Structural studies of the eIF4E–VPg complex reveal adirect competition for capped RNA: Implicationsfor translationLuciana Coutinho de Oliveiraa,1,2, Laurent Volpona,1, Amanda K. Rahardjoa, Michael J. Osbornea,Biljana Culjkovic-Kraljacica, Christian Trahanb, Marlene Oeffingerb,c,d, Benjamin H. Kwoka, and Katherine L. B. Bordena,3

aInstitute of Research in Immunology and Cancer, Department of Pathology and Cell Biology, Université de Montréal, Pavilion Marcelle-Coutu, CheminPolytechnique, Montréal, QC H3T 1J4, Canada; bDepartment for Systems Biology, Institut de Recherches Cliniques de Montréal, Montréal, QC H2W 1R7,Canada; cDépartement de Biochimie et Médecine Moléculaire, Université de Montréal, Montréal, QC H3T 1J4, Canada; and dDivision of ExperimentalMedicine, McGill University, Montréal, QC H3A 1A3, Canada

Edited by Lynne E. Maquat, University of Rochester School of Medicine and Dentistry, Rochester, NY, and approved October 16, 2019 (received for reviewMarch 19, 2019)

Viruses have transformed our understanding of mammalian RNAprocessing, including facilitating the discovery of the methyl-7-guanosine (m7G) cap on the 5′ end of RNAs. The m7G cap is requiredfor RNAs to bind the eukaryotic translation initiation factor eIF4Eand associate with the translation machinery across plant and ani-mal kingdoms. The potyvirus-derived viral genome-linked protein(VPg) is covalently bound to the 5′ end of viral genomic RNA (gRNA)and associates with host eIF4E for successful infection. Divergentmodels to explain these observations proposed either an un-known mode of eIF4E engagement or a competition of VPg forthe m7G cap-binding site. To dissect these possibilities, we resolvedthe structure of VPg, revealing a previously unknown 3-dimensional(3D) fold, and characterized the VPg–eIF4E complex using NMR andbiophysical techniques. VPg directly bound the cap-binding site ofeIF4E and competed for m7G cap analog binding. In human cells, VPginhibited eIF4E-dependent RNA export, translation, and oncogenictransformation. Moreover, VPg formed trimeric complexes witheIF4E–eIF4G, eIF4E bound VPg–luciferase RNA conjugates, andthese VPg–RNA conjugates were templates for translation. Infor-matic analyses revealed structural similarities between VPg and thehuman kinesin EG5. Consistently, EG5 directly bound eIF4E in a sim-ilar manner to VPg, demonstrating that this form of engagement isrelevant beyond potyviruses. In all, we revealed an unprecedentedmodality for control and engagement of eIF4E and show that VPg–RNA conjugates functionally engage eIF4E. As such, potyvirus VPgprovides a unique model system to interrogate eIF4E.

VPg | m7 cap | potyvirus | translation | eIF4E

The eukaryotic translation initiation factor eIF4E plays importantroles in posttranscriptional control in plant and animals (1). Its

association with the methyl-7-guanosine (m7G) “cap” on the 5′ endof RNAs allows eIF4E to recruit transcripts to the RNA processingmachinery (2). To date, the m7G cap is generally accepted as theuniversal 5′ adaptor for RNAs in eukaryotes (3), with the exceptionof (i) the structurally related m3G cap, which is also used bynematodes (4), and (ii) with lower-frequency, nicotinamide ade-nine dinucleotide (NAD) and related analogs that destabilizetranscripts and thus, are probably not involved in active translation(5). Through its m7G cap-binding activity, eIF4E recruits specifictranscripts to the translation machinery in the cytoplasm andpromotes the nuclear export of selected RNAs from the nucleus (6,7). Both activities contribute to modulation of the proteome and inmammals, to its oncogenic activity (6, 7). For instance, eIF4E isdysregulated in many human cancers (6). In humans, targetingeIF4E with a cap competitor, the guanosine analog ribavirin,impairs its biochemical activities correlating with clinical re-sponses in early-phase trials in leukemia, prostate, head, andneck cancers among others (8–13). Thus, the cap-binding activity

of eIF4E can be targeted in patients to provide clinical benefit,highlighting its critical importance.Viruses have paved the way for our understanding of many

aspects of host-cell RNA processing, including m7G capping.Indeed, studies into cytoplasmic polyhedrosis virus (CPV) infec-tion in silkworm and vaccinia virus (VV) in mammalian cells werecritical for the elucidation of the m7G cap structure over 40 y ago(3, 14, 15). Here, we exploited unusual features of potyvirus bio-chemistry to unearth unknown strategies that can be implementedto engage eIF4E. Potyviruses are members of the picorna-likeplant viruses. Their infection of mainstay crops has devastatingeconomic consequences (16). Genetic studies revealed that poty-viruses require host-cell translation machinery to replicate, andspecifically, these reports have associated the potyviral proteingenome linked (viral genome-linked protein [VPg]) with host planteIF4E (17–21). Indeed, mutations in plant eIF4E are associated

Significance

RNA processing including covalent modifications (e.g., the ad-dition of the methyl-7-guanosine [m7G] “cap” on the 5′ end oftranscripts) centrally influences the proteome. For example,eIF4E recruits RNAs for translation by binding the m7G cap.eIF4E is engaged and controlled by the binding of factors to itsdorsal surface while leaving its m7G cap-binding site free forRNA recruitment. Here, we unexpectedly found that a smallviral protein, viral genome-linked protein (VPg), directly bindsthe cap-binding site of eIF4E, indicating that eIF4E can addi-tionally be controlled through direct competition with its cap-binding site. Furthermore, VPg–RNA conjugates also bind eIF4Eand are templates for translation, suggesting that VPg maysubstitute for the m7G cap during infection.

Author contributions: L.C.d.O., L.V., M.J.O., B.C.-K., C.T., M.O., B.H.K., and K.L.B.B. de-signed research; L.C.d.O., L.V., A.K.R., M.J.O., B.C.-K., C.T., and K.L.B.B. performed re-search; L.V., M.J.O., B.C.-K., B.H.K., and K.L.B.B. contributed new reagents/analytic tools;L.C.d.O., L.V., A.K.R., M.J.O., B.C.-K., C.T., and K.L.B.B. analyzed data; and L.C.d.O., L.V.,M.J.O., B.C.-K., C.T., M.O., and K.L.B.B. wrote the paper.

The authors declare no competing interest.

This article is a PNAS Direct Submission.

Published under the PNAS license.

Data deposition: The NMR, atomic coordinates, chemical shifts, and restraints reported inthe paper have been deposited in the Biological Magnetic Resonance Data Bank (http://www.bmrb.wisc.edu/; accession no. 27506) and the Protein Data Bank (https://www.rcsb.org; ID code 6NFW).1L.C.d.O. and L.V. contributed equally to this work.2Present address: NMX Research and Solutions Inc., Laval, QC H7V 5B7, Canada.3To whom correspondence may be addressed. Email: katherine.borden@umontreal.ca.

This article contains supporting information online at www.pnas.org/lookup/suppl/doi:10.1073/pnas.1904752116/-/DCSupplemental.

First published November 11, 2019.

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with potyviral resistance (18–20). VPgs exist in other virus families,such as poliovirus (22). The VPg designation is based on the co-valent linkage of viral RNA to VPg. For the case of potyviruses,the 5′ end of the genomic RNA (gRNA) is covalently attached tothe hydroxyl of tyrosine 64 (potato virus Y [PVY] numbering) (23–25). The genetic interaction between VPg and eIF4E is onlyreported for potyviruses (20), while other virus families typicallyuse these RNA conjugates for replication (22). Consistent withthis, potyviral VPgs only show significant sequence homology witheach other and not with VPgs from other families (SI Appendix,Fig. S1).While genetic studies linked VPg and eIF4E, conclusions from

biochemical studies were highly divergent, leaving the mecha-nism as to how VPg coopts eIF4E activity unsettled. Some groupsreported that PVY VPg binds m7G cap–eIF4E–eIF4G, forminga quaternary complex, which suggests that VPg utilizes a novelsurface on eIF4E for binding and thereby, engaging its activity (17,26). Supporting this model, mutation of the cap-binding site inwheat eIF4E (W123A, W102 in human eIF4E) did not reduce theability of eIF4E to bind VPg but did reduce binding to the 7-methylguanosine diphosphate (m7GDP) cap analog. This sug-gested that VPg bound to a part of eIF4E not previously known tobe involved in its control or engagement (26). By contrast, anotherstudy provided evidence that PVY VPg directly competes for thecap-binding site on plant eIF4E (21). Other reports suggested thatPVY VPg interacts with the eIF4F (eIF4E–eIF4G–eIF4A/B)complex to stimulate cap-independent internal ribosomal entrysite (IRES)-mediated translation, but the exact eIF4F compo-nent required was not ascertained, and whether IRES-mediatedtranslation is relevant to potyviruses is not clear (27). Just as themode of binding to eIF4E is controversial, there are divergentmodels regarding the molecular basis for the VPg–eIF4E in-teraction. One study proposed that residues 41 to 93 of PVY VPgwere used for binding to eIF4E (21), while others reported thatresidues on a predicted long amphipathic helix spanning residues90 to 125 were required (28). Efforts to solve the VPg structurehad been to date unsuccessful. Modeling efforts yielded disparatesolutions, including a helical bundle (29), a long amphipathic helixfor the VPg-binding site for eIF4E (28), and a model based on theFOK1a kinase structure, which included a β-sheet core with flex-ible helices (30). Adding to the confusion, multiple biophysicalstudies proposed that VPg is an intrinsically disordered protein(21, 30–33).While the mechanistic underpinnings of the interplay between

VPg and eIF4E remains controversial, it is clear that eIF4E isrequired for infection and that there is a genetic interaction be-tween eIF4E and VPg. In this way, VPg provides a unique modelsystem to interrogate the modalities required for the engagementand control of eIF4E. No structure of any potyvirus protein exist.Here, we report the structure of the potyvirus protein, VPg, andcharacterized the VPg–eIF4E complex using high-resolution NMRand biophysical methods. We demonstrated that VPg binds thecap-binding site of eIF4E and forms trimeric complexes witheIF4E and eIF4G; furthermore, we demonstrated that VPg–RNAconjugates directly bind eIF4E and were templates for translationconsistent with the requirement of potyviruses for VPg–eIF4Einteractions for their lifecycle. Furthermore, the structural simi-larities of VPg to human proteins suggest that this modality forengagement of eIF4E could be conserved across kingdoms.

Results and DiscussionVPg Adopts a Previously Unknown Protein Fold. We determined thestructure of PVY VPg protein in order to understand its mo-lecular relationship with eIF4E and thus, its role in RNA re-cruitment to the host-cell translation machinery. We studied PVYVPg, since it is the archetypal potyvirus and thus, representative ofthe family (SI Appendix, Fig. S1B). We generated full-length andtruncated forms of PVY VPg protein using bacterial expression

constructs and purified these from the soluble fraction with thequality of the proteins confirmed by SDS/PAGE (Sodium DodecylSulphate-PolyAcrylamide Gel Electrophoresis) (Fig. 1C) (34),NMR (Fig. 1D and SI Appendix, Fig. S2), and mass spectrometry(MS) (SI Appendix, Fig. S3A). We recently reported the NMRassignments for a VPg construct in which the first 37 residues wereremoved to improve stability (VPgΔ37) (34). Here, the 3-dimensional (3D) solution structure of this construct was de-termined by using an automated procedure for iterative nuclearOverhauser effect (NOE) assignment using CYANA (35). Thestructure of VPg is shown in Fig. 1 A and B and SI Appendix, Fig.S3B, and the structural statistics are in SI Appendix, Table S1. Thermsd for the ordered regions was 0.68 Å for the backbone atomsof the top 20 structures.The VPg structure is unlike the previously proposed models.

VPgΔ37 adopts a well-folded core as well as 2 substantial un-structured regions at the N and C termini (residues 38 to 70) and aflexible loop between β4- and β5-strands (residues 145 to 165) (Fig.1A and SI Appendix, Fig. S3B). We note that VPg is not anintrinsically disordered protein. Specifically, there were many long-range NOEs, indicating a tertiary structure (SI Appendix, TableS1), and furthermore, values for the 15N-1H heteronuclear NOEand chemical shift index also indicated the presence of struc-tured elements (34). The VPg structure is composed of a 5-stranded β-sheet with 2 consecutive α-helices between β-strands2 and 3 (Fig. 1A). Our structure of PVY VPg does not resembleVPg structures from other virus families, consistent with the lackof sequence conservation observed (SI Appendix, Fig. S1A). ResidueY64, which is covalently attached to gRNA during infection (23–25),is located within the flexible N terminus but close to the foldeddomain, which starts at residue 72 (Fig. 1A and SI Appendix, Fig.S1B). Inspection of the structure revealed that there were no ele-ments within VPg that possessed any structural similarity to reportedeIF4E-binding motifs (e.g., the helical turn describing the eIF4Econsensus motif [YXXXXLΦ, where X is any residue and Φ is anyhydrophobic] or RING (really interesting new gene) domains [6]).

Elucidation of the eIF4E Binding Site on VPg. We used NMR andpulldown studies to garner information regarding the eIF4E–VPg complex structure (see below). We selected human eIF4Efor these studies because of the extensive knowledge accumu-lated regarding its structure, allosteric effects of ligands, and dy-namics (36–38). Importantly, human cap-bound eIF4E is highlyhomologous in sequence (SI Appendix, Fig. S4A) and structure (SIAppendix, Fig. S4B) to the 3 cap-bound eIF4E plant structuressolved (rmsd ∼ 0.7 Å for 175 atom pairs). Both human and meloneIF4E structures were available in the apo- and cap-bound forms aswell as ternary complexes with eIF4G (38, 39). These structures arehighly homologous, indicating that these were conserved acrosskingdoms. The large positive surface used to bind the phosphates ofthe m7G cap in the unoccupied cap-binding site was conservedacross kingdoms (SI Appendix, Fig. S4C). Given the overall simi-larity between structures coupled with the deeper structural un-derstanding of human eIF4E, we studied the VPg–human eIF4Ecomplex to ascertain how VPg engaged and/or con-trolled eIF4E.We identified the VPgΔ37 and eIF4E interface using a com-

bination of NMR-based strategies and mutagenesis. We used cap-free eIF4E as a starting point for these studies. First, using 50 μM15N-,13C-, or specifically labeled Ile, Leu and Val methyl groups(ILV) VPgΔ37, we monitored the effects of addition of unlabeledeIF4E (150 μM) on signal broadening and chemical shift pertur-bation (CSP) of the amide nitrogens, methyl carbons, and carbonylgroups using the 1H-15N HSQC (heteronuclear single quantumcoherence), the methyl region of the 1H-13C spectra, or the H/Cprojection of the 3D-HNCO experiment, respectively (Fig. 2 Aand B and SI Appendix, Fig. S5). We observed substantial broad-ening for residues located in the loop E108-G119 of VPgΔ37, andin particular, for carbonyls of E108, R109, and Q116 and amides of

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R109, D111, I113, and M115-G119, including the side-chain amideof Q116 (Fig. 2 C and E). Methyl carbons in the region V103-L118 also exhibited broadening (Fig. 2B) in addition to residuesneighboring this loop (L80 and L166). The flexible regions ofVPgΔ37 (i.e., the large loop [N145-E165] and both N [S38-E70]and C termini [A182-E188]) were not altered by eIF4E, indicatingthat they are unlikely to be involved in the interaction. Consistentwith this observation, deletion of the first 62 residues in VPg didnot impair binding to eIF4E (SI Appendix, Fig. S3 C and E).Second, we used transferred cross-saturation (TCS) experiments

(40) to directly detect through space interactions between eIF4Eand VPg. In this experiment, the labeled VPg was in 4-fold excessrelative to unlabeled eIF4E (90 μM eIF4E, 360 μM VPg). Con-sistent with the above NMR data, we observed substantial re-ductions in signal intensity of the amide side chain of residueQ116 and to a lesser extent, of the backbone amides of D111 andE114 (Fig. 2D), indicating that these residues were close in spaceto eIF4E. We then used mutagenesis to validate the NMR data.VPgΔ37 mutants M115A/Q116A and D111K/E114K/Q116K hadweaker affinity for eIF4E compared with wild-type VPgΔ37 asobserved by increased intensities of the NMR signals (SI Appendix,Fig. S6). These spectra were acquired under identical concentra-tions and ratios; thus, differences in intensities in comparingspectra directly reflect affinity. To support the delineation of thebinding surface, we also mutated residues outside of the predictedbinding site and examined their impact. Consistently, these mu-tations (E86K/E87K, D92K, or E98K/E102K) did not significantlyimpact the VPg–eIF4E interaction (SI Appendix, Fig. S6D). Noneof the mutants described above disrupted the VPg structure asobserved by NMR or circular dichroism (CD) (SI Appendix, Fig.S3D). Altogether, TCS, CSP, signal broadening, and mutationalstudies indicate that the α1–α2 loop forms the surface on VPg thatbinds to eIF4E.

Mapping of the VPg Binding Site on eIF4E. We used the samestrategy to determine the binding site of VPgΔ37 on eIF4E. Wemonitored spectral perturbations of 50 μM 15N- or 13C-eIF4Esamples as a function of unlabeled VPgΔ37 addition to a maxi-mum of 150 μM eIF4E (Fig. 3 and SI Appendix, Fig. S7). The most

substantial broadening was detected around the cap-binding site,particularly for backbone residues in the F48-L60 loop (Fig. 3C)which also contains W56, one of the tryptophan residues that bindsthe m7G cap. Other proton amides around the cap-binding pocketof eIF4E were also broadened (i.e., the K95 and the H200). Asshown for VPgΔ37 (Fig. 2D), the empty spaces indicate residuesthat were not quantified due to spectral overlap (Fig. 3C). Con-sistently, we also observed carbon chemical shifts for methylgroups located directly behind the phosphate-binding region ofthe cap-binding site of eIF4E (I63 and L85) (SI Appendix, Fig. S7).Given that these changes were concentrated in the cap-bindingsite, we individually mutated 2 parts of this site: the m7G moiety-binding region, which includes W56, and the phosphate-bindingregion, which includes R157, K159, and K162 that form apositively charged patch. The R157E/K159E/K162E triple mutation(4ETrMut) substantially reduced the interaction of eIF4E withVPgΔ37 as did the W56A mutation (SI Appendix, Figs. S8 A–Cand S9). These findings were confirmed by glutathione S-trans-ferase (GST) pulldown experiments using murine eIF4E, which isonly 4 residues different from human eIF4E (SI Appendix, Fig.S8E). Importantly, none of these mutants altered the overall foldof eIF4E as assessed by HSQC and CD (SI Appendix, Fig. S8 A–D). Importantly, W56 and R157/K159/K162 comprise one endof the cap-binding site and are relatively close in space in boththe apo- and cap-bound eIF4E forms (<11 Å). We note that theW102A mutation, allied to W123A in wheat, barely affects theability of eIF4E to bind VPg (SI Appendix, Fig. S9). In apo-eIF4E, W102 is at the other end of the cap-binding site, whileupon cap binding, this residue moves to be in close proximitywith W56, R157, K159, and K162 (38). In all, our NMR datastrongly indicate that the VPg-binding site on eIF4E is clusteredin the region of the cap-binding site, including W56, R157, K159,and K162.

The VPg–eIF4E Complex.We used the above NMR and mutagenesisdata to generate a model of the eIF4E–VPgΔ37 complex usingthe restraint-driven docking program HADDOCK (41). We notethat the intermediate NMR exchange regimen observed for thecomplex meant that it was impossible to observe NOEs due to

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Fig. 1. Structure of the PVY VPg protein. (A) Cartoon representation of the closest to the average structure in the ensemble for VPgΔ37. The family of best20 structures is shown in SI Appendix, Fig. S3B. Residues starting at F60 shown as residues 37 to 70 are disordered. (B) Surface rendering of VPg structure. Redindicates negative charged area, blue indicates positive, and white are hydrophobic. (C) SDS/PAGE gel of full-length (FL) and VPg truncation constructs usedfor NMR analysis; molecular mass markers are shown. (D) 1H-15N HSQC spectrum of VPgΔ37.

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broadening of signals for residues at the interface. The resultingHADDOCK calculation led to a solution where 150 of 200 struc-tures converged to the same eIF4E–VPgΔ37 complex. Of these150 structures, the backbone rmsd for the complex was ∼0.8 Å,

and the buried surface area for the complex was ∼1,960 Å2 (Fig.4A). In this complex, W56 is buried in a hydrophobic pocketformed by M107, V108, M115, and L118 in the helix–loop–helixstructure of VPg. Also, the positively charged residues (R157 andK162) in the phosphate-binding pocket of eIF4E are facing D111and D112 of VPg (Fig. 4B).To independently validate our model, we performed cross-

linking mass spectrometry (XL-MS) on cross-linked heterodimersthat were isolated by size exclusion chromatography. Accordingto our model, only 3 lysines of VPg are located near the interfaceof the heterodimer (K105, K106, and K138), and hence, a lownumber of cross-links between both proteins was expected. Fur-thermore, most of the cross-linker was absorbed by intramolecularcross-link between VPg K105-K106 (extracted ion chromatogram[XIC] values in SI Appendix, Table S2), thus contributing greatly toa reduction of interprotein cross-link occurrences between VPgand eIF4E. Nevertheless, consistent with our HADDOCK model,the most frequently observed cross-link (15 occurrences), whichalso had the highest score (5.33), bridged K106 of VPgΔ37 withK159 of eIF4E, for which the Euclidean distance measured by theXwalk software (42) was found to be 16.0 Å (SI Appendix, TableS2). This most frequent cross-link was consistent with the magni-tude of line broadening in the different HSQCs, TCS, and muta-genesis data defining the binding sites for both eIF4E and VPg.The second most frequent interprotein cross-link only occurred4 times and was found between eIF4E–K192 and VPg–K47, thelatter of which is in a highly flexible region of VPg. The presence ofthis cross-link is consistent with previous studies that showed thatthe N terminus of VPg was involved in binding to eIF4E (21).Finally, a cross-link between eIF4E–K192 and VPgΔ37–K138 wasobserved but only with 2 occurrences. As observed in the apoform of eIF4E, the K192-containing loop and a fortiori, its sidechain are flexible (Protein Data Bank [PDB] ID code 2GPQ),positioning these 2 lysines between 25 and 37 Å apart (calcu-lated by the Xwalk software). Thus, in some conformations, theselysines would be accessible. Taken together, the XL-MS data sup-port our HADDOCK model based on NMR and mutagenesisrestraints.The VPg–eIF4E complex presented here provides a molecular

basis for understanding the genetic studies of eIF4E and VPg. Thepositions of eIF4E mutants that impart resistance to potyvirus

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Fig. 2. The binding surface on VPg used to interact with eIF4E. (A) 1H-15NHSQC of 50 μM 15N-labeled VPg in the absence (red) or presence (blue) of a3-molar excess of unlabeled eIF4E. (B) Constant time 1H-13C HSQC spectrum ofILV-labeled VPg (50 μM) in the absence (red) or presence of 2-molar excesseIF4E (blue). (C) VPg residues are perturbed by eIF4E binding. Light blue indi-cates 1H CSP or broadening, dark blue indicates 1H methyl CSP, and yellowindicates broadening from the 1H-13C projection of the 3D HNCO spectra (SIAppendix, Fig. S5). (D) Per residue plot of backbone amide line broadening of15N-labeled VPgΔ37 in response to binding of eIF4E (extracted from A). Resi-dues that undergo line broadening below the dashed line are in cyan. Notethat empty spaces correspond to residues that overlap, residues that are notassigned, or Proline residues. (E) Plot of the intensity ratios of the cross-peaks inthe TCS experiment for the backbone (blue) and asparagine/glutamine side-chain (orange) resonances for VPg.

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Fig. 3. Interaction surface used by eIF4E to bind VPg. (A) 1H-15N HSQC of15N-labeled eIF4E (50 μM) in the absence (red) or presence (blue) of 3-molar excessVPg. (B) Broadening and CSPs mapped onto the apo-eIF4E structure (PDB ID code2GPQ) depicted as light blue balls. (C) Per residue plot of backbone amide linebroadening of 50 μM eIF4E in response to binding of VPgΔ37 (extracted from A).Residues that undergo line broadening below the dashed line are in cyan.

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infection in plants are mapped onto our eIF4E–VPg complexstructure (SI Appendix, Fig. S8F). These mutants are generallylocated in the loop containing W56 (using human eIF4E aminoacid numbering) or in the nearby phosphate-binding site (43, 44),which overlays with the proposed VPg-binding site. Interestingly,the engineered wheat W123A mutant (or human W102A), whichdid not disrupt the eIF4E–VPg interaction (27) (SI Appendix, Fig.S9), was found in the upper part of the cap-binding site and wasnot included in the experimentally defined eIF4E-binding surfaceused to interact with VPg, but it is clearly important for m7G capbinding. Additionally, the VPg residues, which interacted witheIF4E, were consistent with substitutions in this region beingcritical for viral pathogenicity (S101-N121 of PVY–VPg) (SI Ap-pendix, Fig. S8G) (43–45).We next determined whether VPgΔ37 bound plant eIF4E in a

similar manner to human eIF4E. Previous biophysical studiesshowed that wheat eIF(iso)4E bound to VPg, and hence, wefocused on this homolog for our studies (27, 46). Given that therewere no NMR assignments available for eIF(iso)4E, we exam-ined the interaction via 1H-15N HSQCs using 35 μM 15N-labeledVPgΔ37 with unlabeled eIF(iso)4E (105 μM). We found that VPgused the same binding region (I113-N121) to bind eIF(iso)4E (SIAppendix, Fig. S10 B and C) as it used to associate with humaneIF4E (Fig. 2), indicating that this was conserved across kingdoms.Superimposing our homology model of wheat eIF(iso)4E onto thewheat eIF4E structure revealed that the orientation of the set ofpositive residues (corresponding to the R157, K159, and K162)and W56 (human numbering) was the same in plant and humaneIF4Es (SI Appendix, Figs. S4 and S10A). Thus, the basic princi-ples for the VPg–eIF4E interaction seem evolutionarily conserved.

VPg Binds eIF4E in the Presence of an eIF4G Peptide. eIF4G is amajor platform protein in the translation initiation complex,recruiting not only eIF4E–RNA complexes but other cofactors to

the small ribosomal subunit in order to initiate translation (47, 48).To determine whether eIF4E–VPg complexes bind eIF4G, wecarried out NMR experiments where VPgΔ37 was added to15N-labeled eIF4E in complex with an eIF4G peptide containing theconsensus binding site (4Gp). Both VPgΔ37 and 4Gp were used ina 3-fold molar excess (150 μM) compared with eIF4E (50 μM);4Gp binds on the dorsal surface of eIF4E (SI Appendix, Fig.S11D), a region distal to the cap-binding and VPg-binding pocketin eIF4E. Interaction of both partners affects eIF4E’s signals dif-ferently in the HSQC, serving as reporters for binding. Indeed, onVPgΔ37 addition, most signals were broadened (SI Appendix, Fig.S11 B and F), while 4Gp addition induces peak shifting only forresidues in the proximity to its binding site on eIF4E. In particular,addition of 4Gp perturbed residues W73, Y34, I35, K36, L75,and L137 on the dorsal surface of eIF4E as expected (SI Ap-pendix, Fig. S11 A and E and Fig. 5C), which were not alteredby VPgΔ37 binding. Overall, on addition of 4Gp to the eIF4E–VPgΔ37 complex, we observed 2 phenomena (SI Appendix, Fig.S11 C and G): 1) signals remain broad, indicating that VPgΔ37 wasstill bound to eIF4E, and 2) residues close to the 4Gp-binding site(see above) were shifted similarly as in the eIF4E/4Gp complex.Thus, our NMR data analysis revealed that eIF4E associates with a4Gp and VPgΔ37 simultaneously.

m7G Cap Analogs Compete for VPg Binding to eIF4E.Given that VPgand the m7G cap analogs bound overlapping surfaces on eIF4E,we explored whether the cap analog m7GDP and VPgΔ37 competedfor binding of 15N-eIF4E by HSQC experiments. Addition of20-fold excess m7GDP to preformed VPgΔ37–eIF4E complexes(50 μM eIF4E, 150 μM VPgΔ37, 1 mM m7GDP) led to thereemergence of eIF4E resonances but now in their m7GDP-boundpositions (Fig. 4C and SI Appendix, Fig. S12 A–C). This indicatedthat the m7GDP cap analog and VPgΔ37 competed for eIF4E onthe same site. Importantly, VPgΔ37 did not bind to m7GDP capitself as demonstrated by the observation that HSQC spectra of15N-labeled VPg protein were unchanged in the presence or ab-sence of the m7GDP (SI Appendix, Fig. S13D). Thus, the m7GDPcap analog competed for VPg binding in vitro. This was confirmedusing the reverse titration in which we monitored changes to the15N transverse relaxation-optimized spectroscopy (TROSY)HSQC of 2H-15N VPg (50 μM) on addition of unlabeled eIF4E(up to 175 μM) in the absence and presence of 1 mM m7GDP.Residues at the proposed VPg-binding site (D111, I113, M115,Q116, L118, G119, and N121) were substantially broadenedupon eIF4E addition to VPg (SI Appendix, Fig. S13A). How-ever, in the presence of 1 mM m7GDP (SI Appendix, Fig. S13 Band C), these peaks reappeared, indicating that m7GDP competesfor the same binding site on eIF4E as VPg. The same experimentwas performed with wheat eIF(iso)4E and yielded equivalentresults (SI Appendix, Fig. S10D). Thus, VPg utilized the cap-binding site on both human and plant eIF4E for recognitionand competed for cap analogs.To gain a quantitative understanding of this interaction, we

determined the dissociation constants for eIF4E–VPg and eIF4Ecap analogs. Unfortunately, NMR titrations of the eIF4E–VPgcomplex via 1H-15N HSQC techniques resulted in disappearanceof peaks, making determination of Kd values challenging for avariety of technical reasons, including the inability to obtain in-formation of the completely broadened state where binding wouldbe saturated. Using isothermal calorimetry (ITC), we determinedthe Kd for the human eIF4E–m7GDP cap analog. Consistent withprevious results in phosphate buffer used in the above NMRexperiments, we obtained a Kd of 0.37 ± 0.02 μM, which isconsistent with previous literature reports for this interac-tion in phosphate buffer (49). When evaluating the Kd for theeIF4E–VPgΔ37 interaction by ITC using the same bufferconditions, we observed no heat release, suggesting that theinteraction is mainly entropic. Thus, we used a fluorescence

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assay, whereby VPg was labeled with N-(iodoacetyl)-N′-(5-sulfo-1-naphtyl)ethylenediamine (IAEDANS) (33), an organicfluorophore that fluoresces at 490 nm, a region where the VPgand eIF4E proteins produced no signal. We observed a Kd of0.26 ± 0.07 μM (Fig. 4D) and a Hill coefficient of ∼1.1, in-dicating a 1:1 ratio of VPg–eIF4E complexes, consistent withour NMR studies. This affinity is very similar to literaturevalues for wheat eIF4E and VPg (∼0.3 μM) (27). Thus, theaffinities of eIF4E for VPg and the m7G cap analog were verysimilar, consistent with the competition that we observed in ourabove NMR studies.

VPg–RNA Conjugates Directly Bind eIF4E and Were Templates forTranslation. Our above studies demonstrated that VPgΔ37 com-peted for m7G cap binding to eIF4E and in this way, could in-terfere with host-cell translation. However, they also suggestedthat VPgΔ37–RNA conjugates engaged eIF4E–eIF4G complexespossibly to be recruited to the translation machinery. As a first stepto test this possibility, we investigated whether RNA conjugationdisrupted the interaction with eIF4E or alternatively, if VPg–RNAconjugates could directly bind to eIF4E. During infection, VPgΔ37is conjugated through the side-chain hydroxyl group of Y64 to the5′ end of the viral gRNA (23–25). Y64 is in a flexible region distalto the eIF4E-binding site, consistent with the possibility thatVPgΔ37 interacts with eIF4E and RNA simultaneously (Fig. 4A).Unfortunately, potyvirus VPg–gRNA conjugates are not availablein purified forms to directly investigate this with the viral gRNA.Using maleimide chemistry (50), we thus conjugated a 19-merfragment of luciferase RNA onto VPgΔ37 (SI Appendix, Fig.S14A). In this case, we generated VPgΔ37 Y64C for conjugationto the 5′ end of the RNA. In this same construct, we mutated theonly naturally occurring cysteine C150, which is found in theflexible loop, to alanine in order to ensure a single conjugation sitefor the protein. This protein was folded as the wild type and bindseIF4E similarly to wild-type VPgΔ37 (SI Appendix, Fig. S14B).The 19-mer RNA segment with a maleimide group on its 5′ endwas then cross-linked to the sulfur side chain of C64 (SI Appendix,Fig. S14A). We verified that the 19-mer RNA was conjugatedusing SDS/PAGE and silver staining, where the VPgΔ37(C150A/Y64C)–RNA conjugate was ∼6 kDa larger as expected. We fur-ther confirmed the presence of RNA by treating the conjugatewith RNase (ribonuclease), which resulted in a reduction in sizecorresponding to the unmodified form of VPgΔ37(C150A/Y64C)(Fig. 5A). Similarly, treatment with proteinase K also disrupted theconjugate, leaving only the free 19-mer RNA band. Using eIF4E–GST pulldown, we observed that the VPgΔ37(C150A/Y64C)–RNA conjugate bound eIF4E but not the GST control (Fig. 5B).Thus, VPg recruited RNA to eIF4E.Given the above findings, we examined the possibility that

VPg–RNA conjugates were templates in in vitro translation as-says. To produce conjugates with full-length messenger RNAs(mRNAs), we had to alter our strategy in order to conjugate full-length luciferase RNAs (∼1,800 nucleotides) to VPgΔ37(C150A/Y64C), which yielded a species of ∼500 kDa. Using in vitrotranscription, guanosine-5′-monophosphorothioate (GMPS) wasincorporated into the 5′ end of luciferase transcripts and sub-sequently coupled to 2,2′-pyridine disulfide using standardmethods (51, 52). A disulfide exchange reaction of the resultingpyridyl-disulfide linkage on the 5′ end of the RNA was usedfor conjugation to VPgΔ37(C150A/Y64C) (53). To monitor theefficiency of conjugation, VPg–RNA conjugates were subjected toagarose gel electrophoresis due to their large size followed byimmunoblotting (54) for the His tag of VPg (SI Appendix, Fig.S14C). Unconjugated VPgΔ37(C150A/Y64C) is shown for com-parison. As observed, all of VPgΔ37(C150A/Y64C) was conju-gated to the luciferase RNA, and no unconjugated RNA wasdetected after the reaction. For comparison, we generatedluciferase transcripts using in vitro transcription without any

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Fig. 5. (A) Silver-stained SDS/PAGE gel showing the conjugation betweenVPgΔ37 (Y64C, C150A) and the 19-mer RNA with a 5′ maleimide group. Noβ-mercaptoethanol or DTT (dithiothreitol) was present in order to preserve theconjugate. VPg, the RNA fragment, and the conjugated form are shown (lanes1, 2, and 4, respectively). Validation of the conjugate is shown by treatmentwith either RNase A (lane 5) or proteinase K (Prot.K; lane 6). The 19-mer RNA isdiffuse because of the percentage of acrylamide used. Position of the RNase Aprotein is shown in lane 3 for comparison. We note the presence of dimericforms through C64 in the absence of reductant. (B) Association of the VPg–RNAconjugate with GST–eIF4E but not GST alone by western blot using an anti-Histag antibody (Upper). As expected, the VPg dimer also binds GST–eIF4E. Anasterisk is shown to highlight a VPg–RNA degradation product that occurredduring the GST pulldown. GST loading is shown below by western blot usingan anti-GST antibody. The samples were run on the same gel, with unrelatedsamples removed for clarity. (C) Model representing eIF4G (orange; PDB IDcode 5T46) derived from the crystal structure of the eIF4E–eIF4G complex withthe VPgΔ37 (green) and eIF4E (blue) as displayed in Fig. 4A. The Y64, which iscovalently attached to gRNA during infection, is shown. Similarly, the VPg–RNAconjugates made in vitro were also at the same position, C64 (in the text). (D)Western blot for Luciferase protein produced in in vitro translation reactionsusing wheat germ lysates and different luciferase RNAs: conjugated to VPg(VPg–RNA conj), uncapped, m7G-capped, and m7G-capped luciferase RNA inthe presence of 10 μM VPg protein (capped RNA + VPg protein). Loading ofdifferent luciferase RNAs was confirmed by qRT-PCR (SI Appendix, Fig. S14D).(E) Quantification of western blots for in vitro translation assays described in D.Data were derived from 3 independent experiments. Mean intensities ± SDs areshown. P values are from Student’s t test (*P < 0.05, **P < 0.01, ***P < 0.001).

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modifications (referred to as uncapped) and also, generatedcapped luciferase transcripts using the VV-capping enzyme.Equal amounts of each luciferase RNA, confirmed by qRT-PCR(SI Appendix, Fig. S14D), were used as templates for in vitrotranslation assays, and the production of luciferase protein wasmonitored by western blot using an antiluciferase antibody withthe experiments carried out 3 independent times (Fig. 5 D and E).We observed ∼2-fold more luciferase protein produced fromm7G-capped luciferase RNA templates than uncapped templates asexpected. The uncapped templates provided a lower bound forbackground translation, where it is well established that translationof uncapped RNAs occurs in in vitro systems but less efficientlythan when RNAs are m7G capped (55). The levels of translationfor VPg-capped luciferase transcripts were nearly identical tom7G-capped RNAs and ∼2-fold higher than observed for uncappedRNA (Fig. 5 D and E). This demonstrated that the VPg conju-gation to the luciferase RNA did not interfere with its translation,indicating that VPg–luciferase RNA conjugates were templatesfor translation. Moreover, VPg–luciferase conjugates were trans-lated with the same efficiency as capped RNAs, suggesting thatVPg could functionally substitute for the m7G cap. These obser-vations are consistent with our identification of VPg–eIF4E–eIF4G complexes (Fig. 6) and VPg–RNA–eIF4E complexes(Fig. 5). We note that the dynamic range of our assay was limited(2-fold between capped/VPg relative to uncapped RNA). Finally,the addition of free VPg (i.e., not conjugated to the RNA) reducedtranslation, consistent with our model of cap competition andprevious reports (49).

VPg Suppressed Cap-Dependent eIF4E Activities in Cells. Next, weexplored the impact of VPg on eIF4E activity in human cells todetermine if the effects observed in vitro were recapitulated incellulo. We postulated that VPg without RNA would potentlyinhibit cap-dependent activities of endogenous eIF4E in humancells by competing for host-cell transcripts. To test this hy-pothesis, we generated stable human osteosarcoma U2Os celllines expressing either full-length VPg–FLAG or vector controls.U2Os cells were selected based on their well-characterized eIF4Eactivities (6, 56, 57). We investigated the effects of VPg on thenuclear functions in mRNA export and cytoplasmic functions intranslation, because both functions require the ability of eIF4E tobind the m7G cap (6, 7).We first examined translational efficiency using polysomal

analysis in VPg–FLAG cells vs. vector controls. We monitoredtranslation and RNA export of 2 well-characterized target tran-scripts of eIF4E: c-Myc and MCL1 (56, 58). VPg overexpressiondid not alter the overall polysomal profile, indicating that it did notinterfere with the formation of ribosomes (Fig. 6A). However, VPgreduced translational efficiency for both MCL1 and c-Myc tran-scripts; in contrast, it did not alter the total RNA levels for eitherof these transcripts, and it did not affect the translation efficiencyof Actin, an RNA insensitive to eIF4E (Fig. 6A and SI Appendix,Fig. S15A). We next explored whether the RNA export activity ofeIF4E, which is also cap dependent, was impaired in VPg–FLAGcells relative to vector controls (Fig. 6B). Cells were fractionatedinto nuclear and cytoplasmic compartments, and RNAs werequantified by qRT-PCR (quality of fractionation was verified bysemi-qPCR) (SI Appendix, Fig. S15C). VPg–FLAG overexpressionimpaired nuclear export of eIF4E target transcripts (e.g., MCL1and c-Myc) by ∼2-fold but not eIF4E-independent transcripts, suchasGAPDH and VEGF RNAs. Again, total levels of RNA were notaltered for any of the examined transcripts as expected (SI Ap-pendix, Fig. S15A). Consistent with VPg inhibiting both export andtranslation of these RNAs, protein levels for MCL1 and c-Mycwere substantially decreased in VPg–FLAG cells relative to vec-tor controls, while β-Actin protein levels were unchanged (Fig.6C). This is also consistent with previous studies in plants thatshowed that potyvirus infection inhibited translation of specific

RNAs (59), likely because only a subset of these is sensitive toeIF4E inhibition with other compensatory mechanisms cominginto play, such as eIF3d-mediated cap-dependent translation forexample (60). Thus, VPg inhibited activity of endogenous eIF4E.To further dissect the role of VPg in eIF4E-dependent trans-

lation in cells, we explored whether VPg, via eIF4E, also interactedwith endogenous eIF4G (Fig. 6D). We immunoprecipitated VPg–FLAG using an FLAG antibody and compared this with vectoror eIF4E–FLAG-expressing cells. We observed that VPg–FLAGimmunoprecipitated with endogenous eIF4E, endogenous eIF4G,

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Fig. 6. VPg represses eIF4E function in human cancer cells. (A) Polysomeanalyses of cells expressing VPg or vector controls indicates that VPg reducestranslation efficiency of c-Myc and Mcl1 RNAs but not Actin, the negativecontrol, without altering the global polysome profile (Lower). (B) VPg inhibitedeIF4E-dependent mRNA export for targets RNAs. RNA levels were measured innuclear and cytoplasmic fractions by qRT-PCR. While the increase in GAPDHwas significant, it was so modest that it seems unlikely to be physiologicallyrelevant. P values are shown. (C) Western blot analysis of the effects of VPgoverexpression on eIF4E targets Mcl1 and cMyc. VPg–FLAG levels are given, andactin is provided as a loading control. Note that VPg does not lower endoge-nous eIF4E protein levels. (D) FLAG immunoprecipitations from cells over-expressing VPg–FLAG, eIF4E–FLAG, or controls. Blots were probed as indicated.(E) VPg overexpression suppresses formation of foci in eIF4E–Myc over-expressing cells. ANOVA (P < 0.0009) was conducted. Experiments were carriedout 3 independent times; means ± SDs are shown in A, B, and E (***P < 0.001).

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and the RNA helicase DDX3, which is part of the active trans-lation complex, but not with 4E-BP1, the inhibitor of eIF4E. Thus,VPg engaged active translation initiation complexes. As expected,eIF4E–FLAG bound to eIF4G, DDX3, and 4E-BP1, while noproteins were found in the negative control FLAG immunopre-cipitations from vector controls. This suggests that eIF4G–eIF4E–VPg–RNA complexes readily formed in human cells, which isconsistent with in vitro translation assays, where addition ofVPgΔ37, when not conjugated to RNA, impeded translation ofluciferase RNAs (Fig. 5 D and E). Taken together, VPg is able toimpair host-cell RNA export and translation by competing witheIF4E for host-cell m7G-capped RNAs as well as during infectionwhen, conjugated to gRNA, VPg is positioned to recruit gRNA tothe translation machinery.Given that VPg inhibited the mRNA export and translation

activities of eIF4E, both of which contribute to its oncogenicpotential and require cap binding (6, 7), we explored the possi-bility that VPg inhibited foci formation in eIF4E-overexpressingcells. In this case, stable U2Os cell lines were generated expressingeIF4E–Myc, VPg–FLAG + eIF4E–Myc, or vector controls. Asexpected, eIF4E–Myc increased foci formation ∼3-fold over vec-tor. VPg–FLAG + eIF4E–Myc cells produced ∼3-fold fewer focirelative to eIF4E–Myc, comparable with vector controls (Fig. 6Eand SI Appendix, Fig. S15B). While the initial focus of our VPgstudies was to discover biochemical principles with regard to en-gagement of eIF4E, it is clear that the unusual properties of VPgcould be exploited in the future to inhibit the oncogenic activity ofeIF4E in human cancer cells.

Human Homologs Based on Structural Similarity with VPg.Given thatpotyvirus VPg did not adopt a fold like VPgs from other virusfamilies (consistent with the lack of sequence homology) (SI Ap-pendix, Fig. S1A) or like known eIF4E-binding partners, we usedthe DALI server to ascertain if the VPg fold was similar to anyothers in the protein databank. The top 10 DALI hits includedhuman kinesin 5 family member KIF11/EG5 (top hit) and pro-karyotic or yeast-derived ribosomal large subunit protein 1 com-plexed to 23S ribosomal RNA or a viral IRES (SI Appendix, TableS3). With regard to the top hit, VPg is about half the size of EG5,where the homologous region includes the motor domain (SI Ap-pendix, Fig. S16A). Given the structural homology, we investigatedwhether EG5 bound eIF4E. Using NMR, we found that EG5 di-rectly bound eIF4E, leading to a similar pattern of spectral broad-ening observed for VPgΔ37 (SI Appendix, Fig. S16 C and E). LikeVPgΔ37, excess m7G cap competed for eIF4E in preformed EG5–eIF4E complexes, leading to reemergence of cross-peaks atcap-bound positions (SI Appendix, Fig. S16D). Furthermore,peak broadening of the 15N-labeled eIF4E TrMut was significantlyreduced compared with wild-type eIF4E on addition of EG5 (SIAppendix, Fig. S16E), again supporting that both EG5 and VPginteract with the cap-binding surface on eIF4E. Interestingly, thisinteraction could have important implications for newly identifiedfunctions of EG5 (i.e., it interacted with the RNA-binding ZBPprotein to traffic actin transcripts to modulate cell motility andadditionally, was found to associate with and traffic ribosomes)(61–63). Thus, our studies into potyviruses revealed a previously un-known eIF4E partner protein EG5 and a means for human pro-teins to engage eIF4E.

ConclusionsThese studies leveraged previous plant genetic investigations toreveal the existence of a fundamentally different modality to en-gage and control eIF4E. The traditional view is that eIF4E is onlycontrolled or engaged by interactions with its dorsal surface asobserved for eIF4G. The notion that eIF4E could be inhibited byproteins competing for the cap-binding site was not previouslyconsidered. Consistent with this, VPg impaired host-cell eIF4E-dependent RNA export and translation through binding the

cap-binding site to prevent host-cell RNA association with eIF4E.Our studies with the human EG5 protein suggest that this modalityis not restricted to potyvirus but rather, is conserved across king-doms. Based on these findings, VPg represents a previouslyunknown class of inhibitor of human eIF4E, which could beleveraged in future for therapeutic purposes.Importantly, previous genetic studies demonstrated the re-

quirement for VPg–eIF4E interactions and thus, presumably, thehost-cell host translation machinery for successful viral infection.Given our data, one possibility is that free VPg (not conjugated toRNA) sequesters eIF4E–eIF4G complexes to allow for eitherIRES-mediated or some other form of translation to be engagedfor VPg–gRNA conjugates. Another nonmutually exclusive pos-sibility based on our findings is that VPg–gRNA conjugates arerecruited by eIF4E–eIF4G complexes to the translation machin-ery. In this model, VPg would substitute for the m7G RNA cap onthe gRNA, mediating a form of m7G cap-independent, eIF4E-dependent translation. In all, our studies demonstrated the exis-tence of a fundamentally different form of cap competitor (VPgand EG5) than m7G cap analogs and identified an unanticipatedbinding surface for proteins to engage eIF4E. Furthermore, ourfindings suggest that VPg could substitute as the cap for potyvirusgRNA, which in physical nature, is a significant departure from them7G cap first identified using CPV and VV over 40 y ago (3, 14,15) or recently discovered adenosine nucleotides (5). Investigatingthese possibilities will be interesting areas of future study.

Materials and MethodsProtein Purification. Unlabeled and labeled constructs of PVY VPg encodingresidues 38 to 188 were expressed and purified as described previously (34).VPg mutants and deletion constructs (38 to 188 and 63 to 188) were gen-erated using Quick Change mutagenesis (Bio Basic Inc.). eIF4E was expressedboth in pET-28a for NMR and biophysics and in pGEX-6p1 for GST pulldownexperiments (64). Wheat eIF(iso)4E was expressed in pET28a and purifiedsimilarly as human eIF4E. Human EG5 was overexpressed in a pGEX-6p1construct. All constructs were verified by sequencing. After overnight induc-tion at 20 °C, pelleted cells were resuspended in Tris Buffer (TB) buffer (10 mMTris, pH 7.5, 250 mMKCl, 0.1% Triton X-100, 1 mMMgCl2, 1 mMDTT) and lysedusing sonication (8 rounds of 10 s at high power using the Sonic DismembratorModel 500 from Fisher); the supernatant of the lysate was added to pre-equilibrated glutathione Sepharose 4B beads for affinity purification. Afterextensive washing with TB buffer containing 500 mM KCl, the protein wascleaved overnight with the Prescission Protease in TB buffer, eluted, and fur-ther purified by gel filtration (Superdex 75). High levels of purity (>95%) wereobtained.

NMR Spectroscopy. NMR experiments were performed on Bruker Avance IIIspectrometers running at 600 or 800MHz equippedwith 5-mmQCIP (quadrupleresonance probe with phosphorus) or 5-mm TCI (triple resonance inverse probe)cryoprobes, respectively. For structure determination, typical sample conditionswere 0.4 to 0.5 mM VPg in 50 mM phosphate buffer, 150 mM NaCl, 1 mM DTT,and 0.02%NaN3 with either 7 or 100%D2O (pH 7.5), and all spectra were ran at20 °C. 3D NOESY (NOE spectroscopy) spectra were acquired using nonuniformsampling (NUS) with 30% Poisson Gap sampling (65). For NMR titrations, con-centrations for labeled and unlabeled proteins were 50 and 150 μM, respec-tively, and when the m7GDP cap analog was added, a 20-molar excess to eIF4Ewas used. Spectra were processed with NMRPipe (66) and SMILE (67) and ana-lyzed with SPARKY (68).

HADDOCK-Derived Complex Structure of VpG–eIF4E. The HADDOCK2.2 webserver(41) was used to generate restraint-driven docking for interaction be-tween eIF4E and VPg using the standard protocols with the VPg structurereported here and eIF4E (PDB ID code 2GPQ). Default HADDOCK settingswere used for the docking, generating a final 200 structures. The finalmodels were clustered based on the fraction of common contacts usinga 0.60 cutoff.

XL-MS. Cross-linked sample was prepared by mixing equimolar amounts ofeIF4E and His-tagged VPgΔ37 (0.15 μmol) with 1 mM DSS (4,4-Dimethyl-4-silapentane-1-sulfonic acid). After a 15-min incubation at room temperature,the reaction was stopped using NH4HCO3 and further purified by gelfiltration chromatography (Superdex-75 column). Proteins were digested

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overnight at 37 °C using a 1:20 trypsin-to-protein ratio, and trypsin wasinactivated with 0.5% trifluoroacetic acid. A series of samples (4 μg per well)was loaded into a 2-mg sorbent 96-well plate Oasis MCX μElution plate(Waters), washed, and eluted according to SI Appendix, Table S4 to enrichfor multiply charged peptides. All samples were loaded at 600 nL min−1 on a17-cm × 75-μm inner diameter PicoFrit fused silica capillary column (NewObjective) and packed in house with Jupiter 5 μm C18 300 Å (Phenomenex).The column was mounted in an Easy-nLC II system (Proxeon Biosystems) andcoupled to an Orbitrap Fusion mass spectrometer (ThermoFisher Scientific)equipped with a Nanospray Flex Ion source (Proxeon Biosystems). MS rawfiles were analyzed using SIM-XL (69, 70). MS2 spectra were visually inspec-ted, and theoretical surface-accessible solvent distances for each compiledcross-link were measured using Xwalk (42).

CD. CD spectra were collected on a Jasco-810 spectropolarimeter using a0.1-cm quartz cuvette (Hellma) on pure proteins at 20 μM at room temperature.Relative ellipticity was converted to mean residue molar ellipticity accordingto Fasman (71).

Fluorescence Spectroscopy. Cysteine residues were reduced by buffer-exchanging VPg into phosphate-buffered saline (PBS) buffer at pH 7.4 (con-taining 50 μM tris(2-carboxyethyl)phosphine [TCEP] and purged with N2) usingan NAP-5 column (Fisher). VPg was then labeled by adding 10-molar excess ofthe organic fluorophore IAEDANS dropwise while stirring. The reaction wasleft at 25 °C for 2 h in the dark and stopped by adding 10 mM DTT. Bufferexchange was performed over an NAP-5 column using a spin column (Amicon10-kDa MWCO [molecular cutoff]; Fisher) to remove unbound dye. Fluores-cence measurements were carried out as follows. Briefly, 0.5 μM labeled VPgwas incubated with increasing concentrations of eIF4E (0 to 2.5 μM) in PBSbuffer containing 1 mMDTT. Fluorescence measurements were performed in a0.3 × 0.3-cm2 fluorescence cuvette (Hellma) using a Cary Eclipse FluorescenceSpectrophotometer (Agilent Tech.). The degree of IAEDANS labeling was de-termined by measuring the absorbance at 336 nm using the extinction coef-ficients 5,700 M−1 cm−1. The IAEDANS-labeled VPg displayed a maximumfluorescence intensity at 484 nm. The fluorescence intensity increased∼45% onassociation of eIF4E. Binding isotherms were fit according to the Hill model.Measurements were carried out at least 3 independent times.

ITC. ITC was performed with aMicrocal ITC200 calorimeter operating at 20 °C.The data were analyzed with MicroCal Origin software. The protein con-centration was 10 μM in the cell, while the m7GDP cap analog was 100 μM inthe syringe. Experiments consisted of 16 injections of 2.5 μL at a rate of0.5 μL s−1 at 180-s intervals. The first injection peak was discarded from theisotherm. The baseline was automatically generated by the MicroCal Originpackage and corrected manually. The binding isotherm was fitted using theOne Set of Sites model in the MicroCal Origin package.

RNA Conjugation. VPgΔ37 was mutated in positions 64 (Y64C) and 150(C150A) to present only 1 cysteine at the position where VPg is known to becovalently attached to gRNA during infection. The sulfhydryl group ofC64 was conjugated with the maleimide functional group of a succinimidyl-4-(N-maleimidomethyl)cyclohexane-1-carboxylate oligonucleotide (GeneLink)as shown in SI Appendix, Fig. S14A. The reaction was performed at roomtemperature with a 20-fold excess of the modified oligonucleotide. Moredetails are given in SI Appendix, Supplementary Methods.

To generate RNA templates suitable for translation, full-length luciferaseRNA (∼1,800 nucleotides) was in vitro transcribed (using the T7 Megascript Kit;Ambion) and 5′ end primed with GMPS (Axxora; Biolog Life Science Institute)(51, 52). GMPS-primed RNAs were than coupled to 2,2′-pyridine disulfide (SIGMA)to produce pyridyl-disulfide linkage on the 5′ end of RNAs. Disulfide ex-change chemistry between the thiol group from VpG and pyridyl-disulfideon the 5′ end on luciferase RNA was used to covalently attach RNA to pu-rified His–VPg C150A/Y64C. A schematic diagram of these reactions is shown

in SI Appendix, Fig. S14E (53), and a detailed protocol is given in SI Appendix,Supplementary Methods. Validation of the conjugates was carried out usingformaldehyde-agarose gel electrophoresis followed by immunoblotting withanti-His antibodies.

In Vitro Translation Assays. Wheat germ lysate system was used to assess therelative translation of VPg–luciferase RNA conjugates, m7G-capped lucif-erase RNA, or uncapped luciferase RNA. More details are in SI Appendix,Supplementary Methods.

Cell Culture and Transfection. U2Os cells (ATCC) were maintained in 5% CO2 at37 °C in DMEM (Dulbecco’s Modified Eagle Medium, Gibco BRL) supple-mented with 10% FBS (Fetal Bovine Serum) and 1% penicillin-streptomycin(Invitrogen). Transfections for stable cell lines were performed using TransIT-LT1 Transfection Reagent (Mirus) as specified by the manufacturer andselected in puromycin-containing medium (10 μg/mL) for pMSCVeIF4E–Mycand/or G418 (1 mg/mL) for 2Flag–VPg overexpressing cell lines. The identityof U2Os cell line has been authenticated using STR profiling (MontrealEpiTerapia Inc.).

Cellular Fractionation, mRNA Export Assay, and Polysomal Profiling. Nucleo-cytoplasmic fractionations for mRNA export assays and polysomal profilingwere done as previously described (56, 58, 72) and detailed in SI Appendix,Supplemental Methods. TRIzol reagent (ThermoFisher Scientific) was addedto each fraction, and RNAs were extracted using a DirectZol RNA Miniprepkit (Zymo Research), including deoxyribonuclease (DNase) treatment. RNAswere reversed transcribed using M-MLV Reverse Transcriptase (ThermoFisherScientific). SensiFastSybr Lo-Rox Mix (Bioline) was used for qRT-PCR analysesby relative standard curve method (Applied Biosystems User Bulletin #2).

Anchorage-Dependent Foci Assays. Experiments were carried out as describedpreviously. A total of 500 cellswere seededper10-cmplate or 100 cells perwell in6-well plates for 14 d, and then, they were stained with Giemsa (Sigma-Aldrich).

Immunoprecipitations. U2Os cells were fixed with 1% paraformaldehyde(PFA) for 10 min at room temperature (RT) and quenched with 0.15 M glycinefor 5 min at RT. Cells were than washed 3 times with cold 1× PBS, lysed bysonication (4 rounds of 5 s at lowest power) in NucleoTrap (NT2) buffer (56),and centrifuged for 10 min at 12,000 × g. Lysates were precleared withSephadex G beads (GE Healthcare) for 30 min at 4 °C, and 0.75 to 1 mg ofprecleared lysates were used for immunoprecipitation with 7 to 10 μg anti-Flag antibody (Sigma) overnight at 4 °C. After incubation, complexes werewashed 6 times with NT2 buffer, eluted by boiling in Tris(hydroxymethyl)aminomethane (Tris) EDTA (ethylenediaminetetraacetic acid) containing 1%sodium dodecyl sulfate (SDS) and 12% β-mercaptoethanol, and analyzed bywestern blot.

Data Availability. The processed spectra and atomic coordinates for VPg weredeposited into thePDBand theBiologicalMagnetic ResonanceDataBank (PDB IDcode 6NFW and Biological Magnetic Resonance Data Bank accession no. 27506).

ACKNOWLEDGMENTS. We are grateful for helpful discussions and VPg andwheat eIF(iso)4E constructs from Dr. Jadwiga Chroboczek (Université Greno-ble Alpes-Centre National de la Recherche Scientifique [UGA-CNRS]) andDr. Karen Browning (University of Texas), respectively. We thank Jose RafaelDimayacyac (Institute of Research in Immunology and Cancer) and Dr. JackKornblatt (Concordia University) for technical assistance. We also thankDr. Tara Sprules at Quebec/Eastern Canada High-Field NMR facility for useof the 800-MHz NMR. K.L.B.B. acknowledges financial support from NIHGrants R01 CA80728 and R01 CA98571, Canadian Institutes of Health Re-search (CIHR) Grant PJT159785, the Canada Research Chair in Molecular Bi-ology of the Cell Nucleus, and the Canadian Foundation for Innovation forupgrades to the 600-MHz instrument.

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