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Originally published In Press as doi:10.1074/jbc.M408294200 on September 8, 2004

J. Biol. Chem., Vol. 279, Issue 46, 48397-48403, November 12, 2004
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First Snapshots of the HIV-1 RNA Structure in Infected Cells and in Virions*

Jean-Christophe Paillart{ddagger}§, Markus Dettenhofer¶||, Xiao-fang Yu¶, Chantal Ehresmann{ddagger}, Bernard Ehresmann{ddagger}, and Roland Marquet{ddagger}

From the {ddagger}Unité Propre de Recherche 9002 du CNRS conventionnée à l'Université Louis Pasteur, Institut de Biologie Moléculaire et Cellulaire, 15 rue René Descartes, 67084 Strasbourg cedex, France and the Department of Molecular Microbiology and Immunology, Johns Hopkins University School of Hygiene and Public Health, Baltimore, Maryland 21205

Received for publication, July 22, 2004 , and in revised form, September 7, 2004.


   ABSTRACT
TOP
 ABSTRACT
INTRODUCTION
 EXPERIMENTAL PROCEDURES
 RESULTS AND DISCUSSION
 REFERENCES
 
With the increasing interest of RNAs in regulating a range of cell biological processes, very little is known about the structure of RNAs in tissue culture cells. We focused on the 5'-untranslated region of the human immunodeficiency virus type 1 RNA genome, a highly conserved RNA region, which contains structural domains that regulate key steps in the viral replication cycle. Up until now, structural information only came from in vitro studies. Here, we developed chemical modification assays to test nucleotide accessibility directly in infected cells and viral particles, thus circumventing possible biases and artifacts linked to in vitro assays. The secondary structure of the 5'-untranslated region in infected cells points to the existence of the various stem-loop motifs associated to distinct functions, proposed from in vitro probing, mutagenesis, and phylogeny. However, compared with in vitro data, subtle differences were observed in the dimerization initiation site hairpin, and none of the proposed long range interactions were observed between the functional domains. Moreover, no global RNA rearrangement was observed; structural differences between infected cells and viral particles were limited to the primer binding site, which became protected against chemical modification upon tRNA3 Lys annealing in virions and to the main packaging signal. In addition, our data suggested that the genomic RNA could already dimerize in the cytoplasm of infected cells. Taken together, our results provided the first analysis of the dynamic of RNA structure of the human immunodeficiency virus type 1 RNA genome during virus assembly ex vivo.


   INTRODUCTION
TOP
 ABSTRACT
 INTRODUCTION
EXPERIMENTAL PROCEDURES
 RESULTS AND DISCUSSION
 REFERENCES
 
In retroviruses, including human immunodeficiency virus type 1 (HIV-1),1 the full-length unspliced transcript is used as a pre-mRNA (for splicing and export), mRNA (for synthesis of Gag and Gag-Pol precursor proteins), and genome for packaging as an RNA dimer into infectious particles. The 5'-untranslated region (UTR) of HIV-1 RNA is the most conserved part of the genome (1). It contains a number of cis-acting sequences recognized by proteins or RNAs that regulate crucial steps of the viral life cycle (2) (see Fig. 1A), transcription, polyadenylation, splicing, nuclear export, translation, RNA dimerization and packaging, and reverse transcription.



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  		FIG. 1.
Schematic representation of the 5'-untranslated region of HIV-1 genomic RNA. A, the regulatory motifs are represented by gray color boxes on the linear map and localized on the secondary structure models proposed by B (2, 5) and C (6–8). The long distance interaction (LDI) and branched multiple hairpin (BMH) conformations have been proposed to be involved in a riboswitch regulating various steps in the viral life cycle (see Introduction). R, repeat in the 5'- and 3'-end of the RNA genome; U5, unique in 5'-end; AUG, translation initiation codon.

 
The secondary structure of the 5'-UTR of HIV-1 genomic RNA has been extensively studied by biochemical, phylogenetic, and mutational approaches (2–7), and there is a consensus concerning the existence of independent structural domains such as the trans-activation region (TAR), the polyadenylation signal hairpin (poly(A)), the primer binding site (PBS) domain, and three stem-loops containing the dimerization initiation site (DIS or SL1), the splice donor site (SD or SL2), and the so-called packaging signal ({Psi} or SL3) (see Fig. 1B).

Depending on the experimental conditions, the 5'-UTR of HIV-1 (B-subtype) can adopt two alternative conformations in vitro (6, 8, 9), a dimerization-competent conformation (BMH, branched multiple hairpins) and a thermodynamically more stable dimerization-incompetent structure (LDI, long distance interaction) in which the DIS loop is masked by the poly(A) domain (see Fig. 1C). It has been suggested that a RNA switch may precisely coordinate the functions associated to the 5'-UTR functions in time and space. Two long range interactions have been identified in the BMH conformer in vitro, a pseudoknot between the apical loop of the poly(A) hairpin and a region within the Gag open reading frame (7, 10), and a base-pairing of nucleotides linking the poly(A) and PBS domains with the Gag start codon region (7, 11) (see Fig. 1C, orange). These long distance interactions can only exist in the BMH conformation, and they could regulate the equilibrium between the BMH and LDI conformers.

Beside these intramolecular interactions, symmetric intermolecular interactions take place in vitro at the DIS (12–14) and could also occur at TAR (15) and poly(A) hairpin (16). In addition, tRNA3 Lys, the reverse transcription primer, is selectively encapsidated into virions and is annealed to the PBS during or after budding (17, 18). In vitro studies suggested that tRNA3 Lys also interacts with viral sequences upstream of the PBS (19–21), but no consensus exists concerning these additional interactions (18). Furthermore, the 5'-UTR binds several viral proteins, such as Tat (22, 23), the nucleocapsid (NC) protein (24) and the viral infectivity factor (25). Notably, the NC protein is a nucleic acid chaperone that activates RNA refolding events (24) and acts, as part of the Gag precursor, as a positive trans-acting factor in genomic RNA packaging (26).

Despite an increased knowledge of in vitro RNA secondary and tertiary structures, the structure of the 5'-UTR has never been directly investigated in tissue cell culture. To test this structure and the complex rearrangements that could take place during viral assembly, we analyzed the RNA conformation directly in infected cells and in viral particles by in situ chemical probing with dimethyl sulfate (Me2SO4) and used our data to test the models derived from in vitro structural studies.


   EXPERIMENTAL PROCEDURES
TOP
 ABSTRACT
 INTRODUCTION
 EXPERIMENTAL PROCEDURES
RESULTS AND DISCUSSION
 REFERENCES
 
Plasmids and Cell Culture
The construction of HXB2NEO, which carries the neomycin phosphotransferase gene (neoR) cloned into the nef region of the genome has been described elsewhere (27). H9 cells chronically infected with HXB2NEO were grown in complete RPMI 1640 medium supplemented with 10% fetal bovine serum and G418 (1 mg/ml).

Reverse Transcriptase Assays
Progeny viruses were quantified using a micro reverse transcriptase assay, as described previously (28).

Immunoblotting
Immunoblotting was performed as described in Ref. 29, except that viral pellets were lysed in radioimmune precipitation assay buffer (140 mM NaCl, 8 mM Na2HPO4, 2 mM NaH2PO4, 1% Nonidet P-40, 0.5% sodium deoxycholate, 0.05% SDS). Proteins were detected with HIV-1-positive patient serum (1:1000).

Native Northern blotting
Virion pellets were resuspended in 300 µl of 10 mM Tris-HCl, pH 7.5, 1 mM EDTA, 100 mM NaCl, 1% SDS, 100 µg of proteinase K/ml for 20 min at 37 °C. After phenol-chloroform extraction and ethanol precipitation, RNA pellets were washed twice with 70% ethanol and dissolved in 10 mM Tris-HCl, pH 7.5, 1 mM EDTA, 1% SDS, and 100 mM NaCl. RNA was fractionated on 0.9% agarose gels in 1x Tris-HCl/borate/EDTA buffer at 100 V for 2–3 h at 4 °C. The agarose gel was incubated at 65 °C for 30 min in 10% formaldehyde, and samples were transferred for2hat4 °C onto a Hybond-XL membrane (Amersham Biosciences) in 17 mM NaH2PO4, 8 mM Na2HPO4. The membrane was exposed to UV light for RNA cross-linking. After a 1-h blocking at 42 °C in 10 ml of ULTRAhybTM (Ambion), the membrane was probed overnight at 42 °C using an [{alpha}32P]dCTP-labeled HIV-1 DNA probe (nucleotide 1–2000) prepared as recommended by the manufacturer (DECAprimeTM II, Ambion). After hybridization, the membrane was washed twice for 5 min with 2x SSC (300 mM NaCl, 30 mM sodium citrate)-0.1% SDS buffer and twice for 15 min with 0.1x SSC-0.1% SDS buffer and submitted to autoradiography.

Analysis of Virion RNA Dimerization
Aliquots of RNA were heated at the indicated temperature for 10 min and quickly chilling in ice. Heat-denatured monomeric and dimeric RNAs were separated by electrophoresis through a 0.9% native-agarose gel and analyzed by Northern blotting as described above.

Ex Vivo RNA Modification
Chronically infected H9 cells were washed twice with 1x phosphate-buffered saline and suspended in 30 µl of 1x phosphate-buffered saline. Progeny viruses were pelleted as above and resuspended in 90 µl of 1x phosphate-buffered saline. Cells and viruses were treated with 3 µl of Me2SO4 (Acros Organics) at 37 °C for 4 and 8 min. Reactions were stopped with 1 ml of TriReagent (Molecular Research Center), and RNA was extracted as indicated by the supplier. Modified bases were detected by extension with an avian myeloblastosis virus reverse transcriptase (Qbiogene) of 5'-[{gamma}32P]ATP-labeled primers as described in Ref. 30. In vitro Me2SO4 modification of 1–500 HxB2 RNA was performed as described previously with the same primers as above (30).


   RESULTS AND DISCUSSION
TOP
 ABSTRACT
 INTRODUCTION
 EXPERIMENTAL PROCEDURES
 RESULTS AND DISCUSSION
REFERENCES
 
In vitro, the 5'-UTR of the HIV-1 genomic RNA folds into a series of independent stem-loop structures (TAR, poly(A), DIS, SD, {Psi}) or domains (PBS) (2–5, 7, 11) maintained by long distance interactions (7, 10, 11) (Fig. 1). Recently, it was proposed that exclusive RNA conformers, LDI and BMH (Fig. 1), may regulate crucial steps in the viral life cycle (6, 8, 9). To test the relevance of these models to the HIV-1 RNA structure ex vivo, we set up conditions to probe the 5'-UTR structure in infected cells and in viral particles. Except for a limited study on murine leukemia virus (31), the results described below represent the first successful in situ probing of a 9–10-kb retroviral RNA. This technique allowed us to 1) circumvent all in vitro possible biases and artifacts due to the renaturation protocols, the ionic strength conditions, the length of the RNA fragments, or the use of mutants that can induce aberrant RNA folding, and 2) provide a snapshot of viral RNA structure in cells (before packaging and tRNA3 Lys annealing) and in mature virions (after packaging and primer annealing). Me2SO4 penetrates cells without permeabilization treatments, thus allowing RNA modification under nearly cell conditions.

Chronically infected H9 cells and purified virions were treated with Me2SO4 (see "Experimental Procedures"), and the RNA was extracted after modification. By using an extension assay with primers complementary to sequences located 3' to the major splice donor site, we were able to specifically detect modified nucleotides in the full-length HIV-1 genomic RNA. Me2SO4 modifies adenines at position N-1 and cytosines at N-3 when the Watson-Crick side of these nucleotides is not base-paired (32). Of note, the H9 cell line does not show any defect in protein synthesis and polyprotein proteolysis (data not shown) (27, 33).

The in Situ Structure of HIV-1 5'-UTR Is Very Similar to the in Vitro Structure
Typical experiments covering the PBS domain and the SL1, SL2, and SL3 hairpins are shown in Fig. 2. These gels show that the quality of the in situ probing data is very good, and we were able to determine the reactivity level of most adenines and cytosines in cells and viruses. These levels were categorized as highly reactive, weakly reactive, and unreactive and reported on the 5'-UTR secondary structure model (Fig. 3). Importantly, our probing experiments on infected cells show that the main structural domains derived from in vitro experiments also exist ex vivo indicating the rather limited structural differences discussed below.



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  		FIG. 2.
Structure-probing data of HXB2NEO RNA in H9 cells and in viral particles on region 110–192 (A), 200–296 (B), and 281–336 (C). Modification of A(N1) and C(N3) by Me2SO4 was performed according to the "Experimental Procedures" for 0 (unmodified control), 5, 10, and 20 min. Sequence lanes (U, A, C, and G) are run in parallel. Regulatory motifs are indicated on each side of the gels. Black triangle, corresponds to protection against Me2SO4 modification observed in viral particles; P,to reverse transcriptase pauses.

 



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  		FIG. 3.
Secondary structure model of the 5'-untranslated region of HIV-1 genomic RNA in infected cells and in virions. The reactivity at the tested Watson-Crick positions (A(N1), C(N3)) are indicated. The functional motifs are indicated above each stem-loop domain. Long distance interactions previously proposed by Abbink and Berkhout (11), Harisson and Lever (3), and Paillart et al. (10) are represented by brown, blue, and green bars, respectively (the RNA partner for the long distance interaction proposed by Paillart et al. is not present on this model). To facilitate discussion, the PBS domain has been dissected in four structural elements named PBS1 (lower stem), PBS2 (middle stem), PBS3 (A-rich loop stem), and PBS4 (the PBS loop). PAS (pink-purple) and A-rich loop (pink) have been proposed to provide secondary interactions with tRNAlys3 upon binding the PBS.

 
The TAR and poly(A) Hairpins—These domains include TAR, the binding site of Tat, and the poly(A) hairpin that is critical for repression of the 5'-copy of the polyadenylation signal (34). These hairpins are strongly constrained among HIV-1 strains (2). In TAR, the predicted bulges were highly reactive, whereas in the loop, the only highly reactive nucleotide was A34 (Fig. 3), which is in keeping with in vitro chemical probing experiments performed in the absence of proteins (5, 35). Nucleotide C23 located in the Tat binding site was strongly modified (Fig. 3). The in vitro binding of a Tat-derived peptide to this three-nucleotide bulge strongly protected this residue against Me2SO4 modification (35). Thus, our results suggested that Tat and the P-TEFb complex (positive transcription elongation factor complex b) that binds the TAR apical loop during transcription trans-activation (36) do not remain associated with the genomic RNA. To note, this stem is involved in the packaging of the genomic RNA (37, 38).

Modifications in the poly(A) hairpin are consistent with the proposed secondary structure model. Noticeably, although nucleotides A76, A77, and A78 located in the 5' part of the poly(A) loop were highly reactive, modification of C80, which is also located in the loop was weak (Fig. 3). The low reactivity of C80 might be the consequence of a long range pseudoknot between nucleotides G79 to C85 and a sequence located in the matrix coding sequence (from G441 to C447) (10). Ex vivo probing of the matrix coding region will be required to further test this interaction.

The PBS Domain—The PBS domain can be separated into four different subdomains that we named PBS1–PBS4 for clarity (Fig. 3). In situ Me2SO4 modification of HXB2NEO RNA did not reveal any significant difference with in vitro experiments performed with HXB2 RNA (39, 40). Indeed, all adenines and cytosines that are unpaired in vitro are modified by Me2SO4 in infected cells, in agreement with in situ Me2SO4 probing of the PBS domain of the NL4.3 isolate (41). Thus, the reactivity profile is generally consistent with the proposed secondary structure model, with the exception of the PBS1 stem that either does not exist as such or is strongly breathing, suggesting a possible interaction with a RNA or protein partner as suggested by Laughrea and co-workers (42).

The Leader Region—This region (240–355) encompasses the DIS, SD, and {Psi} hairpins that are involved in RNA dimerization, splicing, and encapsidation, respectively (Fig. 3) (2, 43).

In vitro, the DIS promotes dimerization of HIV-1 RNA by forming a kissing-loop complex via intermolecular base pairing of the self-complementary sequence in the DIS loop. This complex might eventually be converted into a more stable extended duplex (for review, see Ref. 43). In vitro RNA dimerization protects the self-complementary sequence (nucleotides 257–262) from chemical modification, whereas the surrounding purines display a characteristic modification profile (5, 30). Remarkably, the DIS loop displays the same reactivity pattern in infected cells (Fig. 2B). In particular, hyperreactivity of the last adenine in the DIS loop (A263), which is the hallmark of in vitro RNA dimerization (5, 30), was also observed in infected cells (Fig. 2B). Thus, our results suggest that HIV-1 genomic RNA might already be dimeric in the infected cells. Alternatively, they could indicate that most cellular viral RNA is associated with budding particles. However, the complete absence of Me2SO4 modification of the self-complementary sequence indicates that the DIS structure is homogenous thus favoring the first interpretation. Moreover, fractionation experiments showed that a significant part of the genomic RNA is nuclear (not shown). Furthermore, our data indicated that the PBS of the cellular RNA is not occupied by tRNA3 Lys. As tRNA3 Lys annealing occurs early in the budding process (i.e. before processing of the Gag-Pol precursors (44)), we think little RNA from budding virions is present in our cellular RNA. In any case, our data indicated that intermolecular contacts at the DIS occur very early or even before budding of the viral particles.

A main difference between ex vivo and in vitro probing data resides at nucleotides C267–A271. Whereas these nucleotides are not reactive in vitro (5, 30), they show a weak but reproducible reactivity toward Me2SO4 in infected cells (Figs. 2B and 3). These nucleotides are located in the DIS upper stem, adjacent to the conserved internal loop B formed by nucleotides 247 and 271–273 (Fig. 3) (1). Interestingly, this internal loop has a direct impact on RNA dimerization, RNA packaging, and viral infectivity (4, 45–47). Recent in vitro NMR and biochemical studies suggested that loop B might function by destabilizing the DIS stem (48, 49), possibly facilitating a conversion of the kissing-loop dimer into a more stable duplex. Our data clearly showed that this part of the upper DIS stem is structurally breathing, suggesting a possible interaction with viral and/or cellular proteins or its direct involvement in tertiary RNA folding.

We clearly observed the signatures of the SD and {Psi} stem-loop motifs (Figs. 2C and 3), based on in vitro experiments (2–5, 7). The bulging out nucleotide A296, the opposite nucleotide A286 in the SD stem, and A319 in the {Psi} loop are accessible to Me2SO4 in situ (Fig. 2C). Primer extension from infected cells yielded a major reverse transcriptase pause site. Two possibilities can explain this result. Either reverse transcriptase is stopped by a very stable structure specific for the genomic RNA in cells, or a significant part of the RNA is specifically cut at this position.

The Interdomain Linkers and the Long Distance Interactions—In the secondary structure model, the structured domains described above are joined by single-stranded linkers (Fig. 3). In keeping with this model, C110, C111, C238, A239, A242, C281, A301–A305, A326, A327, A330, A332, A334, and A336 are strongly modified by Me2SO4 in cells. These data argue against the long distance base pairing proposed between nucleotides 105–115 and 334–344 in the context of the BMH conformer (7, 11) (Figs. 1 and 3, orange). Similarly, they do not support base pairing between nucleotides 227–231 and 331–335 (Fig. 3, blue bars) (3, 50). However, one cannot exclude that these interactions exist in a minor RNA population or that they only exist transiently.

No Evidence for the LDI Conformation
In vitro, the 5'-UTR of HIV-1 genomic RNA can adopt alternative conformations named LDI and BMH (Fig. 1) (6, 8, 9). It has been proposed that a NC-mediated LDI-to-BMH switch would inhibit translation and favor dimerization and encapsidation (6, 8, 11). To check the existence of the LDI conformer in cells, we reported our probing data on the parts of the LDI secondary structure that differ from the BMH conformation (Fig. 4). Several nucleotides that are base-paired in the LDI model were highly reactive. The reactivity of C110 and A263 could be explained by the presence of an adjacent bulge. However, the strong reactivities of C96, A241, A255, and A256 are difficult to reconcile with the LDI conformation (Fig. 4B). Conversely, nucleotides A89, A98, A101, C245, and C264 that should be modified in the LDI conformation were completely unreactive. Importantly, those nucleotides would give a detectable signal if they were reactive in a minor population corresponding to 5–10% of the total RNA (32). Concerning SD, {Psi}, and the initiation codon of gag (Fig. 4C), there are fewer discrepancies. However, the LDI conformation does not explain the high reactivity of A296. Taken together our data showed that the major fraction of the genomic RNA associated with cells is in the dimerization and packaging-competent BMH form, and they argued against the existence of the LDI conformer, although we cannot formally rule out that this conformation exists in a minor population or during a short period of time.



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  		FIG. 4.
The LDI model proposed by Abbink and Berkhout (11). A, the regulatory motifs are marked in colors.Me2SO4-modified nucleotides obtained in infected cells and viral particles (this study) were reported on domains (B and C, boxed) undergoing conformational changes between LDI and BMH models (see Fig. 1). Color code for intensity of Me2SO4 modification is the same as in the legend to Fig. 3.

 
The Genomic RNA Undergoes a Few Limited Structural Rearrangements after Budding
To identify structural differences between pre- (in cells) and postbudding genomic RNA, virions were chemically treated with Me2SO4. Our first concern was to establish that the encapsidated HXB2NEO RNA was in native dimeric state. We isolated genomic RNA from HXB2NEO virions and performed a nondenaturing Northern blot analysis. Only one RNA species with a mobility corresponding to the expected RNA dimer was present in untreated samples (Fig. 5, lane C). We measured the thermal stability of the RNA dimer by increasing the temperature from 37 to 65 °C. The dimeric RNA was completely converted into monomers after incubation at 58 °C (Fig. 5), giving a Tm of 54–55 °C, as previously determined with several subtype B isolates (42, 50, 51). These results indicated that the HXB2NEO genomic dimer exhibits wild-type properties.



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  		FIG. 5.
Northern blot analysis of HXB2NEO virion RNA content. The nature of genomic RNA packaged into HXB2NEO viral particles was monitored using melting curves and electrophoresis analysis of viral RNAs. Virion RNA was resuspended in Tris-HCl/EDTA/NaCl buffer (see "Experimental Procedures") and heat-denatured for 10 min at the indicated temperature. Dimers and monomers were electrophoresed in a 0.9% native agarose gel and probed with an HIV-1 DNA fragment corresponding to region 1–2000 of HIV-1 RNA.

 
The {Psi} hairpin is the major packaging signal of HIV-1 RNA (52, 53), but TAR, the poly(A) hairpin, the PBS domain, the DIS, and the SD hairpins all contribute to the packaging efficiency (Ref. 54 and references therein). Thus, major structural rearrangements of HIV-1 RNA could occur during budding and/or the maturation of viral particles. Remarkably, only a few limited structural differences were observed between cells and virions (Figs. 2 and 3).

No difference was observed between cells and virions in the TAR loop, despite the fact that NC induces TAR-TAR dimerization in vitro (15). Similarly, the reactivity of the poly(A) loop was unchanged, although this hairpin has been proposed to participate in the RNA dimerization process (16).

Nucleotides in the PBS that were reactive in cells (C185, C189, A191, and A192) were protected in virions, as expected from tRNA3 Lys placement onto the PBS during or immediately after budding (17). Beside the PBS, tRNA3 Lys has been proposed to interact with the A-rich loop (Fig. 3, nucleotides 169–172, pink) (19, 55) and with the primer activation signal (nucleotides 123–130, pink/purple) (21, 56). However, there is no evidence for any additional interaction between viral RNA with tRNA3 Lys in virions. Indeed, the A-rich loop, which has been shown to interact in vitro with the anticodon loop of tRNA3 Lys in HIV-1 MAL (19, 55), was not protected in virions. Similarly, we observed no Me2SO4 reactivity changes in the PAS, which has been proposed to interact with the T{Psi}C arm of tRNA3 Lys, or in the opposite stem sequence (nucleotides 220–226) in cells and viruses (Fig. 2B). Taken together, these data indicated that HXB2NEO genomic RNA underwent little, if any, structural rearrangement during formation of the reverse transcription initiation complex. This conclusion is in keeping with comparative in vitro and ex vivo structural probing and in vitro functional comparison of the HIV-1 NL4.3 and MAL isolates (41). This study suggests that the interaction between the A-loop and the anticodon loop of tRNA3 Lys only takes place in a subset of HIV-1 isolates, including MAL, whereas no interaction between the viral RNA and the primer tRNA is required outside the PBS for optimal initiation of reverse transcription of subtype B isolates, including NL4.3 and HXB2.

As for other retroviruses (for review see Ref. 43), the dimer of HIV-1 genomic RNA undergoes a thermal stabilization upon proteolysis of the Gag precursor (51, 57). Based on in vitro experiments, it has been proposed that this stabilization could correspond to the transition of the initial kissing-loop complex formed at the DIS into an extended duplex. However, the thermal stabilization of the mature RNA dimer does not necessary imply a transition at the DIS, as other parts of the RNA genome may be involved (for review see Ref. 43). Quite surprisingly, in situ probing of the DIS revealed no structural reactivity differences between cells and virions (Figs. 2B and 3). Because the palindromic sequence in the DIS loop is protected in virions, we conclude that the intermolecular interaction at the DIS is preserved in the mature RNA dimer. However, we cannot make conclusions about the stabilization mechanism. On one hand, additional contacts can occur all along the genomic RNA in regions that we did not probe. On the other hand, Me2SO4 probing is not useful to distinguish the kissing-loop complex from the extended duplex, as the same nucleotides (A255, A256, and A263) are expected to be reactive in the two structures (43). Experiments are in progress to test the formation of an extended duplex in mature virions. Although the SL3 hairpin is essential for RNA packaging (52, 53), only one nucleotide (C316) was protected in this stem loop in virions, as compared with cells (Fig. 2C). This protection could reflect Gag binding to the high affinity purine-rich 317GGAG320 loop, probably through interactions with the NC domain of Gag similar to those described in the structure of the NC-SL3 complex (58).

Our results showed that the overall secondary structure of the 5'-UTR is very similar in infected cells and in virions and also very close to the in vitro RNA structure, except for the absence of long range base pairing. These results are surprising, because the genomic RNA is covered by NC protein in virions but not in cells, and in vitro studies were performed in the absence of proteins (2–5, 7). They do not explain why we do not see extensive NC footprints on genomic RNA in virions. A possible explanation could be protein inactivation by Me2SO4 (59). However, we used short Me2SO4 reaction times (5 or 10 min), while NC protein retains its zinc chelating and RNA binding capabilities after a 30 min reaction (60, 61). Most likely, NC mainly binds the RNA ribose-phosphate backbone, thereby providing protection against nucleases (62). Specific interactions with bases likely take place at a limited number of high affinity binding sites (58), however the preferred sequence is (UG)n (63), and we only tested A and C residues in this study. Finally, it is possible that the genomic RNA is not completely protected by Gag (NC domain) (64), or alternatively, the NC domain may be binding the RNA shortly after protein synthesis within the cytoplasm of infected cells (65), which could explain the similarities between cell and virion RNA structure, supporting the idea that RNA-NC contact points do not differ much between cell and virions.

Concluding Remarks
The 5'-UTR is the most highly conserved region of the HIV-1 genome, and it regulates several key steps in the viral life cycle. Despite numerous previous studies (2–5,7), our work constitutes the first direct investigation of the 5'-UTR structure of the full-length HIV-1 genomic RNA in infected cells and virions. This region displays very similar structures in situ and in vitro (this work and Refs 2–5 and 7). However, we noticed destabilization of the DIS stem in infected cells and virions. Similarly, we did not observed the previously proposed long range base pairings between regions connecting the functional domains (7, 11). These regions might interact with viral and/or cellular factors. In addition, our results also suggest that the genomic RNA is already dimeric in the cytoplasm of infected cells.

When comparing the 5'-UTR structure in cells and virions, we only observed a few limited differences in the PBS and in SL3. In the PBS domain, we observed the interaction of tRNA3 Lys with the PBS but none of the proposed additional interactions (19, 20, 39, 66). This result is in keeping with a recent comparative in vitro and ex vivo comparison of the HIV-1 NL4.3 and MAL isolates (41), which suggested that no interaction between the genomic RNA and tRNA3 Lys is required outside the PBS for optimal initiation of reverse transcription of subtype B isolates. Finally, our results do not favor the proposed LDI conformer (6, 8, 9, 11), and although one cannot exclude that minor conformations are present at low concentration or with a short half-life, we managed to test the major conformation in intact cells and virions without the need for RNA renaturation and avoiding any possible artifacts because of the ionic conditions or the short length of RNA transcripts. These are generally considered as the main weaknesses of the in vitro experiments. This study can be viewed as a starting point to dissect mechanisms involving RNA-RNA interactions such as the hypothetical transition from the DIS kissing-loop complex to an extended duplex, and the maturation of the genomic RNA after particle budding. Structural information might facilitate efforts to develop therapeutic strategies targeting these steps.

Because the LDI conformation is more stable than the BMH (6, 8), the ex vivo structure of the 5'-UTR is not the most thermodynamically stable, suggesting that it is kinetically trapped during transcription. Consistent with a co-transcriptional 5'–3' directional folding, local hairpin structures should be favored to the detriment of long range interactions that would require the melting of preformed structures. This strongly limits the use of algorithms such as Mfold (67) to predict the effects of mutations on the 5'-UTR structure. Indeed, the in situ probing technique described in this study will allow a direct testing of these effects and will allow distinguishing direct effects of mutations from indirect structural effects.


   FOOTNOTES
 
* This work was supported by grants from the Agence Nationale de Recherches sur le SIDA (ANRS) (to C. E. and R. M.). The costs of publication of this article were defrayed in part by the payment of page charges. This article must therefore be hereby marked "advertisement" in accordance with 18 U.S.C. Section 1734 solely to indicate this fact. Back

|| Present address: Dept. of Genetics, Howard Hughes Medical Institute, Harvard Medical School, Boston, MA 02115. Back

§ To whom correspondence should be addressed: UPR 9002-CNRS, Institut de Biologie Moléculaire et Cellulaire, 15, rue René Descartes, 67084 Strasbourg cedex, France. Tel.: 330-3-88-41-70-35; Fax: 330-3-88-60-22-18; E-mail: jc.paillart{at}ibmc.u-strasbg.fr.

1 The abbreviations used are: HIV-1, human immunodeficiency virus type 1; UTR, untranslated region; TAR, trans-activation region; PBS, primer binding site; DIS, dimerization initiation site; BMH, branched multiple hairpins; LDI, long distance interaction; NC, nucleocapsid; SD, splice donor. Back


   ACKNOWLEDGMENTS
 
We thank Dr. Alan Rein for critical reading of the manuscript.



   REFERENCES
TOP
 ABSTRACT
 INTRODUCTION
 EXPERIMENTAL PROCEDURES
 RESULTS AND DISCUSSION
 REFERENCES
 

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