Covalent Cross-Linking within Supramolecular Peptide Structures

Jun 29, 2012 - Photo-induced covalent cross-linking for the analysis of biomolecular interactions. George W. Preston , Andrew J. Wilson. Chemical Soci...
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Covalent Cross-Linking within Supramolecular Peptide Structures George W. Preston,†,‡,§ Sheena E. Radford,‡,§ Alison E. Ashcroft,‡,§ and Andrew J. Wilson*,†,‡ †

School of Chemistry, ‡Astbury Centre for Structural Molecular Biology, and §Institute of Molecular and Cellular Biology, Faculty of Biological Sciences, University of Leeds, Woodhouse Lane, Leeds LS2 9JT, U.K. S Supporting Information *

ABSTRACT: β-Sheet peptide nanostructures (e.g., amyloid fibrils) are recognized as important entities in biological systems and as functional materials in their own right. Their unique physical properties and architectural complexity, however, present a challenge for structure determination at atomic resolution. Covalent cross-linking and mass spectrometry are appealing methods for this endeavor because, potentially, a large amount of information can be extracted from a small sample in a single experiment. Previously, we described preliminary studies on the use of a photoreactive diazirine-containing amino acid to crosslink peptide monomers in nanostructures, together with the integrated separation and analysis of the products using ion mobility spectrometry coupled to conventional mass spectrometry. Here, a pH-switchable system (Aβ16−22, a sequence from the amyloid-β peptide) was used to examine cross-linking chemistry in morphologically distinct supramolecular structures containing, or entirely composed of, diazirine-functionalized peptides. We examine the relationship between cross-linker chemistry, covalent cross-links (identified using chemical derivatization and tandem mass spectrometry), and noncovalent structure, and report differences in the site of cross-linking that can only be explained by supramolecular templating. The results demonstrate the applicability of the approach for obtaining structural restraints in ordered supramolecular assemblies, provided that a considered evaluation of the cross-linked products is undertaken.

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wing to their unique physical properties and association with disease, β-sheet peptide nanostructures (e.g., amyloid fibrils) are of interest to chemists, physicists, biologists, and materials scientists alike.1−3 Amyloid fibrils are frequently investigated because they are implicated in the etiologies of numerous debilitating and terminal diseases,1 while more generally, filamentous peptide nanostructures are studied because of their mechanical properties4 and kinetically controlled assembly.5,6 The molecular architecture of amyloid-like β-sheet nanostructures is known to be assembled from the cross-β motif,7 in which β-strands are aligned perpendicular to the long axis of the filament (Figure 1a). This substructure is a fundamental determinant of the nanostructures’ physical and morphological characteristics, i.e., noncrystalline, unbranched, large, and insoluble. For small peptides, structural information about β-sheet nanostructures has been obtained from crystallography,8−10 while for isotope or spin-labeled peptides, information has been obtained using nuclear magnetic resonance (NMR) spectroscopy11−14 or electron paramagnetic resonance (EPR) spectroscopy.15−17 Though highly informative, these methods rely on elements of conformational homogeneity in the analytes, which are often heterogeneous mixtures.18 Covalent cross-linking, with analysis using mass spectrometry (MS) and tandem mass spectrometry (MS/MS),19−22 is a powerful method for analyzing biomolecular interactions. The approach involves conversion of a reversible noncovalent contact into a stable, irreversible linkage via the formation of one or more covalent bonds. Accordingly, structural © 2012 American Chemical Society

Figure 1. (a) Cartoon representation of the cross-β motif. β-Sheets stack as a laminate with a dry sheet−sheet interface. (b) Covalent cross-linking within a cross-β structure. Upon irradiation, a photoreactive side chain (yellow) could form a cross-link (red) to the side chain of a neighboring molecule.

information is embedded in a covalent form that can be isolated, then identified by MS. Cross-linking is suitable for studying the internal organization of nanostructures because of their apparent rigidity: the supramolecular structures of prion Received: May 16, 2012 Accepted: June 29, 2012 Published: June 29, 2012 6790

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aggregates and amyloid-β (Aβ) fibrils, for example, have been studied using a homobifunctional cross-linking reagent23 and disulfide cross-linking,24 respectively. Although useful, these methods do, however, rely on functional group proximity, i.e., monomers may need to be engineered in order to place the relevant groups within cross-linking distance. Moreover, slow reaction kinetics coupled with dynamic motions in the protein could lead to cross-links that do not accurately reflect native structure. The PICUP (photoinduced cross-linking of unmodified proteins) approach developed by Fancy and Kodadek25 aimed to address such concerns by generating reactive protein radicals in situ. Bitan and Teplow subsequently employed this method to evaluate size distributions in populations of amyloidogenic oligomers,26 while others have used it to cross-link mature aggregates.27 Despite being clearly advantageous with respect to preserving native structure, the use of a diffusible activator in PICUP has implications for the kinetics of the overall process and for the extraction of structural information buried in tightly packed cross-β cores. It is also notable that various other studies have made structural inferences by evaluating the assembly competency of peptide constructs containing predefined cross-links.28−30 Site-specific incorporation of a photoactivated cross-linker can circumvent many of the above limitations, as it offers the advantage of rapid and indiscriminate cross-linking from a defined position in a polypeptide. We31 and others32 have developed photoreactive constructs for cross-linking experiments on peptide nanostructures. In a generalized experiment (Figure 1b), irradiation generates reactive sites within the crossβ framework, via which cross-links then form. Here, we describe the use of the carbene-generating 3-aryl-3(trifluoromethyl)diazirine33 (TFMD) group (cf. other photolabeling strategies34−37) which was chosen on the basis of its relatively small size, coupled with the comparably rapid, indiscriminate reactivity of the carbene.38,39 Previous work31 demonstrated that the complex mixture of products arising from photolysis could be effectively resolved using ion mobility spectrometry−mass spectrometry (IMS− MS) and that cross-linked peptides could be isolated for MS/ MS analysis from within a heterogeneous mixture, even in the presence of species with the same mass-to-charge (m/z) ratio. The present study aimed to investigate how the supramolecular organization of a peptide β-sheet nanostructure could be encoded within a cross-linked peptide via the unique chemistry of the TFMD group. In order to do this, TFMD was placed at different positions in a self-assembling peptide known to display pH-dependent morphological plasticity. We hypothesized that the effects of these changes could be manifested in both the positions of the cross-links and the morphologies of the resulting nanostructures. Information was then extracted from photolysis products using a combination of chemical derivatization, IMS−MS, and MS/MS experiments. The data show a clear effect of supramolecular templating on the formation of covalent bonds between peptides but, interestingly, a lack of correlation between morphology and cross-link position. The results reveal that, with careful attention to both cross-linker chemistry and analytical methodology, a wealth of data with which to analyze noncovalent contacts in supramolecular structures can be obtained.

Supporting Information. Briefly, nanostructures were assembled by adding phosphate buffer to a concentrated solution of peptide in dimethyl sulfoxide, then cross-linked by irradiation under a 6 W ultraviolet (UV) lamp at 365 nm. Cross-linked nanostructures were isolated by centrifugation, then disaggregated in neat hexafluoroisopropyl alcohol (HFIP). IMS−MS and IMS−MS/MS data were acquired on a Waters Synapt G1 high-definition mass spectrometer, for which the IMS buffer gas was N2. For the MS/MS experiments, fragmentation was by collision-induced dissociation, and the collision gas was Ar. For routine characterization, other mass spectra were acquired using a Bruker HCT Ultra ion trap mass spectrometer as part of a liquid chromatography−MS system. Further details of MS instrumentation and methods are provided in the Supporting Information.

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RESULTS Selection and Preparation of Photoreactive Aβ16−22 Constructs. Aβ16−22 (Ac−KLVFFAE−NH2; Chart 1), a selfassembling heptapeptide,40 whose sequence originates from full-length Aβ, was chosen for this analysis because its diphenylalanine motif presented opportunities for conservative incorporation of the TFMD moiety via TFMD-Phe.41,42 Accordingly, the TFMD-functionalized peptides Aβ16−22− F19* and Aβ16−22−F20* (numbering according to full-length Aβ) were prepared by solid-phase peptide synthesis (see Chart 1 and the Supporting Information). Chart 1. Structure of Aβ16−22 and Its TFMD-Functionalized Analoguesa

a

Numbering corresponds to the full-length Aβ peptide.

Peptide Assembly Reveals a Complex Interplay of Structural Features and Nanostructure Morphology. The ability of Aβ16−22 to form morphologically distinct nanostructures, including fibrils, tapes, and nanotubes, is welldocumented.40,43−45 Specifically of interest were the fibril and nanotube morphologies because they are associated with different underlying β-sheet structures40,43 and because these different structures are generated in a pH-dependent fashion: neutral pH conditions promote the assembly of fibrils composed of antiparallel, in-register β-sheets,40 whereas acidic conditions promote the assembly of nanotubes composed of antiparallel, out-of-register (one-residue-shifted) β-sheets.43 Under the assumption that morphology would serve as a useful indicator of molecular alignment, attempts were made to

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EXPERIMENTAL SECTION Full details of all reagents, preparative chemistry, analytical methods, and various control experiments can be found in the 6791

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Figure 2. Transmission electron micrographs showing nanostructures assembled from Aβ16−22 and its TFMD-functionalized analogues under different pH conditions: (a) Aβ16−22, pH 7.0; (b) Aβ16−22−F19*, pH 7.0; (c) Aβ16−22−F20*, pH 7.0; (d) Aβ16−22 and Aβ16−22−F20* coassembled from a 4:1 molar mixture at pH 7.0; (e) Aβ16−22, pH 2.0; (f) Aβ16−22−F19*, pH 2.0; (g) Aβ16−22−F20*, pH 2.0; (h) Aβ16−22 and Aβ16−22−F20* coassembled from a 4:1 molar mixture at pH 7.0. Scale bars represent 100 nm.

(see the Experimental Section and Supporting Information). Upon irradiation, the diazirine groups are consumed to generate the reactive carbene and a linear diazoisomer;49 the former then reacts further, while the latter is considered relatively inert. Importantly, TEM showed that irradiation did not qualitatively alter the morphology of nanostructures containing TFMD groups (Supporting Information Figure S5). Reaction products derived from the nanostructures were isolated by centrifugation, then depolymerized using neat HFIP. Experiments demonstrating the effectiveness of HFIP for this purpose are detailed in the Supporting Information (see Figure S-6 and associated content). We also provide experimental evidence that the products obtained in this manner do not derive from solution-phase cross-linking reactions during photolysis (see Supporting Information Figures S-7, S-14, and associated content). Following depolymerization, samples were introduced into the mass spectrometer, where IMS−MS typically resolved three packets of ions: singly charged monomers (longest tD), doubly charged dimers (intermediate tD), and triply charged trimers (shortest tD). Figure 3a shows the IMS−MS data for the Aβ16−22−F20* homopolymer assembled at pH 7.0. Spectra for the other homopolymer samples are provided in the Supporting Information (Figures S-9 and S-10). Qualitative analysis of the spectra suggested the presence of intermolecular bonds (dimers and trimers, Figure 3) and that the TFMD-functionalized monomers had undergone numerous other transformations with both positive and negative changes in mass (Δm). In unphotolysed control samples, the TFMD groups were largely intact (Supporting Information Figure S-8), indicating that the observed transformations had not occurred in solution prior to assembly and irradiation. Irradiation of nanostructures assembled from mixed peptide stocks generated covalent heterodimers (Figure 3b and Supporting Information Figure S-11), which indicated that peptides Aβ16−22 and Aβ16−22−F20* had coassembled rather than having partitioned into separate nanostructures. Also, in terms of the underlying chemistry, detection of a heterodimer provided clear evidence that cross-linking was not dependent on the cross-reaction of two TFMD groups (or derivatives thereof). ESI-IMS−MS Allows the Identity of Cross-Linked Peptides to be Determined. Overall, our analysis of

replicate wild-type nanostructures with TFMD-functionalized peptides. Initially, the ability of the photoreactive peptides to form homomeric aggregates under the two different pH conditions was tested. Then, in an effort to further preserve elements of native structure, the photoreactive peptides were diluted (1:4) with wild-type Aβ16−22. The self-assembled morphologies of peptide nanostructures formed in these experiments are summarized in Figure 2; a detailed account of the various assembly preparations and the results obtained is given as a Supplementary Results section in the Supporting Information. Briefly, transmission electron microscopy (TEM) showed that Aβ16−22 alone formed pH-dependent nanostructures as has been reported by others,40,43,45 i.e., filaments at neutral pH and nanotubes at pH 2.0 (Figure 2, parts a and e). Interestingly, Aβ16−22−F19* displayed similar behavior at neutral pH (Figure 2b; see also the previous work31) but maintained a filamentous morphology at pH 2.0 (Figure 2f). This suggested that the underlying molecular structure was resisting a switch of alignment. When the position of the TFMD group was changed to position-20 (Aβ16−22−F20*), aspects of pH-dependent morphology reappeared, namely, the formation of wider tapes at acidic pH (compare Figure 2, parts c and g). Of all the experiments attempted, pH dependency was best preserved in a preparation where Aβ16−22−F20* was coassembled with wild-type Aβ16−22: dense mats of filaments were observed at neutral pH (Figure 2d), whereas nanotubes and ribbons were formed at pH 2.0 (see Figure 2h and the Supporting Information). Cross-linking experiments (see later) indicated that these nanostructures were true coassemblies. IMS−MS Permits Isolation of Cross-Linked Peptides Encoding Supramolecular Information. Previously, it was demonstrated that IMS−MS could be used to resolve complex mixtures of photolysis products, including mixtures of oligomeric cross-linked peptides, on account of their m/z values and their physical shapes in a single, rapid experiment.31 IMS involves the separation of gas-phase ions according to their velocities in the presence of a buffer gas under the influence of a weak electric field. The time taken (drift time, tD) is dependent on the ion’s charge and its rotationally averaged collision cross section due to interactions with the buffer gas.46−48 In our experiments, each (mixture of) peptide(s) was assembled into the relevant nanostructure and cross-linked by UV irradiation 6792

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could be envisaged: these can be broadly classified as relatively labile (e.g., imines,50 esters,51 imidates51) and relatively stable (e.g., bonds to aliphatic or aromatic carbon atoms36). This distinction is important because (a) some labile cross-links may only be evident from degradation products, and (b) cross-links of intermediate stability can be subjected to informative derivatization reactions (see later). Experiments to elucidate various aspects of cross-linker reactivity (e.g., toward carboxylic acids) are described in the Supporting Information. Assignments on the basis of Δm could mostly be rationalized by reactions of the carbene with solvents and buffer components (Figure 3 and Supporting Information Table S-1).52 Particularly noteworthy, however, was the detection of a covalent HFIP adduct, which could only have formed postphotolysis. This led us to explore reactivity of the aforementioned diazoisomer,49 whose absence in the IMS−MS spectra suggested some form of degradation. Full details of these experimental considerations can be found in the Supporting Information. MS/MS Sequencing of Cross-Linked Adducts Provides Insight into the Structural Organization Within Aβ16−22 Aggregates. Sequencing of peptides by MS/MS is based on fragmentation at consecutive points (e.g., amide bonds) along the backbone53 to give a sequence of residue masses. For localizing cross-links by MS/MS, the tandem mass spectrum of singly protonated Aβ16−22 (Supporting Information Figure S15a) was used as a reference as all of its amide bonds should be labile. The spectrum is dominated by b and y ions from single amide cleavages (Roepstorff−Fohlman−Biemann nomenclature,53−55 see Supporting Information Figures S-16 and S-17) although we also observed some evidence of b ion sequence scrambling56 (see the supplementary discussion in the Supporting Information). Importantly, comparison of the fragmentation signatures of insertion products from nanostructures in which the same peptide had been assembled at different pHs (Supporting Information Figures S-18, S-19, and S-20) indicated that their covalent structures were identical. This pointed to a similar mode of noncovalent packing under both pH conditions. Similarly, the fragmentation patterns for photolysis products of dif ferent peptides assembled at the same pH were sequence-dependent, indicating that crosslinking is not influenced by the reactive preferences of the carbene and is determined by supramolecular templating. Intramolecular Adducts. To identify cross-links, each covalently linked structure can be considered as a stable core with free peptide chains attached to it (≤2n chains for a covalent n-mer). Hence, intramolecular cross-links should preserve covalent connectivity between cross-linked residues, even when the backbone is fragmented (Figure 4a). The location of cross-links can thus be determined from the fragmentation pattern. Note, however, that the degree of interpretation can be limited by cross-link heterogeneity: a hypothetical case in considered in Figure 4d, and this phenomenon is evidenced in the fragmentation of cyclized Aβ16−22−F19* (from the fibrils assembled at pH 2.0). In this case, a full, uninterrupted b series (b8 → b2; Figure 4b) was observed. Given that aryl-substituted carbenes are not prone to intramolecular rearrangement, 39 the difference in mass corresponding to the cross-linker (F*) is unlikely to be a real measurement of residue mass; rather, it is probably a consequence of there being two or more superimposed fragmentation series. The fragmentation pattern for cyclized Aβ16−22−F20*, when compared to that of Aβ16−22 or cyclized Aβ16−22−F19*, was

Figure 3. IMS−MS data for disassembled nanostructure photolysates: (a) Aβ16−22−F20* assembled at pH 7.0; (b) Aβ16−22 and Aβ16−22− F20* coassembled from a 4:1 molar mixture at pH 7.0. An m/z scale is given on the right-hand vertical axis of each IMS−MS plot; a full mass spectrum is shown on the left-hand vertical axis. Assigned photolysis products are depicted in cartoon form. Chemical structures are given for relevant transformations of the TFMD group; these were assigned manually based on known reactions of the carbene (ref 36). WT denotes the wild-type Aβ16−22 peptide. Drift time (tD) was not calibrated.

TFMD chemistry is represented by the manually assigned structures in Figure 3 and in extensive supporting information (see the Supporting Information). Our findings can be summarized thus: the TFMD group reacts within nanostructures to generate a substantial array of reaction products, and although complex, this array is rich in information about the environment of the cross-linker prephotolysis. Intermolecular and intramolecular cross-links (see Figure 3) were the primary source of information from which noncovalent structure was inferred: the former contain information about the alignment of peptides with respect to one another; the latter are indicative of monomer conformation. Considering the functional groups of the Aβ16−22 peptide and the known reactivity of TFMD-derived carbenes (i.e., insertion reactions36), various types of cross-link 6793

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Intermolecular Adducts. In our analyses of cross-linked peptide dimers, we refer to the chain from which the cross-link originated as α and the peptide to which it formed as β (see Supporting Information Figure S-17 and cf. nomenclature of Schilling et al.57). In addition to the usual considerations associated with MS/MS of covalently linked pairs of peptides (e.g., double fragmentations; see the Supporting Information),21,57 we addressed specific challenges associated with carbene-mediated cross-linking and sequence homology: first, as for cross-linked monomers, potential isomerism in the parent ions leading to mixtures of fragmentation series; second, sequence homology between peptide chains preventing particular backbone fragment ions from being assigned as α or β. Only when fragmentation occurs beyond the position of the cross-linker (N-terminal of the cross-linker for a, b, and c ions; C-terminal for x, y, and z ions) can the resulting ion be unambiguously assigned. Cross-linked dimers extracted from fibrils of Aβ16−22−F19* displayed fragmentation patterns similar to those reported in the earlier work (see Supporting Information Figures S-19 and S-21a), which were indicative of intermolecular cross-linking between F19* residues. For Aβ16−22−F20*, we focused on the doubly charged dimer containing two cross-links (m/z 974; see Figure 3). When the crude photolysate was probed for ester cross-links using NPA (Supporting Information Figure S-22), m/z 974 appeared to undergo conversion to a species with m/z 1004, corresponding to a dimeric aminolysis product. This product has retained its intermolecular cross-link, but its ester cross-link has been broken to give an alcohol-functionalized cross-linker derivative and a modified Glu residue (see structures in Figure 4e). Sequencing revealed that both the modified Glu residue and the cross-linker derivative were within the β-chain, meaning that the ester cross-link had been intramolecular (Figure 5 and Supporting Information Figure S21b). Further fragmentation of the β-chain then localized the intermolecular cross-link to within the N-terminal Ac−KL region (Figure 5). It was also noted that, in the heterodimeric cross-linked structure obtained from the mixed peptide experiment, the amide bond between Lys-16 and Leu-17 also appeared to fragment. In this structure, therefore, the intermolecular cross-link can be localized more precisely to within Ac−K (Supporting Information Figure S-21c).

Figure 4. Sequencing of cyclized monomers: (a) cyclized monomers are considered as stabilized macrocycles with labile polyamide chains attached to them; (b) sequence ions of cyclized peptide Aβ16−22− F19* (from fibrils assembled at pH 2.0); (c) sequence ions of cyclized peptide Aβ16−22−F20* (from nanotapes assembled at pH 2.0). Open triangles denote parent [M + H]+ ions. F* denotes the cross-linker. (d) The occurrence of different intramolecular cross-links in mixtures can, in theory, mask positional information from an MS/MS experiment. (e) Sequencing of the aminolysis product obtained via treatment of cyclized Aβ16−22−F20* with NPA. Spectra with full assignments are provided in Figure S-15 of the Supporting Information.

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DISCUSSION The architectural complexity and physical properties of β-sheet peptide nanostructures often restrict the quality and quantity of structural data that can be acquired from these systems. Using photoreactive constructs, the present study aimed to determine the extent to which covalent cross-linking within nanoscale objects was reflective of supramolecular structure. To address our aim, a switchable three-dimensional molecular framework was employed, consisting of β-sheets with pH-dependent register. Thus, it was envisaged that changing the sequence and/or solution pH would modulate supramolecular structure to yield distinctive cross-links. The subtle effects of modifications to monomer structure on self-assembly were observed as perturbations to the system’s pH dependency. Results of self-assembly experiments revealed that this pH dependency could indeed be recapitulated, but that this required (a) judicious placement of the cross-linker within the primary structure of the monomer, and (b) attention to the formulation of the peptide stock. This result is perhaps unsurprising, given the sensitivity of other Aβ16−22 derivatives

noticeably different (Figure 4c), mainly because b6 and b7 were strongly suppressed or absent. This indicated that the amides on either side of Ala-21 were protected, suggesting that peptide Aβ16−22−F20* had cross-linked internally to Glu-22. Suspecting an ester cross-link, we treated the crude disaggregated product mixture with n-propylamine (NPA), which was found to ring-open lactones with a diagnostic 59 Da increase in mass. When the photolysate from Aβ16−22−F20* aggregates obtained at pH 2.0 was treated with NPA, a decrease in the relative intensity of the intramolecularly cross-linked product was observed, accompanied by appearance of an intense peak corresponding to an aminolysis product which, following MS/ MS analysis (Figure 4e), pinpointed the F20*−E22 cross-link as an ester (see Supporting Information Chart S-4). 6794

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Figure 5. Fragment ions from a doubly cross-linked dimer of Aβ16−22−F20* post NPA treatment. The cartoons illustrate how both cross-links in this structure were localized. α denotes the peptide from which the intermolecular cross-link originated; β denotes the peptide to which the intermolecular cross-link formed. Chemical structures of the modified residues are shown where necessary.

reported recently by others45,58 and the finding that register switching in the underlying β-sheet is dependent on βbranching of the amino acid side chain at position 18.59 For analyses of the covalent structures of cross-linked peptides, three aspects of methodology proved particularly valuable: the resolving power of IMS−MS, chemical derivatization, and rigorous unbiased analysis of the MS/MS data. Previously,31 IMS−MS alone was used to separate photolysis products in the gas phase. In the present study, we added a derivatization step prior to IMS−MS, which had the potential to resolve otherwise intractable mixtures of species with the same combinations of m/z and tD. Accordingly, derivatization with NPA functioned not only as a probe for labile ester crosslinks but also as a label with which to differentiate parts of a molecule on the basis of mass. This label allowed us to restrain both the intra- and the intermolecular cross-links within a doubly cross-linked dimer. We found that the fragmentation patterns of cross-linked peptides were dependent on the position at which the crosslinker was placed (i.e., whether the photoreactive monomer was Aβ16−22−F19* or Aβ16−22−F20*), but not on the assembly conditions (pH 2.0 vs pH 7.0). Sequencing of cross-linked dimers revealed that Aβ16−22−F20* had formed intermolecular cross-links to the N-terminus, while Aβ16−22−F19* had done so within the central hydrophobic core of the sequence. This observation provides robust evidence of supramolecular templating within the nanostructures, because chemical affinity alone would have directed cross-linking to the same site in both systems. Analyses of covalent structure also allow conclusions to be drawn as to the arrangement of peptides within the nanostructures. The intermolecular cross-links (F19* to F19* and F20* to K16) are consistent with the noncovalent interactions within an in-register antiparallel β-sheet (Figure 6a), but not within the alternative register-shifted β-sheet (Figure 6b). Additionally, the intramolecular cross-link (F20* to E22), if taken to be formed in a templated fashion, is consistent the β-strand conformation. We also considered that a photoreactive group at the sheet−sheet interface of a laminate could cross-link peptides in different β-sheets (Figure 6c). Using molecular dynamics simulations, Mehta et al. generated models of β-sheet laminates constructed from both the inregister and out-of-register β-sheets (Figure 6, parts a and b,

Figure 6. Cartoons illustrating β-sheet peptide alignment and interfacial side chain interactions. (a) In-register and (b) out-ofregister β-sheet structures from solid-state NMR studies (refs 40 and 43). (c) Basis for sheet−sheet interfacial cross-linking (cf. Figure 1b). The cross-linker (yellow) interdigitates with other side chains (gray). This organization is translated into a cross-link (red).

respectively).43 We note that the intermolecular cross-link between position-20 and Lys-16 observed for Aβ16−22−F20* is remarkably consistent with these authors’ model for Aβ16−22 in fibrils at near-neutral pH and agrees much less well with the model for nanotubes at acidic pH. In particular, the relative positions of F20* and K16 side chains in both models would prevent them from contacting each other at the sheet−sheet interfaces. Intriguingly, the pattern of cross-links observed in preparations of Aβ16−22−F20* was invariant with morphological switching. There are two possible explanations for this observation: first, that the resolution of cross-link formation is too low to distinguish between two different β-sheet registers; second, that the structures under analysis contain the same β-sheet unit whose higher order packing has some dependency on the solution conditions. Of these possibilities, the former is contradicted by our sequencing data, which suggest a good degree of accuracy. The latter is much more plausible, especially in light of a recent solid-state NMR study60 6795

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(10) Colletier, J.-P.; Laganowsky, A.; Landau, M.; Zhao, M.; Soriaga, A. B.; Goldschmidt, L.; Flot, D.; Cascio, D.; Sawaya, M. R.; Eisenberg, D. Proc. Natl. Acad. Sci. U.S.A. 2011, 108, 16938−16943. (11) Tycko, R. Methods Enzymol. 2006, 413, 103−122. (12) Petkova, A. T.; Ishii, Y.; Balbach, J. J.; Antzutkin, O. N.; Leapman, R. D.; Delaglio, F.; Tycko, R. Proc. Natl. Acad. Sci. U.S.A. 2002, 99, 16742−16747. (13) Jaroniec, C. P.; MacPhee, C. E.; Bajaj, V. S.; McMahon, M. T.; Dobson, C. M.; Griffin, R. G. Proc. Natl. Acad. Sci. U.S.A. 2004, 101, 711−716. (14) van der Wel, P. C. A.; Lewandowski, J. R.; Griffin, R. G. J. Am. Chem. Soc. 2007, 129, 5117−5130. (15) Margittai, M.; Langen, R. Q. Rev. Biophys. 2008, 41, 265−297. (16) Torok, M.; Milton, S.; Kayed, R.; Wu, P.; McIntire, T.; Glabe, C. G.; Langen, R. J. Biol. Chem. 2002, 277, 40810−40815. (17) Cobb, N. J.; Sonnichsen, F. D.; Mchaourab, H.; Surewicz, W. K. Proc. Natl. Acad. Sci. U.S.A. 2007, 104, 18946−18951. (18) Lewandowski, J. R.; van der Wel, P. C. A.; Rigney, M.; Grigorieff, N.; Griffin, R. G. J. Am. Chem. Soc. 2011, 133, 14686− 14698. (19) Sinz, A. Mass Spectrom. Rev. 2006, 25, 663−682. (20) Trnka, M. J.; Burlingame, A. L. Mol. Cell. Proteomics 2010, 9, 2306−2317. (21) Petrotchenko, E. V.; Borchers, C. H. Mass Spectrom. Rev. 2010, 29, 862−876. (22) Schermann, S. M.; Simmons, D. A.; Konermann, L. Expert Rev. Proteomics 2005, 2, 475−485. (23) Onisko, B.; Fernandez, E. G.; Freire, M. L.; Schwarz, A.; Baier, M.; Camina, F.; Garcia, J. R.; Rodriguez-Segade Villamarn, S.; Requena, J. R. Biochemistry 2005, 44, 10100−10109. (24) Shivaprasad, S.; Wetzel, R. Biochemistry 2004, 43, 15310−15317. (25) Fancy, D. A.; Kodadek, T. Proc. Natl. Acad. Sci. U.S.A. 1999, 96, 6020−6024. (26) Bitan, G.; Teplow, D. B. Acc. Chem. Res. 2004, 37, 357−364. (27) Piening, N.; Weber, P.; Hogen, T.; Beekes, M.; Kretzschmar, H.; Giese, A. Amyloid 2006, 13, 67−77. (28) Sciarretta, K. L.; Gordon, D. J.; Petkova, A. T.; Tycko, R.; Meredith, S. C. Biochemistry 2005, 44, 6003−6014. (29) Krishnan, R.; Lindquist, S. L. Nature 2005, 435, 765−772. (30) Sandberg, A.; Luheshi, L. M.; Sollvander, S.; Pereira de Barros, T.; Macao, B.; Knowles, T. P. J.; Biverstal, H.; Lendel, C.; EkholmPetterson, F.; Dubnovitsky, A.; Lannfelt, L.; Dobson, C. M.; Hard, T. Proc. Natl. Acad. Sci. U.S.A. 2010, 107, 15595−15600. (31) Smith, D. P.; Anderson, J.; Plante, J.; Ashcroft, A. E.; Radford, S. E.; Wilson, A. J.; Parker, M. J. Chem. Commun. 2008, 5728−5730. (32) Egnaczyk, G. F.; Greis, K. D.; Stimson, E. R.; Maggio, J. E. Biochemistry 2001, 40, 11706−11714. (33) Brunner, J.; Senn, H.; Richards, F. M. J. Biol. Chem. 1980, 255, 3313−3318. (34) Kotzybahibert, F.; Kapfer, I.; Goeldner, M. Angew. Chem., Int. Ed. 1995, 34, 1296−1312. (35) Dorman, G.; Prestwich, G. D. Biochemistry 1994, 33, 5661− 5673. (36) Brunner, J. Annu. Rev. Biochem. 1993, 62, 483−514. (37) Tamura, T.; Tsukiji, S.; Hamachi, I. J. Am. Chem. Soc. 2012, 134, 2216−2226. (38) Blencowe, A.; Hayes, W. Soft Matter 2005, 1, 178−205. (39) Platz, M. S. Acc. Chem. Res. 1995, 28, 487−492. (40) Balbach, J. J.; Ishii, Y.; Antzutkin, O. N.; Leapman, R. D.; Rizzo, N. W.; Dyda, F.; Reed, J.; Tycko, R. Biochemistry 2000, 39, 13748− 13759. (41) Shih, L. B.; Bayley, H. Anal. Biochem. 1985, 144, 132−141. (42) Fishwick, C. W. G.; Sanderson, J. M.; Findlay, J. B. C. Tetrahedron Lett. 1994, 35, 4611−4614. (43) Mehta, A. K.; Lu, K.; Childers, W. S.; Liang, Y.; Dublin, S. N.; Dong, J. J.; Snyder, J. P.; Pingali, S. V.; Thiyagarajan, P.; Lynn, D. G. J. Am. Chem. Soc. 2008, 130, 9829−9835. (44) Tao, K.; Wang, J.; Zhou, P.; Wang, C.; Xu, H.; Zhao, X.; Lu, J. R. Langmuir 2011, 27, 2723−2730.

on a related peptide system (AAKLVFF): this peptide was found to assemble into filaments or nanotubes in a solventdependent fashion, and the resulting morphologically distinct structures were found to contain the same β-sheet building block.

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CONCLUSION In summary, a versatile peptide system was used to perform a detailed examination of how supramolecular contacts in a unique molecular environment can be translated into covalent bonds by the TFMD cross-linker. Despite complexities associated with the chemistry of TFMD, the results demonstrate the amenability of this cross-linker for investigating amyloid-like structures. The potential for rapid activation, along with the fast reaction of the carbene intermediate, would allow such studies to answer important questions about aggregation pathways (e.g., when does the cross-β fold first develop?) and how conformational properties are correlated with toxicity in amyloid-related oligomers.61,62 The experiments described here demonstrate the applicability of photoinduced cross-linking to answer such questions.

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ASSOCIATED CONTENT

S Supporting Information *

Additional information as noted in text. This material is available free of charge via the Internet at http://pubs.acs.org.

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AUTHOR INFORMATION

Corresponding Author

*E-mail: [email protected]. Notes

The authors declare no competing financial interest.

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ACKNOWLEDGMENTS We thank Dr. Martin Parker and Dr. Bruce Turnbull for helpful discussions, the European Research Council [ERC-StG240324] and the British Biotechnology and Biological Sciences Research Council (BBSRC) for a studentship for G.W.P. The Synapt HDMS was purchased using funds from the BBSRC Equipment Initiative (BB/E012558/1).

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REFERENCES

(1) Chiti, F.; Dobson, C. M. Annu. Rev. Biochem. 2006, 75, 333−366. (2) Cherny, I.; Gazit, E. Angew. Chem., Int. Ed. 2008, 47, 4062−4069. (3) Knowles, T. P. J.; Buehler, M. J. Nat. Nanotechnol. 2011, 6, 469− 479. (4) Knowles, T. P.; Fitzpatrick, A. W.; Meehan, S.; Mott, H. R.; Vendruscolo, M.; Dobson, C. M.; Welland, M. E. Science 2007, 318, 1900−1903. (5) Baldwin, A. J.; Knowles, T. P. J.; Tartaglia, G. G.; Fitzpatrick, A. W.; Devlin, G. L.; Shammas, S. L.; Waudby, C. A.; Mossuto, M. F.; Meehan, S.; Gras, S. L.; Christodoulou, J.; Anthony-Cahill, S. J.; Barker, P. D.; Vendruscolo, M.; Dobson, C. M. J. Am. Chem. Soc. 2011, 133, 14160−14163. (6) Miller, Y.; Ma, B.; Nussinov, R. Chem. Rev. 2010, 110, 4820− 4838. (7) Geddes, A. J.; Parker, K. D.; Atkins, E. D. T.; Beighton, E. J. Mol. Biol. 1968, 32, 343−358. (8) Nelson, R.; Sawaya, M. R.; Balbirnie, M.; Madsen, A. O.; Riekel, C.; Grothe, R.; Eisenberg, D. Nature 2005, 435, 773−778. (9) Sawaya, M. R.; Sambashivan, S.; Nelson, R.; Ivanova, M. I.; Sievers, S. A.; Apostol, M. I.; Thompson, M. J.; Balbirnie, M.; Wiltzius, J. J. W.; McFarlane, H. T.; Madsen, A. O.; Riekel, C.; Eisenberg, D. Nature 2007, 447, 453−457. 6796

dx.doi.org/10.1021/ac301198c | Anal. Chem. 2012, 84, 6790−6797

Analytical Chemistry

Article

(45) Senguen, F. T.; Lee, N. R.; Gu, X. F.; Ryan, D. M.; Doran, T. M.; Anderson, E. A.; Nilsson, B. L. Mol. Biosyst. 2011, 7, 486−496. (46) Uetrecht, C.; Rose, R. J.; van Duijn, E.; Lorenzen, K.; Heck, A. J. R. Chem. Soc. Rev. 2010, 39, 1633−1655. (47) Kanu, A. B.; Dwivedi, P.; Tam, M.; Matz, L.; Hill, H. H. J. Mass Spectrom. 2008, 43, 1−22. (48) Bohrer, B. C.; Mererbloom, S. I.; Koeniger, S. L.; Hilderbrand, A. E.; Clemmer, D. E. Annu. Rev. Anal. Chem. 2008, 1, 293−327. (49) Hashimoto, M.; Hatanaka, Y. Anal. Biochem. 2006, 348, 154− 156. (50) Platz, M.; Admasu, A. S.; Kwiatkowski, S.; Crocker, P. J.; Imai, N.; Watt, D. S. Bioconjugate Chem. 1991, 2, 337−341. (51) Bayley, H. Photogenerated Reagents in Biochemistry and Molecular Biology; Elsevier: Amsterdam, The Netherlands, 1983; Vol. 12. (52) Kanoh, N.; Nakamura, T.; Honda, K.; Yamakoshi, H.; Iwabuchi, Y.; Osada, H. Tetrahedron 2008, 64, 5692−5698. (53) Biemann, K. Annu. Rev. Biochem. 1992, 61, 977−1010. (54) Roepstorff, P.; Fohlman, J. Biomed. Mass Spectrom. 1984, 11, 601−601. (55) Biemann, K. Biomed. Environ. Mass Spectrom. 1988, 16, 99−111. (56) Van Stipdonk, M.; Bleiholder, C.; Osburn, S.; Williams, T. D.; Suhai, S.; Harrison, A. G.; Paizs, B. J. Am. Chem. Soc. 2008, 130, 17774−17789. (57) Schilling, B.; Row, R. H.; Gibson, B. W.; Guo, X.; Young, M. M. J. Am. Soc. Mass Spectrom. 2003, 14, 834−850. (58) Senguen, F. T.; Doran, T. M.; Anderson, E. A.; Nilsson, B. L. Mol. Biosyst. 2011, 7, 497−510. (59) Liang, Y.; Pingali, S. V.; Jogalekar, A. S.; Snyder, J. P.; Thiyagarajan, P.; Lynn, D. G. Biochemistry 2008, 47, 10018−10026. (60) Madine, J.; Davies, H. A.; Shaw, C.; Hamley, I. W.; Middleton, D. A. Chem. Commun. 2012, 48, 2976−2978. (61) Campioni, S.; Mannini, B.; Zampagni, M.; Pensalfini, A.; Parrini, C.; Evangelisti, E.; Relini, A.; Stefani, M.; Dobson, C. M.; Cecchi, C.; Chiti, F. Nat. Chem. Biol. 2010, 6, 140−147. (62) Glabe, C. G. J. Biol. Chem. 2008, 283, 29639−29643.

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