The intergenic IRES of Cricket Paralysis Virus (CrPV-IRES) forms a tight complex with 80S ribosomes capable of initiating the cell-free synthesis of complete proteins in the absence of initiation factors. Such synthesis raises the question of what effect the necessary IRES dissociation from the tRNA binding sites, and ultimately from all of the ribosome, has on the rates of initial peptide elongation steps as nascent peptide is formed. Here we report the first results measuring rates of reaction for the initial cycles of IRES-dependent elongation. Our results demonstrate that 1) the first two cycles of elongation proceed much more slowly than subsequent cycles, 2) these reduced rates arise from slow pseudo-translocation and translocation steps, and 3) the retarding effect of ribosome-bound IRES on protein synthesis is largely overcome following translocation of tripeptidyl-tRNA. Our results also provide a straightforward approach to detailed mechanistic characterization of many aspects of eukaryotic polypeptide elongation.https://doi.org/10.7554/eLife.13429.001
Inside cells, machines called ribosomes make proteins using instructions carried by molecules of messenger RNA (or mRNA). The ribosomes bind to the mRNA and then move along it to assemble the proteins in a process called translation. The first step of translation – when the ribosome binds to the mRNA – is known as initiation. In human and other eukaryotic cells, initiation mainly occurs through a mechanism that requires many proteins called initiation factors to recruit the ribosome to a cap structure formed at one end of the mRNA.
When viruses infect cells, they hijack the ribosomes of the host cell to produce large quantities of viral proteins. However, unlike their host cells, many viruses use a different pathway to initiate translation of their mRNAs. The mRNAs of these viruses have regions known an internal ribosome entry sites (IRESs) that host cell ribosomes can bind to instead.
After initiation, the ribosome progressively assembles the building blocks of proteins (amino acids) into a chain until the new protein is complete. Molecules called transfer RNAs bind to individual amino acids and bring them to the ribosome. Previous research has shown that, prior to initiation, IRESs on Cricket Paralysis Virus mRNAs bind to the ribosome and occupy sites where transfer RNAs would normally bind. However, it was not clear how this affects the elongation process. Zhang et al. now address this question using a cell-free system that allowed them to recreate and observe translation outside of the normal cell environment.
Zhang et al. found that the binding of an IRES to a ribosome slows down the early steps of elongation. A likely explanation for this is that the IRES elements have to be displaced from the ribosome before the incoming transfer RNAs can occupy the three tRNA sites. However, as elongation progresses, the effects of the IRES elements are overcome and the pace of elongation increases significantly. Zhang et al.’s findings provide a convenient approach that could be used for future studies of elongation. This approach could also help researchers find out how abnormalities in translation contribute to human diseases, including muscle-wasting disorders.https://doi.org/10.7554/eLife.13429.002
Initiation of protein synthesis in eukaryotic cells proceeds via two well-established pathways. The cap-dependent pathway involves recognition of 7-methyl-guanosine at the 5’-terminus of mRNA by a preinitiation complex of 40S ribosomal subunit and a host of initiation factors prior to a scanning step that results in initiator aminoacyl-tRNA(aa-tRNA) pairing with a cognate start codon, followed by 60S binding to form the 80S initiator complex (Jackson et al., 2010; Aitken and Lorsch, 2012). The second pathway involves binding of the ribosome to an internal ribosome entry site (IRES), a structure that is present in many virus-encoded mRNAs, as well as in some cellular mRNAs (Fitzgerald and Semmler, 2009). Initiation of protein synthesis from an 80S·IRES complex can take place in the absence of some or even all of the initiation factors required in the cap-dependent pathway (Filbin and Kieft, 2009), depending on the IRES source. The intergenic IRES of Cricket Paralysis Virus (CrPV-IRES) forms a complex with 80S ribosomes that is capable of initiating the synthesis of complete proteins in cell-free assays completely lacking initiation factors (Jan et al. 2003; Pestova and Hillen, 2003). More recently, high resolution structural studieshave shown that, prior to polypeptide chain initiation, the closely related Dicistroviridae IRES structures from CrPV (Fernandez et al., 2014; Muhs et al., 2015) and Taura syndrome virus (Koh et al., 2014) occupy all three tRNA binding sites (E, P, and A) on the ribosome, with the protein coding region beginning immediately downstream from IRES segment occupying the A-site (Figure 1).
CrPV-IRES binds with high affinity (Kd ~ 10 nM) to the 80S ribosome (Jang and Jan, 2010), raising the question of what effect the necessary IRES dissociation from the tRNA binding sites, and ultimately from all of the ribosome as well, has on the rates of initial peptide elongation steps as nascent peptide is formed (Muhs et al., 2015). Since prior to the work reported in this paper nothing had been published concerning the rate of initial oligopeptide synthesis by an 80S·CrPV-IRES complex, it has been unclear whether there is a retarding effect due to the presence of IRES on the ribosome, and, if so, how many cycles of peptide elongation are required before the ribosome begins to form peptide bonds at a higher rate. In considering this question, we make use of the simplified 12-step scheme of initial tetrapeptide synthesis shown in Figure 2, which provides a useful framework for presenting the results described in this paper. In this scheme Steps 1–3 show the reactions required for initial binding of the first tRNA to the A site followed by translocation to the P-site, and reactions 4–6, 7–9, and 10–12 represent three elongation cycles, ending with P-site bound tetrapeptidyl-tRNA, completing the third cycle of polypeptide synthesis. This model makes the reasonable assumption that binding of successive aminoacyl-tRNAs (aa-tRNAs) cognate to the mRNA requires the progressive removal of IRES structures from each of the tRNA binding sites, such that translocation of dipeptidyl tRNA to the P-site (structure 7) requires removal of the IRES from the last of the three tRNA binding sites. In the work reported below, we demonstrate first, that the initial elongation steps are indeed quite slow and are limited by the translocation step of the elongation cycle, and second, that the rate of elongation accelerates following translocation of tripeptidyl-tRNA to the P-site.
In our experiments, eukaryotic ribosomes are prepared from shrimp cysts (Iwasaki and Kaziro, 1979), elongation factors are prepared from yeast, and charged tRNAs are prepared from yeast and E. coli. In addition, the peptide coding sequence attached to the 3’-end of the CrPV-IRES (Figure 1) has been mutated for ease in detection of peptide synthesis via 35S-Met incorporation. In all such mutants the initial wt-codon triplet GCU encoding Ala has been replaced by UUC, encoding Phe, a change that has little effect on the expression of active luciferase in a cell-free protein synthesis assay (Figure 1—figure supplement 1). The initial coding sequences of the mutants used in this work are presented in Supplementary file 1. Collectively, they allow monitoring of the rates of PheMet, PheLysMet, PheValLysMet and PheLyValArgGlnTrpLeuMet synthesis. In presenting the results below, Steps 1–12 and structures 1 – 13 are as described in the scheme for initial tetrapeptide synthesis proposed in Figure 2. Values of t1/2 for Steps 1–12, determined as described, are summarized in Table 2.
We previously have utilized two assays to measure binding of the ternary complex Phe-tRNAPhe·eEF1A·GTP (Phe-TC) to the 80S·CrPV-IRES (80S·IRES) complex (Ruehle et al., 2015). The increase in proflavin-labeled Phe-tRNAPhe fluorescence anisotropy measures binding to either the A- or P-site (structures 3 and 4, respectively, Figure 2). [3H]-Phe-tRNAPhe cosedimentation with the 80S·IRES complex measures accumulation of 4 only, since A-site binding is too labile to survive the ultracentrifugation step (Yamamoto et al., 2007).
In Figure 3 we present time-resolved application of the anisotropy assay that allows us to measure the rates of Phe-TC binding to form Structure 3 from 1. These resultswere fit to the scheme shown in Figure 2, giving values for k1, k-1, and k2 in both the presence and absence of eEF2·GTP that are summarized in Table 1. In the absence of eEF2 (blue trace), the equilibrium position of Step 1, a so-called pseudo-translocation step (Muhs et al., 2015) in which the IRES vacates the A-site, favors Structure 1 over Structure 2 by approximately 20-fold, consistent with recent structural studies (Fernandez et al., 2014; Koh et al., 2014; Muhs et al., 2015). Phe-TC binds to Structure 2 yielding Structure 3, in a process where the rate-limiting step is the conversion of Structure 1 to Structure 2. Preincubation of 80S·IRES complex with 1 µM or 3 µM eEF2·GTP leads to clear biphasic binding of Phe-TC, with the more rapid and slower phases each accounting for ~50% of binding, respectively (red and black traces). These results indicate that, consistent with recent results of Petrov et al. (2016), the equilibrium between Structures 1 and 2 is shifted in the presence of eEF2·GTP, such that approximately half of 80S·IRES is present as 2.Phe-TC binding to 2, resulting in the formation of Structure 3, accounts for the rapid phase in the red and black traces. Further formation of 3 is limited by the slower rate of 1 to 2 conversion. Although added eEF2·GTP decreases all three apparent rate constants, the effect is much greater on k-1 (~50-fold reduction) than on either k1 (~twofold reduction) or k2 (~fourfold reduction). The near identity of the red and black traces, performed at different eEF2·GTP concentrations, suggests that this factor interacts with both 1 and 2, with a dissociation constant significantly less than 1 µM. The large inhibitory effect of eEF2·GTP on k-1 is consistent with its role as a translocase, and with recent results demonstrating that a principal role of EF-G, the prokaryotic equivalent of eEF2, is to inhibit back-translocation (Adio et al., 2015). eEF2·GTP inhibition of k2 may be due, at least in part, to a requirement for eEF2·GDP dissociation prior to Phe-TC binding.
Formation of Structure 4 from Structure 1, as measured by the co-sedimentation assay, requires the presence of eEF2·GTP and proceeds at a considerably slower rate than formation of Structure 3 from Structure 1 (Figure 3), allowing estimation of a t1/2 for Step 3, a second pseudo-translocation step involving conversion of 3 to 4, of 210 ± 10 s. It is this further slow step that accounts for the lack of significant effect of preincubation with eEF2·GTP (5’ or 60’) on the rate of formation of 4 from 1 (Figure 3).
Using ribosomes programmed with the appropriate coding sequence mutants (Supplementary file 1) and [35S]-Met-TC, we employ a rapid mixing and quench assay to measure rates of PheMet, PheLysMet, and PheLysValMet synthesis, with detection and quantification of product by thin layer electrophoresis (TLE) (Figure 4A and Figure 4—figure supplement 1). For PheMet synthesis (Figure 4B) we preform Structure 4 and measure its conversion to Structure 6. We measurePheLysMet synthesis, Structure 9, starting from either Structure 4 or Structure 7 (Figure 4C) and PheValLysMet synthesis, Structure 12, starting from either Structure 7 or Structure 10 (Figure 4D). In all three cases, reactions involving only TC binding and a single peptide bond formation (4 to 6; 7 to 9; 10 to 12) proceed in remarkably similar fashion, each showing biphasic behavior with a rapid phase accounting for 65 ± 10% of reaction proceeding with a t1/2 of ~6–9 s and a slower, minor phase proceeding much more slowly (t1/2 ~220–240 s), possibly corresponding to defective ribosomes. Reactions involving formation of two peptide bonds, as in the conversion of 4 to 9 or 7 to 12 are well approximated as single phase reactions with t1/2 values of 90–110 s. Conversion of 4 to 9 proceeds via Steps 4 – 8, allowing the t1/2 value for the translocation Step 6 to be estimated as 84 s, from the difference between the t1/2 value for the 4 to 9 reaction and the sum of the t1/2 values for the 4 to 6 and 7 to 9 reactions (major phases). Similarly, the t1/2 value for the translocation Step 9 can be estimated as 110 s from the difference between the t1/2 value for the 7 to 12 reaction and the sum of the t1/2 values for the 7 to 9 and 10 to 12 reactions. Since the di-, tri- and tetrapeptides synthesized in the results reported in Figure 4 use different coding sequence mutants, these estimates of translocation t1/2 values depend on the not unreasonable assumption that the identities of the tRNAs undergoing translocation do not have a major influence on the translocation rate. With this caveat, the results presented in Figure 4 lead to the clear conclusion that translocation is the rate limiting step in each of the first two cycles of polypeptide elongation, proceeding from 4 to 10.
In an attempt to resolve the TC binding step (reactions 4, 7, and 10) from the peptide formation step (reactions 5, 8, and 11) we also employed a rapid mixing and quench assay to determine the rates with which [35S]-Met-tRNAMet is able to cosediment with the ribosome following mixing of [35S]-Met-TC with structures 4, 7, or 10. This strategy was successful for [35S]-Met–TC reaction with structure 7 (containing P-site bound PheLys-tRNALys, Figure 4C) or structure 10 (containing P-site bound PheValLys-tRNALys Figure 4D), in which the [35S]-Met-TC cosedimentation rates outpace the rates of peptide bond formation with Met-TC. These rate differentials permit estimates to be made for the t1/2 values of TC binding (Step 7, 3 s; Step 10, 2 s) and peptide bond formation (Step 8, 4 s; Step 11, 7 s). They also provide a clear indication that, within Structures 8, 9, 11 and 12, Met-tRNAMet, PheLysMet-tRNAMet, and PheValLysMet- tRNAMet, whenbound to the A-site, efficiently cosediment with ribosomes, which is typical for A-site bound tRNAs in conventional (non-IRES) elongation complexes (Warner and Rich, 1964; Nwagwu, 1975).
However, for [35S]-Met–TC reaction with structure 4 (containing P-site bound Phe-tRNAPhe), the [35S]-Met-TC cosedimentation rate is much slower than the dipeptide formation rate (Figure 4B). This indicates that PheMet-tRNAMet, and possibly Met-tRNAMet as well, are not bound stably to the ribosome in Structures 5 and 6, and that only PheMet-tRNAMet bound to the P-site (Structure 7) is fully recovered by cosedimentation. As a result, the cosedimentation assay does not permit estimation of the t1/2 values for Steps 4 and 5. It is possible that the lability of the A-site tRNAs in structures 5 and 6 is due to IRES binding to the E-site, which is absent in structures 8, 9 and 11, 12, and may reflect an allosteric A-site: E-site interaction. Evidence for allosteric A-site/E-site interactions has been presented for both bacterial and eukaryotic ribosomes (Nierhaus 1990; Chen et al., 2011; Ferguson et al., 2015), although the general validity of this interaction has been questioned (Semenkov et al., 1996; Petropoulos and Green, 2012).
The results presented in Figure 4 show that translocation proceeds slowly through the first two elongation cycles of nascent protein synthesis, raising the question of how far nascent protein synthesis has to proceed to overcome the retarding effect of ribosome-bound IRES. In Figure 5 we present the results of two experimental approaches demonstrating that translocation of tetrapeptidyl-tRNA proceeds much more rapidly than translocation of tripeptidyl-tRNA.
The first approach makes use of the fact that formation of peptidyl-puromycin proceeds more rapidly with peptidyl-tRNA bound to the P-site than to the A-site, permitting puromycin reactivity to distinguish A-site from P-site peptidyl-tRNA. As shown in Figure 5A, puromycin (1 mM) reacts with A-site bound PheValLys-tRNALys, Structure 9, about 20times more slowly (t1/2 1400 ± 300 s) than it reacts with P-site bound PheValLys-tRNALys(t1/2 76 ± 16 s). The corresponding t1/2 value for puromycin reaction with PheValLys-tRNALys undergoing translocation from the A- to P-site is 170 ± 30 s. This increase of approximately 100 s for translocating PheValLys-tRNALysvs. translocated PheValLys-tRNALys closely matches the t1/2 value of 110 ± 30 s estimated above for the translocation of tripeptidyl-tRNA (Table 2) and can be assigned to the translocating step. In contrast, the rates of puromycin reaction with translocating and translocated PheValLysMet-tRNAMet (Structure 13)are indistinguishable from one another (t1/2 values of 37 ± 4 s and 46 ± 7 s, respectively, Figure 5B), a clear demonstration that translocation of PheValLysMet-tRNAMet proceeds rapidly with respect to puromycin reaction. Our results allow us to estimate an upper limit value of t1/2 for the translocation Step 12 of ≤10 s.
Puromycin reacts at similar rates with translocated PheValLys-tRNALys (Structure 10, t1/2 76 ± 16 s) and PheValLysMet-tRNAMet (Structure 13, t1/2 46 ± 7 s). These rates, while consistent with those reported by others for puromycin reaction with eukaryotic P-site bound Met-tRNAMet (Lorsch and Herschlag, 1999), N-AcPhe-tRNAPhe (Ioannou et al., 1997), and Cy3-Met-tRNAMet (Ferguson et al., 2015), are several hundred-fold slower than those measured for puromycin reaction with prokaryotic P-site bound peptidyl- or fMet-tRNA. This largely explains why the rate reduction for puromycin reaction with A-site vs. P-site bound peptidyl-tRNA is so much more modest for eukaryotic ribosomes (~20-fold, Figure 5A) than for prokaryotic ribosomes (103–104-fold, Pan et al., 2007; Semenkov et al., 1992 ; Sharma et al., 2004; Peske et al., 2004 ).
Above we have demonstrated that, under our conditions, aa-tRNA binding and peptide bond formation proceed with an overall t1/2of 6 – 9 s for each of the three elongation steps we have studied. This relative constancy, coupled with the much slower translocation of tripeptidyl-tRNA (Step 9) vs. tetrapeptidyl-tRNA (Step 12), leads to the prediction that synthesis of a longer peptide that required the tripeptidyl-tRNA translocation step (Step 9) would proceed significantly more slowly than synthesis not requiring this step.
In the second approach we verified this prediction by demonstrating that octapeptide FKVRQWLM formation, as measured by the cosedimentation assay, is much slower when synthesis is initiated with P-site bound PheLys-tRNALys (Structure 7) vs. P-site bound PheLysVal-tRNAVal (Structure 10) (Figure 5C). Indeed, the rates of FKVRQWLM synthesis are only marginally increased when reaction is initiated with P-site bound tetrapeptidyl-tRNA or pentapeptidyl-tRNA as compared with tripeptidyl-tRNA, reinforcing the notion that the retarding effect of ribosome-bound IRES on protein synthesis is largely overcome following translocation of tripeptidyl-tRNA.
The results presented in this paper constitute the first time that rates of reaction have been determined for the initial cycles of IRES-dependent elongation. They demonstrate quite clearly that the first two cycles of elongation proceed much more slowly than subsequent steps, and that these reduced rates arise from slow, rate-determining, pseudo-translocation and translocation steps. Translocation during the first elongation cycle (Step 6) clearly requires displacement of the IRES from the E-site, so it is not unexpected that it would be slow. Less predictable is the slow translocation in the second elongation cycle, (Step 9) after the IRES structure has, presumably, already left the E-site (Figure 1). The slow rate of Step 9 might be due to a full dissociation of IRES from the ribosome during this step, a suggestion that could be tested by appropriately designed structural studies. In any case, our results do clearly demonstrate that, following translocation of tripeptidyl-tRNA from the A- to P-site, the pace of nascent peptide chain elongation picks up dramatically. Further work, comparing quantitatively the rates of successive cycles of nascent peptide elongation following tetrapeptide formation (i.e, cycles 4, 5, 6, 7, etc.) will be required to determine how many cycles are required before any retarding influence of bound CrPV-IRES is completely eliminated.
Our results also clarify an aspect of the initial binding of the first aa-tRNA to the 80S·CrPV-IRES complex. Prior results have shown that initial aa-tRNA binding, in the form of a ternary complex, to an 80S·IRES complex, as measured either by cosedimentation (Fernandez et al., 2014), or by filter binding and toeprinting (Yamamoto et al., 2007), requires eEF2·GTP, leading to the conclusion that initial aa-tRNA binding can only bind to the 80S·IRES complex after an eEF2-dependent translocation event (Fernandez et al., 2014). While we agree with the experimental results, and have in fact reproduced the cosedimentation result in our own work, we disagree with the conclusion. This is because these earlier experiments only measured stable aa-tRNA binding, corresponding to formation of Structure 4 in which aa-tRNA binds to the P-site. However, it is clear from the anisotropy experiment conducted in the absence of added eEF2·GTP (Figure 3, blue trace) that ternary complex binding measured in situ, which can monitor labile binding to the A-site (Structure 3)does not require eEF2·GTP. This is easily understood as an example of Le Chaltelier’s principle, in which the equilibrium between Structure 1 (closed A- site)and Structure 2 (open A- site), which strongly favors Structure 1, is pulled to the right by aa-tRNA binding. Preincubation with eEF2 also shifts the equilibrium to the right, leading to an initial rapid phase of reaction with Phe-TC (Figure 3).
This latter shift, for which the results presented in Figure 3 provide strong inferential evidence, appears to be at odds with earlier toeprinting results showing no shift in IRES position within the 80S·CrPV-IRES complex on addition of eEF2 alone (Pestova et al., 2003; Jan et al., 2003). In agreement with the suggestion of Muhs et al. (2015), we believe it likely that this apparent inconsistency arises from eEF2 dissociation from the ribosome during the toeprinting assay (Pestova et al., 1996), with the consequent favoring of Structure 1. This is because GTP is required for tight binding of eEF2 to the ribosome (Nygård and Nilsson, 1984), but the toeprinting assay is carried out for an extended period of time (45 min) under non-denaturing conditions in the absence of added GTP, conditions that would eventually deplete GTP due to ribosome-dependent eEF2·GTP hydrolysis (Nygård and Nilsson, 1989). In addition, the toeprinting assay is performed at a Mg2+ concentration of 10.5 mM, considerably higher than the 5 mM used in our kinetic studies, which could also affect the 1 to 2 equilibrium position.
How relevant are the present results for in vivo initiation of IRES-dependent protein synthesis? We note three potential concerns. First, our in vitro systemis quite heterogeneous, with ribosomes derived from shrimp cysts, yeast elongation factors, and yeast and E. coli charged tRNAs. However, as reviewed in Koh et al. (2014), IRESs can initiate translation on ribosomes from many eukaryotic organisms, including shrimp (Cevallos and Sarnow, 2005), indicating that the molecular mechanism is not species-specific. CrPV IRESs in particular can initiate translation on ribosomes from yeast (Thompson et al., 2001) to human (Spahn et al., 2004). Furthermore, eukaryotic elongation factors have structures that are very strongly conserved (Soares et al., 2009; Jørgensen et al., 2006), and there is strong evidence that charged tRNAs from one species form functional complexes with both eEF1A and ribosomes from a different species (Jackson et al., 2001; Ferguson et al., 2015). Second, the coding sequences employed in this work are different from that immediately downstream of wt-CrPV-IRES (Supplementary file 1). This is also unlikely to pose a major difficulty, given the strong evidence that mutations in the downstream sequence are, in general, tolerated without substantial effect on initiation of translation (Tsukiyama-Kohara et al., 1992; Wang et al., 1993; Hellen and Sarnow, 2001; Rijnbrand et al., 2001), although mutations of some downstream sequences do give rise to relatively minor changes in IRES activity (Kim et al., 2003; Shibuya et al., 2003; Wang et al., 2013). Third, the elongation rate of even the later cycles of IRES-dependent elongation (Figure 5C) is quite slow (~0.1 s-1). Although this rate is essentially identical to that reported for tripeptide synthesis in a cap-dependent yeast-based in vitro translation system which requires five initiation factors and eEF3 in addition to eEF1A and eEF2 (Acker et al., 2007; Eyler and Green, 2011; Gutierrez et al., 2013), it is 1.5–2 orders of magnitude slowerthan rates of peptide elongation that have been estimated for intact eukaryotic cells at 37°C (3–10 s-1) (Boehlke and Friesen, 1975; Hershey, 1991). There is evidence that, in many eukaryotic cells, the protein synthesis machinery is highly organized, containing several components, including ribosomes, a multi-aminoacyl-tRNA synthetase complex, eEF-1A, and several auxiliary proteins (Negrutskii et al., 1994; Negrutskii and El’skaya, 1998; David et al., 2011). It has been suggested that this organized structure optimizes translation rate by coordinating synthetase activities to facilitate channeling of aa-tRNAs to the elongating ribosomes. Thus, protein synthesis in a permeabilized mammalian cell, in which this structure is likely to be preserved, proceeds 40-fold faster than what is obtained in a cell-free system prepared from the same cells which presumably lacks this structure (Negrutskii et al., 1994). The slow rates measured for both the IRES-dependent and cap-dependent in vitro systems could be due, at least in part, to their lack of aa-tRNA channeling. Such channeling would be unlikely to accelerate the very slow translocation rates in the initial peptide elongation cycles reported in this work, although we cannot exclude the possibility that other proteins present in vivo might have such effects. Future efforts will address this issue. Here, incorporation of some of the features of a recently introduced in vitro protein synthesissystem in which initiation is carried out using the IRES from hepatitis C virus could be useful (Machida et al., 2014).
Detailed mechanistic characterization of many aspects of eukaryotic polypeptide elongation has been held back by the lack of a convenient system for its study. The very simple in vitro IRES-dependent elongation system described here should be useful in overcoming this limitation. As one example, it is generally assumed, based on extensive structural similarities (Jørgensen et al., 2006), that eEF2 functions in catalyzing eukaryotic elongation in much the same way that EF-G catalyzes prokaryotic elongation, but this assumption does not take into account some important structural differences, including the fact that eEF2 is subject to post-translational modifications not found in EF-G, with clear consequences for activity but, as yet, little understanding of mechanism (Dever and Green, 2012; Mittal et al., 2013 ; Greganova et al., 2011; Liu et al., 2012). The CrPV-IRES based system should permit detailed rate and structural dynamic studies of eEF2 catalytic function, of the kind that have proved so useful in elucidating EF-G function in bacterial protein synthesis (Pan et al., 2007; Chen et al., 2011; Chen et al., 2013; Holtkamp et al., 2014; Salsi et al., 2015).
The wt CrPV Phe-IRES vector, as well as several variants in which the first Ala codon is replaced by a Phe codon, were the kind gifts of Dr. Eric Jan.This replacement, which has little effect on the initiation of translation (see Discussion and Figure 1—figure supplement 1), was made as a matter of convenience, since tRNAPhe was available to us and the appropriate tRNAAla acceptor was not. The vectors encoding the PheMet, PheValMet, PheValLysMet, and PheLysValArgGlnTrpLeuMet were generated by PCR insertion of corresponding sequences (Supplementary file 1) into the CrPV Phe-IRES vector. All cloned sequences were verified by standard sequencing methods using appropriate primers.
For in vitro transcription of full-length mRNA for the Luciferase assay, the WT and mutated Phe-IRES plasmids were linearized with XbaI, which cleaves the plasmids after the firefly luciferase coding region. mRNA was transcribed in vitro using the AmpliScribe T7 transcription kit (EPICENTRE) according to the manufacturer. For in vitro transcription of short-length mRNAs, the mutated IRES plasmids were linearized with NarI, which cleaves 33 nt downstream of the ATG start codon of the luciferase coding region.
In vitro translation of firefly luciferase with WT and mutated F-IRES mRNA (1 μg in 50 μL of reaction mixture) was performed using the Flexi Rabbit Reticulocyte Lysate System (Promega) according to the manufacturer. IRES mRNA was omitted in the control reaction. Fluc activities (Figure 1—figure supplement 1) were determined using a plate reader (Envision 2103, Perkin-Elmer) to detect the luminescence signal.
Shrimp (A. salina) 80S ribosomes were prepared from dried, frozen cysts as previously described (Iwasaki and Kaziro, 1979) with some modifications. After the shrimp cysts were ground open, debris was removed by centrifugation at 30,000xg for 15 min and crude 80S ribosomes were precipitated from the supernatant by addition of 4.5% (w/v) PEG 20K (Ben-Shem et al., 2011). 40S and 60S subunits were resolved on 10–30% sucrose gradients after puromycin treatment. E. coli 30S subunits were prepared as described (Grigoriadou et al., 2007). eEF1A was purified from yeast according to published methods (Thiele et al., 1985). His6-eEF2 was isolated from an overexpressing yeast strain (TKY675) generously provided by Dr. Terri Kinzy, and purified as described (Jørgensen et al., 2002). Proflavin-labeled Phe-tRNAPhe, denoted Phe-tRNAPhe(prf), was prepared as previously described (Wintermeyer and Zachau 1974, Betteridge et al., 2007). Yeast tRNAPhe was purchased from Sigma. Other isoacceptor tRNAs were prepared from bulk tRNA (Roche) from either E. coli (tRNAGln, tRNALys, tRNAMet) or yeast (tRNAArg, tRNALeu, tRNATrp, tRNAVal) by hybridization to immobilized complementary oligoDNAs, as described (Barhoom et al., 2013 ; Liu et al., 2014). E. coli and yeast tRNAs were charged with their cognate amino acids as described (Pan et al., 2006, 2009).
All complexes were prepared in buffer 4 (40 mM Tris-HCl pH 7.5, 80 mM NH4Cl, 5 mM MgOAc2, 100 mM KOAc, 3 mM 2-mercaptoethanol) at 37°C. For the preparation of ternary complexes (TC, aa-tRNA·eEF1A·GTP) and 80S·IRES complexes containing either Phe-tRNAPhe or peptidyl-tRNA bound in the P-site, buffer 4 was supplemented with 1 mM GTP and 1 mM ATP. All TC complexes were prepared by incubating the relevant charged tRNA (1.6 μM, based on amino acid stoichiometry) with eEF1A (8 μM) for 5 min. 80S·IRES complexes were formed by incubation of shrimp 40S (0.8 µM) and 60S (1.6 µM) subunits with the appropriate IRES (2.4 µM) for 5 min. 80S·IRES complexes containing Phe-tRNAPhe or peptidyl-tRNA bound in the P-site were formed by mixing 80S·IRES complexes (0.8 µM) with 1 μM eEF2 and the appropriate TCs (1.6 µM for each) for 15–40 min. To determine radioactively labeled aa-tRNA binding stoichiometries, 40 µL samples were subjected to ultracentrifugation at 4°C (540,000xg) for 40 min through a 1.1 M sucrose cushion. Excess bacterial 30S bacterial ribosome subunits (600 pmol/15 ± 5 µL) were added as carrier to enhance pelleting and allow facile calculation of complex recovery. Control experiments carried out in the absence of IRES or of both IRES and 80S ribosomes demonstrated that only negligible amounts of labeled peptidyl-tRNA cosedimented due to binding to 30S subunits (Figure 4—figure supplement 2). The pellets were gently washed twice with buffer 4 and dissolved in 100 μL of buffer 4 for A260 determination. Ribosome recoveries typically varied between 60 and 80%.
Unless otherwise noted, all reactions were performed at 37°C in buffer 4 supplemented with 1 mM GTP. All kinetic results reported are the averages of 2–4 independent determinations, performed on different days. No systematic effort was made to carry out duplicate experiments using independently made stock reagent solutions, although this was sometimes done. Error bars in figures are shown as average deviations.
Phe-tRNAPhe(prf)·eEF1A·GTP ternary complex was rapidly mixed with 80S·FVKM-IRES complex in the presence or absence of eEF2·GTP using a KinTek stopped-flow spectrofluorometer model SF-300X. Proflavin labeled Phe-tRNAPhe was excited at 462 nm and monitored using a pair of 495 nm long-pass filters. A T-shape configuration was utilized such that instrument-specific polarizers were attached to both the excitation and the two emission light paths. In each independent measurement, 15–20 shots (rapid mixing of samples) were averaged to provide the time course of anisotropy change. The g-factor and anisotropy value were calculated using the instrument software as described (Lakowicz 1999, Ameloot et al., 2013). Experimental data were processed and analyzed by Felix software (from PTI).
80S·IRES complex (0.8 μM) with no tRNA bound (Figure 2) was rapidly mixed with [3H]-Phe-TC in the presence of eEF2·GTP in a KinTek Corporation RQF-3 Rapid Quench-Flow Instrument the reaction mixture was quenched at various times with 0.5 M MES buffer (pH 6.0), and the stoichiometry of ribosome-bound [3H]-Phe-TC was determined by ultracentrifugation as described above for complex characterization. Similar procedures were used to determine the kinetics of [35S]-Met-TC binding to 80S·IRES complexes containing Phe-tRNAPhe or peptidyl-tRNA in the P-site (Figure 4).
80S·IRES complexes containing either Phe-tRNAPhe or the appropriate peptidyl-tRNA in the P-site, prepared using the standard procedure (see Complex preparations above), were rapidly mixed with [35S]-Met-TC (1.6 µM) and additional TCs as required (all 1.6 µM) in a KinTek Corporation RQF-3 Rapid Quench-Flow Instrument, and the reaction mixture was quenched at various times with 0.8 M KOH. [35S]-Met-containing peptide was released from tRNAMet by further incubation at 37°C for 3 hr. The pH of the samples were adjusted with acetic acid to pH 2.8, lyophilized, suspended in water, and centrifuged to remove particulates, which contained no 35S. The supernatant was analyzed by thin layer electrophoresis as previously described (Youngman et al., 2004), using the same running buffer, and the labeled peptide was located by autoradiography. The identities of PheMet, PheLysMet, and PheValLysMet, (Figures 4A, Figure 4-figure supplement 1, Figure 5C) were confirmed by their comigrations with authentic samples obtained from GenScript (Piscataway, NJ). A further demonstration of tetrapeptide identity was provided by matrix-assisted laser desorption/ionization (MALDI) mass spectrometric analysis (Ultraflex III TOF/TOF, Bruker: Phe-Val-Lys-Met(Na+), calculated, 546.7; found 546.6. In addition, the 80S·IRES complex containing P-site bound Phe-Val-Lys-Met-tRNA was reacted for 40 min with 10 mM puromycin (37°C, buffer 4 plus 1 mM GTP). The resulting puromycin adduct, Phe-Val-Lys-Met-puro(H+), released into solution, was also identified by MALDI: calculated, 978.7; found, 978.9.
80S·FKVRQWLM-IRES complexes containing the appropriate peptidyl-tRNA in the P-site, prepared using the standard procedure (see Complex preparations above)were mixed with [35S]-Met-TC (1.6 µM) and additional TCs as required (all 1.6 µM), for various times followed by quenching with 0.5 M MES (pH 6.0) buffer. PheLysValArgGlnTrpLeuMet octapeptide synthesis was measured by [35S]-Met cosedimentation with 80S·FKVRQWLM-IRES complexes. Over the time scale of these measurements (60 –600 s, Figure 5C), all [35S]-Met-tRNAMet stably bound to the ribosome undergoes a peptide transfer reaction (see Table 2). PheLysValArgGlnTrpLeuMet peptide was released from tRNAMet using the base treatment described above for other peptidyl tRNAs and its identity was confirmed by its comigration during thin layer electrophoresis with an authentic sample obtained from GenScript (Piscataway, NJ) (Figure 5C—figure supplement 1).
Rates of puromycin adduct formation were measured for FVK-tRNALys bound in the A-site, and for both FVK-tRNALys and FVKM-tRNAMet either pre-bound in the P-site or undergoing translocation from the A-site to the P-site. In all cases, reaction mixtures were quenched with 0.5 M MES (pH 6.0) buffer. The quenched samples were next ultracentrifuged and the radioactivity co-sedimenting with the ribosome, which decreases as more puromycin adduct is formed, was determined. All incubations and reactions were carried out at 37°C. All complexes were reacted with puromycin (1 mM, final concentration) for various times before quenching. No decreases in radioactivity co-sedimenting with the ribosome were observed in the absence of added puromycin.
80S·FVKM-IRES complex with FVK-tRNA pre-bound at A-site was formed by incubating the 80S·FVKM-IRES complex containing FV-tRNA at the P-site (Structure 7, POST-2, 0.4 µM), purified by sedimentation through a sucrose cushion, with 0.8 μM [3H]-Lys-TC for one minute. The resulting complex was then reacted with puromycin.
Two procedures were employed, which yielded equivalent results. Procedure 1: 80S·FVKM-IRES complex containing FV-tRNA at the P-site (Structure 7, POST-2, 0.4 µM), purified as described above, was incubated with 0.8 μM [3H]-Lys-TC and eEF2.GTP (1.0 µM) for 15 min at 37°C. The resulting complex was then reacted with puromycin. Procedure 2: 80S·FVKM-IRES complex (0.8 µM) was preincubated for 15 min with eEF2·GTP (1 µM) and Phe-TC, Val-TC, and [3H]-Lys-TC (all TCs present at 1.6 µM). The resulting complex was then reacted with puromycin.
80S·FVKM-IRES complex (0.8 µM) was preincubated for 15 min with eEF2·GTP (1 µM) and Phe-TC, Val-TC, Lys-TC and [35S]-Met-TC (all TCs present at 1.6 µM). The resulting complex was then reacted with puromycin.
80S·FVKM-IRES complex (0.8 µM) was preincubated for 15 min with eEF2·GTP (1 µM) and Phe-TC and Val-TC, both present at 1.6 µM, yielding PheVal-tRNAVal bound in the P-site. This complex was then mixed for 1 min with ([3H]-Lys-TC (1.6 µM) to form PheValLys-tRNALys bound in the A-site, which was then reacted with puromycin in the presence of additional added eEF2·GTP (final concentration 1 µM).
80S·FVKM-IRES complex (0.8 µM) was preincubated for 15 min with eEF2·GTP (1 µM) and Phe-TC, Val-TC, and Lys-TC (all TCs present at 1.6 µM), yielding PheValLys-tRNALys bound in the P-site. This complex was then mixed for 1 min with ([35S]-Met-TC (1.6 µM) to form PheValLysMet-tRNAMet bound in the A-site, which was then reacted with puromycin in the presence of additional added eEF2·GTP (final concentration 1 µM).
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Alan G HinnebuschReviewing Editor; National Institute of Child Health and Human Development, United States
In the interests of transparency, eLife includes the editorial decision letter and accompanying author responses. A lightly edited version of the letter sent to the authors after peer review is shown, indicating the most substantive concerns; minor comments are not usually included.
Thank you for submitting your work entitled "One, two, three, go – initiating polypeptide elongation in an IRES-dependent system" for consideration by eLife. Your article has been favorably evaluated by John Kuriyan (Senior editor) and three reviewers, one of whom, Alan Hinnebusch, is a member of our Board of Reviewing Editors.
The reviewers have discussed the reviews with one another and the Reviewing Editor has drafted this decision to help you prepare a revised submission.
Summary of work:
This paper conducts in vitro experiments using a purified system to study the rates of initial oligopeptide synthesis by an 80S·CrPV-IRES complex. The results suggest that the first two elongation cycles involved in producing a di- or tripeptide are slow and are limited by the translocation step, and that the rate of elongation accelerates following translocation of tripeptidyl-tRNA to the P-site (the second true translocation event). The IRES was suitably modified to replace the 1st Ala codon with a Phe codon, allowing fluorescence anisotropy measurements of the rate of proflavin-modified Phe-tRNAPhe binding to either the A- or P-site, and to allow incorporation of labeled Met-tRNAMet to monitor production of FM, FKM, FVKM, and FKVRQWLM synthesis. Sedimentation of complexes through sucrose gradients was used to assay stable binding of Met-tRNA or peptidyl-tRNAs containing labeled Met to the ribosome. The anisotropy results in Figure 3 suggest that EF2 shifts the equilibrium of the first pseudotranslocation step in which the IRES is moved in the decoding center to vacate the A-site, and thereby increases the rate of Phe-tRNAPhe binding to the A site in a biphasic reaction. The sedimentation assay indicates that the second pseudotranslocation step, in which Phe-tRNA is translocated from the A to the P site is much slower than the combination of the first pseudotranslocation plus Phe-tRNAPhe binding to the A site. They go on to use a rapid mixing and quench assay to measure rates of FM, FKM, and FKVM synthesis, with detection and quantification of product by thin layer electrophoresis (TLE), using the appropriate pre-formed substrates for addition of labeled Met to Phe-tRNAPhe or the relevant peptidyl tRNA. Results in Figure 4 indicate that the rates of aa-tRNA binding to the A site and formation of a peptide bond are very similar for all 3 substrates that produce a di-, tri-, or tetrapeptide, and the calculated rates of translocation in these reactions indicate that the rates of translocation for the dipeptidyl- and tripeptidyl-tRNAs are slow and rate-limiting in forming the tri- and tetrapeptides, respectively. Using the sedimentation assay, they could also measure the very rapid rates of ternary complex binding to the A site in the 2nd and 3rd elongation cycles producing the tri- and tetrapeptides, but found that A-site bound TC or FM-tRNA was too unstable to capture for the intermediates 5-6 containing the IRES in the E site, which they speculate might involve an allosteric effect of the IRES in destabilizing A-site binding. By then measuring rates of puromycin reaction with either the tri- or tetrapeptidyl tRNAs either under conditions where prior translocation from the P-site to the A-site is required or where the peptidy-tRNAs are prebound to the A site, they deduce that the rate of the 3rd translocation of tetrapeptidyl-tRNA from the P to A site is very rapid compared to the 2nd or 3rd translocation events.
The following issues raised in the referees' comments shown below need to be addressed in the revised manuscript.
Ref. #1: All of the requests for clarification of reaction conditions or interpretations of results should be addressed with appropriate revisions of text.
Ref.#2: The request for complete details about fitting the kinetic data in comment #1 should be honored.
As described in comment #2, the need for modifying the IRES sequence should be more explicitly stated in Materials and methods, as it would have been technically feasible to carry out all experiments with the WT sequence. In addition, it is important to compare the rates of IRES-mediated translation given by the native sequence versus your modified IRES sequence either in a cell-free extract or in your reconstituted system.
To respond to points 3. & 4., it would be sufficient to revise the text to remove statements about allostery; for point 5, to explain why you monitored the puromycin reaction as you did in Figure 5A-B. For points 6-10, clarify the relevant issues with the appropriate revisions of text, and provide the requested control to rule out Met-tRNA binding to 30S subunits.
Ref. #3: Points 1.-3 concern your conclusion that eEF2 does not stimulate the rate of TC binding in step 2 and rather only shifts the equilibrium in step 1 towards structure 2, which is at odds with other published work. As this is a hypothetical interpretation of the biphasic kinetics in Figure 3, it is important that you attempt to provide evidence for this deduced effect of eEF2 on the equilibrium of step 1 using toe-printing analysis of the complexes. It is also necessary to justify the statement that k-1 is much smaller than k2, which is not self-evident, and explain how it could be that eEF2 would shift the equilibrium to structure 2 without altering the t1/2 of step 1. You must also discuss the discrepancy with the previous results, particularly those of Ruehle et al.
You should clarify the issues raised in points 4-6 with the appropriate revisions of text.
The results are significant in providing strong evidence that the two pseudotranslocation steps (steps 1 and 3 in Figure 2) as well as the two conventional translocation steps (steps 6 and 9) involved in producing a tripeptide and translocating it to the P site occur slowly and are rate-limiting for the first two elongation cycles of CrPV IRES-directed translation, and that the next translocation step occurs much more rapidly and does not appear to be rate-limiting for production of longer polypeptides. The slow rate of the 2nd translocation step is surprising because the IRES should no longer occupy the decoding sites, so they may be detecting a novel intermediate with the IRES still bound to the ribosome outside of the decoding center and affecting a ribosome conformational change involved in translocation. The results are also significant in providing evidence against the previous conclusion by others that EF2 is required for the first pseudotranslocation that moves the IRES out of the A site (step 1) to permit recruiting the first TC to the A-site, as here they found no effect of EF2 on the rate of this reaction when it is monitored directly by fluorescence anisotropy measurements as opposed to stable TC binding to the P site assayed by sedimentation, which occurs only after the second pseudotranslocation reaction (step 3). Finally, the results using the CrPV IRES are valuable in laying the groundwork for studying eukaryotic elongation in a simplified purified system without the need for initiation factors to form the first peptide bound. There are a number of instances where the reaction conditions or interpretations of results need to be more carefully described, as follows:
1) In the second paragraph of the subsection “Rates of oligopeptide formation and Met-tRNAMet cosedimentation”: This section is very confusing as the composition of the aa-tRNA or peptidyl-tRNA species in question, and whether it is bound to the A or P sites, is generally unclear. Particularly, in the clause "…These rate differentials provide a clear indication that peptidyl- Met-tRNAMet and Met-tRNAMet bound to the A-site (Structures 8, 9 and 11, 12), as well as peptidyl-Met-tRNAMet binding to the P-site (Structures 10 and 13), efficiently…", the clarity could be improved by mentioning the specific peptidyl-tRNAs by their amino acid compositions for each of the structures, as "peptidyl-Met-tRNAMet" is imprecise nomenclature.
2) In the subsection “Rates of oligopeptide formation and Met-tRNAMet cosedimentation”: Also, it's not perfectly clear whether the instability of Met-tRNA binding to the A-site observed for structure 3 and 5, and FM-tRNA to structure 6 using the co-sedimentation assay is unusual; whereas stable binding of both Met-tRNA and peptidyl-tRNA to the A-site in structures 8, 9, 11-12 is typical of A-site-bound ligands in conventional (non-IRES) elongation complexes; and this distinction needs to be carefully spelled out.
3) In the second paragraph of the subsection “Translocation of tetrapeptidyl-tRNA (Step 12) is much more rapid than of tripeptidyl-tRNA (Step 9)”: requires citation of a figure for puromycin synthesis rates.
4) In the second paragraph of the subsection “Translocation of tetrapeptidyl-tRNA (Step 12) is much more rapid than of tripeptidyl-tRNA (Step 9)” and Figure 5A-B: more detail is required about how the experiment in Figures 5A-B was done in terms of starting reactants.
5) In the last paragraph of Results: more detail is required to understand how binding of peptidyl tRNA harboring labeled Met is distinguished from Met-tRNA binding to the A-site in these assays.
6) Error estimates have not been defined in any of the figures or tables and this needs to be rectified.
The paper by Cooperman et al. provides a first kinetic description of the initial steps of translation by eukaryotic ribosomes after the initiation at an IRES. There are very few papers on mechanistic aspects of eukaryotic translation in general and the present work provides very interesting, significant insights. It is a real challenge to obtain clean kinetic data for the eukaryotic system and this paper provides valuable information on the kinetic properties of eukaryotic ribosomes. The technical quality of the data is high (I particularly like the quality of the stopped-flow traces, they are really excellent) and the conclusions are warranted. My only reservation is that many technical aspects are not clearly described. Given that this paper delineates new kinetic approaches to study eukaryotic translation, it is particularly important to provide a very detailed description of methods. This really has to be carefully revised. I personally do not like the title, because it is not informative at all; a more rigorous title would be better.
1) Fitting of the kinetic data in this paper is mostly non-exponential and would require fitting to a model by numerical integration. This part is completely missing in Materials and methods. This must be described in all details required to understand and reproduce the calculations.
2) Among all potential criticism of the experimental system (Discussion, third paragraph), I am particularly worried about the altered coding sequence. It is not clear why the native Ala codon was replaced by Phe, as unlabeled amino acids are used anyway and the tRNAs are prepared by a method which should readily yield tRNA(Ala), i.e. one could label tRNA(Ala) with Prf (Kothe and Rodnina, 2007). This means that in principle one could use the native sequence. Furthermore, the measurements and rate determinations would be even more straightforward if [14C] and [3H]-labeled amino acids were used instead of [35S]Met; in this case one could use the native sequence. The need for modifying the sequence should be more explicitly stated in Materials and methods. Clearly, the whole set of experiments cannot be repeated with Ala as a 1st amino acid; however, it would be highly desirable to compare IRES-mediated translation with a native sequence and a modified sequence in the cell-free translation system (commercial or reconstituted from components).
3) The "instability" of aa-tRNA binding in A site is not likely to be caused by the dissociation of aa-tRNA from the A site prior to peptide bond formation. As authors show, peptide bond formation is rapid (Table 1); so as soon as aa-tRNA binds, it will be rapidly incorporated into peptide. Instead, peptidyl-tRNA tends to drop-off easily (see Semenkov 2000 and Konevega 2004). Although this dissociation is relatively slow, it normally explains the drop-off during centrifugation (which takes minutes to hours). In any case, there is no evidence for the allosteric interactions between the A and E sites. The statements on the allosteric interplay should be removed, as they only weaken the paper (subsection “Rates of oligopeptide formation and Met-tRNAMet cosedimentation”, third paragraph) (By the way, the so-called "allosteric effect" is simply caused by the presence of deacylated tRNA in aa-tRNA preps, which chases the labeled tRNA from the E site. There are several other groups, in addition to Rachel Green, who provided strong evidence against the model).
4) The instability issues addressed in point 3 could be overcome by using nitrocellulose filtration assays, which are really very commonly used to study ribosome studies. Are there any technical issues which preclude the use of nitrocellulose filter in this case? If yes, this should be described in Materials and methods.
5) The authors use puromycin reaction to identify the P-site position of the peptidyl-tRNA, which is a very reliable and well-established method. It is therefore surprising that the authors did not use the time-resolved puromycin assays (which they have established for the prokaryotic ribosomes) to determine the rates of translocation in a more direct fashion than described in Table 1? If there is some technical issue specific for eukaryotic ribosomes, it would be important to indicate this in Materials and methods.
6) Discussion: "Further work will be required to determine how many cycles of elongation are required before any retarding influence of bound IRES is completely eliminated": I am confused here. Table 1 shows that k12 is very fast, i.e. the 5th round is already rapid. This should be clarified.
7) In the subsection “Complex preparations. TCs and various 80S·IRES complexes”. 30S subunits were added to the centrifugation assays as a carrier. However, 30S subunits can bind [35S]Met (at least to some extent) even in the absence of the mRNA. Are there controls for that?
8) Figure 1 legend. "in the both cases" – it is not clear which two cases are meant. It is also not clear why the incubation with EF2 is so long (1-2 hours) – is it really necessary? I guess the authors just wanted to be on the safe side, but this has to be clearly stated in Materials and methods.
9) Figure 4 legend; "or just [35S]Met-TC”. My expectation is that in the absence of the preceding Lys-TC Met should not be incorporated at all. This has to be clarified.
10) In the subsection “Kinetic measurements”, "averages of 2-4 independent determinations" – do the authors mean technical replicates or independent experiments (i.e. biological replicated)?
Reviewer #2 (Additional data files and statistical comments):
The statistical information is appropriate except for the clarification needed in the subsection “Kinetic measurements” (point 10 in the review).
Zhang and colleagues have investigated the effect of the CrPV IRES RNA on the first rounds of translation elongation including the initial pseudo-translocation steps. They have performed for the first time experiments to determine the rates of the various main steps of the elongation cycles. They show that the IRES initial significantly slows down the translocation steps are down until the tetrapeptide stage is reached. The CrPV is an important minimal model system for translation initiation in eukaryotes The results are novel and interesting. However, the taking into account the following points is recommended before the paper can be published.
1) According to Figure 3, blue curve, the binding of tRNA to the A-site (structure 2 in the nomenclature of the authors) reaches nearly the same level with or without eEF2. However, in previous experiments binding of tRNA was dependent on eEF2 or at least significantly increased (Yamamoto et al., 2007; Fernandez et al., 2014; Ruehle et al., 2015). The experiments by Ruehle et al. have been done by colocalization using fluorescence. The discrepancy should be discussed. Furthermore, the authors should perform experiments to establish the nature of the complex in a more direct manner, e.g. by toeprinting of the complex.
2) The authors state that k-1 is much smaller than k2 and was assumed to be negligible. How has this been determined? In fact, a toeprint signal indicative of translocation cannot be obtained by incubation of binary 80S-IRES complexes with eEF2 alone, but requires an A-site ligand, too (Jan et al., 2003; Muhs et al., 2015). This suggest that the translocated complex (structure 2) is kinetically labile and contradicts the present statement that the back-translocation is negligible. Again, the authors should do experiments, e.g. toeprinting, to provide additional evidence that pre-incubation with eEF2-GTP shifts the equilibrium between structures 1 and 2 from 95:5 to 50:50 as stated.
3) The explanation about the effect of eEF2 on the equilibrium between structure 1 and 2 (Discussion) is curious. They propose that eEF2 shifts the equilibrium towards structure 2 at the same time do not measure a significant effect on the t1/2 of step 1. How this is possible? Do the authors propose that k-1 is becoming smaller, i.e. that eEF2 slows down back-translocation without accelerating forward translocation?
4) Can the authors rule out that back-translocation or peptidyl-tRNA drop off influences the following steps towards tetrapeptide synthesis?
5) The first sentence of the paper "Initiation of protein synthesis in eukaryotic cells proceeds via two well-established pathways" is to some extent misleading. As the authors acknowledge later factor requirement of different IRES RNAs is diverse and the structure and mechanism of different IRES RNAs is also different. So there are several pathways leading to internal initiation.
6) The peptide coding sequence is not really part of the IRES. Therefore, it is misleading to talk about mutant IRES, when the changes are in the ORF.
7) At the beginning of the Results part the authors should specify in which system they are working.
8) The green line is not mentioned in the legend to Figure 3.
9) A-site/E-site allostery has been recently reported by Ferguson et al., 2015, Mol Cell for human 80S ribosomes. This is relevant for the respective discussion in the third paragraph of the subsection “Rates of oligopeptide formation and Met-tRNAMet cosedimentation”.
[Editors' note: further revisions were requested prior to acceptance, as described below.]
Thank you for resubmitting your work entitled "Kinetics of initiating polypeptide elongation in an IRES-dependent system" for further consideration at eLife. Your revised article has been favorably evaluated by John Kuriyan (Senior editor), a Reviewing editor, and two reviewers.
The manuscript has been improved but there are some remaining issues raised by Reviewer #3 that need to be addressed before acceptance. In considering these new comments, Reviewer #2 also asks that you carefully review the Materials and methods to insure that they contain all of the critical information that would be required to duplicate the work.
The authors have significantly improved the manuscript by satisfactorily addressing the comments and concerns of the reviewers. The story they prevent extends our knowledge on the function IRES-dependent systems and can be published in eLife.
In the revised version Zhang and colleagues have addressed many points raised by the reviewers and have improved their paper. However, there are still some points where discussion should be extended.
1) The authors now confirm that the effect of eEF2 is to inhibit k-1 and write that this is "consistent with its role as a translocase". However, the accepted role of EF-G/eEF2 is to accelerate translocation, not to inhibit back-translocation. This warrants additional discussion.
2) The authors have pre-incubated 80S-FVKM-IRES complexes with eEF2-GTP for 1 – 2 hr. How they can rule out that GTP consumption has an impact on the experiment? Is it possible that accumulation of eEF2-GDP during pre-incubation results in a significant fraction of 80S FVKM-IRES-eEF2 complexes (between Structures 1 and 2) to slow down k2.
3) The authors write that the anisotropy assay in Figure 3 reports on formation of Structure 3 from 1. However, as the co-sedimentation assay reports, during the measurement time there should be also formation of Structure 4 at later time.
4) The authors state that A-site-bound Phe-tRNAPhe is labile. Can they estimate the off-rate k-2. Is it valid to neglect k-2 for numerical integration?https://doi.org/10.7554/eLife.13429.017
- Haibo Zhang
- Martin Y Ng
- Yuanwei Chen
- Barry S Cooperman
The funder had no role in study design, data collection and interpretation, or the decision to submit the work for publication.
We thank Eric Jan for gifts of wt and variant CrPV Phe-IRES vectors and for helpful discussions.
- Alan G Hinnebusch, Reviewing Editor, National Institute of Child Health and Human Development, United States
© 2016, Zhang et al.
This article is distributed under the terms of the Creative Commons Attribution License, which permits unrestricted use and redistribution provided that the original author and source are credited.