Structural mapping of oligomeric intermediates in an amyloid assembly pathway

Abstract
Introduction
Results
Discussion
Materials and methods
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Abstract

Transient oligomers are commonly formed in the early stages of amyloid assembly. Determining the structure(s) of these species and defining their role(s) in assembly is key to devising new routes to control disease. Here, using a combination of chemical kinetics, NMR spectroscopy and other biophysical methods, we identify and structurally characterize the oligomers required for amyloid assembly of the protein ΔN6, a truncation variant of human β₂-microglobulin (β₂m) found in amyloid deposits in the joints of patients with dialysis-related amyloidosis. The results reveal an assembly pathway which is initiated by the formation of head-to-head non-toxic dimers and hexamers en route to amyloid fibrils. Comparison with inhibitory dimers shows that precise subunit organization determines amyloid assembly, while dynamics in the C-terminal strand hint to the initiation of cross-β structure formation. The results provide a detailed structural view of early amyloid assembly involving structured species that are not cytotoxic.

https://doi.org/10.7554/eLife.46574.001

Introduction

Oligomers have been the focus of amyloid research over decades because of their pivotal role in assembly and their potential cytotoxicity (Chiti and Dobson, 2017). Numerous aggregation-prone proteins (or their fragments) form oligomers (Benilova et al., 2012; Wei et al., 2011; Laganowsky et al., 2012; Apostol et al., 2013), some of which are cytotoxic (Laganowsky et al., 2012; Ono et al., 2009; Fusco et al., 2017), while others are not (Mannini et al., 2014). Many groups have attempted to elucidate the structure(s) of amyloid oligomers with different biological properties (Chiti and Dobson, 2017). However, their ephemeral nature, dynamic signature, and heterogeneity in mass and conformation provide significant experimental challenges. Hence, our current understanding of the structure of oligomers is often limited to low-resolution models (Fusco et al., 2017; Vestergaard et al., 2007; Campioni et al., 2010; Cremades et al., 2012), or to oligomers assembled from non-natural amino acids, short peptides, or protein fragments (Laganowsky et al., 2012; Apostol et al., 2013; Sangwan et al., 2017). Establishing a relationship between the oligomers observed and the mechanism of amyloid formation is also an important, but challenging, task. In some cases, oligomers have been shown to be ‘off-pathway’ since they have to dissociate for amyloid formation to proceed (Wu et al., 2010; Baskakov et al., 2002; Souillac et al., 2003; Bieschke et al., 2010). Characterization of such species, however, does not provide insight into the structural mechanism by which initially unstructured (e.g. Aβ42, α-synuclein) or natively structured proteins (e.g. lysozyme, transthyretin, antibody light chains, β₂-microglobulin (β₂m)) undergo conformational conversion to form the parallel in-register cross-β structure of amyloid (Iadanza et al., 2018a). Other oligomers have been shown to be on-pathway (Fusco et al., 2017; Cremades et al., 2012), or to form via secondary nucleation processes that enhance the rate of fibril formation (Cohen et al., 2013). Proteins in an oligomeric or aggregated form have also been characterized kinetically, thermodynamically and biophysically (Cohen et al., 2013; Cohen et al., 2018; Lenton et al., 2017). However, a detailed understanding of both the structural properties of oligomers and their role in assembly is needed in order to understand the structural mechanism(s) of amyloid formation and the origins of cytotoxicity, as well as to design inhibitors of the assembly process.

Here, we describe an integrative approach which uses kinetic modeling to identify oligomers formed on-pathway to fibril formation, NMR spectroscopy and other biophysical methods to determine their structural properties, and cellular assays to determine their cytotoxicity. The strategy employed can be applied to other assembling protein systems and draws on the powers of NMR to provide detailed structural information about individual precursors in dynamic equilibrium within complex mixtures of assembling species, and kinetic modeling to ascribe their role in amyloid formation. To exemplify this combined structural and kinetic approach we focus on the naturally occurring variant of human β₂m, known as ΔN6. This variant lacks the N-terminal six amino acids and is formed by natural proteolytic truncation of the wild-type (WT) protein (Esposito et al., 2000; Eichner et al., 2011). WT human β₂m (named herein as hβ₂m) forms amyloid deposits in the joints of patients undergoing long term hemodialysis (Gejyo et al., 1985). However, hβ₂m does not aggregate into amyloid fibrils at physiologically relevant pH and temperature on an experimentally accessible timescale in vitro (the pH in normal and diseased joints ranges from 5.5 to 7.4; Floege and Ehlerding, 1996). Addition of Cu²⁺ ions, detergents, organic solvents, glycosaminoglycans or collagen can drive hβ₂m amyloid formation at neutral pH (Platt and Radford, 2009; Stoppini and Bellotti, 2015; Yamamoto et al., 2005; Benseny-Cases et al., 2019). These reagents partially unfold the native protein, facilitating cis-trans isomerization of Pro32 that initiates assembly (Eichner et al., 2011; Platt and Radford, 2009; Yamamoto et al., 2005; Chiti et al., 2001). By contrast with the intransigence of hβ₂m to form amyloid in vitro, ΔN6 is highly amyloidogenic, forming fibrils rapidly in vitro in the absence of additives at pH 6–7 (Karamanos et al., 2016). ΔN6 forms ~ 30% of β₂m in amyloid plaques in patients with dialysis-related amyloidosis (Bellotti et al., 1998). Previous studies have shown that ΔN6 can induce amyloid formation of hβ₂m at near-neutral pH in vitro (Eichner et al., 2011) and can co-assemble with the WT protein into amyloid fibrils (Sarell et al., 2013). The structure of hβ₂m in amyloid fibrils formed in vitro at low pH (pH 2.0) has also been solved recently using solid-state NMR and cryo-EM, revealing a parallel in-register cross-β structure typical of amyloid, which differs dramatically from the all anti-parallel immunoglobulin fold of the native precursor (Iadanza et al., 2018b). The atomic structure(s) of hβ₂m amyloid fibrils formed in vivo, and those of ΔN6 formed in vitro or ex vivo, however, are not yet known.

Several examples of oligomers (dimers, tetramers and hexamers) of WT hβ₂m have been reported previously (Calabrese et al., 2008; Eakin et al., 2006; Mendoza et al., 2011; Mendoza et al., 2010; Halabelian et al., 2015; Colombo et al., 2012; Rennella et al., 2013; Liu et al., 2011; Karamanos et al., 2014), with one report of a domain swapped dimer of ΔN6 stabilized by addition of a nanobody (Domanska et al., 2011). Since hβ₂m is inert to aggregation at physiological pH and temperature in vitro, the oligomerization of the protein was stimulated by mutation and/or the addition of Cu²⁺ ions (Calabrese et al., 2008; Eakin et al., 2006; Mendoza et al., 2011; Mendoza et al., 2010), or by linkage of monomers via non-native disulfide bonds (Halabelian et al., 2015; Colombo et al., 2012). Although some of these oligomers form under conditions in which WT hβ₂m may eventually form fibrils, the role of individual oligomeric species in the aggregation mechanism remains unclear. The oligomers formed in the initiating stages of aggregation of ΔN6 into amyloid also remain obscure.

Here, we show that amyloid formation of ΔN6 occurs via a remarkably specific assembly mechanism involving the transient formation of dimers and hexamers. Exploiting NMR methods able to analyze dynamic and lowly populated states (Anthis and Clore, 2015), we characterize these assemblies, yielding a structural model of the initiating events in ΔN6 aggregation in atomic detail. The results reveal the formation of head-to-head dimers that pack into symmetric hexamers that retain a native-like immunoglobulin fold and are not cytotoxic. The hexamers appear to be primed for further conformational change into the cross-β structure of amyloid by dynamic unfurling of their C-terminal β-strands. The results portray a detailed atomic view of the early stages of ΔN6 assembly that may enable the development of routes to combat disease by targeting the specific protein-protein interactions that define the early stages of assembly.

Results

Fibril elongation occurs through an oligomeric state

Previous results have shown that ΔN6 assembles rapidly into amyloid fibrils in vitro at pH 6.2, but not at pH 8.2 (Eichner et al., 2011), suggesting that lowering the pH increases the population of aggregation-prone species. Such species may also be relevant in vivo given the acidic microenvironment of the joints of DRA patients (Eichner et al., 2011; Bellotti et al., 1998; Karamanos et al., 2014). At pH 6.2 (close to its pI of 5.8) ΔN6 is dynamic, but retains a native-like immunoglobulin fold (Eichner et al., 2011). To determine the kinetic mechanism by which ΔN6 aggregates into amyloid fibrils, experiments were performed in which ΔN6 fibril seeds (20 μM monomer equivalent concentration) were incubated with different concentrations of ΔN6 monomers (20 μM to 500 μM) and the rate of amyloid formation was monitored by the fluorescence of thioflavin T (ThT). All experiments were performed at pH 6.2 at a total ionic strength of 100 mM (see Materials and methods). The simplest kinetic mechanism in which monomers add to the fibril ends would result in a linear dependence of the initial rate of fibril elongation versus the monomer concentration, with saturation at high monomer concentrations (Buell et al., 2014; Buell et al., 2010; Xue et al., 2009). Such behavior is observed for seeded assembly of acid unfolded monomers of hβ₂m, which initially lack persistent structure (Platt et al., 2008), into amyloid fibrils at pH 2.0 (Figure 1a,b). By contrast, ΔN6 showed more complex behavior, with a clear non-linearity in the initial rate of elongation versus monomer concentration, in which rapid seeded growth occurs only above ~200 μM ΔN6 (Figure 1c,d). This indicates that fibril elongation by ΔN6 must involve addition of one or more oligomeric species to the fibril ends under the conditions employed.

Figure 1

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Dependence of the fibril elongation rate on the concentration of soluble protein.

Seeded elongation assays for (a) hβ₂m at pH 2.0 monitored by ThT fluorescence. 20 μM of preformed seeds of hβ₂m (formed at pH 2.0) and varying amounts of soluble protein were added, as indicated in the key. Note that the protein does not aggregate under these conditions in the absence of seeds on this timescale (Xue et al., 2008). The dashed line shows the initial rate of each reaction. (b) The initial rate of fibril elongation (shown in units of ThT fluorescence (a.u.)/h) versus the concentration of hβ₂m added. The dashed line represents a prediction using a monomer addition model (see Table 4). (c) Seeded elongation assays for ΔN6 using 20 μM preformed seeds formed from ΔN6 at pH 6.2 as a function of the concentration of soluble ΔN6 added. Open blue symbols denote the ThT fluorescence signal of 500 μM ΔN6 in the absence of seeds. The dashed line shows the initial rate of each reaction. (d) The initial rate of fibril elongation (shown in units of ThT fluorescence (a.u)/h) versus the concentration of soluble ΔN6 added. The dashed line shows the dependence of the elongation rate (in units of ThT fluorescence (a.u)/h) on the concentration of monomer assuming a monomer addition model (see Table 4). The elongation rate for monomer addition shows a hyperbolic behavior as a function of monomer concentration, with a linear dependence at lower monomer concentrations, followed by a saturation phase at higher monomer concentrations. The simulation in (b) (dashed line) uses a slower microscopic elongation rate (k_e) (Table 4) than that used in panel (d) and therefore saturation is not achieved by 410 μM protein in (b), but is in (d). Five replicates are shown for each protein concentration. Error bars show the standard deviation between all replicates.

https://doi.org/10.7554/eLife.46574.002

Native-like dimers and hexamers form during Δn6 assembly

The concentration-dependence of ΔN6 elongation could be explained by an oligomer(s) acting as the elongation unit. To explore whether oligomeric species of ΔN6 are formed under the conditions employed, sedimentation velocity analytical ultracentrifugation (AUC), size exclusion chromatography (SEC), cross-linking, and NMR experiments were performed. These approaches report on the conformational properties and molecular weight distribution of the assemblies formed at different ΔN6 concentrations. Sedimentation velocity AUC experiments showed that ΔN6 forms discrete oligomers at pH 6.2, with monomers, dimers and higher order species with a sedimentation coefficient (S value) consistent with 6–9-mers (although the rapid equilibration of the species present prevents accurate determination of their mass and population) (Figure 2a). To investigate the molecular mass of the species present, ΔN6 was cross-linked after different incubation times in the absence of fibril seeds using 1-ethyl-3-(3-dimethylaminopropyl)-carbodiimide hydrochloride (EDC) (see Materials and methods) and the resulting species examined using SDS-PAGE (Figure 2b). This revealed the presence of hexamers during assembly (Figure 2b). The population of the hexameric species is decreased at later time points, presumably because it is consumed into fibrils (Figure 2b). Analytical SEC of ΔN6 at different protein concentrations without cross-linking revealed only monomers and dimers (Figure 2—figure supplement 1a), consistent with the higher order assemblies dissociating upon dilution on the column. However, when cross-linking was performed prior to SEC, higher molecular weight oligomers were observed, with these species being more abundant when higher protein concentrations were used (Figure 2—figure supplement 1b,c). At the highest concentration of ΔN6 used (500 μM) cross-linking resulted in the rapid formation of high molecular weight aggregates that elute in the void volume (Figure 2—figure supplement 1b,c). The population of these aggregates increases with time, accompanied by depletion of the oligomers, consistent with these species being capable of assembly into amyloid (Figure 2—figure supplement 1d).

Figure 2 with 2 supplements see all

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ΔN6 oligomer formation.

(a) Sedimentation velocity AUC of ΔN6 at different concentrations, as indicated by the key. Note that the higher order species decrease in intensity at high protein concentrations (>200 μM) consistent with the formation of large aggregates that sediment rapidly before detection (see also Figure 2—figure supplement 1d). (b) SDS–PAGE of cross-linked ΔN6 (80 μM) at different time-points during de novo fibril assembly in the absence of fibril seeds (see Materials and methods). Note that dimers are not observed, presumably as they are not resilient to the vigorous agitation conditions used to accelerate fibril formation in these unseeded reactions, or are not efficiently cross-linked by EDC under the conditions used (see Materials and methods). A negative stain electron micrograph of ΔN6 after 100 hr of incubation is shown below. Scale bar – 500 nm. (c) The methyl region of the ¹H NMR spectrum of ΔN6 at 400 μM (left) or 10 μM ΔN6 (right). (d) Per residue combined ¹H-¹⁵N chemical shift differences between the ¹H-¹⁵N HSQC spectrum of ΔN6 at 10 μM and 400 μM. Blue dots represent residues for which assignments are missing in both spectra. The dashed line represents one standard deviation (σ) of chemical shifts across the entire dataset. Residues that show chemical shift differences > 1σ are shown in yellow,>2σ are colored red, and residues for which the chemical shift difference is not significant (<1σ) are colored gray. Residues that are broadened beyond detection in the spectrum obtained at 400 μM are colored in magenta (see also Figure 2—figure supplement 2a). Residues are numbered according to the sequence of the WT protein. Arg 97 is hydrogen bonded to residues in the N-terminus and presumably is indirectly affected by the interaction. (e) The structure of ΔN6 (2XKU; Eichner et al., 2011) colored in the same scheme as (d). Pro32 is shown in blue space-fill. The buffer used in all experiments was 10 mM sodium phosphate pH 6.2 containing 83.3 mM NaCl (to maintain a constant ionic strength of 100 mM for all experiments), 25°C.

https://doi.org/10.7554/eLife.46574.003

¹H-NMR and ¹H-¹⁵N HSQC NMR spectra of ΔN6 were next acquired to examine the properties of the oligomers formed. Significant changes in chemical shift and linewidth of individual resonances at different concentrations of ΔN6 were observed, consistent with the finding that ΔN6 self-assembles into higher molecular weight species at pH 6.2 (Figure 2c and Figure 2—figure supplement 2a). The residues most affected lie in the A strand and the BC, DE and FG loops, suggesting that these regions form the intermolecular interfaces in the higher molecular weight species (Figure 2d,e). Consistent with these observations, measurement of the rotational correlation time (τ_c) and diffusion coefficient of the sample, which reflect the average size and shape of the molecules formed, showed a linear dependence on ΔN6 concentration, consistent with protein oligomerization in which the resulting species are in dynamic exchange (Figure 2—figure supplement 1e,f). Together these results show that ΔN6 assembles into dimers and hexamers that are assembly competent, in dynamic exchange, and assemble via interfaces which are located in the apical region of the protein that surrounds Pro32 (Figure 2e).

To estimate the dissociation constants for dimer and hexamer formation, the chemical shifts and residual dipolar couplings (RDCs) of individual resonances were measured as a function of ΔN6 concentration from 10 to 410 μM (Figure 2—figure supplement 2a–e). Significant chemical shift differences were observed when the ΔN6 concentration was increased from 10 μM to 50 μM without significant line broadening (Figure 2—figure supplement 2c, panels i-iii). Increasing the protein concentration to 100 μM caused a decrease in the chemical shift differences (Figure 2—figure supplement 2c, panel iv), which then increase again in magnitude at 200 μM and 410 μM, accompanied by significant line broadening (Figure 2—figure supplement 2a and c, panels v,vi). This complex behavior is consistent with a monomer-dimer-hexamer equilibrium in which the monomers and dimers have different chemical shifts, while the chemical shifts of dimers and hexamers are similar (an assumption that is supported by our structural models, see below), and the exchange rate between monomers and hexamers is significantly faster than that between monomers and dimers. Therefore, the monomer-dimer equilibrium dominates the equilibrium (and the observed chemical shift) at low concentrations (50 μM). At higher concentrations the dimer is depleted relative to the hexamer and the chemical shift observed becomes a complex combination of the population of each species, the exchange rate between each species, and the difference in chemical shift of each residue in each assembly. Fitting the chemical shift data to a monomer – dimer – hexamer model yields a K_d for dimer formation of ≤50 μM, while that of hexamer formation is ~10 ± 5 x 10⁻⁹ M² (Benilova et al., 2012) (see Materials and methods and Figure 2—figure supplement 2b,d), indicating that once dimers form they have a high affinity for one another. Importantly, the monomer – dimer – hexamer model with the estimated affinities adequately describes the observed increase in the τ_c and the observed diffusion coefficient versus protein concentration (Figure 2—figure supplement 1e,f), independently supporting the model derived. Increasing the K_d for dimer formation to >100 μM results in unrealistically low values for the hexamerization K_d (Figure 2—figure supplement 2d). Moreover, measurement of RDCs versus protein concentration results in a biphasic curve (Figure 2—figure supplement 2e), consistent with a multi-equilibrium process. Using these data, the RDCs of the dimer species can be calculated for a range of estimated K_d values (see Materials and methods). Fitting the dimer RDCs to the structure of ΔN6 (2XKU; Eichner et al., 2011), shows significantly poorer fits to the predicted RDC values assuming a dimer K_d higher than 50 μM (Figure 2—figure supplement 2f). To explain the chemical shift and RDC data, therefore, the dimer K_d must be ≤50 μM.

Specific interfaces determine aggregation

To map the interfaces involved in ΔN6 oligomer formation in more detail, intermolecular paramagnetic relaxation enhancement (PRE) experiments were performed. The PRE depends on the distance between a paramagnet and adjacent nuclei and can provide distance information about (transient) binding interfaces for nuclei that are within ~20 Å of the paramagnetic center (Clore and Iwahara, 2009), quantified by the effect of the spin label on the relaxation rates of each amide proton (the H_N-Γ₂ PRE rate). ¹⁴N-ΔN6 was spin-labeled with (1-oxyl-2,2,5,5-tetramethyl-D3-pyrroline-3-methyl) methanethiosulfonate (MTSL) by creating Cys variants at positions 20, 33, 54 or 61. Each protein (60 μM) was then mixed with ¹⁵N-ΔN6 (60 μM) at pH 6.2. At this total protein concentration, the PREs observed are dominated by the monomer-dimer equilibrium (35% of molecules are monomer, 51% of ΔN6 molecules are in dimers and 14% of ΔN6 molecules are in hexamers, determined from the K_d values measured above). These experiments (Figure 3) showed increased H_N-Γ₂ rates for residues in the A strand and the BC, DE and FG loops when the spin label is attached to residues 33, 54, or 61, suggestive of a head-to-head interaction involving the apical regions of the protein (Figure 3a–c and e). In accord with this conclusion, when MTSL is attached at position 20 at the distal side of the protein (Figure 3e), the H_N-Γ₂ rates are vastly decreased (Figure 3d).

Figure 3 with 2 supplements see all

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Identification of interacting surfaces in ΔN6 dimers.

Intermolecular PRE data for the self-association of ΔN6. ¹⁵N-ΔN6 (60 μM) was mixed with an equal concentration of (a) ¹⁴N-(S33C)ΔN6-MTSL; (b) ¹⁴N-(L54C)ΔN6-MTSL; (c) ¹⁴N-(S61C)ΔN6-MTSL; or (d) ¹⁴N-(S20C)ΔN6-MTSL in 10 mM sodium phosphate buffer, pH 6.2 containing 83.3 mM NaCl (a total ionic strength of 100 mM). The resulting Γ₂ rates are color-coded according to the amplitude of the PRE effect (see scale bar: gray-insignificant (<20 s⁻¹), yellow->20 s⁻¹, red->50 s⁻¹, pH 6.2, 25°C). Blue dots in the plots are residues for which resonances are not assigned (na) at pH 6.2. Red crosses indicate high H_N-Γ₂ rates for which an accurate value could not be determined. Control experiments showed that the small PREs arising from¹⁴N-(S20C)ΔN6-MTSL arise from non-specific interactions with MTSL itself. Solid black lines depict fits to the PRE data for the dimer structure shown in Figure 4a. Note the poor fits for some residues which are sensitive to hexamer formation (14% of ΔN6 molecules) under the conditions used. Residues are numbered according to the WT sequence and the position of β-strands (2XKU; Eichner et al., 2011) is marked above each plot. (e) The structure of ΔN6 (2XKU; Eichner et al., 2011) with the BC loop shown in magenta, the DE loop in green and the FG loop in yellow. The MTSL attachment sites are highlighted as spheres.

https://doi.org/10.7554/eLife.46574.006

To determine whether the head-to-head dimers are critical for aggregation, the AUC, PRE and fibril growth experiments were also performed at pH 8.2 where ΔN6 does not assemble into amyloid fibrils even after extended incubation times (Figure 3—figure supplement 1a). The sedimentation velocity AUC experiments revealed that monomers and tetramers are formed at pH 8.2, but not hexamers, with the equilibrium in favor of the monomer (Figure 3—figure supplement 1b). Consistent with this, the τ_c of 600 μM ΔN6 at pH 8.2 is ~12 ns, in marked contrast with the τ_c of ~50 ns predicted for 600 μM ΔN6 at pH 6.2 (Figure 2—figure supplement 1e). Finally, intermolecular PRE experiments at pH 8.2 showed small Γ₂ rates irrespective of the site of MTSL labeling (Figure 3—figure supplement 1c–d), suggesting that the monomers bind with different affinity and/or via different interfaces at this pH. To investigate these hypotheses, CPMG relaxation dispersion NMR experiments were performed. These experiments are able to detect excited states populated to as little as 1% of the total protein in solution (Hansen et al., 2008). Concentration-dependent CPMG profiles of residues in the B strand, D strand, DE loop, E strand and EF loop were observed at pH 8.2 (Figure 3—figure supplement 2a–d), indicating that the binding interface for tetramer formation differs substantially from the loop-loop interactions in the apical region of the protein that dominate assembly at pH 6.2, despite the fact that ΔN6 retains an immunoglobulin-like fold at both pH values (Figure 3—figure supplement 2e,f). As a consequence of the altered interface that forms at pH 8.2, hexamers and fibrils do not form. Together these results indicate that the head-to-head dimers formed at pH 6.2 are uniquely able to assemble into the hexamers that are crucial for fibril assembly.

Different dimer structures determine amyloid inhibition and propagation

To generate dimer structures consistent with the experimental data obtained, simulated annealing molecular dynamics calculations were performed. The calculations converged to two dimer structures (Figure 4a, Figure 4—figure supplement 1 and Table 1). In the lowest energy model (model A), the ΔN6 monomers are arranged in an extended conformation with the N-terminal residues M6 and I7 (WT numbering), along with the BC, DE and FG loops forming the interface (Figure 4a). The inhibitory dimer of ΔN6:murine β₂m (mβ₂m) was previously determined using a similar approach (Karamanos et al., 2014). This dimer also has a head-to-head subunit arrangement but is characterized by a more acute angle between ΔN6 subunits in which the monomers interact predominantly through the BC and DE loops (Karamanos et al., 2014) (Figure 4b, Video 1). Thus, distinct protein dimers formed from closely related sequences (mβ₂m and hβ₂m are 70% identical and 90% similar in sequence) give rise to fundamentally different outcomes of assembly.

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Structural models of ΔN6 dimers.

Structural models of (a) the lowest energy ΔN6 homodimer (dimer A) and (b) the ΔN6-mβ₂m heterodimer that inhibits ΔN6 fibril assembly (Karamanos et al., 2014). Interface residues (identified as those residues that have any pair of atoms closer than 5 Å) are shown in a ball and stick representation on one subunit and are colored in space fill in gold in (a) or red in (b) on the surface of the second subunit. ΔN6 is shown in the same pose (blue) in (a) and (b). The BC, DE and FG loops are shown in magenta, green and yellow, respectively, and the position of attachment of MTSL for the PRE experiments (residues 20, 33, 54 and 61) is highlighted in spheres. PDB files are publicly available from the University of Leeds depository (https://doi.org/10.5518/329). See also Video 1.

https://doi.org/10.7554/eLife.46574.009

Table 1

Agreement between experimental and back-calculated intermolecular PREs for the two dimer structures (dimer A and dimer B (see Figure 5—figure supplement 3).

RMS values are shown comparing the measured versus the predicted values from the structure PREs measured from S33, L54 and S61. Data from position S20 were not used as they arise from non-specific interactions with MTSL.

https://doi.org/10.7554/eLife.46574.011

PRE term	RMS dimer A	RMS dimer B
S33C(ΔN6)-ΔN6 (s⁻¹)	18.65	15.10
L54C(ΔN6)-ΔN6 (s⁻¹)	29.02	27.44
S61C(ΔN6)-ΔN6 (s⁻¹)	19.44	23.27
*High PREs (Å)	2.78	3.79

*High PREs refer to PREs in the BC loop (measured from S33, L54 and S61) that (due to their large value) could not be measured accurately and therefore are incorporated as loose distance restraints.

Video 1

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posterframe for video — Comparison of productive and inhibitory dimers.

The ΔN6 subunit in each dimer (this study) is shown as a dark blue cartoon, while the second ΔN6 monomer in the productive dimer and the mβ₂m subunit in the inhibitory dimer (Karamanos et al., 2014) are shown as light blue and red, respectively. The BC, DE, FG loops are colored in magenta, green and yellow, respectively, while the intra-dimer interface residues are shown as sticks on both subunits.

https://doi.org/10.7554/eLife.46574.012

Structural models of on-pathway hexamers

Although the majority of the intermolecular PREs can be satisfied by the dimer A structure, the fits are not perfect (Figure 3a–d), presumably since ~14% of ΔN6 molecules form hexamers at the concentration of ΔN6 employed (120 μM). The PRE experiments were thus repeated at higher concentrations (320–400 μM) of ΔN6, wherein > 40% of ΔN6 molecules are predicted to be in hexamers. These experiments revealed a pattern of H_N-Γ₂ rates similar to those obtained at 120 μM ΔN6 (Figure 5—figure supplement 1a–d), with the highest H_N-Γ₂ rates involving the N-terminus, BC, DE and FG loops, suggesting that similar interfaces are formed in the dimer and hexamer. CPMG experiments at 180 μM ΔN6 (26% of ΔN6 molecules are monomer, 48% are dimer, and 26% are hexamer) and 480 μM ΔN6 (13% of ΔN6 molecules are monomer, 32% are dimer and 55% are hexamer) showed that residues in the apical regions of ΔN6, surrounding Pro 32, are also in concentration-dependent exchange at both ΔN6 concentrations at pH 6.2, in support of this conclusion (Figure 5—figure supplement 2).

The ordered nature of assembly (monomer, dimer, hexamer) and the identification of the interfaces involved, allowed us to generate models for the hexameric species (Figure 5—figure supplement 3). The measured PREs were converted into distances and simulated annealing molecular dynamics calculations were performed to create hexamer structures consistent with the experimental PRE and chemical shift data using the lowest energy dimer model (dimer A shown in Figures 4a and 5 - Figure 5—figure supplement 3a), as well as the less favorable dimer model B (Figure 4—figure supplement 1e and Figure 5—figure supplement 3b), as starting points. Note that the structure calculation strategy employed does not require knowledge of the dimer and hexamer populations (see Materials and methods). Starting from dimer A (Figure 4a) the structure calculation resulted in a hexamer in which the three dimers trimerize to form a compact daisy-like structure (Figure 5a–c). The PREs back-calculated from this model are consistent with the experimental data (Figure 5—figure supplement 4). Importantly, hexamer structures generated from dimer B (Figure 4—figure supplement 1e) resulted in poorer fits to the PRE profiles (Materials and methods and Table 2).

Figure 5 with 6 supplements see all

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Structural model of ΔN6 hexamers.

(**a–c**) Sphere representations of the hexamer model formed from dimer A rotated by 90° in each view. Subunits belonging to the same dimer are colored in different tones of the same color. (d) The monomer-monomer (intra-dimer) interface is highlighted in green on the surface of the dimer formed from subunits 1a and 1b (within dimer A), with the other dimers shown as cartoons. (e) The inter-dimer interface is colored red on the surface of the dimer formed from subunits 1a and 1b, with the dimers shown as cartoons. (f) As in (e), but showing the dimer formed from subunits 1a and 1b, superposed with the mβ₂m subunit in the inhibitory ΔN6-mβ₂m dimer (Karamanos et al., 2014) (green cartoon). The ΔN6-ΔN6 and ΔN6-mβ₂m dimers were aligned on the ΔN6 subunit 1b. Schematics of the assemblies are shown at the bottom colored as in (**d–f**). Note that the BC, DE and FG loops are highlighted as thicker chains in blue, green and cyan, respectively, in d-f. PDB files are publicly available from the University of Leeds depository (https://doi.org/10.5518/329). See also Video 2.

https://doi.org/10.7554/eLife.46574.013

Table 2

Agreement between experimental and back-calculated intermolecular distances for different hexamer structures.

RMS values are shown comparing the measured versus the predicted distances from each structural model for distances measured from S33, L54 and S61. Data from position S20 were not used as they arise from non-specific interactions with MTSL. See also Figure 5—figure supplement 3.

https://doi.org/10.7554/eLife.46574.020

PRE term	Hexamer 1 RMS (Å)	Hexamer 2(i) RMS (Å)	Hexamer 2(ii) RMS (Å)	Hexamer 2(iii) RMS (Å)
S33C(ΔN6)-ΔN6	2.34	2.68	2.58	2.53
L54C(ΔN6)-ΔN6	1.25	2.33	2.26	1.87
S61C(ΔN6)-ΔN6	2.22	2.7	2.68	3.11

In the hexamer models generated from dimer A the dimeric subunits are arranged in a helical manner twisted by ~120°, creating a hexamer that is ~60 Å in diameter and 75 Å in length. This hexamer model is consistent with the collision cross-section (CCS) of ΔN6 hexamers measured using the lowest charge state (15+) (the most native-like species; Vahidi et al., 2013; see Materials and methods) detected using Electrospray Ionization Ion Mobility Spectrometry – Mass Spectrometry (ESI-IMS-MS), but the measured CCS is inconsistent with hexamers derived using dimer B (Figure 5—figure supplement 5a). The monomer-monomer and dimer-dimer interfaces in the best fit hexamer structure (Figure 5a–e) involve similar, but not identical, regions, with the inter-dimer interface extending further into the β-sheet containing the A, B, E and D β-strands, while the intra-dimer interface is formed mostly through the BC and DE loops (Figure 5d–e and Video 2). The formation of dimers generates a hydrophobic surface which becomes buried in the hexamer (Figure 5e, Figure 5—figure supplement 5b and Table 3). Consistent with this, the cross-linked hexamers show a small (1.3-fold) increase in fluorescence in the presence of the hydrophobic dye 8-anilino-1-naphthalenesulfonic acid (ANS), that is much smaller than the ~100 fold increase in ANS fluorescence observed for a typical ‘molten globule’ state (Semisotnov et al., 1991), but similar in magnitude to ANS bound to the highly structured on-pathway folding intermediate of Im7 (monitored using the trapped equilibrium mimic of this species, Im7 L53AI54A (Spence et al., 2004) (Figure 5—figure supplement 5d). The ΔN6 dimers show a similar increase in ANS fluorescence as the hexamers despite having a larger exposed hydrophobic surface area, possibly because ANS binds more weakly or has a lower quantum yield when dimer-bound (Figure 5—figure supplement 5c–d). The interface formed in the inhibitory ΔN6-mβ₂m dimer overlaps with the surface required for hexamerization, but not for ΔN6-ΔN6 dimerization (Figure 5f), rationalizing why mβ₂m is able to inhibit amyloid formation (note that the K_d of the mβ₂m:ΔN6 complex is 70 μM (Karamanos et al., 2014), similar to that (~50 μM) estimated here for ΔN6 homo-dimerization). The dimers and hexamers were incubated with SH-SY5Y cells, a cell line that is commonly used in studies of amyloid toxicity (Laganowsky et al., 2012; Fusco et al., 2017; Campioni et al., 2010; Jakhria et al., 2014; Leri et al., 2016; Giorgetti et al., 2008), and which has been shown previously to take up monomeric and fibrillar β₂m (Jakhria et al., 2014). Interestingly, there was no evidence for cytotoxicity in assays for 3-(4,5-dimethylthiazol-2-yl)−2,5-diphenyltetrazolium bromide (MTT) reduction, lactate dehydrogenase release, reactive oxygen species formation and cellular ATP level (see Materials and methods) (Figure 5—figure supplement 6). However, rapid dissociation of the uncross-linked oligomers, prevention of conversion to a cytotoxic form by cross-linking, or cytotoxicity requiring different cell types or prolonged exposure (>24 hr) to the oligomers cannot be ruled out.

Table 3

Analysis of dimer and hexamer interfaces.

The buried surface area is calculated as the sum for the two subunits for each complex. Interface residues were identified as those residues that lose at least 10% of accessible surface area upon oligomer formation.

https://doi.org/10.7554/eLife.46574.021

	ΔN6 dimer A	ΔN6 hexamer
Buried Surface Area (Å²)	1233	4201
% Charged residues in the interface	28	18
% Hydrophobic residues in the interface	44	54

Video 2

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Hexamer dynamics may prime further assembly into amyloid

The hexamer shown in Figure 5 retains a native-like immunoglobulin fold in which the β-strands are anti-parallel. Hence, a major conformational rearrangement has still to occur for ΔN6 to form amyloid fibrils in which the β-strands stack in a parallel in-register structure (Debelouchina et al., 2010) (R. Silvers, Y. Su, R.G. Griffin, and S.E. Radford, unpublished). Hints of how this conformational change may be initiated were obtained by quantitative analysis of the CPMG data shown in Figure 6, Figure 5—figure supplement 2 and Figure 6—figure supplement 1. Globally fitting these data for residues which lie in the dimer and/or hexamer interfaces (residues 26, 34, 35, 37, 51, 59, 65, 66, 67, 83, 84, 85, Figure 5d–e) to a two-state fast exchange model yields an exchange rate, k_ex^bind, of 1790 ± 290 s⁻¹ (Figure 6a,b). Distinct CPMG profiles were observed, however, for residues 87, 89, 91 and 92 which lie in the G strand of monomeric ΔN6 and which are not involved in the dimer-dimer interfaces (they show no significant concentration-dependent chemical shifts, nor PREs are observed for these residues at low or high protein concentration (Figure 6—figure supplement 1a). The CPMG data for these residues presumably report on conformational changes that result from hexamerization rather than the direct binding event itself. The CPMG data indicate that these residues exchange with a lowly populated (2%) excited state with an interconversion rate, k_ex^G, of 205 ± 150 s⁻¹, 10-fold slower than k_ex^bind (Figure 6c–d and Figure 6—figure supplement 1a–e). Therefore, a distinct process, possibly local unfolding of the C-terminal β-strand, occurs when the hexamer is formed that is driven by the free energy of hexamer formation (ΔG°_hexamer ~4 kJ/mol). At ΔN6 concentrations of 480 μM k_ex^G is increased to 1170 ± 196 s⁻¹ (Figure 6—figure supplement 1f–i), consistent with increased hexamer formation enhancing the observed rate of dynamics of the G strand. Hexamer formation thus potentially destabilizes the G-strand of ΔN6, causing local unfolding of this region of the polypeptide chain (although further experiments measuring the sign of the chemical shift change would substantiate this conclusion). This may then lead to more catastrophic structural reorganization of the hexamer into the parallel in-register structure of amyloid (note that the G-strand sequence forms a β-strand in the ΔN6 fibril core; Su et al., 2014). Whether structural conversion occurs within the hexamer, at the fibril end, or requires further, more elaborate molecular steps such as active participation of the fibril surface, or disassembly into smaller structural units prior to fibril assembly, remains to be seen.

Figure 6 with 1 supplement see all

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G-strand unfurling may occur upon hexamer formation.

¹⁵N CPMG relaxation dispersion data at 750 MHz (magenta) and 950 MHz (red) (180 μM ΔN6, pH 6.2 (26% ΔN6 molecules are monomers, 48% are in dimers, 26% are in hexamers) for residues (a) 51, (b) 37, (c) 89, and (d) 92. Residues 37 and 51 report on intermolecular interactions that describe dimer and/or hexamer formation (schematic, top left), while residues 89 and 92 do not lie in an interface and report instead in the dynamics of the G strand in the different assemblies formed. The position of all five residues used in the cluster analysis of G strand dynamics is shown in spheres on the structure of ΔN6 (blue cartoon, top right). Pro32 is shown as a magenta sphere. Solid lines represent global fits to the Bloch-McConnell equations (Materials and methods) for each cluster of residues. The extracted parameters of the global fit for the two processes (k_ex^bind and k_ex^G) are indicated above the plots.

https://doi.org/10.7554/eLife.46574.023

A unified model of Δn6 polymerization

As a final test of the validity of the model of ΔN6 assembly proposed we assessed the ability of the structural, kinetic and thermodynamic parameters deduced above to describe the observed rates of fibril formation measured using ThT fluorescence, as well as the τ_c values versus ΔN6 concentration measured by NMR, and the fibril yield. Using the dimer and hexamer structural models shown in Figures 4 and 5 and the estimated K_d values for their formation, all of the derived experimental data could be recapitulated (Figure 7). Fitting the seeded fibril growth data to different kinetic models that assume (i) monomers to add to the fibril ends (Figure 7—figure supplement 1a); (ii) monomers are in exchange with a monomeric excited state that is responsible for growth (Figure 7—figure supplement 1b); or (iii) dimers are the elongation units (Figure 7—figure supplement 1c), fail to describe the seeding data (Materials and methods and Table 4). By contrast, a model assuming addition of hexamers describes the ThT kinetic profiles well (Figure 7a), while a more complicated monomer-dimer-tetramer-hexamer model does not improve the fit significantly (Figure 7—figure supplement 1d). The populations of monomer, dimer and hexamer, together with the derived structural models, are also consistent with the observed dependence of τ_c on protein concentration (Figure 7b). Finally, the amount of hexamer formed (in the absence of seeds) is also predictive of the fibril yield (Figure 7c,d) consistent with the hexamer being required for fibril formation. This conclusion is also supported by the appearance of hexamers early during assembly in the absence of seeds and their disappearance as fibrils form (Figure 2b).

Figure 7 with 1 supplement see all

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The monomer-dimer-hexamer model describes the thermodynamics and kinetics of fibril elongation.

(a) Global fits (blue solid lines) to the fibril elongation kinetics monitored by ThT fluorescence assuming a hexamer addition model at different concentrations of soluble ΔN6 (dots) (Materials and methods and Table 4). The concentrations of ΔN6 are colored according to the key. The average of five replicates is shown. (b) Protein correlation times (τ_c) measured using NMR (red) and back-calculated values (green) using the populations of monomers, dimers and hexamers predicted from the monomer-dimer-hexamer model and the correlation times of the dimers and hexamer structural models shown in Figures 4 and 5. (c) The fibril yield (after 100 hr) of each elongation reaction. SDS-PAGE analysis of the whole reaction (shown in (a)) before centrifugation (W) or of the supernatant (S/N) after centrifugation at the different concentrations of ΔN6, as indicated. (d) Bar-charts showing the % of insoluble material (gray) measured using densitometry of the gel shown in (c). The % hexamer population in the absence of seeds (black) predicted by the monomer-dimer-hexamer model at each ΔN6 concentration correlates with the % insoluble material (gray). Note that the fibril yield is low since fibrils cannot form when the monomer concentration falls significantly below the K_d for dimer formation (50 μM).

https://doi.org/10.7554/eLife.46574.025

Table 4

Reaction schemes, rate equations and rate constants for the fibril elongation models tested.

X represents the species that add onto the fibril ends.

https://doi.org/10.7554/eLife.46574.027

Module	Variant	Reaction scheme	Rate equations	Rate constants
Pre-polymerization	No Pre-polymerization (Monomer addition)	$X = X_{1}$	$\frac{d [X]}{d t} = \sum_{i = 2}^{N} {- k}_{e} F_{i - 1} X_{} + k_{e}^{'} F_{i}$	$k_{1} = k_{e}$ $k_{1}^{'} = k_{e}^{'},$
	Monomer conformational exchange	$X_{1} \overset{k_{1}}{\underset{k_{1}}{⇄}} X_{1}^{'}$ $X = X_{1}^{'}$	$\frac{d [X_{1}]}{d t} = - k_{1} X_{1} + k_{1}^{'} X_{1}^{'}$ $\frac{d [X]}{d t} = {\begin{matrix} k_{1} X_{1} - k_{1}^{'} X_{1}^{'} \\ + \sum_{i = 2}^{N} - k_{e} F_{i - 1} X + k_{e}^{'} F_{i} \end{matrix}$	$k_{1}, k_{e}$ $k_{1}^{'}, k_{e}^{'},$
	Dimer addition	$X_{1} + X_{1} \overset{k_{1}}{\underset{k_{1}}{⇄}} X_{2}$ $X = X_{2}$	$\frac{d [X_{1}]}{d t} = - 2 k_{1} X_{1} X_{1} + 2 k_{1}^{'} X_{2}$ $\frac{d [X]}{d t} = {\begin{matrix} k_{1} X_{1} X_{1} - 2 k_{1}^{'} X \\ + \sum_{i = 2}^{N} - k_{e} F_{i - 1} X + k_{e}^{'} F_{i} \end{matrix}$	$k_{1}, k_{e}$ $k_{1}^{'}, k_{e}^{'},$
	Hexamer addition	$X_{1} + X_{1} \overset{k_{1}}{\underset{k_{1}^{'}}{⇄}} X_{2}$ $X_{2} + X_{2} + X_{2} \overset{k_{1}}{\underset{k_{1}^{'}}{⇄}} X_{6}$ $X = X_{6}$	$\frac{d [X_{1}]}{d t} = - 2 k_{1} X_{1} X_{1} + 2 k_{1}^{'} X_{2}$ $\begin{array}{ll} \frac{d [X_{2}]}{d t} = k_{1} X_{1} X_{1} - k_{1}^{'} X_{2} - \\ 3 k_{2} X_{2} X_{2} X_{2} + 3 k_{2}^{'} X_{6} \end{array}$ $\frac{d [X]}{d t} = {\begin{matrix} k_{2} X_{2} X_{2} X_{2} - k_{2}^{'} X \\ + \sum_{i = 2}^{N} - k_{e} F_{i - 1} X + k_{e}^{'} F_{i} \end{matrix}$	$k_{1}, k_{2}, k_{e}$ , $k_{1}^{'}, k_{2}^{'}, k_{e}^{'},$
	Monomer-Dimer-Tetramer-Hexamer	$X_{1} + X_{1} \overset{k_{1}}{\underset{k_{1}^{'}}{⇄}} X_{2}$ $X_{2} + X_{2} \overset{k_{2}}{\underset{k_{2}^{'}}{⇄}} X_{4}$ $X_{2} + X_{4} \overset{k_{3}}{\underset{k_{3}^{'}}{⇄}} X_{6}$	$\frac{d [X_{1}]}{d t} = - 2 k_{1} X_{1} X_{1} + 2 k_{1}^{'} X_{2}$ $\begin{array}{ll} \frac{d [X_{2}]}{d t} = k_{1} X_{1} X_{1} - k_{1}^{'} X_{2} - \\ 2 k_{2} X_{2} X_{2} + 2 k_{2}^{'} X_{4} - \\ k_{3} X_{4} X_{2} + k_{3}^{'} X_{6} \end{array}$ $\begin{array}{ll} \frac{d [X_{4}]}{d t} = k_{2} X_{2} X_{2} - k_{2}^{'} X_{4} - \\ k_{3} X_{4} X_{2} + k_{3}^{'} X_{6} \end{array}$	$k_{1}, k_{2}, k_{3} k_{e}$ $k_{1}^{'}, k_{2}^{'}, {k_{3}^{'}, k}_{e}^{'},$
		$X = X_{6}$	$\frac{d [X]}{d t} = {\begin{matrix} k_{3} X_{4} X_{2} - k_{3}^{'} X \\ + \sum_{i = 2}^{N} - k_{e} F_{i - 1} X + k_{e}^{'} F_{i} \end{matrix}$
Polymerization		$\begin{array}{ll} X X \\ ↓ ↓ \\ F \overset{k_{e}}{\underset{k_{e}^{'}}{⇄}} F_{1} \overset{k_{e}}{\underset{k_{e}^{'}}{⇄}} F_{2...} F_{N} \\ ↓ ↓ \\ X X \end{array}$	$\frac{d [F]}{d t} = - k_{e} X_{} F_{1} + k_{e}^{'} F_{2}$ $\begin{array}{ll} \frac{d [F_{i}]}{d t} = k_{e} X_{} F_{i - 1} - k_{e}^{'} F_{i} - k_{e} \\ X_{} F_{i} + k_{e}^{'} F_{i + 1} 2 \leq i < N \end{array}$ $\frac{d [F_{N}]}{d t} = k_{e} X_{} F_{i - 1} - k_{e}^{'} F_{i}$

Discussion

Understanding the molecular details of oligomer formation is vital if we are to understand why proteins aggregate into amyloid and why different species have different toxicities (Iadanza et al., 2018a; Lu et al., 2013). Here, we present a general strategy, summarized in Figure 8—figure supplement 1, which allows the identification of oligomeric intermediates in amyloid assembly and enables their structural characterization. By combining the powers of NMR to detect lowly populated species in dynamic exchange, with complementary techniques such as AUC and cross-linking, oligomeric intermediates can be identified and structurally characterized in atomic detail. Importantly, to link these intermediates to the mechanism of aggregation, the derived affinities, stoichiometries and structural models can then be used to globally model the time course of fibril assembly. The strategy presented is not only applicable to protein aggregation, but to any weakly self-associating protein system. Given that the balance between monomers, dimers, higher molecular weight oligomers and fibrils could depend critically on the experimental conditions, including the pH, temperature, protein concentration, amount of seed added, buffer composition and ionic strength, the same protein, or a closely related protein variant, could assemble via different mechanism(s) under different conditions. Indeed, aggregation of many amyloidogenic proteins, including hβ₂m (Iadanza et al., 2018b), is known to result in polymorphic fibrils (Close et al., 2018; Fitzpatrick et al., 2017; Colvin et al., 2016; Zhang et al., 2019) that could extend via different mechanisms. The approach described here can distinguish between such different assembly pathways and may be able to shed light on the role of individual oligomeric species in aggregation and the origins of amyloid polymorphism.

Using the workflow derived, we show that elongation of ΔN6 amyloid seeds proceeds via a specifically organized hexamer (Figure 8). This finding contrasts with the more common view of monomer addition to fibril ends that has been observed for Aβ40/42 (Cohen et al., 2018), α-synuclein (Buell et al., 2014), huntingtin exon 1 (Vitalis et al., 2009) and for unfolded hβ₂m at pH 2.0 (Figure 1a) (Xue et al., 2008), while oligomers are thought to play critical roles in the primary/secondary nucleation phases of the assembly of these proteins (Cohen et al., 2018). By contrast with these initially disordered proteins, the monomeric precursor of ΔN6 assembly is structured, a scenario that accounts for more than 20 of the 70 human proteins known to cause amyloid disease (Sipe et al., 2016). Other amyloid precursors that are initially structured show an inability to self-seed (e.g. transthyretin; Hurshman et al., 2004), or display a non-classical dependence of the elongation rate on protein concentration (e.g. light chains; Blancas-Mejía et al., 2017). Whether these and other structured protein precursors assemble by a mechanism akin to that of ΔN6 could be answered by applying the integrated kinetic and structural approach described here to further examples of this set of proteins.

Figure 8 with 2 supplements see all

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Fibril formation in atomic detail.

Schematic representation of the mechanism of amyloid formation for ΔN6. During folding of hβ₂m, a highly dynamic intermediate with a flexible A strand is populated prior to formation of the native-like intermediate termed Ι_T, which has a native-like fold but contains a non-native *trans* X-Pro32 bond. The latter species is mimicked by ΔN6 and formed in vivo by proteolytic degradation of the WT protein (Bellotti et al., 1998). Only I_T/ΔN6 is primed for aggregation, while the intermediate with the flexible A strand is not able to assembly directly into amyloid (Karamanos et al., 2016). As reported here, ΔN6 forms elongated head-to-head dimers (upper image, center) which assemble into hexamers. Alternative dimers involving interactions between the ABED β-sheets in adjacent molecules formed at pH 8.2 (lower image, center) do not associate further into fibrils. Murine β₂m (mβ₂m) also interacts with ΔN6 at pH 6.2 to form head-to-head heterodimers. The subunit orientation is different in this heterodimer (Karamanos et al., 2014), occluding the hexamer interface and inhibiting assembly (central image). ΔN6 hexamers can elongate fibrillar seeds and show enhanced dynamics in the G strand which could represent the first step towards the major structural reorganization required to form the parallel in-register amyloid fold. How this final step occurs, however, remains to be solved.

https://doi.org/10.7554/eLife.46574.028

Here, we show that ΔN6 dimers and hexamers with well-defined interfaces involving the apical regions of the protein are required for fibrils to form under the conditions employed (Figure 8). By contrast, formation of other interfaces, such as that observed here for ΔN6 at pH 8.2 and the previously reported mβ₂m:ΔN6 heterodimer (Karamanos et al., 2014) are not able to assemble into amyloid fibrils (Figure 8). The arrangement of subunits in the ΔN6 dimer and hexamer observed here is different to that in a previously reported structure of a domain swapped ΔN6 dimer (Figure 8—figure supplement 2c). However, the G strand that is responsible for the domain swap is dynamic in the hexamer structure presented here, consistent with this edge β-strand being able to dissociate from the β-sandwich to form both structures. A variant (H13F) of hβ₂m has also been reported to form hexamers in the presence of Cu²⁺ ions (Calabrese et al., 2008) (Figure 8—figure supplement 2a,b). In the crystal structure of this species, the dimers and hexamers interact in a side-to-side or head-to-head manner to create a ring-like assembly, in marked contrast with the daisy-like organization of monomers in the ΔN6 hexamers shown in Figure 5. Real-time NMR studies of the folding of hβ₂m have also revealed protein concentration-dependent exchange-broadening in the apical loops of its transient folding intermediate I_T (Rennella et al., 2013), an observation that has been attributed to head-head oligomers, in agreement with the data presented here for ΔN6 which structurally mimics I_T (Eichner et al., 2011). The interfaces observed in the ΔN6 dimer and hexamer also differ from the canonical inter-sheet stacking between immunoglobulin domains in antibodies, suggesting that the structural features described here are specific to the dimers and hexamers involved in amyloid assembly. Taken together, the results show that β₂m can form different protein-protein interactions, only a specific set of which results in species capable of assembly into amyloid.

Although many studies have attributed the toxicity of amyloid to oligomeric species (Chiti and Dobson, 2017), we show here that the dimers and hexamers of ΔN6 are not cytotoxic, at least under the conditions employed, possibly because they are structured and bury substantial hydrophobic surface area. Interestingly, the oligomerization of ΔN6 has been linked to increased toxicity in Caenorhabditis elegans models (Diomede et al., 2012). Since amyloid formation can proceed via multiple pathways, it is possible that the cytotoxic species of ΔN6 formed in the C. elegans body wall muscle are different to those formed here in vitro. For several proteins, cytotoxicity has been ascribed to off-pathway oligomers that accumulate in the lag time of assembly, consistent with amyloid formation being protective for the cell (Bieschke et al., 2011). Interconversion between different forms of oligomers may also be required for cytotoxicity (Fusco et al., 2017; Cremades et al., 2012). Such a process could be compromised in the cross-linked species of ΔN6 used here.

In summary, by taking advantage of the power of NMR spectroscopy to visualize transient species, and combining these experiments with detailed analysis of the kinetic, thermodynamic and hydrodynamic properties of the aggregating ensemble of species, we have been able to determine an atomic structural model of two oligomeric species required for amyloid formation of ΔN6 at pH 6.2, and have generated a model that describes a potential mechanism of fibril elongation from these states. Our findings portray an assembly mechanism that is remarkably well-defined, involving the formation of specific protein-protein interfaces that are unique to the initiating stages of amyloid assembly. Substantial conformational changes have still to occur, however, for the hexameric intermediate to form the cross-β structure of amyloid. How this is achieved remains an open question, but could involve binding to the fibril ends and/or fibril surfaces. Most importantly, the results reveal a remarkable specificity to the early stages of ΔN6 amyloid assembly that involves the formation of well-defined oligomeric species via specific interfaces, the precise details of which determine the course of assembly. These findings suggest new avenues to combat disease by specific targeting of the early intermediates in the amyloid cascade which, at least for ΔN6, involve specific interactions between non-native, assembly-competent states.

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Dependence of the fibril elongation rate on the concentration of soluble protein.

ΔN6 oligomer formation.

Identification of interacting surfaces in ΔN6 dimers.

Structural models of ΔN6 dimers.

Agreement between experimental and back-calculated intermolecular PREs for the two dimer structures (dimer A and dimer B (see Figure 5—figure supplement 3).

Comparison of productive and inhibitory dimers.

Structural model of ΔN6 hexamers.

Agreement between experimental and back-calculated intermolecular distances for different hexamer structures.

Analysis of dimer and hexamer interfaces.

ΔN6 assembles into dimers and hexamers.

G-strand unfurling may occur upon hexamer formation.

The monomer-dimer-hexamer model describes the thermodynamics and kinetics of fibril elongation.

Reaction schemes, rate equations and rate constants for the fibril elongation models tested.

Fibril formation in atomic detail.

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Theodoros K Karamanos

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Matthew P Jackson

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Antonio N Calabrese

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Sophia C Goodchild

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Emma E Cawood

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Gary S Thompson

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Arnout P Kalverda

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Eric W Hewitt

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Sheena E Radford

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