The Ribosomal Stalk Binds to Translation Factors IF2, EF-Tu, EF-G and RF3 Via a Conserved Region of the L12 C-Terminal Domain
doi:10.1016/j.jmb.2006.10.025 J. Mol. Biol. (2007) 365, 468–479
The Ribosomal Stalk Binds to Translation Factors IF2, EF-Tu, EF-G and RF3 via a Conserved Region of the L12 C-terminal Domain
Magnus Helgstrand1, Chandra S. Mandava2, Frans A. A. Mulder1,3 Anders Liljas3, Suparna Sanyal2⁎ and Mikael Akke1⁎
1Department of Biophysical Efficient protein synthesis in bacteria requires initiation factor 2 (IF2), Chemistry, Lund University, elongation factors Tu (EF-Tu) and G (EF-G), and release factor 3 (RF3), each Box 124, SE-22100 of which catalyzes a major step of translation in a GTP-dependent fashion. Lund, Sweden Previous reports have suggested that recruitment of factors to the ribosome and subsequent GTP hydrolysis involve the dimeric protein L12, which 2Department of Cell and forms a flexible “stalk” on the ribosome. Using heteronuclear NMR Molecular Biology, Uppsala spectroscopy we demonstrate that L12 binds directly to the factors IF2, University, Box 596, BMC, EF-Tu, EF-G, and RF3 from Escherichia coli, and map the region of L12 SE-75124 Uppsala, Sweden involved in these interactions. Factor-dependent chemical shift changes 3Department of Molecular show that all four factors bind to the same region of the C-terminal domain Biophysics, Lund University, of L12. This region includes three strictly conserved residues, K70, L80, and Box 124, SE-22100 E82, and a set of highly conserved residues, including V66, A67, V68 and Lund, Sweden G79. Upon factor binding, all NMR signals from the C-terminal domain become broadened beyond detection, while those from the N-terminal domain are virtually unaffected, implying that the C-terminal domain binds to the factor, while the N-terminal domain dimer retains its rotational freedom mediated by the flexible hinge between the two domains. Factor- dependent variations in linewidths further reveal that L12 binds to each factor with a dissociation constant in the millimolar range in solution. These results indicate that the L12-factor complexes will be highly populated on the ribosome, because of the high local concentration of ribosome-bound factor with respect to L12. © 2006 Elsevier Ltd. All rights reserved. *Corresponding authors Keywords: L12; ribosome; GTPase; NMR spectroscopy; protein synthesis
Introduction In the past few years, structural studies by X-ray crystallography and cryo-electron microscopy (cryo- EM) have revolutionized our understanding of Ribosomes are molecular machines, whose flex- 2–9 ible components enable the dynamic process of ribosome function at atomic resolution. As protein synthesis that includes movement along the increasingly more structural information on the mRNA, tRNA translocation and peptidyl transfer.1 ribosome becomes available, the focus of the field is shifting towards understanding the detailed – motions necessary to accomplish translation.10 14 As a general rule, G-protein translation factors Present address: F. A. A. Mulder, Department of promote the transitions between different structural Biophysical Chemistry, University of Groningen, states of the ribosome, and these processes often Nijenborgh 4, 9747 AG Groningen, the Netherlands. involve GTP hydrolysis in a ribosome-dependent Abbreviations used: CTD, C-terminal domain; EF-G, manner. Protein L12 (also denoted L7/L12) is the elongation factor G; EF-Tu, elongation factor Tu; EM, main component of the prominent “stalk” of the 50 S electron microscopy; HSQC, heteronuclear ribosomal subunit in bacteria, and has been impli- single-quantum correlation; IF2, initiation factor 2; NTD, cated as a key player in factor recruitment and – N-terminal domain; RF3, release factor 3. factor-dependent GTP hydrolysis,15 17 as outlined E-mail addresses of the corresponding authors: further below. Functionally analogous proteins are [email protected]; [email protected] present in eukaryotes and archaea, but these do not
0022-2836/$ - see front matter © 2006 Elsevier Ltd. All rights reserved. L12 binds to IF2, EF-Tu, EF-G, and RF3 469 show any amino acid sequence homology to the release of inorganic pyrophosphate after GTP – L12.18 20 L12 forms a symmetric dimer mediated hydrolysis.17 A recent cryo-EM reconstruction of EF- by its N-terminal domains (NTD), to which each C- G bound to the ribosome suggests a direct interac- terminal domain (CTD) is connected flexibly via an tion of the CTD of L12 with the G' domain of EF-G.37 – unstructured hinge segment.21 23 High-resolution There is indirect evidence for interactions between structures are available for the isolated CTD of L12 and EF-Tu as well. It has been shown that Escherichia coli L12,24 as well as for the intact E. coli specific mutations in the L12 CTD affect the binding L12 dimer.21 L12 is the only multi-copy protein of of the EF-Tu–GTP–aminoacyl-tRNA ternary com- the ribosome, with two or three dimers binding to plex to the ribosome.38 These data, together with the the ribosome via protein L10, depending on the structural homology between the L12 CTD and EF- organism.16 Recently, the structure of the complex Ts, have been used to generate a model for the between L10 and the dimeric NTDs of L12 from structure of the EF-Tu–L12 complex,38 based on the Thermotoga maritima was determined, which re- crystal structure of the EF-Tu–EF-Ts complex.39 It is vealed that the L12 NTD dimers bind to helix α8 an open question whether L12 also stimulates the of L10. Apparently, the length of α8 determines the GTPase activity of IF2 and RF3. A recent cryo-EM number of L12 dimers that bind to the ribosome in a structure of the initiation complex shows one L12 given organism; helix α8 is longer in Thermotoga L10 CTD intercalated between L11 and the G-domain of and contains one additional (third) binding site for a IF2.9 Because IF2, EF-Tu, EF-G, and RF3 have two dimer of L12, compared with the case in E. coli.16 homologous domains and occupy overlapping sites – Notably, the L12 dimers are highly disordered in all on the ribosome,9,31,32,40 42 it is expected that ribo- high-resolution crystal structures of the 70 S or 50 S some-bound L12 can interact with all of these particles.2,6,7,25 However, from cryo-EM reconstruc- factors. tions of the ribosome in various functional states, it Here, we demonstrate that L12 binds to IF2, EF- appears that the stalk acquires different conforma- Tu, EF-G, and RF3, through a strictly conserved tions during translation. Modeling of the L10– region of its CTD. Estimated affinities between the (L12NTD)4 complex into the electron densities of factors and L12 in solution indicate that the different cryo-EM structures suggests that the corresponding complexes will be highly populated orientation of helix α8 varies between functional on the ribosome. states.16 Furthermore, a body of structural and biochemical data has shown that parts of the hinge region and the entire CTD are mobile in ribosome- Results – bound L12,22,23,26 28 and that its conformation and flexibility are altered dramatically upon binding of Using heteronuclear NMR spectroscopy, we the translation factors EF-G10,23,29,30 or EF-Tu31,32 to investigated the transient interactions between L12 the ribosome. Evidence from chemical cross- and the translation factors IF2, EF-Tu, EF-G, and linking33 and immuno-EM34 studies shows that RF3, as well as bovine serum albumin (BSA), which the CTD contacts different parts of the ribosome, served as a negative control. Separate titration series including the 30 S neck and head regions, as well as were performed for each of IF2, EF-Tu, EF-G, RF3 a position close to the EF-G binding site near L11.35 and BSA, where the unlabeled protein was titrated Thus, there is ample evidence that L12 can attain into a solution of 15N-labeled L12, and a 1H-15N different conformational states during different heteronuclear single-quantum coherence (HSQC) steps of translation. spectrum was acquired at each titration point. In bacteria, translation involves four main G- Interactions between L12 and factor were identified proteins that catalyze different steps. These are: on the basis of changes in chemical shifts and initiation factor 2 (IF2), which is required for linewidths of cross-peaks in the HSQC spectrum of initiation; elongation factor Tu (EF-Tu), which L12, as a function of added factor. The chemical shift carries amino acid-charged tRNAs to the ribosome; is exquisitely sensitive to the magnetic environment elongation factor G (EF-G), which catalyzes tRNA of the nuclear spin, and the linewidth depends on translocation; and release factor 3 (RF3), which the size of the molecular system. Changes in releases the class I release factors after removal of chemical shifts and linewidths provide residue- the nascent peptide.1 All of these factors possess the specific information on changes in the local envir- well-conserved G-domain, which binds and hydro- onment and on the size of the molecular complex, lyzes GTP to carry out their functions, and the because each backbone N-H amide group in L12 succeeding β-barrel domain (domain II). It has been corresponds to a single cross-peak in the 1H-15N shown that L12 alone can stimulate a low level of HSQC spectrum. Thus, these data map out which GTP hydrolysis by EF-G.36 Furthermore, ribosomes parts of L12 interact with each factor. The response depleted of L12 are impaired in the GTPase was very similar in all titration series, and followed activation of both EF-Tu and EF-G.15 The molecular the general features described in detail below for the mechanism of this GTPase activation is not fully interaction between EF-G and L12. The HSQC understood; it has been suggested that L12 increases spectra obtained in the presence of IF-2, EF-Tu, the catalytic rate of GTP hydrolysis,36 or increases RF3 and BSA are available as Supplementary Data the rate of binding EF-G to the ribosome rather than (Figures S1–S4), and the results are summarized affecting the GTP hydrolysis step per se,16 or controls below. 470 L12 binds to IF2, EF-Tu, EF-G, and RF3
Changes in the 1H-15N HSQC of L12 upon addition of EF-G
Figure 1(a)–(d) show 1H-15N HSQC spectra of L12, acquired in the presence of various concentra- tions of EF-G. The spectral changes include both subtle and dramatic effects. At 0.4–0.8 molar equivalents of EF-G (with respect to the L12 dimer), a subset of cross-peaks in the 1H-15N HSQC of L12 shows gradual changes in chemical shifts from those obtained in the absence of EF-G (Figure 1(a)–(c)). These cross-peaks correspond to residues V66–V68, K70, L80, and E82, which are located on a continuous surface region of the L12- CTD. The chemical shift changes become accentu- ated with increasing concentrations of EF-G. Con- comitantly, all cross-peaks from the CTD become increasingly broadened, and eventually most cross- peaks disappear from the spectrum at the point where 1.2 equivalents of EF-G, with respect to L12 dimer, have been added (Figure 1(d)). In contrast, the cross-peaks from the hinge region and the NTD remain virtually unaffected by the presence of EF-G. These differential changes in line broadening serve as an important verification of binding, as explained in detail below (see Binding affinities). The present results clearly establish that factor-bound and free L12 are in fast to intermediate exchange, because (i) the chemical shifts of the residues in the binding interface change gradually upon addition of factor, and (ii) the CTD cross-peaks disappear well before the addition of one equivalent of factor with respect to L12 monomer, which indicates that each L12 CTD is exchanging between free and factor-bound states within the time-scale of the HSQC experiment.
Interactions between translation factors and L12
Figure 2 presents close-up views of a representa- tive set of residues in L12 that experience chemical shift changes upon addition of factors. In addition, Figure 2 includes one representative residue from each of the NTD and CTD (M17 and K108, respectively), whose chemical shifts do not change. Figure 2(a) shows the chemical shift changes observed for residues K70 (left-hand panel) and L80 (middle panel) during titration with IF2. Comparisons with residues M17 and K108 (right- hand panel) demonstrate clearly that the observed chemical shift changes are significant, even though these are relatively small (∼0.04 ppm in the 1H dimension). Figure 3(a) summarizes the factor- dependent chemical shift changes in L12 upon addition of IF2. In the IF2–L12 complex, the largest chemical shift perturbations occur for residues K70, 1 15 L80 and E82, with minor changes observed for the Figure 1. H- N HSQC spectra of the L12 dimer with neighboring residues V66 and V68, and also for V20 different concentrations of added EF-G. Cross-peaks are in the NTD. color-coded according to the domain structure of L12: green, NTD; red, hinge region; blue, CTD. (a) No factor Figure 2(b) shows the behavior of residues V66 added; (b) 0.4 equivalent of EF-G (with respect to L12 and E82 during titration with EF-Tu, and Figure 3(b) dimer); (c) 0.8 equivalent; (d) 1.2 equivalents. Black circles summarizes the data for all residues. The largest indicate the peak positions in the absence of EF-G for those chemical shift changes are observed for residue E82, cross-peaks that move significantly upon addition of with smaller changes observed for V66 and V68. In EF-G. The concentration of L12 dimer was 0.5 mM. L12 binds to IF2, EF-Tu, EF-G, and RF3 471
Figure 2. Close-up views of selected cross-peaks in superimposed 1H-15N HSQC spectra of the L12 dimer, obtained at different equivalents (molar ratios), x, of added factor with respect to L12 dimer. Two representative cross-peaks exhibiting significant chemical shift changes are shown for each titration series (left-hand and middle panels), together with two representative cross-peaks, M17 and K108, that are unaffected by factor addition (right-hand panel). (a) Addition of IF2, showing residues K70 (left) and L80, (middle); x=0, 0.4, 0.8, 1.2, 1.6. (b) Addition of EF-Tu, showing residues V66 (left) and E82 (middle); x=0, 0.1, 0.2, 0.3, 0.4. (c) Addition of EF-G, showing residues V67 (left) and L80 (middle); x=0, 0.4, 0.8, 1.2, 1.8. (d) Addition of RF3, showing residues G79 (left) and L80 (middle); x=0, 0.4, 0.8, 1.1, 1.7. The cross-peak contours are colored according to the increasing molar ratio of the added factor in the order yellow, red, brown, black. this dataset, there is a drift in pH with factor of EF-G. Figure 3(c) summarizes the quantitative addition (presumably due to insufficient buffering analysis of the data obtained for the interaction capacity of the solutions) that results in a change in between EF-G and L12. Again, the largest shift chemical shift across the entire sequence, as exem- changes in L12 induced by the addition of EF-G are plified by residues M17 and K108 in the right-hand observed for residues V66–V68, K70, L80, and E82. panel of Figure 2(b). The magnitude, but not the Figure 2(d) shows the chemical shift titrations direction, of the chemical shift changes due to the displayed by residues G79 and L80 upon addition of drift in pH varies between residues, thus making RF3 to L12. The largest chemical shift perturbations this data set noisier than the others. However, the occur for residues L80 and E82, with minor changes chemical shift changes associated with the pH drift observed for E27 and G79 (Figure 3(d)). In this case, do not mask those associated with factor binding, as the chemical shift change for E27 in the NTD is evident from Figure 3(b). clearly significant, but most likely due to titration of Figure 2(c) shows expanded regions of the spectra a nearby charged group, e.g. the carboxylate side- displayed in Figure 1, highlighting the titrating chain, rather than to interactions with RF3. This chemicalshifts ofresidues V67and L80uponaddition issue is further addressed below. 472 L12 binds to IF2, EF-Tu, EF-G, and RF3
Control experiment: titration of L12 with BSA formed a negative control experiment by titrating L12 with BSA. Upon addition of BSA to L12, the To verify that the identified interactions between largest chemical shift perturbation occurs for residue the factors and L12 do not reflect promiscuous E27 in the NTD (Figure 3(e)), paralleling the binding due to e.g. hydrophobic effects, we per- observations for RF3. There was no significant chemical shift change in the CTD upon addition of BSA.
Non-specific chemical shift changes
In each titration series, minor chemical shift changes appear for a number of residues in the NTD (Figure 3). In the majority of cases, these changes are insignificant and are simply due to noise, but in a few cases the changes are significant. These aberrant results can be explained by minor changes in salt concentration, buffering capacity, or electrostatic screening due to altered concentrations of protein43 throughout the titration series. How- ever, these caveats do not pose any problem when interpreting the data in terms of factor binding, because the linewidths serve as a separate control of factor interactions, as explained below.
Differential line broadening upon factor binding
Factor binding leads to uniform broadening (decrease in peak height) of all cross-peaks from the CTD, while the linewidths of the cross-peaks from the NTD and the hinge region remain unaffected (Figure 4). This demonstrates that the CTD attains the slow rotational correlation time characteristic of a large molecular mass complex. In contrast, the NTD, which is attached flexibly to the CTD via the hinge region, experiences virtually the same degree of rapid rotational averaging as in the free state. These results provide firm evidence that the CTD binds to the G-protein factors, whereas the NTD does not. Importantly, the differential line broadening thus serves as an independent verifica- tion of factor binding, and enables us to discriminate chemical shift changes that are due to factor binding from those that result from non-specific interactions or electrostatic effects due to titrating groups. Both in the absence and in the presence of factors, a single set of cross-peaks is observed from the two copies of the L12 dimer, demonstrating that at each titration point the two CTDs experience the same magnetic environment on average. This fact reflects fast exchange between the free and factor-bound states
Figure 3. Factor-dependent chemical shift changes, δω/δx, plotted versus residue number. (a) IF2, (b) EF-Tu, (c) EF-G, (d) RF3, (e) BSA. The chemical shift change was evaluated as the slope, δω/δx, of a straight line fit to the data pairs (ω, x), where ω is the chemical shift and x is the molar ratio of factor versus dimeric L12. Cross-peaks are color-coded according to the domain structure of L12: green, NTD; red, hinge region; blue, CTD. Chemical shift changes for 1H and 15N are represented by boxes and circles, respectively. Values for 15N were scaled by a factor of 0.2 to make them comparable to 1H values. Note that the vertical scale differs between panels. L12 binds to IF2, EF-Tu, EF-G, and RF3 473 and equal probability of factor binding for the two Binding affinities CTDs of the symmetric dimer. Thus, the present results do not indicate that two CTDs bind The factor-dependent line broadening of L12 simultaneously to one factor molecule. Furthermore, enables us to estimate the affinity between L12 and it should be noted that addition of BSA to L12 does each G-protein factor. The linewidths of the cross- not result in any additional line broadening (Figure peaks from the free state contain contributions due 4(e)) clearly indicating that the observed interactions to relaxation in the bound state. It should be noted between L12 and the translation factors are specific. that chemical exchange does not contribute to the linewidths for the large majority of residues, which do not exhibit chemical shift changes upon binding of the CTD to the factor. Using a two-state model (free and factor-bound states), approximate trans- verse relaxation rates were calculated for each of the concentrations used in the experimental titrations. Dissociation constants (KD)wereobtainedby iteratively fitting Equations (1)–(4) to the experi- mental data for the CTD of L12, shown in Figure 5. These fits are constrained by the transverse relaxa- tion rates calculated on the basis of the rotational τ diffusion correlation times of the free state ( c,L12), which is known from experiment,22 and the factor- τ bound state ( c,F), which is estimated from an τ empirical relation between c and the molecular mass of the complex.44 The results are presented in Table 1. In each case, the factor-dependent line- widths of residues in the L12 CTD correspond to a KD value in the millimolar range. It should be noted that these KD values are rough estimates that τ depend on the estimated c,F of the bound states, which are subject to systematic errors. The errors in KD reported in Table I are the standard asymptotic errors of the fits, which reflect the quality of the fits, but do not include systematic errors. Plausible τ sources of systematic errors in c,F include signifi- cant inter-domain motions or rotational diffusion anisotropy. We evaluated the effects of systematic τ errors in c,F on the fitted value of KD: an error of τ ±20% in c,F corresponds to an error of ±50% in KD, with shorter correlation times yielding lower values of KD (i.e. higher affinities). Overall, the values of KD estimated here should be accurate to within one order of magnitude.
Discussion
The CTD of ribosome-bound L12 displays vir- tually the same degree of flexibility as in the free state.22,23 Thus, our present results on the L12-factor
Figure 4. Cross-peak intensities of residues in L12 plotted versus residue number. Data are shown for varying molar ratios (x) of factor to L12 dimer, with symbols according to increasing value of x in the order blue diamond, red square, yellow triangle, green circle, and purple cross. (a) IF2, x=0, 0.4, 0.8, 1.2, 1.6. (b) EF-Tu, x=0, 0.1, 0.2, 0.3, 0.4. (c) EF-G, x=0, 0.4, 0.8, 1.2, 1.8. (d) RF3, x=0, 0.4, 0.8, 1.1, 1.7. (e) BSA, x=0, 0.4, 0.9, 1.3, 1.8, 2.2. Cross-peak intensities were evaluated as peak heights. Data were normalized using the N-terminal residue, which is considered unaffected by factor binding. The domain structure of L12 is indicated by the colored bar above the x-axis: green, NTD; red, hinge region; blue, CTD. 474 L12 binds to IF2, EF-Tu, EF-G, and RF3
Figure 5. Cross-peak linewidths plotted versus molar ratio (x) of factor to L12 dimer. Linewidths were evaluated as described in the text for a subset of cross-peaks that are well resolved and observable over the entire concentration range of added factor in all four titrations. The increased dispersion in linewidths at larger values of x is primarily the result of increased uncertainty in the measurement as the lineshapes become broader. complexes in solution are expected to serve as valid the same helix-loop-helix motif (Figure 6, left-hand models for the corresponding interactions on the panels) in the CTD of L12. In general, the major ribosome. Previous reports have shown that the effects are observed for residues L80 and E82, translational GTPases bind to overlapping sites near located at the N-terminal tip of helix 5, and for – the base of the L12 stalk on the ribosome.9,31,32,40 42 V66, V68 and K70 in the N-terminal half of helix 4. Our present results show clearly that the translation As evident from the right-hand panels of Figure 6, factors also share an overlapping interaction surface the identified residues define a continuous surface on the CTD of L12. Figure 6 highlights in red the on the L12 CTD, except for the interaction with EF- residues in the L12 CTD that exhibit significant Tu. Notably, the residues forming the identified chemical shift changes upon factor binding. Yellow binding surface are highly conserved (Supplemen- color indicates those residues that are unaffected by tary Data Figure S5), and match partly with those factor addition, as well as a single residue (K65) for (K65, V66, I69, K70, R73, and K84) identified by site- which data are missing due to spectral overlap. In directed mutagenesis to interact with EF-Tu and EF- each case, the chemical shift perturbations pinpoint G.17,38 The apparent differences between the bind- ing surfaces for the different factors may suggest that the detailed interactions vary slightly between Table 1. Dissociation constants for the interactions the different L12-factor complexes. However, this between translation factors and L12 conclusion cannot be reached at present, because differences among the factors in their amino acid Translation factor KD (mM) composition of the binding surface can potentially IF-2 2.5±0.3 generate different chemical shift changes on L12, EF-Tu 0.2±0.1 even if the binding surface of the latter is identical in EF-G 0.4±0.1 all four cases. The same argument also rationalizes RF3 0.4±0.1 the apparent discrepancies between the present The reported errors in the fitted values of KD were evaluated as results and those derived from site-directed the standard asymptotic error of the fit. Systematic errors are not mutagenesis.17,38 Furthermore, the mutagenesis included; see the text for a detailed discussion of these. studies do not detect the binding interface between L12 binds to IF2, EF-Tu, EF-G, and RF3 475
Figure 6. Residues forming the factor-binding surface on the CTD of L12. Residues with significant chemical shift perturbations are colored red and labeled with residue number. The left-hand panels show ribbon diagrams of the backbone trace, while the right-hand panels show the contact surface. Binding surfaces are shown of (a) IF2, (b) EF-Tu, (c) EF-G, and (d) RF3. This Figure was prepared using Molmol.54 476 L12 binds to IF2, EF-Tu, EF-G, and RF3
L12 and the factor directly, but rather report the implicate a role for L12 in the recruitment of factors effects on binding that a given mutation exerts, to the ribosome, prior to factor activation by GTP which may be indirect. Thus, despite different binding. Addressing the potential effects of L12 on experimental conditions, the two methods confirm GTP hydrolysis requires that the factors be in the each other by identifying largely the same binding GTP-bound state. It is conceivable that the interac- surface. The present NMR data further generalize tion and affinity between L12 and a translational G- these observations by demonstrating that all the factor depend on whether the factor is bound to GTP major translational G-proteins interact with the CTD or GDP. Comparative analyses of each factor in of L12 in essentially the same way. different states (free and nucleotide-bound) will be Early studies recognized that the identified the subject of future studies. region on L12 was of potential functional In conclusion, we have demonstrated that the importance.24 The fact that a conserved region of CTD of L12 binds directly to the G-protein transla- L12 interacts with four different translational tion factors IF2, EF-Tu, EF-G, and RF3 using a GTPases suggests directly that the latter share a conserved binding surface. The novel observations common, and possibly conserved, structural feature presented here should provide important points of involved in the interaction. Recent cryo-EM results reference for model building of complexes between suggested that the CTD of L12 interacts with the G' L12 and IF2, EF-Tu, EF-G, and RF3, as well as for domain of EF-G.37 In light of our results, the detailed interpretation of three-dimensional recon- conclusion from the cryo-EM study is unexpected, structions based on cryo-EM images. because the G' domain is quite unique to EF-G. By contrast, all four translation factors share the G domain as well as domain II. Helix D of the G Materials and Methods domain in EF-Tu is involved in binding EF-Ts, and is most likely to interact with L12, as demonstrated NMR sample preparation previously.38 Our present results identifying the same binding surface for four G-factors on L12 The 15N-labeled ribosomal protein L12 from E. coli was strengthen the view that only a conserved domain produced and purified as described.22 The NMR samples on the G-factors would interact with L12, which contained L12 in polymix buffer45 at pH 6.8. The remains to be verified experimentally. concentration of the 15N-labeled L12 sample was approxi- The present data yielded estimates of the binding mately 0.5 mM, as measured by the Bradford assay. affinities between L12 dimer and each factor, which range between 0.2 mM and 2.5 mM (Table 1). It Expression and purification of G-protein factors should be noted that the estimated values of KD are approximate because they were derived using a His-tagged factors IF2, EF-Tu and EF-G were over- simplistic analysis, as discussed above. Previous expressed in E. coli BL21(DE3). The plasmids carrying His- studies have determined the dissociation constant tagged IF2 and EF-Tu were a kind gift from A. C. Forster.48 for the EF-Tu–L12 complex to be 10 μM,36 which is The His-tagged proteins were purified on a Ni-NTA in reasonable agreement with the value determined column using an imidazole elution gradient. The purity of here, considering that the previous study was EF-Tu was greater than 99% after this single purification performed in the presence of GTP. The millimolar step. EF-G and IF2 were further purified on a QFF column using a NaCl elution gradient. Protein solutions were dissociation constants should be compared to the concentrated using Ultra-spin concentrators (Amicon). local concentration of factor once it is bound on the E. coli RF3 cloned in the plasmid pOSEX3 was over- ribosome, which is of the order of at least 1 mM, expressed in E. coli MKH13 by salt induction and purified suggesting that the L12-factor complexes will be as described.49 All factors were dialyzed against polymix highly populated on the ribosome. It seems unlikely buffer45 and stored at −80 °C. Due to their limited stability that all copies of L12 interact with the G-protein in the absence of GDP,46,47 both EF-Tu and RF3 were factor that is bound to the ribosome. Indeed, the purified in the GDP-bound state. present identification of a single factor-binding site on the L12 CTD suggests strongly that a single CTD NMR spectroscopy binds to the ribosome-bound factor at a given time. The other copies of L12 could potentially interact All experiments were performed at 30.0(±0.1) °C on a with G-protein factors not yet positioned in their Varian Unity Inova 600 MHz spectrometer equipped with final binding sites on the ribosome. an inverse broadband probehead and single-axis pulsed- 1 15 The present experiments were performed with field gradient capabilities. H- N HSQC reference spectra samples prepared in polymix buffer,45 without were typically recorded with spectral widths of 8000 Hz, sampled over 512 complex points in the 1H dimension, added guanine nucleotides. Under these conditions, 15 IF2 and EF-G are most likely free of nucleotides, and 1650 Hz, sampled over 128 complex points in the N dimension. Chemical shift assignments of L12 were whereas EF-Tu and RF3 contain bound GDP, as a obtained as described.22,50,51 consequence of their high affinities for GDP in the 15N-labeled L12 was titrated with each of the proteins free (non-ribosome-bound) state, with KD values in IF2, EF-Tu, EF-G, RF3 and BSA. Each titration series 46,47 the nanomolar range. The present conditions typically included five additions of protein and an 1H-15N thus represent the situations where the factors are in HSQC spectrum was acquired at each titration point. their resting states. To this extent, our results Spectra were thus obtained for samples with molar ratios L12 binds to IF2, EF-Tu, EF-G, and RF3 477 of factor to L12-dimer of x=0, 0.4, 0.8, 1.2 and 1.6 for IF2; 0, 43182 Da; EF-G, 77450 Da; and RF3, 59443 Da; yielding τ 0.1, 0.2, 0.3 and 0.4 for EF-Tu; 0, 0.4, 0.8, 1.2 and 1.8 for EF- approximate values of c,F of: IF2, 49 ns; EF-Tu, 22 ns; EF-G, G; 0, 0.4, 0.8, 1.1 and 1.7 for RF3; and 0, 0.4, 0.9, 1.3, 1.8 and 40 ns; and RF3, 30 ns. The rotational diffusion correlation τ 22 2.2 for BSA. time for the L12 CTD in the free state is c,L12 =5.9 ns. Igor Pro (WaveMetrics, Inc.) was used to fit a global value for K and residue-specific scaling factors (A) as a function of NMR data processing and analysis D the amount of translation factor added to the L12 sample, relating the calculated linewidths to the experimental ones. 52 All data were processed using nmrPipe. The raw data The model given by Equations (1)–(4) was fitted to the 1 were apodized using a shifted sine bell in both the H and linewidths of 14 well-resolved amide 1H-15N cross-peaks 15 N dimensions. The final size of each matrix was (K100, D101, A103, E104, K108, L110, E111, A113, G114, 2048×256 real points after zero filling and Fourier A115, V117, E118, V119 and K120) that did not show any transformation. Cross-peak positions and intensities chemical shift differences between the free and factor- 53 were evaluated using CCPN analysis, and linewidths bound states. Hence, chemical exchange contributions to were extracted using nmrPipe. the transverse relaxation rates can be safely ignored. Linewidths were measured in both the 1Hand15N Determination of dissociation constants dimensions, resulting in 28 data points at each titration point for each factor. The same set of cross-peaks was used for all four factor titrations. The linewidths of the L12 CTD depend on the relative populations of free and factor-bound CTDs. Hence, these data report on the binding affinities between L12 and the factor in question. The dissociation constant is given by the equilibrium concentrations of factor-bound and free L12 dimer: Acknowledgements ½ ½ ¼ L12 F ð Þ KD ½ 1 Supported by research grants from the Swedish L12 : F Research Council to M.A., S.S. and A.L., from the in which [L12] denotes the concentration of free L12 dimer, Swedish Foundation for Strategic Research to M. [F] is the factor concentration and [L12:F] is the concentra- A., from the Göran Gustafsson Stiftelse to S.S., tion of factor-bound L12 dimer. Equation (1) is based on and from EU framework program 4 to A.L. F.A.A. the assumption that a single factor molecule binds to each M. was supported by a Marie Curie fellowship dimer of L12. Assuming further that factor binding (HPMF-CT-2001-01245) awarded by the European involves a single CTD in each L12 dimer, the relative Commission. populations of free and factor-bound CTDs are given by:
2½L12 þ½L12 : F p ¼ ð2aÞ Supplementary Data F 2ð½L12 þ½L12 : F Þ Supplementary data associated with this article ½L12 : F can be found, in the online version, at doi:10.1016/ p ¼ ð2bÞ B 2ð½L12 þ½L12 : F Þ j.jmb.2006.10.025
Thus, KD can be expressed in terms of the relative populations of free and factor-bound L12 CTDs: References ð2pF 1Þ½F KD ¼ ð3Þ 2pB 1. Liljas, A. (2004). Structural Aspects of Protein Syn- thesis, World Scientific Publishing Co., Singapore. and [F] can be expressed in terms of K and the total D 2. Ban, N., Nissen, P., Hansen, J., Moore, P. B. & Steitz, concentrations of L12 and factor. If the exchange rate T. A. (2000). The complete atomic structure of the large between the bound state and the free state is fast, the ribosomal subunit at 2.4 Å resolution. Science, 289, resulting linewidths can be approximated as: 905–920. 3. Wimberly, B. T., Brodersen, D. E., Clemons, W. M., Jr, R2 ¼ AðpBðsc;L12 þ sc;FÞþpFsc;L12Þð4Þ Morgan-Warren, R. J., Carter, A. P., Vonrhein, C. et al. τ τ where c,L12 and c,F are the correlation times for (2000). Structure of the 30S ribosomal subunit. Nature, rotational diffusion of free L12 and free translation factor, 407, 327–339. respectively. A is a residue-specific scaling factor that 4. Schluenzen, F., Tocilj, A., Zarivach, R., Harms, J., takes into account local variations in the relaxation rate, Gluehmann, M., Janell, D. et al. (2000). Structure of which are caused by a range of factors, e.g. fast internal functionally activated small ribosomal subunit at 3.3 fluctuations, the density of surrounding 1H spins, etc. angstroms resolution. Cell, 102, 615–623. Equation (4) is based on the assumption that the single 5. Gabashvili, I. S., Agrawal, R. K., Spahn, C. M., non-bound CTD of the L12 dimer in the factor-bound Grassucci, R. A., Svergun, D. I., Frank, J. & Penczek, state contributes in the same way to the linewidth (i.e. P. (2000). Solution structure of the E. coli 70S ribosome has the same relaxation properties) as each of the two at 11.5 Å resolution. Cell, 100, 537–549. τ CTDs in the free state. For each factor, the value of c,F 6. Yusupov, M. M., Yusupova, G. Z., Baucom, A., was approximated from an empirical relation,44 scaled to Lieberman, K., Earnest, T. N., Cate, J. H. & Noller, the present temperature and water viscosity using Stokes' H. F. (2001). Crystal structure of the ribosome at 5.5 Å law. The molecular masses are: IF2, 97350 Da; EF-Tu, resolution. Science, 292, 883–896. 478 L12 binds to IF2, EF-Tu, EF-G, and RF3
7. Schuwirth, B. S., Borovinskaya, M. A., Hau, C. W., gions of intact Escherichia coli ribosomes. Proc. Natl Zhang, W., Vila-Sanjurjo, A., Holton, J. M. & Cate, Acad. Sci. USA, 101, 10949–10954. J. H. D. (2005). Structures of the bacterial ribosome at 24. Leijonmarck, M. & Liljas, A. (1987). Structure of the 3.5 Å resolution. Science, 310, 827–834. C-terminal domain of the ribosomal protein-L7 8. Petry, S., Brodersen, D. E., Murphy, F. V., Dunham, protein-L12 from Escherichia coli at 1.7 Å. J. Mol. Biol. C. M., Selmer, M., Tarry, M. J. et al. (2005). Crystal 195, 555–580. structures of the ribosome in complex with release 25. Harms, J., Schluenzen, F., Zarivach, R., Bashan, A., factors RF1 and RF2 bound to a cognate stop codon. Gat, S., Agmon, I. et al. (2001). High resolution Cell, 29, 1255–1266. structure of the large ribosomal subunit from a 9. Allen, G. S., Zavialov, A., Gursky, R., Ehrenberg, M. & mesophilic eubacterium. Cell, 107, 679–688. Frank, J. (2005). The cryo-EM structure of a translation 26. Gudkov, A. T., Gongadze, G. M., Bushuev, V. N. & initiation complex from Escherichia coli. Cell, 121, Okon, M. S. (1982). Proton nuclear magnetic reso- 703–712. nance study of the ribosomal protein L7/L12 in situ. 10. Valle, M., Zavialov, A., Sengupta, J., Rawat, U., FEBS Letters, 138, 229–232. Ehrenberg, M. & Frank, J. (2003). Locking and 27. Cowgill, C. A., Nichols, B. G., Kenny, J. W., Butler, P., unlocking of ribosomal motions. Cell, 114, 123–134. Bradbury, E. M. & Traut, R. R. (1984). Mobile domains 11. Spahn, C. M., Gomez-Lorenzo, M. G., Grassucci, R. A., in ribosomes revealed by proton nuclear magnetic Jorgensen, R., Andersen, G. R., Beckmann, R. et al. resonance. J. Biol. Chem. 259, 15257–15263. (2004). Domain movements of elongation factor eEF2 28. Hamman, B. D., Oleinikov, A. V., Jokhadze, G. G., and the eukaryotic 80S ribosome facilitate tRNA Traut, R. R. & Jameson, D. M. (1996). Rotational and translocation. EMBO J. 23, 1008–1019. conformational dynamics of Escherichia coli ribosomal 12. Wilson, K. S. & Noller, H. F. (1998). Molecular protein L7/L12. Biochemistry, 35, 16672–16679. movement inside the translational engine. Cell, 92, 29. Gudkov, A. T. & Gongadze, G. M. (1984). The L7/L12 337–349. proteins change their conformation upon interaction 13. Ogle, J. M., Carter, A. P. & Ramakrishnan, V. of EF-G with ribosomes. FEBS Letters, 176,32–36. (2003). Insights into the decoding mechanism from 30. Agrawal, R. K., Heagle, A. B., Penczek, P., Grasucci, recent ribosome structures. Trends Biochem. Sci. 28, R. A. & Frank, J. (1999). EF-G dependent GTP 259–266. hydrolysis induces translocation accompanied by 14. Sanyal, S. C. & Liljas, A. (2000). The end of the large conformational changes in the 70S ribosome. beginning: structural studies of ribosomal proteins. Nature Struct. Biol. 6, 643–647. Curr. Opin. Struct. Biol. 10, 633–636. 31. Stark, H., Rodnina, M. V., Rinke-Appel, J., Brima- 15. Mohr, D., Wintermeyer, W. & Rodnina, M. V. (2002). combe, R., Wintermeyer, W. & van Heel, M. (1997). GTPase activation of elongation factors Tu and G on Visualization of elongation factor Tu on the Escherichia the ribosome. Biochemistry, 41, 12520–12528. coli ribosome. Nature, 389, 403–406. 16. Diaconu, M., Kothe, U., Schlunzen, F., Fischer, N., 32. Valle, M., Zavialov, A., Li, W., Stagg, S. M., Sengupta, Harms, J. M., Tonevitsky, A. G. et al. (2005). Structural J.,Nielsen,R.C.et al. (2003). Incorporation of basis for the function of the ribosomal L7/12 stalk in aminoacyl-tRNA into the ribosome as seen by cryo- factor binding and GTPase activation. Cell, 121, electron microscopy. Nature Struct. Biol. 10, 899–906. 991–1004. 33. Traut, R. R., Oleinikov, A. V., Makarov, E., Jokhadze, 17. Savelsbergh, A., Mohr, D., Kothe, U., Wintermeyer, W. G., Perroud, B. & Wang, B. (1993). Structure and & Rodnina, M. V. (2005). Control of phosphate release function of Escherichia coli ribosomal protein L7/L12: from elongation factor G by ribosomal protein L7/12. effect of cross-links and deletions. In The Translational EMBO J. 24, 4316–4323. Apparatus: Structure, Function, Evolution (Nierhaus, 18. Wahl, M. C. & Möller, W. (2002). Structure and K. H., Franceschi, F., Subramanian, A. R., Erdmann, function of the acidic ribosomal stalk proteins. Curr. V. A. & Wittmann-Liebold, B., eds), pp. 521–532, Protein Pept. Sci. 3,93–106. Plenum Publishing Corp., New York. 19. Krokowski, D., Tchorzewski, M., Boguszewska, A. & 34. Olson, H. M., Sommer, A., Tewari, D. S., Traut, R. R. & Grankowski, N. (2005). Acquisition of a stable Glitz, D. G. (1986). Localization of two epitopes of structure by yeast ribosomal PO protein requires protein L7/L12 to both the body and stalk of the large binding of P1A-P2B complex: in vitro formation of the ribosomal subunit. Immune electron microscopy using stalk structure. Biochim. Biophys. Acta, 1724,59–70. monoclonal antibodies. J. Biol. Chem. 261, 6924–6932. 20. Remacha, M., JimenezDiaz, A., Santos, C., Briones, E., 35. Dey, D., Bochkariov, D. E., Jokhadze, G. & Traut, R. R. Zambrano, R., Gabriel, M. A. R. et al. (1995). Proteins (1998). Cross-linking of selected residues in the N- and P1, P2, and P0, components of the eukaryotic C-terminal domains of Escherichia coli protein L7/L12 ribosome stalk. New structural and functional aspects. to other ribosomal proteins and the effect of elonga- Biochem. Cell Biol. 73, 959–968. tion factor Tu. J. Biol. Chem. 273, 1670–1676. 21. Bocharov, E. V., Sobol, A. G., Pavlov, K. V., Korzhnev, 36. Savelsbergh, A., Mohr, D., Wilden, B., Wintermeyer, D. M., Jaravine, V. A., Gudkov, A. T. & Arseniev, A. S. W. & Rodnina, M. V. (2000). Stimulation of GTPase (2004). From structure and dynamics of protein L7/ activity of translation elongation factor G by riboso- L12 to molecular switching in ribosome. J. Biol. Chem. mal protein L7/L12. J. Biol. Chem. 275, 890–894. 279, 17697–17706. 37. Datta, P. P., Sharma, M. R., Qi, L., Frank, J. & Agrawal, 22. Mulder, F. A. A., Bouakaz, L., Lundell, A., Venkata- R. K. (2005). Interaction of the G' domain of elongation ramana, M., Liljas, A., Akke, M. & Sanyal, S. (2004). factor G and the C-terminal domain of ribosomal Conformation and dynamics of ribosomal stalk protein L7/L12 during translocation as revealed by protein L12 in solution and on the ribosome. cryo-EM. Mol. Cell, 20, 723–731. Biochemistry, 43, 5930–5936. 38. Kothe, U., Wieden, H. J., Mohr, D. & Rodnina, M. V. 23. Christodoulou, J., Larsson, G., Fucini, P., Connell, (2004). Interaction of helix D of elongation factor Tu S. R., Pertinhez, T. A., Hanson, C. L. et al. (2004). with helices 4 and 5 of protein L7/12 on the ribosome. Heteronuclear NMR investigations of dynamic re- J. Mol. Biol. 336, 1011–1021. L12 binds to IF2, EF-Tu, EF-G, and RF3 479
39. Kawashima, T., BerthetColominas, C., Wulff, M., 47. Zavialov, A. V., Buckingham, R. H. & Ehrenberg, M. Cusack, S. & Leberman, R. (1996). The structure of (2001). A posttermination ribosomal complex is the the Escherichia coli EF-Tu center dot EF-Ts complex at guanine nucleotide exchange factor for peptide re- 2.5 angstrom resolution. Nature, 379,511–518. lease factor RF3. Cell, 107,115–124. 40. Agrawal, R. K., Penczek, P., Grassucci, R. & Frank, J. 48. Forster, A. C., Cornish, V. W. & Blacklow, S. C. (2001). (1998). Visualization of elongation factor G on the Pure translation display. Anal. Biochem. 333, 358–364. Escherichia coli 70S ribosome: the mechanism of 49. Mortensen, K. K., Hansen, H. F., Grentzmann, G., translocation. Proc. Natl Acad. Sci. USA, 95, 6134–6138. Buckingham, R. H. & Sperling Petersen, H. U. (1995). 41. Stark, H., Rodnina, M. V., Wieden, H. J., van Heel, M. Osmo-expression and fast two-step purification of & Wintermeyer, W. (2000). Large-scale movement of Escherichia coli translation termination factor RF-3. Eur. elongation factor G and extensive conformational J. Biochem. 234, 732–736. change of the ribosome during translocation. Cell, 100, 50. Bocharov, E. V., Gudkov, A. T. & Arseniev, A. S. (1996). 301–309. Topology of the secondary structure elements of 42. Klaholz, B. P., Myasnikov, A. G. & van Heel, M. (2004). ribosomal protein L7/L12 from E. coli in solution. Visualization of release factor 3 on the ribosome FEBS Letters, 379, 291–294. during termination of protein synthesis. Nature, 427, 51. Bocharov, E. V., Gudkov, A. T., Budovskaya, E. V. & 862–865. Arseniev, A. S. (1998). Conformational independence 43. Linse, S., Jönsson, B. & Chazin, W. J. (1995). The effect of N- and C-domains in ribosomal protein L7/L12 of protein concentration on ion binding. Proc. Natl and in the complex with protein L10. FEBS Letters, Acad. Sci. USA, 92, 4748–4752. 423, 347–350. 44. Kreishman-Deitrick, M., Egile, C., Hoyt, D. W., Ford, 52. Delaglio, F., Grzesiek, S., Vuister, G. W., Zhu, G., J. J., Li, R. & Rosen, M. K. (2003). NMR analysis of Pfeifer, J. & Bax, A. (1995). NMRPipe: a multidimen- methyl groups at 100-500 kDa: model systems and sional spectral processing system based on UNIX Arp2/3 complex. Biochemistry, 42, 8579–8586. pipes. J. Biomol. NMR, 6, 277–293. 45. Jelenc, P. C. & Kurland, C. G. (1979). Nucleoside 53. Vranken, W. F., Boucher, W., Stevens, T. J., Fogh, triphosphate regeneration decreases the frequency R. H., Pajon, A., Llinas, P. et al. (2005). The CCPN of translation errors. Proc. Natl Acad. Sci. USA, 76, data model for NMR spectroscopy: development of 3174–3178. a software pipeline. Proteins: Struct. Funct. Genet. 59, 46. Wagner, A., Simon, I., Sprinzl, M. & Goody, R. S. 687–696. (1995). Interaction of guanosine nucleotides and their 54. Koradi, R., Billeter, M. & Wüthrich, K. (1996). analogs with elongation-factor Tu from Thermus MOLMOL: a program for display and analysis of thermophilus. Biochemistry, 34, 12535–12542. macromolecular structures. J. Mol. Graph. 14,51–55.
Edited by J. Doudna
(Received 4 July 2006; received in revised form 27 September 2006; accepted 9 October 2006) Available online 13 October 2006