If you don't remember your password, you can reset it by entering your email address and clicking the Reset Password button. You will then receive an email that contains a secure link for resetting your password
If the address matches a valid account an email will be sent to __email__ with instructions for resetting your password
Self-association of WT β2-microglobulin (WT-β2m) into amyloid fibrils is associated with the disorder dialysis related amyloidosis. In the familial variant D76N-β2m, the single amino acid substitution enhances the aggregation propensity of the protein dramatically and gives rise to a disorder that is independent of renal dysfunction. Numerous biophysical and structural studies on WT- and D76N-β2m have been performed in order to better understand the structure and dynamics of the native proteins and their different potentials to aggregate into amyloid. However, the structural properties of transient D76N-β2m oligomers and their role(s) in assembly remained uncharted. Here, we have utilized NMR methods, combined with photo-induced crosslinking, to detect, trap, and structurally characterize transient dimers of D76N-β2m. We show that the crosslinked D76N-β2m dimers have different structures from those previously characterized for the on-pathway dimers of ΔN6-β2m and are unable to assemble into amyloid. Instead, the crosslinked D76N-β2m dimers are potent inhibitors of amyloid formation, preventing primary nucleation and elongation/secondary nucleation when added in substoichiometric amounts with D76N-β2m monomers. The results highlight the specificity of early protein–protein interactions in amyloid formation and show how mapping these interfaces can inform new strategies to inhibit amyloid assembly.
). During its recycling process, the heavy chain of MHC-1 (bound to the cell membrane) is internalized by the host cell, whereas β2m monomers are released into the serum and subsequently eliminated through degradation by the kidneys (
). This variant has a dramatic effect on the properties of the protein, despite its location in a solvent-exposed loop (Fig. 1A). The D76N-β2m variant retains a native Ig fold (RMSD compared with WT-β2m of 0.59 Å (Cα atoms) (
). The self-assembly mechanisms of WT-β2m and ΔN6-β2m have been investigated previously and shown to be initiated by partial unfolding to form an intermediate in which the native cis Pro32 isomerizes to the trans form, known as IT (
). The structure of IT is mimicked by ΔN6-β2m, which also retains its all antiparallel β-stranded Ig structure, has a trans Pro32, a repacked hydrophobic core and destabilization of the hydrogen bonds across its Ig fold (
). There is also no information currently available about oligomers formed early in D76N-β2m amyloid formation, including how they relate to the on-pathway dimers described for ΔN6-β2m, or the inhibitory dimers formed between ΔN6-β2m and the nonaggregating murine β2m (mβ2m) (
To investigate dimeric species formed from D76N-β2m, we here employ NMR and crosslinking mass spectrometry (MS) to identify, trap, and characterize dimers formed during D76N-β2m self-assembly. NMR is a versatile technique that allows exploration of the structure, dynamics, and the interactome of biomolecules in solution (
). NMR in combination with paramagnetic relaxation enhancement (PRE) is ideal for identifying and structurally characterizing transient intermediates in biomolecular interactions with atomistic resolution, and herein, we use this approach to investigate transient intermolecular dimers formed for D76N-β2m (
). A number of paramagnetic probes are available to explore these transient states and S-(1-oxyl-2,2,5,5-tetramethyl-2,5-dihydro-1H-pyrrol-3-yl) methyl methanesulfonothioate (MTSL) is the most widely utilized in the field due to its relative ease of coupling to the target protein via native or introduced Cys residues (
), allowing us also to investigate their role in aggregation. The results revealed an interaction interface for D76N-β2m dimers that is distinct from that observed previously for the on-pathway dimers of ΔN6-β2m. The crosslinked D76N-β2m dimers are not able to assemble into amyloid and are potent inhibitors of D76N-β2m assembly. The results highlight the significance of investigating short-lived intermolecular species in solution to better understand the roles they play in biological mechanisms (
). They also uncover an efficient strategy to inhibit amyloid assembly by targeting early dimers as kinetic traps of the assembly process.
Exploring transient dimers of D76N-β2m in solution
To initiate our studies of D76N-β2m assembly, we measured the rate of aggregation of the protein into amyloid using thioflavin T (ThT) fluorescence and compared the behavior of the protein with that of WT-β2m. The experiments were performed at pH 6.2 in 25 mM sodium phosphate buffer supplemented with 115 mM NaCl—conditions identical to those used previously to determine the aggregation mechanism of ΔN6-β2m (
), D76N-β2m aggregates into amyloid rapidly under these conditions, with a Thalf (time to reach 50% of the maximum ThT signal) of 17.3 ± 2.4 h and reaching a plateau after ∼20 h, while WT-β2m did not form ThT-positive fibrils within the 42 h timescale of this experiment (Fig. 1B). Consistent with this, negative stain electron micrographs of the samples at the end of the experiment clearly displayed the presence of amyloid fibrils for D76N-β2m, while no detectable fibrils were observed for WT-β2m (Fig. 1B (inset)). Given the very similar native/IT structures of the proteins (
), the dramatic differences in their amyloid potential suggests that early oligomers formed from the two proteins might differ significantly in conformation.
To explore the nature of transient noncovalent oligomeric species formed from D76N-β2m, 2D 1H-15N heteronuclear single quantum coherence (HNHSQC) NMR spectra were acquired at protein concentrations spanning 25 μM to 200 μM (in 25 mM sodium phosphate, 115 mM NaCl, pH 6.2, 25 °C). Surprisingly, and in marked contrast to the behavior of ΔN6-β2m (
), no significant changes in HN chemical shift or peak intensity were observed as a function of protein concentration over this range (Fig. 1C), indicating that dimeric or higher order oligomeric species, if formed, are not detected under these conditions.
Given that oligomers of D76N-β2m were not detected in the experiments aforementioned, we next used NMR PREs to explore whether lowly populated, transient oligomeric states are formed that could play a role in governing the aggregation of D76N-β2m. NMR PREs are ideal to detect transiently formed intermolecular species, as they are able to detect species that populate between 0.2% to 0.5% of the total conformers in solution, especially when they are in dynamic equilibrium on a fast NMR timescale (
). Accordingly, MTSL was covalently coupled to a cysteine residue introduced into different locations on the surface of 14ND76N-β2m (NMR inactive, bait). The variants created were S20C (AB loop), S33C (BC loop), S57C (DE loop), and S88C (FG loop) (Fig. 1A). The new Cys residues were inserted into solvent-exposed loops in the protein, in order to avoid alterations in the structure of the native protein. Sequence changes in the major aggregation-prone region of the protein (residues 60–66), shown previously to be the only region to affect the rate of aggregation into amyloid using mutational scanning, were also avoided (
). Indeed, control experiments in which each Cys-containing protein was modified with (diamagnetic) MTSL showed that all variants are natively folded at the temperature of the NMR PRE experiments (25 °C), despite being thermodynamically destabilized (Figs. S1 and S2). Each MTSL-labeled protein at natural nitrogen abundance (14N) was then mixed 1:1 with 15ND76N-β2m (NMR active, target) and intermolecular PREs resulting from transient noncovalent interactions were measured for each variant (using 100 μM of each protein) (see schematics in Fig. 2). No significant intermolecular PREs were observed when 14N-D76N-β2m/S20C-MTSL was mixed with 15ND76N-β2m (Fig. S3), suggesting that if dimers form, they do not involve this region of the protein. By contrast, when 14N-D76N-β2m/S33C-MTSL, 14N-D76N-β2m/S57C-MTSL, or 14N-D76N-β2m/S88C-MTSL were each mixed individually with 15ND76N-β2m PREs were observed, with the strongest involving three regions in 15ND76N-β2m: a) residues ∼33 to 37 (the BC loop), b) residues 52 to 65 (D/E strands and DE loop) and c) residues 83 to 88 (FG loop) (Figs. 2 and S4). This suggests that one or more oligomer(s), probabilistically dimers, are transiently formed that involve interactions at these interfaces. In addition, residues 7 to 9 and 11 to 15 in the target protein also show a significant PRE effect (Ip/Id < 0.8), especially when the 14N-D76N-β2m/S57C-MTSL and 14N-D76N-β2m/S88C-MTSL probes were used (Fig. 2).
Mapping the dimer interface using crosslinking
To characterize the interface formed in the transient D76N-β2m dimers in more detail, we exploited a tag-transfer photo-crosslinking strategy, using a cleavable MTS-diazirine heterobifunctional crosslinker developed previously (Fig. 3A) (
). This crosslinker is first attached to a specific Cys on the surface of one of the interacting partners in a protein–protein interaction, with the attached diazirine moiety allowing subsequent rapid and nonspecific crosslinks to heavy atoms, which lie within <10 Å of the Cα of the Cys modified with the crosslinker (
). Subsequent reduction and alkylation of the crosslinked species then enables identification of crosslinked sites via their covalent modification (addition of 145.06 Da) (Fig. 3A). Combined with the use of an LED illuminating system (
). D76N-β2m-S57C was chosen for this analysis since residue 57 is intimately involved in the dimer interface, according to the magnitude of NMR PREs observed from this residue (Fig. 2). Accordingly, D76N-β2m-S57C was modified with MTS-diazirine, and the modified protein (200 μM) was then crosslinked for 30 s before analysis of the products using SDS-PAGE (Fig. 3B). As predicted, proteins migrating with the mass expected of dimers (∼24 kDa) resulted from the crosslinking. Addition of DTT (20 mM) subsequent to crosslinking resulted in a lack of dimers and reappearance of monomers, consistent with successful tag transfer (Fig. 3B). The dimeric and monomeric crosslinked species were then purified using gel filtration (Fig. 3C), subjected to in-solution digestion with trypsin (after treatment with DTT), and identification of crosslinked products using LC-MS/MS. Comparison of the two samples allowed intramolecular versus intermolecular crosslinks to be identified. The results of this analysis (Figs. 3D, S5, Tables S1 and S2) yielded intermolecular crosslinks from residue 57 on one monomer to residues in the dimer interface located in the AB loop (E15), BC loop (His31, Ile35 and Glu36), and the D-, E-, and F-strands (His51, Ser52, Ser55, Tyr63, Leu64, Val82 and Asn83) in an adjacent protein. These sites are consistent with the intermolecular PRE-NMR results presented previously, suggesting that these regions of the protein form the epicenter of the dimer interface.
Model of D76N-β2m dimers
In order to extract structural information on the transient dimers of D76N-β2m, rigid-body in silico docking was performed in explicit solvent using the NMR PRE and crosslinking data from S33C, S57C, and S88C as constraints (Table S3). All constraints were treated as nonambiguous. Data from S20C were not used for deriving the structural model, as no significant NMR PREs resulted from this site. The results revealed that three clusters (formed of four representative structures) satisfied the experimental constraints (Tables S4 and S5). In all of these clusters, the monomers involved in the model dimers adopt a ‘top-to-side’ orientation with respect to each other. The dimerization interface is dominated by residues in the BC loop (Ile35 and Glu36), D-strand (His51, Ser52, Asp53, Leu54, Ser55, Phe56 and Asp59), and E-strand (Leu54 and Leu55) from one monomer and BC (Ser33), DE (Ser57), and FG (Ser88) loops from the other monomer (Fig. 4). These dimers differ significantly from the on-pathway ‘head to head’ homodimers of ΔN6-β2m (
). Notably, the distance of Cys20 to the dimer interface in the models generated is >30 Å and hence is outside the sensitivity of the PRE, consistent with a lack of significant PREs from this site (Fig. S3).
D76N-β2m crosslinked dimers are effective inhibitors of amyloid formation
Previous studies have shown that dimers of WT-β2m crosslinked by disulfide bonds at residue 33 (Fig. S6) accelerate amyloid formation of the normally aggregation-resilient protein (at least in the presence of the cosolvent trifluoroethanol) (
). We therefore analyzed how the purified crosslinked dimers (XL-Ds) of D76N-β2m/S57C affect amyloid formation, by incubating the dimers either alone or in the presence of monomeric D76N-β2m and monitoring amyloid formation versus time via ThT fluorescence (20 μM monomeric protein in 25 mM sodium phosphate, 115 mM NaCl, pH 6.2). Under these conditions, D76N-β2m forms fibrils rapidly (Thalf of ca. 17.3 ± 2.4 h, with the maximal ThT signal after ∼20 h) (Fig. 5). Surprisingly, fibril formation was arrested over the total incubation time when 10% (w/w) crosslinked dimer was added (Fig. 5A), with significant retardation in the kinetics of fibril growth (0.8-fold increase in Thalf) occurring when as little as 1.25% (w/w) dimer was added (Fig. S7, B and D). Notably, the XL-Ds alone were unable to assemble into ThT-positive amyloid (Fig. S7, B and D) but did aggregate into material able to be pelleted by centrifugation (Fig. S8) when incubated alone at high concentration, presumably since the conformational changes required to form a cross-β structure are not possible from the crosslinked state. In marked contrast with the behavior of the XL-Ds on fibril formation, addition of the crosslinked monomers marginally increased the rate of fibril growth, demonstrating that the inhibition is specific to the dimeric species (Fig. S7, A and C).
To confirm the inhibition of fibril growth by the XL-Ds, insoluble and soluble material after 120 h of incubation of each sample was separated by centrifugation and the percent insoluble material quantitatively determined by SDS-PAGE (Experimental procedures). The results showed that D76N-β2m incubated without XL-Ds was found exclusively in the insoluble fraction after 120 h of incubation (Fig. S8, A and B). D76N-β2m incubated with the intramolecularly crosslinked monomers also showed no decrease in aggregate yield. However, in samples that contained >5 % (w/w) XL-Ds, little, if any, insoluble material could be detected (Figs. 5B, S8,A, and B), demonstrating the unique ability of the XL-Ds to arrest amyloid assembly.
Defining the inhibition mechanism of D76N/57C-β2m dimers
To examine in more detail the mechanism by which XL-Ds are able to arrest amyloid assembly of D76N-β2m, the soluble products of the inhibition reaction were examined using HNHSQC NMR. For these experiments, monomeric 15N-labeled D76N-β2m was mixed with 10 % (w/w) 14N-labeled XL-Ds and a HNHSQC NMR spectrum immediately recorded. The sample was then removed from the NMR tube, placed in a 96-well plate, and incubated under identical conditions to those used for the ThT kinetic analysis. After 38 h, the sample was removed and a HNHSQC NMR spectrum again recorded. The results revealed no effect on the chemical shifts and only a minor increase in linewidth (∼12% reduction across all resonances) of the monomeric 15N-labeled protein when the samples were immediately mixed, consistent with a weak interaction between the 15N-monomers and XL-Ds (Fig. S9). The chemical shifts and linewidths of the 15N-labeled protein product at the end of the incubation were also unchanged relative to the starting material (Fig. 5C). These results demonstrate that the XL-Ds kinetically inhibit aggregation by transient binding to monomers, rather than by stabilizing unproductive larger oligomeric species, as has been observed with small molecule inhibitors of β2m amyloid formation (
To explore whether the XL-Ds are also able to kinetically inhibit fibril elongation, fibril growth was measured in the presence of seeds formed from preformed D76N-β2m fibrils (Experimental procedures). The Thalf of fibril formation is decreased (∼2.5- and 2.7-fold) in the presence of 3% or 10% (w/w) fibril seeds, respectively, consistent with elongation and/or secondary nucleation mechanisms enhancing the rate of fibril formation (Fig. 6). When supplemented with 10% (w/w) XL-Ds, fibril growth in the presence of seeds was again inhibited (relative to the same reactions in the absence of XL-Ds), with the magnitude of inhibition depending on amount of seeds added (0.8- and 1.6-fold increase in Thalf in the presence of 3 % and 10% (w/w) fibril seeds and 10% (w/w) crosslinked dimer, respectively, relative to the unseeded reactions plus dimer). Clearly, therefore, XL-Ds are able to retard seeded fibril growth, consistent with binding to monomers and/or the fibrils themselves.
To determine whether the XL-Ds bind to preformed fibrils, a pelleting assay was performed in which preformed fibril seeds were incubated with XL-Ds and binding measured by centrifugation to separate fibril bound versus free XL-Ds (Experimental Procedures). The results (Fig. S10A) revealed that the XL-Ds do not interact stably with fibrils. Finally, surface plasmon resonance experiments, in which D76N-β2m monomers were immobilized onto the chip and XL-Ds passed over the surface, were used to confirm that the XL-Ds do indeed bind to monomeric D76N-β2m (Experimental procedures) (Fig. S10B).
Together with the data presented previously, the results indicate that the XL-Ds are able to inhibit primary nucleation— similar behavior to the action of some molecular chaperones (
). Additionally, elongation and/or secondary nucleation processes involved in the fibrillation of D76N-β2m are also inhibited by XL-Ds. Such kinetic inhibition presumably results from transient binding of the XL-Ds to D76N-β2m monomers, reducing the effective concentration of assembly competent monomers for participation in fibril growth events. Such a mechanism would also explain the effectiveness of the XL-Ds to inhibit assembly when added in substoichiometric amounts. However, other mechanisms of action, including interaction with oligomeric species not visible in the experiments used here, cannot be ruled out.
Protein self-assembly is a ubiquitous, and often complex, process involved in an array of functions in biology (
), slowing amyloid formation by kinetically inhibiting the assembly steps of nucleation, fibril elongation, and/or secondary nucleation are more tractable. Strategies to inhibit fibril assembly using small molecules (
Here, we describe the ability of stabilized dimers of an aggregating protein to inhibit its own self-assembly. We show that the mechanism of action of the inhibitory dimers is by transient binding to assembly competent monomers, which competes with the protein–protein interactions required for amyloid assembly. While populated only transiently and rarely in solution, the formation of D76N-β2m dimers presumably contributes to the observed rate of amyloid formation, with XL-Ds slowing assembly by binding to monomers and also by disfavoring the structural transitions needed to generate the amyloid fold. Analysis of the assembly mechanism of ΔN6-β2m into amyloid using NMR PREs, combined with detailed kinetic analysis using ThT fluorescence and other methods, identified head-to-head elongated dimers as essential precursors of amyloid formation, with the rate of amyloid formation dependent on the formation of these key initiating species (
). Interestingly, the on-pathway ΔN6-β2m dimers contain some interacting residues in common with those of the D76N-β2m inhibitory dimers identified here (namely the BC loop and partially the DE loop) but are structurally distinct in that the D76N-β2m dimers are asymmetric and involve mainly the D-strand in one monomer and BC-, DE-, and FG-loops on the other monomer (Figs. 4 and S6). These structural differences presumably define the diametrically opposed outcomes of assembly of these two dimer folds. Inhibitory dimers formed between ΔN6-β2m and mβ2m have also been reported previously (
). Inhibition of amyloid formation of ΔN6-β2m has also been achieved by covalent attachment of a small molecule fragment to residue 52 in the D strand, which results in the formation of off-pathway tetramers (
). Given this information, the inhibitory potential of the D76N-β2m dimers determined here can also be rationalized, at least in part, since they sequester interactions known to be essential for amyloid formation in their ΔN6-β2m counterparts. While further work will be needed to determine the structure of on-pathway dimers and higher order oligomers of D76N-β2m, the finding that regions involved in early assembly are shared in D76N-β2m and ΔN6-β2m suggests that the two proteins assemble via similar pathways, despite their involvement in distinct diseases that affect different individuals (with/without renal dysfunction), and result in amyloid deposition in different regions of the body (
A second striking finding of the results presented here is the efficient nature of the inhibition caused by D76N-β2m XL-Ds, with complete arrest of assembly over 120 h being achieved with ratios of D76N-β2m:dimers of only 1:0.1 (w/w). We showed that inhibition occurs via kinetic competition, with dimer–-monomer interactions competing with productive monomer–monomer and monomer–fibril interactions that define primary nucleation, secondary nucleation, and fibril elongation. While the affinity of these different interactions for D76N-β2m remain unknown, previous studies of ΔN6-β2m:mβ2m heterodimers and ΔN6-β2m homodimers show Kds ranging from 68 to 494 μM (
), similar in magnitude with the weak intermolecular D76N-β2m interaction observed here. These results highlight the ability of weak binding to effect inhibition in these kinetically controlled assembly processes. Discovery of new amyloid inhibitors, therefore, will not necessarily require tight binding, as is the general case in ligand discovery strategies in medicinal chemistry (
). Instead, as we portray here, mapping the interacting interfaces in early oligomers, using NMR PREs and chemical crosslinking can provide an excellent starting point to generate strategies to disrupt amyloid fibril formation.
By contrast with the inhibition of amyloid formation shown here for D76N-β2m XL-Ds, crosslinking β2m or other proteins with disulfide bonds has been shown to accelerate amyloid formation, dependent on the location of the disulfide bond introduced (
). By contrast with the highly defined crosslinks involving disulfide bond formation, crosslinking via diazirines provides the opportunity of mapping more promiscuous or transient interactions. As we showed previously, it is also important to ensure that the covalent addition of new chemical moieties onto a protein sequence does not affect its structure or aggregation, as changes in solubility, local or global stability, and the inherent amyloid propensity of the sequence can all result from posttranslational or other chemical modifications. We demonstrate the structural integrity of labeled proteins herein using CD, NMR, and thermal stability measurements. Importantly, previous studies of 56 variants of D76N-β2m, obtained using random mutagenesis and screening for sequences with altered aggregation potential, have revealed that the presence of a single region, spanning residues 60 to 66, alongside Asn at residue 76, is essential for amyloid formation, with mutations elsewhere in the protein having little effect (
). Importantly, the inhibitory dimers identified here do not involve this critical region (which forms the E-strand in the native structure (Fig. 1A)).
Finally, our work has important implications for the amyloid field, by highlighting the potential of stabilizing early assembly intermediates as routes to inhibit amyloid formation. Such findings may provide new avenues to combat disease, by stabilizing lowly populated species of the same system and reusing their stable equivalents as possible therapeutic reagents. At a more fundamental level, the results highlight the specificity of the early interactions that drive β2m amyloid assembly, with different dimers able to promote or prevent amyloid formation in this family of proteins, dependent on the interactions made and their stability.
The gene encoding WT-β2m and D76N-β2m are inserted in pINK expression vector (
). Monomeric protein was purified using size-exclusion chromatography, and the purity and fidelity of the sequence were determined using 12% SDS-PAGE and electrospray ionization MS (ESI-MS), respectively. ESI-MS also confirmed the formation of the disulfide bond in all samples. Monomeric proteins were divided into aliquots and frozen using liquid nitrogen and stored at −20 °C.
The S20C, S33C, S57C, and S88C variants of D76N-β2m were produced by site-directed mutagenesis and purified as aforementioned, except that an additional anion-exchange (Q-Sepharose) chromatography step under denaturing conditions (25 mM Tris–HCl, 8 M urea, pH 8.0) was included at the beginning of the purification. All Cys variants were refolded by flash dilution (1:10) in 25 mM Tris–HCl, 300 mM NaCl, 500 mM arginine, pH 8.0. The sample was then dialyzed three times in 25 mM Tris–HCl, pH 8.0.
D76N-β2m at natural nitrogen abundance (14N-D76N-β2m) or labeled with 15N (15N-D76N-β2m) were thawed at room temperature (RT) and then further purified by gel filtration. Monomeric fractions from size-exclusion chromatography (Superdex75 PG-26/600) were collected and immediately used for fibrillation assays under the following conditions: 20 μM of D76N-β2m (in the absence or presence of various concentrations of 14N-D76N-β2m-S57C–crosslinked dimer or monomer), 10 μM of ThT in 25 mM sodium phosphate, 115 mM NaCl, pH 6.2, in a 96-well plate (clear flat bottom, Corning 3631). The plates were sealed with a plastic film, incubated at 37 °C for 38, 90, or 120 h, depending on the conditions to be evaluated, with shaking at 600 rpm (Fluostar Omega plate reader). Amyloid formation was monitored by ThT fluorescence (excitation at 440 nm and emission at 475 nm). Fibril seeds were created by treating the samples at the end of a fibrillation reaction for 1 min of bath sonication at fixed frequency (Ultrawave instrument, model U100H).
Aggregate yield was determine using Tris-Tricine-SDS PAGE (5:1 ratio of acrylamide and bis-acrylamide, respectively) following separation of insoluble/soluble material by centrifugation (23,000g for 10 min). Pellets were treated with 8 M urea for 30 min in the same buffer before analysis by SDS-PAGE. In the case of the soluble material, no urea was added prior to SDS-PAGE. Gels were stained with instant blue solution (Abcam: ab119211), imaged, and band quantified using densitometry with UVITEC transilluminator (Q9Aliance).
After the fibrillation, samples were collected and stored at 4 °C for analysis by electron microscopy. Ten microliters of fibril solution was added to a carbon-coated grid (home produced, without glow discharge) for 2 min, followed by staining with 2% (w/v) uranyl acetate. The grids were washed twice with water; electron micrographs were acquired on T12 microscope (Gatan US4000 equipped with 4k CCD camera and 120 keV Lab6 electron source) at the Electron Microscopy facility in the Astbury Biostructure Laboratory University of Leeds.
HNHSQC NMR and data processing
HNHSQC spectra, obtained using eight scans, 2048 and 164 complex points in the direct and indirect dimensions, respectively, were acquired on Bruker 750 MHz or 950 MHz NMR spectrometers, equipped with TCI (1H, 2H, 13C, 15N channels) cryoprobes (Bruker Avance III HD console, acquisition software, Topspin 3.2). All spectra were recorded at 25 °C, pH 6.2, in 25 mM sodium phosphate, 115 mM NaCl supplemented with 3% (v/v) D2O using protein concentrations ranging from 20 to 200 μM. NMRPipe (
). NMR assignments were taken from BMRB (access code: 50302), which were obtained under identical conditions to those employed here.
14N-D76N-β2m Cys variants (S20C, S33C, S57C, and S88C) were thawed at RT. The samples were incubated with 20 mM DTT for 30 min at RT. DTT was then removed from solution using a 5 ml centrifugal desalting column (Zebra 7K/MWCO, 89882), which was equilibrated with 50 mM sodium phosphate and 150 mM sodium chloride, pH 7, prior to use. Eluted protein fractions were quantified and the protein concentration was adjusted to 200 μM. The sample was then mixed with 2 mM (final concentration) of MTSL (CAS: SC-208677) in 500 mM guanidine HCl. The reaction mixture was incubated for 4 h at RT and then centrifuged at 44,000g for 1 h to remove any protein aggregates. β2m-Cys-MTSL labeled variants were isolated by gel filtration (Superdex75 PG-26/600), and monomeric fractions were collected and stored at 4 °C for further use. The mass of the modified proteins was confirmed by ESI-MS recorded using a Xevo QToF G2-XS mass spectrometer (Waters UK) operated in positive ion mode. The MS spectra show complete labeling of the proteins (observed masses of D76N-β2m of 12,059.46 ± 0.28 Da, 12,059.78 ± 0.11 Da for S57C-MTSL and S88C-MTSL, respectively (expected mass of 12060 Da).
Changes in the stability of D76N-β2m as result of insertion of cysteine and MTSL at residues 20, 33, 57, and 88 were evaluated by thermal denaturation using far UV CD (Chirascan Plus). For these experiments, 20 μM of protein, dissolved in 25 mM sodium phosphate, 115 mM NaCl, pH 6.2 was used. The temperature ramp was from 20 °C to 80 °C, at a rate of 1 °C/min. A far UV CD spectrum (195–260 nm) of each protein was acquired at each temperature during the ramp. The data were fitted to two state equilibrium in CDPal software (https://github.com/PINT-NMR/CDpal) (
Where E is the mean residue ellipticity, ΔHm is the change of enthalpy at the midpoint of denaturation (Tm), ΔCp is the change of heat capacity, R the universal gas constant, and T is the temperature (Kelvin).
NMR PRE experiments
To measure intermolecular PREs, 100 μM 15N-D76N-β2m was mixed with 100 μM of 14N-D76N-S/C-MTSL variants and HNHSQC spectra were recorded (paramagnetic conditions) at 25 °C in 25 mM sodium phosphate, 115 mM NaCl, pH 6.2. Ascorbic acid dissolved in the same buffer at pH 6.2 was then added to a final concentration of 1 mM and left at RT for 1 h. A second HNHSQC spectrum was then collected (diamagnetic conditions). The NMR data were processed as described previously. Note that no significant change in pH is observed after addition of ascorbic acid (<0.1 units) indicated by measuring the pH of the solution and analysis of the chemical shifts (HN) of His residues. Analysis of chemical shift perturbations of the diamagnetic samples were determined using the equation:
These experiments showed no significant change in structure of each monomer, compared with unmodified D76N-β2m (Fig. S2). The extent of the intermolecular PRE effect was evaluated by calculating the ratio (Ip/Id) of resonance intensity of the paramagnetic (Ip) and diamagnetic (Id) samples. The conformational ensembles of MTSL shown in different figures were generated using the MTSLWizard module in PyMol (
14N-D76N-β2m-S57C (200 μM protein) was labeled with the MTS-diazirine photo-crosslinker (Fig. 3A) using a protocol similar to that of MTSL labeling. The reaction was left to progress for 4 h at RT. Then, MTS-diazirine–labeled monomeric protein was recovered by gel filtration. The mass of the modified proteins was confirmed by ESI-MS (11,905 ± 0.37 Da, expected mass for the labeled monomer 11,905 Da).
Photo-crosslinking was achieved at RT using 200 μM 14N-D76N-β2m-S57C-diazirine in 25 mM sodium phosphate and 115 mM NaCl, pH 6.2. The sample was photo-crosslinked for 30 s at 365 nm using a home-built LED illumination device (
). Crosslinked species were isolated by gel filtration (Superdex75 Increase 10/300) equilibrated with 25 mM sodium phosphate and 115 mM NaCl, pH 6.2. Monomeric and dimeric crosslinked conformers were verified by 12% Tris-Tricine-SDS-PAGE (5:1 ratio, acrylamide and bis-acrylamide, respectively). Finally, samples were stored at 4 °C.
Identification of crosslinked sites using trypsin digestion and MS
The isolated pure crosslinked dimer and monomer samples were first reduced by incubation with 10 mM DTT at 57 °C for 1 h with shaking. The samples were left to cool to RT, followed by alkylation with 55 mM iodoacetic acid at RT for 45 min in the dark with shaking. Then, trypsin (20 ng/μl in 25 mM ammonium bicarbonate) was added in a 1:50 ratio (w/w of protease:β2m) and the mixture was incubated at 37 °C for 18 h with shaking. The reaction was stopped by adding 5 μl of 1% (v/v) TFA. The digested peptides were purified by reverse phase chromatography on Sep-Pak C18 column. Peptides were eluted from the column with 500 μl 50% (v/v) acetonitrile and 0.1% (v/v) formic acid. Finally, the peptides were evaporated to dryness and reconstituted in 20 μl 0.1% (v/v) aqueous TFA prior to MS analysis.
The peptide-containing solution (3 μl) was injected into a reverse-phase in house-packed C18) capillary column (75 μm × 200 mm) and separated by gradient elution of 5% to 95% (v/v) acetonitrile with 0.1% (v/v) formic acid at a flow rate of 250 nl/min. The separated peptides were eluted directly from the column and then infused into an Orbitrap Velos (ThermoFisher Scientific) mass spectrometer using an electrospray capillary voltage of 2.7 kV. The mass spectrometer was operated in positive ion mode. Data acquisition was performed in data-dependent acqusition mode and fragmentation was performed by using ion-trap. Up to 20 most intense ions per precursor scan were selected for MS/MS. Dynamic exclusion of 30 s was used. Peptide MS/MS data processing and modification localization were performed by using PEAKS Studio X+ (Bioinformatic Solutions Inc).
Modeling dimers of D76N-β2m using docking
Two molecules of D76N-β2m (Protein Data Bank: 4FXL) (
) were used as input for flexible molecular docking. One molecule was modified by the insertion of three cysteines at positions 33, 57, and 88. PRE (HN) and crosslinking data were then used as experimental restraints: a radius of 7.5 Å (covering the distance between the sulfhydryl group and the nitroxy group in MTSL) or 12 Å (covering the distance between sulfhydryl [bait molecule] and nitrogen [target molecule] atoms for crosslinking) was employed. The second molecule was not modified. PRE restraints were organized into three groups according their Ip/Id ratio value (Ip/Id 0.2–0.6, 0.05–0.2 and < 0.05 [resonance undetectable]) (Table S3). 4FXL-Cys (modified monomer, A) and 4FXL (unmodified monomer, B) were used as input structures for molecular docking using HADDOCK2.4, allowing flexibility in all segments (
). The docking process was performed in three sequential steps: rigid body, semiflexible, and finally, water refinement. Two-hundred structures were analyzed and then clustered in three families (with four representative members). Sampling and clustering parameters during docking are summarized in Table S4, and the number of restraints satisfied/not satisfied by the final lowest energy structures are shown in Table S5.
Binding of XL-Ds to fibril seeds
D76N-β2m fibrils were freshly generated in 25 mM sodium phosphate and 115 mM NaCl, pH 6.2, as described previously. Fibrils were then recovered by centrifugation at 13,000 rpm (benchtop microfuge) and their concentration adjusted to 20 μM (monomer equivalent concentration) in the same buffer. XL-Ds (2 μM) were added and incubated for 1 h at RT by centrifugation (13,000 rpm, 10 min), and the XL-D remaining in solution were quantified by measuring the absorbance at 280 nm.
Binding of XL-D to monomers
Surface plasmon resonance was used to assay monomer:XL-D interactions. For this assay, a cysteine was inserted into D76N-β2m immediately after the initiating N-terminal methionine (Cys0). The sample was then labeled with EZ-Link-Maleimide-PEG11-Biotin (ThermoFisher: 21911) using a protocol similar to that used for labeling with MTSL or diazirine, as described previously. Binding of XL-D to monomers was monitored using a Biacore T200 instrument (Cytiva). D76N-C0-Biotin-β2m was immobilized onto a streptavidin sensor chip (Cytiva-BR100398) and XL-D was passed over the immobilized monomer. All samples were in 25 mM sodium phosphate and 115 mM NaCl, pH 6.2. Nine concentrations of XL-D were tested between 5.8 nM and 1.5 μM (Fig. S10B). The association of XL-D was monitored for 60 s, followed by 360 s of dissociation at a flow rate of 30 μl/min at 25 °C.
The authors declare there they have conflict of interest with the contents of this article.
We thank members of our laboratories for helpful discussions and Nasir Khan for his excellent technical support. We also thank Arnout Kalverda for his NMR technical and scientific support and Iain Manfield for assistance with SPR. We acknowledge funding from Wellcome ( 109984 ) (Roberto Maya-Martinez, Nicolas Guthertz). We are also grateful to the University of Leeds and Wellcome ( 094232 ) for funding for the Chirascan CD spectrometer and for access to the Astbury Biostructure Laboratory EM and BioNMR Facilities. The SPR was funded by Wellcome ( 062164/Z/00/Z ). Waters MClass UPLC and Xevo G2-XS QTOF MS instruments were funded by the BBSRC ( BB/M012573/1 ). This research was funded in whole, or in part, by Wellcome . For the purpose of open access, the author has applied a CC BY public copyright license to any Author Accepted Manuscript version arising from this submission.
S. E. R. conceptualization; R. M. M., Y. X., N. G., T. K. K., and M. W. methodology; R. M. M., Y. X., N. G., and M. W. investigation; F. S., A. L. B., and S. E. R. resources; R. M. M. and S. E. R. writing–original draft; Y. X., N. G., T. K. K., M. W., F. S., and A. L. B. writing–review & editing; Y. X. visualization; F. S., A. L. B., and S. E. R. supervision; F. S., A. L. B., and S. E. R. funding acquisition.
Funding and additional information
SER holds a Royal Society Professorial Fellowship (RSRP∖R1∖211057).