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. Author manuscript; available in PMC: 2017 Apr 5.
Published in final edited form as: Structure. 2016 Apr 5;24(4):509–517. doi: 10.1016/j.str.2016.02.019

Long-range communication between different functional sites in the picornaviral 3C protein

Yan M Chan 1, Ibrahim M Moustafa 2, Jamie J Arnold 2, Craig E Cameron 2, David D Boehr 1,*
PMCID: PMC4824962  NIHMSID: NIHMS765560  PMID: 27050688

Summary

The 3C protein is a master regulator of the picornaviral infection cycle, responsible for both cleaving viral and host proteins, and interacting with genomic RNA replication elements. Here we use nuclear magnetic resonance spectroscopy and molecular dynamics simulations to show that 3C is conformationally dynamic across multiple timescales. Binding of peptide and RNA lead to structural dynamics changes at both the protease active site and the RNA binding site, consistent with these sites being dynamically coupled. Indeed, binding of RNA influences protease activity, and likewise, interactions at the active site affect RNA binding. We propose that RNA and peptide binding reshapes the conformational energy landscape of 3C to regulate subsequent functions, including formation of complexes with other viral proteins. The observed channeling of the 3C energy landscape may be important for regulation of the viral infection cycle.

Introduction

Viral genomes tend to be quite compact, but have evolved to maximize their information content. One solution to the limited coding capacities of RNA viruses is to combine multiple functions into single proteins. The different functions of these viral proteins may require them to interact with multiple viral RNA sequences and/or viral and host proteins (e.g. (Bedard and Semler, 2004; Daijogo and Semler, 2011; Sola et al., 2011; Steil and Barton, 2009; Yin et al., 2007; Yu et al., 2013). In general, it is thought that not all functions occur simultaneously and that ligand binding to one active/binding site may shut down the other. How this communication occurs is not known.

The picornavirus 3C protein is a great model system for understanding the regulation of multiple functions in a viral protein. The 3C protein can also be found as a domain within the 3CD and 3BCD polyproteins (Cameron et al., 2010). The 3C(D) protein is the major protease responsible for most of the polyprotein cleavage events (Cameron et al., 2010; Jore et al., 1988; Parsley et al., 1999; Ypma-Wong et al., 1988), and is also important for regulating the host cell's response through cleavage of critical host cell proteins (Chase et al., 2014; de Breyne et al., 2008; Kuyumcu-Martinez et al., 2004; Rozovics et al., 2012; Walker et al., 2013; Yalamanchili et al., 1997; Zhou et al., 2013). The 3C protein and/or its precursors also interact with cis-acting replication elements (CREs) located within the viral genome (Steil and Barton, 2009). Interactions with these RNA structures are important for regulating replication and translation events (Blair et al., 1998; Franco et al., 2005; Gamarnik and Andino, 1998; Nayak et al., 2006; Parsley et al., 1997; Shih et al., 2004; Walker et al., 1995; Yang et al., 2004). It is not completely clear if the two functions (i.e. protease and RNA-binding) of 3C are interrelated. In Norovirus (NV) 3C-like protease, RNA binding was found to impair the protease catalytic rate (Viswanathan et al., 2013), but similar studies have not been done with picornaviral 3C proteins. However, amino acid changes at the RNA binding site of Enterovirus 71 (EV71) 3C decreased the protease activity, but amino acid changes at the protease active site had little impact on RNA binding (Shih et al., 2004). The protease activity of the HRV 14 3C was likewise affected by amino acid changes at the RNA binding site (Walker et al., 1995).

The goals of the current research were to determine if protein/peptide and RNA binding modulate the RNA-binding and protease activities of poliovirus (PV) 3C, respectively, and if so, to delineate long-range interactions important for this regulation. Regulation of the two functions of 3C through binding of the other ligand may be important in the virus infection cycle. In this manuscript, we indeed demonstrate that RNA and peptide binding alter proteolytic activity and RNA binding, respectively. We also identify millisecond conformational exchange events by NMR R2 relaxation dispersion experiments near the protease active site that are modulated by RNA binding. These findings suggest that RNA and peptide binding selects different subsets of 3C conformations, thereby regulating subsequent 3C activities.

Results

Mutual regulation of the two functions of PV 3C

Previous studies indicated that RNA binding impairs the catalytic rate of the NV 3C-like protease (Viswanathan et al., 2013). These studies and others (Shih et al., 2004; Walker et al., 1995) suggested that the two functions of 3C may be interrelated. To test this hypothesis, we analyzed the protease activity and RNA binding functions of PV 3C in the presence of the other ligand. In our studies, we used two versions of 3C: proteolytically-active (3Cactive) and proteolytically-inactive (3Cinactive) protein in which the active site cysteine has been changed to an alanine and the remaining cysteine has been changed to a serine (i.e. 3Cinactive has the substitutions C147A and C153S). Besides the protease activity, we do not expect significant functional and/or structural dynamic differences between 3Cactive and 3Cinactive. The C153S substitution is on the surface of the protein and not near either the protease active site or the predicted RNA binding site. For RNA binding, we used either the oriI (origin of replication Internal) RNA or the stem left (SL) region of the oriI RNA (Figure 1). For the peptide, we used either the CA peptide, which corresponds to the cleavage site between 2C and 3A of the polyprotein, or the AB peptide, which corresponds to the cleavage site between 2A and 2B of the viral polyprotein (Figure 1).

Figure 1.

Figure 1

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The poliovirus 3C protease has multiple functions. (A) The structure of the 3C protease (PDB 1L1N) showing the locations of the proposed RNA binding site (in green spheres) and the catalytic triad (in black spheres). (B) 3C interacts with viral RNA control sequences including the internal origin of replication (oriI) site. The stem left (SL) RNA that is used in these studies corresponds to the nucleotide sequence shown in blue. (C) 3C catalyzes the hydrolysis of peptide bonds in the viral polyprotein, including between 2C and 3A, and 2A and 2B. These cleavage sites are represented by the CA and AB peptides respectively. The two C-terminal Arg residues (underlined) are added to the peptide to increase solubility, but they are not predicted to interact with 3C. See also Figure S1.

To probe the effects of SL RNA binding on 3Cactive protease activity, we adopted a FRET-based (fluorescence resonance energy transfer) protease assay in which proteolytic activity releases the N-terminus attached fluorophore from the C-terminus attached quencher group (Hata et al., 2000) (Figure 1). We performed a series of experiments with different SL RNA concentrations, allowing us to perform ANOVA (one way analysis of variance) and post hoc Tukey HSD (honest significant difference) statistical tests. The presence of the SL RNA had a small, but statistically significant (i.e. ANOVA p-value ∼ 0.01; HSD p-value for the largest SL RNA concentration ∼0.01) effect on the catalytic turnover rate constant (kcat) when using a derivative of the CA peptide as substrate, but did not have a statistically significant effect on the Michaelis-Menten constant (KM) (Table 2).

Table 2. RNA binding has a small influence on 3C protease activity.

3Cactive : SL RNA ratio KM (μM) kcat (sec-1) kcat/KM (105 sec-1M-1)
1:0 62.73 ± 31.34 31.78 ± 8.76 5.07 ± 1.52
1:1 48.96 ± 20.71 34.04 + 7.32 6.95 ± 1.73
1:2 53.61 ± 16.23 38.31 ± 6.08 7.17 ± 1.40
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We were also interested in probing the effect of peptide on RNA binding. As such, we used fluorescence polarization experiments to monitor RNA binding, which is fluorescein-labeled on the 3′ end, to 3Cinactive in the absence and presence of peptide. For these experiments, we monitored binding of SL RNA and oriI RNA in the presence/absence of CA and AB peptides (Figure 1). Again, we performed a series of experiments with different concentrations of CA and AB peptides allowing us to perform ANOVA and Tukey HSD statistical tests. The presence of the CA peptide did not significantly alter the dissociation constant (Kd) associated with SL RNA binding (Table 1; ANOVA p-value ∼ 0.61), but led to a small decrease in the oriI Kd (ANOVA p-value ∼ 0.02). In contrast, the presence of the AB peptide resulted in more substantial changes in both SL and oriI RNA binding (Table 1). The presence of the AB peptide led to a decrease in SL RNA binding affinity (ANOVA p-value ∼ 0.0001), but an increase in oriI RNA binding affinity (ANOVA p-value ∼ 0.0006; HSD p-value for the largest peptide concentration ∼0.0005). These results suggest that the effect of peptide binding on RNA affinity is peptide-sequence dependent, and these effects are also dependent on the sequence/structure of the RNA itself.

Table 1. Peptide binding influences RNA binding affinity.

RNA peptide 3Cinactive:peptide ratio Kd for RNA (μM) Kd (-peptide)/Kd (+peptide)
SL CA 1:0 0.76 ± 0.14
1:1 1.23 ± 0.16 0.62
1:5 0.87 ± 0.16 1.14
AB 1:1 1.74 ± 0.44 0.44
1:5 1.63 ± 0.42 0.47
oriI CA 1:0 12.83 ± 4.65
1:2 10.84 ± 7.95 1.18
1:5 9.76 ± 3.50 1.31
AB 1:1 9.01 ± 3.08 1.42
1:2 6.67 ± 0.86 1.92
1:5 4.01 ± 0.43 3.20
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Poliovirus 3C is a monomer in solution

Our studies indicated that RNA and peptide binding modulate the protease and RNA binding activities of PV 3C, respectively. To gain more insight into the regulation of these two functions, we characterized these interactions through nuclear magnetic resonance (NMR) methodology. The interpretation of our solution-state NMR experiments on PV 3C is dependent on the oligomeric state of the protein. According to dynamic light scattering (DLS) and NMR analyses, 3Cactive and 3Cinactive are both monomeric in solution. The theoretical translation diffusion coefficients for monomeric and dimeric 3C are 9.9 × 10−11 m2/s and 7.5 × 10−11 m2/s, respectively (Amero et al., 2008). The translational diffusion coefficient for 3Cinactive was estimated to be 9.6 × 10−11 m2/s by NMR and 9.3 × 10−11 m2/s by DLS. Similarly, the translational diffusion coefficient for 3Cactive was estimated to be 9.9 × 10-11 m2/s by NMR and 9.4 × 10-11 m2/s by DLS. Addition of the reducing agent β-mercaptoethanol to 3Cactive (which has two Cys residues) did not substantially change the measured translational diffusion coefficients (9.7 × 10-11 m2/s by NMR and 9.3 × 10-11 m2/s by DLS). These results are consistent with previous NMR studies (Amero et al., 2008) and in contrast to the finding that PV 3C crystallizes as a dimer (Mosimann et al., 1997). These results are reminiscent of previous findings where NV 3C-like protease was found to be a dimer in the X-ray crystal structure (Zeitler et al., 2006), but a monomer in the NMR solution-state structure (Takahashi et al., 2013).

RNA and peptide binding lead to long-range NMR chemical shift perturbations

To gain some insight into the long-range effects of peptide and RNA binding, we analyzed NMR chemical shift perturbations (i.e. through 1H-15N HSQC spectra) induced upon binding peptide or RNA. For these experiments, we only used the SL RNA because previous DLS experiments had indicated that in the presence of the full 29-nt oriI RNA PV 3C forms a multimer, which would severely complicate NMR analysis; this does not occur with the SL RNA (Amero et al., 2008).

We identified resonances with appreciable chemical shift changes as those showing chemical shift changes at least one standard deviation above the average chemical shift change across all resonances. Appreciable chemical shift changes induced by SL RNA binding are associated with amino acid residues in two clusters (Figure 2A). One cluster is close by the KFRDI motif, which was identified by previous NMR experiments (Amero et al., 2008; Claridge et al., 2009) and mutational studies (Andino et al., 1993; Blair et al., 1998; Hammerle et al., 1992; Leong et al., 1993; Nayak et al., 2006; Shih et al., 2004; Walker et al., 1995) as being important for RNA binding. This cluster included residues 81-89 on the h3 helix, residues on the N-terminal h1 helix (adjacent to h3), residues 29-32 on the β2 strand (nearby h3), and residues 156 and 157 on the β11 strand (nearby h3). It should also be noted that upon binding SL RNA, some residues on the h3 helix showed two resonances (Figure 3; e.g. Asp85 and His89), which is consistent with conformational exchange on the slow NMR timescale. This effect is not due to the use of subsaturating concentrations of RNA; under these conditions, we expect that 99.6% of 3C is bound with SL RNA (based on Kd of 0.76 μM; see Table 1). The other cluster of residues show small chemical shift changes and correspond to residues nearby the protease active site, including Asn165 and Phe170 on the C-terminal side and His40 and Glu63 on the N-terminal side of the center cleft between the two β-barrel sub-domains. These findings are similar to those for HRV-14 3C, which also suggested that structural changes around the protease active-site were induced upon binding RNA (Claridge et al., 2009).

Figure 2.

Figure 2

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RNA and peptide binding perturb the chemical environment around the other ligand's binding site. 1H-15N HSQC spectra were compared to 3Cinactive without and 3Cinactive bound with (A) SL RNA or (B) CA peptide. NMR chemical shift perturbations were calculated using the equation Δδcombined=(ΔδH2+(ΔδN/5)2)1/2 where ΔδH and ΔδN are the chemical shift differences between 3C with and without the appropriate ligand for the backbone amide proton and nitrogen, respectively. (C) Residues showing substantial chemical shift perturbations in the presence of RNA and/or peptide are plotted as spheres on the 3C X-ray crystal structure (PDB 1L1N), where those with chemical shift perturbations one and two standard deviations above the average are shown respectively in light blue (orange) and darker blue (red) for SL RNA (peptide) binding. Residues that show substantial chemical shift pertubations for both SL RNA and peptide binding are shown in purple. NMR spectra were collected at 298 K with ∼215 μM 3Cinactive in the presence/absence of 215 μM SL RNA or 215 μM CA peptide using a buffer consisting of 10 mM HEPES pH 7.5 and 50 mM NaCl. See also Figure S2.

Figure 3.

Figure 3

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The conformational ensemble of 3C is different when bound with both peptide and RNA compared to when it is bound with only one ligand. (A) Spectral overlays of specific resonances from 1H-15N HSQC spectra collected for ligand-free 3Cinactive (black), and 3Cinactive bound with SL RNA (blue) or CA peptide (red) or both (purple). (B) The NMR spectra of the ternary complex bound with peptide and SL RNA also depends on the order of ligand binding. The resonances are colored purple and cyan for when SL RNA and CA peptide are added first respectively. (C) Locations of residues associated with two or more resonances in the ternary complexes bound with peptide and RNA are shown as magenta colored spheres. NMR spectra were collected at 298 K with ∼215 μM 3Cinactive in the presence/absence of 215 μM SL RNA or 215 μM CA peptide using a buffer consisting of 10 mM HEPES pH 7.5 and 50 mM NaCl. See also Figure S3.

For CA peptide binding, residues associated with substantial chemical shift changes also appear in two clusters (Figure 2B). One cluster is located in the region between the two β-barrel sub-domains where the catalytic triad lies, including residues 23-25, 40-41 on the N-terminal β-barrel subdomain, and residues 102, 116, 132, 137, 142, 148, 165, 168-171 on the C-terminal β-barrel sub-domain. The other cluster is nearby the predicted RNA binding site, including Arg176 and Thr180 on the C-terminal h4 helix, and His89 on the h3 helix adjacent to the C-terminal h4 helix. Binding of AB peptide resulted in a similar pattern of chemical shift perturbations as that induced by the CA peptide, although the chemical shift perturbations were generally of lower magnitude (Figure S2).

RNA and peptide select different sets of 3C conformations

It is intriguing that both SL RNA and peptide binding induced chemical shift changes to resonances belonging to two clusters of amino acid residues (Figure 2C). One cluster is nearby the predicted ligand-binding site, and the other cluster is nearby the predicted binding site of the other ligand. These findings are intriguing considering that we have shown that RNA/peptide binding can affect the interactions of 3C with the other ligand (i.e. peptide/RNA; Tables 1 and 2). To gain more insight into these potential cooperative effects, we formed the binary complexes with either RNA or peptide, and then titrated in peptide or RNA, respectively. Remarkably, some of the resonances associated with these ternary complexes were at different chemical shift positions than that observed for either of the binary complexes (including catalytic residues His40 and Glu71) (Figure 3A). In fact, many of these residues were associated with two resonances, again suggestive of conformational exchange on the slow NMR timescale. Perhaps even more remarkable was the finding that the chemical shift positions of these resonances depended on whether SL RNA or CA peptide was added first (Figure 3B). Many of the residues associated with multiple resonances in the ternary complexes (Figure 3C) are near the 3C dimer interface as observed by crystallographic studies (Mosimann et al., 1997) and/or near interfaces proposed to be important for interacting with other PV proteins (Shen et al., 2008).

RNA binding selects a different ensemble of conformations

The appearance of two resonances for some amino acid residues in the RNA binary or ternary complexes suggested that there are multiple conformations of 3Cinactive in solution exchanging on the slow NMR timescale. To probe this further, we performed NMR R2 relaxation dispersion experiments, which report on protein structural dynamics on the μs-ms timescale (Loria et al., 2008) (Figure 4A). Unfortunately, we were not able to perform these experiments in the presence of CA peptide, because we could not achieve saturating conditions with the peptide due to its low solubility.

Figure 4.

Figure 4

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The binding of RNA selects for a different subset of 3C conformations. (A) Example NMR R2 relaxation dispersion curves collected at 1H Larmor frequencies of 600 (black) and 850 (red) MHz. Residues displaying Rex values greater than 5 s-1 at 850 MHz for (B) ligand-free 3Cinactive and (C) 3Cinactive bound with SL RNA are plotted as colored spheres. (D) Comparison of the dynamic chemical shift changes (Δω) derived from the R2 relaxation dispersion curves between ligand-free 3Cinactive and SL RNA-bound 3Cinactive. It should be noted that some residues that show Δω > 4.5 ppm are not plotted. Residues with substantially different Δω in the presence/absence of SL RNA are also plotted as blue spheres in (C). The R2 relaxation dispersion experiments were conducted at 295 K using a buffer consisting of 25 mM potassium phosphate pH 8.0 and 150 mM NaCl. See also Table S1.

One parameter that can be gleaned from the R2 relaxation dispersion experiments is Rex, which is the contribution to R2 from conformational exchange, and indicates those regions of the protein undergoing conformational exchange on the μs-ms timescale. For 3C without ligand, the resonances with substantial Rex (i.e. > 5 s-1) are associated with residues at the center cleft near the protease active-site and residues near the N-terminal h1 helix (Figure 4B, S3). The corresponding amino acid residues include Tyr6, Asn14 and Ile15 on the N-terminal h1 helix, Phe25 on the β2 strand adjacent to the N-terminal h1 helix, Asn105 on the loop adjacent to the h1 helix, Lys175, Phe179, and Gln181 on the h4 helix of the C-terminus, Leu37 and residues 60-75 on the N-terminal β-barrel sub-domain, and Gln122, Gly123, Ala133, Met160, Ala171 and Ala172 on the C-terminal β-barrel sub-domain. Similar patterns of conformational exchange were observed when 3Cinactive was bound with SL RNA, although there were a few additional regions that showed conformational exchange (including residues 13, 100-103, 108, 116, 150 and 162).

Data from the R2 relaxation dispersion experiments can also give more quantitative information about the kinetics and thermodynamics of conformational exchange. Assuming conformational exchange between two conformations (say conformations A and B), R2 relaxation dispersion experiments can yield the exchange rate constant kex that is the sum of the forward (kA→B) and reverse (kB→A) rate constants, the populations of the exchange conformations (pA, pB), and the dynamic chemical shift difference between the exchanging conformations (Δω = δωA - δωB) (Loria et al., 1999). Generally, experimental data for different residues are fit with global kex and pa/pb values, and residue-specific Δω values. The global kex and pB values for 3Cinactive without RNA (kex = 480±22 s-1, pb = 0.020±0.001) and with SL RNA (kex = 518±19 s-1, pB = 0.023±0.001) were very similar. It should be noted that residues undergoing conformational exchange are generally not those that show substantial chemical shift perturbations upon binding SL RNA (Figure 2A; Table S1), so comparisons of Δω values can give some insight into how the binding of SL RNA affects the nature of the higher energy protein conformation (Figure 4D). Many of the Δω values correlate between the apo and RNA-bound states, however, there are a group of residues in which the Δω values for the SL RNA binary complex are substantially higher than the Δω values for 3Cinactive without RNA. These residues tend to be spatially close to the protease active site (Figure 4C; residues shown in blue). Many of these residues, or residues nearby, show two or more resonances in the RNA/peptide ternary complexes (Figure 3). It is also interesting that SL RNA binding increased kcat by a similar amount (∼1.2 fold) as it increased the back rate constant for the R2 relaxation dispersion experiments (kB→A = 9.6 s-1 and 11.9 s-1 for without and with SL RNA, respectively). Altogether, these results suggest that binding of SL RNA selects for a different subset of 3C conformations, and these conformations may interact with peptides at the active site differently than 3C without RNA.

Nanosecond dynamics of 3C are also modulated by RNA binding

The NMR experiments indicated that 3C is highly dynamic on the microsecond-millisecond timescale. We also probed 3Cactive structural dynamics on the nanosecond timescale using all-atoms molecular dynamic (MD) simulations and experimentally through further NMR experiments. For the MD simulations, the 3Cactive monomer extracted from the crystal structure (PDB 1L1N) was immersed in a box filled with water solvent and subjected to 100 ns MD simulation. Analysis of the MD trajectory revealed that, on average, the 3Cactive structure did not show substantial deviations from the starting crystal structure during simulation (RMSD = 1.52 Å). Nevertheless, the amino terminal residues 1-13 showed large deviations (Figure 5A); the first five residues appeared more extended towards solution in comparison to the crystal structure. Many of these residues were identified as being associated with substantial chemical shift changes upon binding RNA.

Figure 5.

Figure 5

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Nanosecond dynamics nearby the protease active-site and RNA binding site. (A) Shown is the average 3Cactive structure (grey), calculated from the last 50 ns of the MD simulation, superimposed on the 3C crystal structure (light blue, PDB 1L1N). Apart from the N-terminal residues (aa 1-13), the two structures superimpose well with an RMSD of 1.52 Å. (B) The calculated per-RMSD across the last 50 ns of the trajectory is plotted as a function of residue. Residues corresponding to the peaks are indicated by red arrows and labeled. The N-terminal residues appeared show the largest amplitude dynamics during the simulation. (C) The per-RMSD in (B) is mapped onto the average 3C structure. Residues corresponding to the highest 25% of per-RMSD data (>1.6) are colored red, residues corresponding to the lowest 25% of per-RMSD data (<0.8) are colored green, and residues with per-RMSD values in the range 0.8-1.6 are colored grey. (left) Residues that showed largest perturbations in chemical shifts due to peptide binding are displayed as spheres and labeled; these residues, with the exception of His168, revealed a small-to-moderate dynamics during simulation. In this view, the peptide-binding site is at the front. (right) A different view in which the RNA-binding site is at the front. Residues that showed largest perturbations in chemical shifts upon RNA-binding are displayed as spheres and labeled; these residues revealed moderate-to-large dynamics during the simulation. See also Figure S4.

To obtain information on the structural dynamics of individual amino acids, we performed per-residue RMSD analysis (per-RMSD) across the last 50 ns of the trajectory (Figure 5B). In this analysis, the average positional deviations of individual residues, including heavy sidechain atoms, were calculated relative to the starting structure. The N-terminal residues 1-13 revealed the largest deviations with per-RMSD values in the range 2.7-14.1 Å. Most of the 3C residues, 75% of the total residues, exhibited per-RMSD values lower than 1.6 Å. The residues that showed relatively large deviations are distributed across the entire 3C structure with peaks at positions 22, 31, 45, 52, 65, 81, 93, 113, 130, 143, 167 and 180 (Figure 5B). Dynamics of residues around positions 22, 65, 130, 143 and 167 are expected to affect the binding of the peptide substrate. Whereas, dynamics of residues around positions 31, 52, 81 and 113 could impact RNA binding. Mapping of the per-RMSD values on the 3C structure revealed that almost all residues involved in peptide-binding that showed large chemical shift perturbations exhibited low-to-moderate amplitude nanosecond dynamics (Figure 5C), whereas, residues involved in RNA-binding exhibited moderate-to-large amplitude nanosecond dynamics (Figure 5C).

To help experimentally verify some of the MD results, we also attempted to collect T1 longitudinal and T2 transverse relaxation times, but unfortunately, the experimental data were not of sufficient quality for Lipari-Szabo model-free analysis (Lipari and Szabo, 1982a, b). Nonetheless, we were able to gain some qualitative insight by analyzing the steady-state 1H-15N heteronuclear Overhauser effects (hetNOEs) (Figure S4), which tend to correlate with order parameters (S2). According to these results, the most flexible region is the loop between the β6 strand and the α3 helix that connects the two β-barrel sub-domains. The MD simulations also show that this region is dynamic on the nanosecond timescale (Figure 5).

With the NMR experiments, we were also able to evaluate dynamic changes induced by the presence of RNA. Addition of SL RNA led to some local changes to the hetNOE values for 3Cinactive. Phe25 and Gly29 on the interface of the N-terminal β-barrel sub-domain and Val162 and Ala172 on the interface of the C-terminal β-barrel sub-domain had increased hetNOE values in the presence of SL RNA, suggestive of decreased picosecond-nanosecond dynamics, whereas Gln65, Glu71, Glu121, Arg134, Leu136 and Ala173 had decreased hetNOE values (i.e. increased dynamics) in the presence of SL RNA.

Discussion

Viral genomes are typically small. One way viruses maximize the information content in their genomes is to encode multifunctional proteins like the 3C protease. One way to rationalize the multiple functions of 3C is to envision an ensemble of conformations, whose members are responsible for different functions (Boehr, 2012). For example, different conformations may be responsible for interacting with different RNA or protein binding partners; similar scenarios have been invoked for ubiquitin (Lange et al., 2008) and other proteins (Boehr et al., 2009) to help explain how these proteins interact with such divergent binding partners. Here, we have shown that 3C fluctuates into different conformations across multiple timescales, and binding partner interactions alter the thermally accessible conformations to regulate 3C function.

NMR chemical shift perturbation analyses indicated that RNA/peptide binding leads to structural dynamics changes at both the predicted binding site and around the binding site of the other ligand for 3Cinactive (Figure 2). Some residues were associated with multiple resonances when RNA/peptide was bound, suggesting conformational exchange on the slow NMR timescale (Figure 3). Intriguingly, many of these residues had different chemical shift positions depending on the order of RNA/peptide binding (Figure 3). RNA binding also induced changes in the ps-ns (Figure S4) and ms-ms timescale dynamics of 3Cinactive (Figure 4). With RNA bound, residues around the protease active site fluctuated into a different conformation, as assessed by the R2 relaxation dispersion studies (Figure 4). The higher energy conformation might be important for interacting with protein substrates, according to a conformational selection mechanism (Boehr et al., 2009; Kumar et al., 2000; Miller and Dill, 1997; Tsai et al., 1999). These results suggest that binding of RNA (peptide) selects a different subset of conformations, which then impacts how peptide (RNA) will subsequently interact with 3C (Figure 6). In fact, peptide binding influences RNA affinity (Table 1) and RNA affects proteolytic activity (Table 2).

Figure 6.

Figure 6

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Conformational channeling in the 3C protein. Schematics of the conformational energy landscapes on 3C in the absence and presence of peptide and RNA ligands. Importantly, the ligand binding order determines the lowest energy conformation in the ternary complex, as also suggested by the chemical shift perturbations in Figure 3. This channeling of the free energy landscape may also affect how 3C interacts with other viral and host factors.

Binding of peptide/RNA thus appears to change the structural dynamics of 3C, which can then regulate how 3C interacts with other binding partners. In this context, it was especially interesting that the AB peptide had a more substantial effect on RNA binding than the CA peptide (Table 1). These differences may be related to the different functions of the P2 and P3 proteins within the polyprotein (Figure S1). The AB peptide is representative of the cleavage sites within the P2 domain of the polyprotein. The P2 proteins are important for host protein interactions (2A) and generation of vesicles and replication organelles (2B and 2BC) (Gradi et al., 1998; Rust et al., 2001; Trahey et al., 2012; Ventoso et al., 1998). The CA peptide is representative of the cleavage site that releases P3 proteins from P2 proteins; P3 proteins are most notable for their roles in genome replication and post-translational processing (Cameron et al., 2010). These findings suggest that many of the P2 and P3 protein functions are separated temporally, and so, their proteolytic release by 3C(D) may be regulated differentially by RNA interactions.

Changes to which conformations are thermally accessible may also impact interactions with other viral or host proteins. Many of the residues that show differences in the picosecond-nanosecond (Figure S4) and microsecond-millisecond (Figure 4, S3) dynamics upon binding RNA have been implicated as being important for peptide binding and protease activity (Blair et al., 1996; Mosimann et al., 1997). Other residues are important in forming higher order complexes with other viral proteins. For example, a structural model involving a 3C dimer and a 3D monomer binding to oriI RNA has been proposed to be important for initiation of RNA synthesis (Shen et al., 2008). Many of the 3C residues that form intersubunit interactions within this complex are conformationally dynamic on the millisecond timescale, as shown either by the R2 relaxation dispersion experiments (Figure 4) or by the presence of multiple resonances in the ternary complexes (Figure 3), including residues important for 3C-3D interactions such as Phe25, Lys108 and Asn165 and residues important for 3C-3C interactions such as Lys60, Ala61 and Asn69. These results suggest that conformational fluctuations on the millisecond timescale may be preparing 3C to form a dimer, as it does in the X-ray crystal structure, or interact with 3D and potentially other binding partners.

It should also be kept in mind that the 3C protein can be found as part of the 3BCD and 3CD polyproteins. The 3BC(D) protein may be involved in initiation of genome replication; 3CD is thought to be the major protease and the major CRE-binding protein (Cameron et al., 2010). In fact, 3CD generally has higher proteolytic activity and different protease specificity compared to 3C (Parsley et al., 1999; Ypma-Wong et al., 1988). However, the reasons for these differences are unclear. The crystal structure of 3CD appears simply to be a composite of the 3C and 3D domains with no additional protein interactions that may help to explain these functional differences (Marcotte et al., 2007). The covalent attachment of the 3D domain to the C-terminus of 3C may alter the internal motions of 3C to influence RNA binding ability and protease activity. The ability to fit the R2 relaxation dispersion data to global kinetic and thermodynamic parameters suggests that the millisecond motions in 3C are concerted, and so changes in the N- and C-termini would likely lead to dynamic changes elsewhere in the protein, especially at the active site and RNA binding site. It has been previously proposed that dynamic fluctuations may be responsible for functional differences between 3C and 3CD (Cameron et al., 2009). Dynamic regions in 3C may be involved in other biomolecular interactions; targeted studies of these dynamic regions may reveal novel biomolecular interactions important in regulating the multifunctional roles of 3C in viral infection.

Experimental Procedures

Protein overexpression and purification

Following transformation of Escherichia coli BL21(DE3) pCG1 cells with plasmids encoding 3Cactive or 3Cinactive (i.e. C147A/C153S), cells were overexpressed using autoinducible minimal media as described previously (Studier, 2005). Protein purification was carried out as previously described (Amero et al., 2008). Protein concentration was measured by a NanoDrop spectrophotometer (Thermo Scientific) or SpectroMax M2 spectrophotometer (Molecular Devices) using the theoretical extinction coefficient at 280 nm of 8940 M-1cm-1. More detailed information about protein overexpression and purification can be found in the Supplemental Experimental Procedures.

Dynamic light scattering experiments

Dynamic light scattering experiments were performed with a Viscotek 802 Instrument (Malvern Instruments). Data was analyzed using the mass model on OmniSIZE 3.0 (Malvern Instruments).

NMR analyses

1H-15N HSQC NMR spectra were recorded at 25°C on either a 600 MHz or 850 MHz Bruker Avance III spectrometer equipped with a 5 mm “inverse detection” triple resonance (1H/13C/15N) single axis gradient TCI cryoprobe. 15N R2 relaxation dispersion datasets were obtained simultaneously on both 600 MHz and 850 MHz Bruker Avance III spectrometers equipped with TCI cryoprobes. 15N R2 relaxation dispersion experiments were performed using the pulse sequences as described previously (Loria et al., 1999). Parameter fittings were performed using the GLOVE program (Sugase et al., 2013). More detailed information can be found in the Supplemental Experimental Procedures.

Pulsed-field gradient experiments were performed with the 850 MHz Bruker instrument equipped with a diffusion probe using pulse sequences described previously (Price, 1997). NMR titration and diffusion experiments were performed in 10 mM HEPES pH 7.5, 50 mM NaCl at 298 K. R2 relaxation dispersion experiments were performed in 25 mM potassium phosphate pH 8.0 and 150 mM NaCl at 293 K.

Fluorescence-based assays

Fluorescence polarization experiments were performed at 25°C on a Beacon 2000 fluorescence polarization analyzer (Life Technologies). Up to 20 μM of 3Cinactive was incubated at 25°C for 30 seconds with 0.2 nM 3′-fluorescein-labeled SL or oriI RNA. The buffer used was 10 mM HEPES pH 7.5, 10 mM NaCl. The fluorescence protease assay for 3Cactive was previously described (Hata et al., 2000). Assays were performed at 30°C on a SpectraMax M2 microplate reader using a 96-well microtitre plate. The buffer used for the assays consisted of 10 mM HEPES pH 7.5, 50 mM NaCl. The volume of the assays was 200 μL. Peptide was dissolved in dimethylsulfoxide (DMSO) immediately before use; the DMSO concentration made up no more than 5%(v/v) of the final sample mixture. Peptide concentration was determined by weight and volume. Data was obtained with excitation at 328 nm and emission at 393nm. More details can be found in the Supplemental Experimental Procedures.

MD Simulations

MD simulations were performed similar to that described previously (Moustafa et al., 2011) using the AMBER12 package (Case et al., 2005; Roe and Cheatham, 2013) with amber 99SB force field (Hornak et al., 2006). The 3C monomer (Chain A) of the crystal structure 1L1N was used as the initial coordinates for the simulation. The 3C molecule was immersed in a truncated octahedron solvent box filled with TIP3P water molecules; the distance between any protein atom and the edge of the solvent box was kept at 20 Å. The charge of the simulated structure was neutralized by adding the appropriate number of counter ions. The all-atom NPT simulation was carried out at 300 K and the Berndsen thermostat was used to control the temperature during simulation. The total simulation time was 100 ns using a 1 fs integration time-step; any bonds involving H-atoms were constrained using the Shake algorithm. The non-bonded interactions were calculated under periodic boundary conditions using a cutoff distance of 9 Å; the electrostatic interactions were calculated using Particle Mesh Ewald (PME) method. Analysis of the MD trajectory was done as described previously (Moustafa et al., 2011), utilizing the last 50 ns of the trajectory using Ptraj and Cpptraj of the AMBER package (Roe and Cheatham, 2013).

Supplementary Material

supplement

Highlights.

  • - binding to a viral protein influences structural dynamics at another binding site

  • - ligand binding also influences the function of the other binding site

  • - control of the energy landscape may regulate myriad functions of this viral protein

  • - protein structural dynamics effectively increases the genomic information content

Acknowledgments

The authors would like to thank Dr. Adriano Z. Zambom for advice with the statistical analyses, and Drs. Julia Fecko and Neela Yennawar for help with preliminary biophysical experiments. The authors would also like to thank Drs. Nihal Altan-Bonnet, Ann Cali and Peter Takvorian for discussions on viral proteins. This study was supported by grants from the US National Institutes of Health (R01AI091985 to N.A-B., A.C., P.T. and D.D.B.; R01AI053531 to C.E.C.; R01AI104878 to D.D.B.).

Footnotes

Author Contributions: Conceptualization, Y.M.C., I.M.M., C.E.C. and D.D.B.; Methodology, Y.M.C., I.M.M., J.J.A., C.E.C. and D.D.B.; Software, Y.M.C. and I.M.M.; Formal Analysis, Y.M.C., I.M.M. and D.D.B.; Investigation, Y.M.C. and I.M.M.; Data Curation, I.M.M., C.E.C. and D.D.B.; Writing – Original Draft, Y.M.C., I.M.M. and D.D.B.; Writing – Review & Editing, Y.M.C., I.M.M., J.J.A., C.E.C. and D.D.B.; Visualization, Y.M.C., I.M.M. and D.D.B.; Supervision, J.J.A., C.E.C and D.D.B.; Project Administration, C.E.C. and D.D.B.; Funding Acquisition, C.E.C. and D.D.B.

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References

  1. Amero CD, Arnold JJ, Moustafa IM, Cameron CE, Foster MP. Identification of the oriI-binding site of poliovirus 3C protein by nuclear magnetic resonance spectroscopy. J Virol. 2008;82:4363–4370. doi: 10.1128/JVI.02087-07. [DOI] [PMC free article] [PubMed] [Google Scholar]
  2. Andino R, Rieckhof GE, Achacoso PL, Baltimore D. Poliovirus RNA synthesis utilizes an RNP complex formed around the 5′-end of viral RNA. EMBO J. 1993;12:3587–3598. doi: 10.1002/j.1460-2075.1993.tb06032.x. [DOI] [PMC free article] [PubMed] [Google Scholar]
  3. Bedard KM, Semler BL. Regulation of picornavirus gene expression. Microbes Infect. 2004;6:702–713. doi: 10.1016/j.micinf.2004.03.001. [DOI] [PubMed] [Google Scholar]
  4. Blair WS, Nguyen JH, Parsley TB, Semler BL. Mutations in the poliovirus 3CD proteinase S1-specificity pocket affect substrate recognition and RNA binding. Virology. 1996;218:1–13. doi: 10.1006/viro.1996.0160. [DOI] [PubMed] [Google Scholar]
  5. Blair WS, Parsley TB, Bogerd HP, Towner JS, Semler BL, Cullen BR. Utilization of a mammalian cell-based RNA binding assay to characterize the RNA binding properties of picornavirus 3C proteinases. RNA. 1998;4:215–225. [PMC free article] [PubMed] [Google Scholar]
  6. Boehr DD. Promiscuity in protein-RNA interactions: conformational ensembles facilitate molecular recognition in the spliceosome: conformational diversity in U2AF(6)(5) facilitates binding to diverse RNA sequences. Bioessays. 2012;34:174–180. doi: 10.1002/bies.201100152. [DOI] [PubMed] [Google Scholar]
  7. Boehr DD, Nussinov R, Wright PE. The role of dynamic conformational ensembles in biomolecular recognition. Nat Chem Biol. 2009;5:789–796. doi: 10.1038/nchembio.232. [DOI] [PMC free article] [PubMed] [Google Scholar]
  8. Cameron CE, Moustafa IM, Arnold JJ. Dynamics: the missing link between structure and function of the viral RNA-dependent RNA polymerase? Curr Opin Struct Biol. 2009;19:768–774. doi: 10.1016/j.sbi.2009.10.012. [DOI] [PMC free article] [PubMed] [Google Scholar]
  9. Cameron CE, Oh HS, Moustafa IM. Expanding knowledge of P3 proteins in the poliovirus lifecycle. Future Microbiol. 2010;5:867–881. doi: 10.2217/fmb.10.40. [DOI] [PMC free article] [PubMed] [Google Scholar]
  10. Case DA, Cheatham TE, Darden T, Gohlke H, Luo R, Merz KM, Onufriev A, Simmerling C, Wang B, Woods RJ. The Amber biomolecular simulation programs. Journal of Computational Chemistry. 2005;26:1668–1688. doi: 10.1002/jcc.20290. [DOI] [PMC free article] [PubMed] [Google Scholar]
  11. Chase AJ, Daijogo S, Semler BL. Inhibition of poliovirus-induced cleavage of cellular protein PCBP2 reduces the levels of viral RNA replication. J Virol. 2014;88:3192–3201. doi: 10.1128/JVI.02503-13. [DOI] [PMC free article] [PubMed] [Google Scholar]
  12. Claridge JK, Headey SJ, Chow JY, Schwalbe M, Edwards PJ, Jeffries CM, Venugopal H, Trewhella J, Pascal SM. A picornaviral loop-to-loop replication complex. J Struct Biol. 2009;166:251–262. doi: 10.1016/j.jsb.2009.02.010. [DOI] [PMC free article] [PubMed] [Google Scholar]
  13. Daijogo S, Semler BL. Mechanistic intersections between picornavirus translation and RNA replication. Adv Virus Res. 2011;80:1–24. doi: 10.1016/B978-0-12-385987-7.00001-4. [DOI] [PubMed] [Google Scholar]
  14. de Breyne S, Bonderoff JM, Chumakov KM, Lloyd RE, Hellen CU. Cleavage of eukaryotic initiation factor eIF5B by enterovirus 3C proteases. Virology. 2008;378:118–122. doi: 10.1016/j.virol.2008.05.019. [DOI] [PMC free article] [PubMed] [Google Scholar]
  15. Franco D, Pathak HB, Cameron CE, Rombaut B, Wimmer E, Paul AV. Stimulation of poliovirus RNA synthesis and virus maturation in a HeLa cell-free in vitro translation-RNA replication system by viral protein 3CDpro. Virol J. 2005;2:86. doi: 10.1186/1743-422X-2-86. [DOI] [PMC free article] [PubMed] [Google Scholar]
  16. Gamarnik AV, Andino R. Switch from translation to RNA replication in a positive-stranded RNA virus. Genes Dev. 1998;12:2293–2304. doi: 10.1101/gad.12.15.2293. [DOI] [PMC free article] [PubMed] [Google Scholar]
  17. Gradi A, Svitkin YV, Imataka H, Sonenberg N. Proteolysis of human eukaryotic translation initiation factor eIF4GII, but not eIF4GI, coincides with the shutoff of host protein synthesis after poliovirus infection. Proc Natl Acad Sci U S A. 1998;95:11089–11094. doi: 10.1073/pnas.95.19.11089. [DOI] [PMC free article] [PubMed] [Google Scholar]
  18. Hammerle T, Molla A, Wimmer E. Mutational analysis of the proposed FG loop of poliovirus proteinase 3C identifies amino acids that are necessary for 3CD cleavage and might be determinants of a function distinct from proteolytic activity. J Virol. 1992;66:6028–6034. doi: 10.1128/jvi.66.10.6028-6034.1992. [DOI] [PMC free article] [PubMed] [Google Scholar]
  19. Hata S, Sato T, Sorimachi H, Ishiura S, Suzuki K. A simple purification and fluorescent assay method of the poliovirus 3C protease searching for specific inhibitors. J Virol Methods. 2000;84:117–126. doi: 10.1016/s0166-0934(99)00138-x. [DOI] [PubMed] [Google Scholar]
  20. Hornak V, Abel R, Okur A, Strockbine B, Roitberg A, Simmerling C. Comparison of multiple amber force fields and development of improved protein backbone parameters. Proteins-Structure Function and Bioinformatics. 2006;65:712–725. doi: 10.1002/prot.21123. [DOI] [PMC free article] [PubMed] [Google Scholar]
  21. Jore J, De Geus B, Jackson RJ, Pouwels PH, Enger-Valk BE. Poliovirus protein 3CD is the active protease for processing of the precursor protein P1 in vitro. J Gen Virol. 1988;69(Pt 7):1627–1636. doi: 10.1099/0022-1317-69-7-1627. [DOI] [PubMed] [Google Scholar]
  22. Kumar S, Ma BY, Tsai CJ, Sinha N, Nussinov R. Folding and binding cascades: Dynamic landscapes and population shifts. Protein Science. 2000;9:10–19. doi: 10.1110/ps.9.1.10. [DOI] [PMC free article] [PubMed] [Google Scholar]
  23. Kuyumcu-Martinez NM, Van Eden ME, Younan P, Lloyd RE. Cleavage of poly(A)-binding protein by poliovirus 3C protease inhibits host cell translation: a novel mechanism for host translation shutoff. Mol Cell Biol. 2004;24:1779–1790. doi: 10.1128/MCB.24.4.1779-1790.2004. [DOI] [PMC free article] [PubMed] [Google Scholar]
  24. Lange OF, Lakomek NA, Fares C, Schroder GF, Walter KFA, Becker S, Meiler J, Grubmuller H, Griesinger C, de Groot BL. Recognition dynamics up to microseconds revealed from an RDC-derived ubiquitin ensemble in solution. Science. 2008;320:1471–1475. doi: 10.1126/science.1157092. [DOI] [PubMed] [Google Scholar]
  25. Leong LE, Walker PA, Porter AG. Human rhinovirus-14 protease 3C (3Cpro) binds specifically to the 5′-noncoding region of the viral RNA. Evidence that 3Cpro has different domains for the RNA binding and proteolytic activities. J Biol Chem. 1993;268:25735–25739. [PubMed] [Google Scholar]
  26. Lipari G, Szabo A. Model-free approach to the interpretation of nuclear magnetic resonance relaxation in macromolecules. 1. Theory and range of validity. Journal of the American Chemical Society. 1982a;104:4546–4559. [Google Scholar]
  27. Lipari G, Szabo A. Model-free approach to the interpretation of nuclear magnetic resonance relaxation in macromolecules. 2. Analysis of experimental results. Journal of the American Chemical Society. 1982b;104:4559–4570. [Google Scholar]
  28. Loria JP, Berlow RB, Watt ED. Characterization of enzyme motions by solution NMR relaxation dispersion. Acc Chem Res. 2008;41:214–221. doi: 10.1021/ar700132n. [DOI] [PubMed] [Google Scholar]
  29. Loria JP, Rance M, Palmer AG., 3rd A TROSY CPMG sequence for characterizing chemical exchange in large proteins. J Biomol NMR. 1999;15:151–155. doi: 10.1023/a:1008355631073. [DOI] [PubMed] [Google Scholar]
  30. Marcotte LL, Wass AB, Gohara DW, Pathak HB, Arnold JJ, Filman DJ, Cameron CE, Hogle JM. Crystal structure of poliovirus 3CD protein: virally encoded protease and precursor to the RNA-dependent RNA polymerase. J Virol. 2007;81:3583–3596. doi: 10.1128/JVI.02306-06. [DOI] [PMC free article] [PubMed] [Google Scholar]
  31. Miller DW, Dill KA. Ligand binding to proteins: The binding landscape model. Protein Science. 1997;6:2166–2179. doi: 10.1002/pro.5560061011. [DOI] [PMC free article] [PubMed] [Google Scholar]
  32. Mosimann SC, Cherney MM, Sia S, Plotch S, James MN. Refined X-ray crystallographic structure of the poliovirus 3C gene product. J Mol Biol. 1997;273:1032–1047. doi: 10.1006/jmbi.1997.1306. [DOI] [PubMed] [Google Scholar]
  33. Moustafa IM, Shen H, Morton B, Colina CM, Cameron CE. Molecular dynamics simulations of viral RNA-dependent RNA polymerases link conserved and correlated motions of functional elements to fidelity. J Mol Biol. 2011 doi: 10.1016/j.jmb.2011.04.078. [DOI] [PMC free article] [PubMed] [Google Scholar]
  34. Nayak A, Goodfellow IG, Woolaway KE, Birtley J, Curry S, Belsham GJ. Role of RNA structure and RNA binding activity of foot-and-mouth disease virus 3C protein in VPg uridylylation and virus replication. J Virol. 2006;80:9865–9875. doi: 10.1128/JVI.00561-06. [DOI] [PMC free article] [PubMed] [Google Scholar]
  35. Parsley TB, Cornell CT, Semler BL. Modulation of the RNA binding and protein processing activities of poliovirus polypeptide 3CD by the viral RNA polymerase domain. J Biol Chem. 1999;274:12867–12876. doi: 10.1074/jbc.274.18.12867. [DOI] [PubMed] [Google Scholar]
  36. Parsley TB, Towner JS, Blyn LB, Ehrenfeld E, Semler BL. Poly (rC) binding protein 2 forms a ternary complex with the 5′-terminal sequences of poliovirus RNA and the viral 3CD proteinase. RNA. 1997;3:1124–1134. [PMC free article] [PubMed] [Google Scholar]
  37. Price WS. Pulsed-field gradient nuclear magnetic resonance as a tool for studying translational diffusion.1. Basic theory. Concepts in Magnetic Resonance. 1997;9:299–336. [Google Scholar]
  38. Roe DR, Cheatham TE. PTRAJ and CPPTRAJ: Software for Processing and Analysis of Molecular Dynamics Trajectory Data. Journal of Chemical Theory and Computation. 2013;9:3084–3095. doi: 10.1021/ct400341p. [DOI] [PubMed] [Google Scholar]
  39. Rozovics JM, Chase AJ, Cathcart AL, Chou W, Gershon PD, Palusa S, Wilusz J, Semler BL. Picornavirus modification of a host mRNA decay protein. MBio. 2012;3:e00431–00412. doi: 10.1128/mBio.00431-12. [DOI] [PMC free article] [PubMed] [Google Scholar]
  40. Rust RC, Landmann L, Gosert R, Tang BL, Hong W, Hauri HP, Egger D, Bienz K. Cellular COPII proteins are involved in production of the vesicles that form the poliovirus replication complex. J Virol. 2001;75:9808–9818. doi: 10.1128/JVI.75.20.9808-9818.2001. [DOI] [PMC free article] [PubMed] [Google Scholar]
  41. Shen M, Reitman ZJ, Zhao Y, Moustafa I, Wang Q, Arnold JJ, Pathak HB, Cameron CE. Picornavirus genome replication. Identification of the surface of the poliovirus (PV) 3C dimer that interacts with PV 3Dpol during VPg uridylylation and construction of a structural model for the PV 3C2-3Dpol complex. J Biol Chem. 2008;283:875–888. doi: 10.1074/jbc.M707907200. [DOI] [PMC free article] [PubMed] [Google Scholar]
  42. Shih SR, Chiang C, Chen TC, Wu CN, Hsu JT, Lee JC, Hwang MJ, Li ML, Chen GW, Ho MS. Mutations at KFRDI and VGK domains of enterovirus 71 3C protease affect its RNA binding and proteolytic activities. J Biomed Sci. 2004;11:239–248. doi: 10.1007/BF02256567. [DOI] [PubMed] [Google Scholar]
  43. Sola I, Mateos-Gomez PA, Almazan F, Zuniga S, Enjuanes L. RNA-RNA and RNA-protein interactions in coronavirus replication and transcription. RNA Biol. 2011;8:237–248. doi: 10.4161/rna.8.2.14991. [DOI] [PMC free article] [PubMed] [Google Scholar]
  44. Steil BP, Barton DJ. Cis-active RNA elements (CREs) and picornavirus RNA replication. Virus Res. 2009;139:240–252. doi: 10.1016/j.virusres.2008.07.027. [DOI] [PMC free article] [PubMed] [Google Scholar]
  45. Studier FW. Protein production by auto-induction in high density shaking cultures. Protein Expr Purif. 2005;41:207–234. doi: 10.1016/j.pep.2005.01.016. [DOI] [PubMed] [Google Scholar]
  46. Sugase K, Konuma T, Lansing JC, Wright PE. Fast and accurate fitting of relaxation dispersion data using the flexible software package GLOVE. J Biomol NMR. 2013;56:275–283. doi: 10.1007/s10858-013-9747-5. [DOI] [PMC free article] [PubMed] [Google Scholar]
  47. Takahashi D, Kim Y, Lovell S, Prakash U, Groutas WC, Chang KO. Structural and inhibitor studies of norovirus 3C-like proteases. Virus Research. 2013;178:437–444. doi: 10.1016/j.virusres.2013.09.008. [DOI] [PMC free article] [PubMed] [Google Scholar]
  48. Trahey M, Oh HS, Cameron CE, Hay JC. Poliovirus infection transiently increases COPII vesicle budding. J Virol. 2012;86:9675–9682. doi: 10.1128/JVI.01159-12. [DOI] [PMC free article] [PubMed] [Google Scholar]
  49. Tsai CJ, Ma BY, Nussinov R. Folding and binding cascades: Shifts in energy landscapes. Proceedings of the National Academy of Sciences of the United States of America. 1999;96:9970–9972. doi: 10.1073/pnas.96.18.9970. [DOI] [PMC free article] [PubMed] [Google Scholar]
  50. Ventoso I, MacMillan SE, Hershey JW, Carrasco L. Poliovirus 2A proteinase cleaves directly the eIF-4G subunit of eIF-4F complex. FEBS Lett. 1998;435:79–83. doi: 10.1016/s0014-5793(98)01027-8. [DOI] [PubMed] [Google Scholar]
  51. Viswanathan P, May J, Uhm S, Yon C, Korba B. RNA binding by human Norovirus 3C-like proteases inhibits protease activity. Virology. 2013;438:20–27. doi: 10.1016/j.virol.2013.01.006. [DOI] [PubMed] [Google Scholar]
  52. Walker EJ, Younessi P, Fulcher AJ, McCuaig R, Thomas BJ, Bardin PG, Jans DA, Ghildyal R. Rhinovirus 3C protease facilitates specific nucleoporin cleavage and mislocalisation of nuclear proteins in infected host cells. PLoS One. 2013;8:e71316. doi: 10.1371/journal.pone.0071316. [DOI] [PMC free article] [PubMed] [Google Scholar]
  53. Walker PA, Leong LE, Porter AG. Sequence and structural determinants of the interaction between the 5′-noncoding region of picornavirus RNA and rhinovirus protease 3C. J Biol Chem. 1995;270:14510–14516. doi: 10.1074/jbc.270.24.14510. [DOI] [PubMed] [Google Scholar]
  54. Yalamanchili P, Weidman K, Dasgupta A. Cleavage of transcriptional activator Oct-1 by poliovirus encoded protease 3Cpro. Virology. 1997;239:176–185. doi: 10.1006/viro.1997.8862. [DOI] [PubMed] [Google Scholar]
  55. Yang Y, Rijnbrand R, Watowich S, Lemon SM. Genetic evidence for an interaction between a picornaviral cis-acting RNA replication element and 3CD protein. J Biol Chem. 2004;279:12659–12667. doi: 10.1074/jbc.M312992200. [DOI] [PubMed] [Google Scholar]
  56. Yin J, Liu Y, Wimmer E, Paul AV. Complete protein linkage map between the P2 and P3 non-structural proteins of poliovirus. J Gen Virol. 2007;88:2259–2267. doi: 10.1099/vir.0.82795-0. [DOI] [PubMed] [Google Scholar]
  57. Ypma-Wong MF, Dewalt PG, Johnson VH, Lamb JG, Semler BL. Protein 3CD is the major poliovirus proteinase responsible for cleavage of the P1 capsid precursor. Virology. 1988;166:265–270. doi: 10.1016/0042-6822(88)90172-9. [DOI] [PubMed] [Google Scholar]
  58. Yu L, Takeda K, Markoff L. Protein-protein interactions among West Nile non-structural proteins and transmembrane complex formation in mammalian cells. Virology. 2013;446:365–377. doi: 10.1016/j.virol.2013.08.006. [DOI] [PubMed] [Google Scholar]
  59. Zeitler CE, Estes MK, Prasad BVV. X-ray crystallographic structure of the Norwalk virus protease at 1.5-angstrom resolution. Journal of Virology. 2006;80:5050–5058. doi: 10.1128/JVI.80.10.5050-5058.2006. [DOI] [PMC free article] [PubMed] [Google Scholar]
  60. Zhou Z, Mogensen MM, Powell PP, Curry S, Wileman T. Foot-and-mouth disease virus 3C protease induces fragmentation of the Golgi compartment and blocks intra-Golgi transport. J Virol. 2013;87:11721–11729. doi: 10.1128/JVI.01355-13. [DOI] [PMC free article] [PubMed] [Google Scholar]

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