Nucleic Acids Research, 2001, Vol. 29, No. 15 3145-3153
© 2001 Oxford University Press
Bulged residues promote the progression of a loop–loop interaction to a stable and inhibitory antisense–target RNA complex
UPR 9002 du CNRS, Institut de Biologie Moléculaire et Cellulaire, 15 rue Rene Descartes, 67084 Strasbourg Cedex, France and 1Institute of Cell and Molecular Biology, Biomedical Center, Uppsala University, Box 596, Husargatan 3, S-75124 Uppsala, Sweden
Received May 8, 2001; Revised and Accepted June 20, 2001.
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ABSTRACT |
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 TOP ABSTRACT INTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONSUPPLEMENTARY MATERIALREFERENCES |
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In several groups of bacterial plasmids, antisense RNAs regulate copy number through inhibition of replication initiator protein synthesis. These RNAs are characterized by a long hairpin structure interrupted by several unpaired residues or bulged loops. In plasmid R1, the inhibitory complex between the antisense RNA (CopA) and its target mRNA (CopT) is characterized by a four-way junction structure and a side-by-side helical alignment. This topology facilitates the formation of a stabilizer intermolecular helix between distal regions of both RNAs, essential for in vivo control. The bulged residues in CopA/CopT were shown to be required for high in vitro binding rate and in vivo activity. This study addresses the question of why removal of bulged nucleotides blocks stable complex formation. Structure mapping, modification interference, and molecular modeling of bulged-less mutant CopA–CopT complexes suggests that, subsequent to loop–loop contact, helix propagation is prevented. Instead, a fully base paired loop–loop interaction is formed, inducing a continuous stacking of three helices. Consequently, the stabilizer helix cannot be formed, and stable complex formation is blocked. In contrast to the four-way junction topology, the loop–loop interaction alone failed to prevent ribosome binding at its loading site and, thus, inhibition of RepA translation was alleviated.
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INTRODUCTION |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONSUPPLEMENTARY MATERIALREFERENCES |
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Many antisense RNAs are regulators of plasmid copy number (1). Plasmid R1 belongs to the IncFII family of plasmids which shares the overall genetic organization with respect to replication control functions. In plasmid R1, replication is negatively controlled at the translational level by binding of the antisense RNA (CopA) to its target site (CopT) in the leader region of the repA mRNA,
80 nt upstream of the repA start codon (2). Synthesis of the replication initiator protein RepA requires translation of a short leader peptide (tap), encoded upstream of repA. Binding of CopA prevents tap translation by occluding ribosome binding at the tap ribosome binding site (RBS), and thereby repA expression is inhibited (3–5). Interestingly, several copy number control antisense RNAs which all inhibit translation of replication initiator proteins have structural properties similar to those of CopA. These RNAs are fully complementary to their target site and contain a long stem–loop structure. For proper control, the intracellular concentration of the antisense RNA must be a measure of the plasmid concentration: this is ensured by constitutive synthesis and a short half-life of these RNAs (6). Furthermore, efficiency of in vivo control is correlated with high rates of antisense RNA binding to its target site [in the order of 106 M–1s–1 (7)]. This class of antisense RNAs does not require trans-acting proteins to promote inhibition. Thus it can be speculated that the structure of these antisense RNAs has evolved to be optimized for the critical steps that determine their regulatory functions. Kinetic studies have shown that the initial step of antisense RNA binding involves a loop–loop interaction mediated by a limited number of Watson–Crick base pairs, the so-called kissing complex [R1 (8); pMU720 (9); ColIb-P9 (10,11)]. In plasmid R1, we have recently shown that this loop–loop interaction is rapidly converted into a stable and inhibitory complex formed through a hierarchy of distinguishable intermediates, and that the formation of a full duplex in vitro is too slow to be of biological relevance (12–14). The major product of the binding reaction adopts an unusual structure characterized by a four-helix junction. Its formation involves extensive breakage of intramolecular base pairs to promote the formation of two intermolecular helices (see Fig. 1) (13). This conversion is essential for the formation of an irreversible complex and for the activity of the antisense RNA in vivo (14). We proposed previously that the four-way junction structure could promote a side-by-side helical alignment to allow formation of a third long intermolecular helix, involving the 5' tail of CopA and the complementary region of CopT (13). Recent data indicated that a topology, strikingly similar to that of the stable CopA–CopT complex, may be present in complexes of all R1-related plasmids (11,15). This structure completely and irreversibly inhibits repA translation, mainly by occluding ribosome binding [e.g. pMU720 (16); R1 (5); Col1b-P9 (17)].
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In addition to the recognition loops and the single-stranded regions required for stable complex formation, these antisense and target RNAs carry internal loops or unpaired residues in the major stem close to the hairpin loop. This is characteristic for antisense RNAs with long hairpins for which, unlike RNAI/RNAII of ColE1 (18,19), no protein is involved in regulation. The upper stem structure-disrupting elements can have several functional implications. They protect the RNAs against the double strand-specific RNase III since removal of bulged residues rendered CopA sensitive to cleavage in vivo and in vitro (20). Furthermore, their deletion strongly affected control in vivo and rapid formation of stable antisense/target RNA complexes in vitro [R1 (21); pUM720 (22); Col1b-P9 (23)]. Importantly, the positioning of these bulged residues close to the apical loop is critical since an internal loop placed at the bottom part of the stem did not restore inhibitory activity (21).
In the present paper, we show that removal of the bulged residues in CopA and CopT interferes with progression of the initial loop–loop helix towards the four-way junction. Instead, the loop–loop interaction formed by bulge-less CopA and CopT complex becomes arrested in a state that involves pairing of all loop bases, and does not convert to a stable complex. This complex fails to prevent ribosome binding at tap RBS.
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MATERIALS AND METHODS |
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Oligodeoxyribonucleotides
Oligodeoxyribonucleotides were bought from IBA-NAPS. Primers for the generation of transcription templates were for CopT302: T7GG, 5'-GAA ATT AAT ACG ACT CAC TAT AGG TTA AGG AAT TTT GTG GCT GG-3' (T7 RNA polymerase promoter sequence underlined) and SeqP/II, 5'-CGG ATT CGG GTT CTT TA-3'; for CopT74: GW46, 5'-GAA ATT AAT ACG ACT CAC TAT AGG CCC CGA TAA TCT TCT TCA ACT T-3' and GW47, 5'-TGT TGG CTA TAC GGT TTA AGT-3'; for CopA: T7-SA, 5'-GAA ATT AAT ACG ACT CAC TAT AGT AGC TGA ATT GTT GGC TAT ACG-3' and T7-EA, 5'-AAA GCA AAA ACC CCG ATA ATC TTC-3'. The primers for CopI were T7-SI, 5'-GAA ATT AAT ACG ACT CAC TAT AGG GCC CCG GTA ATC TTT TCG T-3' and T7-EI, 5'-AAA CCC CGA TAA TCT TCT TCA-3'. SeqP/II was also used for primer extension in the toeprinting experiments.
DNA templates and RNA synthesis
CopT, CopA, CopI and mutant RNAs (Lo, Up, L/U) were synthesized by T7 RNA polymerase from PCR-generated DNA fragments as described (20). PCR fragments were generated from plasmid pGW58 carrying the wild-type copA/copT region (3), and from mutant plasmids (20). Transcription of CopT yields a run-off product of 302 nt initiated with GG instead of the GU sequence of the wild-type repA mRNA. A shorter CopT74 corresponding to the complementary sequence of the antisense RNA was also produced, the RNA starts with GG instead of AG, i.e. CopA RNA contains a 5' terminal G instead of an A residue. Neither of these nucleotide changes affects structure or binding properties. CopA and CopI mutants contained either a U52-A64:2 base pair inserted in the upper stem (Lo), or a C47:2-G69 base pair instead of a bulged guanine 69 (Up), or both (L/U) (Fig. 1) (20). The complementary changes were also introduced in CopT mutants. RNAs were purified from 8% polyacrylamide–urea gels.
5' End-labeling of dephosphorylated RNA was performed with T4 polynucleotide kinase and [
-32P]ATP (24). Labeled RNAs were purified by polyacrylamide–urea gel electrophoresis, eluted and precipitated twice with ethanol. Before use, the RNAs were dissolved in water and renatured by incubation at 90°C for 2 min, followed by slow cooling at 20°C in TMN buffer (20 mM Tris-acetate pH 7.5, 10 mM Mg-acetate, 100 mM sodium acetate).
Enzymatic and chemical probing
Antisense RNA binding was carried out at 37°C for 15 min in TMN buffer with end-labeled CopT (3 x 10–8 M) and a 5-fold excess of unlabeled CopA or CopI (1.5 x 10–7 M), or with end-labeled CopA or CopI (4 x 10–8 M) and a 5-fold excess of unlabeled CopT (2 x 10–7 M). Full duplexes between CopT and CopA species were formed by incubation at 90°C for 2 min followed by slow cooling to 37°C in TMN buffer. RNase V1 hydrolysis, Pb2+-induced hydrolysis and modification [N7]G by lead- and nickel-complex (NiCR) were done as previously described (13). Cleavage positions were identified by running, in parallel, RNase T1, RNase U2 and alkaline ladders of the end-labeled RNA (25). Incubation controls were done to detect nicks in the RNA.
Phosphate modification interference with ethylnitrosourea
5' End-labeled RNA (10 nM) was first modified in 50 µl of buffer containing 50 mM sodium cacodylate pH 8.0, 1 mM EDTA with 16 µl of an ethylnitrosourea (ENU)-saturated ethanol solution for 1 min at 90°C. Reactions were stopped by ethanol precipitation with 0.3 M sodium acetate. Pellets were dissolved in TMN buffer, and complexes were formed with 1–5 nM of the unlabeled complementary RNA. An aliquot of labeled RNA was submitted to the same treatment except that ENU was omitted. Free and complexed RNAs were separated by electrophoresis at 4°C on 8% non-denaturing polyacrylamide gels in 45 mM Tris-borate pH 8.3. The corresponding bands were identified by autoradiography, cut out, and the RNA was eluted overnight at 4°C in 500 mM ammonium acetate pH 6, 0.1 mM EDTA, followed by precipitation. Ethylated phosphates were identified by incubating the modified RNA in 10 µl of 0.1 M Tris–HCl pH 9.0 at 50°C for 10 min. The control experiment was submitted to the same treatment. After precipitation, the RNA fragments were separated by denaturing gel electrophoresis on 15% polyacrylamide gels. Cleavage positions were identified by running in parallel RNase T1, and alkaline ladders of the end-labeled RNA.
Toeprinting assays
Toeprinting was performed essentially as previously described (5). The [32P]5' end-labeled primer SepP/II was first annealed to CopT RNA in 20 mM Tris-acetate pH 7.5, 60 mM NH4Cl, 3 mM ß-mercaptoethanol, heated at 90°C for 1 min and cooled on ice for 1 min. After the addition of 10 mM Mg-acetate, the RNA was incubated at 20°C for 20 min. Initiation complex formation was performed at 37°C from 30 s to 15 min in a reaction mixture containing 10 mM Tris-acetate pH 7.4, 60 mM NH4Cl, 10 mM Mg-acetate, 6 mM ß-mercaptoethanol and, unless otherwise stated, 40 nM CopT RNA, 125 nM 30S subunits and 1 µM tRNAfMet. In the toeprinting inhibition experiments, antisense RNA/target RNA complexes were formed prior to ternary complex formation at 37°C for 15 min (as described above) in the presence of a 5-fold excess of CopA or CopI species, followed by the addition of 30S and tRNA. Concentrations of the antisense RNA was chosen to get an optimal formation of the kissing complex. Primer extension reactions were subsequently performed at 37°C for 15 min by adding 2 U of AMV reverse transcriptase (Life Sciences). Reactions were stopped by phenol extraction and ethanol precipitation.
Graphic modeling
The 3D model of CopT-L/U (nucleotides C88 to A178) interacting with CopA-L/U (nucleotides G1 to U87) was built using several algorithms (26) incorporated in the program MANIP (27). The generated model was subjected to restrained least-squares refinement using the programs NUCLIN and NUCLSQ (26) in order to ensure geometry and stereochemistry with allowed distances between interacting atoms and to avoid steric conflicts. The color views were generated with the program DRAWNA (28).
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RESULTS |
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The loop–loop interaction is essential to initiate pairing in the wild-type and mutant complexes
Mutations in CopA and CopT which generated fully helical upper stems decreased the rate and stability of stable CopA–CopT complex formation by more than one order of magnitude (21). These effects suggested the topology of mutant and wild-type complexes to be different. Here, we used chemical modification interference experiments to probe the initial interaction in the wild-type CopA–CopT, CopI–CopT and mutant CopI-L/U–CopT-L/U complexes. CopI is a truncated CopA, unable to form the stable complex with CopT, but capable of forming the extended kissing complex (8,13). In the L/U mutant RNAs, the bulged nucleotides have been replaced by standard Watson–Crick base pairs (Fig. 1) (20,21). Phosphate groups were modified by treatment with N-ethyl-N-nitrosourea. The protocol used ensured ethylation of the non-bridging oxygens to, on average, less than one modification per molecule. Such an approach had been successfully used to probe the loop–loop interaction involved in the dimerization process of HIV-1 RNA (29). Modified antisense or target RNAs were allowed to form complexes with their counterparts. Free RNA and RNA–RNA complex, were separated by gel electrophoresis under native conditions and eluted from the gel. Sites of modifications in each fraction were then identified by alkaline treatment which induces RNA scission at the modified phosphate. Cleavages present in the free RNA species but absent from the ones in the complex indicate the positions at which modification prevents complex formation (negative interference). Consequently, the free RNA fraction is enriched for RNA species that carry modifications that interfere with complex formation, and thus the cleavages corresponding to the positions of negative interference are enhanced (Fig. 2, lanes F).
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Strong inhibition of wild-type CopA–CopT complex formation was observed upon ethylation of phosphates 112–114 in CopT, and 58–59 in CopA (Fig. 2A and B). Interestingly, the strongest interference pinpointed a subset of nucleotides in both CopT and CopA loops (Fig. 2D) in which many copy number mutations had been mapped [5'-GGCG in CopT; e.g. (30)]. In addition, ethylation of phosphates 111, 115–120 in CopT, and phosphates 57 and 60 in CopA, resulted in partial interference. Identical results were obtained with the truncated antisense RNA, CopI (result not shown). The mutant complex formed between CopI-L/U and CopT-L/U was also analyzed (Fig. 2C). Compared to the wild-type case, a single and extended region of strong negative interference was observed, comprising phosphates 57–62 within the loop of CopI-L/U (Fig. 2D). In addition, partial interference was observed upon modification of phosphates 54–56 in the 5' part of the CopI-L/U stem. Given the number of modifications that prevent complex formation, this negative interference may also reflect local perturbation of the CopA/I and CopT loop structures.
These experiments indicate that, in both the wild-type and L/U mutant cases, modification of phosphates within the recognition loops interfered with complex formation. This suggests that removal of bulges did not impair formation of the initial loop–loop interaction. However, the differences in the interference patterns observed between the wild-type and mutant complex suggest that these two complexes adopt different topologies.
Removal of the bulged residues prevents the formation of the intermolecular helices
We previously showed that the initial loop–loop interaction is converted to an unusual four-way junction structure (13). The initial base pairs formed are disrupted to permit helix progression into the upper stems, resulting in the formation of two intermolecular helices B and B' (Fig. 1C). RNase V1 probing of CopA–CopT or CopI–CopT complexes provided a characteristic signature for helices B and B' (Fig. 3A) (13): new or enhanced cleavages were induced in CopA or CopI at consecutive positions U52–C56 (helix B), and in CopT at C104–A105 (helix B'). In addition, the intermolecular helix C, which greatly enhances CopA–CopT complex stability, was characterized by the occurrence of several V1 cuts in the 3' end of CopT, as well as by the disappearance of V1 cleavages in CopA at positions G7–A9 and G16–A19, resulting from melting of helix I (Fig. 3A). Thus, RNase V1 was used to probe the structure of the homologous antisense/target RNA complexes containing mutations of either the lower bulge (Lo), upper bulge (Up) or both (L/U) (Fig. 3B–D), as well as the heterologous complexes formed by wild-type CopA and mutant CopT RNAs (results not shown).
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None of the mutant complexes presented the characteristic RNase V1 pattern observed for the wild-type complex. In the three homologous mutant complexes (CopA-Up/CopT-Up, CopA-Lo/CopT-Lo, CopA-L/U/CopT-L/U), helix C did not form properly. This is indicated by cleavages at positions 7–9 and 17–19 in CopA-Lo and CopA-L/U bound to their target CopT, which remained unchanged as compared to free mutant CopA (Fig. 3B and D), whereas CopT-Up induced only weak protections at positions 17–19 in CopA-Up (Fig. 3C). Several new cuts were detected in the hairpin loops of CopA-Up (C54–C56), of CopA-Lo (U55 and C56) and of CopA-L/U (U55–C56, A60 and A61) upon binding of CopT-Up, CopT-Lo and CopT-L/U, respectively (Fig. 3). In addition, enhanced cleavages were observed on the 5' side of the major stem of CopA-Lo and CopA-L/U. Conversely, in all three homologous mutant complexes, binding of CopA variant did not change the pattern of RNase V1 cleavage in the corresponding CopT target. In particular, no enhanced cleavages at C104–A105 were detected in mutant CopT, indicating that helix B' is not formed in the mutant complexes (results not shown).
Thus, replacing the bulged residues by Watson–Crick base pairs in the antisense and target RNAs induced dramatic changes in the overall topology of the CopA–CopT complex: the two intermolecular helices B and B' were not formed, and thus formation of the third intermolecular helix C could not occur. These data are in good agreement with the dramatic effects of these three mutations on the rate of stable CopA–CopT complex formation (21).
A fully base paired loop–loop interaction in the mutant CopA–CopT complexes
To obtain more detailed information on the accessibility of nucleotides within the CopA-L/U/CopT-L/U complex, NiCR-induced hydrolysis was used (13). Lead-induced cleavages occur mainly in single-stranded regions and in helices of low stability. NiCR modifies [N7]G and is very sensitive to stacking of base rings. Thus, N7 of purines within a helix are not reactive unless the deep groove is widened (31).
We showed previously that Pb2+-cleavages in the CopA–CopT complex were restricted to unpaired residues located in the loops (C58–A61) connecting the two intermolecular helices B and B' (13). In free CopA-L/U RNA, lead cleaved mainly in the hairpin loop and in the 5' single-stranded region of the RNA (Fig. 4) (21). Binding of either CopT-L/U or CopT-wt induced strong protections at residues U55–A61 in the apical loop whereas minor protections were detected in the 5' domain of CopA-L/U (Fig. 4A). This is supported by the NiCR probing experiment (Fig. 4B). In the major stem–loop of CopA-L/U, only the N7 position of G57 was modified whereas all guanines within the stem were unreactive. Notably, G69 which is base paired in CopA-L/U was not modified, whereas it was reactive in CopA-wt (Fig. 4B). The only change induced by binding of CopT-L/U was restricted to the complete protection of G57 at N7 in CopA-L/U (Fig. 4B). In agreement with results from the RNase V1 probing, the stabilizer helix C was not formed in the CopA-L/U/CopT-L/U complex since position N7 of guanines 12, 23–24, 30, 32–33 in the 5' domain of CopA-L/U were still reactive in the complex (Fig. 4B). Binding of CopT-Up also induced protection restricted to G57 at N7 in CopA-Up whereas G69 at N7 remained reactive towards NiCR (data not shown). By contrast, most of the N7 positions of guanines, with the notable exception of G57, showed protection towards NiCR in CopA-wt when bound to CopT-wt (Fig. 4B). In full duplexes (formed artificially), all the guanines at N7 of CopA-wt were fully protected by binding of CopT (Fig. 4B). Thus, these experiments allow us to distinguish between the full duplex, the extended kissing in CopA–CopT complex, and the loop–loop interaction in CopA-L/U/CopT-L/U.
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Altogether these data indicate that removal of the bulged residues within the major stems of CopA and CopT resulted in a complex characterized by a fully base paired loop–loop helix whose formation was incompatible with subsequent formation of the stabilizer helix C.
A loop–loop interaction is insufficient to prevent ribosome binding
CopA-mediated inhibition works by steric interference: the CopA–CopT complex prevents binding of ribosomes at the tap translation initiation site (5). Even a truncated CopA, resembling CopI, inhibited RepA synthesis in vivo, although at somewhat reduced efficiency (32). CopI, although unable to form a stable complex with CopT, forms the four-way junction structure (8,13), and was capable of interfering transiently with ribosome binding in vitro (5). Since removal of the bulged residues inhibited the activity of the mutant antisense RNAs in vivo (21), we tested here whether the loop–loop interaction in the three mutant CopI–CopT complexes could interfere with ribosome binding.
Toeprinting analysis was used to monitor the formation of translation initiation complexes on mRNAs in vitro (33). Previous experiments showed that CopI-dependent inhibition of ribosome binding at the tap initiation site was only detectable after a short incubation time. Therefore, complexes between CopI and CopT carrying either mutation Lo, Up or L/U were pre-formed, at RNA concentrations allowing essentially quantitative formation of the kissing complex during pre-incubation. Thereafter, the ternary mRNA/30S/tRNAfMet complex was formed by adding 30S subunits and tRNAfMet. Ternary complex formation was quantified over time by monitoring the signal of the tap toeprint (Fig. 5). For reference, the same experiments were carried out with the wild-type CopI/CopT and CopA/CopT complexes, respectively. As shown previously, inhibition by CopA is irreversible within the time interval assayed, whereas CopI-dependent inhibition was progressively lost with 50% inhibition after 2 min incubation (Fig. 5) (5). In contrast, the three homologous mutant CopI–CopT complexes were completely inactive in inhibition of initiation complex formation at the tap RBS (Fig. 5).
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DISCUSSION |
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A common characteristic feature of most, if not all, prokaryotic antisense and target RNAs is the presence of well-defined stem–loop(s) as key structure elements. In many antisense regulatory systems, RNAs initiate binding by transient and reversible interactions between complementary loops and, depending on their particular topological constraints, use different pathways to proceed rapidly to more stable and inhibitory complexes (e.g. 9,11,34,35). Formation of sufficiently stable complexes is important to obtain maximal inhibition rates, and to achieve quasi-irreversible inhibition kinetics.
For CopA/CopT of plasmid R1, we have recently shown that the initial loop–loop interaction becomes disrupted when intermolecular base pairing is propagated into the upper stems, creating a four-way junction topology (Fig. 1C) (13). The same sequence of events is also supported for the truncated antisense RNA (CopI) which only contains the recognition stem–loop structure (Fig. 1). The four-way junction topology is essential to promote the formation of the additional intermolecular helix between the single-stranded regions of the two RNAs that results in the stable CopA–CopT complex (14). The question addressed here concerns the driving force for conversion of the initial loop–loop to the extended kissing complex. Both CopA and CopT carry, in the upper part of the recognition stem, several unpaired nucleotides. Replacement of these residues by Watson–Crick base pairs decreased binding rate in vitro and control in vivo (21). Given the magnitude of impairment, it could not be excluded that these mutant RNAs followed different binding pathways which did not initiate by a loop–loop interaction but instead by contacts within the 5' single-stranded region of CopA. The experiments presented here indicate that removal of the bulged residues does not prevent the initial formation of a loop–loop helix, but abolishes the subsequent transition towards the four-way junction. Structure probing of the three homologous mutant RNA complexes indicates that helices B and B' do not form (Figs 3 and 4). Instead, the data suggest that an intermolecular helix is formed between all 6 nucleotides in the complementary loop sequences, and that stem sequences do not participate in helix propagation. Interestingly, an extensively paired loop–loop helix was proposed for complexes between stem–loops of RNAI and RNAII of ColE1 (19,36). RNAI regulates ColE1 plasmid replication by inhibiting maturation of the pre-primer, RNAII. Binding is initiated by at least two loop–loop interactions followed by a series of reactions that progressively lead to the formation of a stable duplex (34,37). The NMR structures of two kissing complexes (6 and 7 bp loop–loop base pairs) have been solved (38–40). Both structures show full base pairing between the loops, and are characterized by a pronounced bend at the loop–loop helix towards the major groove. Interestingly, ethylation of any phosphate within the 6-nucleotide loop sequence of CopI-L/U prevents nucleation and initiation events for binding to CopT-L/U (Fig. 2). The dimerization of HIV-1 RNA which involves a loop–loop interaction between 6 complementary nucleotides in a 9-membered loop, is also inhibited by the ethylation of most of the phosphates of the loop nucleotides (29). However, in the wild-type CopA/CopT complex, where the initial interactions are transient, interference was restricted to the 5' phosphates of the GCCA sequence of CopA and CopI, as well as to the complementary sequence of CopT. Weak interference observed at phosphates 114–120 of CopT implicates these residues in formation of helix B which is an early step required to commit the interacting RNAs to stable complex formation (14). Taken together, our study strongly argues for a striking difference in topology between complexes formed by the bulge-less mutant and wild-type RNAs.
In contrast to the ColE1 case where extensive loop–loop helices are involved in the inhibitory pathway, the loop–loop helix in the context of CopA-L/U/CopT-L/U is not productive since the long intermolecular stabilizer helix C cannot form (Fig. 3) (21). The suggested overall topology of the loop–loop helix tentatively explains this defect. Based on the known NMR loop–loop structures, a tertiary model for the CopA-L/U/CopT-L/U complex was built by graphic modeling, taking into account stereochemical constraints and probing data (Fig. 6). Despite the presence of a pronounced bend at the loop–loop helix, the two complementary regions that normally interact to give the stabilizer helix C are too far apart in space for efficient pairing (>100 Å; Fig. 6). In the ColE1 plasmid, the stem–loop structures of RNAI/II are shorter than in CopA/T, and bulged residues comparable to those in CopA/T are lacking (19). Also, bending at the loop–loop helix is further enhanced by Rom protein, most likely facilitating the distal contact between the 5' tail of RNAI and its complement (38) which initiates full pairing. In plasmid R1 and related plasmids, no protein is known to play such a role. Instead, all these plasmids carry unpaired residues or bulged loops in the upper stem regions of antisense and target RNAs although their positions are not strictly conserved (15). These elements are essential for rapid in vitro binding and in vivo control [R1 (21); pMU720 (22); Col1b-P9 (23)]. In all these systems, the rapid conversion of the initial loop contacts to the extended kissing complexes is crucial for efficient inhibition. This step requires substantial melting of the upper stem regions of both antisense and target RNAs (9,11,13). We show here that, in the CopA/CopT system, this step is driven via the bulged residues; unpaired residues in the upper stems favor inter-strand helix progression over imperfect intra-strand base pairing. Bulged residues in long hairpin structures were also shown to be essential in other antisense regulated systems which initiate antisense/target binding via loop–linear RNA pairing (41–43).
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Previous work showed that CopA inhibits the synthesis of RepA indirectly. Translation of a leader peptide, Tap, encoded immediately downstream of the CopA target site and in front of the repA reading frame is required for repA translation (4,44). CopA prevents ribosome binding at the tap ribosome loading site (5). Even CopI, although it only forms the four-way junction structure, transiently interferes with ribosome binding in vitro (Fig. 5) (5) and inhibits repA expression in vivo albeit at lower efficiency (32). Here we show that the CopI–CopT complexes carrying mutations Lo, Up and L/U do not inhibit ribosome binding at the tap RBS (Fig. 5). Since previous work has ruled out distal structure rearrangements in CopT as the mode of inhibition (12,13,45), the likely difference between inhibition (wild-type CopI–CopT) and lack of inhibition (bulge mutant CopI–CopT) should be due to their different structures. Even though CopI binds far upstream of the tap RBS, the bulky topology of the wild-type complex can sterically interfere with initiation, at least transiently. When full CopA is tested, the stabilizer helix C then renders inhibition irreversible. The suggested structure for the bulge mutant CopI–CopT complex (Fig. 6B) might be oriented so that ribosome access fails to be blocked. However, detailed structural analysis of the four-way junction topology will be required to elucidate the molecular mechanism by which a distal kissing complex can interfere with ribosome binding.
Intermolecular interactions between RNAs play fundamental roles in gene expression. Most of these are based on sequence complementarity, and loop–loop interactions are often implicated. In vitro selection approaches designed to select aptamers directed against RNA predominantly resulted in the isolation of sequences complementary to a hairpin loop in a defined structural context (46,47). It appears that, depending on the biological context of the RNA–RNA interaction, RNA structures have evolved either to ‘freeze’ a complex once formed or, alternatively, to convert initial interactions by propagating helices along topologically feasible pathways (reviewed in 48). In antisense regulation, formation of stable RNA–RNA complexes requires a propagation of the initial loop–loop helix to proceed rapidly to sufficiently stable and inhibitory complexes. The features that govern the high efficiency of natural antisense RNA are thus related to the secondary and tertiary structures of the interacting partners as well as of the complexes formed.
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SUPPLEMENTARY MATERIAL |
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Coordinates of the molecular model of CopA-L/U/CopT-L/U complex are available as supplementary material at NAR Online.
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ACKNOWLEDGEMENTS |
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We thank Hervé Moine for stimulating discussions and critical reading of the manuscript and G. Bec for advice. This work was supported by grants from the Centre National de la Recherche Scientifique (CNRS), and by the Swedish Natural Science Research Council (NFR, E.G.H.W.) and the Swedish Research Council for Engineering Sciences (TFR, E.G.H.W.).
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FOOTNOTES |
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* To whom correspondence should be addressed. Tel: +33 3 88 41 70 51; Fax: +33 3 88 60 22 18; Email: p.romby{at}ibmc.u-strasbg.fr
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REFERENCES |
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