Abstract
The assembly of 30S ribosomes requires the precise addition of 20 proteins to the 16S ribosomal RNA. How early binding proteins change the ribosomal RNA structure so that later proteins may join the complex is poorly understood. Here we use single-molecule fluorescence resonance energy transfer (FRET) to observe real-time encounters between Escherichia coli ribosomal protein S4 and the 16S 5′ domain RNA at an early stage of 30S assembly. Dynamic initial S4–RNA complexes pass through a stable non-native intermediate before converting to the native complex, showing that non-native structures can offer a low free-energy path to protein–RNA recognition. Three-colour FRET and molecular dynamics simulations reveal how S4 changes the frequency and direction of RNA helix motions, guiding a conformational switch that enforces the hierarchy of protein addition. These protein-guided dynamics offer an alternative explanation for induced fit in RNA–protein complexes.
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Acknowledgements
This work was supported by grants from the National Institutes of Health (R01 GM60819 to S.A.W.; R01 GM65367 to T.H.) and from the National Science Foundation (NSF) (PHY0822613 to T.H. and MCB12-44570 to Z. L.-S.). Supercomputer computing time was provided by NSF XSEDE (TG-MCA03S027). T.H. is an investigator with the Howard Hughes Medical Institute.
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H.K., S.C.A., Z.L.-S., T.H. and S.A.W. designed the research. H.K., S.C.A., K.R. and M.M. conducted experiments, S.C.A. and M.M. provided samples, K.C. performed molecular dynamics simulations, H.K. and K.R. analysed the data and H.K. and S.A.W. wrote the paper with input from other authors.
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Extended data figures and tables
Extended Data Figure 1 Modification of the 5′ domain RNA preserves its structure.
a, The secondary structures of the wild-type and extended 5′ domain RNAs were probed by selective 2′-hydroxyl acylation analysed by primer extension (SHAPE)60. The 5′ domain RNA (2 pmol) was annealed to unlabelled oligonucleotides and folded in HKM20 buffer (80 mM K-HEPES, pH 7.6, 300 mM KCl, 6 mM 2-mercaptoethanol, 20 mM MgCl2) before treatment with 3 mM N-methylisatoic acid (NMIA) at 42 °C for 26 min. Modifications were detected by primer extension and quantified as described previously19. Results of SHAPE chemical probing of the free RNA structure for the wild-type 5′ domain (left) and the 5′ domain with h16 and h3 extensions after annealing with oligonucleotides (right). Saturation of grey indicates reactivity with NMIA. Dashed circles indicate nucleotides that were not detected in our primer extension assay. The results show that the extensions added for the fluorescent labelling of the rRNA do not significantly perturb the rRNA folding. b, Native PAGE folding assay of 5′dom-h3h16. Fluorescently labelled oligonucleotide (h3P5–Cy5; 25 nM) was annealed to an equimolar concentration of extended 5′ domain RNA in 10 μl CE buffer (20 mM Na-cacodylate, 0.5 mM Na2EDTA) for 5 min at 70 °C and 5 min at 25 °C. The RNA–oligonucleotide complex was then folded at 37 °C for 30 min in varying [MgCl2] (0–20 mM) before electrophoresis on a native 8% polyacrylamide gel containing 10 mM MgCl2. The folding midpoint was 0.5 ± 0.1 mM MgCl2, similar to that of the wild-type 5′ domain RNA (1.4 ± 0.2 mM) reported previously61.
Extended Data Figure 2 Annealing of labelled oligonucleotides to the 5′ domain RNA.
a, b, 32P-labelled oligonucleotides were annealed to the 3′ extension of h3 of 5′dom-h3 (h3P5, DNA) (a) or to the extended loop of h16 of 5′dom-h3h16 (h16P2-2, RNA) (b). Annealing reactions were performed in 80 mM K-HEPES pH 7.6, 330 mM KCl, 6 mM 2-mercaptoethanol at 25 °C. Binding data were fit to the quadratic form of a two-state binding isotherm. Apparent dissociation constants were ≤0.4 nM and 2.6 ± 0.2 nM for h3 and h16 oligonucleotides, respectively. Equilibrium constants are the average and s.d. of two or more independent trials. The lengths of the labelled oligonucleotides were varied to optimize affinity with the extended 5′ domain RNA, while avoiding perturbations to S4 binding (see Extended Data Fig. 1).
Extended Data Figure 3 S4 labelling and its binding to the rRNA.
E. coli ribosomal protein S4 was overexpressed, purified and labelled with Cy3 or Cy5 fluorescent dyes as described in Methods. a, SDS–PAGE of unlabelled protein stained with Coomassie (left) or labelled with Cy5 (right). b, c, Ensemble titration of the modified 5′ domain RNAs in HKM20 buffer shows that S4–Cy5 binds with similar affinity as the wild-type S4–rRNA complex. Extended 5′ domain RNAs annealed with h3P5–Cy3 and/or h16 oligonucleotides were titrated with S4–Cy5 in a 500-μl cuvette, and the fluorescence emission was recorded from 550 to 700 nm with 540-nm excitation (Fluorolog-3, Horiba). Excitation and emission slits were fixed at 2 nm and 5 nm, respectively. The sample was incubated at 37 °C for 1 min after each addition. Two or more independent measurements were averaged and titration curves were fitted to a quadratic binding expression. Equilibrium dissociation constants were 5′dom-h3, 0.11 ± 0.02 nM (statistical error of the fit parameter) and 5′dom-h3h16, 0.2 ± 0.1 nM, at 37 °C, and were comparable to that of the 5′ domain RNA with wild-type E. coli S4 (0.9 nM)17.
Extended Data Figure 4 Exchange kinetics of docked and flipped complexes.
a, Sample FRET traces are shown for mutant rRNAs in 20 mM Mg2+. b, The cumulative histograms of the dwell times in the high- and low-FRET conformations were calculated for wild-type 5′ domain RNA, C507G mutant and Δh18loop mutant, and fit with both mono-exponential and bi-exponential decay functions. One of the triplicate sets of data is demonstrated for each (refer to Methods for number of traces). Significantly lower χ2 values suggest that the data are best fit with two exponential terms, except for transitions from the low-FRET to the high-FRET state of the wild-type complex, which was well fit by a single exponential decay function. c, d, All of the dwell-time histograms were fit with bi-exponential decay and the fitting parameters were compared between the wild-type and the mutants. Both components of the transition from the high- to low-FRET state were faster in the mutants than in the wild-type complex. The lifetime of the wild-type low-FRET state had a single component; for the mutants we observed an additional slow component in the lifetime.
Extended Data Figure 5 Sample traces of S4 binding and binding trials.
a, In 20 mM Mg2+, the binding occurred mostly at the low-FRET state. Infrequently, we observed the dissociation and secondary binding of S4. b, At 4 mM Mg2+, we often observed unsuccessful and transient binding of S4. Arrows indicate the transient fluorescence signals from unstable binding. These traces also exhibit the mid-FRET spike at the beginning of successful binding events. c, The portion of the molecules that form a stable complex on the first try to the total molecules that form stable complex within 5 min was plotted with varying [Mg2+]. The error bars represent the 95% confidence interval assuming a binary distribution. The number of molecules used was 76, 81, 318 and 170 for 2, 4, 10 and 20 mM Mg2+, respectively.
Extended Data Figure 6 Progression of FRET population at different [Mg2+].
From the synchronized maps of FRET distribution as shown in Fig. 2, the relative populations of low (0–0.35)-, mid (0.35–0.55)- and high (0.55–0.9)-FRET states were plotted at different [Mg2+]. In 20 and 10 mM Mg2+, the bound complexes started with large low-FRET population that converted to high-FRET population within 5–10 s. At 4 and 2 mM Mg2+, there were considerable mid- and high-FRET populations in the beginning, reflecting the broad initial FRET distribution (Fig. 2e, f). This quickly converted to the low-FRET population, which was then followed by slow conversion to the high-FRET population within several seconds. The number of molecules used was 112, 239, 116 and 275 for 20, 10, 4 and 2 mM Mg2+, respectively.
Extended Data Figure 7 Sample traces at 10-ms time resolution.
Single-molecule traces at higher time resolution demonstrate the heterogeneous and fluctuating behaviour of the encounter complex (S4–Cy3 and 5′dom-h3–Cy5). The change of FRET in different molecules cannot be described as a single behaviour. In general, the initial FRET distribution over the complexes is broad and the FRET signal converges to the relatively stable low FRET before the transition to the high FRET.
Extended Data Figure 8 Dynamics of free 5′dom-h3h16.
a, Schematic of extensions in h16 and h3 with labelled oligonucleotides. b, Sample FRET traces showing the fluctuation between two distinct states. The frequency and distribution of these fluctuations varied between molecules. c, Mg2+ dependence of the molecular heterogeneity. Histograms show the relative high-FRET population for each molecule (157, 162 and 74 traces for 20, 5 and 1 mM Mg2+, respectively). At higher [Mg2+], more molecules stay in the high-FRET state for longer periods of time. High FRET between h16 and h3 does not necessarily correspond to the native structure of the 5′ domain RNA in complex with protein S4 that is represented by high FRET between S4 and h3.
Extended Data Figure 9 Examples of switching between different h16–h3 dynamics.
Arrows indicate when the fluctuation dynamics switch between stable high FRET, stable low FRET and alternating high- and low-FRET behaviours. The RNA was labelled as in Extended Data Fig. 8.
Extended Data Figure 10 S4 binding trajectories from hybrid MD-Gō simulations.
a, Simulated FRET between S4 and h3 from representative binding trajectories displays various binding pathways. b, Density map constructed from 61 successful binding trajectories. The trajectories were synchronized at the moment when the first native contact between S4 and the 5WJ is established (dotted lines in a). c, Sample trajectories of successful binding, showing how folding of h16 and h3 is induced by S4 binding.
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Kim, H., Abeysirigunawarden, S., Chen, K. et al. Protein-guided RNA dynamics during early ribosome assembly. Nature 506, 334–338 (2014). https://doi.org/10.1038/nature13039
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DOI: https://doi.org/10.1038/nature13039
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