Intrinsically disordered proteins (IDPs) studied by EPR and in-cell EPR

Intrinsically disordered proteins (IDPs) are a class of proteins lacking a stable three-dimensional structure in solution. Nonetheless, these proteins are native and fulfill many important biological functions, among them cell signaling, recognition and regulation.¹  IDPs are highly prevalent in humans: Genome analyses predict that around 25% of all human proteins are disordered from end to end, while up to 40% contain unstructured regions.²  Due to their abundance as well as their unique structural and dynamical flexibility, IDPs are key players in many biological pathways, capable of specific interactions with a multitude of binding partners and therefore often binding to or serving as hubs in protein interaction networks.^3,4  Due to the same reasons, many members of the IDP family are known to be associated with a variety of human diseases, among them prominently cancers, cardiovascular diseases, diabetes and neurodegenerative diseases.^5–7  All of this has made IDPs a research field of tremendous importance and interest since the turn of the century.

In the free energy landscape of proteins with a native globular fold there is a pronounced free energy minimum that stabilizes a distinct 3D fold, which often represents the unique functional form of a globular protein (Fig. 1A). The decrease in entropy associated with restrictions of the conformational freedom during folding is compensated by the formation of many intramolecular contacts.^8,9  However, the free energy landscape of an IDP looks distinctly different (Fig. 1B): It is characterized by the lack of a global free energy minimum, but shows many local minima instead, which are separated by small energy barriers that allow quick and frequent interconversion between the accessible states. Consequently, the conformational ensemble of an IDP in solution is heterogeneous and characterized by a dynamic exchange between many accessible structures. This non-folding behavior is encoded in the amino acid composition of IDPs: These proteins contain less hydrophobic (Ile, Leu, Val) and aromatic (Trp, Tyr, Phe) amino acid residues, but a significantly larger proportion of small and hydrophilic amino acid residues (Arg, Gly, Gln, Ser, Pro, Glu, Lys) and are richer in structure-breaking amino acid residues (Pro, Gly) than typical globular proteins.^7,8,10,11  Although IDPs cannot spontaneously fold into a compact globular structure, the presence of interaction partners can alter the IDP free energy landscape in a way that more pronounced energy minima appear: Upon interaction with a partner, IDPs often undergo a structural reorganization that defines a state of clearly reduced free energy, which is thermodynamically stabilized (Fig. 1B).

While some IDPs undergo a folding process as a whole upon interaction with a binding partner, more commonly specific recognition motifs in the disordered protein adopt secondary or tertiary structural elements upon interaction with a binding partner.^12–14  As a mechanism of these disorder-to-order transitions, two major models are discussed in the literature – the ‘conformational selection’ model and the ‘induced folding’ model.¹⁴  The first model is based on the assumption that the binding partner selects a specific conformation resembling the bound conformation from the wide ensemble of coexisting conformations the IDP adopts when free in solution (Fig. 1B). The latter model is based on the assumption that the IDP binds to its interaction partner in the fully disordered state and folds while bound to the partner, i.e., folding is induced by the partner. In reality, one or the other process may occur, or also some combination of the two models.¹⁴  The full dimension of the structural flexibility of IDPs becomes obvious in cases, where one recognition domain of an IDP can adopt various structural folds upon interaction with different binding partners. One prominent example is the C-terminal disordered region of tumor suppressor p53, which can adopt helical, β-strand or irregular structure upon interaction with different partners.^14,15  In contrast to a concise disorder-to-order transition, some IDPs may also stay largely disordered and still show fast interconversion between coexisting conformations while in functional complex with a binding partner. Such heterogeneous protein complexes are referred to as ‘fuzzy complexes’.^16–18 

Similarly to the formation of functional complexes with protein partners, a profound indentation in the free energy landscape of IDPs can be caused by formation of non-functional complexes of IDPs, like oligomers, amorphous aggregates and amyloid fibrils (Fig. 1B).⁸  Since IDPs are involved in many crucial cellular processes, e.g., signaling and regulation, misfolding of these proteins is often pathogenic.^5–7  While some IDPs may exhibit an intrinsic propensity for forming a pathologic conformation, others are misfolding due to external factors, e.g., due to disrupted chaperone-interaction, point mutations, interaction with other proteins, toxins and small molecules or impaired post-translational modifications.⁶  The largest group of misfolding diseases is characterized by the formation of stable, insoluble, highly organized filamentous protein aggregates known as amyloid fibrils, which accumulate in tissues or organs.^7,19,20  Prominent among these amyloidogenic diseases are neurodegenerative diseases, where filamentous protein deposits are accumulating in the patient's brain. Striking examples are the IDPs α-synuclein, which is found as aggregated inclusions in the Lewy bodies in the brain of Parkinson's disease (PD) patients,^21–23  and tau, which is a hallmark of Alzheimer's disease (AD) in its aggregated form.^24–26 

Their prevalence in humans, their importance in a multitude of biological processes as well as their significance for health and disease makes IDPs a highly relevant and important subject of research. However, their structural and dynamical flexibility as well as the versatility of their shapes and appearances described above makes research on IDPs a challenging matter. Traditionally, protein structures are elucidated using NMR spectroscopy or X-ray crystallography.^27,28  However, they are not straightforwardly applicable to obtain a conclusive analysis of the heterogeneous conformational ensemble that is characteristic for IDPs comprising a variety of quickly interconverting structures. Nonetheless, various methods of NMR spectroscopy, e.g., chemical shift dispersion, paramagnetic relaxation enhancement (PRE), and residual dipolar couplings (RDC), are widely applied for the investigation of IDPs.^29–31  However, NMR techniques are subjected to limitations when it comes to investigation of large protein complexes, e.g., with lipids or other binding partners. Thus, investigation of IDPs in their relevant macromolecular context using NMR has its boundaries. Many other biophysical methods have been employed to investigate IDPs, among them circular dichroism (CD) spectroscopy,³²  fluorescence resonance energy transfer (FRET),^33,34  and many more.^13,26  Since it is challenging to obtain meaningful experimental results on IDPs, computational approaches like prediction of intrinsic disorder or MD-simulations using experimental restraints, e.g., from PRE, are frequently used in IDP research.^13,35  Commonly, the macroscopic morphology of IDPs in the fibril state is judged by transmission electron microscopy (TEM).

A powerful experimental method that has become increasingly important in IDP research is electron paramagnetic resonance (EPR) spectroscopy. EPR is a versatile spectroscopy technique that allows us to study the dynamical and structural changes of proteins, in particular of IDPs.³⁶  As it can only detect unpaired electrons, EPR spectroscopy is virtually background-free. EPR spectroscopy allows measurements of proteins of any size in arbitrarily complex environments, e.g., in the presence of binding partners like proteins, lipids or small molecules, as well as under arbitrary environmental conditions and even in the cell.^37,38 

EPR spectroscopy is sensitive to unpaired electrons only. Thus, in order to perform EPR measurements with many proteins, a paramagnetic center has to be introduced as a spin label via site-directed spin labelling (SDSL).³⁹  Depending on the application, a suitable spin label has to be selected from a variety of possibilities. Some examples are shown in Fig. 2. For a recent review, the interested reader is referred to, e.g., Roser et al.⁴⁰ 

The spin labels most commonly used for EPR spectroscopy on proteins are nitroxide radicals. Among them, the methanethiosulfonate spin label (MTSL, see Fig. 2A), which is attached to a native or engineered cysteine residue in the protein via disulfide bonding, is most frequently used.⁴¹  However, in the reducing environment of a cell, both the N–O-moiety as well as the disulfide bond with the cysteine are susceptible to reduction.⁴¹  In order to ensure a stable attachment of the spin label to a cysteine, various linker-moieties have been developed, e.g., the maleimido moiety as used in 3-maleimido-proxyl (see Fig. 2B), which shows enhanced pH stability and is therefore often used for investigation of biological systems.^40,41  For cysteine-specific attachment of a spin label, all native cysteine residues in the protein, which are not to be labelled, need to be replaced by non-reactive amino acids (often Ala, Ser) prior to labelling. A different and much more elegant way of introducing a spin label into a protein for in-cell EPR is via genetically encoded spin labelling, where the spin label is introduced as an unnatural amino acid with bio-orthogonal reactivity during the biosynthesis of the protein in vivo, making the technique especially suited for in-cell EPR.^42,43  The corresponding unnatural amino acid SLK-1 is shown in Fig. 2C. When using this spin labelling approach, native cysteines present in the protein need not be substituted, implicating a more undisturbed protein than with cysteine-specific spin labelling. This also applies for a proposed spin labelling strategy based on click chemistry targeting unnatural amino acids.⁴⁴  Approaches for stabilizing the nitroxide radical itself against reduction include the sterical shielding of the unpaired electron.⁴⁵  A promising approach is the use of pyrrolidine-based nitroxides with ethyl groups slowing down the reduction process.^45,46  As, in general, the stability of nitroxides against reduction is limited, other spin labels have been developed in recent years that pave the way towards in-cell EPR: Recently, a family of Gd³⁺-based spin labels has been introduced, which show superior stability in the cellular environment and exhibit a high sensitivity for EPR spectroscopy.⁴⁷  Successful in cellula EPR distance measurements at Q- and W-band frequencies (34.5 and 95 GHz, respectively) have been reported with Gd³⁺-chelates like Gd³⁺-PyMTA⁴⁸  (see Fig. 2D) or Gd³⁺-DOTA-M.⁴⁹  Moreover, trityl radicals have been shown to be promising spin labels for in-cell EPR due to their favorable reduction characteristics.⁵⁰  A different approach to spin labelling is the coordination of Cu²⁺-ions by two strategically placed histidine residues. EPR distances determined between two Cu²⁺-ions have been shown to be very narrow and readily relatable to the protein backbone structure.^51,52 

In the investigation of spin-labelled protein oligomers, e.g., amyloid fibrils, several spin labels from various protein monomers might come into close contact. If contributions from inter-molecular spin–spin interactions are unwanted in the experiment, spin-labelled protein needs to be diluted with diamagnetic protein (Fig. 3).

Different methods of EPR spectroscopy allow accessing various types of information about the protein under investigation. There are two main experimental EPR techniques: On the one hand, in continuous-wave (cw) EPR spectroscopy the sample is subjected to continuous irradiation with microwave energy. Several excellent reviews describe the applications and developments in cw EPR for the investigation of proteins.^39,54–57  Typical cw EPR experiments are performed at X-band microwave frequencies (9 GHz) and deliver information about the following sample parameters: (i) the mobility of the spin label side chain, (ii) the polarity of the spin label microenvironment, (iii) the solvent accessibility of the spin label side chain and (iv) distances to other paramagnetic centers. In the corresponding cw EPR experiments a site scan, i.e., performing the EPR experiment with several samples, where the spin label is attached to different positions of interest in the protein, helps to create a comprehensive picture of a protein region.

For the determination of the spin label mobility (i), the cw EPR experiment is performed at ambient temperature, or particularly at a physiologically relevant temperature. Most commonly, information about the reorientational dynamics of a spin label side chain is extracted from the EPR spectra via spectral shape analysis using full spectral simulations including the determination of the rotational correlation time τ_corr using, e.g., the EasySpin software package.⁵⁸  Cw EPR line shapes in X-band typically resolve spin label dynamics on a 100 ps to 100 ns timescale, which reflect side chain motions, backbone dynamics as well as tertiary contacts.^39,59  Changes in the dynamics of a spin label side chain, e.g., due to interaction with a binding partner or conformational transitions of the protein can be detected with cw EPR spectroscopy: Even if the rate of conformational exchange is beyond the EPR timescale, the environment of the spin label is usually subjected to distinct changes, which allow the detection of distinct components in the EPR spectrum reflecting the different conformational states of the protein.³⁹  Often, such EPR spectra contain not only one but several contributions from a superposition of various conformational states of the protein conformational ensemble. Multi-component spectral simulations allow to monitor and to quantify fractions of multiple structural states that coexist or exchange with each other.⁶⁰ 

The polarity of the spin label microenvironment (ii) is reflected in the hyperfine-tensor component A_zz as well as the g-tensor component g_xx. While a polar environment shifts A_zz to higher values, g_xx is decreased. A_zz can easily be obtained from X-band cw EPR spectra recorded in frozen solution samples.⁶¹  In room-temperature spectra effects of micro-environmental polarity are reflected in the isotropic hyperfine splitting a_iso. The micro-environmental polarity can deliver information about the protein fold, secondary structure elements or embedding of the protein in a lipid layer.⁵⁴ 

The accessibility (iii) of the spin label side chain to paramagnetic quenchers can be monitored using cw EPR power saturation curves. While metal ion complexes like NiEDDA and CrOx report on the accessibility of the spin label side chain from the bulk water phase, molecular oxygen reports on the accessibility from a lipid phase.⁵⁶  In case of interaction of the protein with a lipid phase, accessibility information can be used to judge immersion of labelled protein sites into a lipid layer or exposure to the solvent.⁶²  Periodical patterns in the accessibility also may be interpreted in terms of secondary structural elements like an α-helix.

Cw EPR spectroscopy also gives access to (iv) spin–spin distances in the range of 8–25 Å.⁶³  This information manifests itself in spectral broadening due to magnetic dipolar interaction in EPR spectra recorded of frozen solution samples and can be extracted by deconvolution techniques. However, the distance range and thus the applicability of this technique is limited and pulsed EPR spectroscopy provides the means to access much larger distance ranges.

There is a variety of pulsed EPR spectroscopic methods available that provide access to complementary information to cw EPR methods. The most widely applied pulsed EPR technique in the context of proteins and IDPs is pulsed EPR distance measurements using double electron–electron resonance (DEER) spectroscopy, also known as PELDOR (pulsed electron double resonance), which has been reviewed in several excellent articles.^64–67  Spin–spin distances between 1.8 and 6 nm in membrane proteins and up to 10 nm in deuterated soluble proteins or even 16 nm in favorable cases are accessible with DEER.^65,68  The experiment is commonly performed at cryogenic temperatures (e.g., 50 K) in frozen glassy solution. Since the distance information is frequency-encoded, the result of a DEER experiment provides not only a mean distance, but a precise distance distribution, which allows to understand also non-homogeneous conformational ensembles of proteins and to follow structural transitions upon, e.g., changes in the environmental conditions or interaction with arbitrary binding partners.⁶⁹  As discussed above, IDPs feature a very broad conformational ensemble, which might even remain fuzzy in the presence of specific binding partners.^16–18  In these cases, DEER time traces suffer from a lack of clear dipolar signal modulations and thus, the standard DEER analysis procedures fail.^70,71  For such situations, a modulation depth-based data analysis procedure has been established with the IDP osteopontin by Kurzbach et al.: Based on the extraction of an effective modulation depth Δ_eff at a specific dipolar evolution time t it is possible to judge if the average spin–spin distance increases or decreases, since a lower effective modulation depth Δ_eff at the time t corresponds to a larger spin–spin distance (Fig. 4).^71,72  Thus, conclusions about transitions in the conformational ensemble of an IDP become possible, even though no clear disorder-to-order transition may take place.

The modulation depth of DEER traces is also the source of a different kind of information: Oligomerization of proteins can be monitored using EPR distance measurements with modulation depth calibration^73,74  or monitoring modulation depth vs. flip angle.⁷⁵  Thus, it becomes possible to extract useful information about protein–protein interactions.

Another pulsed EPR method called electron spin echo envelope modulation (ESEEM) can be used to measure the presence of NMR active nuclei in the environment of a spin label. In the protein context, this is a powerful approach to measure exposure of a protein site to solvent deuterium.

Ideally, protein structure, conformational transitions, protein function and protein interactions are monitored in the most relevant environment, i.e., inside the living cell. Macromolecular crowding as well as the presence of numerous potential interaction partners may significantly alter the behavior of proteins under investigation compared to an in vitro experimental setup. In general, EPR spectroscopy is suitable for observing proteins work inside the cell and recent research has focused more and more on the development of corresponding experimental protocols.³⁷  Gd³⁺-chelates in combination with microinjection of oocytes of Xenopus laevis^48,49  or hypotonic swelling of HeLa cells⁷⁶  proved to be well suited for intracellular EPR distance measurements. The introduction of MTSL or 3-carboxy-PROXYL into the cell in combination with the oxidizing agent K₃Fe(CN)₆ also resulted in EPR spectra and DEER traces of good quality.^77,78 

In this article we review the research that has been conducted in the field of IDPs using EPR spectroscopy in the recent years. In particular, we focus on two prominent examples of IDPs, the ‘Parkinson protein’ α-synuclein and tau, a key player in Alzheimer's disease. Both proteins have been subject of many spectroscopic and especially EPR studies. We will review the latest of these studies and discuss, which developments may become important for future EPR studies on IDPs.

In its physiological function, the protein tau binds and stabilizes microtubules and has little tendency for aggregation.⁷⁹  Under pathological conditions however, anomalous aggregates of tau are linked to various neurodegenerative diseases referred to as tauopathies.^80,81  This heterogeneous group of dementias and movement disorders also includes Alzheimer's disease (AD), hallmarked by neurofibrillary tangles (NFTs) of tau filaments that form deposits inside the diseased neurons.²⁵  While most cases of disease are sporadic, tau was also linked to inherited forms of frontotemporal dementia and Parkinsonism caused by mutations in the tau gene MAPT and involving abundant filamentous tau inclusions in the brain.^82,83 

The mechanisms of tau aggregation in vivo as well as the pathways underlying tau-related neurodegeneration are still enigmatic: Although the link between tau and several diseases is well-established, the question, whether NFTs are the primary neurotoxic tau species, is still a matter of much debate.⁸⁴  The correlation between the appearance of NFTs and disease progression in AD brains was taken as evidence that tau fibrils cause neurodegeneration. However, recent research points in the direction that soluble, pre-filament tau may constitute the most neurotoxic protein species.^84,85  Thus, in order to develop effective tau-directed therapeutic strategies against tau-based diseases, it is of prime importance to understand the pathomechanisms of tau-mediated neurodegeneration.⁸⁶ 

Human tau is encoded in the MAPT gene, which is expressed mainly in the axons of the central nervous system (CNS). By alternative splicing, six tau isoforms are generated, which differ by the presence or absence of two N-terminal inserts (N1, N2) and the presence of either three (3R) or four (4R) microtubule binding repeats in the C-terminal half of tau (Fig. 5A).^24,87  The longest tau isoform consists of 441 amino acids, and whenever residue numbers are given in this article, they refer to this isoform. The overall amino acid composition of tau is rather hydrophilic, which prevents tau from folding into a globular structure and makes it an IDP.^88,89  Tau can be subdivided into two major domains, the C-terminal assembly domain, which has a basic character, and the N-terminal projection domain, which contains a predominantly acidic stretch. Tau is overall basic, but is rather a dipole with two domains of opposite charge.^90,91  Despite its generally disordered character, tau was shown to adopt a broad conformational ensemble in solution featuring an overall ‘paperclip’ shape, where the C- and N-terminus approach each other and the microtubule binding region (MTBR) (Fig. 5B).^34,92 

Physiological tau is mainly found in neurons, where it binds to microtubules with the MTBR and flanking regions, while the N-terminal projection domain projects away from the microtubules (Fig. 5B).²⁴  By binding to several tubulin dimers tau stabilizes microtubules, thus regulating microtubule assembly and dynamics as well as axonal transport.^93–95 

Tau inclusions as found in AD patients’ brains consist of paired helical filaments (PHFs) and straight filaments (SFs) formed from all tau isoforms.^96–99  The protease-resistant core of tau filaments is established by the repeat region of tau which folds into β-sheet structures, while the N- and C-terminal regions of tau remain disordered and project away from the core as the so-called ‘fuzzy coat’.^100,101  Aggregation was shown to initiate from two short hexapeptide motifs PHF6* (²⁷⁵VQIINK²⁸⁰) and PHF6 (³⁰⁶VQIVYK³¹¹) located in repeats R2 and R3 of the MTBR, respectively.¹⁰²  Only recently, the structure of tau filaments extracted from AD brain was solved with atomic resolution by cryo-electron microscopy by Fitzpatrick and coworkers.⁹⁹  They showed that the filament core region is C-shaped and consists of intramolecular anti-parallel cross-β structures running perpendicular to the filament axis and that all six tau isoforms are arbitrarily stacked along the filament axis. Interestingly, the fibril core was shown to include repeats R3 and R4 of the MTBR.⁹⁹  For most studies concerned with the characterization of tau aggregates, filaments are not extracted from brain tissue, but generated in vitro. Tau aggregation can be induced by negatively charged cofactors that compensate the positive charge in the repeat region, like polyanions such as heparin and RNA, or fatty acids.^80,103–105  Most commonly, tau fibrils are prepared by the addition of heparin, which has been suggested to induce the formation of AD-like tau filaments.¹⁰⁴ 

Tau is subject to manifold post-translational modifications (PTMs). In particular, phosphorylation at up to 85 potential sites is crucial in regulating the physiological functions of tau, including microtubule binding.¹⁰⁶  Hyperphosphorylation reduces microtubule binding and is assumed to drive tau aggregation, since hyperphosphorylated tau from AD brains can self-assemble in vitro.^107,108  However, since phosphorylation alone does not seem to be sufficient for aggregation, the issue is still a matter of debate.⁸⁰ 

An important early contribution of EPR spectroscopy to tau research aimed for understanding the architecture of tau filaments along the fiber axis with SDSL cw EPR spectroscopy and line shape analysis using MTSL as a spin label. It was shown that β-strands, which assemble in the repeat region of individual tau molecules, are aligned in parallel with each other such that the same amino acids stack right on top of each other along the axis of a heparin-induced tau filament.¹⁰⁹  This was implied by very characteristic single-line EPR spectra found for aggregated tau with spin labels attached at positions 301–320 in the repeat R3 (Fig. 6). The loss of the two outer nitroxide hyperfine lines was confirmed to result from spin exchange due to orbital overlap of several spin labels, enabled by the stacking of the same amino acids from different tau molecules along the fibril axis.¹¹⁰  In contrast, EPR spectra recorded of tau, which was spin-labelled outside the filament core (positions 400–404) showed that this region remains structurally dynamic upon aggregation.

Independent support for the parallel in-register arrangement of β-strands was provided by quantitative analysis of dipolar broadening in cw EPR spectra of fibrils of singly spin-labelled tau and α-synuclein. The dipolar spin–spin interaction was converted into distance distributions, which are in agreement with the parallel in-register β-sheet arrangement.¹¹¹ 

The parallel in-register arrangement of β-strands in the filament core region is not limited to tau, but rather a shared structural characteristic of amyloid fibrils, that has been confirmed (also with methods of EPR spectroscopy) for many other proteins, including α-synuclein and prion proteins.^110,112 

Despite all similarities in the general architecture, amyloid fibrils can still appear in different conformations, even when formed from the same protein.¹¹³  While the overall arrangement of β-strands running perpendicular to the filament axis is maintained, a large variety is possible in core sizes and β-strand interactions. For example, the introduction of point mutations in repeat R2 of tau was shown to induce local structural changes by EPR spectroscopy, while the overall parallel in-register arrangement of β-strands in the fibril core was unchanged.¹¹⁴ 

Using diamagnetic dilution experiments and cw SDSL EPR it was shown that tau constructs of 3R tau and 4R tau co-assemble with each other into heterogeneous filaments with a core of parallel, in-register arranged β-strands from soluble tau.¹¹⁵  Implied in this finding is a huge number of possible tau fibril conformers: tau may either form 4R fibrils, or 3R fibrils, or 4R/3R heterogeneous fibrils with a variety of ratios between 4R and 3R tau. Different fibril morphologies of tau are disease-specific: while AD filaments contain 4R and 3R tau, most diseases are characterized by 4R or 3R tau fibrils only.^82,116 

In the prion concept, different filament strains characterized by structural polymorphism are thought to be responsible for phenotypic diversity in disease. They spread by mechanisms of intercellular propagation and templated-assisted conversion of monomeric protein onto the filament seeds, while the strain-specific properties are maintained (Fig. 7).²⁵ 

In the past years, evidence was provided that tau has prion-like properties: Fibril seeds were shown to have the ability to propagate between cells to distant brain regions and assemble wild-type tau into filaments.^117,118  Moreover, distinct tau strains were shown to stably reproduce their conformation during propagation with high fidelity defining different tauopathies.^82,119 

Tau filaments consisting of 3R and 4R tau are characterized by an asymmetric seeding barrier, i.e., 3R tau filaments can recruit 4R tau, while 4R tau filaments cannot recruit 3R tau.¹²⁰  EPR distance measurements using DEER in combination with SDSL and extensive molecular dynamics simulations were used in order to investigate the molecular basis of the seeding barrier.¹²¹  Monomers of 3R and 4R tau fragments containing only the repeat region (known as K19 and K18 tau, respectively) were grown onto seeds formed from 3R or 4R tau and intramolecular distances were determined between spin labels located in repeat R3 of the fibril core. 4R tau grown onto 4R seeds resulted in distance distributions with several contributions, indicating heterogeneous fibril conformers with at least three distinct conformers. In contrast, when 4R tau and 3R tau were grown onto 3R tau seeds, homogeneous fibrils were indicated by narrow distance distributions. The results suggest conformational variations in R3 of 3R or 4R tau filaments. Moreover, they show a structural plasticity of tau as the initial seeds imprint their conformation onto the recruited tau monomers as evident in the case of 4R monomers which grow into different fibril conformers on different seeds.¹²¹  The emergence of structural heterogeneity as well as the conformation-based seeding barrier together with the template-based conversion of monomers are important characteristics of prion-like behavior.¹²¹ 

The influence of single point mutations on the populations of fibril conformers of 4R tau filaments (prepared with truncated K18 tau) was further investigated by DEER spectroscopy.¹²²  Structurally heterogeneous 4R tau seeds were used as initial templates for filament assembly of mutated tau monomers carrying spin labels at positions 311 and 328. Several tau mutations were tested, among them some of well-known relation to diseases (ΔK280,¹²³  P301S,¹²⁴  S320F,¹²⁵  Q336R¹²⁶ ). While several of the mutated tau constructs reproduced the intramolecular distance distribution and thus the ratio of fibril conformers of wild-type tau (Fig. 8B), some mutated tau derivatives, in particular some of the disease-related ones, induced large shifts in the populations of fibril conformers (Fig. 8C). As the fibrils are generated by template-assisted growth onto the tau seeds, the proposed mechanism responsible for the change in the fibril conformer populations is seed selection due to changed conformational compatibilities of fibrils formed from point-mutated tau: mutants showing no interference with any of the original conformers will grow onto the wild-type seeds conserving the conformational ensemble. Mutants with incompatibilities to some of the original conformers will change the overall composition of the ensemble, possibly amplifying minor subpopulations (Fig. 8D). The mechanism of seed selection as shown in this study in vitro could provide a plausible explanation of how hereditary mutations might change the initial ensemble of tau conformational composition and lead to the emergence of changed fibril ensembles.

The influence of a change in incubation conditions during fibril growth on the conformational ensemble of 3R and 4R tau filaments (prepared with truncated K19 and K18 tau, respectively) was also assessed by means of intramolecular DEER spectroscopy using the spin label positions 311/322 (in 3R tau) and 311/328 (in 4R tau) for determination of intramolecular distances.¹²⁷  A multicycle seeding scheme was used in order to test the relative stability of individual fibril conformers. While fibrils formed from 3R tau were basically unaltered over 15 cycles of seeding, 4R tau fibril populations changed during consecutive cycles of seeding. Homogeneous 4R fibrils were dominantly formed upon stirring in the first cycle of growth, quiescent growth in the following cycles led to formation of a heterogeneous ensemble of fibril conformers. This change in the fibril structural ensemble is suggested to be a result of the alteration of growth conditions from stirring to quiescent, which could lead to selective amplification of distinct conformers by a mechanism related to the fragilities and growth rates of different fibril conformers: under stirring conditions, fibril conformers with increased fragilities (but decreased growth rates) might have a selective advantage over more stable (faster growing) species due to the production of many new fibril ends. When removing the selective pressure (i.e., stirring), the population ensemble evolves towards the latter conformation. The authors show that fibril conformer populations depend on the subtle balance between fracture and growth.¹²⁷ 

Only very recently, Fichou and coworkers questioned the idea that heparin-induced tau filaments are similar to the PHFs and SFs extracted from AD brains: They performed a study comparing intramolecular DEER distance distributions of heparin-induced tau filaments with corresponding simulations based on the AD-filament structure of tau published by Fitzpatrick et al.^99,128  They used a truncated version of tau lacking the N-terminal part, but containing all repeats and the C-terminus. As a reference they also measured DEER distances of tau in solution and compared those distances with the corresponding simulations based on a tau solution structural ensemble published previously.¹²⁹  While Fichou and coworkers find that the DEER distance distributions recorded of tau in solution are in adequate agreement with the published ensemble, larger discrepancies between experiment and simulations appeared in the case of heparin-induced tau filament samples. For all tested doubly spin-labelled tau derivatives they found distance distributions considerably broader than expected from the simulations, indicating that heterogeneous tau fibril conformers are present in heparin-induced tau fibrils, instead of only PHFs and SFs as found in AD-filaments. Moreover, the average inter-spin distances were larger than predicted, suggesting that the key features of AD-filaments, like the C-shape and the cross-β structure, are not the dominant conformations in heparin-induced tau fibrils.¹²⁸  Upon aggregation with heparin a stretching of intramolecular tau inter-spin distances in repeats R1, R2 and R3 is observed in the distance distributions, consistent with the formation of a cross-β structure. With ESEEM experiments of doubly spin-labelled tau variants the authors determined a homogeneous reduction in the accessibility to deuterium atoms of the solvent across all four repeats of the tau MTBR, suggesting that all four repeats are part of the tightly packed heparin-induced filament. This is in contrast to the AD-filaments, where only R3 and R4 are part of the core region.⁹⁹  The authors suggest that heparin, which is a fairly extended disordered chain, cannot serve as template for uniform tau aggregates, but rather generates polymorphic tau fibrils.¹³⁰  However, since tau strains have been shown to be disease-specific, for the meaningful biological analysis of specific tau strains in terms of diseases it might be necessary to employ tau filaments seeded from structurally defined templates, instead of polymorphs resulting from heparin-induced tau aggregation.¹¹⁹ 

In the past years some EPR studies have focused on the investigation of very early events in the tau aggregation pathway, monitoring tau conformational states and populations in solution and during earliest stages of aggregation into insoluble fibrils. As there is evidence that primary tau toxicity might be exerted by soluble oligomers formed in the early stages of fibril formation, knowledge about their structure and properties is important for developing tau-based therapeutics.

A tau fragment containing 13 residues of the R2 region, including hexapeptide PHF6*, was used to test the hypothesis that subtle changes in the solution conditions, such as changes in buffer type or presence of osmolytes (TMAO vs. urea), influence the aggregation pathways and end products.¹³¹  A cysteine-substitution allowed spin labelling at the N-terminal end of the fragment and thus cw EPR measurements in order to access solvent accessibility, packing and mobility of the spin label, as well as the fractions of mobile and fibrillated tau, before and after aggregation initiation. The EPR results together with Overhauser dynamic nuclear polarization (ODNP)-derived surface water diffusivity, ion mobility mass spectrometry (IM-MS), transmission electron microscopy (TEM) and thioflavin T (ThT) fluorescence show that both urea and TMAO strongly influence oligomer formation. While urea breaks apart inter-tau contacts, shifting the tau population towards monomers, and hinders downstream inter-tau fibrillation also in the presence of heparin, TMAO facilitates tau oligomerization already in the absence of heparin. Also the change of buffer type, which is often considered a minor factor in sample preparation, was shown to influence the conformational ensemble of the tau peptide and shifted it to more aggregation-prone populations in comparison to water, which in turn influenced the downstream aggregation of tau peptides after addition of heparin.¹³¹  Taken together, the results show that the solution state of tau before initiation of aggregation influences the whole pathway of protein aggregation and the propensity to form fibrils.

Using a truncated tau version that comprises all four repeats and the C-terminal region, structural transitions of tau in the early stages of aggregation in solution were probed by ODNP as well as cw EPR line shape analysis as a function of aggregation time.¹³² 

Several key positions across R3 and the C-terminus were spin-labelled one at a time for this purpose. Multi-component EPR spectra at different time points after initiation of aggregation consist of (i) a mobile, solvent-exposed component corresponding to monomers or small oligomers of tau, (ii) an immobile component, corresponding to tau immobilized at an inter-tau interface, and in the case of not spin-diluted samples (iii) a spin-exchanged single-line component (Fig. 9A,B). The results of this study suggest that as soon as 5 min after initiation of aggregation 40–70% of tau variants carrying a spin label in the R3 region are involved in dynamic and partially structured aggregation intermediates, reflected by immobilization of the spin label side chains. Approximately 5% of all R3 spin-labelled tau are already involved in β-sheet arrangements (Fig. 9C).

Importantly, over time, only the fractions of embedded tau change, while spectral simulations show that the EPR parameters of the individual components remain unchanged. At this early time in the aggregation process, no fibrillary assemblies are observable by cryo-TEM (Fig. 9D). This indicates that a major tau population of partially structured intermediates forms within minutes after aggregation initiation, which are on-pathway species towards formation of mature fibrils long before actual fibrils are detected in the sample. After maturation of the fibrils the β-sheet content in the R3 region plateaus at 70–80%. In contrast, the C-terminal spin-labelled residues show no spin-exchanged component also after >24 h incubation with heparin, indicating the persistence of disorder in this protein region. Furthermore, the authors find that, once formed, the β-sheet structures do not exchange subunits but form stable entities already in stages of aggregation as early as 5 min.

DEER spectroscopy was also used to detect how aggregation initiation acts on the conformational ensemble of soluble tau in a time-dependent manner by monitoring the end-to-end distances of the key fibril-forming hexarepeats PHF6 and PHF6* in freeze-quenched tau samples.¹³³  Eschmann and coworkers analyzed the distances flanking the PHF6 and PHF6* segments in truncated tau fragments containing all four repeats and the C-terminus in their solution state as well as after addition of heparin. In all cases they observe a relatively compact conformational ensemble of tau in solution characterized by an average distance of just above 3 nm (Fig. 10). Upon initiation of aggregation they observe a stretching of the distances spanning both hexarepeats consistent with a conformational extension of these regions to just above 4 nm. Time-resolved DEER experiments show that all distance distributions recorded within 12 h after heparin addition can be described by a 2-state model of conformers, i.e., by a compact and an extended conformation, without the need for intermediates. The compact conformation is attributed to the stable solution ensemble of tau conformers, while the extended conformation is characteristic for a stretched out conformation as can be found in β-sheets of mature tau fibrils. Importantly, already 1 min after addition of heparin roughly 50% of all tau has undergone stretching (Fig. 10D), while 10 min after heparin addition roughly 90% are in the extended conformation (Fig. 10E), which is still a solution state consisting predominantly of monomer and dimer species well before the macroscopic appearance of fibrils in the sample. After 1 h the transition in the conformational ensemble towards the compact state is complete. In conclusion, upon initiation of aggregation the population of tau shifts towards an extended state which reflects conformers on the pathway to fibrils, well before β-sheet signatures or fibrils are detectable.¹³³  However, it is not clear whether the formation of the extended β-sheet-like conformation precedes the formation of dynamic aggregation intermediates as suggested by Pavlova and coworkers, or vice versa.¹³²  The results of this study suggest that tau aggregation occurs according to the nucleation conformational transformation model, in which a conformational transformation from a stable tau solution structure to a distinct aggregation-prone structure, as monitored here with EPR spectroscopy, is required to induce the stacking of proteins into fibrils.^134,135 

In addition to the polyanion heparin, RNA can also induce the aggregation of tau protein.^105,136  RNA is a component of every cell, rendering it a highly relevant interaction partner for tau. RNA was shown to drive fibril aggregation and tau fibrils that aggregated in the presence of RNA were shown to adopt a parallel, in-register β-sheet alignment along the fibril axis with characteristic exchange-broadened single-line cw EPR spectra, suggesting that the general fibril architecture is independent of the cofactor used for aggregation induction.¹³⁷  Also the asymmetric seeding barrier, where 3R tau cannot grow onto 4R tau seeds was shown to be preserved when inducing aggregation with RNA.¹³⁷  While RNA was found to bind to the tau fibrils formed, heparin replaces RNA on the fibril surface when added to the sample.¹³⁷ 

In a recent study, tau was shown to undergo liquid–liquid phase separation (LLPS) in the presence of RNA, forming droplets of high protein density.¹³⁸  Such protein-rich structures are known as complex coacervates, which are typically formed by two oppositely charged polyelectrolytes in solution, in this case of the polycation tau and the polyanion RNA.^139,140  The formation of the droplets is shown to be tunable by salt concentration, tau:RNA ratios and temperature. Despite the molecular crowding inside the droplet, tau was shown to maintain native-like mobility and a locally compact conformation around the PHF6⁽*⁾ regions as is typical for the tau conformation in dilute solution.¹³⁸  Thus, a change between the solution state and the coacervate state of tau is reversible and tunable by physiologically viable parameters.¹³⁸  After prolonged residence in the complex coacervate phase, a low-level β-sheet formation was observed.

Lipid membranes have been shown to enhance tau aggregation in vitro.^141–143  It is known that tau interacts with lipid micelles via its MTBR region under the formation of helical structures localized in R1, R3 and R4 of a truncated 3R tau construct containing only the repeat region (K19).¹⁴⁴  With SDSL of tau and mobility as well as accessibility analysis of cw EPR spectra, the interaction of tau with POPC/POPS vesicles was confirmed and a helical nature of the respective regions in tau was found in the bound state.¹⁴⁵  Cw EPR data also imply that the tau amphipathic helical segments interact with the lipid layer periphery and are not deeply immersed into the lipid layer. With DEER distance measurements, distances between the individual helical segments and between the helices and the linker regions were determined. The results suggest that individual helix segments are connected by highly flexible linker regions that allow adaption of the relative orientation of the helices to the spatial constraints posed by the lipid structure.¹⁴⁵  The induction of helical structure upon membrane binding has also been shown for other IDPs, e.g., for α-synuclein, and implications of helical segments in the context of protein aggregation are discussed.¹⁴⁶  A more detailed discussion of IDP-membrane interactions can be found in the α-synuclein section of this chapter.

The human α-synuclein (α-syn) consists of 140 amino acids and is highly abundant in the human brain. Its physiological function is not fully understood, but it is enriched in presynaptic nerve terminals and implicated in neurotransmitter release and vesicle trafficking.^147–149  α-syn consists of (i) an N-terminal amphipathic region, (ii) a central hydrophobic non-amyloid-β component (NAC) region, and (iii) a highly acidic C-terminal region (Fig. 11). α-syn displays remarkable structural versatility: it is intrinsically disordered but can readily adopt various conformations, e.g., β-sheet structure in aggregates or α-helical structure when bound to lipids.^150–153 

α-syn is known to pathologically self-assemble into amyloid fibrils and plaques, which are found in Lewy bodies, the pathological hallmark of PD, but it is also associated with other neurodegenerative diseases.²⁵  Like tau fibrils, α-syn fibrils are arranged in a cross-β conformation with β-sheets running perpendicular to the fibril axis. The fibril core ranges approximately from residues 30–100 as indicated by EPR experiments.^111,155  As for other amyloid fibrils, a parallel in-register arrangement of α-syn along the fiber axis has been revealed with cw EPR spectroscopy as described in detail for tau.¹¹¹ 

Like for other amyloid proteins and tau, α-syn has been shown to have prion-like properties, which allow α-syn-seeds to induce aggregation of endogenous α-syn to form aggregates.¹⁵⁶  α-syn was shown to generate polymorphic fibril structures, and two α-syn fibril strains have been shown to exhibit cell-to-cell transmission and to induce distinct pathologies.¹⁵⁷ 

Various point mutations of α-syn (e.g., A30P, A53T, or E46K) in the N-terminal membrane recognition domain are associated with early-onset Parkinson's disease,^158–160  indicating the importance of membrane binding for the function of α-syn.¹⁵⁴  The N-terminus of α-syn contains seven copies of an 11-residue pseudorepeat, which mediates binding to negatively charged lipids.^161,162  Upon membrane binding, α-syn undergoes a structural transformation towards an α-helix,¹⁶³  which was found by EPR spectroscopy and NMR spectroscopy to be continuous and extended on the surface of unilamellar vesicles¹⁶⁴  or to adopt a horseshoe-like conformation of two helices connected by a loop region on SDS micelles,¹⁶⁵  while the C-terminal region remains unstructured.

As discussed above for tau, in general for amyloid diseases there is a lot of evidence that protein aggregation is the cause for neurodegeneration. Recent results suggest, that the primary neurotoxic species may not be the mature fibril, but prefibrillar oligomers or protofibrils.¹⁶⁶  There is evidence that amyloid toxicity may be caused by membrane permeabilization by pore-like early intermediates as shown for α-syn.¹⁶⁷  The molecular basis of PD appears to be tightly coupled to the structural diversity of α-syn.¹⁶⁸ 

The intrinsically disordered nature of the protein contravenes the typical approach of standard high-resolution structural biology methods. EPR spectroscopy in combination with SDSL is a complementary and valuable tool for unraveling the structural ensemble of α-syn. Since human α-syn does not contain any cysteine residues, most commonly site-directed cysteine mutagenesis and cysteine-reactive radicals are used for SDSL. The usual procedure to check whether the spin labelling does affect the functionality of the protein is not applicable, because the function of α-syn is still unknown. Therefore, typical control experiments are CD spectroscopy or measurements of the aggregation kinetics before and after spin labelling.¹⁶⁹ 

Apart from classical spin labelling EPR studies, there are EPR investigations on α-syn, which exploit the fact that α-syn binds Cu²⁺ ions at several positions along the protein: the paramagnetic Cu²⁺ and the resulting EPR signal was employed to analyze the copper-α-syn interaction in various studies.^{168,170–173}  In conclusion, the C-terminal part of α-syn displays low affinity vs. copper, while the N-terminus binds copper with a high affinity of about 0.1 nM with His50 coordinating the bound copper. Dudzik et al. studied the coordination of copper by membrane-bound α-syn with EPR.¹⁷⁴  In this study, copper was found to bind exclusively to the N-terminus (Met1-Asp2, 1eq.) and did not alter the α-helical membrane-bound conformation of α-syn. Drew investigated the Cu²⁺/α-syn binding in solution and found that the dominant copper coordination mode of α-syn is associated with the formation of α-syn oligomers, in which Cu²⁺ ions occupy intermolecular bridging sites of α-syn dimers and trimers.¹⁷⁵ 

While early oligomers of α-syn are considered to be neurotoxic,¹⁷⁶  fibrils might play a role in spreading the disease from peripheral to central neurons.¹⁷⁷  High-resolution information is mainly available for several short fibril-forming peptides,^178–181  while fibrillary structures for longer proteins are often less understood.^110,182  Structural models of α-syn fibrils were proposed¹⁸³  based on solid-state NMR and cryo-electron microscopic data. However, long-range distance constraints were missing before the first high-resolution structure of full length α-syn fibrils was published (PDB entry 2N0A).¹⁵⁶  Pornsuwan et al. successfully detected several intramolecular long-range distance constrains at the molecular level on the fold of α-syn in fibrils by EPR distance measurements.¹⁸⁴ 

In order to infer the side-chain direction of the spin label within one strand and to conclude on the spatial arrangement of the labels and the strands, they exploited a pair-labelling strategy, i.e., measuring two distances from one labelled residue to two adjacent ones, respectively (Fig. 12). Based on the obtained distance constraints, the authors refrained from proposing a structural model and only concluded that the two inner strands containing residues 54 and 64 might arrange in a more complex fashion than proposed before. Even though a high-resolution structure of full length α-syn is available, polymorphism and coexisting different fibril types need to be taken into account in future studies.^{157,185–187} 

It is likely, that the physiological function of α-syn, e.g., as protein hub, is connected to α-syn's membrane binding affinity.¹⁸⁸  Additionally, it has been observed that the presence of lipid membranes influences the fibrillation kinetics.¹⁸⁹  Depending on the protein : lipid ratio, aggregation is enhanced or reduced. Therefore, α-syn-membrane interactions are studied intensively.

Many scientists introduce their conference talks on α-syn by presenting the high-resolution NMR structure 1XQ8 of monomeric, micelle-bound α-syn (Fig. 13B) (this often happens even when lipids are not involved at all in the presented research). The 1XQ8 structure was obtained by solution NMR, which is restricted in the size of the complex under study, so that small SDS micelles as membrane mimics were used. NMR structure determination revealed, besides the unbound C-terminal tail, an α-helical region featuring a break, resulting in two anti-parallel α-helices within the N-terminal region, called ‘horseshoe’.¹⁶⁵  This model was confirmed by 13 distance constraints obtained by EPR distance measurements between spin-labelled cysteines.¹⁹⁰ 

SDS micelles, however, might be too small to mimic actual organelle membranes. They have typical diameters of 5 nm and therefore may have artificially constrained the protein into the horseshoe structure. Here, the advantage of EPR distance measurements not being restricted in the size of the complex under study, can be exploited. Subsequent EPR studies were performed using small unilamellar vesicles (SUVs), which can accommodate α-syn in an extended alpha-helical conformation. However, it was found by control experiments using dynamic light scattering, that SUVs are eventually disrupted upon interaction with α-syn, depending on the protein:lipid ratio.¹⁹¹ 

Large unilamellar vesicles turned out to be stable up to protein : lipid ratios of at least 1 : 200 and were used for subsequent EPR distance measurements unravelling the membrane bound conformation of α-syn. Quantitative binding is found at least for residues 9–69. The helix conformation was found as a mixture of one single, extended helix as well as a horseshoe-like conformation of two helixes with a loop region in between (Fig. 13).¹⁹² 

Kumar et al. studied the α-syn conformation bound to SUVs mimicking the inner mitochondrial and the neuronal plasma membrane.¹⁹³  They conclude, that their experimental distance constraints obtained by DEER suggest a coexistence of two distinct conformations: firstly, an extended helical conformation and secondly, a broken helix with a larger opening angle between the two helices than the horseshoe-conformation found before on SDS-micelles as well as POPG SUVs and LUVs. This is another hint of the structural variability of α-syn when interacting with membranes.

Besides EPR distance measurements, the rotational dynamics at physiological temperatures of spin labels attached at different sites of α-syn can be obtained by EPR spectral shape analysis and can be used to monitor the local degree of membrane binding in the proximity of the labelled site, e.g., as a function of the composition of artificial membranes. The rotational mobility mainly reflects the residual mobility of the spin label. Upon interaction with macromolecular partners, e.g., membranes, reduced mobility is observed. Often, spectral simulations taking two components with different spin label mobilities are used for data analysis. The slow component can be attributed to binding of α-syn to the membrane in the vicinity of the labelled residue, so that the corresponding fraction reflects the local degree of binding of this region of the protein. This approach gives a more differentiated view of the α-syn-membrane interaction than a global binding affinity measurement.

Using this approach, Robotta et al. showed that binding of α-syn to membranes is initiated in the N-terminal region.¹⁹⁴  At lower surface charge densities, i.e., for membrane compositions with less negatively charged lipids, the binding affinity of regions close to the N-terminus is stronger than that of regions distal from the N-terminus in sequence. This implies, that different binding modes exist for α-syn, i.e., membrane binding involving different stretches of residues (e.g., binding with residues ∼9–69 or ∼9–27). Besides the coexistence of different helical conformations, this underlines the extreme conformational flexibility of α-syn.

Using a similar approach, Kumar et al. focused on the influence of phosphorylation at positions 87 and 129 on the membrane binding behavior of α-syn.¹⁹⁵  The serine residues at positions 87 and 129 were exchanged by an aspartic acid one at a time, to mimic phosphorylation at the respective position. Alanine was used as non-phosphorylated reference. The ‘phosphorylation’ at position 87 negatively affects the membrane binding behavior of all investigated labelling positions (27, 56, 63, 69, 76, 90), whereas ‘phosphorylation’ at position 129 has no effect on the binding affinity of α-syn. Also the influence of point mutations A30P and A53T on the membrane binding behavior of α-syn was investigated with EPR spectroscopy.³²  Robotta et al. studied ‘wild-type’ α-syn (nevertheless containing spin-labelled cysteines) as well as A30P and A53T mutants labelled at nine different positions (9, 18, 27, 35, 41, 56, 69, 90 and 140) in the presence of LUVs with systematically varied lipid compositions. Their results show that the disease mutation A30P does not only reduce the overall binding affinity of α-syn to LUVs, but is structurally defective and locally affects membrane binding.

It is worth mentioning that studies on point mutations using SDSL EPR require (additionally to the point mutation of interested) site-directed mutagenesis and labelling, which also might influence structure and dynamics. Therefore, it is important to directly compare the results of the SDSL-study with and without the point mutation of interest.

The α-syn-membrane interaction can also be observed from the perspective of the membrane by using spin-labelled lipids as spin probes instead of spin-labelled α-syn. Following this approach, Pantusa et al. focussed on the influence of α-syn and the disease mutants A30P, A53T and E46K on the lipids of vesicles.¹⁹⁶  They used anionic dimyristoylphophatidylglycerol (DMPG) LUVs containing 1 mol% n-PCSL (phosphatidylcholine lipids) with a nitroxide reporter group attached at selected carbon atom position. The spectra reflect the lipid chain order and the membrane dynamics. Their findings suggest that α-syn binds to the membrane without deep penetration into the membrane. A30P showed the smallest influence on the membrane, whereas E46K shows similar influence as wild-type α-syn.

The fact that SDSL EPR spectroscopy is virtually background-free, enables studying not only α-syn interactions with artificial membranes, like phospholipid vesicles, but also with isolated organelles, e.g., mitochondria. DEER distance measurements on α-syn were performed in the presence of mitochondria isolated from human HEK 293 cells.¹⁹⁷  The DEER data revealed a distance distribution in the presence of mitochondria, which can be described as a superposition of α-helical α-syn, i.e., bound to artificial membranes and intrinsically disordered α-syn in solution. The relative fractions of both components nicely agree with the bound and unbound fractions determined by cw EPR. This result demonstrates how α-syn binds onto isolated mitochondria in α-helical conformation. The binding profiles for two different artificial membrane compositions mimicking the inner and outer mitochondrial membrane revealed the binding of α-syn solely to the vesicles mimicking the inner mitochondrial membrane (IMM), which has a higher cardiolipin content than the outer membrane. Taken together, the results suggest that α-syn binds α-helically to the inner mitochondrial membrane.

Increasing the level of complexity, spin label EPR spectroscopy can even be performed in the cell. In order to study the intracellular conformation of α-syn, Cattani et al. micro-injected singly spin-labelled, initially monomeric α-syn, including samples with the disease mutants A30P and A53T, into oocytes of Xenopus leavis as model cells. Using intermolecular in-cell DEER experiments intracellular diffusion of α-syn upon microinjection was monitored. To study the (dis-)order of α-syn in the cell, in-cell cw EPR at room temperature was performed.¹⁶⁹  Importantly, it was shown by an independent control experiment that the intracellular micro-viscosity is very similar to aqueous solution. The α-syn in-cell study found no spectral changes compared to spectra measured in aqueous, buffered solution. However, the obtained signal-to-noise ratio, which is limited by the intracellular spin label stability does not allow to exclude a small fraction (up to 20%) of either α-helical or aggregated α-syn in the oocytes. Additionally, there was no difference detectable between the wild-type α-syn and the disease mutations A30P and A53T. The Goldfarb group were the first to perform distance measurements of N-terminally acetylated α-syn doubly spin-labelled with Gd³⁺-DOTA at positions 24/122 and at positions 42/122, which was introduced into A2780 cells via electroporation.¹⁹⁸  The obtained distances revealed the intracellular preservation of the disordered structure of α-syn as observed in buffer. However, again it was not possible to exclude a small fraction (up to 20%) of either α-helical or aggregated α-syn in the cell. An intriguing observation in this study was that α-syn seems to interact with some unidentified molecules in cellula as suggested by a drop in NMR signals across the protein.¹⁹⁸  This highlights, how the presence of all kinds of possible interaction partners may influence the structure and function of a protein inside a cell.

Many dysfunctional pathways and contributing factors have been implicated in IDP neurotoxicity, among them hyperphosphorylation, oligomerization, fibrillation, propagation and strain differences.^86,199  In order to develop mechanism-based therapeutics against IDP-related diseases, we need to understand the molecular mechanisms underlying their non-functional self-aggregation and characterize toxic and native IDP species as well as their endogenous cofactors regulating relevant structural transformations. EPR spectroscopy has been shown to be a powerful tool for the investigation of IDPs. It was used, e.g., for the elucidation of fibril architecture and fibril polymorphs, the processes of fibril formation, as well as for the analysis of the conformational ensemble during interaction with lipid phases. By delivering a wealth of information, EPR spectroscopy has become indispensable in IDP research.

Nevertheless, a careful analysis of the studies summarized herein reveals some limitations of the EPR investigations performed. First, usually EPR sensitivity is adequate for studying non-aggregated IDPs at physiological concentrations (some 10 µM). However, many studies on IDP fibrils are conducted with diamagnetic diluted samples. While diamagnetic dilution allows for measuring only intramolecular contributions, it requires very high protein concentrations (e.g., 800 µM)¹³³  in order to reach a sufficient signal-to-noise ratio in the common EPR experiment. Such high protein concentrations are usually non-physiological and thus may promote a non-physiological shift in the conformation equilibrium of the soluble protein or influence its self-aggregation propensity. High-sensitivity EPR spectroscopy might be the key to reducing molar sample concentrations to the physiologically relevant regime even for diamagnetically diluted samples.

Second, many EPR studies have not been performed with the full-length protein, but with shorter fragments thereof. This is advantageous since it results in greatly accelerated aggregation kinetics,²⁰⁰  while fibril properties are still ‘full-length-like’^115,201  and it is a valuable approach for understanding the molecular mechanisms of certain processes. However, since studies have shown that the populations of IDP conformational ensembles and aggregation depend on subtle variations in the sample conditions,¹³¹  even small effects accompanying protein truncation might influence the research results downstream, in particular their interpretation in terms of biological processes.

Furthermore, IDPs are highly regulated proteins, which are subject to numerous PTMs, e.g., phosphorylation, acetylation, ubiquitination or glycosylation, influencing the IDP energy landscapes.²⁰²  Many of these PTMs have been discussed to influence the aggregation behavior of IDPs and thus play important roles in IDP pathology.^80,203–205  In vitro experiments are often performed in the absence of PTMs or they investigate the influence of specific modifications. However, it is not clear, if such isolated modifications resemble the physiologically relevant state of an IDP.

In order to produce biologically meaningful results in the context of human diseases, it is crucial to further develop in-cell EPR in order to perform EPR experiments with IDPs under the most relevant environmental conditions, i.e., inside the cell. The cellular environment is characterized by molecular crowding and features a huge variety of interaction partners that may modulate the protein structural ensemble and processes of structural reorganization, as well as the oligomerization behavior, resulting fibril conformers and the interaction kinetics with partners like other proteins.²⁰⁶  As a result, in cellula experiments may deliver different findings than the in vitro complement.

However, the application of in-cell EPR is still demanding. Apart from the need for spin labels resistant to the reducing intracellular environment, effective ways of inserting the protein of interest into the cell also need to be developed. This might be done either by employing transfection techniques like electroporation or by the introduction of alternative in-cell spin labelling strategies, e.g., via unnatural amino acids that allow in vivo expression of spin-labelled protein.^42–44 

Recent studies showed that the signal-to-noise ratio in in-cell EPR experiments is crucial for the detection of all relevant protein subpopulations: Cattani et al. estimated that up to ∼20% of a dynamically restricted protein conformation could be present in samples of α-syn in Xenopus leavis oocytes without being detected in the EPR spectra.¹⁶⁹  This finding emphasizes the need for suitable labelling strategies as well as for high-sensitivity EPR spectroscopy. Promising spectroscopic approaches for high-sensitivity EPR distance measurements are laser-induced magnetic dipole spectroscopy (Laser-IMD),²⁰⁷  as well as the emerging applications of shaped pulses for DEER spectroscopy, which allow sensitivity enhancement by increasing the excitation bandwidth as well as improving spin dynamics control.^208,209 

With the development of new spectroscopic techniques, EPR has the potential to further make significant contributions to answering the still-open questions concerning structure and function of intrinsically disordered proteins.