Allosteric Regulation of RNA Affinity by Motif V-VI Coupling in West Nile Virus NS3 Helicase

Allosteric Regulation of RNA Affinity by Motif V-VI Coupling in West Nile Virus NS3 Helicase | bioRxiv /* */ /* */ <!-- <!-- /*! * yepnope1.5.4 * (c) WTFPL, GPLv2 */ (function(a,b,c){function d(a){return"[object Function]"==o.call(a)}function e(a){return"string"==typeof a}function f(){}function g(a){return!a||"loaded"==a||"complete"==a||"uninitialized"==a}function h(){var a=p.shift();q=1,a?a.t?m(function(){("c"==a.t?B.injectCss:B.injectJs)(a.s,0,a.a,a.x,a.e,1)},0):(a(),h()):q=0}function i(a,c,d,e,f,i,j){function k(b){if(!o&&g(l.readyState)&&(u.r=o=1,!q&&h(),l.onload=l.onreadystatechange=null,b)){"img"!=a&&m(function(){t.removeChild(l)},50);for(var d in y[c])y[c].hasOwnProperty(d)&&y[c][d].onload()}}var j=j||B.errorTimeout,l=b.createElement(a),o=0,r=0,u={t:d,s:c,e:f,a:i,x:j};1===y[c]&&(r=1,y[c]=[]),"object"==a?l.data=c:(l.src=c,l.type=a),l.width=l.height="0",l.onerror=l.onload=l.onreadystatechange=function(){k.call(this,r)},p.splice(e,0,u),"img"!=a&&(r||2===y[c]?(t.insertBefore(l,s?null:n),m(k,j)):y[c].push(l))}function j(a,b,c,d,f){return q=0,b=b||"j",e(a)?i("c"==b?v:u,a,b,this.i++,c,d,f):(p.splice(this.i++,0,a),1==p.length&&h()),this}function k(){var a=B;return a.loader={load:j,i:0},a}var l=b.documentElement,m=a.setTimeout,n=b.getElementsByTagName("script")[0],o={}.toString,p=[],q=0,r="MozAppearance"in l.style,s=r&&!!b.createRange().compareNode,t=s?l:n.parentNode,l=a.opera&&"[object Opera]"==o.call(a.opera),l=!!b.attachEvent&&!l,u=r?"object":l?"script":"img",v=l?"script":u,w=Array.isArray||function(a){return"[object Array]"==o.call(a)},x=[],y={},z={timeout:function(a,b){return b.length&&(a.timeout=b[0]),a}},A,B;B=function(a){function b(a){var a=a.split("!"),b=x.length,c=a.pop(),d=a.length,c={url:c,origUrl:c,prefixes:a},e,f,g;for(f=0;f<d;f++)g=a[f].split("="),(e=z[g.shift()])&&(c=e(c,g));for(f=0;f<b;f++)c=x[f](c);return c}function g(a,e,f,g,h){var i=b(a),j=i.autoCallback;i.url.split(".").pop().split("?").shift(),i.bypass||(e&&(e=d(e)?e:e[a]||e[g]||e[a.split("/").pop().split("?")[0]]),i.instead?i.instead(a,e,f,g,h):(y[i.url]?i.noexec=!0:y[i.url]=1,f.load(i.url,i.forceCSS||!i.forceJS&&"css"==i.url.split(".").pop().split("?").shift()?"c":c,i.noexec,i.attrs,i.timeout),(d(e)||d(j))&&f.load(function(){k(),e&&e(i.origUrl,h,g),j&&j(i.origUrl,h,g),y[i.url]=2})))}function h(a,b){function c(a,c){if(a){if(e(a))c||(j=function(){var a=[].slice.call(arguments);k.apply(this,a),l()}),g(a,j,b,0,h);else if(Object(a)===a)for(n in m=function(){var b=0,c;for(c in a)a.hasOwnProperty(c)&&b++;return b}(),a)a.hasOwnProperty(n)&&(!c&&!--m&&(d(j)?j=function(){var a=[].slice.call(arguments);k.apply(this,a),l()}:j[n]=function(a){return function(){var b=[].slice.call(arguments);a&&a.apply(this,b),l()}}(k[n])),g(a[n],j,b,n,h))}else!c&&l()}var h=!!a.test,i=a.load||a.both,j=a.callback||f,k=j,l=a.complete||f,m,n;c(h?a.yep:a.nope,!!i),i&&c(i)}var i,j,l=this.yepnope.loader;if(e(a))g(a,0,l,0);else if(w(a))for(i=0;i (function(w,d,s,l,i){w[l]=w[l]||[];w[l].push({'gtm.start':new Date().getTime(),event:'gtm.js'});var f=d.getElementsByTagName(s)[0];var j=d.createElement(s);var dl=l!='dataLayer'?'&l='+l:'';j.src='//www.googletagmanager.com/gtm.js?id='+i+dl;j.type='text/javascript';j.async=true;f.parentNode.insertBefore(j,f);})(window,document,'script','dataLayer','GTM-M677548'); Skip to main content Home About Submit ALERTS / RSS Search for this keyword Advanced Search New Results Allosteric Regulation of RNA Affinity by Motif V-VI Coupling in West Nile Virus NS3 Helicase View ORCID Profile Priti Roy , View ORCID Profile Martin McCullagh doi: https://doi.org/10.1101/2025.07.04.663198 Priti Roy Department of Chemistry, Oklahoma State University , Stillwater, OK, 74074, USA Find this author on Google Scholar Find this author on PubMed Search for this author on this site ORCID record for Priti Roy Martin McCullagh Department of Chemistry, Oklahoma State University , Stillwater, OK, 74074, USA Find this author on Google Scholar Find this author on PubMed Search for this author on this site ORCID record for Martin McCullagh For correspondence: martin.mccullagh{at}okstate.edu Abstract Full Text Info/History Metrics Supplementary material Preview PDF Abstract The rise of flaviviral diseases, including West Nile virus (WNV), presents a growing threat to global public health and underscores the urgent need for new therapeutic strategies. The non-structural protein 3 helicase (NS3h) of the Orthoflavivirus genus, including WNV, is essential for viral replication and a promising antiviral target. Previously [Roy et al. , Nucleic Acids Research , 52, 13, 2024, 7447–7464], we showed that the motif VI loop (VIL) in WNV NS3h functions as a nucleotide valve, regulating ADP affinity during hydrolysis. In this study, we uncover an ATP-dependent coupling between nucleotide affinity at motif VIL and RNA affinity at motifs IVa and V, suggesting a coordinated mechanism of ssRNA translocation. Using microsecond-scale all-atom molecular dynamics simulations of hydrolysis-cycle intermediates, we find that key VIL residues (R461, R464) correlate strongly with RNA phosphate affinity of motif V. Structural analyses reveal an ATP-sensitive interaction between E413 (motif V) and R461 (motif VIL) that modulates the conformation of the motif V 3 10 -helix, thereby influencing RNA binding. This dynamic interaction is lost in catalytically deficient VIL mutants, which have been experimentally shown to impair hydrolysis and attenuate viral replication. These findings provide mechanistic insights into NS3h function and identify new opportunities for structure-based antiviral design. Introduction West Nile Virus (WNV) remains a growing global public health concern. As an arthropodborne (arbo) virus, WNV is maintained in a transmission cycle between vertebrate reservoir hosts and mosquito vectors. Human infection can result in a wide spectrum of outcomes, ranging from mild fever to severe neurological complications and, in some cases, kidney disease. 1 – 3 WNV has caused multiple outbreaks 4 , 5 and continues to expand into new geographic regions, with an increasing number of countries reporting cases. 6 Despite significant advances in vaccine development over the past two decades, 7 effective treatments for WNV infection remain elusive, and no antiviral drugs have been approved for clinical use. 7 A promising approach to antiviral drug development focuses on targeting essential viral proteins 8 , 9 which requires a comprehensive understanding of the structural mechanisms of viral protein functions. The helicase domain of the non-structural protein 3 (NS3h) of WNV is one such target. WNV is a positive-sense single-stranded RNA (ssRNA) virus belonging to the the Orthoflavivirus genus. Following infection, the viral RNA is translated into viral proteins by the host cellular machinery. 10 Viral replication proceeds through synthesis of a complementary negative-sense RNA strand, forming double-stranded RNA (dsRNA) intermediates. The separation of these intermediates, an essential step in replication, is driven by the helicase activity of NS3h. NS3h is highly conserved across Orthoflavivirus members such as DENV, ZIKV, and JEV, and mutations in this domain have been linked to increased virulence and adaptive evolution in WNV. 11 NS3h unwinds dsRNA by translocating along the negative-sense strand in a 3 ′ →5 ′ direction, powered by ATP hydrolysis. NS3h has therefore emerged as a high-value candidate for structure-based drug development. 12 – 15 A detailed understanding of its structural mechanism is critical for the rational design of WNV-specific and broad-spectrum antivirals. Functionally, NS3h coordinates RNA translocation and ATP hydrolysis through a structurally coupled mechanism. A DEAH-box helicase of superfamily 2 (SF2), NS3h is a monomeric protein comprising three domains (see Figure 1 ). Domains I and II, which adopt a Rossmann fold, form the active site for nucleoside triphosphatase (NTPase or ATPase) activity, commonly referred to as the ATP-pocket. Domain III, positioned opposite domains I and II, contributes to the formation of an RNA-binding cleft, or RNA-cleft. Although the ATP- and RNA-binding sites are spatially separated by ∼ 30 °A (center-of-mass distance), they exhibit strong functional interdependence: RNA binding enhances ATPase activity in the ATP-pocket, 16 and ATP hydrolysis in turn drives conformational changes that mediate RNA translocation within the cleft. 17 , 18 This reciprocal regulation suggests the existence of a structural mechanism that enables allosteric communication between the two sites. Download figure Open in new tab Figure 1: Schematic representation of the ATPase cycle of NS3 helicase (NS3h). Here we depict the intermediates states that NS3h transitions through during ATP hydrolysis cycle. The cycle starts when ssRNA enters RNA-cleft of the Apo NS3h and generates ssRNA state. After ssRNA binding, ATP enters the ATP-pocket and forms ssRNA+ATP state. Following hydrolysis, protein moves to ssRNA+ADP+P i state. Next, P i leaves the ATP-pocket and gives rise to ssRNA+ADP state. Finally, ADP releases and protein re-enter the ssRNA state to resume next cycle. Free energy released from this cycle drives translocation in 3 prime to 5 prime direction along the ssRNA to unwind dsRNA. Mounting evidence suggests that the motif VI loop acts as a key regulatory element linking ATP hydrolysis to RNA translocation in flaviviral helicases. Our previous work combining molecular simulations and biochemical assays demonstrated that the motif VI loop in WNV NS3h modulates nucleotide binding enthalpy in a hydrolysis state-dependent manner, functioning as a nucleotide valve. 19 We also observed ATP-dependent changes in RNA affinity, a phenomenon reported in other members of the NS3 helicase family. 16 , 17 , 20 , 21 Together, these observations support the hypothesis that ATP-dependent nucleotide affinity at motif VI loop may be coupled to changes in RNA affinity within the RNA-cleft. This putative coupling likely involves motif V , which lies in close spatial proximity to motif VI loop and is structurally poised to mediate communication between the ATP-pocket and RNA-binding site. Motif V comprises a β -strand, a 3 10 -helix that interacts with ssRNA, and a coil facing the ATP-pocket. Prior studies have shown that ATP binding induces shifts in motif V toward the nucleotide-binding site in DENV NS3h, 22 and that hydrogen bonding within its 3 10 -helix contributes to RNA unwinding and replication. 23 Structural analyses of ZIKV NS3h revealed conformational variability in the coil region of motif V , 18 and MD simulations suggest it samples distinct conformations across the hydrolysis cycle. Although our prior work found no direct enthalpic signature of hydrolysis state dependence in motif V , we hypothesize that it serves as a structural conduit, mediating ATP-dependent communication from motif VI to the RNA-cleft. The present study tests this hypothesis by analyzing ATP-dependent structural correlations between motifs V and VI , aiming to define a mechanistic link between ATP hydrolysis and RNA translocation, and to inform the development of NS3h-targeted antivirals. Methods Starting Structures and System Preparation In this study, the D471E and D471L mutants, bound to ATP, ADP+P i , and ADP, were generated by modifying previously modeled wild-type WNV NS3h structures in complex with ssRNA. 19 Specifically, the side-chain terminal atoms of D471 were replaced with those of glutamic acid (D471E) and leucine (D471L), respectively, while maintaining the over-all protein architecture. The resulting mutant structures were solvated with TIP3P water molecules. 24 To neutralize the system and maintain an ionic strength of 0.15 M, Na +1 and Cl −1 ions were added. A cubic box with a buffer distance of 20 °Afrom the protein surface was used. The dimensions of the resulting box were 107 °Aalong each axis. The simulations were conducted at neutral pH. For details on histidine protonation states, refer to the wild-type system preparation described in Roy et al., 2024. 19 Simulation Protocol Mutant hydrolysis states were simulated similar to wild-type simulations 19 with GPU-enabled AMBER18 software. 25 In brief, each mutant state was minimized in the conjugate-gradient method, followed by heating at 310 K. After equilibration, the production run was performed for 5 µ s in the NPT ensemble. The integration time step was 2 fs and the coordinates were saved every 10 ps. Equation of motion was solved using ff14SB 26 and ff99bsc0 χ OL3 27 , 28 as potential energy values for protein and ssRNA, respectively. The ATP, 29 ADP, 29 and Mg 2+ 30 parameters were used, while the P i (H 2 PO 4 − ) parameter was adopted from a previous study by our group. 20 In aggregates, we generated approximately 40 µ s of simulated frame and analyzed 60 µ s of simulated frame, including previously reported wild-type simulation data. 19 Analyses Binding Enthalpy, E inter Non-bonded linear interaction energy (E inter ) between the protein and substrates i.e. the phosphate moieties of the ssRNA or ADP is computed from the NPT ensemble of each system which we denote as binding enthalpy. We computed electrostatic and van der Waals interactions as non-bonded interactions, using Cpptraj lie module with a long-range cutoff of 12 °A. Pearson Correlation The Pearson correlation measures the linear dependency of two variables. We computed the correlation coefficient which measures the extent of negative or positive correlation ranging from −1 to 1: The ⟨X⟩ and ⟨Y⟩ denote the ensemble averages. Pearson’s correlation was evaluated for substrate binding affinities and Cartesian positions. Hydrogen Bond (H-bond) A hydrogen bond is considered when the two participating heavy atoms are electronegative and a hydrogen atom is attached to one of the heavy atom. The hydrogen bound electronegative atom acts as a donor, while the other atom acts as an acceptor. We used (i) a cutoff distance of 3 °Abetween the donor and acceptor atom, (ii) a cutoff distance of 1.1 °Abetweenthe donor and donor hydrogen atoms and (iii) an angle value higher than the 120 ◦ formed between the donor, hydrogen and acceptor atoms. Model Corroboration Model hydrolysis states of the mutants are comparable to those of the corresponding wild-type states. Comparison of RNA contact sampling between mutants and wild-type shows that a large number of wild-type conserved contacts sample 70% - 100% in mutants (Supporting Information Table S1-S4). Similarly, we observe sampling of conserved ATP, ADP+P i and ADP contacts, with a sampling probability higher than 70% for most of the wild-type contacts (Table S5-S7). However, a few conserved contacts are lost or samples below 20% in mutants. The octahedral coordination of Mg 2+ is also maintained in the mutant hydrolysis states (Table S8). We computed the backbone dihedral angles of the wild-type and mutants and presented as a 2D histogram in the Supp. Info Figure S1. Mutations are incorporated by only changing the sidechain terminal moiety, and thus, these alterations would not have any influence on the secondary and tertiary structure. Similarly, the distributions of mutant states are comparable to those of wild-type states, largely sampling the α -helix region. Results and Discussion The translocation activity of NS3h proceeds through a non-equilibrium, multistate hydrolysis cycle that couples RNA binding and ATP turnover. Crystal structures and biochemical studies have identified a sequence of conformational states corresponding to distinct steps in this ATPase cycle which we show in Figure 1 . 16 , 17 , 31 , 32 The cycle begins when single-stranded RNA (ssRNA) binds within the RNA-cleft, while the ATP-binding site remains unoccupied; we refer to this state as ‘ssRNA’. RNA binding enhances nucleotide affinity, 33 promoting ATP binding and formation of a pre-hydrolysis complex, ‘ssRNA+ATP’. ATP hydrolysis produces the post-hydrolysis-I state, ‘ssRNA+ADP+P i ’, which then releases in-organic phosphate to yield the post-hydrolysis-II state, ‘ssRNA+ADP’. Finally, ADP release returns the protein to the initial ‘ssRNA’ state. These conformational transitions correspond to elementary steps of helicase motion, likely involving single-nucleotide translocation events. 32 , 34 We adopt these state definitions for our analysis, which builds on prior work demonstrating ATP-dependent changes in nucleotide and RNA binding affinity. 19 In this context, we hypothesize that the motif VIL acts as a dynamic regulator that couples nucleotide affinity to RNA translocation during the hydrolysis cycle. This hypothesis is motivated by our previous study, which combined molecular dynamics simulations and biochemical assays to show that motif VIL plays a key role in controlling nucleotide entry, stabilization, and release. 19 Although motif VIL is spatially distant from the ATP-binding site, mutation of a key residue impaired hydrolysis and viral replication, consistent with long-range allosteric regulation. We previously proposed that motif VIL acts as a nucleotide valve, modulating access to the ATP site across different hydrolysis states. 19 Here, we investigate whether this valve-like behavior is structurally and functionally coupled to RNA affinity, thereby linking ATP turnover to translocation. Our analysis is organized into three parts: (1) the correlation between nucleotide (ADP) and RNA affinity; (2) residue-level interactions that bridge the ATP- and RNA-binding sites; and (3) structural coupling between motif VIL and RNA-binding motifs that varies across hydrolysis states. Correlation between nucleotide affinity at motif VI and RNA affinity WNV NS3h contains eight conserved sequence motifs ( I , Ia , II , III , IV , IVa , V , and VI loop [ VIL ]) that coordinate ATP hydrolysis and RNA translocation through substrate binding in two functionally distinct regions: the ATP-pocket and the RNA-cleft ( Figure 2 A). Motifs I , II , and III form the nucleotide-binding pocket and catalyze ATP hydrolysis, 35 – 37 while motifs IV and IVa reside in the RNA-cleft and participate in RNA unwinding and replication. 37 , 38 Motif V is positioned between these regions and has been implicated in allosteric coupling. 18 , 37 , 38 Motif Ia , which spans both regions, and motif VIL , the so-called arginine finger, are of particular interest: the former shows ATP-dependent motion in related helicases, 39 , 40 and the latter stabilizes nucleotide binding and is essential for catalysis. 16 , 19 Based on structure and prior functional studies, we classify motifs I , II , III , V , and VIL as ATP-pocket motifs and motifs Ia , IV , IVa , and V as RNA-cleft motifs. To assess whether ATP hydrolysis dynamically coordinates substrate binding in these regions, we investigated ATP-dependent coupling between nucleotide and RNA affinity across the hydrolysis cycle. Download figure Open in new tab Figure 2: ATP-dependent correlation between nucleotide and RNA affinity in WNV NS3h. Binding affinity is defined as the interaction energy between a substrate and a motif residue within the protein. For nucleotides, we consider ADP (excluding the γ -phosphate group of ATP); for RNA, we quantify affinity based on interactions with phosphate moieties of single-stranded RNA (RNA(P)). (A) Structural model of WNV NS3 helicase (NS3h), highlighting locations of conserved motifs. NS3h contains eight conserved sequence motifs (I, Ia, II, III, IV, IVa, V, and VIL), all residing within domains I and II. (B–D) Residue-wise cross-correlation plots comparing nucleotide (ADP) and RNA(P) binding affinities for three hydrolysis states: (B) ssRNA+ATP (pre-hydrolysis), (C) ssRNA+ADP+P i (post-hydrolysis I), and (D) ssRNA+ADP (post-hydrolysis II). Regions with marked changes in correlation across states are outlined with rectangular boxes. Nucleotide and RNA binding affinities in NS3h become increasingly coupled as the hydrolysis cycle progresses, indicating that ATP hydrolysis drives coordinated substrate engagement across distant binding sites. Prior work has shown hydrolysis state-dependent nucleotide binding affinity in WNV NS3h, 19 as well as ATP-dependent changes in RNA affinity in both DENV4 and WNV helicases. 19 , 20 These findings suggest that affinity for nucleotide and RNA may be coupled in a hydrolysis-state-dependent manner. To investigate this coupling, we computed residue-level cross-correlations between nucleotide and RNA binding affinities for all conserved motifs in three hydrolysis states: ssRNA+ATP, ssRNA+ADP+P i , and ssRNA+ADP ( Figure 2B–D ). RNA affinity was defined as the interaction energy between motif residues and the phosphate moieties of the RNA backbone (RNA(P)). To ensure consistency across states, nucleotide affinity was calculated using interaction energies with ADP, which is present in all three states. This analysis revealed that nucleotide and RNA affinities become increasingly correlated as the hydrolysis cycle progresses. In the ssRNA+ATP state, correlation coefficients ranged from –0.095 to 0.073 ( Figure 2B ), indicating weak coupling. In contrast, the correlation range broadened significantly in the ssRNA+ADP+P i (–0.396 to 0.406; Figure 2C ) and ssRNA+ADP (–0.417 to 0.433; Figure 2D ) states. This ATP-dependent increase in coupling strength supports a model in which nucleotide binding at the ATP-pocket influences RNA affinity within the cleft, and vice versa. Among ATP-pocket motifs, I and VIL show the strongest coupling to RNA affinity, but with distinct state-dependent profiles—motif VIL , in particular, exhibits behavior consistent with an allosteric regulatory role. Motif I , a core component of the ATP-binding pocket, shows increasing correlation with RNA(P) affinity as the cycle progresses. In the ss-RNA+ATP state, correlation coefficients between motif I and RNA(P) range from –0.095 to 0.073, indicating weak coupling ( Figure 2B ). This correlation becomes more pronounced in the ssRNA+ADP+P i state (–0.254 to 0.252; Figure 2C ), and shifts further toward positive correlation in the ssRNA+ADP state (–0.038 to 0.433; Figure 2D ). In contrast, motifs II , III , and V show no significant correlation with RNA affinity in any hydrolysis state, suggesting they primarily support hydrolysis rather than substrate coupling. Motif VIL displays a distinct pattern: no correlation in the ssRNA+ATP state ( Figure 2B ), but strong bidirectional correlations emerge in the ssRNA+ADP+P i state (–0.396 to 0.406; Figure 2C ), which then shift toward predominantly negative correlations in the ssRNA+ADP state (–0.417 to 0.036; ( Figure 2D )). This ATP-dependent behavior of motif VIL supports its proposed role as a nucleotide valve that dynamically couples nucleotide affinity to RNA binding through the course of the hydrolysis cycle. ATP-dependent coupling between nucleotide and RNA affinity is also evident in the RNA-cleft motifs, particularly motifs IV , IVa , and V . These motifs show increasing correlation with the nucleotide affinity of motif I and motif VIL as the hydrolysis cycle progresses. In the ssRNA+ATP state ( Figure 2B ), correlations between nucleotide affinity at motif I and RNA(P) affinity are generally weak across RNA-cleft motifs (e.g., –0.093 to 0.0 for Ia , –0.080 to 0.033 for IV ), and no notable correlation is observed for motif VIL . In the ssRNA+ADP+P i state ( Figure 2C ), stronger correlations emerge. RNA affinity of motifs IV and IVa show the highest correlation with motif I nucleotide affinity (ranging from –0.254 to 0.252), while motif V also shows moderate coupling. Strikingly, motif VIL shows emergent bidirectional correlations with RNA affinity in motifs IV , IVa , and V , including a peak correlation of 0.406 with motif IVa . By the ssRNA+ADP state ( Figure 2D ), the coupling landscape diverges, revealing distinct roles for motifs I and VIL . Correlation between motif I and motif V reaches a maximum (up to 0.433), whereas correlations with motif VIL become predominantly negative, particularly for motifs IVa and V . These trends highlight a reorganization of coupling between nucleotide- and RNA-binding regions during the hydrolysis cycle and underscore the potential role of motif V as a key mediator in this allosteric communication. Motif VIL exhibits more pronounced and state-specific coupling to RNA affinity than motif I , suggesting a unique mechanism for linking ATP hydrolysis to RNA engagement. From the ssRNA+ATP to the ssRNA+ADP+P i state, the correlation between motif VIL ’s ADP affinity and RNA(P) affinity increases sharply—particularly with RNA-cleft motifs IV , IVa , and V —exceeding the magnitude of change observed for motif I . As the system transitions to the ssRNA+ADP state, motif I ’s correlation with RNA affinity diminishes for most RNA-cleft motifs, persisting only for motif V . In contrast, motif VIL maintains a strong correlation with motif V and shows moderate coupling with motif IVa , while correlations with motif IV is lost. These state-specific patterns are consistent with our previous findings that highlighted motif VIL ’s central role in modulating nucleotide affinity across the hydrolysis cycle. 19 Taken together, these suggest that motif VIL may function as a conduit for transferring free energy from ATP hydrolysis to RNA translocation via selective coupling with RNA-binding elements. Disruption of the ATP-dependent correlation between motif VIL and RNA affinity impairs helicase function in WNV NS3h. Molecular dynamics simulations of hydrolysis-state models for two motif VIL mutants—D471E (Figure S2) and D471L (Figure S3)—reveal a loss of the strong ADP–RNA affinity correlation observed in the wild-type enzyme. These mutants, previously shown to exhibit severely reduced nucleotide binding affinity, 19 fail to maintain the coordinated interaction between ADP binding and RNA affinity. Notably, the correlation between motif I ADP affinity and RNA(P) affinity varied across the mutants, in some cases strengthening and in others weakening relative to wild-type. However, this variability did not compensate for the loss of motif VIL function, suggesting that the ATP-dependent coupling mediated specifically by motif VIL is critical for the enzymatic activity. Together, these findings establish motif VIL as a critical structural element for coupling ATP hydrolysis to RNA translocation in WNV NS3h. Correlation between nucleotide affinity at motif VI and RNA affinity at motif V ATP-dependent changes in phosphate-specific RNA affinity at motifs IVa and V identify key contacts that may coordinate with nucleotide binding at motif VIL during translocation. In particular, motifs IVa and V exhibit notable changes in affinity for the first three RNA phosphates (P 1 –P 3 ) across the hydrolysis cycle. To identify the most functionally relevant RNA-binding sites, we calculated changes in interaction energy (ΔE inter ) between each RNA-cleft motif and individual phosphate groups across successive states. Figure S4 shows phosphate-specific ΔE inter values for motifs Ia , IV , IVa , and V , revealing significant shifts in affinity at P 1 and P 3 for motif IVa , and at P 2 for motif V . Figure 3 presents residue-level phosphate affinity changes over the course of the hydrolysis cycle: each row (from top to bottom) shows transitions corresponding to ATP binding (A–C), hydrolysis (D–F), P i release (G–I), and ADP release (J–L). These data identify T409 and D410 in motif V as key contributors to changes in P 2 affinity, while R388 and K389 in motif IVa drive affinity shifts at P 1 and P 3 . Representative structures in Figure 4 illustrate the evolving positions of these residues and phosphates across hydrolysis states, providing a structural basis for ATP-dependent coupling between RNA- and nucleotide-binding sites. Download figure Open in new tab Figure 3: ATP-dependent changes in RNA phosphate (RNA(P)) affinity for motifs IVa and V. Each panel shows the residue-wise change in RNA phosphate affinity (ΔE inter , in kcal·mol −1 ) for key contacts between protein motifs and ssRNA phosphates during transitions between hydrolysis states. Left, middle, and right columns correspond to changes in affinity between motif V and P 2 , motif IVa and P 1 , and motif IVa and P 3 , respectively. Values of ΔE inter reflect the difference in ensemble-averaged interaction energies computed over full NPT trajectories for each state. Error bars represent the propagated standard deviation of the difference between two ensemble means, where each mean is computed from 50,000-frame chunks. (A–C) Transition from ssRNA to ssRNA+ATP. (D–F) Transition from ssRNA+ATP to ssRNA+ADP+P i . (G–I) Transition from ssRNA+ADP+P i to ssRNA+ADP. (J–L) Transition from ssRNA+ADP (previous cycle) to ssRNA (next cycle). Thicker lines indicate residues exhibiting significant affinity changes during a given transition. See Analysis section for computational details. Download figure Open in new tab Figure 4: Representative structures of the four hydrolysis states illustrating motif IVa and V interactions with RNA phosphates. Key residues from motif IVa (R388 and K389) and motif V (T409 and D410) are shown, as these exhibit significant changes in RNA phosphate affinity (P 1 , P 2 , and P 3 ) across the hydrolysis cycle. Motif IVa and its residues are colored orange; motif V and its residues are colored magenta. The remainder of the protein is shown in translucent grey. The ssRNA is depicted as an ice-blue ribbon, with its phosphate moieties shown in CPK representation (oxygen: red, phosphorus: green). (A) Representative structure of the ssRNA state, with two overlaid snapshots demonstrating R388 sampling both the ‘up’ conformation (near P 1 ) and the ‘down’ conformation (near P 3 ). D410 is positioned near P 2 , while K389 and T409 remain distant from RNA phosphates. (B) Structure of the ssRNA+ATP state showing post-ATP binding rearrangements. R388 continues to sample both conformations, while D410 detaches from P 2 and T409 moves closer to occupy its position. (C) Structure of the ssRNA+ADP+P i state following hydrolysis. R388 transitions to favor binding P 1 , and K389 also forms a close interaction with P 1 . Both motif V residues are detached from the RNA. (D) Structure of the ssRNA+ADP state following P i release. K389 detaches from P 1 , while R388 maintains the ‘up’ conformation. ATP binding induces asymmetric conformational responses in motifs V and IVa , revealing distinct mechanisms of phosphate engagement. Upon ATP binding, motif V forms stable contacts with phosphate P 2 , whereas motif IVa continues to fluctuate between interactions with P 1 and P 3 . Specifically, motif V residue T409 exhibits a calculated ΔE inter of −21.25 ± 1.42 kcal·mol −1 with P 2 , while D410 shows a stronger interaction of −30.16 ± 8.17 kcal·mol −1 ( Figure 3A ). Notably, this apparent increase in affinity for D410 reflects a reduction in repulsive interactions: its interaction energy shifts from 22.24±3.70 kcal·mol −1 in the ssRNA state to 7.83 ± 1.91 kcal·mol −1 in the ssRNA+ATP state. In contrast, motif IVa does not exhibit a net change in affinity for P 1 ( Figure 3B ), although large fluctuations are observed in R388’s interaction with both P 1 (ΔE inter = −3.36 ± 47.86 kcal·mol −1 ) and P 3 (ΔE inter = 7.34 ± 41.10 kcal·mol −1 ) ( Figure 3C ). These findings suggest that ATP binding stabilizes motif V near P 2 , while R388 in motif IVa dynamically samples both an ‘up’ conformation near P 1 and a ‘down’ conformation near P 3 ( Figure 4B ). Following ATP hydrolysis, the RNA-cleft reorganizes: motif V detaches from phosphate P 2 , while motif IVa transitions into a stably bound conformation at P 1 . In this ssRNA+ADP+P i state, we observed a notable weakening in motif V –P 2 interactions, with T409 showing a ΔE inter of 14.38±8.90 kcal·mol −1 ( Figure 3D ), indicative of reduced affinity. D410 remains distant from P 2 and exhibits no significant change, consistent with sustained low repulsion. In contrast, motif IVa forms stronger and more consistent interactions with P 1 : R388 and K389 exhibit ΔE inter values of −37.23 ± 33.81 and −39.90 ± 14.57 kcal·mol −1 , respectively ( Figure 3E-F ). The large uncertainty for R388 reflects its prior fluctuation in the ATP-bound state. Additionally, we observe a weakening of R388–P 3 affinity (ΔE inter = 21.62 ± 18.82 kcal·mol −1 ), indicating a shift away from P 3 and toward stable binding at P 1 . Together, these changes define a hydrolysis-dependent structural reorganization, with motif V detached from P 2 and motif IVa adopting an ‘up’ conformation stabilized at P 1 ( Figure 4C ). A similar hydrolysis-induced R388 ‘up’ conformation was reported in simulations of DENV4 NS3 helicase, 20 although in that system the ‘up’ conformation was also favored in the ATP-bound state—an important contrast to our findings in WNV NS3h. Following P i release, motif IVa begins to disengage from phosphate P 1 , while motif V remains detached from P 2 . During the transition from the ssRNA+ADP+P i state to the ssRNA+ADP state, we observed no significant change in P 2 affinity for either T409 or D410 of motif V ( Figure 3G ), suggesting that T409 maintains a weak interaction and D410 remains non-repulsive. For motif IVa , R388 shows no net change in affinity for P 1 , consistent with a continued strong interaction ( Figure 3H ), and remains weakly associated with P 3 (ΔE inter = 16.66 ±8.04 kcal·mol −1 ; Figure 3I ). In contrast, K389 shows a notable weakening in affinity for P 1 , without a compensatory increase in affinity for P 3 , indicating a net loss of RNA interaction. The smaller error bars in Figures 3H and 3I , relative to those in the earlier state ( Figures 3E–F ), suggest reduced fluctuations in the ssRNA+ADP+P i state. Taken together, these changes yield a partially detached motif IVa , in which R388 retains its ‘up’ conformation at P 1 , while K389 becomes RNA-free. Motif V remains dissociated from P 2 throughout this state ( Figure 4D ). Following ADP release, motif V re-engages partially with P 2 , while motif IVa resumes interactions with both P 1 and P 3 . During the transition from the ssRNA+ADP state to the ssRNA state, we observed a significant change in the interaction energy between D410 and P 2 (ΔE inter = 35.20 ± 8.05 kcal·mol −1 ) ( Figure 3J ), while no net change was detected for other motif V residues. This shift reflects a weakening of D410’s interaction with P 2 , driven by an increase in repulsive affinity in the ssRNA state (22.24 ± 3.70 kcal·mol −1 ) relative to the ssRNA+ADP state (10.20 ± 2.73 kcal·mol −1 ). This repulsion results from D410 reattaching to P 2 while T409 detaches ( Figure 4A ). For motif IVa , R388 exhibits substantial changes in phosphate affinity: a weakening of interaction with P 1 (ΔE inter = 41.52 ± 35.21 kcal·mol −1 ) and a strengthening with P 3 (ΔE inter = −45.62 ± 37.41 kcal·mol −1 ; Figure 3K-L ). The large error bars associated with R388 reflect continued fluctuation between P 1 and P 3 , indicating that this residue resumes sampling of both the ‘up’ and ‘down’ conformations in the ssRNA state ( Figure 4A ). Together, these state-specific conformational changes provide the structural basis for directional phosphate engagement during translocation. The ATP-dependent RNA phosphate affinity and structural shifts of motifs IVa and V support an inchworm-style mechanism for RNA translocation during the hydrolysis cycle. Considering the 3 ′ to 5 ′ translocation direction characteristic of SF2 helicases, we propose the following model. The cycle begins with ATP binding, during which motif IVa releases phosphate P n (corresponding to P 3 ) and advances toward P n+2 (P 1 ), while motif V engages P n+1 (P 2 ). After hydrolysis, motif IVa binds to P n+2 , and motif V detaches from P n+1 . Upon P i release, motif IVa prepares to release P n+2 , and motif V remains unbound. Following ADP release, motif IVa advances toward P n+4 while motif V re-approaches P n+2 . This coordinated progression suggests that motif IVa steps from P n to P n+2 by traversing P n+1 with assistance from motif V . The observed alternation between stable attachment and diffusive sampling, particularly in motif IVa , aligns with a small-step inchworm mechanism. Similar translocation behavior has been proposed in other Orthoflavivirus helicases, 16 , 20 , 38 although a Brownian ratchet model has also been suggested for ZIKV NS3 helicase. 18 ATP-dependent coupling between motif VIL and RNA-cleft contacts is mediated through specific phosphate interactions that shift over the hydrolysis cycle. To pinpoint the RNA-phosphate interactions most associated with motif VIL ’s nucleotide affinity, we performed residue-level correlation analysis across hydrolysis states ( Figure 5A ). We further decomposed these correlations by examining interactions with the two primary ADP-contacting residues of motif VIL , R461 ( Figure 5B ) and R464 ( Figure 5C ). Among all contacts, the RNA phosphate affinity of D410 (motif V ) and K389 (motif IVa ) show only weak correlation with motif VIL ’s ADP affinity ( Figure 5A ). For R388 (motif IVa ), the correlation coefficients between its P 1 affinity and motif VIL ADP affinity are –0.19, 0.06, and –0.31 in the ssRNA+ATP, ssRNA+ADP+P i , and ssRNA+ADP states, respectively. This behavior is primarily driven by coupling to R461 ( Figure 5B ), suggesting an allosteric link between R388 and R461. For R388’s interaction with P 3 , the correlation coefficients were 0.19, 0.27, and –0.28 across the same states, influenced by both R461 and R464 ( Figure 5B–C ). In contrast, the P 2 affinity of T409 (motif V ) exhibited the strongest and most consistent correlation with motif VIL ’s ADP affinity, increasing in magnitude along the hydrolysis cycle. The correlation coefficients are –0.10 (ssRNA+ATP), –0.33 (ssRNA+ADP+P i ), and –0.43 (ssRNA+ADP), reflecting increasingly tight inverse coupling. This trend is primarily mediated by R464 in the ssRNA+ADP+P i state and by R461 in the ssRNA+ADP state, underscoring the functional significance of T409( V )– VIL coupling. Download figure Open in new tab Figure 5: Correlation between RNA phosphate affinity and ADP interaction energy of motif VIL residues. Key RNA-binding residues were identified from ATP-dependent changes in RNA phosphate affinity (see Figure 3 ). The selected phosphate-residue pairs include: P 2 –T409 (motif V ), P 2 –D410 (motif V ), P 1 –R388 (motif IVa ), P 1 –K389 (motif IVa ), P 3 –R388 (motif IVa ), and P 3 –K389 (motif IVa ). (A) Correlation coefficients between the RNA phosphate interaction energies of these residue pairs and the ADP interaction energy of motif VIL , computed for the ssRNA+ATP, ssRNA+ADP+P i , and ssRNA+ADP states. (B, C) Correlations between RNA phosphate affinities of the same residues and the ADP interaction energies of R461 and R464, respectively—two residues previously shown to dominate the ADP binding of motif VIL . 19 Disruption of the coupling between motif VIL and RNA phosphate affinity in motif V impairs the conformational transitions required for RNA translocation. In simulations of motif VIL loop mutant, we observed no ATP-dependent changes in phosphate affinity at the key RNA-cleft contacts of motifs V and IVa (Figure S5), in contrast to the behavior of the wild-type enzyme. This absence of phosphate affinity modulation coincides with the loss of correlation between the P 2 affinity of T409 (motif V ) and the ADP affinity of motif VIL (Figure S6). In the wild-type enzyme, this correlation supports a functional pathway for signal transmission from the ATP-pocket to the RNA-cleft. Based on these observations, we propose that motif V serves as both a receiver of nucleotide-affinity changes from motif VIL and a responder that modulates RNA phosphate affinity to enable translocation. These findings underscore the central role of motif VIL – V coupling in enabling efficient, directional RNA translocation through hydrolysis-state-dependent phosphate affinity shifts. Allosteric correlation between motif VI and RNA affinity of motif V ATP-dependent structural coupling between motif VIL and motif V mirrors their functional coordination in substrate binding. To evaluate whether changes in RNA and nucleotide affinity are accompanied by coordinated structural motion, we calculated the positional correlation between the centers of mass of backbone and sidechain atoms in motifs VIL and V ( Figure 6 ). In the ssRNA state, correlation coefficients range from –0.22 to 0.60 for backbone atoms ( Figure 6A ) and –0.37 to 0.52 for sidechain atoms ( Figure 6E ), indicating moderate coupling. These correlations weaken in the ssRNA+ATP state, with backbone coefficients between –0.07 and 0.44 ( Figure 6B ) and sidechain correlations from –0.33 to 0.41 ( Figure 6F ). In the ssRNA+ADP+P i state, the correlation partially recovers, with backbone values ranging from –0.30 to 0.52 ( Figure 6C ) and sidechain correlations from –0.33 to 0.59 ( Figure 6G ). Finally, in the ssRNA+ADP state, the correlation strengthens further, especially for the backbone (–0.16 to 0.60; Figure 6D ), while sidechain values range from –0.39 to 0.46 ( Figure 6H ). These results indicate that structural coordination between motifs VIL and V is dynamically modulated by the hydrolysis state and parallels the trends observed in their substrate binding affinity. Download figure Open in new tab Figure 6: Identification of key correlated residue pairs between motif V and motif VIL via positional correlation analysis. Positional correlations of the centers of mass for backbone (BB; top row) and sidechain (SC; bottom row) atoms were computed after global alignment of each structure within its respective hydrolysis state ensemble. Panels represent: (A, E) ssRNA, (B, F) ssRNA+ATP, (C, G) ssRNA+ADP+P i , and (D, H) ssRNA+ADP states. Orange squares highlight residue pairs with correlation coefficients greater than 0.55 or less than –0.55, indicating strong positive or negative correlations, respectively. R461–E413 interaction serves as a key structural link between motifs VIL and V during ATP hydrolysis. In our structural correlation analysis, residue pairs exhibiting correlation coefficients greater than |0.55| were marked as strongly coupled and are highlighted in orange in Figure 6 . Among these, the R461–E413 and R464–A416 pairs are particularly notable, with strong correlations emerging in the ssRNA+ADP+P i and ssRNA+ADP states. Notably, the nature of these correlations shifts from sidechain–sidechain to backbone–backbone interactions between states. Structural proximity between R461 and E413 (compared to R464–A416) suggests R461–E413 as a more plausible site of direct interaction (Figure S7). To further probe this interaction, we quantified hydrogen bond (H-bond) occupancy between the R461 sidechain (SC) and both the backbone (BB) and sidechain (SC) of E413. No significant H-bonding was observed in the ssRNA or ssRNA+ATP states (Figure S8). In the ssRNA+ADP+P i state, R461(SC) formed an H-bond with E413(BB) in 72.10 ± 4.78% of frames, while no bonding with E413(SC) is detected ( Figure 7A ). Upon transition to the ss-RNA+ADP state, this interaction shifted: R461(SC)–E413(BB) H-bond occupancy dropped to 6.06 ± 3.95%, whereas R461(SC)–E413(SC) H-bonding increased to 54.94 ± 14.73% ( Figure 7B ). This switching of interaction partners coincides with enhanced ADP affinity of R461 observed in this transition. 19 Importantly, this conformational switch is absent in simulations of the motif VIL mutant (Figure S9), further supporting its functional relevance. Download figure Open in new tab Figure 7: Correlation between R461 and E413 modulates ATP-dependent structural sampling of the motif V 3 10 helix. (A, B) Hydrogen bond (H-bond) sampling between the sidechain of R461 (acceptor) and either the backbone or sidechain of E413 (donor) in the ssRNA+ADP+P i (A) and ssRNA+ADP (B) states. No H-bond formation is observed between these residues in the ssRNA or ssRNA+ATP states (Figure S8). Insets show structural snapshots illustrating the relative positions of E413 and R461. (C) A representative structure highlighting the motif V 3 10 helix, spanning residues D410 to E413. (D–G) Two-dimensional free energy surfaces projected along the ϕ and ψ dihedral angles of E413 for the (D) ssRNA, (E) ssRNA+ATP, (F) ssRNA+ADP, and (G) ssRNA+ADP+P i states. Contours are drawn at intervals of 0.5 k B T, from 0 to 4 k B T. ATP-dependent shifts in sampling indicate structural heterogeneity in the 3 10 helix regulated by the R461–E413 interaction. ATP-dependent interaction between E413 (motif V ) and R461 (motif VIL ) influences the structural heterogeneity of the 3 10 helix in motif V . Residue E413 resides within this helix, which also includes D410 and its preceding residue T409—both of which interact with ssRNA ( Figure 7C ). To evaluate how the E413–R461 interaction affects the 3 10 helix conformation, we computed two-dimensional free energy surfaces in the ϕ – ψ dihedral space ( Figure 7D-G ). In the ssRNA state, the 3 10 helix samples two minima located at (−55 ◦ , −33.5 ◦ ) and (−61 ◦ , −6.5 ◦ ), indicating a broader distribution in ϕ angles ( Figure 7D ). The latter minimum is also sampled in the ssRNA+ATP state ( Figure 7E ), though a distinct intermediate at (−95 ◦ , 0 ◦ ) replaces the second minimum observed in the ssRNA state. In the ssRNA+ADP+P i state, the helix adopts a new minimum at (−59 ◦ , −24.5 ◦ ), along with two additional intermediates at (−71 ◦ , −44.3 ◦ ) and (−103 ◦ , 0 ◦ ) ( Figure 7G ), suggesting a structural shift linked to R461–E413 interaction. Following P i release, in the ssRNA+ADP state—where the R461–E413 interaction site shifts—the helix samples only a single minimum at (−61 ◦ , −20.9 ◦ ) ( Figure 7F ). In motif VIL mutants, which lack this ATP-dependent switching of the R461–E413 interaction, no change in 3 10 helix sampling is observed across hydrolysis states (Figure S10). Conclusions This study investigated the ATP-dependent coupling mechanism that links nucleotide affinity in the ATP-binding pocket to single-stranded RNA (ssRNA) affinity in the RNA-cleft of the West Nile Virus (WNV) NS3 helicase (NS3h). By elucidating the structural basis of this coupling, we aimed to improve our understanding of the ATP hydrolysis-driven translocation process—an essential step in viral genome replication. Using previously generated simulation data for WNV NS3h bound to ssRNA in distinct ATP hydrolysis states (ssRNA, ssRNA+ATP, ssRNA+ADP+P i , and ssRNA+ADP), 19 we built upon earlier findings that identified the motif VI loop as a nucleotide valve controlling ADP affinity across the catalytic cycle. To assess whether this coupling mechanism is functionally conserved, we further analyzed motif VI loop mutants (D471E, D471L), which are known to disrupt nucleotide binding and attenuate viral replication. Our correlation analyses revealed that both motif I and the motif VI loop exhibit coupling between ADP affinity and ssRNA phosphate affinity. However, the motif VI loop uniquely displays an ATP-dependent shift in this correlation, particularly with phosphate interactions involving motif IVa and V . At the residue level, R388 and K389 of motif IVa , along with T409 and D410 of motif V , undergo a sequential change in ssRNA phosphate affinity across the hydrolysis cycle, consistent with the translocation of one phosphate unit per cycle. Notably, the ssRNA affinity of motif V is strongly correlated with the ADP affinity of R461 and R464 in motif VI , previously identified as arginine fingers essential for NTPase activity and nucleotide gating. Further structural analysis revealed a hydrogen bond–mediated connection between E413 (motif V ) and R461 (motif VI ), forming an allosteric bridge that links ATP-pocket dynamics to the RNA-cleft. This interaction modulates the structural heterogeneity of the 3 10 -helix in motif V , a conformational feature directly involved in ssRNA binding. Importantly, this E413–R461 interaction and its downstream structural effects are absent in the motif VI loop mutants, reinforcing its functional significance. Taken together, our findings define a mechanistic conduit by which ATP-dependent conformational changes originating at motif VI propagate through E413 to reshape motif V and modulate ssRNA affinity. These insights not only advance the molecular understanding of viral RNA helicase function but also identify a structurally and energetically conserved allosteric pathway that could serve as a promising target for antiviral drug development against WNV and related Orthoflaviviruses . Acknowledgement Research reported in this manuscript was supported by the National Institute for Allergic and Infectious Diseases of the National Institute of Health under award number R01AI166050. Computational resources for this project were provided by: (1) the High Performance Computing Center at Oklahoma State University supported in part through the National Science Foundation Grant OAC-1531128 and (2) Purdue Anvil under ACCESS project number BIO220160. Funder Information Declared National Institutes of Health, https://ror.org/01cwqze88 , R01AI166050 References (1). ↵ Murray , K. ; Baraniuk , S. ; Resnick , M. ; Arafat , R. ; Kilborn , C. ; Cain , K. ; Shallenberger , R. ; York , T. ; Martinez , D. ; Hellums , J. et al. Risk factors for encephalitis and death from West Nile virus infection . Epidemiology & Infection 2006 , 134 , 1325 – 1332 . OpenUrl CrossRef PubMed Web of Science (2). Roberts , J. A. ; Kim , C. Y. ; Hwang , S. A. ; Hassan , A. ; Covington , E. ; Heydari , K. ; Lyerly , M. ; Sejvar , J. J. ; Hasbun , R. ; Prasad , M. , et al. Clinical, Prognostic, and Longitudinal Functional and Neuropsychological Features of West Nile Virus Neuroinvasive Disease in the United States: A Systematic Review and Meta-Analysis . Annals of Neurology 2025 , (3). ↵ Maxmen , A. The hidden threat of West Nile virus . Nature 2012 , 489 , 349 . OpenUrl CrossRef PubMed (4). ↵ Murray , K. O. ; Mertens , E. ; Desprès , P. West Nile virus and its emergence in the United States of America . Veterinary research 2010 , 41 , 67 . OpenUrl CrossRef PubMed (5). ↵ Bakonyi , T. ; Ferenczi , E. ; Erdélyi , K. ; Kutasi , O. ; Csörgő , T. ; Seidel , B. ; Weissenböck , H. ; Brugger , K. ; Bán , E. ; Nowotny , N. Explosive spread of a neuroinvasive lineage 2 West Nile virus in Central Europe, 2008/2009 . Veterinary microbiology 2013 , 165 , 61 – 70 . OpenUrl CrossRef PubMed (6). ↵ Jani , C. ; Kakoullis , L. ; Abdallah , N. ; Mouchati , C. ; Page , S. ; Colgrove , R. ; Chen , L. H. West Nile virus: another emerging arboviral risk for travelers? Current Infectious Disease Reports 2022 , 24 , 117 – 128 . OpenUrl CrossRef PubMed (7). ↵ Dutta , S. K. ; Langenburg , T. A perspective on current flavivirus vaccine development: a brief review . Viruses 2023 , 15 , 860 . OpenUrl CrossRef PubMed (8). ↵ Bifani , A. M. ; Chan , K. W. K. ; Borrenberghs , D. ; Tan , M. J. A. ; Phoo , W. W. ; Watanabe , S. ; Goethals , O. ; Vasudevan , S. G. ; Choy , M. M. Therapeutics for flaviviral infections . Antiviral Research 2023 , 210 , 105517 . OpenUrl CrossRef PubMed (9). ↵ van den Elsen , K. ; Chew , B. L. A. ; Ho , J. S. ; Luo , D. Flavivirus nonstructural proteins and replication complexes as antiviral drug targets . Current opinion in virology 2023 , 59 , 101305 . OpenUrl CrossRef PubMed (10). ↵ Walter , Z. ; Li , M. ; Molho , M. ; Berish , L. ; Isopi , A. ; O’Mara , M. ; Dittmar , M. ; Nwaezeapu , C. ; Richards , A. ; McCullagh , M. et al. An integrated proteomics approach identifies phosphorylation sites on viral and host proteins that regulate West Nile virus infection . Cell reports 2025 , 44 . (11). ↵ Brault , A. C. ; Huang , C. Y. ; Langevin , S. A. ; Kinney , R. M. ; Bowen , R. A. ; Ramey , W. N. ; Panella , N. A. ; Holmes , E. C. ; Powers , A. M. ; Miller , B. R. A single positively selected West Nile viral mutation confers increased virogenesis in American crows . Nature genetics 2007 , 39 , 1162 – 1166 . OpenUrl CrossRef PubMed Web of Science (12). ↵ Frick , D. N. Helicases as antiviral drug targets . Drug news & perspectives 2003 , 16 , 355 . OpenUrl CrossRef PubMed (13). Fang , J. ; Li , H. ; Kong , D. ; Cao , S. ; Peng , G. ; Zhou , R. ; Chen , H. ; Song , Y. Structure-based discovery of two antiviral inhibitors targeting the NS3 helicase of Japanese encephalitis virus . Scientific reports 2016 , 6 , 34550 . OpenUrl CrossRef PubMed (14). Mastrangelo , E. ; Pezzullo , M. ; De Burghgraeve , T. ; Kaptein , S. ; Pastorino , B. ; Dallmeier , K. ; de Lamballerie , X. ; Neyts , J. ; Hanson , A. M. ; Frick , D. N. et al. Ivermectin is a potent inhibitor of flavivirus replication specifically targeting NS3 helicase activity: new prospects for an old drug . Journal of Antimicrobial Chemotherapy 2012 , 67 , 1884 – 1894 . OpenUrl CrossRef PubMed Web of Science (15). ↵ Selvaratnam , L. ; Willson , T. M. ; Schapira , M. Structural Chemistry of Helicase Inhibition . Journal of Medicinal Chemistry 2025 , (16). ↵ Anindita , P. D. ; Halbeisen , M. ; Řeha , D. ; Tuma , R. ; Franta , Z. Mechanistic insight into the RNA-stimulated ATPase activity of tick-borne encephalitis virus helicase . Journal of Biological Chemistry 2022 , 298 . (17). ↵ Appleby , T. C. ; Anderson , R. ; Fedorova , O. ; Pyle , A. M. ; Wang , R. ; Liu , X. ; Brendza , K. M. ; Somoza , J. R. Visualizing ATP-dependent RNA translocation by the NS3 helicase from HCV . Journal of molecular biology 2011 , 405 , 1139 – 1153 . OpenUrl CrossRef PubMed (18). ↵ García-Martínez , A. ; Zinovjev , K. ; Ruiz-Pernía , J. J. ; Tunõń , I. Conformational changes and ATP hydrolysis in Zika helicase: the molecular basis of a biomolecular motor unveiled by multiscale simulations . Journal of the American Chemical Society 2023 , 145 , 24809 – 24819 . OpenUrl CrossRef PubMed (19). ↵ Roy , P. ; Walter , Z. ; Berish , L. ; Ramage , H. ; McCullagh , M. Motif-VI loop acts as a nucleotide valve in the West Nile Virus NS3 Helicase . Nucleic Acids Research 2024 , gkae500 . (20). ↵ Davidson , R. B. ; Hendrix , J. ; Geiss , B. J. ; McCullagh , M. Allostery in the dengue virus NS3 helicase: Insights into the NTPase cycle from molecular simulations . PLoS Computational Biology 2018 , 14 , e1006103 . OpenUrl CrossRef (21). ↵ Pérez-Villa , A. ; Darvas , M. ; Bussi , G. ATP dependent NS3 helicase interaction with RNA: insights from molecular simulations . Nucleic Acids Research 2015 , 43 , 8725 – 8734 . OpenUrl CrossRef PubMed (22). ↵ Mastrangelo , E. ; Bolognesi , M. ; Milani , M. Flaviviral helicase: insights into the mechanism of action of a motor protein . Biochemical and Biophysical Research Communications 2012 , 417 , 84 – 87 . OpenUrl CrossRef PubMed (23). ↵ Du Pont , K. E. ; McCullagh , M. ; Geiss , B. J. Conserved motifs in the flavivirus NS3 RNA helicase enzyme . Wiley Interdisciplinary Reviews: RNA 2022 , 13 , e1688 . OpenUrl CrossRef (24). ↵ Jorgensen , W. L. ; Chandrasekhar , J. ; Madura , J. D. ; Impey , R. W. ; Klein , M. L. Comparison of simple potential functions for simulating liquid water . The Journal of Chemical Physics 1983 , 79 , 926 – 935 . OpenUrl CrossRef PubMed Web of Science (25). ↵ Lee , T.-S. ; Cerutti , D. S. ; Mermelstein , D. ; Lin , C. ; LeGrand , S. ; Giese , T. J. ; Roitberg , A. ; Case , D. A. ; Walker , R. C. ; York , D. M. GPU-accelerated molecular dynamics and free energy methods in Amber18: performance enhancements and new features . Journal of Chemical Information and Modeling 2018 , 58 , 2043 – 2050 . OpenUrl CrossRef PubMed (26). ↵ Maier , J. A. ; Martinez , C. ; Kasavajhala , K. ; Wickstrom , L. ; Hauser , K. E. ; Simmerling , C. ff14SB: improving the accuracy of protein side chain and backbone parameters from ff99SB . Journal of Chemical Theory and Computation 2015 , 11 , 3696 – 3713 . OpenUrl CrossRef (27). ↵ Banás , P. ; Hollas , D. ; Zgarbová , M. ; Jurecka , P. ; Orozco , M. ; Cheatham III , T. E. ; Sponer , J. ; Otyepka , M. Performance of molecular mechanics force fields for RNA simulations: stability of UUCG and GNRA hairpins . Journal of Chemical Theory and Computation 2010 , 6 , 3836 – 3849 . OpenUrl CrossRef (28). ↵ Zgarbová , M. ; Otyepka , M. ; Sponer , J. ; Mládek , A. ; Banáš , P. ; Cheatham III , T. E. ; Jurecka , P. Refinement of the Cornell et al. nucleic acids force field based on reference quantum chemical calculations of glycosidic torsion profiles . Journal of Chemical Theory and Computation 2011 , 7 , 2886 – 2902 . OpenUrl CrossRef (29). ↵ Meagher , K. L. ; Redman , L. T. ; Carlson , H. A. Development of polyphosphate parameters for use with the AMBER force field . Journal of Computational Chemistry 2003 , 24 , 1016 – 1025 . OpenUrl CrossRef PubMed Web of Science (30). ↵ Li , P. ; Roberts , B. P. ; Chakravorty , D. K. ; Merz Jr , K. M. Rational design of particle mesh Ewald compatible Lennard-Jones parameters for+ 2 metal cations in explicit solvent . Journal of Chemical Theory and Computation 2013 , 9 , 2733 – 2748 . OpenUrl CrossRef (31). ↵ Luo , D. ; Xu , T. ; Watson , R. P. ; Scherer-Becker , D. ; Sampath , A. ; Jahnke , W. ; Yeong , S. S. ; Wang , C. H. ; Lim , S. P. ; Strongin , A. et al. Insights into RNA unwinding and ATP hydrolysis by the flavivirus NS3 protein . The EMBO journal 2008 , 27 , 3209 – 3219 . OpenUrl Abstract / FREE Full Text (32). ↵ Gu , M. ; Rice , C. M. Three conformational snapshots of the hepatitis C virus NS3 helicase reveal a ratchet translocation mechanism . Proceedings of the National Academy of Sciences 2010 , 107 , 521 – 528 . OpenUrl Abstract / FREE Full Text (33). ↵ Swarbrick , C. M. ; Basavannacharya , C. ; Chan , K. W. ; Chan , S.-A. ; Singh , D. ; Wei , N. ; Phoo , W. W. ; Luo , D. ; Lescar , J. ; Vasudevan , S. G. NS3 helicase from dengue virus specifically recognizes viral RNA sequence to ensure optimal replication . Nucleic acids research 2017 , 45 , 12904 – 12920 . OpenUrl CrossRef PubMed (34). ↵ Dumont , S. ; Cheng , W. ; Serebrov , V. ; Beran , R. K. ; Tinoco Jr , I. ; Pyle , A. M. ; Bustamante , C. RNA translocation and unwinding mechanism of HCV NS3 helicase and its coordination by ATP . Nature 2006 , 439 , 105 – 108 . OpenUrl CrossRef PubMed (35). ↵ Matusan , A. E. ; Pryor , M. J. ; Davidson , A. D. ; Wright , P. J. Mutagenesis of the Dengue virus type 2 NS3 protein within and outside helicase motifs: effects on enzyme activity and virus replication . Journal of virology 2001 , 75 , 9633 – 9643 . OpenUrl Abstract / FREE Full Text (36). Benarroch , D. ; Selisko , B. ; Locatelli , G. A. ; Maga , G. ; Romette , J.-L. ; Canard , B. The RNA helicase, nucleotide 5 ′ -triphosphatase, and RNA 5 ′ -triphosphatase activities of Dengue virus protein NS3 are Mg2+-dependent and require a functional Walker B motif in the helicase catalytic core . Virology 2004 , 328 , 208 – 218 . OpenUrl CrossRef PubMed (37). ↵ Du Pont , K. E. ; Davidson , R. B. ; McCullagh , M. ; Geiss , B. J. Motif V regulates energy transduction between the flavivirus NS3 ATPase and RNA-binding cleft . Journal of Biological Chemistry 2020 , 295 , 1551 – 1564 . OpenUrl Abstract / FREE Full Text (38). ↵ Sampath , A. ; Xu , T. ; Chao , A. ; Luo , D. ; Lescar , J. ; Vasudevan , S. G. Structure-based mutational analysis of the NS3 helicase from dengue virus . Journal of virology 2006 , 80 , 6686 – 6690 . OpenUrl Abstract / FREE Full Text (39). ↵ Weber , R. ; McCullagh , M. Role of ATP in the RNA translocation mechanism of SARS-CoV-2 NSP13 helicase . The Journal of Physical Chemistry B 2021 , 125 , 8787 – 8796 . OpenUrl CrossRef PubMed (40). ↵ Lawal , M. M. ; Roy , P. ; McCullagh , M. Role of ATP hydrolysis and product release in the translocation mechanism of SARS-CoV-2 NSP13 . The Journal of Physical Chemistry B 2024 , 128 , 492 – 503 . OpenUrl CrossRef PubMed View the discussion thread. Back to top Previous Next Posted July 04, 2025. Download PDF Supplementary Material Email Thank you for your interest in spreading the word about bioRxiv. NOTE: Your email address is requested solely to identify you as the sender of this article. Your Email * Your Name * Send To * Enter multiple addresses on separate lines or separate them with commas. You are going to email the following Allosteric Regulation of RNA Affinity by Motif V-VI Coupling in West Nile Virus NS3 Helicase Message Subject (Your Name) has forwarded a page to you from bioRxiv Message Body (Your Name) thought you would like to see this page from the bioRxiv website. Your Personal Message CAPTCHA This question is for testing whether or not you are a human visitor and to prevent automated spam submissions. Share Allosteric Regulation of RNA Affinity by Motif V-VI Coupling in West Nile Virus NS3 Helicase Priti Roy , Martin McCullagh bioRxiv 2025.07.04.663198; doi: https://doi.org/10.1101/2025.07.04.663198 Share This Article: Copy Citation Tools Allosteric Regulation of RNA Affinity by Motif V-VI Coupling in West Nile Virus NS3 Helicase Priti Roy , Martin McCullagh bioRxiv 2025.07.04.663198; doi: https://doi.org/10.1101/2025.07.04.663198 Citation Manager Formats BibTeX Bookends EasyBib EndNote (tagged) EndNote 8 (xml) Medlars Mendeley Papers RefWorks Tagged Ref Manager RIS Zotero Tweet Widget Facebook Like Google Plus One Subject Area Biophysics Subject Areas All Articles Animal Behavior and Cognition (7642) Biochemistry (17715) Bioengineering (13907) Bioinformatics (42003) Biophysics (21470) Cancer Biology (18624) Cell Biology (25533) Clinical Trials (138) Developmental Biology (13390) Ecology (19935) Epidemiology (2067) Evolutionary Biology (24356) Genetics (15617) Genomics (22529) Immunology (17753) Microbiology (40432) Molecular Biology (17200) Neuroscience (88681) Paleontology (667) Pathology (2840) Pharmacology and Toxicology (4828) Physiology (7653) Plant Biology (15161) Scientific Communication and Education (2046) Synthetic Biology (4304) Systems Biology (9826) Zoology (2271)