RNA recognition by a Staufen double-stranded RNA-binding domain
2000; Springer Nature; Volume: 19; Issue: 5 Linguagem: Inglês
10.1093/emboj/19.5.997
ISSN1460-2075
AutoresAndres Ramos, Stefan Grünert, Jan Adams, David Micklem, Mark R. Proctor, Stefan M.V. Freund, Mark Bycroft, Daniel St Johnston, Gabriele Varani,
Tópico(s)RNA Research and Splicing
ResumoArticle1 March 2000free access RNA recognition by a Staufen double-stranded RNA-binding domain Andres Ramos Andres Ramos MRC Laboratory of Molecular Biology, Hills Road, Cambridge, CB2 2QH UK Present address: Molecular Structure Division, NIMR, The Ridgeway, Mill Hill, London, NW7 1AA UK Search for more papers by this author Stefan Grünert Stefan Grünert Wellcome/CRC Institute and the Department of Genetics, University of Cambridge, Tennis Court Rd, Cambridge, CB2 1QR UK Present address: IMP, Dr, Bohrgasse 7, 1030 Wien, Austria Search for more papers by this author Jan Adams Jan Adams Wellcome/CRC Institute and the Department of Genetics, University of Cambridge, Tennis Court Rd, Cambridge, CB2 1QR UK Search for more papers by this author David R. Micklem David R. Micklem Wellcome/CRC Institute and the Department of Genetics, University of Cambridge, Tennis Court Rd, Cambridge, CB2 1QR UK Search for more papers by this author Mark R. Proctor Mark R. Proctor Cambridge Centre for Protein Engineering, Hills Road, Cambridge, CB2 2QH UK Search for more papers by this author Stefan Freund Stefan Freund Cambridge Centre for Protein Engineering, Hills Road, Cambridge, CB2 2QH UK Search for more papers by this author Mark Bycroft Mark Bycroft Cambridge Centre for Protein Engineering, Hills Road, Cambridge, CB2 2QH UK Search for more papers by this author Daniel St Johnston Daniel St Johnston Wellcome/CRC Institute and the Department of Genetics, University of Cambridge, Tennis Court Rd, Cambridge, CB2 1QR UK Search for more papers by this author Gabriele Varani Corresponding Author Gabriele Varani MRC Laboratory of Molecular Biology, Hills Road, Cambridge, CB2 2QH UK Search for more papers by this author Andres Ramos Andres Ramos MRC Laboratory of Molecular Biology, Hills Road, Cambridge, CB2 2QH UK Present address: Molecular Structure Division, NIMR, The Ridgeway, Mill Hill, London, NW7 1AA UK Search for more papers by this author Stefan Grünert Stefan Grünert Wellcome/CRC Institute and the Department of Genetics, University of Cambridge, Tennis Court Rd, Cambridge, CB2 1QR UK Present address: IMP, Dr, Bohrgasse 7, 1030 Wien, Austria Search for more papers by this author Jan Adams Jan Adams Wellcome/CRC Institute and the Department of Genetics, University of Cambridge, Tennis Court Rd, Cambridge, CB2 1QR UK Search for more papers by this author David R. Micklem David R. Micklem Wellcome/CRC Institute and the Department of Genetics, University of Cambridge, Tennis Court Rd, Cambridge, CB2 1QR UK Search for more papers by this author Mark R. Proctor Mark R. Proctor Cambridge Centre for Protein Engineering, Hills Road, Cambridge, CB2 2QH UK Search for more papers by this author Stefan Freund Stefan Freund Cambridge Centre for Protein Engineering, Hills Road, Cambridge, CB2 2QH UK Search for more papers by this author Mark Bycroft Mark Bycroft Cambridge Centre for Protein Engineering, Hills Road, Cambridge, CB2 2QH UK Search for more papers by this author Daniel St Johnston Daniel St Johnston Wellcome/CRC Institute and the Department of Genetics, University of Cambridge, Tennis Court Rd, Cambridge, CB2 1QR UK Search for more papers by this author Gabriele Varani Corresponding Author Gabriele Varani MRC Laboratory of Molecular Biology, Hills Road, Cambridge, CB2 2QH UK Search for more papers by this author Author Information Andres Ramos1,2, Stefan Grünert3,4, Jan Adams3, David R. Micklem3, Mark R. Proctor5, Stefan Freund5, Mark Bycroft5, Daniel St Johnston3 and Gabriele Varani 1 1MRC Laboratory of Molecular Biology, Hills Road, Cambridge, CB2 2QH UK 2Present address: Molecular Structure Division, NIMR, The Ridgeway, Mill Hill, London, NW7 1AA UK 3Wellcome/CRC Institute and the Department of Genetics, University of Cambridge, Tennis Court Rd, Cambridge, CB2 1QR UK 4Present address: IMP, Dr, Bohrgasse 7, 1030 Wien, Austria 5Cambridge Centre for Protein Engineering, Hills Road, Cambridge, CB2 2QH UK ‡A.Ramos and S.Grünert contributed equally to this work *Corresponding author. E-mail: [email protected] The EMBO Journal (2000)19:997-1009https://doi.org/10.1093/emboj/19.5.997 PDFDownload PDF of article text and main figures. ToolsAdd to favoritesDownload CitationsTrack CitationsPermissions ShareFacebookTwitterLinked InMendeleyWechatReddit Figures & Info The double-stranded RNA-binding domain (dsRBD) is a common RNA-binding motif found in many proteins involved in RNA maturation and localization. To determine how this domain recognizes RNA, we have studied the third dsRBD from Drosophila Staufen. The domain binds optimally to RNA stem–loops containing 12 uninterrupted base pairs, and we have identified the amino acids required for this interaction. By mutating these residues in a staufen transgene, we show that the RNA-binding activity of dsRBD3 is required in vivo for Staufen-dependent localization of bicoid and oskar mRNAs. Using high-resolution NMR, we have determined the structure of the complex between dsRBD3 and an RNA stem–loop. The dsRBD recognizes the shape of A-form dsRNA through interactions between conserved residues within loop 2 and the minor groove, and between loop 4 and the phosphodiester backbone across the adjacent major groove. In addition, helix α1 interacts with the single-stranded loop that caps the RNA helix. Interactions between helix α1 and single-stranded RNA may be important determinants of the specificity of dsRBD proteins. Introduction The double-stranded RNA-binding domain (dsRBD) is among the most common RNA-binding motifs, and is found in single or multiple copies in many eukaryotic and prokaryotic proteins involved in RNA processing, maturation and localization (Green and Matthews, 1992; St Johnston et al., 1992). Three-dimensional structures of dsRBDs from several proteins have shown that the domain folds into a compact αβββα structure (Bycroft et al., 1995a; Kharrat et al., 1995; Nanduri et al., 1998). As in the other two major eukaryotic RNA-binding protein domains (Varani, 1997), the α-helical surface of the dsRBD structure packs through a conserved hydrophobic core against an antiparallel β-sheet. In vitro studies have shown that dsRBD proteins bind to dsRNA, but not to single-stranded RNA or DNA, nor dsDNA (St Johnston et al., 1992; Bass et al., 1994; Clarke and Matthews, 1995; Bevilacqua and Cech, 1996). These studies have shown that dsRBDs bind any dsRNA of sufficient length, regardless of its base composition, and therefore they represent general dsRNA-binding modules. The dsRBD was first identified in the Drosophila protein Staufen, which contains five copies of this motif (St Johnston et al., 1992). Staufen plays an essential role in the formation of the anterior–posterior axis in Drosophila and represented the first protein factor to be identified as critical for mRNA localization (St Johnston, 1995). Staufen protein associates with oskar mRNA during oogenesis and is required for its transport to the posterior pole of the oocyte, where it defines where the abdomen and germline will develop (Ephrussi et al., 1991; Kim-Ha et al., 1991; St Johnston et al., 1991). After the egg has been laid, Staufen accumulates at the anterior pole of the egg, and anchors the anterior determinant bicoid mRNA (St Johnston et al., 1989; Ferrandon et al., 1994). Staufen plays a role in RNA localization in somatic cells as well, by associating with prospero mRNA during the asymmetric divisions of the embryonic neuroblasts, and by mediating its segregation to the smaller daughter cell produced by this division (Broadus et al., 1998; Schuldt et al., 1998). In common with most other systems where mRNA localization has been studied, the cis-acting signals required for oskar, bicoid and prospero localization all reside within the 3′-untranslated regions (3′-UTR) of these mRNAs (MacDonald and Struhl, 1988; Kim-Ha et al., 1993). Staufen protein associates in vivo with the 3′-UTRs of bicoid and prospero mRNAs to form ribonucleoprotein particles (Ferrandon et al., 1994; Schuldt et al., 1998). The bicoid RNA sequences required for this interaction have been mapped to three largely double-stranded regions (Schuldt et al., 1998). However, it remains to be proven whether Staufen interacts directly with these RNAs in vivo and, if so, how Staufen recognizes these specific transcripts. The binding of the dsRBD to dsRNA represents an example of protein–nucleic acid recognition distinct from the other common RNA-binding motifs characterized so far (Varani, 1997). To determine the nature of the dsRBD–dsRNA interaction, we have conducted extensive mutagenesis on the third dsRBD from Staufen (dsRBD3) and have used nuclear magnetic resonance (NMR) to determine the structure of the complex between this domain and an RNA stem–loop containing an optimal Staufen-binding site. We have mutated five critical interfacial residues located at the RNA–protein interface into full-length Staufen protein. These mutations abolish the RNA-binding activity of dsRBD3 in vitro and prevent Staufen-dependent RNA localization in vivo. The present results provide a description at the atomic level of the interactions between the dsRBD and RNA and demonstrate their physiological significance for Staufen-dependent RNA localization. Results RNA binding by Staufen dsRBD3 The third dsRBD from Staufen (dsRBD3) binds dsRNA with micromolar affinity, and conforms particularly well to the consensus sequence of the dsRBD motif (Gibson and Thompson, 1994). We therefore chose this domain to analyse the structural basis of dsRBD–RNA interaction. As a first step, we determined the minimal and optimal length of dsRNA required for binding by a dsRBD by performing North-western blots with RNA hairpin substrates containing double-helical stems of increasing length. RNAs containing >8 bp of dsRNA bind to dsRBD3, but optimal binding is observed with RNAs of 12 bp or longer (Figure 1A). Since further increases in the length of the double-helical region do not improve binding, we conclude that dsRBD3 binds optimally to stem–loops containing 12 bp. Disruption of the helical structure of the RNA by the introduction of unpaired bases significantly reduces binding. These results are consistent with studies on polypeptides derived from the two dsRBDs of RNA-activated protein kinase (PKR) (Schmedt et al., 1995; Bevilacqua and Cech, 1996). The full-length polypeptide binds to RNAs that contain at least 16 bp, but each dsRBD was found to cover ∼11 bp of RNA. Figure 1.Biochemical characterization of Staufen dsRBD3 interaction with RNA stem–loops. North-western blots showing (A) binding of wild-type dsRBD3 to RNA stem–loops and (B) alanine-scanning mutagenesis of dsRBD3. The top panel shows the positions of the amino acids that were substituted, with a diagram of the secondary structure of the domain underneath. The numbering refers to the general dsRBD alignment scheme; Pro1 corresponds to Pro579 in Drosophila Staufen. Green residues correspond to mutated amino acids required for the correct folding of the domain, while red residues are surface exposed. A black arrow indicates no RNA binding; grey arrow, reduced binding; no arrow indicates that the mutation had no effect. The lower panel shows a representative blot of the alanine substitution mutants probed with double-stranded VA1 RNA. The same amount of protein was present in each lane (data not shown). Download figure Download PowerPoint The identity of amino acids within dsRBD3 involved in RNA recognition was established by systematic alanine-scanning mutagenesis using the same North-western assay (Figure 1B). Several mutations involved amino acids whose identity is crucial for the structure of the dsRBD. Ile8, Phe18, Ala57 and Ala58 form part of the hydrophobic core of the domain. Mutations in Leu21, Arg22, Glu23 and Glu24 were introduced to disrupt the β-bulge within the first strand of the β-sheet, a highly conserved feature that is also present in ribosomal protein S5, a protein that is very similar in both sequence and structure to the dsRBD (Bycroft et al., 1995). As expected, mutation of each of these amino acids strongly reduces or abolishes RNA binding. Mutations in Arg12 and Phe32 are also likely to fall into this class, even though these amino acids are partially exposed on the surface of the domain. Arg12 caps the N-terminal α-helix, and its replacement with alanine might disrupt RNA binding by extending the helix into the following tight turn. Evidence presented below indicates that Phe32 anchors loop 2 and loop 4. The most informative mutations were changes in surface residues that affect RNA binding without altering the conformation of the domain, as demonstrated by circular dichroism. Mutations of this type cluster in three regions of the domain: Ser3, Gln4 plus His6, and Glu7 within the N-terminal helix α1; His28 and Lys30 within loop 2; and Lys50, Lys51 and Lys54 within loop 4 and the beginning of helix α2. It is notable that mutations in Lys50, Lys51 or Lys54 abolish RNA binding, whereas two non-basic amino acids in this loop, Val52 and Ser53, can be mutated to alanine without loss of binding. This suggests that electrostatic interactions mediated by basic residues play an important role in dsRBD–RNA recognition. Staufen dsRBD3 binds RNA using a highly conserved surface and without altering the RNA conformation Having established the biochemical properties of dsRBD3–RNA recognition, we used high-resolution NMR spectroscopy to characterize this interaction in structural detail. A stem–loop of 12 bp capped by an exceptionally stable C(UUCG)G loop was chosen to represent an optimal substrate, as defined by the experiments reported in Figure 1A. Since the dsRBD–dsRNA interaction is not sequence specific, the double-helical region was made fully symmetrical to simplify the NMR spectral analysis. Many protein resonances broadened considerably at subsaturating ratios of RNA when dsRBD3 was titrated with RNA, then sharpened up again when the RNA was added in stoichiometric amounts (Figure 2). This behaviour is found when the interconversion between free and bound forms occurs with intermediate exchange kinetics. This result strongly suggests that the off rate of binding is in the millisecond time scale, consistent with a micromolar dissociation constant and with the on rate being diffusion limited. Essentially complete spectral assignments were obtained for the bound form of dsRBD3 in the presence of RNA by applying standard procedures utilizing 15N- and 13C-15N-labelled dsRBD3 samples mixed with unlabelled RNA. Changes in chemical shift upon RNA binding define the footprint of the RNA on dsRBD3. The regions of the protein where large changes in the NMR spectrum occur upon RNA binding cluster at the N-terminus of the protein, in loop 2 and loop 4 and in the region where α1 packs against α2 and the β-sheet. No significant changes were observed for β2 and β3 or in the C-terminal region of the protein. Two conserved lysine residues within loop 4 (Lys50 and Lys51) are particularly interesting. The backbone amide resonances of Lys50 and Lys51 are invisible in the free protein spectra, presumably due to the accessibility of solvent molecules to this exposed region of the structure. However, the same resonances become visible upon complex formation. These residues are protected from exchange with solvent by the RNA, confirming their role in RNA recognition revealed by the alanine-scanning experiment. Figure 2.1H-15N HSQC spectrum of the free protein (A) and of a sample containing an ∼2-fold protein excess (B). Most protein resonances broaden beyond detection when the protein is in excess, and only resonances from the unfolded regions at either end of the domain remain visible. Under stoichiometric conditions (1:1), resonances from the protein become sharp again (C). Download figure Download PowerPoint The NMR data demonstrate that the folding of the domain does not change significantly upon RNA binding. Residues that display significant changes in the NMR spectrum of dsRBD3 upon RNA binding were therefore mapped on the structure of the free protein domain (Bycroft et al., 1995). The results unambiguously demonstrate that the face of the dsRBD formed by the N-termini of both helices, and by loops 2 and 4 along the edge of the first strand of the β-sheet, represents the RNA-binding surface of dsRBD3. Thus, the results of the alanine-scanning mutagenesis and the NMR footprint identify the same face of Staufen dsRBD3 as the surface where RNA recognition occurs. This protein surface contains exposed residues that are almost completely conserved among Staufen proteins from Drosophila to humans (Figure 3). Figure 3.Evolutionary conservation of the amino acids required for RNA binding in Staufen dsRBD3. (A) Alignment of dsRBD3 sequences from Staufen homologues of Drosophila melanogaster (Dm), Drosophila virilis (Dv), Musca domestica (Md), Mus musculus (Mm), Homo sapiens (Hs) and Caenorhabditis elegans (Ce). Grey boxes highlight positions where two or more other species have the same amino acid as Drosophila melanogaster. (B) Comparison of the results of the NMR analysis of the dsRBD3–RNA complex, the alanine-scanning mutagenesis and the positions of amino acids that have been highly conserved during evolution. Download figure Download PowerPoint Essentially complete spectral assignments were obtained for both free and bound RNA using isotopically labelled RNA samples. Remarkably, only a few residues displayed significant changes in their NMR properties upon formation of the complex, and the changes were generally of modest magnitude. This result demonstrates that dsRBD3–RNA interaction occurs with only small rearrangements of a preformed RNA structure. In order to establish the orientation of dsRBD3 with respect to the RNA, we measured residual dipolar couplings in a partially oriented sample. Dipolar interactions assume finite values when the sample is partially oriented, and these values provide absolute information on the orientation of NH and CH bonds (Tjandra and Bax, 1997). Residual NH dipolar couplings for dsRBD3 in complex with RNA show negative values for amino acids within the two α-helices and positive values for the β-strands (data not shown). When residual coupling constants were measured for the RNA in the complex, we found instead positive couplings for base NH and CH bonds. In double-stranded nucleic acids, the bases are approximately perpendicular to the double helix axis. Therefore, positive values of CH and NH couplings within the RNA, compared with the negative couplings for the protein α-helices NHs, show that the protein is bound to the RNA with the α-helices approximately parallel to the RNA double-helical axis. The dynamic character of the Staufen dsRBD–RNA interaction The analysis of 15N NMR relaxation properties for dsRBD3 was used to study the existence of conformational flexibility in the free and RNA-bound protein domain. As shown in Figure 4, 1H-15N heteronuclear NOEs are ∼0.7–0.8 in the well-folded core of the dsRBD. However, low heteronuclear NOE values are observed for the flexible tail at the end of the construct, reflecting complete disorder. Lower than average heteronuclear NOEs are also observed for residues within loop 2 and loop 4 both in the free and RNA-bound dsRBD3, reflecting residual conformational flexibility. Analysis of additional relaxation parameters reveals the existence of conformational exchange within loop 2, loop 4 and the N-terminus of helix α2 (data not shown). Furthermore, some NH resonances within loop 2 and loop 4 could not be analysed in the complex due to exchange broadening. These results demonstrate that loop 2 and loop 4, two of the three regions of the protein that form the RNA interface (see below), are highly mobile in the free protein and retain significant conformational flexibility in the complex. Figure 4.Heteronuclear 1H-15N NOE for free (A) and RNA-bound (B) Staufen dsRBD3. Download figure Download PowerPoint Structure of the Staufen dsRBD3–RNA complex The structure of dsRBD3 in complex with the stem–loop RNA was determined using a protocol very similar to that used in the determination of the structure of the U1A complex (Allain et al., 1996; Howe et al., 1998). No assumption was made at any stage of the data collection or structure calculation about the nature of the interaction or about the protein or RNA structures. The structure was based on the identification of intermolecular NOE interactions and on the definition of the relative orientation of the protein and RNA achieved by measuring residual dipolar couplings in partially oriented samples. The majority of NOE contacts involved sugar resonances in the sugar–phosphate backbone, suggesting that the domain does not contact the RNA bases intimately. However, the observation of NOE contacts from Ade3 H2 demonstrates that the protein binds the minor groove of the double helix, as suggested (Bevilacqua and Cech, 1996). Experimental and structural statistics are summarized in Table I. A stereo view of the structure is shown in Figure 5A and superposition of 10 low energy structures is shown in Figure 5B. Figure 5.(A) Stereo view of a low energy structure of the dsRBD3–RNA complex; Phe32 and five interfacial residues critical for Staufen function are shown explicitly. (B) Superposition of 10 converged structures and the free protein (orange); this image was prepared by superimposing the protein backbone between residues 1 and 65, but excluding loop 2; residues involved in intermolecular contacts with the RNA are shown explicitly in yellow for one representative structure. Download figure Download PowerPoint Table 1. Experimental constraints and structural statistics Protein RNA (a) Distance and dihedral angle NMR constraints Total 665 588 Intra-residue 202 215 Sequential 175 inter residue 206 Medium/long range 210 Hydrogen bonds 24 35 Dihedral restraints 0 127 Residual dipolar couplings 54 5 Protein–RNA distance restraints 10 Total No. constraints 1263 (b) Structure statistics NOE violations number >0.2 Å 3 ± 2.2 maximum violation 0.94 Å Angle violations number >5° 0.1 ± 0.3 Mean deviation from ideal covalent geometry bond length 0.005 Å bond angles 0.79° impropers 0.41° (c) R.m.s. deviations from average structure (Å) Protein–regular secondary structure (1–20, 30–47, 51–66) backbone 0.87 Å heavy atoms 1.53 Å RNA (superposed on all heavy atoms for the indicated residues) all residues 2.44 Å tetraloop 1.19 Å double-helical stem 2.08 Å Protein–RNA (all heavy atoms for indicated residues) all ordered regions 2.56 Å complex interface loop 2 2.53 Å loop 4 2.14 Å helix α1 1.56 Å The structure of the dsRBD3–RNA complex is of lower precision than that of the U1A complex (Allain et al., 1996; Howe et al., 1998). This is a consequence of the smaller number of intermolecular distance constraints, which could only be partially compensated by introducing absolute orientational information derived from residual dipolar couplings (Bayer et al., 1999). The small number of intermolecular NOEs is due to three distinctive properties of the dsRBD–RNA interaction, reflecting the weak, non-specific association between dsRBD and RNA. First of all, the majority of intermolecular interactions between Staufen dsRBD3 and RNA involve the RNA backbone, where there are relatively few resonances, and these are difficult to assign to specific nucleotides due to spectral overlap. Secondly, the intermolecular dsRBD3–RNA interface is small and involves relatively few protein residues. The area buried upon complex formation is only ≈1450 Å2, 12% of the total surface area. In contrast, protein side chains and RNA bases form an intricate intermolecular interface in the U1A complex that buries a much larger surface (Allain et al., 1996). Thirdly, the interface retains significant conformational flexibility (Figure 4), and this is likely to quench at least some intermolecular NOE interactions. Staufen dsRBD3 contacts the RNA stem–loop through the same sites identified by alanine-scanning mutagenesis and NMR chemical shift analysis (Figure 3): helix α1, loop 2 and loop 4 plus the N-terminal part of helix α2. The distance between loop 2 and loop 4 corresponds to the spacing between the minor groove and the phosphate across the intervening major groove in A-form RNA. The distance between the loop 2–RNA interactions and the tetraloop, the site of helix α1–RNA contacts, is 12 bp. This spacing is in perfect agreement with the optimal substrate length (Figure 1A). Helix α1 interacts with the C(UUCG)G tetraloop. This interaction is well defined by the experimental data (Figure 6C), and the relaxation data confirm that this region of the protein is rigid. However, only few intermolecular contacts can be interpreted as specific to the UUCG sequence. Ser3 interacts with the 2′-OH and phosphate oxygen of C13, the last nucleotide on the 5′ side of the stem. Glu7 interacts with the 2′-OH of U15 and stacks with the aromatic ring of G17, while Lys11 makes electrostatic interactions with the phosphate of residue 16. Ile10 is in van der Waals contact with G17. Intermolecular interactions involving loop 2 and loop 4 are less precisely defined, due to the residual conformational flexibility in this region of the RNA–protein interface revealed by NMR relaxation measurements. Residues within loop 2 interact with 2′-OH and phosphate oxygens within the minor groove close to the bottom of the double-helical stem (Figure 6A). The side chain amide of Lys30 interacts with 2′-OH groups in the minor groove of the RNA, while the heteroaromatic ring of His28 is positioned almost perpendicularly with respect to a phosphate oxygen. In the majority of converged structures, the phosphate oxygen is directed towards the centre of the ring of His28. The position of loop 4 with respect to the RNA is defined indirectly by the interactions observed between helix α1 and the UUCG loop, and between loop 2 and the RNA minor groove. These intermolecular interactions and the structure of the protein unambiguously position loop 4 near the phosphodiester backbone across the major groove from the site of loop 2 interactions with the minor groove. Three critical lysine residues within loop 4 and the N-terminus of helix α2, Lys50, Lys51 and Lys54, interact with phosphate oxygens and one 2′-OH group (Figure 6B) across the major groove from the sites of loop 2–minor groove interaction. The side chains of Lys50 and Lys51 bridge the major groove by interacting with RNA phosphates across the major groove from each other, while Lys54 reinforces these contacts by interacting with the phosphate immediately following the site of interaction of Lys51. Figure 6.Intermolecular interactions between dsRBD and the RNA stem–loop in the superposition of 10 converged structures; one structure is represented in orange for clarity. (A) Interaction between loop 2 and the minor groove of the double-helical stem; Ala27 and His28 from the conserved GPAH sequence and the Lys30 side chains are shown explicitly; 2′-OH groups in close proximity to amino acids side chains are highlighted in red. (B) Interactions between loop 4 and the N-terminus of helix α2 and RNA phosphates (in red). (C) Interaction between helix α1 and the UUCG tetraloop. Download figure Download PowerPoint Comparison of the structure of dsRBD3 free and in the RNA complex confirms that the structure of the protein does not change significantly on RNA binding, with the exception of loop 2. The rotation of loop 2 (towards the RNA in Figure 5B) is necessary to allow interactions between this region of the protein and the RNA. The RNA double-helical region preserves the A-form structure throughout the double-helical stem, and the UUCG tetraloop is in its well characterized conformation in the presence or absence of the protein. The only significant change in RNA structure upon protein binding is a kink at the stem–loop junction, resulting in the bent appearance of the RNA in the complex (Figure 5A). The presence of this distortion is supported indirectly by the observation of significantly shifted resonances in this region of the RNA. The bend allows the interaction between helix α1 and the tetraloop to occur at the same time as the contacts between loop 2 and the RNA minor groove. dsRBD mutagenesis in vivo The biochemical and structural data on dsRBD3 described above provide a framework to analyse whether the RNA-binding activity of this domain is required for Staufen function. Five highly conserved basic amino acids within loop 2 and loop 4 (His28, Lys30, Lys50, Lys51 and Lys54) are required for RNA binding in vitro, and lie at the RNA–protein interface where they interact with the RNA (Figure 5A). To generate a form of domain 3 that is completely null for RNA binding, we replaced all five of these amino acids with uncharged or negatively charged residues. 1H-15N HSQC spectra of the bacterially expressed quintuple mutant dsRBD3 are very similar to that of the wild-type protein (data not shown), demonstrating that mutant and wild-type proteins adopt the same conformation. Consistent with this observation, the domain displayed normal solubility and stability when expressed in Escherichia coli, but its in vitro RNA-binding activity was abolished. The DNA encoding this mutant domain was inserted into a staufen cDNA in place of the wild-type domain, and then transformed into the Drosophila germline in a vector that directs expression of the transgene in the female ovary (Micklem et
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