540 nature chemical biology | vol 9 | SEPTEMBER 2013 | www.nature.com/naturechemicalbiology

articlepublished online: 14 july 2013 | doi: 10.1038/nchembio.1294

HBV is a highly infectious human pathogen that causes seri-ous liver disease. Worldwide, around 350 million people have chronic HBV infections (causing 600,000 deaths annually)1.

Although excellent vaccines exist, there are no effective therapies for chronic HBV infections. Approved therapies are broad-spectrum reverse transcriptase inhibitors that also target the HBV polymerase, but these have low efficacy and side effects and cause resistance1.

HBV replication within hepatocytes involves coordinated inter-actions between viral proteins, nucleic acids and host proteins. Virology has identified the key biomolecules and respective func-tions needed for replication2–5. X-ray crystallography and cryo-electron microscopy yielded beautiful structural insights on HBV capsids, nucleocapsids and virions6–8. A structural view of replication mechanisms has proved elusive as HBV proteins are challenging to study in vitro, thus hampering development of new therapies1.

Infectious HBV virions comprise an icosahedral nucleocapsid formed by the capsid-forming core protein HBc, within which the viral polymerase, genome and host proteins are encapsulated4. Nucleocapsids are, in turn, enveloped by three viral surface proteins: the small (S), middle (M) and large (L) surface proteins (Fig. 1a)2,5. Although S and M have single transmembrane orientations, L adopts two distinct orientations, i-preS L and e-preS L (Fig. 1b). In i-preS L, preS domains are cytoplasmically oriented and have essential roles in nucleocapsid envelopment2. In e-preS L, preS domains are located within the endoplasmic reticulum lumen9, eventually dis-playing preS1 on the exterior of the virion envelope, where it medi-ates infection via a cell-surface receptor (or receptors)5,10. PreS1 also regulates genome amplification after infection, has transactivation activity in host cells, regulates viral particle secretion and interacts with host chaperones to establish the dual topology of L11.

HBV nucleocapsids interact with surface proteins at the cyto-plasmic face of a membrane compartment distal to the trans-Golgi

network, thus generating enveloped virions for export via vesicular trafficking11–13. Recruitment and sorting of protein cargoes destined for different cellular compartments is performed by adaptor pro-tein (AP) complexes, a family of heterotetrameric proteins14. HBV is proposed to exploit adaptor protein complexes and associated transport pathways for envelopment and export of virions11,12.

Yeast two-hybrid and co-immunoprecipitation studies show i-preS L (Fig. 1b) interacts with γ2-adaptin but apparently not with γ1-adaptin12 (its closest homolog and part of the adaptor protein-1 (AP-1) complex). It is not yet known which adaptor protein complex contains γ2-adaptin, but γ1- and γ2-adaptin are not functionally interchangeable and localize to distinct cellular compartments13,15–18. Immunofluorescence microscopy shows that γ2-adaptin normally adopts perinuclear locations within hepatocytes but is recruited to the Golgi by L when in its i-preS L conformation (which displays its preS1 domain in the cytoplasm; Fig. 1b). Similarly, the γ2-adaptin EAR domain (hereafter γ2-EAR; Fig. 1c,d) mediates binding to and localization together with HBV L12. siRNA knockdown of γ2-adaptin has no effects on host protein trafficking, HBV protein expression, genome replication or nucleocapsid formation, but it inhibits nucle-ocapsid envelopment13,19,20. Further, the γ2-adaptin HEAD domain binds HBV nucleocapsids, and γ2-adaptin localizes together with HBV L and nucleocapsids in immunofluorescence experiments20. This complex ballet of interactions mediates virion export by the host ESCRT machinery.

We used a multidisciplinary strategy to obtain structural and thermodynamic insights on the preS1–γ2-EAR interaction. To the best of our knowledge, these are the first atomic-resolution insights into the interaction of an HBV protein with a human binding part-ner needed for viral replication. Whereas γ2-EAR is a globular folded protein, preS1 is an intrinsically disordered protein (IDP)21–23. Unlike many IDPs, preS1 did not have substantial preformed

1School of Medicine and Medical Science, University College Dublin, Dublin, Ireland. 2School of Biomolecular and Biomedical Science, University College Dublin, Dublin, Ireland. 3Astbury Centre for Structural Molecular Biology, University of leeds, leeds, UK. 4School of Biochemistry and Immunology, Trinity College Dublin, Dublin, Ireland. 5Centre for Synthesis and Chemical Biology, University College Dublin, Dublin, Ireland. 6Medical Research Council laboratory of Molecular Biology, Cambridge, UK. 7These authors contributed equally to this work. 8Present addresses: École Normale Supérieure de lyon, Centre de Résonance Magnétique Nucleaire à Très Hauts Champs, UMR 5280 Centre National de la Recherche Scientifique–École Normale Supérieure, villeurbanne, France (G.J.P.R.) and Clare Hall laboratories, Cancer Research UK, Herts, UK (v.E.P.). *e-mail: neil.ferguson@ucd.ie

the hepatitis b virus pres1 domain hijacks host trafficking proteins by motif mimicrymaike c jürgens1,7, judit Vörös2,7, gilles j p rautureau2,8, dale a shepherd3, Valerie e pye4,8, jimmy muldoon5, christopher m johnson6, alison e ashcroft3, stefan m V Freund6 & neil Ferguson1*

Hepatitis B virus (HBV) is an infectious, potentially lethal human pathogen. However, there are no effective therapies for chronic HBV infections. Antiviral development is hampered by the lack of high-resolution structures for essential HBV protein-protein interactions. The interaction between preS1, an HBV surface-protein domain, and its human binding partner, g2-adaptin, sub-verts the membrane-trafficking apparatus to mediate virion export. This interaction is a putative drug target. We report here atomic-resolution descriptions of the binding thermodynamics and structural biology of the interaction between preS1 and the EAR domain of g2-adaptin. NMR, protein engineering, X-ray crystallography and MS showed that preS1 contains multiple g2-EAR–binding motifs that mimic the membrane-trafficking motifs (and binding modes) of host proteins. These motifs local-ize together to a relatively rigid, functionally important region of preS1, an intrinsically disordered protein. The preS1–g2-EAR interaction was relatively weak and efficiently outcompeted by a synthetic peptide. Our data provide the structural road map for developing peptidomimetic antivirals targeting the g2-EAR–preS1 interaction.

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secondary structure, and γ2-EAR binding did not induce further secondary structure formation. PreS1 contains multiple motifs that mimic the γ-adaptin recognition motifs of host cargo proteins that assist transport between membrane compartments24. These motifs allowed preS1 to bind γ2-EAR (and therefore HBV to gain access to the host protein-export apparatus). The essential γ2-EAR–preS1 interaction was efficiently outcompeted by a high-affinity peptide derived from phage-display experiments24. Our detailed insights on the thermodynamics, structural biology and inhibition of the γ2-EAR–preS1 interaction provide an ideal road map for developing peptidomimetic antivirals targeting HBV.

RESulTSAtomic-resolution mapping of the g2-EAR–preS1 interactionThe 1H,15N HSQC spectrum of γ2-EAR was well resolved (Fig. 2a) and independent of protein concentration, consistent with light-scattering data showing that apo–γ2-EAR was strictly monomeric (Supplementary Results, Supplementary Fig. 1). A ten-fold molar excess of unlabeled preS1 elicited large chemical shift perturbations

(CSPs) in a subset of 15N-labeled γ2-EAR amide resonances (Fig. 2a). These CSPs colocalized to three clusters of γ2-EAR resi-dues (714–727, 746–762 and 772–779; Supplementary Fig. 2a) that defined a discrete binding surface (Fig. 2b) in the same location as the peptide-binding sites of other EAR domains25,26.

Amide cross-peaks in 1H,15N HSQC spectra of apo-preS1 had narrow line widths and lacked dispersion in the 1H dimension, features typical of unfolded proteins (Fig. 2c). The 13Cα chemical-shift index (Supplementary Fig. 2b), an indicator of secondary structure27, confirmed that preS1 had only short segments of partially populated α-helical or extended structure. Thus, the 108-residue preS1 domain is an IDP, consistent with earlier spectroscopy stud-ies of a 119-residue variant from a different viral subtype21. γ2-EAR was titrated into 13C,15N-labeled preS1 at increasing molar ratios to identify CSPs elicited by binding (Fig. 2c). A single cross-peak per preS1 amide was observed, indicating fast exchange between free and bound states on the NMR timescale. 1H,15N HSQC spectra of preS1 in complex with γ2-EAR were similar to that of apo-preS1 (at all of the molar ratios examined), indicating that binding did not induce major structural rearrangements in preS1 (Fig. 2c).

PreS1 residues with large CSPs induced by γ2-EAR binding (>2 s.d. from the mean) were mainly located in a single contigu-ous tract (residues 29–41; Supplementary Fig. 2c). NMR titrations for these residues fitted to a single-binding-site model and yielded similar, weak apparent binding affinities (mean Kd ~150 ± 40 μM; Fig. 2d), explaining the fast exchange observed in HSQC spectra of preS1–γ2-EAR complexes (Fig. 2c). NMR titrations of preS1 Gly12 and Asp16 amides, which also had large CSPs, showed that this region bound γ2-EAR more weakly than the motif within residues 29–41 (Supplementary Fig. 2d).

Basic thermodynamics of the g2-EAR–preS1 complexAlthough it is challenging to measure ΔHbinding and ΔSbinding for weak interactions using isothermal titration calorimetry (ITC)28, accurate Kd values can be determined when binding saturation is ensured by concatenating sequential titrations (Fig. 2e). The wild-type preS1–γ2-EAR interaction was enthalpically favorable and entropically unfavorable with an apparent Kd of 120 ± 10 μM (Supplementary Table 1). This value agreed well with values determined from NMR titrations (Fig. 2d) and affinities of other EAR domain–peptide interactions (0.7 ≤ Kd ≤ 230 μM)29,30. The apparent binding stoichi-ometry (n), determined from >40 ITC experiments (Supplementary Table 1), ranged from 0.4–1.0, suggesting that preS1 bound one or two γ2-EAR molecules.

identifying g2-EAR residues involved in binding to preS1We mutated solvent-exposed γ2-EAR residues whose amides had large CSPs (>1 s.d. from the mean (Fig. 2b)) when bound to preS1 to assess their contribution to binding energetics (Supplementary Fig. 3a,b). Of 17 γ2-EAR residues examined, only 6 had lower affinity for preS1 when mutated (Ala716, Lys719, Ser720, Pro754, Arg756 and Arg760; Supplementary Table 1). These residues, thermo-dynamic determinants of binding, localized together to the peptide-binding surface identified by CSPs (Fig. 2b) and found in other EAR domains25,26,29. All of the other mutated residues gave rise to similar affinities to the wild-type protein (Supplementary Table 1), indicating that they contributed little to preS1 binding (Fig. 2b).

identifying which regions of preS1 bind g2-EARPrevious studies identified a five-residue motif that confers peptide or protein binding to human EAR domains, which can be written as follows: Asp/Glu0, Phe/Trp1, X2, X3, Φ4 (where X is any residue, and Φ is phenylalanine, tryptophan, tyrosine, leucine, iso-leucine or methionine)24. Positions 1 and 4 are hydrophobic side chains forming a ‘two-pin plug’ motif packing onto EAR domain surface cavities24,29,31. γ2-EAR binding induced large CSPs in preS1

dC

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Figure 1 | Domain structure and topology for HBV surface proteins and g2-adaptin. (a) S, M and l start from different in-frame initiation codons (arrows) but share a common C terminus2,5. (b) S and M are predicted to have four membrane-spanning regions, with the N and C termini located in the endoplasmic reticulum (ER) lumen (corresponding to the virion exterior after virus envelopment and maturation). Asn146 of S exists in glycosylated (branched structure) and nonglycosylated forms in S, M and l. Asn4 of the preS2 domain is glycosylated only in M. l has two conformations, i-preS and e-preS, that differ in the orientation of preS domains across the endoplasmic reticulum membrane2,5. (c) Reported γ2-adaptin domains and boundaries20. (d) Ribbon diagram showing superimposed crystal structures of γ1-EAR (PDB 1GYU25, residues 703–821, dark gray), γ2-EAR (PDB 4BCX, residues 665–785, black) and GGA3-EAR (PDB 1P4U26, residues 579–723, light gray). EAR domains have two β-sheets packed together in an immunoglobulin-like fold. The structure of apo–γ2-EAR is very similar to that of the EAR domains of γ1-adaptin and GGA3 (Cα r.m.s. deviation of 1.0 Å and 0.9 Å, respectively). This structural similarity is unsurprising given that γ1-EAR, the closest relative of γ2-EAR, shares 45% sequence identity (and 50% identity for binding-site residues, where these are identified as solvated residues with large CSPs induced by binding full-length proteins43).

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residues 29–41 (29NPDWDFNPNKDTW41; Supplementary Fig. 2c). This region contained two motifs resembling the canonical motif: Site 1, 31DWDFN35, where Asp31 is residue 0 and Trp32 and Asn35 adopt canonical ‘pin’ positions, and Site 2, 33DFNPN37, where Asp33 is residue 0, and Phe34 and Asn37 adopt canonical pin positions. To test which site (or sites) bound γ2-EAR, we measured the affin-ity of γ2-EAR for three preS1-derived peptides: preS1(29–40), encompassing Sites 1 and 2; preS1(29–36), spanning Site 1; and preS1(33–40), spanning Site 2 (Supplementary Table 1). PreS1 (29–40) and preS1(29–36) bound γ2-EAR with similar affinities (Kd ~1 mM), whereas no binding was detected for preS1(33–40). Notably, the affinity with which preS1(29–40) and preS1(29–36) bound γ2-EAR was lower by a factor of ten than that of full-length preS1 (Supplementary Table 1).

We used ITC to characterize the affinity of 18 preS1 variants for binding to γ2-EAR (Supplementary Table 1 and Supplementary Fig. 3c,d). Four mutated residues (D31A, W32A, D33A and F34A) had considerably lower affinities for γ2-EAR, which, together with our peptide-binding studies, showed that the main binding motif lay within Site 1. The N35A and N35G mutants had wild-type Kd values, showing that Asn35 did not act as the second pin or contribute to γ2-EAR binding energetics (Supplementary Table 1). Introducing the N35F mutation or inserting a glycine residue between Trp32 and Asp33 (preS1INS), making Site 1 better match the canonical motif24, increased the affinity of preS1 for γ2-EAR (Kd ~45 μM and ~20 μM, respectively; Supplementary Table 1). However, alanine scanning strongly suggested that Trp32 and Phe34 were the first and second pins of Site 1.

γ2-EAR binding induced large CSPs in the amides of preS1 Gly12 and Asp16 (greater than two standard deviations from the mean, Supplementary Fig. 2c) and mapped to Site 3 (9NPLGFFPDHQLD20),

a region with two correctly spaced hydrophobic residues (under-lined) and multiple acidic groups. ITC showed that binding of preS1(9–16), a Site 3 peptide, to γ2-EAR was weaker by a factor of ~20 than that of full-length preS1 (Supplementary Table 1), consistent with NMR titrations (Supplementary Fig. 2d). Thus, preS1 Site 3 is an additional, lower-affinity γ2-EAR binding motif. Notably, two excellent algorithms for identifying bind-ing motifs in disordered proteins, ANCHOR and SLiMPred32,33, clearly identified Site 3 as a putative binding motif. By contrast, the predicted scores for Site 1 were only of borderline importance (Supplementary Fig. 4).

γ2-EAR binding induced CSPs in amides of some preS1 residues downstream of Sites 1 and 2 (Supplementary Fig. 2c). However, these CSPs were below the significance threshold we normally use (>2 s.d. from mean CSP values). To determine the origin of these CSPs, we studied preS1(42–108), a variant where the N-terminal region containing Sites 1, 2 and 3 was deleted. PreS1(42–108) bound γ2-EAR with a Kd of ~750 μM (Supplementary Table 1), showing that an additional γ2-EAR–binding motif exists downstream of resi-due 42, consistent with yeast two-hybrid screens12. On the basis of our findings and ANCHOR predictions32 of putative binding motifs within preS1 residues 41–80 (Supplementary Fig. 4), we identified two putative γ2-EAR–binding motifs, Site 4 (50GAFGLGFTP58) and Site 5 (59PHGGLLGWSP68). These sites lacked the acidic residues present in Site 1 and Site 3 and in canonical motifs. Unfortunately, preS1(50–58), a Site 4 peptide, was insoluble, which prevented us from using ITC and NMR to determine whether it bound γ2-EAR. preS1(59–68), however, was soluble and bound γ2-EAR with a Kd of ~5.2 mM (Supplementary Table 1). On the basis of peptide-binding data, the rank order of γ2-EAR binding affinity was determined to be Site 1 > Site 3 > Site 5. Site 4 may also be a low-affinity motif, but

Asn37

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ole

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ant

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Figure 2 | Mapping the g2-EAR–preS1 interaction using NMR spectroscopy and iTc. (a) overlay of 1H,15N HSQC spectra of γ2-EAR in the apo form (blue) and with a ten-fold molar excess of preS1 (red). Expanded regions show resonances representative of those that showed either large or no CSPs upon binding to preS1. (b) CSPs induced by preS1 binding mapped onto the γ2-EAR surface (PDB 4BCX). Residues with mean CSPs >1 (orange) or >2 (red) s.d. are indicated. Asterisks indicate Gln724 and Gln726, bystander residues that had CSPs arising from their proximity to the preS1-binding site. (c) overlay of 1H,15N HSQC spectra of apo-preS1 (blue) and in the presence of a four-fold molar excess of γ2-EAR (red). Expanded regions show CSPs of preS1 residues after addition of unlabeled γ2-EAR at different molar ratios: apo-preS1 (blue), 1:1 (yellow), 1:2 (purple), 1:3 (cyan), 1:4 (red). The 15N dimension is truncated as no CSPs were detected in resonances upfield of 110 p.p.m. (d) NMR titration determined from c. The solid black line indicates a simulated binding curve generated using the Kd value determined for the wild-type preS1 and γ2-EAR interaction using ITC (Supplementary Table 1). (e) Representative example of concatenated sequential ITC titrations. The first and second titrations for the preS1 N35G mutant are marked with a 1 and 2, respectively. The solid line is the fit to a single-site binding model. These data show the value of concatenating data sets to obtain better-defined binding curves (especially for interactions with micromolar affinity). DoF, degrees of freedom.

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we could not verify this experimentally. CSPs for Sites 4 and 5 were too small to obtain robust Kd values from NMR titrations34.

CSP studies on 15N-labeled γ2-EAR in the presence of preS1 (29–40), preS1(29–36), preS1(9–16) or preS1(59–68) showed that each peptide induced CSPs in the amide resonances (Fig. 3) most affected by binding full-length preS1 (Fig. 2b). Thus, Site 1, Site 3 and Site 5 peptides bind the same cleft on γ2-EAR, making bind-ing mutually exclusive. Further, Site 1 will saturate at much lower concentrations of γ2-EAR or γ2-adaptin. Although G12A and D16A should lower the affinity of Site 3 for γ2-EAR (Supplementary Table 1), these effects were masked in most experiments (where Site 1 binding predominated at the molar ratios used; Fig. 2c–e).

Structural analysis of g2-EAR–peptide interactionsThere are no reported structures of γ2-EAR protein-peptide inter-actions. To better understand γ2-EAR ligand recognition, we deter-mined high-resolution crystal structures of apo–γ2-EAR alone and in complex with a peptide derived from phage-display experi-ments (–2GEEWGPWV5)24,29,30 and preS1(29–36), the Site 1 peptide (Supplementary Table 2). The phage peptide bound γ2-EAR via a two-pin plug motif (Fig. 4a) without inducing structural changes in γ2-EAR compared to apo–γ2-EAR (Cα r.m.s. deviation ~0.3 Å). Trp1 packed between the aliphatic side chains of γ2-EAR Arg756 and Lys758, whereas Trp4 packed between Lys758 and Arg760 (Fig. 4b). The phage peptide backbone of Gly2 and Pro3 distorted outwards, thus orienting the aromatic pins for γ2-EAR binding (Fig. 4a), as in other EAR domain–peptide complexes26,29,31.

As preS1(29–36) bound weakly to γ2-EAR, its occupancy in the crystal structure was low (making interpretation of peptide electron density challenging). Nonetheless, our data showed that preS1 Trp32 and Phe34 mimicked a two-pin plug motif (Fig. 4c) when bound to γ2-EAR and packed into the same γ2-EAR surface cleft as the phage peptide (Fig. 4a–c). The key differences were that (i) Trp32, the first pin of preS1(29–36), adopts a different orientation than Trp1 of the phage peptide (compensated by rearrangements of the γ2-EAR Lys758 side chain) and that (ii) preS1(29–36) pins were separated by one residue, not two as in the canonical motif (Fig. 4a–c)24. The peptide-binding cleft of γ2-EAR was highly basic, had charge complementarity with peptide acidic residues and mapped to the region with the largest CSPs induced by binding full-length preS1 (Fig. 4c,d).

Peptides can inhibit preS1–g2-EAR interactions in vitroGiven the similarity in binding modes of preS1- and phage-derived peptides (Figs. 3 and 4), we hypothesized that the phage peptide

preS1 wild type

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Site 1 peptide

Site 5 peptideSite 3 peptide

Figure 3 | cSPs induced in g2-EAR amide resonances by binding to preS1 and preS1-derived peptides. γ2-EAR CSPs (>2 s.d.) induced by binding full-length wild-type preS1 are indicated in red on the surface of the wild-type γ2-EAR structure. Residues whose amides had the largest CSPs upon addition of a four-fold molar excess of preS1-derived peptides or preS1(42–108) are shown in red on the surface of the wild-type γ2-EAR structure. The residues most affected by binding (Ala716, val717, Arg756, lys758, leu759 and leu761) localized together with known peptide-binding sites of other EAR domains25,26,29,30,49. The CSPs for the Site 5 peptide were the smallest observed as this peptide had the lowest affinity for γ2-EAR, which therefore led to the lowest occupancy of the peptide–γ2-EAR complex (at the molar ratios used in these experiments). Each preS1-derived ligand also induced large CSPs in amide resonances of buried residues within the γ2-EAR peptide-binding cleft (not visible in this representation as they are below the protein surface).

Arg756

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Figure 4 | crystal structures of wild-type g2-EAR in complex with peptide ligands. (a) Surface representation of wild-type γ2-EAR (in gray) in complex with a phage peptide24 (PDB 2YMT; Supplementary Table 2). The two tryptophan residues of the phage peptide (displayed as sticks) plug into the binding surface on γ2-EAR (gray). (b) Close-up showing how the phage peptide two-pin plug motif24,26 packs against aliphatic regions of γ2-EAR basic side chains. The Fo–Fc difference density map of the phage peptide (orange) is shown in green and contoured at 2 σ. (c) Surface representation of the structure of wild-type γ2-EAR in complex with the preS1(29–36) peptide (PDB 3ZHF; Supplementary Table 2). large CSPs induced when full-length preS1 bound to1H,15N-labeled γ2-EAR are indicated in orange and red (1 and 2 s.d. from the mean, respectively, as in Fig. 2b) with the peptide shown as sticks. (d) Electrostatic surface potential calculated for wild-type γ2-EAR in complex with preS1(29–36) (in a similar orientation as that in Fig. 4c). This shows clearly that the γ2-EAR peptide-binding cleft is highly basic (blue) and has favorable charge complementarity with the acidic residues (red) on the peptide. This helps explain the observed changes in affinity when charged residues were mutated to neutral ones (Supplementary Table 1).

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should inhibit the interaction between full-length preS1 and γ2-EAR. The phage peptide bound γ2-EAR four orders of magnitude tighter than full-length preS1 (apparent Kd ~10 nM and 120 μM, respectively) and efficiently displaced full-length preS1 from γ2-EAR in competition experiments (Supplementary Fig. 5).

g2-EAR binding motifs map to a rigid region of preS1We used NMR to characterize the backbone dynamics of preS1 to see whether it had additional properties not recapitulated by Site 1 peptides, which explained why they bound γ2-EAR more weakly than full-length preS1. 15N-1H heteronuclear NOE enhancements, which probed fast, picosecond backbone dynamics35, showed that residues 11–44 (encompassing Site 1, Site 2 and Site 3), though not as rigid as a folded protein, were less dynamic than expected for an IDP (Fig. 5a), a trend confirmed by 15N transverse relaxation data (Supplementary Fig. 6). This conformationally restrained region of preS1 had only transiently populated structural elements (Supplementary Fig. 2b).

Testing conformational selection using MSIon mobility spectrometry–mass spectrometry (IMS-MS) is a gas-phase technique for studying the shape and conformation of biomolecules and complexes thereof. IMS-MS can separate differ-ent conformers with identical m/z, providing they have detectable differences in their relative mobility through the ion-mobility drift cell (arising from orientationally averaged collision cross-sections (CCSs))36–38. Although gas-phase polypeptide conformations may not always recapitulate those in solution, the short time lag between the transfer of proteins from the liquid to the gas phase, combined with near-instantaneous IMS-MS analyses, means that solution conformations are likely to be broadly preserved39. Consequently, IMS-MS has emerged as an exciting technique for characterizing the conformational dynamics of IDPs, providing complementary insights to those obtained using NMR, small-angle X-ray scattering and crystallography40,41.

γ2-EAR had a single m/z distribution (Supplementary Fig. 1c) and IMS-MS arrival time distribution (Fig. 5b), indicating a compact

species (consistent with being folded). PreS1 had a more complex m/z distribution (Supplementary Fig. 1d), and IMS-MS revealed multiple conformers with distinct CCS (Fig. 5b). Although most preS1 conformers were extended, there was a highly populated con-formation (~24% of the total) that was more compact than folded γ2-EAR, a polypeptide of similar mass (Fig. 5b).

We then used IMS-MS to assess whether γ2-EAR preferentially bound certain preS1 conformers. However, IMS-MS could not detect the weak interaction between wild-type preS1 and γ2-EAR. Thus, we engineered a preS1–phage peptide chimera, in which the higher-affinity phage peptide γ2-EAR–binding motif (Fig. 4a,b) replaced Site 1 of full-length preS1 (Supplementary Fig. 1e). IMS-MS dem-onstrated that the chimera populated comparable conformers (with similar CCS values) as wild-type preS1, suggesting near-identical behavior (Fig. 5b). The preS1 chimera bound to wild-type γ2-EAR and formed a 1:1 complex in solution (Fig. 5c) and in the gas phase (Supplementary Fig. 1f), where it could be studied by IMS-MS (Fig. 5d). The m/z spectrum of the complex had two charge-state dis-tributions, indicating two distinct conformer families with distinct, resolvable CCS values, thus showing that γ2-EAR bound extended and compact preS1 conformations. Notably, the CCS versus charge state plots for conformers of the preS1 chimera–γ2-EAR com-plex (Fig. 5d) qualitatively mirrored those of apo-preS1 (Fig. 5b). Thus, γ2-EAR binding does not seem to alter the conformational properties of preS1.

DiScuSSiONTo the best of our knowledge, this is the first atomic-resolution struc-tural and thermodynamic analyses of an HBV protein (preS1) inter-acting with a cognate human binding partner (γ2-EAR) required for HBV replication12,19,20. HBV has evolved conserved motifs (Sites 1, 3 and 5) that mimic host two-pin plug motifs24,26 and allow preS1 to bind γ2-EAR or γ2-adaptin and ‘piggyback’ on the host membrane-trafficking apparatus to facilitate nucleocapsid envelopment11,12,19,20. It is argued that human short linear motifs (SLiMs) are a functional Achilles’ heel that viruses exploit, as viruses can rapidly evolve and mimic even evolutionarily dynamic SLiMs of hosts, in some cases

Figure 5 | Backbone dynamics and conformational diversity of preS1. (a) 15N-1H heteronuclear NoE enhancements of preS1 at 20 °C. ordered regions of globular proteins typically have values between 0.8–1.0. Although highly dynamic regions of preS1 had negative values (for example, N and C termini), residues 11–44 had values of ≤0.4, suggesting that this region rapidly interconverted between ordered and extended conformations. Potential γ2-EAR– binding sites (Sites 1 to 5) examined in this study are indicated. (b) m/z charge state ions [(M+nH)n+] versus CCS plot (determined from IMS-MS) for wild-type preS1 (black), γ2-EAR (red) and the preS1–phage peptide chimera (blue). lines joining points indicate distinct conformational states. Whereas there was one compact γ2-EAR conformer, there were at least four preS1 conformers. The preS1–phage peptide chimera behaved essentially identically to wild-type (WT) preS1. Inset, the IMS-MS arrival time distribution of the 8+ charge state of preS1 (solid line) indicates multiple conformational species with distinct CCS and fits to the sum of four Gaussian profiles (dashed lines). (c) Size-exclusion chromatography with multi-angle laser light scattering (SEC-MAlS) analysis of apo–γ2-EAR (red), apo-preS1 phage peptide chimera (blue) and a 1:1 complex of these proteins (black). An arrow indicates low quantities of unbound γ2-EAR. (d) This plot shows two conformational ensembles for the γ2-EAR–preS1 chimera complex: compact (CCS = 2,053 ± 61 Å2, lowest charge state) and extended (2,388 ± 60 Å2, 11+ charge state). Inset, the IMS-MS arrival time distribution for the complex (11+ charge state) indicates compact and extended conformers. Error bars on IMS figures show the s.d. obtained from three independent measurements.

d

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even duplicating SLiMs to provide ‘functional robustness’42 (as we found for preS1).

We believe that Site 1 is the preferred γ2-EAR binding motif in vivo as it will saturate preferentially at lower concentrations of γ2-EAR or γ2-adaptin. A single-site model best described the binding stoichiometry for γ2-EAR–preS1 interactions at molar ratios ≤4:1 (Fig. 2e), conditions where preS1 binding to the lower- affinity Sites 3 and 5 did not intrude substantially. However, when Site 1 mutations reduced the affinity of γ2-EAR–preS1 interactions (Supplementary Table 1), Site 3 and 5 binding became more appar-ent (and yielded non-unity n values in ITC experiments). Multiple, functionally redundant motifs provide a ‘Kd floor’, ensuring that even mutation-prone viruses can always exploit valuable functions of the host machinery.

Although the γ2-EAR–phage peptide structure (Fig. 4a) showed the binding mode of other EAR domain-peptide complexes26,29, our protein engineering, crystallographic and NMR experiments also showed that preS1 Site 1 (31DWDFN35) bound γ2-EAR by mimicking a two-pin plug binding motif26. PreS1, however, man-ages this using only one spacer residue (Fig. 4a–c). Conversely, the γ1-EAR–binding motif in γ-BAR contains three spacer residues43. These data, combined with our identification of other noncanonical motifs (Sites 3 and 5), suggest that EAR-domain binding motifs are perhaps more plastic than first thought24,26. A reappraisal of EAR-binding determinants may help identify new host cargo proteins.

Acidic residues near two-pin plug motifs are important for γ1-EAR ligands24. We saw similar effects for the preS1–γ2-EAR inter-action (Supplementary Table 1). Our structural studies (Fig. 4c,d) suggest that these effects arise from favorable electrostatic interac-tions between ligand acidic residues and basic residues lining EAR domain peptide-binding clefts. PreS1 D31A and D33A mutations weakened binding (Supplementary Table 1), presumably through removal of favorable electrostatic interactions. Similarly, the γ2-EAR K719Q, R756A, R756Q and R760A mutants, which lowered the basicity of the peptide-binding surface when present as single point mutations, also reduced the affinity of γ2-EAR for preS1. The K38A mutation in PreS1, which reduced charge repulsion with basic γ2-EAR residues, increased preS1 affinity for γ2-EAR.

Although bioinformatics algorithms are successful at identify-ing molecular recognition features undergoing disorder-to-order transitions upon binding, unstructured regions conserved across protein families and SLiMs23,42, even the best algorithms did not robustly identify preS1 Site 1 (Supplementary Fig. 4). Thus, we searched for additional properties of preS1 that might assist motif identification in other IDPs.

NMR backbone dynamics revealed a conformationally restrained region of preS1 (residues 11–44; Fig. 5a) within which lay Sites 1 and 3. This rigidity was remarkable for an IDP, especially one that had (at most) only transiently populated secondary structures (Supplementary Fig. 2b). Even Site 5, the lowest-affinity γ2-EAR–binding motif we identified, lay within a region less dynamic than most regions of preS1. Inspection of Sites 1, 3 and 5 showed that each motif was flanked by proline residues (PDWDFNP, PLGFFP and PHGGLLGWSP, respectively). Although it is premature to place too much emphasis on the absolute rigidity of these motifs, the backbone dynamics of γ2-EAR–binding motifs and other regions of preS1 fall into discrete clusters flanked by proline residues with coherent dynamic properties (Fig. 5a). We speculate that these ‘dynamic clusters’ arise from conformational restrictions, wherein flanking prolines act as motif ‘start’ and ‘stop’ signals (with interven-ing sequences determining specificity and affinity).

Markedly, recent virology studies show that Sites 1 and 3 are functionally important loci for preS1-mediated uptake of HBV virions into host hepatocytes (where the e-preS L conformer dis-plays preS1 on the virion exterior; Fig. 1b)44. Similarly, neutralizing antibodies have nonoverlapping epitopes between preS1 residues

21–32, another relatively rigid region of preS1. It is not yet clear whether sequence composition, dynamics or both cause the rigid region of preS1 to be functionally diverse and antigenic, but our observations are notable. We speculate that interaction motifs may often be located in dynamic clusters pinched off between proline residues. If this proves to be general, it could help identify interac-tion motifs as proline residues are common in IDPs, and backbone dynamics data are rapidly acquired, easy to interpret and readily benchmarked against other sequence features.

Although we anticipated that preS1 should have extended con-formations, to our surprise, it also populated a conformer more compact than folded γ2-EAR (Fig. 5b). A reasonable assumption was that γ2-EAR would bind equally well to extended and compact preS1 conformations as its recognition motifs were short (Figs. 4 and 5a), which is exactly what we observed in IMS-MS experi-ments (Fig. 5d). We found no evidence of γ2-EAR binding inducing secondary structure formation in preS1 (Fig. 2c), but some local ordering of the polypeptide backbone and side chains within bind-ing motifs is likely.

IMS-MS is a powerful method for directly testing binding mod-els such as conformational selection, where a protein may show binding preferences for ‘open’ or ‘closed’ conformations of partner proteins. Similarly, IMS-MS may reveal whether there are coarse-grain structural features present in IDPs that influence binding (even though such ‘structure’ does not fit the classical structure-function paradigm)22. Such structure may explain why full-length preS1 bound γ2-EAR tighter than Site 1 peptides or bound preS1(42–108) tighter than the Site 5 peptide. However, it is also possible that flanking preS1 sequences make additional, cryptic contributions to binding that are not identified in this study.

As this research was inspired by cellular and virology research, it is appropriate to consider the wider biological context of our findings. If γ2-adaptin is part of a heterotetrameric adaptor pro-tein complex14 (as seems likely), this complex will contain another adaptin molecule with its own EAR domain. This adaptor protein complex could bind preS1 Sites 1 and 3 (or Sites 1 and 5), with Site 1 being the preferred γ2-adaptin motif (Fig. 6). Initial binding to Site 1 would enhance the affinity of the adaptor protein complex for Site 3 or 5 (or vice versa, depending on the specificity of each EAR domain). Alternatively, the adaptor protein complex could bridge multiple i-preS L chains by binding their preS1 domains. These binding modes would enhance the affinity and specificity of the

Site 3 Site 1 Site 1preS1

? ?γ2 γ2

Site 5

i-preS

Figure 6 | Models of g2-adaptin interactions leading to envelopment of HBV nucleocapsids. γ2-adaptin may function as part of an adaptor protein complex14 in which the EAR domains of two adaptin molecules, γ2-adaptin and ?-adaptin (an as-yet-undefined adaptin), bind Site 1 and Site 3 or Site 1 and Site 5 within a single molecule of i-preS l. γ2-adaptin HEAD is reported to interact with the HBv capsid-forming core protein20. The HEAD domains of each adaptin are connected to their respective EAR domains via flexible hinge regions (in gray)14. This flexible linker may facilitate formation of the multiprotein complex needed for nucleocapsid envelopment. Potential sites for therapeutic targeting are indicated by red crosses. In this article, we showed that a phage-derived peptide efficiently inhibited the preS1-γ2-EAR interaction in vitro (Supplementary Fig. 5).

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γ2-adaptin–preS1–nucleocapsid interactions compared to individ-ual interactions acting in isolation.

These interaction models (Fig. 6) are appealing as they help explain how the γ2-adaptin HEAD domain can interact with the HBV nucleocapsid while at the same time presenting its C-terminal EAR domain to interact with i-preS L. This spatial coordination of key molecular interactions between HBV and host proteins should efficiently recruit HBV nucleocapsids for envelopment and export, consistent with reported functional and colocalization studies16,19,20. Notably, earlier virology studies aimed at mapping the sequence determinants of virion uptake show that deletions spanning preS1 Site 1, Site 3 or Site 5 impair HBV virion secretion and clearly ablate virus infectivity45,46. These findings support the functional impor-tance of the γ2-EAR binding sites that we report here (with the residual secretion remaining when only one site was deleted, con-sistent with multiple motifs conferring functional redundancy42). Thus, preS1 fits well the paradigm of a multifunctional viral IDP using multiple SLiMs to mediate more functionality than is possi-ble for a similar-sized globular protein, helping to offset functional limitations that could arise from the spatial constraints imposed by the tiny HBV genome42.

As siRNA knockdown of γ2-adaptin ablates HBV nucleocapsid envelopment19,20, the γ2-adaptin–preS1 interaction is a poten-tial therapeutic target. We showed that a high-affinity peptide efficiently outcompeted the preS1–γ2-EAR interaction in vitro (Supplementary Fig. 5), and potent antiviral peptides that target HBV are reported44,47,48. Thus, our structural and thermodynamic insights provide an ideal road map for developing peptidomimetic inhibitors of the γ2-EAR–preS1 interaction. It remains to be seen whether this can be achieved without interfering with normal interactions of host adaptor proteins. However, as preS1 also uses noncanonical motifs, it may be possible to specifically target preS1 interactions without deleterious side effects. Potent, well-tolerated inhibitors of γ2-adaptin–preS1 interactions could provide a route toward improved combination therapies for chronic HBV infec-tions—a much-needed outcome for the millions suffering from this potentially lethal disease.

received 25 January 2013; accepted 12 June 2013; published online 14 July 2013

METHODSMethods and any associated references are available in the online version of the paper.

Accession codes. Protein Data Bank. The crystal structure coordi-nates for the following structures were deposited under the acces-sion codes in parentheses: apo–γ2-EAR (4BCX), γ2-EAR in complex with the phage-derived peptide (2YMT) and γ2-EAR in complex with preS1 Site 1 peptide (3ZHF).

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acknowledgmentsWe would like to thank K. Nakayama (University of Kyoto) for the γ2-adaptin clone; J. Lyons (Trinity College Dublin) and Deutsches Elektronen-Synchrotron–European Molecular Biology Laboratory beamline scientists for assistance with X-ray data collection; N. Moran (Royal College of Surgeons in Ireland) for access to biophysical instrumentation; M. Caffrey (Trinity College Dublin) for access to X-ray apparatus; and D. Shields and C. Mooney for bioinformatics help. This work was supported by Science Foundation Ireland (SFI)–President of Ireland Young Researcher Award (09/YI/B1682 to N.F.), Stokes Lecturer Award (07/SK/B1224a to N.F.), Royal Irish Academy–Royal Society Exchange award (IE111031 to N.F. and A.E.A.), SFI grant (07/IN.1/B1836) and US National Institutes of Health grants (GM75915, P50GM073210 U54GM094599) to M. Caffrey (funding V.E.P. and X-ray generator). D.A.S. was funded by an Engineering and Physical Sciences Research Council PhD Studentship. The Synapt mass spectrometer was purchased with Biotechnology and Biological Sciences Research Council UK funds (BB/E012558/1).

author contributionsM.C.J. purified proteins, performed ITC and NMR experiments, determined crystal structures, analyzed data and wrote the paper; J.V. purified proteins, performed ITC and NMR experiments, analyzed data and wrote the paper; G.J.P.R. performed and analyzed NMR experiments and wrote the paper; J.M. performed NMR experiments; D.A.S. performed and analyzed MS experiments and wrote the paper; V.E.P. determined crystal structures and wrote the paper; C.M.J. performed and analyzed calorimetry experi-ments; A.E.A. designed and interpreted MS experiments and wrote the paper; S.M.V.F. performed and analyzed NMR experiments and wrote the paper; N.F. directed research, designed and interpreted experiments and wrote the paper.

competing financial interestsThe authors declare no competing financial interests.

additional informationSupplementary information is available in the online version of the paper. Reprints and permissions information is available online at http://www.nature.com/reprints/index.html. Correspondence and requests for materials should be addressed to N.F.

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50. Sattler, M., Schleucher, J. & Griesinger, C. Heteronuclear multidimensional NMR experiments for the structure determination of proteins in solution employing pulsed field gradients. Prog. Nucl. Magn. Reson. Spectrosc. 34, 93–158 (1999).

ONliNE METHODSReagents. All reagents were of AnalR grade and purchased from Sigma Chemical Co. and Thermo-Fisher Scientific. N-acetylated and C-amidated peptides were synthesized by ChinaPeptides (Shanghai, China). The mass and identity of pro-teins and peptides were confirmed by MALDI mass spectrometry. Protein and peptide concentrations were determined from UV absorbance and calculated molar extinction coefficients (with corrections for light-scattering contributions). Quikchange kits were used for site-directed mutagenesis (Stratagene, CA).

Protein expression and purification. DNA encoding the first 108 residues of the L surface antigen protein (subtype adyw) was subcloned into a modified pRSETa vector (Invitrogen, CA) to yield pPRES1(1–108). E. coli C41 (DE3) cells transformed with pPRES1(1–108) were grown for 3 h at 37 °C and induced overnight with 1 mM IPTG. The cell pellet was harvested by centrifugation and resuspended in Buffer A (50 mM Tris-HCl, pH 8.0, 200 mM NaCl, 10 mM imidazole, 1 mg/ml lysozyme, protease inhibitor tablets (Roche, CA) and 1,000 Kunitz units of DNaseI). Fusion protein containing preS1 was isolated from clarified cell lysates using Ni-affinity chromatography. The fusion protein was cleaved off using TEV protease during an overnight dialysis in 20 mM Tris-HCl, pH 8.0. The fusion protein and TEV protease were removed using a second Ni-chromatography step. Flow-through fractions (containing preS1) were concentrated using ammonium sulfate fractionation. The pellet thus obtained was resuspended in 0.2 M ammonium bicarbonate before injection onto a preparative Superdex75 gel filtration column (GE Healthcare, PA). PreS1 fractions were identified using SDS-PAGE, pooled and freeze-dried before use.

DNA encoding residues Ala665–Gln785 of the human γ2-EAR domain was subcloned into a modified pRSETa vector to generate pG2EAR. γ2-EAR con-structs were overexpressed in E. coli C41 (DE3) grown at 22 °C with an over-night induction with 0.75 mM IPTG. Cell pellets were resuspended in Buffer A and lysed using sonication. The clarified cell lysate was applied to a Ni-affinity column and histidine-tagged fusion protein eluted in 50 mM Tris-HCl, pH 8.0, 200 mM NaCl, 450 mM imidazole. The fusion protein was cleaved off using an overnight incubation with TEV protease. The digested fusion protein and protease were removed using a second Ni-affinity step, and the flow-through containing γ2-EAR was concentrated using centrifugal concentrators. γ2-EAR constructs were further purified using a Superdex75 gel filtration column (GE Healthcare, PA). Fractions containing pure γ2-EAR domain were identified using SDS-PAGE, pooled and stored at 4 °C until needed.

Multi-angle laser light scattering. Static light-scattering measurements, to determine the oligomeric state and purity of recombinant proteins, were performed at 20 °C using a Shimadzu HPLC system connected to Wyatt Technologies Dawn Heleos II light scattering detector and Optilab REX refrac-tometer (Wyatt, CA). Size-exclusion chromatography was performed using an analytical Superdex75 column (GE Healthcare, PA) inside a column oven. To minimize temperature variations, all of the instruments had Peltier tem-perature controllers connected together using insulated PEEK tubing. Sample injections were 15–100 μl of protein solutions containing preS1 and γ2-adaptin EAR domains at various final concentrations (up to a maximum concentra-tion of 2.4 mM and 1.2 mM, respectively). SEC-MALS data were collected and analyzed using Wyatt Astra software (Wyatt, CA) to determine molar mass and monodispersity of species eluting from columns.

NMR spectroscopy. NMR spectra were recorded at 10 °C and 20 °C using 600 MHz Varian VNMRS (Agilent Technologies, CA) and 700 MHz Bruker Avance (Bruker BioSpin, MA) NMR spectrometers equipped with triple-resonance probes. The sample buffer for γ2-EAR NMR studies was 20 mM Tris-HCl buffer, pH 7.0, 100 mM NaCl, 1 mM DTT, 0.05% (w/v) NaN3, 0.1 μM DSS and 10% (v/v) D2O, whereas 30 mM MES buffer pH 6.0, 0.05% (w/v) NaN3 and 0.1 μM DSS was used for preS1 experiments. γ2-EAR was used at protein concentrations of 20–600 μM, whereas preS1 was used at 20–2,000 μM, concentrations where each protein was strictly monomeric (Supplementary Fig. 1). For NMR titrations, 1H,15N HSQC spectra were acquired using 40 μM 13C,15N-labeled preS1, to which unlabeled γ2-EAR was added at different molar ratios. Spectral acquisition and data processing were performed using the manufacturer’s software.

Backbone assignments for γ2-EAR and preS1 were based on standard triple resonance experiments50: 1H,15N HSQC, HNCACB, CBCACONH, HNCO, HNCACO; 15N backbone dynamics data were obtained from Pseudo3D 15N heteronuclear T1, T2 and 15N-1H NOE enhancement experiments derived

from the Bruker pulse sequence library. 1H chemical shifts were referenced relative to the DSS internal standard and heteronuclei via indirect referenc-ing. Backbone resonance assignments were obtained for 96% of γ2-EAR and 100% of preS1 residues (counting only nonproline residues). γ2-EAR residues Asn731, Val781 and the vector-derived Gly-2Gly-1Ser0 could not be unambiguously assigned.

ITC. ITC measurements were performed at 10 °C using a MicroCal ITC200 calorimeter (GE Healthcare, PA). Prior to experiments, samples were dia-lyzed against 20 mM sodium phosphate buffer, pH 7.0, 100 mM NaCl, 0.05% (w/v) sodium azide. For protein-protein interaction titrations, ~1.6 mM preS1 stock was titrated into 160 μM γ2-EAR, whereas for peptide-protein titrations, ~5 mM peptide was titrated into 500 μM γ2-EAR. Identical parameters were used for all of the experiments (20 injections, stirring at 1,000 r.p.m., 2-μl injections every 160 s). We used ‘double titrations’ to ensure binding satura-tion: a single run was followed by a second titration into the same cell solu-tion (after the first titration, the excess solution was aspirated from the dead volume above the ITC cell, and the syringe was refilled using exactly the same stock of titrant). The two titration curves were concatenated using ConCat32 software (GE Healthcare, PA). Reference titrations were performed to correct for heat of dilution effects. Concatenated, referenced titrations were analyzed using Origin7.0 (OriginLab Corp, MA).

X-ray crystallography. Wild-type γ2-EAR in 30 mM MES, pH 6.0, was concen-trated to ~600 μM before protein crystallization trials. Vapor diffusion crystal-lization screens were performed at 20 °C using a Mosquito robot (TTP Labtech Inc, MA). Diffraction-quality γ2-EAR crystals grew in 100 mM MES–imidazole, pH 6.5; 0.02 M 1,6-hexanediol; 0.02 M 1-butanol; 0.02 M (R,S)-1,2-propanediol; 0.02 M 2-propanol; 0.02 M 1,4-butanediol; 0.02 M 1,3-propanediol; 10% (w/v) PEG 20000; and 24% (v/v) PEG400 (using a 2:1 ratio of protein to mother liquor). Crystals were cryo-cooled in mother liquor using 30% (v/v) PEG 400 as cryoprotectant. Data for apo-γ2-EAR and the γ2-EAR–phage peptide crystals were collected in-house on a MicroMax rotating anode X-ray generator and R-axis IV++ image plate detector (Rigaku Americas Corporation, TX, wave-length 1.5418 Å). Data for the γ2-EAR/preS1(29–36) peptide crystal was col-lected on the EMBL–DESY beam-line P14 on a PILATUS 6M (25 Hz, 450-μm sensor thickness, wavelength 1.2395 Å) detector. All of the data were collected at 90 K. Data were anisotropic and had high mosaic spread (average 1.8°). Data were processed using XDS51 and Scala52. Initial phases were obtained by molecular replacement using the program BALBES53 (apo–γ2-EAR) or Phaser54 (γ2-EAR–peptide complexes), and initial building was performed by ARP/wARP55 implemented in the CCP4 suite56. The final structures were iteratively built and refined using Coot57, Refmac5 (ref. 58) and PHENIX59 and had good geometry (98% of all residues were in the favorable region of the Ramachandran plot) and fit to the electron density. Data and refinement statis-tics are given in Supplementary Table 2.

Electrospray ionization IMS-MS. All mass spectra and IMS-MS data were acquired using a Waters Synapt HDMS orthogonal acceleration quadrupole-traveling wave time-of-flight mass spectrometer (Micromass UK Ltd, UK). The source ionization and transmission voltages and instrumental pressures on the Synapt HDMS mass spectrometer were optimized to preserve noncovalent interactions and minimize protein unfolding36,60. No collision-induced dissoci-ation was observed. Samples were analyzed using nanoelectrospray ionization with platinum- or gold-plated borosilicate capillaries fabricated in-house using a P-97 micropipette puller (Sutter Instrument Company, CA) and a Polaron SC7620 sputter coater (Quorum Technologies, UK). A capillary voltage of 1.5 kV, cone voltage of 20 V, trap voltage of 6 V and transfer voltage of 4 V were used. The source pressure was maintained at 2.8 mbar with a trap flow rate of 0.1 ml/min. PreS1 and γ2-EAR protein samples (typically 10 μM) were electro-sprayed from solutions of 150 mM ammonium acetate (pH 8). All m/z spectra were calibrated using cluster ions generated from a separate introduction of cesium iodide solution (40 mg/ml aq.). The Synapt HDMS IMS device was cali-brated using denatured cytochrome c and myoglobin as described elsewhere36. Data were analyzed using MassLynx 4.1 software (Micromass UK Ltd, UK).

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