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Kaposi's sarcoma–associated herpesvirus produces a highly abundant, nuclear noncoding RNA, polyadenylated nuclear (PAN) RNA, which contains an element that prevents its decay. The 79-nucleotide expression and nuclear retention element (ENE) was proposed to adopt a secondary structure like that of a box H/ACA small nucleolar RNA (snoRNA), with a U-rich internal loop that hybridizes to and protects the PAN RNA poly(A) tail. The crystal structure of a complex between the 40-nucleotide ENE core and oligo(A)9 RNA at 2.5 angstrom resolution reveals that unlike snoRNAs, the U-rich loop of the ENE engages its target through formation of a major-groove triple helix. A-minor interactions extend the binding interface. Deadenylation assays confirm the functional importance of the triple helix. Thus, the ENE acts as an intramolecular RNA clamp, sequestering the PAN poly(A) tail and preventing the initiation of RNA decay.
Kaposi's sarcoma–associated herpesvirus (KSHV) is the causative agent of Kaposi's sarcoma (KS), the most common AIDS-associated cancer (1, 2). Although largely controlled in the developed world by antiretroviral treatment against HIV, KS has become one of the most prevalent cancers in Africa (3). KSHV is a γ-herpesvirus that exists in either a latent or lytic state. During the lytic phase, KSHV produces PAN (polyadenylated nuclear) RNA, a 1.1-kb noncoding RNA with a 5′ cap and 3′ poly(A) tail (4-6) that is retained in the nucleus of infected cells. Although the function of PAN RNA is not known, it accumulates to extremely high levels, amounting to as much as 80% of the polyadenylated RNA in the cell (4, 6). The expression and nuclear retention element (ENE), a 79-nucleotide (nt) element located near the 3′ end of PAN RNA, is responsible for this accumulation (7). The ENE prevents deadenylation and decay of PAN RNA through direct, cis-acting sequestration of the poly(A) tail (8). The ENE also abrogates rapid nuclear decay when inserted into polyadenylated mRNAs lacking introns (8), apparently through protection from deadenylation, the first step in degradation of eukaryotic mRNAs (9, 10). Secondary structure prediction and analyses of mutants using cellular assays suggested that the ENE forms a hairpin containing a U-rich internal loop flanked by short helices, reminiscent of box H/ACA small nucleolar RNA (snoRNA) hairpins (Fig. 1A) (8, 11). The internal loops of box H/ACA snoRNAs base pair with target regions in ribosomal RNA (rRNA) to direct the conversion of specific uridine residues to pseudouridine (12). By analogy, the ENE internal loop was predicted to base pair with the poly(A) tail of PAN RNA (Fig. 1A), protecting it from exonucleolytic digestion (11).
To elucidate the mode of interaction between the ENE and the poly(A) tail of PAN RNA, we have determined the crystal structure of a complex formed between the 40-nt ENE core and oligo(A) RNA and studied it biochemically (13). Electrophoretic mobility shift assays showed that the isolated ENE is capable of binding oligo(A) in trans (fig. S1). U to C mutations in the internal loop greatly decrease the interaction between the ENE and oligo(A), whereas truncations of the ENE that retain the helix-loop-helix core (Fig. 1A) have no effect (fig. S1). These results are consistent with those obtained from in vivo and in vitro assays of ENE mutants (11). Nuclear magnetic resonance (NMR) studies confirmed that the ENE core alone adopts a helix-loop-helix conformation (fig. S2A). Increased spectral complexity in the imino region of one-dimensional 1H spectra upon A10 addition suggested that the loop becomes involved in hydrogen-bonding interactions with the oligo(A) (fig. S2B). Crystallization trials were then performed using several ENE core sequences that varied in the helix length mixed with a variety of short A-rich RNA oligonucleotides. Optimal crystallization occurred with the 40-nt ENE core construct and A9 RNA. Data from crystals soaked in buffer solutions containing iridium hexamine were used to determine the phases, and the final structure was refined to 2.5 Å resolution (<I/σI> = 1.0; <I/σI> = 2.0 at 2.65 Å resolution) with working and free R factors (Rwork and R/Rfree) of 21.8%/23.3%, respectively (figs. S3 to S5 and table S1).
The crystal structure reveals that the ENE core forms a triple-stranded complex with its bound A9 oligonucleotide (Fig. 1). The ENE assumes the expected secondary structure, with Watson-Crick stems flanking a U-rich internal loop (10). Instead of forming base pairs around this loop, however, the A9 RNA adopts an extended conformation and interacts with both the ENE loop and the lower stem. Nucleotides A5 to A9 of the A9 oligonucleotide simultaneously engage both sides of the internal loop in an extended U-A•U major-groove triple helix (Fig. 2A). Here, the five consecutive A nucleotides form Watson-Crick base pairs with the five consecutive U nucleotides on the 3′ side of the loop [U27 to U31 (14)]. The 5′ side of the ENE internal loop lies in the major groove of the resulting helix, with the Watson-Crick face of nucleotides U8 to U12 base pairing with the Hoogsteen face of the A nucleotides (Fig. 2B). Hydrogen bonds are also observed between the A9 phosphate backbone and the ribose hydroxyl groups of the Hoogsteen U strand. The base triples formed are nearly planar, and there is no direct contact between the two U strands of the internal loop (Fig. 2, A and B). The helical axis of the complex is nearly straight through the transitions between the triple helix and the flanking ENE stems. The additional two nucleotides of the 3′side of the ENE internal loop (A32 and U33) bulge out, allowing continuous stacking between the lower stem and the A and Hoogsteen strands of the triple helix. Despite the presence of the Hoogsteen strand in the A:U major groove, a deep groove remains (fig S6).
The binding interface is augmented by a triad of A-minor interactions formed between the bound A9 oligonucleotide and the lower stem of the ENE. Nucleotides A2 to A4 contact the three consecutive G-C base pairs that close the lower stem (Fig. 2C). As previously described for 23S rRNA in the high-resolution structure of the 50S ribosome (15), the three As of the triad penetrate with increasing depth into the G:C minor groove from the 5′ to 3′ direction, such that the initial interaction is a type III, the next is a type II, and the last is a type IA-minor interaction (Fig. 2D) (15, 16).
The interactions made between the ENE core and A9 RNA are strikingly different from those determined for the box H/ACA snoRNAs and their target rRNA sequences (Fig. 3, A and B). In the snoRNA structures, the substrate strand is sharply kinked, forming parallel Watson-Crick helices with the two strands of the snoRNA internal loop, which then stack coaxially on the flanking snoRNA hairpin helices (17, 18). The crystal structure described here involves an intermolecular interaction between the core ENE and A9 RNA. However, as the ENE has only been observed to protect the PAN RNA poly(A) tail in an intramolecular manner (8), these elements formally adopt a pseudoknot structure in the context of the full-length RNA (see supporting online material). Consistent with this, the ENE A9 complex resembles H-type pseudoknots observed in telomerase RNA (Fig. 3, C and D) (19) and the S-adenosylmethionine-responsive riboswitch SAM-II (fig. S7) (20). Although the topology of the surrounding stems is different in the H-type pseudoknots, the structures of the major-groove triple helices can be superimposed with that of the ENE complex with root mean square deviations (RMSDs) of ~1 Å. Felsenfeld, Davies, and Rich first studied RNA triple helices composed of poly(U)-poly(A)•poly(U) more than half a century ago (21, 22). Denaturation studies later suggested that DNA hairpins with T-rich loops likewise bind oligo(dA) through formation of T-A•T triple helices (23). Analogous interactions between RNA hairpins and A-rich sequences were also proposed (23). However, the ENE is the first natural example of the use of a U-rich internal loop to capture and sequester a poly(A) RNA sequence.
We used deadenylation assays to confirm the functional importance of the ENE:A9 triple helix described here (13). Mutation of a single U to C in either side of the ENE U-rich pocket had previously been shown to decrease protection of PAN RNA's poly(A) tail from deadenylation in nuclear extract (11). Simultaneous U to C mutations (mutating one nucleotide from each side of the pocket) were as deleterious as deletion of the entire ENE. Because the two U nucleotides mutated in that experiment contact the same A in the crystal structure (Fig. 4A), we postulated that the loss of ENE function arising from disruption of a single U-A•U base triple might be restored by replacement with a nearly isosteric C-G•C base triple (Fig. 4B) (24). Native gel shift analysis supported this proposal, as ENEs containing simultaneous U to C mutations contacting the same A in the crystal structure can bind an oligo(A) molecule containing a single A to G substitution (A7GA2) (fig. S8, A and C), whereas ENE constructs with mutations to C in U residues that contact two different A nucleotides in the crystal structure cannot (fig. S8, B and C).
We thus transcribed PAN RNA deadenylation substrates that contained either wild-type or double-mutant ENE or that lacked the ENE altogether (see legend to Fig. 4C). Each of the substrates terminated in a 60-nt poly(A) tail with or without a single A to G substitution (A60 or A57GA2). We assessed these constructs for ENE-dependent protection from deadenylation in nuclear extract (Fig. 4C, left and middle panels). As expected, the wild-type ENE protected the A60 tail from deadenylation, whereas the A60 tails of constructs containing no ENE (Δ ENE) or the double-mutant ENE (U903C, U949C) were not protected from deadenylation (Fig. 4C, left panels) (10). In contrast, the double-mutant ENE effectively protected the A57GA2 poly(A) tail, which has a single A to G substitution close to its 3′ end (Fig. 4C, center panel). The presence of a G residue in the poly(A) tail does not on its own confer resistance to deadenylation, as the A57GA2 poly(A) tail of a construct lacking the ENE was not protected (Fig. 4C, bottom middle panel). These results, supported by the native gel shift data described above (fig. S8), indicate that the triple helix observed in the structure is critical to ENE function.
Finally, we tested the ability of the double-mutant ENE to protect a poly(A) tail containing a more internal G substitution (A19GA40) (Fig. 4C, right panel). In nuclear extract, this poly(A) tail was rapidly deadenylated to a size consistent with formation of a triple helix that includes the predicted C-G•C base triple (Fig. 4C, right middle panel). The ability of the double-mutant ENE to locate a single G within a long stretch of A nucleotides is striking. The deadenylation data also demonstrate that the ENE does not require the 3′ terminus of the poly(A) tail for binding, despite the involvement of the 3′ end of the A9 RNA in the final base triple of the ENE core:A9 structure. The results further argue that no specific register is required for the interaction of the wild-type ENE with the PAN RNA poly(A) tail. The presence of multiple binding sites for the ENE along the poly(A) sequence may contribute to its ability to protect tails of various lengths from deadenylation by cellular exonucleases (Fig. 4A). How the ENE may collaborate with poly(A)–-binding proteins that are known to coat the poly(A) tails of RNA polymerase II transcripts in vivo (25) remains to be determined.
The key feature of the core ENEL:A9 crystal structure is a functionally important U-A•U major-groove triple helix, which is extended by A-minor interactions. The structure reveals an intramolecular clamp mechanism for recognition of poly(A) RNA and suggests how the ENE sequesters the PAN poly(A) tail from degradation by cellular deadenylases. Since viruses routinely borrow strategies from their hosts, we predict that similar mechanisms may protect some cellular noncoding RNAs from rapid turnover.
We thank S. Borah, K. Tycowski, K. Herbert, and K. Riley for critical reading of the manuscript and the entire Steitz laboratories for helpful discussion.We thank S. Strobel and G. Conn for the generous gifts of iridium(III) hexamine and 3′-hepatitis delta virus (HDV) plasmid, respectively. Special thanks to Y. Zuo and G. Blaha for crystallography assistance, P. Moore and E. Paulson for NMR assistance, and M.-D. Shu, D. Mishler, and K. Durniak for technical assistance. X-ray data were collected at the National Synchrotron Light Source (X29A) at Brookhaven National Laboratory (13). Financial support for this research was provided by NIH grant CA16038 to J.A.S. and NIH grant GM022778 to T.A.S. The content is solely the responsibility of the authors and does not necessarily represent the official views of NIH. R.M.M.-F. is supported by a Jane Coffin Childs Memorial Fund Postdoctoral fellowship. J.A.S. and T.A.S. are investigators of the Howard Hughes Medical Institute. Coordinates and structure factors have been deposited in the Protein Data Bank under accession code 3P22.
Supporting Online Material
Materials and Methods
Figs. S1 to S8