Long-range RNA–RNA interactions between distant regions of the hepatitis C virus internal ribosome entry site element

Esther Lafuente1, Ricardo Ramos1 and Encarnación Martínez-Salas1

Centro de Biología Molecular ‘Severo Ochoa’, Consejo Superior de Investigaciones Científicas-Universidad Autónoma de Madrid, Cantoblanco 28049 Madrid, Spain1

Author for correspondence: Encarnación Martínez-Salas. Fax +34 91 3974799. e-mail emartinez{at}cbm.uam.es


   Abstract
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Abstract
Introduction
Methods
Results
Discussion
References
 
Efficient internal initiation of translation from the hepatitis C virus (HCV) internal ribosome entry site (IRES) requires sequences of domain II, but the precise role of these sequences is still unknown. In this study, the formation of RNA–RNA complexes in the HCV IRES was evaluated. Using transcripts that contain the sequences of the structural HCV IRES domains II, IIIabcd, IIIabc, IV and IIIef-IV, specific long-range interactions between domains II and IV, as well as domains II and IIIabcd, have been found. These interactions were readily detected in a gel mobility-shift assay and required the presence of magnesium ions. A high concentration of nonspecific competitors, an 80 nt fragment of 18S rRNA or poly(I:C), did not interfere with the formation of RNA complexes. Interestingly, an RNA oligonucleotide bearing the sequence of stem–loop IIId interacted with domain II but not with domain IV or IIIef-IV, strongly suggesting that the interaction between domains II and IIIabcd was mediated by the IIId hairpin. Interaction between domains IIIabcd and IV was barely detected, consistent with the result that the apical part of domain III folds independently of the rest of the IRES. Moreover, the addition of stem–loop IIIef sequences to domain IV significantly reduced its ability to interact, which is in agreement with the formation of a compact RNA structure of domain IV with IIIef. The interactions observed in the absence of proteins between domains II and IV as well as stem–loop IIId and domain II may be transient, having a regulatory role in the translation efficiency of the HCV IRES.


   Introduction
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Abstract
Introduction
Methods
Results
Discussion
References
 
Internal ribosome entry site (IRES) elements are cis-acting mRNA sequences that promote internal entry of the ribosome in the appropriate framework to allow translation initiation at the correct start codon. The structural organization of the IRES element appears to be an essential determinant of its activity. However, secondary structural requirements for initiation of translation differ even among related IRES, like those present in picornavirus. This difference is more remarkable when comparison is extended to the hepatitis C virus (HCV) or classical swine fever virus (CSFV) IRES (Jackson, 2000 ). Hence the same function seems to be accomplished with different structural organization.

Experimental evidence in support of tertiary structure generated by RNA–RNA interactions is available for several IRES (Wang et al., 1995 ; Kanamori & Nakashima, 2001 ). A pseudoknot structure in the HCV IRES was shown to be required for IRES activity, as mutations that destabilized tertiary interactions between residues of loop IIIf with complementary residues in domain IV were accompanied by a strong reduction in translation initiation (Honda et al., 1996 ). Recently, interactions between distant residues of domain II have been claimed as another tertiary structure element of the HCV IRES (Lyons et al., 2001 ).

Long-range RNA–RNA interactions have been shown to occur in vitro between functional domains of the foot-and-mouth disease virus (FMDV) IRES (Ramos & Martínez-Salas, 1999 ). These interactions are strand-specific and depend on RNA concentration, ionic conditions and temperature. The RNA–RNA interactions observed in vitro between separated domains of the FMDV IRES, in the absence of proteins, suggest that the IRES adopts a specific folding pattern depending upon environmental conditions. Notably, the central domain of the FMDV IRES (named domain I or 3) seems to play a key role in directing the folding of the molecule. Domain 3 is the only domain that interacts efficiently with all the other domains, suggesting that it holds the other domains of the IRES. Consistent with this essential role, mutations in conserved motifs in the distal loop of this domain impaired the activity of the element (López de Quinto & Martínez-Salas, 1997 ; Robertson et al., 1999 ). Furthermore, it is of note that the binding site for several eukaryotic initiation factors (eIFs) and other RNA-binding proteins are located outside of the central region, in the distal domains 2 and 4–5 (López de Quinto et al., 2001 ).

The HCV IRES has been shown to adopt different folding patterns in response to different ionic concentrations (Kieft et al., 1999 ). Additionally, mutations in loop IIId cause a structural reorganization of the HCV IRES, as measured by RNase T1 sensitivity and Fe(II)–EDTA cleavage, concomitantly with the reduction in IRES activity (Jubin et al., 2000 ). In the absence of eIFs or the 40S ribosomal subunit, the HCV IRES seems to adopt a unique structure, which is stable at physiological salt concentrations. A structural element containing stem–loops IIIa, b and c facilitates binding of eIF3 (Kieft et al., 2001 ). On the other hand, subdomains IIId, e and f, together with the pseudoknot structural element, constitute the binding site for the 40S ribosomal subunit (Spahn et al., 2001 ). Recently, it has been shown that discontinuous fragments in domains III and IV in the RNA isolated from 80S RNA complexes are protected from RNase A cleavage (Lytle et al., 2001 ). Remarkably, sequences in domain II are required for IRES activity but the specific role of these sequences is still unknown.

Here we show that domain II of the HCV IRES can establish specific, long-range interactions with sequences located in domain IV, towards the 3' end of the IRES. In addition, stem–loop IIId contributes to the interaction observed between domains II and IIIabcd, while the apical part of domain III seems to fold independently of the other domains. Consistent with this, interaction between domains IIIabcd and IV is barely detected by means of this assay. On the other hand, stem–loop IIIef induces the formation of a compact structure with domain IV, which leads to a significant reduction in the ability of domain IV to interact with other IRES regions.


   Methods
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Abstract
Introduction
Methods
Results
Discussion
References
 
{blacksquare} Generation of separated structural domains of the HCV IRES.
The region of RNA corresponding to the HCV IRES was subdivided into five transcripts, in addition to that representing the whole IRES. The MFOLD program of the GCG package was used to predict the folding pattern of the separate domains, which conserved the predicted structure they assume in the context of the full-length IRES (Fig. 1A, B) (Honda et al., 1999 ). cDNAs corresponding to these five domains were subcloned into the pGEM3 vector (Promega). For that purpose, primers complementary to the 5' and 3' ends of each domain were used in a standard PCR that included the p156 plasmid, which harbours the HCV genotype 1b IRES sequence (N. Ibarrola & E. Martínez-Salas, unpublished data) as template. Each pair of primers consisted of a sense oligonucleotide (ODN2S, ODN3S and ODN4S), which included an EcoRI site, and an antisense oligonucleotide (ODN2As, ODN3As and ODN4As), which included an HindIII site (Table 1).



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Fig. 1. (A) Schematic representation and activity of the HCV IRES. Domain numbering is taken from Honda et al. (1999) . The panel on the right shows the translational activity of HCV IRES in reticulocyte lysates, as compared to that of the FMDV IRES, in bicistronic RNAs of the form CAT–IRES–luciferase. The mobility of the luciferase and CAT proteins is indicated by an arrow. (B) Schematic representation of the predicted structure of transcripts corresponding to the different HCV IRES domains used in this work.

 

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Table 1. Oligonucleotides used in this work

 
The complete IRES region of HCV, spanning nt 40–383 of the viral RNA (Honda et al., 1999 ), was also cloned into the same vector using ODN2S and ODN4As. The plasmid DIV was generated after digestion of domain IIIef-IV with SmaI and religation of the large fragment. This construct allows the synthesis of a transcript encompassing nt 125–130 fused to the stem–loop present in residues 318–383. This RNA maintains the C-rich region present between domains II and III fused to domain IV but lacks stem–loop IIIef (Fig. 1B).

The sequence of the entire length of the IRES under study was obtained using automatic sequencing (ABI PRIM Dye Terminator Cycle Sequencing Ready Reaction kit; Perkin Elmer). Additional nucleotides derived from the vector cloning sites do not seem to modify the predicted structure of the subcloned domains. No complementarity between the sense transcripts was observed in the additional residues.

{blacksquare} In vitro transcription.
Sense RNAs were transcribed in vitro from 0·5–1 µg of the linearized plasmids using 50 U T7 RNA polymerase (New England Biolabs), 50 mM DTT, 0·5 mM rNTPs, 20 U RNasin (Promega) or using the MEGAshortscript kit (Ambion). Plasmids were linearized with HindIII, with the exception of the IIIabc transcript, which was obtained after NheI digestion. When needed, RNA transcripts were labelled to a specific activity of 3 µCi/µg using [{alpha}32P]CTP (400 Ci/mmol). Reactions were incubated for 10 min with 1 U RQ1 DNase (Promega) and unincorporated [32P]CTP was eliminated by exclusion chromatography in TE (10 mM Tris and 1 mM EDTA, pH 8)-equilibrated Sephadex G 50–80 (Sigma) columns. RNA was extracted with phenol–chloroform, ethanol-precipitated and resuspended to an appropriate concentration in RNase-free water. The transcript of 136 nt, including a fragment of 80 nt from human 18S rRNA, was generated by T7 RNA polymerase-mediated transcription from pTRI-18S DNA template (Ambion).

{blacksquare} RNA–RNA interaction assay.
RNA molecules were mixed in a volume of 4 µl, heated at 95 °C for 3 min before adding 1 µl of a fivefold-concentrated binding buffer to reach a final concentration of 50 mM sodium cacodylate, pH 7·5, 300 mM KCl and 10 mM MgCl2 (Ramos & Martínez-Salas, 1999 ; Ferrandon et al., 1997 ). Independent RNA denaturation procedures did not modify the pattern of complex formation relative to mixed denaturation (R. Ramos & E. Martínez-Salas, unpublished data). Poly(I:C) (Pharmacia), yeast tRNA (Swartzmann) or a transcript including 80 nt from human 18S rRNA (transcribed from pTRI-18S) were used as nonspecific competitor molecules (800 nM each) in binding reactions, which also contained the specific interactor (800 nM) and the probe of interest (20 nM).

RNA–RNA complexes were allowed to form for 30 min at 37 °C and analysed immediately by electrophoresis in nondenaturing gels (Ramos & Martínez-Salas, 1999 ; Ferrandon et al., 1997 ; Paillart et al., 1996 ; Fedor & Uhlenbeck, 1990 ). Gels were run at room temperature for 20 min at 23 V/cm in TBM buffer (45 mM Tris, pH 8·3, 43 mM boric acid and 0·1 mM MgCl2). RNA Century molecular mass markers (Ambion) (0·5 µg) were loaded in parallel. Prior to drying the gel, RNA was stained with ethidium bromide and photographed to record the mobility of each transcript under study. Dried gels were exposed for autoradiography, as well as to a phosphorImager plate, to quantify the intensity of the retarded bands. Data were represented as the percentage of the RNA complex of interest relative to the input probe, averaged from at least three independent experiments.

In competition- and oligonucleotide-binding assays, the molar ratio of primer to transcript ranged from 0·25 to 1. When required, oligonucleotides were labelled at the 5' end using [{gamma}-32P]ATP (3000 Ci/mmol) and polynucleotide kinase (New England Biolabs) during 1 h at 37 °C. Unincorporated [{gamma}-32P]ATP was eliminated by exclusion chromatography in TE-equilibrated columns.

{blacksquare} In vitro IRES activity.
HpaI-linearized plasmids were transcribed in vitro with T7 RNA polymerase to produce bicistronic transcripts of the form CAT–IRES–luciferase. In vitro translations in nuclease-treated reticulocyte lysates (Promega) were programmed with p156-derived bicistronic transcript harbouring the HCV-1b IRES. The transcript derived from pBIC, which contains the FMDV IRES (López de Quinto & Martínez-Salas, 1999 ), was included for comparison. In both cases, 1 µg RNA, heated at 70 °C for 5 min, was translated for 1 h at 30 °C in 25 µl of 50% reticulocyte lysate in the presence of 25 µCi [35S]methionine (PRO-MIX L-35S; Amersham). Translation products were treated with 50 µg/ml RNase A, mixed with disruption buffer and resolved by 15% SDS–PAGE.


   Results
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Abstract
Introduction
Methods
Results
Discussion
References
 
RNA–RNA interactions between domains of the HCV IRES
We have previously shown the existence of long-range RNA–RNA interactions between separated domains of the FMDV IRES (Ramos & Martínez-Salas, 1999 ). To investigate whether this situation also applied to other IRES elements, we monitored the formation of RNA complexes using separate domains of the HCV IRES. Prior to its use for RNA-binding assays, IRES activity was verified in reticulocyte lysates using a bicistronic RNA of the form CAT–HCV IRES–luciferase (Fig. 1A). Luciferase translation was readily detected from both the HCV and FMDV IRES; the FMDV IRES was used as a positive control. Translation of the first cistron, CAT, was efficient in both transcripts. As the sequences used in the HCV IRES include part of the core-coding region, luciferase is translated from two initiator codons, as already noticed in similar constructs with different IRES of HCV (Sáiz et al., 1999 ). The higher molecular mass polypeptide is a fusion protein initiated at the HCV IRES AUG codon.

Then, the RNA region corresponding to the HCV IRES was divided in five transcripts, named II, IIIabcd, IIIabc, IIIef-IV and IV (Fig. 1B). In order to preserve the structure of each domain, each transcript was designed to contain stable stem–loop structures, according to mutational analysis carried out previously (Honda et al., 1996 , 1999 ). The transcript corresponding to the 5' end of the IRES encompasses nt 43–119 (stem–loop II). The central region was cloned in two forms, one containing nt 134–290 (stem–loop IIIabcd) and a second one, IIIabc, devoid of stem–loop IIId. The 3' region of the IRES was also designed in two forms. The first one, IIIef-IV, encompasses nt 290–383, including stem–loop IIIef and domain IV, fused to residues 125–134. This transcript was designed to mimic the RNA structure that allows the formation of the pseudoknot shown by Wang et al. (1995) . The second transcript contained domain IV alone (nt 125–130 fused to nt 318–383).

Following denaturation, pairs of RNA consisting of one 32P-labelled and one unlabelled transcript were incubated in binding buffer. Analysis of complex formation was carried out in native acrylamide gels, as described previously (Ramos & Martínez-Salas, 1999 ). Using domain II as probe, stable RNA–RNA complexes were detected with the different HCV transcripts tested, whose mobility changed according to the pair of RNA used in the assay (Fig. 2A). A weak dimerization of probe II increased in intensity when the concentration of transcript II was 800 nM (Fig. 2A, compare lanes 1 and 2), was always observed.



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Fig. 2. Requirement of magnesium ions for RNA–RNA interactions between separated domains of HCV IRES. (A) Autoradiography of complexes formed using 20 nM labelled domain II and 800 nM of the indicated unlabelled transcripts following electrophoretic separation in a 6% nondenaturing acrylamide gel run in TBM buffer. The lane marked ‘-’ shows the probe alone. (B) Effect of EDTA in complex disassociation: preassembled RNA complexes were analysed in TBE buffer. The TBE gel stained with ethidium bromide prior to drying and autoradiography is also shown (C).

 
Formation of RNA–RNA complexes required the presence of Mg2+ in the binding buffer as well as during the electrophoresis. Thus, the presence of EDTA in the running buffer during electrophoretic separation of preassembled complexes between domain II and the different transcripts led to a severe decrease in the retarded complexes relative to the gels containing Mg2+ ions (Fig. 2A, B, compare TBM with TBE autoradiography). The presence of the corresponding unlabelled transcripts, which remained unbound, was readily detected by ethidium bromide staining of the same gel (Fig. 2C).

The specificity of the interactions of the different transcripts with probe II was assessed by the lack of formation of retarded complexes with a transcript that included 80 nt of the 18S rRNA (pTRI-18S) or poly(I:C). Addition of these RNA molecules (800 nM) simultaneously with the probe and the specific interactor to the incubation mixture did not interfere with the formation of shifted bands observed in their absence (Fig. 3). In some experiments, the complex retarded with the full-length IRES led to the formation of a doublet rather than a single complex (Fig. 2A, compare the lanes labelled IRES with those in Fig. 3).



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Fig. 3. RNA–RNA interactions between separated domains of HCV IRES. 32P-labelled domain II (20 nM) was incubated in binding buffer with unlabelled RNA molecules (800 nM) corresponding to the sense sequences of domains II, IIIabcd, IIIabc, IIIef-IV, IV or IRES, in either the presence or the absence of the indicated nonspecific competitor RNA [800 nM 18S rRNA or poly(I:C)]. The lane marked ‘-’ shows the probe alone. Retarded complexes were fractionated in a 6% acrylamide gel in TBM buffer.

 
Domains II and IV cross-interact while the apical part of domain III barely interacts with them
Next we examined the complexes formed with labelled domain IV (Fig. 4A). Albeit with different intensity, retarded complexes were observed with all the RNAs tested: transcripts II, IIIabcd, IIIabc, IIIef-IV, itself and the whole IRES. A significant amount of the probe was present in the form of homodimers when a high concentration of transcript IV was added to the binding reaction. Moreover, the mobility of the IV–IV homodimer was similar to that of IV–II heterodimers (Fig. 4A). Yeast tRNA or poly(I:C) (800 nM) were coincubated with the unlabelled RNA and probe IV to assess the specificity of the retarded complexes (data not shown).



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Fig. 4. (A) Domain IV interaction with the HCV IRES transcripts. Probe IV (20 nM) was incubated in binding buffer with the indicated unlabelled RNA molecules (800 nM). (B) Increasing amounts of transcript IIIef-IV competes out IV–IIIabcd complex formation. Numbers at the top of the gel indicate the concentration of transcript IIIef-IV. Probe IV (20 nM) was used in the presence (+) or absence (-) of 800 nM transcript IIIabcd. The mobility of the respective retarded complexes, based on staining of the same gel, is marked on the left. (C) Incubation of transcripts II, IIIabcd and IV does not induce the formation of trimeric complexes. Probe IV (20 nM) was incubated alone (-), with IIIabcd transcript (800 nM) or with increasing amounts of domain II, as indicated at the top of the gel. Probe IV homodimer and the heterodimer II–IV comigrate.

 
Quantitative analysis of the retarded complexes formed between combinations of the different transcripts confirmed that their intensity varied with the pair of IRES domains used in the assay (Table 2; see also Figs 2A, 3 and 4A). Using 20 nM of each labelled domain and 800 nM of the unlabelled domain as probe, efficient formation of complexes was observed between domains II and IV and vice versa. In the latter, the total intensity of the retarded complex has been corrected for the amount of self-dimerization, as shown by probe IV at 20 nM. The efficiency of the interactions observed with probes IIIabc and IIIabcd showed a lack of symmetry as compared to the efficiency observed when they were used as the unlabelled RNA with probes II, IIIef-IV or IV (Table 2).


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Table 2. RNA–RNA interactions between HCV IRES domains

 
The intensity of complexes IIIef-IV with IV was a little higher than with the other domains. Domain IV showed the strongest interactions, as it interacted efficiently with domain IIIef-IV and itself, as well as with the whole IRES (Table 2; Fig. 4A). However, addition of sequences corresponding to stem–loop IIIef to domain IV led to a decrease in the efficiency of RNA–RNA interactions. These results strongly suggested that stem–loop IIIef forms a compact structure with sequences in domain IV, in agreement with the pseudoknot structure proposed by Wang et al. (1995) . In contrast, domain IIIabcd was a weak interactor, displaying only moderate binding with domain II. Interestingly, removal of stem–loop IIId from domain IIIabcd led to a significant reduction in its ability to interact with domain II (Table 2).

Competition studies between different HCV domains were carried out to determine whether formation of retarded complexes could be diminished by the presence of the other HCV IRES domains. Using domain IV as probe, increasing amounts of transcript IIIef-IV competed out efficiently the interaction between transcripts IIIabcd and IV (Fig. 4B). These results were fully consistent with those shown in Table 2, indicating not only that the interaction between transcripts IIIabcd and IV was weak but also that it was rapidly displaced by the interaction between transcripts IIIef-IV and IV.

Mixtures of transcripts II, IIIabcd and IV, representing most of the IRES domains, did not give raise to trimeric complexes (Fig. 4C) even under conditions of high concentrations of transcripts II and IIIabcd. This result suggested that formation of each heterodimer complex, II–IV and IIIabcd–IV, occurred independently of the third component in the mixture. Moreover, increasing concentrations of domain II reduced only slightly the intensity of the IIIabcd–IV complex, indicating that domain II was not an efficient competitor of the interaction between domains IV and IIIabcd. It has to be noticed that domain IV formed a homodimer that comigrated with complex II–IV.

The HCV IRES transcripts were able to self-dimerize to a lower extent than the FMDV IRES transcripts (Ramos & Martínez-Salas, 1999 ). The exception was domain IV, which reached values of about 30% (Table 2), although still below the self-dimerization capacity observed for the FMDV IRES domain 3 under similar conditions of RNA concentration and ionic strength. Remarkably, dimerization of domain IV was efficiently competed out by the presence of transcript IIIef-IV (Fig. 4B).

Hairpin IIId can interact with domain II
As shown in Table 2, a significant decrease in the interactions between the two versions of domain III, IIIabcd and IIIabc, and domain II was observed in the transcript devoid of stem–loop IIId. This result suggested that stem–loop IIId could contribute to the interactions mediated by domain IIIabcd with domain II. To test this possibility, a IIId sense RNA oligonucleotide (sIIId) was used to determine whether this short structure was able to interact with any of the HCV transcripts. Remarkably, using equimolar amounts of RNA to oligonucleotide, the 5' end-labelled sIIId interacted with domain II (Fig. 5). Two discrete bands appeared to be labelled after incubation of this RNA oligonucleotide with domain II, which may correspond to the monomer and the dimer of this transcript bound to the sIIId oligonucleotide, according to ethidium bromide-staining data.



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Fig. 5. Binding sites of stem–loop IIId. A 5' end 32P-labelled RNA oligonucleotide sIIId (20 nM) harbouring the sequence of stem–loop IIId (top panel) was incubated in binding buffer with equimolar concentrations of the transcripts, indicated at the top of the gel, and the complexes were fractionated through a 6% acrylamide gel in TBM buffer. The lower panel shows the percentage of label present in the retarded complex, averaged from the data from three independent experiments.

 
The observation that the oligonucleotide containing hairpin IIId produced a retarded band with transcript II was consistent with the decrease observed in binding of IIIabc to transcript II, relative to that of IIIabcd. Thus, the absence of stem–loop IIId in transcript III was responsible for the significant reduction in the intensity of the interaction between transcripts II and IIIabc.

In addition, interaction of the sIIId oligonucleotide with transcript IIIabc was observed (Fig. 5), suggesting the involvement of this hairpin in the interactions detected between the apical and basal part of domain III. The specificity of this contact was confirmed by the lack of binding with transcripts IIIef-IV and IV. Furthermore, labelled sIIId barely interacted with transcript IIIabcd (Fig. 5). This result could be interpreted as a direct involvement of IIId residues in RNA interactions between IIIabc and IIId or as a modified structure of IIIabcd transcript relative to IIIabc, which precluded IIId from binding.

Then, an antisense version of the IIId stem–loop sequence (asIIId; Table 1) was used to study the contribution of stem–loop IIId to the interactions between the apical and basal part of domain III. As expected, the asIIId oligoribonucleotide interacted very efficiently with transcript IIIabcd but not with transcript IIIabc (Fig. 6A). Additionally, asIIId interacted with transcript IIIef-IV, albeit to a lower extent.



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Fig. 6. Binding of IIId antisense oligonucleotide (asIIId) to its target sequence does not modify the interaction between transcripts IIIabcd and IIIef-IV. (A) Specific binding of the 5' end-labelled asIIId oligonucleotide to its target sequence in the HCV IRES, within a concentration range of 10–2·5 nM. (B) The molar excess of asIIId oligonucleotide indicated at the top of the gel was used to compete out the interactions between probe IIIef-IV (20 nM) and transcripts IIIabcd or IIIabc (800 nM). The gel stained with ethidium bromide shows the unlabelled transcript. As a consequence of close to 100% binding of transcript IIIabcd to the asIIId oligonucleotide, a 1:1 molar ratio shows a retarded mobility of the transcript, following both ethidium bromide staining and autoradiography.

 
Using domain IIIef-IV as probe, the interaction between domains IIIabcd and IIIef-IV was not interfered by increasing amounts of asIIId (Fig. 6B, left panel). Oligonucleotide asIIId was efficiently bound to transcript IIIabcd, as deduced from the shift of IIIabcd RNA (Fig. 6B, right panel). Thus, the IIId antisense oligonucleotide remained bound to its target sequence when transcript IIIabcd was interacting with probe IIIef-IV. Transcript IIIabc, which did not interact with asIIId, was used as a negative control in competition assays.

These results were consistent with the weak interaction between domains IIIabcd and IV, indicating that the apical part of domain III folds independently of stem–loops IIIef and IV. In agreement with this result, a mixture of three transcripts, domains II, IIIabcd and IV, did not yield trimeric complexes (Fig. 4C). Taking the results of binding and competition assays together, we concluded that the apical part of domain III folds independently of stem–loops IIIef and IV. Notably, stem–loop IIId was responsible for the interaction observed between domains II and IIIabcd but it did not seem to interact with domain IV or IIIef-IV.


   Discussion
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Abstract
Introduction
Methods
Results
Discussion
References
 
In this study, we present experimental evidence for long-range RNA interactions within distal regions of the HCV IRES by means of a gel mobility-shift assay. This technique has been used before to demonstrate long-distance interactions in several RNA molecules, including the 3' UTR of bicoid mRNA (Ferrandon et al., 1997 ), the dimerization signal of genomic human immunodeficiency virus type 1 RNA (Paillart et al., 1996 ), the hammerhead catalytic RNA (Fedor & Uhlenbeck, 1990 ) or the FMDV IRES (Ramos & Martínez-Salas, 1999 ).

The interactions shown here between domains II and IV in the absence of proteins provide support for the essential role played by domain II in HCV IRES activity. It is possible that these interactions are transient, being displaced by RNA-binding proteins or the small ribosomal subunit. However, domain IIIabc barely interacted with the rest of the IRES, indicating that it folds independently of domains II, IIIef-IV and IV, which is in agreement with data recently reported (Spahn et al., 2001 ; Beales et al., 2001 ).

On the other hand, the 40S ribosomal subunit produces an RNase T1 footprint in a guanine residue of the apical loop of domain II as well as three guanine residues of stem–loop IIId (Kieft et al., 2001 ). Thus, some proximity or association between these RNA regions is likely. Consistent with this, the reduction observed when stem–loop IIId was removed from transcript IIIabcd suggested that stem–loop IIId was able to interact with domain II. We tested this hypothesis by incubating a labelled RNA oligonucleotide carrying the IIId sense sequence with domain II. Remarkably, efficient binding was observed with sequences of domain II but not with domain IV or IIIef-IV, which is in agreement with the possibility mentioned above. These results were also consistent with the lack of competition shown by the antisense oligonucleotide asIIId. Binding of asIIId to its target sequence did not interfere with the interaction between transcripts IIIabcd and IIIef-IV. The IIId antisense oligonucleotide, which binds very efficiently to domain IIIabcd, remained bound to its target sequence when the latter was forming a complex with transcript IIIef-IV, which is in agreement with the conclusion that IIIabcd folds independently of stem–loop IIIef.

Cross-interaction was observed between domains II and IV, including the stem–loop that contains the initiator codon and the unstructured region at the beginning of the coding sequence. However, in some instances, the efficiency, but not the pattern, of the interactions observed display a lack of symmetry. This asymmetry is only observed with interactions exerted by probes IIIabc and IIIabcd, where the interactions are weaker. The reason for this asymmetry in the intensity of the interactions is not known but it can be attributed to the difference in concentration of probe and interactor RNA used in the assay.

The absence of stem–loop IIIef from transcript IV resulted in a significant increase in the interaction of transcript IV with the other RNAs. Therefore, stem–loop IIIef induced the formation of a compact RNA structure with domain IV, leading to a strong reduction in interactions with residues from outside the stem–loop. Although the exact residues involved in the interaction between domains IIIef and IV are not known yet, formation of a pseudoknot structure (Wang et al., 1995 ) may contribute to the significant reduction in binding between domain IV and the rest of the IRES.

Compared to FMDV IRES domain 3, the HCV transcripts were able to weakly self-interact. Of note is that domain IIIabcd is unable to self-dimerize, in spite of having a very long stem–loop at its base. Under our experimental conditions, the best self-interactor is domain IV, which could reach values of about 30%, close to those obtained with the whole HCV IRES transcript but far below the strong self-dimerization shown by domain 3 of FMDV (Ramos & Martínez-Salas, 1999 ). It is not known yet whether self-dimerization has something to do with IRES trans-complementation. However, it is interesting to note that there are published data reporting the inability of HCV IRES mutants to complement in trans, as opposed to FMDV IRES (Tang et al., 1999 ).

We have shown here cross-interactions between domains II and IV, corresponding to the 5' and 3' distal regions of the HCV IRES. This is also in contrast to the FMDV IRES, where the central domain was the one interacting strongly with all the others, at least in the absence of proteins (Ramos & Martínez-Salas, 1999 ). Therefore, a very different structural organization adopted by FMDV and HCV IRES allows efficient internal initiation of translation.

The IRES of FMDV and HCV adopt different structural organizations that reflect a different manner to promote internal initiation. Accordingly, a different pattern of RNA–protein interaction is observed for each of these IRES elements (López de Quinto et al., 2001 ; Kieft et al., 2001 ; Pestova et al., 1998 ; López de Quinto & Martínez-Salas, 2000 ; Buratti et al., 1998 ; Sizova et al., 1998 ; Kolupaeva et al., 2000 ; Pilipenko et al., 2000 ). Results of toe-print analysis indicated that 48S initiation complex formation driven by the HCV and CSFV IRES required eIF2-GTP/Met-tRNAi, eIF3 and 40S subunits (Pestova et al., 1998 ). As a consequence of the use of different mechanisms to initiate translation, HCV IRES does not require eIF4G to assemble a 48S initiation complex, whereas FMDV does (reviewed by Martínez-Salas et al., 2001 ). This observation poses the question of how the RNA present in the HCV IRES recognizes the translational machinery. A direct contact between the HCV IRES and the 40S ribosomal subunit has been demonstrated recently (Spahn et al., 2001 ). In agreement with the IRES structural model derived from the latter study, the apical part of domain III, which binds eIF3 (Kieft et al., 2001 ; Buratti et al., 1998 ; Sizova et al., 1998 ), folds independently of the rest of the IRES.

In summary, the results shown here indicate that the apical part of domain III forms a structural element separate from domains II and IV. While domains II and IV can interact, at least in the absence of proteins, stem–loop IIIef induces the formation of a compact structure with domain IV, which strongly reduces its binding to other regions of the IRES. Furthermore, we have also shown that the interaction observed between domains IIIabcd and II is facilitated by stem–loop IIId. It has been shown that IIId sequences form part of the structural motif involved in the recognition of the ribosomal subunit (Jubin et al., 2000 ; Kieft et al., 2001 ; Spahn et al., 2001 ; Lytle et al., 2001 ). Thus, it is likely that the interactions we have observed between domains II and IIId may play a regulatory role in the activity of this IRES element, modulating its capacity to interact with the ribosome.


   Acknowledgments
 
We are grateful to N. Ibarrola for the preparation of construct p156, which harbours the HCV IRES; R. Moreno-Otero for HCV sample supply; J.-J. Toulmé for the synthesis of RNA oligonucleotides sIIId and asIIId and E. Cano for excellent laboratory assistance. We also thank J.-J. Toulmé, S. López de Quinto and C. Gutiérrez for helpful suggestions on the manuscript. This work was partially supported by grants PM98.0122 from DGES, HF1999–0023 concerted action between France and Spain, 08.2/0024/1997 from CAM and by an institutional grant from Fundación Ramón Areces.


   References
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Abstract
Introduction
Methods
Results
Discussion
References
 
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Received 25 October 2001; accepted 18 January 2002.