation, strongly affected


by aggregating conditions that are still uncharacterized but that can result in different final aggre-gation products. Interestingly, all pathological mutations in the aS sequence, excluding A30P, arelocated in the core region of the fibril, suggesting that they can have some consequences not onlyin the assembly of aS fibrils but also in the formation of the oligomeric species.If even small differences in the protofibrillar and fibrillar aggregation products could be

relevant to their potential effects in vivo, a fine and detailed characterization of even the tiniestdissimilarities is pivotal for designing the most appropriate therapeutic strategy.Moreover, an in-depth characterization of aS fibrils purified from LBs extracted from parkin-

sonian brains or from cellular models for aS fibril formation is still lacking, although TEM imagesof aS fibrils have been available since the discovery of aS presence in LBs (Spillantini et al. 1998).These data will allow the exploitation and testing of the knowledge acquired through in vitrostudies in physiologically relevant models.

6. Aggregation in cell models

Recently, efforts to link the in vitro aggregation properties of aS to their biological and pathologicalsignificance have increased tremendously, along with interest in recasting these biophysical datainto biologically relevant processes.The main techniques used to study protein aggregation in cells typically require fixation, label-

ling with antibodies (suitable for TEM or fluorescence microscopy) and fluorescent dyes recog-nizing amyloid fibrils. These methods allow the detection of aS aggregates, but little detailedinformation can be obtained on the kinetics of the aggregation process in the cellular environ-ment and on the structural properties of these cellular aggregates.Moreover, the study of aS aggregation in cells presents additional problems that should be

outlined here: (i) aS overexpression itself is not always enough to trigger aggregation in cells,the process is often accelerated by exogenous agents (i.e., seeds obtained by sonication of aSfibrils or exogenous co-expression with synphilin) (Buttner et al. 2010; Engelender et al. 1999;Luk et al. 2009); and (ii) early oligomeric species are rarely observed, mainly because their insta-bility and transient nature impedes their accumulation to detectable levels.Generally, discriminating among different aS aggregation intermediates and fibrils in cells re-

mains a difficult task because the resolution of conventional optical microscopy is low (200–300 nm)compared with the objects observed (fibril length is of the order of micrometres, but theirdiameters are ∼10 nm). Advanced optical microscopy techniques, such as fluorescence lifetime,fluorescence recovery after photobleaching, and fluorescence anisotropy, may provide additionalinformation. However, these data require interpretation to become relevant for describing specificaS aggregation intermediates. Conversely, super resolution techniques require cell fixation andeither specific expensive dyes for staining, in the case of STED microscopy (for example,Atto), or photoactivatable dyes or proteins, in the case of PALM, to obtain a sensible improve-ment in the quality of the data. Nevertheless, the recent use of advanced and super resolutionfluorescence or dye-free microscopy techniques that allow a study of the dynamics of the aggre-gation process in a cellular environment have provided novel results, as described in the nextparagraphs.

6.1 Aggregation kinetics and aggregate properties in cells

Several studies employ immunocytochemistry and the conventional fluorescence microscopytechnique to identify aS aggregation in cells. These assays require cells transiently transfected


with aS or stable cell lines overexpressing aS. Moreover, aS aggregation in cells is triggered byexogenous agents, such as recombinant aS seeds or oxidative stress (Luk et al. 2009; Nonakaet al. 2010; Roberti et al. 2007; Waxman & Giasson, 2010). A few days after treatment(from 2 to 7 depending on the experimental conditions), cells are fixed and immunostaining isperformed to visualize aS with fluorescence microscopy.

6.1.1 aS fused with fluorescent proteins (FPs) or tags

As an alternative, aS fused to FPs can be used (McLean et al. 2001). FPs are valuable tools,being detectable in live cells, with the caveat that they are much larger than aS (enhancedgreen fluorescent protein – EGFP – is ∼28 kDa, while aS is 14.6 kDa). Moreover, FPs have aβ-barrel structure, whereas aS, at least, the fraction that seems to be more aggregation prone,is in an unfolded state in the cell cytoplasm.The possibility that aS fused to a FP could behave differently from the wt, could hamper the

use of these constructs to study aggregation in live cells or in living organisms (McLean et al.2001). McLean et al. observed that aS fused to EGFP (or other tags) at the C-term can formlarge inclusions when overexpressed in mouse-cultured primary neurons, whereas no detectableaggregation was present when neurons were transfected with untagged aS, suggesting thatmanipulation of C-term aS may enhance its aggregation propensity, possibly altering the long-range intramolecular interactions normally present. Moreover, using the same experimentalsetup, the authors observed differences in cellular aggregation between PD pathological mutantsand wt aS fused with EGFP. A53T aS-EGFP and wt aS-EGFP showed inclusions in ∼5% oftransfected cells, while A30P aS-EGFP did not cause the formation of large inclusions.However, other research groups successfully used aS-EGFP variants to investigate aS behav-

iour and aggregation in immortalized cells (Outeiro et al. 2008; van Ham et al. 2010); the reasonfor such discrepancies may be the use of primary neuron cultures, which are likely to be moresensitive to variations in protein homeostasis due to aS overexpression. aS overexpression caninterfere with pathways that control the amount of proteins present within a cell. Accordingly,aS was shown to hamper protein clearance mechanisms (Ebrahimi-Fakhari et al. 2012), and simi-lar to chimaeric amyloidogenic proteins, aS was proposed to sequester essential cellular proteinsduring aggregation (Olzscha et al. 2011). McLean et al. concluded that overexpression of aS inprimary neuronal culture remains a good model for studying aS aggregate formation in cells.An unceasing effort is on-going to improve the study of the aS aggregation process and to

determine to what extent fluorescence probes or proteins fused to aS are suitable tools for study-ing aS aggregation in PD cell or animal models by live imaging. A first attempt in this directioncame from Roberti et al. who generated a construct in which aS was fused to a tag containing fourcysteines (C4 tag) inserted in a 12-mer peptide (FLNCCPGCCMEP). C4 tags can covalently bindbiarsenical compounds (termed FlAsH and ReAsH, depending on fluorescence excitation andemission wavelengths), which because of their high selectivity, are expected to label the proteindirectly in the cytoplasm of live cells after permeating the cell membrane. FlAsH and ReAsH be-come fluorescent only under the structural constraints imposed by binding to the C4 tag (Robertiet al. 2007). Since the C4 tag plus the biarsenical dye have a combined molecular weight of ∼1.3kDa, it is likely that they have negligible effects on aS structural features and its aggregation pro-cess. The aggregation properties of aS-C4 compared with those of the wt aS in a ThT fluores-cence assay in vitro, showed no relevant differences, confirming that the C4 tag and its boundfluorophore are a suitable probe for studying aS aggregation. The caveat of using FlAsH ligands


is that they do not bind only to the tetracysteine tag but could also stain proteins rich in cysteines.Furthermore, the staining is typically less efficient than conventional staining methods, such asimmunocytochemistry (Hearps et al. 2007; Stroffekova et al. 2001).A second method has been proposed by van Ham et al. who expressed and purified a recom-

binant form of aS fused to a yellow fluorescent protein (aS-YFP). They then used biochemicaland biophysical techniques in vitro to demonstrate that aS and aS-YFP (or other similar FPs)showed similar aggregation kinetics and products (van Ham et al. 2010).Clearly, further investigation is required to characterize the possible effects of fused FPs

or staining with biarsenical compounds not only on aS aggregation but also on aS functionand behaviour in the cellular environment (degradation, stability, post-translational modifications,etc.).

6.1.2 TEM and ThS staining on fixed cells

To overcome the limitations of optical microscopy, Nonaka et al. performed TEM onaS-overexpressing cells, in which aggregation of overexpressed or endogenous aS was triggeredby addition of seeds to the culture medium (Nonaka et al. 2010). TEM microscopy resolutionis approximately three orders of magnitude higher than that of conventional optical microscopytechniques (∼0.2–2 nm), and in fact, the authors were able to identify aS amyloid fibrils in cellcytoplasm. Recently, we were also able to image aS fibrils being phagocytosed by macrophagesand subsequently internalized into lysosomes by TEM (Codolo et al. 2013), further supportingthat TEM imaging is a powerful technique for visualizing fibrillar aS in cells. The limitation isthat TEM does not provide any information about the presence of other aggregation intermedi-ates or oligomers because of their heterogeneity, which makes them difficult to distinguish fromother intracellular structures.The presence of β-sheet fibrillar forms of aS triggered by seeds was also observed by ThS

measurements. Staining with an antibody against aS phosphorylated at Ser129 was used to vali-date the proposed cell model (Luk et al. 2009; Waxman & Giasson, 2010) as this phosphorylatedform of aS is highly enriched in LBs (Fujiwara et al. 2002). The limitation of these techniques isthat they require cell fixation and are therefore not suitable for live imaging.

6.1.3 Advanced fluorescence microscopy techniques applied to the study of aS aggregationin cells

aS aggregation in cell models was also studied by advanced fluorescence microscopy techniques,i.e., FRET, fast recovery after photobleaching (FRAP) and fluorescence lifetime imagingmicroscopy (FLIM). In 2006, a combination of FLIM and FRET was used to study aS confor-mations and self-interactions within cells (Klucken et al. 2006). The technique allows detection ofmolecular proximity and oligomerization by measuring the fluorescence lifetime of a fluorophore.In this particular case, human H4 neuroglioma cells were transiently transfected with an equimo-lar ratio of aS plasmids tagged with V5 or haemagglutinin (HA), fixed, and stained with primaryantibodies against V5 and HA and with secondary antibodies conjugated with two different fluor-ophores (Cy3 and Alexa 488, respectively). The fluorescence lifetime of an FRET donor fluor-ophore (Alexa 488) is shortened when it is close to (<10 nm) an FRET acceptor (Cy3), and thedegree of shortening is proportional to the donor–acceptor distance. Using this setup, the authorswere able to detect aS oligomers within intact cells, although they could not observe large aggre-gates. They also performed FLIM measurements and showed that the lifetime of the aS-HA


staining fluorophore Alexa 488 was ∼2.2 ns in the absence of the corresponding acceptor Cy3,while in the presence of aS-V5 stained with Cy3 at the C-term, the lifetime of Alexa 488decreased to 1.4 ns. The authors proposed two different interpretations: (i) aS monomers inthe cell cytoplasm fold and bring the N-term and the C-term of the same aS molecule intoclose proximity, causing a strong interaction; and (ii) the C-term of an aS molecule interactswith the N-term of another aS molecule, possibly at the early stages of the oligomerization pro-cess. These results correlate very well with some of the data on aS folding (Section 2), furthersuggesting that some of the reported conformational modifications may truly be associatedwith the early steps of the aggregation process. No information on the later steps of aS oligomer-ization was provided by this type of experiment in cells.Biomolecular fluorescence complementation (BiFC) is a method used to validate protein–

protein interactions; fragments of the FP sequence are fused with the proteins of interest, andwhen an interaction occurs, the fragments come into close proximity and the FPs reassembleand regain their fluorescence emission properties. This technique allowed Outeiro et al. (2008)to visualize aS oligomerization in living cells. They generated BiFC aS mutants, in which aS isfused with GFP fragments that become fluorescent only when aS molecules come together.Using this setup, they identified aS dimers and higher oligomeric species formed within cells,including the nuclear region. The authors also demonstrated that these low molecular weightspecies are essential for the formation of larger aggregates and can be related to the observedcytotoxicity. The induced toxicity decreased upon oligomer stabilization through a chaperoneprotein (heat-shock protein 70), which has been previously reported to reduce aS aggregationand toxicity in PD cell and animal models (Auluck et al. 2002; Klucken et al. 2004).Although these studies provided new clues into the initial steps of the aggregation process, they

are not informative enough to fully describe aS aggregation in cells. The difficulties in the deter-mination of the kinetics of the aggregation process and the dimensions and structural propertiesof the aggregation intermediates are major drawbacks in working with aS aggregation in cells.The biarsenical fluorophores mentioned above were used to study aS aggregation in

live SH-SY5Y cells by FRET (Roberti et al. 2007). Imaging over time with confocal microscopyallowed observation of aS aggregates in SH-SY5Y transfected with aS-C4 and stained with FlAsHand ReAsH 48 h after transfection (in this experiment, the aggregation of aS was triggeredby oxidative stress). The authors found a high variability in pixel-by-pixel FRET efficiency(10–80%), which was related to the proximity of aS-C4 stained molecules in different cells.They were able to define two ensembles by analysing the FRET efficiency distributions. Thefirst ensemble is characterized by FRET efficiencies between 60 and 80% and represents aSaggregates, while the second one, with efficiencies between 30 and 45%, corresponds to the non-aggregated form of the protein. This method, however, does not allow the determination of thesize of aS aggregates, but it gives a hint about the amount of aggregates and their distribution inlive cells and may be used to detect the effects of aS inhibitors or enhancers.Using the same aS-C4 plasmid to overexpress aS in SH-SY5Y cells and staining the protein

with ReAsH, the same authors investigated aS aggregation in living cells by FRAP and confocalfluorescence anisotropy (CFA) (Roberti et al. 2011). FRAP allows the monitoring of molecularmobility through fluorescence recovery of labelled aS over time after photobleaching.Therefore, FRAP data provide information on the mobile fraction of a protein and on its dif-fusion coefficient, D. In transfected cells, the regions without aggregates show complete recovery,with a biphasic behaviour characterized by a very fast recovery, which is associated with free pro-tein (D=10 μm2 s−1), followed by a slower recovery due to membrane-bound protein and to


small aggregates (D=0.08–0.15 μm2 s−1). Conversely, where large aggregates are present, most ofthe protein is immobilized; the small FRAP recovery observed in this case is most likely due tosmall aggregates exchanging with the larger aggregates (D=0.03–0.04 μm2 s−1).CFA provides information on changes in the size of aS aggregates; fluorescence anisotropy

r measurements were used to generate anisotropy maps and total fluorescence intensity If imagesof cells or regions of interest within them by classifying pixels in the images using If and rvalues. On this basis, the authors suggested the presence of four different aS species in thecell cytoplasm: monomeric free aS, membrane-bound aS and dim or bright aS aggregates.The FRET principle was exploited by Schierle et al. to study the kinetics of aS oligomerization.

Starting from the observation that aS aggregates can act as FRET acceptors for various fluores-cence proteins, they fused aS with a yellow fluorescent protein (aS-YFP) and measured FRETbetween the growing oligomers (acceptors) and the donor YFP (Kaminski Schierle et al.2011). After the in vitro characterization of the FRET sensor response to the aggregation process,the technique was used to study aS aggregation in the SH-SY5Y cell line. aS-YFP was introducedinto cells through electroporation, and measurements of the excited state lifetime allowed moni-toring of FRET pixel-by-pixel as a function of time after electroporation. The average lifetimewas ∼3020±20 ps for extracellular aS and 2970±70 ps for cytoplasmic monomeric proteinat the beginning of the process. The lifetime value decreased with time to 1480±40 ps after4 days. FRET efficiency, and consequently donor YFP lifetime, in this system is strongly corre-lated to aggregate size; however, a quantitative evaluation of oligomer size is not possible usingthis method.Super resolution methods, as mentioned before, can be applied to the study of aS aggregation

not only in vitro but also in cell models. The application of these methods required the delivery ofaS labelled with rhodamine spiroamide RSA577 mixed with wt aS in a 1:3 molar ratio into HeLacells and cell fixation after aggregate appearance (Roberti et al. 2012). Using PALM, Roberti et al.were able to identify the formation of spheroidal aggregates that were assigned to two differentgroups. The first group consisted of aggregates with diameters <40 nm, while the second in-cluded aggregates with diameters between 40 and 200 nm. Their results are considerably higherresolution compared with the images that can be obtained using confocal microscopy. However,PALM imaging requires fixation, mainly because of the large number of frames that are needed toreconstruct a high-resolution image; therefore, it does not allow the study of aS aggregation in livecells. However, to perform a sort of kinetic analysis of aS aggregation, cell samples can be fixed atdifferent time points for later analysis.The new techniques described here for studying aS aggregation in live cells confirmed some

in vitro observations, such as the identification of oligomeric species as the early products ofthe aggregation process. Moreover, they suggest that aS oligomers do not always proceed toform amyloid fibrils in cells, a process that is promoted in the presence of seeds. The level ofcharacterization of aggregates in fixed or live cells needs to be improved; a refinement of theavailable methods both in term of structural properties and available resolution must be reachedto tackle the link between aS aggregate properties and toxicity in cells.

6.2 aS aggregates toxicity

As gene dosage of SNCA is linked to pathology and aS is deposited in LBs, it seems reasonablethat point mutations in SNCA act through a gain of function rather than through a loss offunction.


Several different toxic mechanisms have been correlated to the aS aggregation process and toits oligomeric and fibrillar products. Given the heterogeneity of the aggregation products, it isreasonable to expect that cytotoxicity may occur through different mechanisms.aS could be defined as a polymorphic protein, plastically changing its conformation, secondary

structure and interactors (proteins or lipids). Given this premise, it is not surprising to find a largevariety of noxious effects ascribed to aS in its different observed conformations.Among the most studied toxicity mechanisms ascribed to aS is the ability of monomeric or

oligomeric aS to interact in a disruptive way with lipid membranes (Reynolds et al. 2011) or toform well-characterized pores similar to those observed in the presence of pore-forming toxins(Kim et al. 2009; Tosatto et al. 2012; Zakharov et al. 2007). To explore the pore-forming capacityof aS in its various forms in intact cells, aS was overexpressed in both SH-SY5Y and HEK293cells, in which aS can aggregate. Specifically, aggregate-like aS species observed in SH-SY5Y byfluorescence microscopy were localized in part of the cell membrane, with a concomitant increasein conductance (Feng et al. 2010). Increased conductance was also independently found inHEK293T cells overexpressing aS (Tsigelny et al. 2007). Based on these convergent observations,and in agreement with in vitro studies (Kim et al. 2009; Schmidt et al. 2012; van Rooijen et al.2009a), increased membrane conductance could be predicted to result in ion leakage withexpected deleterious effects. Formation of membrane pores may support Na+ diffusion fromthe extracellular environment to the cytoplasm, driven by its favourable electrochemical potential,with consequent unwanted electrical responses in neuronal cells. Ca2+ may also diffuse into thecytoplasm from the extracellular space, causing cytotoxicity. Furthermore, under the assumptionthat aS oligomers induce an increase in plasma membrane conductance, it is reasonable to predictthat they could also damage intracellular membranes, i.e., vesicles containing neurotransmitters(Mosharov et al. 2006) and mitochondrial membranes. The latter have been independentlyreported to be damaged by aS overexpression. Indeed, aS overexpression in HeLa cells inducesmitochondrial fission and variation in mitochondria morphology (Nakamura et al. 2011). Theauthors could not verify that the mitochondrial-associated aS was oligomeric, but they wereable to demonstrate aS oligomer interaction with mitochondrial-like membranes in vitro. Moreto the point, aS overexpression in PC12 cells (a pheocytochroma-derived dopaminergic cellline) leads to mitochondrial depolarization and subsequent mitochondrial-mediated cell death(Smith et al. 2005; Tanaka et al. 2001). Although a few studies suggest the formation of oligomericaS pores in vitro, their existence in vivo still awaits direct experimental evidence.The presence of toxic aS forms affects the physiology not only of membranes but also of other

cellular structures. In this respect, aS overexpression was shown to induce endoplasmatic reticu-lum (ER) stress in PC12 cells (Smith et al. 2005) and Golgi fragmentation in COS-7 cells (Gosaviet al. 2002). The observed ER stress induced by aS was further confirmed by the finding of aSoligomers within the ER in an A53T transgenic mouse model of PD (Colla et al. 2012).Both the ubiquitin–proteasome system and the autophagic lysosomal pathway have been

suggested as targets of aS aggregation products. These systems are involved in the clearanceof monomeric, oligomeric or aggregated aS, among other proteins, and have been shown tobe impaired in cell and animal models of aS aggregation, possibly because they are saturatedby an excessive amount of misfolded protein. However, the literature regarding this field iswide and sometimes controversial (Cuervo et al. 2004; Ebrahimi-Fakhari et al. 2012; Mak et al.2010; Plotegher & Civiero, 2012).A related aS toxicity mechanism that has been recently proposed is the sequestration of essen-

tial proteins during the aggregation process. As proof of concept, this sequestration mechanism


was shown for amyloid-forming chimaeric proteins in a recent paper (Olzscha et al. 2011).An analogous process was shown to occur for aS in primary neurons treated with seeds, inwhich synaptic proteins are sequestered into the aggregates (Volpicelli-Daley et al. 2011), andcan be predicted for other chaperone proteins present in cell cytoplasm. Molecular crowdinginduced by aS aggregates and the ability of aS to co-aggregate with other proteins or chaperones(Dedmon et al. 2005a; Xu et al. 2002) found in LBs (Shirakashi et al. 2006; Xia et al. 2008) furthersubstantiate this hypothesis.Finally, it has been shown that neuroinflammatory mechanisms (Alvarez-Erviti et al. 2011a;

Maguire-Zeiss & Federoff, 2010), oxidative stress and ROS production (Cremades et al. 2012;Freeman et al. 2013) can be induced by the presence of aS oligomers and fibrils.Interestingly, Codolo et al. showed that recombinant aS fibrils trigger inflammation in human

monocytes (Codolo et al. 2013), likely through a pathway that is general for needle-shaped objects,such as amyloid beta fibrils (Halle et al. 2008) as well as asbestos and carbon nanotubes (Palomakiet al. 2011). In addition, a specific form of aS oligomer directly released from cells was shown toactivate microglia (Kim et al. 2013), while other aS aggregates obtained from recombinant aS didnot result in an inflammatory response. Interestingly, both papers suggest that the activation ofthis process is mediated by an interaction with the membrane protein toll-like receptor 2.Although it is widely accepted that aS aggregation is toxic to cells, albeit that the nature of these

toxic species is still controversial, another possibility is that aS aggregation instead causes a de-crease in the available functional aS and a consequent loss of function, which leads, in turn,to cell death. aS has in fact been suggested to play an important role at the presynapse throughinteraction with the SNARE complex during vesicle exocytosis and neurotransmitter release(Burre et al. 2010; Nemani et al. 2010). Indeed, a recent report showed that aS oligomerizationis responsible for the inhibition of SNARE-mediated vesicles docking (Choi et al. 2013).Figure 4 summarizes the most important toxic mechanisms related to aS aggregation in

cells that could be addressed by biophysical methods. For example, organelles’ integrity in thepresence of aS aggregates could be studied by TEM, FRET or other microscopy techniques(Nakamura et al. 2008, 2011), or electrophysiological studies could monitor membrane permea-bilization due to aS oligomers (Kim et al. 2009). These major advancements in biophysics couldprovide new tools for addressing these open issues.A relevant question that remains unanswered in the etiopathogenesis of PD is when aS

oligomerization or aggregation is the cause of toxicity and when toxicity is, instead, the resultof upstream pathological events that affect aS physiological functionality and induce aSaggregation.

6.3 Misfolded aS transmission in cell models

Braak staging for PD indicates that LBs first appear in the olfactory bulb and that only later indisease progression do they form in other brain regions (Braak et al. 2003). This result suggeststhat the transmission of misfolded or aggregated aS from unhealthy to healthy neurons could bethe culprit for the spread of the pathology and the progression of the disease. This hypothesisgained further support from clinical evidence (Kordower et al. 2008; Li et al. 2008) showingthe presence of LBs in naïve neurons grafted in parkinsonian brains 10 years after surgery.The evaluation of this hypothesis requires many of the experimental approaches that fall in the

realm of biophysics. Several cellular studies have now confirmed that oligomeric aS can transfer


from a donor to a receiving cell (Desplats et al. 2009; Hansen et al. 2011). Although some resultssuggest that the transfer may occur via endocytosis, they did not provide any direct evidence.The transmission of fragments of recombinant aS fibrils was found to be responsible

for enhanced aS aggregation. The fragments serve as ‘seeds’ and trigger aS aggregation in therecipient cells (Nonaka et al. 2010), as evaluated by TEM and fluorescence microscopy imaging.Volpicelli-Daley et al. (2011) further showed that pre-formed recombinant aS fibrils were taken

up by cultured primary mouse neurons. These fibrils affected neural excitability and connectivity,as reported by calcium imaging of the hippocampal neuronal networks. Moreover, aS inclusionformation caused both the sequestration of some synaptic proteins and neuronal cell death.Volpicelli-Daley et al. further showed intracellular propagation of fibrils from soma to neuritesand vice versa using microfluidic devices as precast templates to guide the growth of neurites.

Proteasome impairment

Autophagic lysosomalpathway impairment

Axonal transport

Oligomers basedpermeabilization

ROS production

Monomers basedpermeabilization

Mitochondrial damage

Monomeric synuclein

Monomeric synuclein ubiquitinated

Monomeric synuclein alpha-Helix

Oligomeric synuclein

Fibrillar synuclein

Oligomers basedER stress

Exosome transmission

Nanotubes transmission

Fig. 4. Summary of the most recurrent toxic mechanisms proposed related to aS aggregation andtransmission: membrane damages (Feng et al. 2010; Mosharov et al. 2006), mitochondrial morphologyalterations, dysfunctions and damages (Devi et al. 2008; Nakamura et al. 2011; Smith et al. 2005; Tanakaet al. 2001), ER stress (Smith et al. 2005), oxidative stress and ROS production (Cremades et al. 2012;Freeman et al. 2013) and clearance mechanisms impairment (Cuervo et al. 2004; Ebrahimi-Fakhari et al.2012; Mak et al. 2010). Among the aS aggregates transmission mechanism exosome transmission(Alvarez-Erviti et al. 2011b; Danzer et al. 2012), axonal transport (Freundt et al. 2012) and nanotubestransport (Gousset et al. 2009) are reported.


In vivo fibril propagation was recently documented by immunohistological experiments: trans-mission from neuron to neuron in mouse brains after fibril injection induced the spread ofPD pathology to untreated brain regions. Interestingly, these mice showed some of the symptomsthat can be ascribed to PD (Luk et al. 2012).Despite these recent studies, the precise, and most likely multiple, mechanisms that cause aS

spreading from neuron to neuron or aS uptake by cells from the extracellular space are not clear.Several different hypotheses have been proposed: (i) direct cell-to-cell transfer due to theincreased protein concentration in the extracellular space, possibly caused by cell death; (ii) exo-somes or vesicles that protect the protein from the action of proteolytic enzymes and carry it torecipient cells; (iii) molecular chaperones that mediate aS transmission; (iv) misfolded aS thatmoves through the slow movement of axonal transport; and (v) membrane nanotubes that inter-connect cells over long-distances serving as highways for aS aggregates (Steiner et al. 2011).Hints about the possible mechanism came from two recent papers. The first proposed that

overexpressed aS can be transmitted from transfected SH-SY5Y cells to normal SH-SY5Ycells by exosomes (Alvarez-Erviti et al. 2011b; Danzer et al. 2012). The second showed that fibril-lar aS can be transferred along axons via anterograde axonal transport, with a velocity that wasinversely proportional to the aggregate size (Freundt et al. 2012).Among the other mechanisms, tunnelling nanotubes, which allow the movement of organelles

and vesicles (Rustom et al. 2004), seem to be promising candidates, mostly because of their pro-ven involvement in prion protein spreading. Indeed, it has been shown, by time-lapse confocalimaging of live neurons, that tunnelling nanotubes are responsible for the transfer of endogenousand exogenous prion proteins confined in lipid vesicles (Gousset et al. 2009).This emerging interest is related not only to the mechanisms that govern the transfer of aS

but also to the definition of those aS species that are specifically transferred. The latter representthe true key to understanding the pathological process as those species are likely to carry thestructural information required to spark aS aggregation in healthy neurons. These unansweredquestions are perfect candidates for future biophysical investigations

7. Conclusions

As can be seen from the contents of this review, a plethora of information is available on thekinetics of the aS aggregation process in vitro and on the properties of stable aggregation products.However, less is understood about aS oligomers, which appear to be heterogeneous, transient,numerically less abundant and, when generated in vitro, highly variable in nature. However,they are likely to be the most toxic species.Recently, the use of more powerful advanced microscopy techniques to study aS aggregation

in cells has begun to emerge. In the near future, the newly available methods should allow thecorrelation of specific aS aggregation intermediates to precise toxic events. The new methods,combined with appropriate experimental strategies, have the potential to unravel the sequentialcascade of non-toxic and toxic events triggered by aS oligomeric and fibrillar species, thus iden-tifying the primary and secondary toxic mechanisms and their timing in the progression of thedisease.Another relevant issue that biophysicists should consider is the translation of advanced fluor-

escence microscopy techniques successfully used to study aS aggregation in vitro into more physio-logical environments – live cells as the first step, with whole organisms, such as zebrafishand mice, as the ultimate goal.



8. Acknowledgements

This work was supported by grants obtained from the University of Padova (grantno. PRAT2010-CPDA103503) and the Italian Ministry of Education, University and Research(grant no. PRIN2010-M2JARJ), the Rientro dei Cervelli Program (lncentivazione alla mobilitàdi studiosi stranieri e italiani residenti all’estero) from the Italian Ministry of Education.We thank Daniele Ottaviani for reading the manuscript and for the useful discussion.

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Biophysical groundwork as a hinge to unravel the biology of α-synuclein aggregation and toxicity

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