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Structural basis of the non-coding RNA RsmZ acting as a protein sponge


MicroRNA and protein sequestration by non-coding RNAs (ncRNAs) has recently generated much interest. In the bacterial Csr/Rsm system, which is considered to be the most general global post-transcriptional regulatory system responsible for bacterial virulence, ncRNAs such as CsrB or RsmZ activate translation initiation by sequestering homodimeric CsrA-type proteins from the ribosome-binding site of a subset of messenger RNAs. However, the mechanism of ncRNA-mediated protein sequestration is not understood at the molecular level. Here we show for Pseudomonas fluorescens that RsmE protein dimers assemble sequentially, specifically and cooperatively onto the ncRNA RsmZ within a narrow affinity range. This assembly yields two different native ribonucleoprotein structures. Using a powerful combination of nuclear magnetic resonance and electron paramagnetic resonance spectroscopy we elucidate these 70-kilodalton solution structures, thereby revealing the molecular mechanism of the sequestration process and how RsmE binding protects the ncRNA from RNase E degradation. Overall, our findings suggest that RsmZ is well-tuned to sequester, store and release RsmE and therefore can be viewed as an ideal protein ‘sponge’.

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Figure 1: Sequential and specific binding of the RsmE protein to the RsmZ sRNA.
Figure 2: Solution structures of both conformers of the 70-kDa complex between RsmZ(1−72) and three RsmE homodimers.
Figure 3: Fine-tuning of the binding affinity of RsmZ for RsmE results from both negative and positive binding cooperativity.
Figure 4: RsmE progressively protects the sRNA RsmZ from degradation.
Figure 5: The life of the ncRNA RsmZ as a protein sponge.

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Protein Data Bank

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Coordinates, NMR and EPR restraints have been deposited in the Protein Data Bank under accession numbers 2mf0 and 2mf1.


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We thank G. A. Mackie for providing the RNase E (1−529) clone; N. Diarra Dit Konté and B. Alila for the help with RNA production; M. Blatter for help with structure calculations; G. Wider, T. Stahel, C. Maris and F. Damberger for help with NMR spectroscopy setup; G. Braach, Y. Nikolaev and M. Sattler for discussions. This work was supported by the Swiss National Science Foundation (SNF) grant numbers 3100A0-118118, 31003ab-133134 and 31003A-149921 to F.H.-T.A. and the SNF-NCCR structural biology Iso-lab.

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Authors and Affiliations



O.D. prepared protein and RNA samples for NMR and EPR spectroscopy, collected and analysed the NMR experiments, designed the structure calculation protocol and performed structure calculations, conducted and analysed ITC experiments, gel shift and RNase E cleavage assays, designed the project and the experiments and wrote the manuscript. E.M. designed, performed and analysed in vitro translation and E. coli whole-cell extract RNase protection assays. M.Y. measured and analysed EPR data. M.S. performed initial NMR experiments. G.J. analysed EPR data and secured funding. F.H.-T.A. designed and supervised the study, secured funding, and wrote the manuscript. All authors discussed the results, commented on and approved the manuscript.

Corresponding author

Correspondence to Frédéric H.-T. Allain.

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The authors declare no competing financial interests.

Extended data figures and tables

Extended Data Figure 1 General protocol for the structure determination of the full complex structure.

(1) The structures of the separate binding site complexes are solved. (2) Comparing the chemical shifts of the resonances in the isolated binding site complexes and the full complex reveals which parts of the isolated binding site complexes retain their structure in the full complex. Very similar A(N)GGAX binding motifs in all the five binding sites led to severe spectral overlap and thus required segmental labelling of a single binding site on the RsmZ sRNA at a time. (3) The global structure of the full complex is obtained by constraining the different domains with several EPR long-range distances measured between spin-labels attached at several positions on the RsmZ sRNA.

Extended Data Figure 2 Chemical shift comparison of RNA in isolated domain complexes or within the full 70-kDa complex.

a, Overlay of the protein amide 1H–15N HSQC spectra of the isolated stem–loop complexes (SL1, violet; SL2, green; SL3, orange; SL4, cyan; and GGA39–41, red) and the 70-kDa RsmZ(1–72)/RsmE 1:3 complex (black). The free protein is shown in blue. b, The 1H–13C transverse relaxation optimized spectroscopy (TROSY) spectra of the RNA aromatic adenine H2–C2 resonances of the segmentally labelled RNAs in complex (in colour) were shifted on both axes by 0.5 times the one-bond amide scalar coupling 1JNH to be superimposed on the 1H–13C HSQC spectra of the isolated binding sites in complex (black). The colours of the spectra correspond to the isotopically labelled RNA regions within the 70-kDa complex in the corresponding samples: SL1, 1–16; SL2, 17–40; SL3, 41–58; SL4, 59−72; 9nt-GGA39−41, 36–44. Note that nucleotides (nt) 36–40 are isotopically labelled in both SL2 and 9nt-GGA39–41, whereas nucleotides 41–44 are isotopically labelled in both SL3 and 9nt-GGA39–41 constructs (red labels in the SL2 and SL3 spectra). Double peaks due to the two conformations in the 70-kDa complex present in solution are indicated by arrows. See Supplementary Discussion for further explanation. c, Shown are the G35 and G55 imino regions in the 1H–15N HSQC spectra of the full complex indicating two conformations. d, Depiction of chemical shift agreement between RNA in isolated domain and within the full 70-kDa complex. The RNA nucleotides having at least one assigned resonance with a combined chemical shift (ΔCCS) difference <0.12 p.p.m. or <0.2 p.p.m. between the isolated domain and full complex are coloured in red or orange, respectively. Black balls between two base-paired nucleotides indicate a protected imino observed in the 1H–15N HSQC spectra of the full complex. The smaller black ball representing the G71 imino indicates that this imino was only weakly present. The regions for which the structures of the isolated domains were used as building blocks for the full complex are boxed. See Supplementary Methods for further explanation.

Extended Data Figure 3 Overview of measured EPR distances.

a, Location of all eight spin-label positions shown on the secondary structure of RsmZ (top) or on the three-dimensional structures of both conformations L and R (bottom). A ball represents the centre of the spin label distribution at the corresponding site. A common A(N)GGAX motif (where N and X stand for any nucleotide) is present at every binding site. The corresponding nucleotides N or X were replaced by a thiouridine residue to which the spin-label was attached. Because the residues N and X are only unspecifically recognized by the protein7,10, the complex formation is not disturbed, evidenced by native PAGE gels confirming the formation of the 1:3 full complex and by double electron-electron resonance (DEER) measurements of each isolated binding site in complex with one RsmE protein dimer18. b, Distance distributions obtained from DEER data (black). Grey and red vertical lines show the distance ranges considered in the EPR constraint determination (grey, 100% area; red, 70% area; see Methods). Red vertical lines indicate the regions that were taken as the EPR distance constraints. Asterisks indicate noise artefacts (B-E) and the features that appeared to be unstable upon variation of the background model. Note that for B-E, C-F and E-G, distance distributions are possibly affected by partial aggregation.

Extended Data Figure 4 Potential tertiary interactions stabilize the global fold in both conformations, which cannot directly interconvert.

af, Conformer L (ac) and conformer R (df). Protein chains A/B, C/D and E/F belong to RsmE protein dimers 1, 2 or 3, respectively. a, The RNA backbones of SL1 and SL3 approach each other up to 5.5 Å and thus, the high local charge density is probably stabilized by ions. b, The RNA backbones of SL2 and SL3 approach each other. Besides potential ions, the positively charged side chain of R31E (dimer 3) and the protein backbone amide of Q28A (dimer 1) might help to neutralize the high charge density. c, Potential hydrogen bonds and one stacking interaction between the conserved A41 and A18 could rationalize the packing of the third domain (GGA39–41, red residues) towards the RNA linker between SL1 and SL2 (A18). d, A base-stacking interaction between A41 and A43 might stabilize the positioning of the third domain with respect to SL3. e, In several low-energy structures, the conserved C17 and A18 are stacking on the U1–A16 SL1 stem-closing base pair. While stacking, C17 and A18 form a non-canonical cSWC base pair. These base-pair and stacking interactions orient the stem of SL1 in respect to the RNA linker residues C17–A18 directly preceding SL2. Furthermore, the RNA backbone linker between SL1 and SL2 approaches the RNA backbone of SL3. This high charge density could be stabilized by the K7B protein side chain (dimer 1). f, The RNA backbone at the 3′ end of SL4 could contact the K7F side-chain residue (dimer 3), thus also contributing to stabilization of the position of the third dimer. Overall, these potential interactions rationalize the presence of two well-defined structures. g, Two different possible RNA backbone conformations explain the formation of two conformations that cannot interconvert unless all proteins have dissociated.

Extended Data Figure 5 NMR evidence for a structural and/or dynamic change upon binding of one SL2 RNA molecule to the RsmE protein homodimer.

Top, a schematic representation of the free protein, the singly RNA bound protein and the doubly bound protein form is depicted. The dots represent a certain protein resonance in the free protein (blue), on the free binding site when the other protein binding site is bound (magenta), in the bound form when the other protein binding site is unbound (orange) or in the bound form when the other protein binding site is also bound (red). Bottom, a zoom into a representative region of the protein 1H–15N HSQCs at all the titration points is shown (from top to bottom: free protein, 0.66, 1.33 and 2 equivalents of SL2 RNA). The resonance peaks of isoleucine I34 are labelled in colour according to the four possible unbound or bound states (same colour code as schematic representation (top)).

Extended Data Figure 6 ITC binding data and gel shift assays.

a, b, Binding of the first two RsmE dimers to RsmZ increases the binding affinity of the third dimer. a, Gel shift assay of an RNA construct only containing the first four stem–loops of RsmZ, RsmZ(1−72), shows that the binding of the third dimer has around 300 nM Kd when the first two dimers are already bound. b, When disrupting the four binding sites of the first two dimers (using SL1234_ΔGGA_GGA39–41, see Supplementary Methods), the binding affinity of the third dimer is reduced 25-fold. The affinity was determined by ITC, because no binding was detectable by gel shift assays. c, ITC titration curves showing that the SL23 linker is longer than required to bind the first dimer with the highest possible affinity (see Fig. 3b). d, ITC titration curve of RsmE binding to a truncated form of the 5′ UTR of the hcnA mRNA missing the low-affinity GGA motifs nos. 1 and 2.

Extended Data Figure 7 Conservation of RsmZ sRNAs in Pseudomonas.

a, Secondary structure of the P. fluorescens Pf-5 RsmZ sRNA (which is identical to the CHA0 strain). b, Multiple sequence alignment of RsmZ sRNAs in different Pseudomonas species. The alignment was adapted manually according to the structural knowledge of the RsmZ sRNA. The GGA-binding motifs including all loop nucleotides are shown in red, nucleotides located in stem regions are coloured in green, and base-paired terminator nucleotides in purple. Nucleotides conserved within all the species are marked by stars. c, In vitro translation assay demonstrating a very similar translation activation potential for the RsmZ sRNAs in P. fluorescens and P. aeruginosa. The slightly smaller translation activation of RsmZ in P. aeruginosa is likely due to the missing GGA85–87 motif. d, Conservation of RNase E cleavage sites in RsmZ sRNAs in Pseudomonas. The three RNase E cleavage sites are conserved in Pseudomonas, except that in P. aeruginosa the GGA85–87 cleavage site is missing because the terminator directly follows the GGA76–78 motif. All RNase E cleavage sites overlap with GGA-binding motifs that bind dimer 2, dimer 3 and dimer 4 (boxed nucleotides). The nucleotides located in a stem are shown in bold and are underlined. The RNase E consensus cleavage site proposed by Kaberdin is shown below the multiple sequence alignments51. Nucleotides matching the RNase E cleavage consensus sequence are shown in green, the non-matching nucleotides in red. The two main cleavage sites in the single-stranded region between SL4 and the terminator are single-stranded and match the RNase E cleavage consensus sequence. The cleavage site in the loop of SL1 contains some nucleotides 3′ to the cleavage site, which are not single-stranded and do not match the cleavage consensus sequence. This is in agreement with the RNase E cleavage site in SL1 being less active compared to the two main cleavage sites between SL4 and the terminator. e, Sequential RNase E degradation of the RsmZ sRNA in vitro.

Extended Data Figure 8 Selecting the correct cluster combinations by comparing measured and back-calculated distance distributions.

a, How to obtain the modelled distance distribution for a single spin label pair (AC as an example) for one specific structure? The modelled distance distribution is calculated by plotting the occurrence of distances between all the radical positions of spin label A to all the radical positions of spin label C. On the top, the ‘radical clouds’ (spin-label distributions) for spin labels A and C are superimposed onto one structure of cluster L2 (left) or cluster R (right). Bottom, the measured (blue) distance distribution is superimposed on the modelled one of the L2 structure (cyan) and the R structure (magenta). b, Overview of measured (blue) and modelled (red) distance distributions for both possible cluster combinations for some representative spin label pairs (note that the AG and AC distance pairs are the best indicators to discriminate between the different cluster combinations (see Supplementary Methods)). The modelled distance distribution for the two clusters present are summed up and shown as a single distribution (red curve). All the distances are shown in Å. The best cluster combination (L2-R) is boxed in green. An overview of all 21 spin-label pairs for both possible solutions is shown in Extended Data Fig. 9c, d.

Extended Data Figure 9 Supplementary DEER data.

a, DEER primary time domain data (black) with 3D background fits (red). b, DEER form factor traces (black) and their best model free fits (red) with use of Tikhonov regularization (obtained with DEER analysis). c, d, Measured (blue) versus modelled spin label distributions (red) for cluster L1-R (c) or L2-R (d) combinations (see Supplementary Methods and Extended Data Fig. 8 for explanations). Asterisks indicate noise artefacts (B-E) and the features that appeared to be unstable upon variation of the background model. Note that for BE, CF and EG, distance distributions are possibly affected by partial aggregation.

Extended Data Table 1 NMR and refinement statistics for complexes

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This file contains Supplementary Methods, a Supplementary Discussion and additional references. (PDF 427 kb)

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Duss, O., Michel, E., Yulikov, M. et al. Structural basis of the non-coding RNA RsmZ acting as a protein sponge. Nature 509, 588–592 (2014).

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