Functional Interactions between the G′ Subdomain of Bacterial Translation Factor EF-G and Ribosomal Protein L7/L12
2007; Elsevier BV; Volume: 282; Issue: 51 Linguagem: Inglês
10.1074/jbc.m707179200
ISSN1083-351X
AutoresRoxana Nechifor, Marat B. Murataliev, Kevin S. Wilson,
Tópico(s)Bacterial Genetics and Biotechnology
ResumoProtein L7/L12 of the bacterial ribosome plays an important role in activating the GTP hydrolytic activity of elongation factor G (EF-G), which promotes ribosomal translocation during protein synthesis. Previously, we cross-linked L7/L12 from two residues (209 and 231) flanking α-helix AG′ in the G′ subdomain of Escherichia coli EF-G. Here we report kinetic studies on the functional effects of mutating three neighboring glutamic acid residues (224, 228, and 231) to lysine, either singly or in combination. Two single mutations (E224K and E228K), both within helix AG′, caused large defects in GTP hydrolysis and smaller defects in ribosomal translocation. Removal of L7/L12 from the ribosome strongly reduced the activities of wild type EF-G but had no effect on the activities of the E224K and E228K mutants. Together, these results provide evidence for functionally important interactions between helix AG′ of EF-G and L7/L12 of the ribosome. Protein L7/L12 of the bacterial ribosome plays an important role in activating the GTP hydrolytic activity of elongation factor G (EF-G), which promotes ribosomal translocation during protein synthesis. Previously, we cross-linked L7/L12 from two residues (209 and 231) flanking α-helix AG′ in the G′ subdomain of Escherichia coli EF-G. Here we report kinetic studies on the functional effects of mutating three neighboring glutamic acid residues (224, 228, and 231) to lysine, either singly or in combination. Two single mutations (E224K and E228K), both within helix AG′, caused large defects in GTP hydrolysis and smaller defects in ribosomal translocation. Removal of L7/L12 from the ribosome strongly reduced the activities of wild type EF-G but had no effect on the activities of the E224K and E228K mutants. Together, these results provide evidence for functionally important interactions between helix AG′ of EF-G and L7/L12 of the ribosome. Elongation factors (EF) 3The abbreviations used are:EFelongation factorCTDC-terminal domainmant-GTP2′,3′-O-(N′-methylanthraniloyl)-GTPTKtriple lysine mutantPiinorganic phosphate. 3The abbreviations used are:EFelongation factorCTDC-terminal domainmant-GTP2′,3′-O-(N′-methylanthraniloyl)-GTPTKtriple lysine mutantPiinorganic phosphate. Tu and G are the bacterial counterparts of universal translation factors, members of the G protein superfamily (1Bourne H.R. Sanders D.A. McCormick F. Nature. 1991; 349: 117-127Crossref PubMed Scopus (2683) Google Scholar), which hydrolyze GTP to GDP and inorganic phosphate (Pi). The ribosome activates the latent GTPase activities of EF-Tu and EF-G. These factors, in turn, regulate protein synthesis by promoting specific molecular movements in the ribosome during protein synthesis. EF-Tu delivers aminoacyl-tRNA substrates to the ribosome, dependent on codon-anticodon pairing. EF-G promotes ribosomal translocation, involving movement of two tRNAs and mRNA in the ribosomal cavity, following the formation of each peptide bond. elongation factor C-terminal domain 2′,3′-O-(N′-methylanthraniloyl)-GTP triple lysine mutant inorganic phosphate. elongation factor C-terminal domain 2′,3′-O-(N′-methylanthraniloyl)-GTP triple lysine mutant inorganic phosphate. GTP hydrolysis by EF-Tu and EF-G occurs on their G domains, which are similar to one another in their amino acid sequences and their core tertiary structures. The hydrolysis reaction involves in-line nucleophilic attack of a water molecule on the γ-phosphorus of GTP (2Eccleston J.F. Webb M.R. J. Biol. Chem. 1982; 257: 5046-5049Abstract Full Text PDF PubMed Google Scholar). On EF-Tu (and possibly also EF-G), this reaction is catalyzed by a conserved histidine residue (3Daviter T. Wieden H.J. Rodnina M.V. J. Mol. Biol. 2003; 332: 689-699Crossref PubMed Scopus (122) Google Scholar), whose side chain is believed to rotate to a position next to the water molecule (4Berchtold H. Reshetnikova L. Reiser C.O. Schirmer N.K. Sprinzl M. Hilgenfeld R. Nature. 1993; 365: 126-132Crossref PubMed Scopus (514) Google Scholar). In contrast to EF-Tu, EF-G contains a G′ subdomain, which is invariably inserted between α-helices DG and EG (5Czworkowski J. Wang J. Steitz T.A. Moore P.B. EMBO J. 1994; 13: 3661-3668Crossref PubMed Scopus (358) Google Scholar). The G′ subdomain is also present in the same location in EF-2, the eukaryotic cytoplasmic homolog of EF-G (6Jorgensen R. Ortiz P.A. Carr-Schmid A. Nissen P. Kinzy T.G. Andersen G.R. Nat. Struct. Biol. 2003; 10: 379-385Crossref PubMed Scopus (136) Google Scholar). However, the G′ subdomains of EF-G and EF-2 are unrelated in their amino acid sequences and tertiary structures. The functional significance of the G′ subdomain in either factor has not been determined. How the bacterial ribosome activates GTP hydrolysis by EF-Tu and EF-G remains obscure. Early research identified L7/L12, 4L7 and L12 refer to the N-acetylated and unacetylated forms of the same protein. 4L7 and L12 refer to the N-acetylated and unacetylated forms of the same protein. a protein component of one of the peripheral stalks of the ribosome, as an important contributor to GTPase activation of both factors (7Kischa K. Moller W. Stoffler G. Nat. New Biol. 1971; 233: 62-63Crossref PubMed Scopus (103) Google Scholar, 8Hamel E. Koka M. Nakamoto T. J. Biol. Chem. 1972; 247: 805-814Abstract Full Text PDF PubMed Google Scholar). More recent studies identified residues of the C-terminal domain (CTD) of L7/L12, important for GTPase activation and rapid association of both factors with the ribosome (9Kothe U. Wieden H.J. Mohr D. Rodnina M.V. J. Mol. Biol. 2004; 336: 1011-1021Crossref PubMed Scopus (60) Google Scholar, 10Diaconu M. Kothe U. Schlunzen F. Fischer N. Harms J.M. Tonevitsky A.G. Stark H. Rodnina M.V. Wahl M.C. Cell. 2005; 121: 991-1004Abstract Full Text Full Text PDF PubMed Scopus (310) Google Scholar). Nucleotides of 23 S RNA (11Cundliffe E. Hill W.E. Dahlberg A. Garrett R.A. Moore P.B. Schlessinger D. Warner J.R. The Ribosome: Structure, Function, and Evolution. American Society for Microbiology, Washington, D. C.1989: 479-490Google Scholar) and unidentified ribosomal components (12Mohr D. Wintermeyer W. Rodnina M.V. Biochemistry. 2002; 41: 12520-12528Crossref PubMed Scopus (126) Google Scholar) have also been implicated in GTPase activation of these factors. The sites on EF-Tu and EF-G that interact with L7/L12 are only beginning to be elucidated. For EF-Tu, mutational studies identified negatively charged residues in its helix DG that may interact with positively charged residues of L7/L12 (9Kothe U. Wieden H.J. Mohr D. Rodnina M.V. J. Mol. Biol. 2004; 336: 1011-1021Crossref PubMed Scopus (60) Google Scholar). For EF-G, no functional studies of this type have been reported so far. However, two topographic studies indicated that the G′ subdomain of EF-G is proximal to L7/L12. First, residues 209 and 231, flanking helix AG′, of Escherichia coli EF-G were cross-linked to L7/L12 (13Nechifor R. Wilson K.S. J. Mol. Biol. 2007; 368: 1412-1425Crossref PubMed Scopus (18) Google Scholar). In the same study, no cross-links were detected from two other EF-G residues (156 and 160) of helix DG, whose sequence is completely different from the corresponding helix of EF-Tu. Second, a cryo-EM study localized EF-G residue 209 near the base of the ribosomal stalk, which was interpreted as an interaction with the CTD of L7/L12 (14Datta P.P. Sharma M.R. Qi L. Frank J. Agrawal R.K. Mol. Cell. 2005; 20: 723-731Abstract Full Text Full Text PDF PubMed Scopus (71) Google Scholar). This study was undertaken to investigate the potential functional roles of the G′ subdomain of EF-G. We present results from kinetic experiments that provide evidence for interactions between residues of helix AG′ of EF-G and L7/L12, which are important for activating GTP hydrolysis and ribosomal translocation. Materials—E. coli 70 S ribosome and phage T4 gene 32 mRNA were prepared as described (15Wilson K.S. Nechifor R. J. Mol. Biol. 2004; 337: 15-30Crossref PubMed Scopus (30) Google Scholar). L7/L12 was specifically stripped from the ribosome by an ethanol-NH4Cl washing procedure (12Mohr D. Wintermeyer W. Rodnina M.V. Biochemistry. 2002; 41: 12520-12528Crossref PubMed Scopus (126) Google Scholar). The extent of L7/L12 stripping was assessed by immunoblotting using a L7/L12 polyclonal antibody (13Nechifor R. Wilson K.S. J. Mol. Biol. 2007; 368: 1412-1425Crossref PubMed Scopus (18) Google Scholar). E. coli tRNAfMet and tRNAPhe were purchased from Sigma. The mRNA, 5′-AAGGAGGUAAAAAUGUUUGCU(N6)-3′, was synthesized by Dharmacon and conjugated at its 3′ end with pyrene as described (16Studer S.M. Feinberg J.S. Joseph S. J. Mol. Biol. 2003; 327: 369-381Crossref PubMed Scopus (97) Google Scholar). Mant-labeled nucleotides were purchased from Invitrogen. Buffers—Buffer 1 consists of 50 mm Tris-HCl (pH 8.0), 100 mm KCl, 10 mm MgCl2, 0.1 mm EDTA, 1 mm phenylmethylsulfonyl fluoride, 10 mm β-mercaptoethanol in water. Buffer 2 consists of 80 mm HEPES-KOH (pH 7.7), 50 mm NH4Cl, 10 mm MgCl2, 1 mm dithiothreitol in water. Buffer 3 consists of 50 mm HEPES-KOH (pH 7.7), 100 mm NH4Cl, 20 mm MgCl2, 1 mm dithiothreitol in water. The AlF4- complex consists of 100 μm AlCl3, 10 mm NaF in buffer 2. GTP Binding and Hydrolysis Assays—All fluorescence measurements described below were made by a QM-6 fluorimeter (Photon Technology International). Binding of mant-GTP to EF-G was measured by titrating mant-GTP (1 μm) in 2 ml of buffer 2 with EF-G (0-15 μm). After adding each EF-G aliquot, mixing with a stir bar, and allowing the solution to reach equilibrium, fluorescence (excitation 362 ± 2 nm; emission 438 ± 4 nm) was recorded. Multiple-turnover GTP hydrolysis was assayed as described (13Nechifor R. Wilson K.S. J. Mol. Biol. 2007; 368: 1412-1425Crossref PubMed Scopus (18) Google Scholar), with the following modifications. EF-G (0.04 μm) and ribosomes (variable concentration) were preincubated (37 °C, 10 min) in 9 μl of buffer 2. Reactions (10 μl) were started by adding 1 μl of 500 μm GTP (containing 0.05 μCi of [γ-32P]GTP). Samples (2 μl) were withdrawn after appropriate time intervals, during the linear kinetics of the reaction. Samples were quenched and analyzed by TLC as described (13Nechifor R. Wilson K.S. J. Mol. Biol. 2007; 368: 1412-1425Crossref PubMed Scopus (18) Google Scholar). Data of GTP molecules hydrolyzed per EF-G molecule/s (vo) as a function of ribosome concentration [R] were calculated and fitted (via SigmaPlot2000 software) to: vo = kcat × [EF-G] × [R]/(KMR + [R]). Multiple-turnover mant-GTP hydrolysis was detected by the fluorescence change between mant-GTP and mant-GDP bound to EF-G. The fluorescence of mant-GTP (120 μm) in 250 μl of buffer 2 was monitored by exciting the fluorophore at 362 nm and detecting its emission at 438 nm. EF-G (25 μm) and vacant ribosomes (0.7 μm) were added in successive steps and mixed by pipetting. Samples (20 μl) were removed from the cuvette and precipitated with 0.2 m HCl. The supernatants of the samples were concentrated in a vacuum centrifuge to ∼8 μl, and 2 μl of each sample (∼500 pmol) was analyzed by TLC (17Wilden B. Savelsbergh A. Rodnina M.V. Wintermeyer W. Proc. Natl. Acad. Sci. U. S. A. 2006; 103: 13670-13675Crossref PubMed Scopus (101) Google Scholar). Single-turnover mant-GTP hydrolysis was monitored in a stopped-flow device (MiniMixer, KinTek), which contained two reactant syringes. Syringe A contained 1 ml of 24 μm vacant ribosomes in buffer 2 at ∼22 °C. Syringe B contained 1 ml of 20 μm EF-G and 10 μm mant-GTP (same buffer and temperature). In some experiments, AlF4- complex (100 μm) was included in syringe B. Samples (∼200 μl) from each syringe were rapidly mixed together (dead time of ∼3.5 ms) and injected into a cuvette in our fluorimeter. Fluorescence (excitation 362 ± 2 nm; emission 438 nm ± 4) was monitored over 40-400 s, with measurements taken every 50-1000 ms, depending on the rate constant of the EF-G protein being tested. Data for each reaction were fitted to a single exponential equation, Ft = F∞ + ΔFmax × exp(-kobs × t), where F0 is the initial fluorescence at time = 0; Ft is the fluorescence at time t; ΔFmax is the maximum fluorescence change (F0 - F∞), and kobs is the observed reaction rate constant. Ribosomal Translocation Assays—Multiple-turnover ribosomal translocation was monitored by the toeprinting method (18Hartz D. McPheeters D.S. Traut R. Gold L. Methods Enzymol. 1988; 164: 419-425Crossref PubMed Scopus (325) Google Scholar). A pretranslocation complex (1 μm) was formed in 90 μl of buffer 3 with E. coli ribosomes (1 μm) containing phage T4 gene 32 mRNA (0.8 μm) and uncharged tRNAfMet (1.2 μm) and tRNAPhe (1.2 μm) bound in the P and A sites, respectively, of the ribosome (15Wilson K.S. Nechifor R. J. Mol. Biol. 2004; 337: 15-30Crossref PubMed Scopus (30) Google Scholar). EF-G (0.1 μm) and GTP (500 μm) were added, and reactions were incubated at 37 °C. Samples (10 μl) were removed after various time intervals (as indicated) and analyzed by toeprinting (15Wilson K.S. Nechifor R. J. Mol. Biol. 2004; 337: 15-30Crossref PubMed Scopus (30) Google Scholar). Single-turnover ribosomal translocation was monitored by the change in fluorescence of mRNA labeled with pyrene at its 3′ end (16Studer S.M. Feinberg J.S. Joseph S. J. Mol. Biol. 2003; 327: 369-381Crossref PubMed Scopus (97) Google Scholar). A pretranslocation complex (0.5 μm) was formed in 5 ml of buffer 3, similar to the above complex except substituting pyrene-mRNA (0.4 μm). This complex was divided into five equal aliquots, which were loaded successively into syringe A of the stopped-flow device. Syringe B contained the various EF-G proteins (1.25-12 μm) and GTP (2 mm) in 1.2 ml of buffer 3. Reactions were started by rapidly mixing 200 μl samples from each syringe. Fluorescence (excitation 332 ± 4 nm; emission 377 ± 8 nm) was monitored over 200-300 s, with averaged measurements taken every 60-200 ms, depending on the rate constant of the EF-G protein being tested. Data were fitted to a double exponential equation: Ft = F∞ +ΔF1 × exp(-k1 × t) + ΔF2 × exp(-k2 × t). Observed translocation rates as a function of EF-G concentration were fitted to Reaction 1. A1+B1k1[A1]⇄k-1A1·B1k2→A2·B1k3→A2·B2REACTION 1 where A1 = EF-G·GTP; A2 = EF-G·GDP; B1 = pretranslocational complex; B2 = post-translocational complex; kobs = A1/(C1 × A1 + C2); C1 = (k2 + k3)/k2 × k3; and C2 = (k-1 + k2)/k1 × k2. To investigate the function of the G′ subdomain, we first genetically replaced the entire subdomain of E. coli EF-G (residues 166-261) with a single Gly residue. The N- and C-terminal ends of the G′ subdomain are closely juxtaposed in the x-ray crystal structures of Thermus thermophilus EF-G proteins (5Czworkowski J. Wang J. Steitz T.A. Moore P.B. EMBO J. 1994; 13: 3661-3668Crossref PubMed Scopus (358) Google Scholar, 19Connell S.R. Takemoto C. Wilson D.N. Wang H. Murayama K. Terada T. Shirouzu M. Rost M. Schuler M. Giesebrecht J. Dabrowski M. Mielke T. Fcini P. Yokoyama S. Spahn C.M. Mol. Cell. 2007; 25: 751-764Abstract Full Text Full Text PDF PubMed Scopus (155) Google Scholar), suggesting that this deletion would not perturb the folding of the rest of EF-G. We introduced this genetic deletion into a plasmid encoding E. coli EF-G with a C-terminal His6 tag (15Wilson K.S. Nechifor R. J. Mol. Biol. 2004; 337: 15-30Crossref PubMed Scopus (30) Google Scholar). Although the mutant protein could be produced at high levels in E. coli, it aggregated in cellular inclusion bodies, and we were unable to refold it after purification under denaturing conditions. We then replaced single amino acid residues of the G′ subdomain. In selecting these mutations, we considered three sources of information as follows: (i) our previous cross-linking results indicating the proximity of L7/L12 to EF-G residues 209 and 231, which flank helix AG′ (13Nechifor R. Wilson K.S. J. Mol. Biol. 2007; 368: 1412-1425Crossref PubMed Scopus (18) Google Scholar); (ii) an alignment of the amino acid sequences of the G′ subdomain from bacterial and mitochondrial phyla (supplemental Fig. 1); and (iii) a study identifying conserved, positively charged residues of L7/L12 important for EF-G GTPase activity (10Diaconu M. Kothe U. Schlunzen F. Fischer N. Harms J.M. Tonevitsky A.G. Stark H. Rodnina M.V. Wahl M.C. Cell. 2005; 121: 991-1004Abstract Full Text Full Text PDF PubMed Scopus (310) Google Scholar), suggesting that L7/L12 interacts with conserved, negatively charged residues of EF-G. The G′ subdomain is generally poorly conserved in relation to other EF-G domains. We chose to mutate the three most conserved, negatively charged residues corresponding to Glu-224, Glu-228, and Glu-231 of E. coli EF-G. In the structures of T. thermophilus EF-G proteins, residues equivalent to Glu-224 and Glu-228 are located within helix AG′, and the residue equivalent to Glu-231 is at the N terminus of helix BG′ (Fig. 1). The side chains of all three residues are exposed to solvent. We replaced these Glu residues with oppositely charged Lys residues, either individually or in combination, hoping that these mutations would repel interactions with L7/L12. The His6-tagged EF-G proteins (four G′ mutants and wild type) were produced and purified under native conditions (15Wilson K.S. Nechifor R. J. Mol. Biol. 2004; 337: 15-30Crossref PubMed Scopus (30) Google Scholar). G′ Mutations Hinder Ribosome-activated GTP Hydrolysis—We first examined the GTP hydrolysis activities of the purified EF-G proteins, activated by vacant ribosomes (lacking tRNA and mRNA). Experiments were performed under multiple-turnover conditions of fixed EF-G concentration (0.04 μm), saturating GTP concentration (500 μm), and increasing ribosome concentration. As shown in Fig. 2A, the initial velocity (vo) of GTP hydrolysis catalyzed by wild type EF-G increased with increasing ribosome concentrations up to ∼2 μm, followed by a drop in vo at higher ribosome concentrations, because of turnover inhibition effects as reported previously (20Rohrback M.S. Bodley J.W. Biochemistry. 1976; 15: 4565-4569Crossref PubMed Scopus (27) Google Scholar). Based on fitting the Michaelis-Menten equation to the data up to 2 μm ribosome, wild type EF-G and the E231K mutant displayed nearly the same turnover (kcat = 9.9 and 7.7 s-1, respectively) and apparent ribosome binding (Km = 0.91 and 1.5 μm). In contrast, the other three G′ mutants, E224K, E228K, and the triple Lys mutant (TK), were severely defective, to the extent that their vo values were not measurable at any ribosome concentration above a control reaction containing only the ribosome. The latter three G′ mutants were specifically defective in ribosome-activated GTP hydrolysis. In the absence of the ribosome, the basal GTPase activities of all four G′ mutants varied between ∼3 × 10-4 and 3 × 10-3 s-1, slightly faster than ∼4 × 10-4 s-1 for the wild type protein, at the same EF-G and GTP concentrations indicated above (data not shown). Under multiple-turnover conditions, the defects in ribosome-activated GTP hydrolysis could, in principle, be manifested at one or several kinetic steps in the uncoupled GTPase reaction cycle (20Rohrback M.S. Bodley J.W. Biochemistry. 1976; 15: 4565-4569Crossref PubMed Scopus (27) Google Scholar). We could exclude effects on the initial binding of GTP to EF-G, as all four G′ mutants bound a fluorescent substrate derivative, mant-GTP, with affinities (Kd values: 2.2-3.7 μm) that were indistinguishable from the wild type (Fig. 2B). To try to localize the defects of the G′ mutants to specific steps in the reaction cycle, we sought a GTPase kinetic assay performed under single-turnover conditions. Using mant-GTP as a fluorescent reporter, we observed that its fluorescence increased substantially when wild type EF-G was added (Fig. 3A), as reported previously (21Savelsbergh A. Mohr D. Wilden B. Wintermeyer W. Rodnina M.V. J. Biol. Chem. 2000; 275: 890-894Abstract Full Text Full Text PDF PubMed Scopus (77) Google Scholar). When we subsequently added a limiting amount of the ribosome, the fluorescence gradually dropped to a lower plateau. This suggested that mant-GTP was slowly hydrolyzed to mant-GDP, although it was bound to EF-G, activated by a limiting amount of ribosome that was recycled. To test this hypothesis, we removed three samples (S1, S2, and S3) at different time points during the fluorescence decay. These samples were immediately quenched by acid precipitation and subsequently analyzed by TLC (Fig. 3B). According to the mobilities of known molecular markers resolved on the same TLC, S1 (taken after EF-G addition) contained only mant-GTP; S2 (taken after ribosome addition, near the end of the fluorescence decay) contained mostly mant-GDP and small amounts of mant-GTP, and S3 (taken in the fluorescence plateau) contained only mant-GDP. Thus, these experiments clearly show that the fluorescence decrease was because of mant-GTP hydrolysis. The fluorescence decrease of the mant group presumably reflects a conformational change in EF-G. It is unclear whether this occurs during cleavage of the phosphoanhydride bond or during the subsequent release of the Pi product from EF-G. To characterize the mechanism further, we monitored single-turnover mant-GTP hydrolysis by using a stopped flow device that rapidly mixed EF-G·mant-GTP with a stoichiometric amount of the ribosome. We compared reactions in the presence or absence of AlF4-, an analog of the planar transition state structure of the γ-phosphate leaving group, which binds tightly to many G protein·GDP complexes. Under single-turnover conditions, the fluorescence signal decayed exponentially with much faster kinetics and similar in amplitude (Fig. 3C). When AlF4- was included with EF-G·mant-GTP before mixing them with the ribosome, the rapid fluorescence decrease was followed by an increase, which was slower and smaller in amplitude. As a positive control, when EF-G·mant-GDP and AlF4- were mixed with the ribosome, only a larger fluorescence increase was observed. As a negative control, when EF-G·mant-GDP was mixed with the ribosome, a smaller fluorescence increase was observed. Collectively, these results suggest that the initial fluorescence decrease encompasses both mant-GTP hydrolysis and Pi release from EF-G. The subsequent fluorescence increase is because of AlF4- binding to EF-G·mant-GDP, which follows Pi release. The small fluorescence increase seen in the negative control most likely reflects binding of EF-G·mant-GDP to the ribosome. We then applied the above assay to compare the G′ mutants and wild type EF-G proteins in catalyzing mant-GTP hydrolysis (Fig. 4). Under single-turnover conditions, wild type EF-G catalyzed this reaction with kobs of 1.8 s-1 (Table 1). This value is ∼10-fold slower than previously reported for Pi release (20 s-1), and much slower than GTP hydrolysis (80 s-1), as measured by other methods under comparable conditions (22Savelsbergh A. Katunin V.I. Mohr D. Peske F. Rodnina M.V. Wintermeyer W. Mol. Cell. 2003; 11: 1517-1523Abstract Full Text Full Text PDF PubMed Scopus (247) Google Scholar). These comparisons further suggest that the fluorescence decrease is not caused by the hydrolysis reaction itself, but rather because of a slower EF-G conformational change that follows Pi release. Alternatively, the slower rate may arise from the mant group partially interfering with the hydrolysis reaction.TABLE 1Functional effects of stripping L7/L12 from the ribosome Values represent the averages ± S.D. of three or four independent reactions (stopped flow injections).+EF-GMant-GTP hydrolysis, kobsmRNA translocation, kobs+L7/L12–L7/L12+L7/L12–L7/L12s–1s–1Wild type1.8 ± 0.10.149 ± 0.0070.58 ± 0.020.067 ± 0.004E224K0.028 ± 0.0010.027 ± 0.0010.055 ± 0.0030.051 ± 0.006E228K0.027 ± 0.0010.024 ± 0.0010.059 ± 0.0020.050 ± 0.005E231K0.8 ± 0.10.088 ± 0.0030.27 ± 0.010.057 ± 0.006TK0.009 ± 0.0010.022 ± 0.0010.029 ± 0.0010.029 ± 0.001 Open table in a new tab Regardless of the precise explanation, the G′ mutants showed substantially reduced rates of mant-GTP hydrolysis, in comparison to wild type EF-G (Table 1). The most defective single G′ mutants were E224K and E228K, whose kobs values were both 65-fold lower than wild type. The E231K mutant was the most active, with kobs only 2-fold lower than wild type, in accord with the multiple-turnover GTP hydrolysis assay (Fig. 2A). The TK mutant was the most defective of all, with kobs that was 200-fold lower than wild type. However, the TK mutant still displayed significant ribosome-activated GTPase activity, relative to controls of the ribosome alone (Fig. 4) and the basal GTPase activity of the TK mutant in the absence of the ribosome. In summary, results from multiple- and single-turnover assays both identified mutants E224K and E228K as having large defects in GTP hydrolysis. The single-turnover assays provide the more direct and accurate measurements, but they are ambiguous with regard to effects on hydrolysis or Pi release. It is also possible that the apparent defects in mant-GTP hydrolysis are manifested because of an impaired association of G′ mutants with the ribosome. The latter possibility is addressed below. G′ Mutations Reduce Ribosomal Translocation Kinetics—Previous studies have indicated that GTP hydrolysis by EF-G is strongly coupled to ribosomal translocation, the coordinated movement of tRNA and mRNA in the ribosomal cavity. Upon binding of EF-G·GTP to a pretranslocational ribosome complex, GTP hydrolysis occurs rapidly (23Rodnina M.V. Savelsbergh A. Katunin V.I. Wintermeyer W. Nature. 1997; 385: 37-41Crossref PubMed Scopus (389) Google Scholar), which is thought to induce conformational rearrangements in the complex that drive the subsequent steps of Pi release from EF-G·GDP and translocation (22Savelsbergh A. Katunin V.I. Mohr D. Peske F. Rodnina M.V. Wintermeyer W. Mol. Cell. 2003; 11: 1517-1523Abstract Full Text Full Text PDF PubMed Scopus (247) Google Scholar, 24Katunin V.I. Savelsbergh A. Rodnina M.V. Wintermeyer W. Biochemistry. 2002; 41: 12806-12812Crossref PubMed Scopus (89) Google Scholar). Although translocation can occur in the presence of nonhydrolyzable GTP analogs, GTP hydrolysis strongly promotes translocation both kinetically and thermodynamically (24Katunin V.I. Savelsbergh A. Rodnina M.V. Wintermeyer W. Biochemistry. 2002; 41: 12806-12812Crossref PubMed Scopus (89) Google Scholar, 25Pan D. Kirillov S.V. Cooperman B.S. Mol. Cell. 2007; 25: 519-529Abstract Full Text Full Text PDF PubMed Scopus (162) Google Scholar). Given the effects of the G′ mutations on GTP hydrolysis activated by the vacant ribosome, we reasoned that these mutations might also exert effects on translocation if, as in the wild type situation, GTP hydrolysis and translocation remain strongly coupled mechanisms. We assembled a pretranslocational ribosome complex that contained a model mRNA and two tRNAs bound in the A and P sites of the ribosome (15Wilson K.S. Nechifor R. J. Mol. Biol. 2004; 337: 15-30Crossref PubMed Scopus (30) Google Scholar). We first measured translocation in the toeprinting assay, which monitors the position and movement of the ribosome relative to its bound mRNA, as detected by reverse transcription of a DNA primer annealed to the mRNA downstream of the ribosome (18Hartz D. McPheeters D.S. Traut R. Gold L. Methods Enzymol. 1988; 164: 419-425Crossref PubMed Scopus (325) Google Scholar). Because reverse transcription is much slower than ribosomal translocation, we examined translocation reactions under multiple-turnover conditions of limiting EF-G (0.1 μm), saturating GTP (500 μm), and pretranslocational complex (1 μm). Fig. 5 compares ribosomal translocation profiles catalyzed by wild type EF-G and the TK mutant. At the start of reactions, the pretranslocational complex was indicated by two primer extension products, a major one stopping at +17 and minor one at +16, relative to the initiation codon of the mRNA. Following addition of wild type EF-G and GTP, the +17 product was converted rapidly (within the 1st min) to a +19 product, indicating the post-translocational complex. The +16 product was converted more slowly to the same +19 product. Following addition of TK and GTP, both +16 and +17 products were converted to the +19 product at about the same kinetics and much slower than the wild type reaction. The E224K and E228K mutants catalyzed slower translocation similar to the TK mutant, whereas E231K catalyzed faster translocation between the rates of wild type and TK (supplemental Fig. 2). To measure accurately the kinetics of translocation under single-turnover conditions, we took advantage of a fluorescence assay that monitors the movement of pyrene attached to the 3′ end of a short mRNA that just spans the ribosome (16Studer S.M. Feinberg J.S. Joseph S. J. Mol. Biol. 2003; 327: 369-381Crossref PubMed Scopus (97) Google Scholar). A pretranslocational complex (0.5 μm) containing pyrene-labeled mRNA and the same two tRNAs was rapidly mixed with wild type EF-G (1.25 μm) and saturating GTP. The fluorescence decay associated with translocation followed distinctly biphasic kinetics (supplemental Fig. 3). In the first phase of the reaction, kobs was 0.58 s-1, whereas kobs of the second phase was 0.056 s-1. It should be noted that the rate constant of the faster phase is comparable with the original report (16Studer S.M. Feinberg J.S. Joseph S. J. Mol. Biol. 2003; 327: 369-381Crossref PubMed Scopus (97) Google Scholar) but slower than reported in other studies (22Savelsbergh A. Katunin V.I. Mohr D. Peske F. Rodnina M.V. Wintermeyer W. Mol. Cell. 2003; 11: 1517-1523Abstract Full Text Full Text PDF PubMed Scopus (247) Google Scholar, 26Feinberg J.S. Joseph S. RNA (N. Y.). 2006; 12: 580-588Crossref PubMed Scopus (13) Google Scholar). This discrepancy has been recently attributed to the C-te
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