Translation of the hepatitis C virus (HCV) polyprotein is ini-tiated at an internal ribosome entry site (IRES) element inthe 5′ untranslated region of HCV RNA. The HCV IRES ele-ment interacts directly with the 40S subunit, and biochemicalexperiments have implicated RNA elements near the AUGstart codon as required for IRES–40S subunit complex for-mation. The data we present here show that two RNA stemloops, domains IIId and IIIe, are involved in IRES–40S sub-unit interaction. The structures of the two RNA domainswere solved by NMR spectroscopy and reveal structural fea-tures that may explain their role in IRES function.
Initiation of translation of the hepatitis C virus genome ismediated by an internal ribosome entry site (IRES) element inthe 5′ untranslated region (5′ UTR) of HCV genomic RNA1,2.Normal translation initiation in eukaryotes occurs by recogni-tion of the 5′ cap structure of the mRNA by eukaryotic initia-tion factor 4E (eIF4E) followed by assembly of other initiationfactors and the 40S ribosomal subunit, and subsequent scan-ning of the 5′ UTR to the first AUG initiation codon3. In HCV,IRES-mediated initiation eliminates the requirement for the 5′cap structure and scanning. The 40S subunit is recruiteddirectly to the vicinity of the start codon by interaction withthe IRES element; only a subset of the total translation initia-tion factors is required for this process2. As such, IRES-mediat-ed initiation in HCV and related pestiviruses is reminiscent ofprokaryotic translational initiation, in which the Shine-Dalgarno interaction between messenger RNA and 16S riboso-mal RNA recruits 30S subunits directly to the start codon, andonly three initiation factors are required4. How the IRESdirectly interacts with the 40S ribosomal subunit remainsunclear.
The HCV IRES element is a complex RNA secondary struc-ture consisting of nucleotides 44–354 in the 5′ UTR5–8.Biochemical experiments have demonstrated the roles of par-ticular subdomains to IRES function. In particular, mutagene-sis and functional data support a pseudoknot structure nearthe AUG start codon9,10. The sites of interaction with the 40Ssubunit have been mapped by toe-printing, which yieldedstrong, 40S-dependent primer extension stops in the IRES sub-domains including and adjacent to the pseudoknot structure4.Mutations within two hairpin loops, domain IIId and IIIe, dis-rupt IRES-mediated initiation11,12. Biochemical and smallangle X-ray scattering experiments suggested that the IRES hasa metal-dependent tertiary structure, which may be requiredfor interaction with 40S subunits12.
nature structural biology • volume 7 number 12 • december 2000 1105
We present here biochemical and functional data that sup-port the direct role of domain IIId and IIIe hairpins in IRESinteraction with the 40S subunit and we have determined thestructures of these two RNA domains by NMR spectroscopy.Our results further support the role of IRES RNA structure inrecognition of the 40S ribosomal subunit.
HCV IRES chemical footprintingThe HCV IRES binds directly to the 40S ribosomal subunit inthe absence of external factors4. To map the regions of the IRESthat may interact with 40S subunits, we performed chemicalprobing on the IRES RNA (domain IV and the 3′ half ofdomain III) in the absence or presence of ribosomal subunits.The chemical reactivity of the unbound IRES is consistentwith the well-characterized secondary structure of the IRES(Fig. 1a). The reactivity of the bases depends on the Mg2+ con-centration, consistent with the previously defined Mg2+-dependent formation of tertiary structure for the IRES12.Upon binding of the folded IRES to 40S subunits, two regionsof the IRES show strong changes in reactivity (Fig. 1b). TheWatson-Crick faces of guanosines in the hairpin loops ofdomains IIId and IIIe are strongly reactive to kethoxal in theunbound IRES, and are almost completely protected upon 40Sbinding. In contrast, three adenosines in the stem of domainIIId show increases in reactivity upon 40S subunit binding.These loops are required for IRES function in vivo andin vitro11,12 and our chemical probing experiments suggest thatthey are involved in IRES–40S subunit interaction.
NMR spectroscopyTo understand further the role of domains IIId and IIIe in IRESfunction, we determined both structures using NMR spec-troscopy (Fig. 1c). For domain IIIe, a 14-residue RNA oligonu-cleotide, corresponding to nucleotides 291–302 was studiedusing homonuclear NMR methods. The spectra were readilyassigned without isotopic labeling, and a total of 271 NOE and88 dihedral torsion angle restraints were obtained. For domainIIId, a 29-residue RNA oligonucleotide, corresponding tonucleotides 253–279, was studied by NMR spectroscopy. Acombination of homonuclear and heteronuclear 2D and 3DNMR methods yielded a total number of 705 NOE and 200dihedral torsion angle restraints. Measurement of intra basepair 2JNN couplings across hydrogen bonds was employed toestablish base pairing schemes13. For both hairpin loops, onlyNMR-derived restraints were used for structure calculations.
Structure of HCV IRES domain IIIeThe domain IIIe hairpin loop adopts a novel tetraloop foldand is well defined by the NMR data (heavy atom root meansquare (r.m.s.) deviation = 0.89 Å; Fig. 2a). The helical stem ofdomain IIIe is terminated by a U294-G299 wobble pair, fol-lowed by a loop closing, sheared G295-A298 base pair (Fig.2b). Similar pairing interactions are observed in the GNRAtetraloops and in purine-rich hexaloops14,15. The loopsequence (–GAUA–), however, does not conform to the stan-dard GNRA motif and adopts a different structure. The basesof A296 and U297 point towards the major groove and are notinvolved in RNA backbone contacts that would stabilize theloop fold; the two central nucleotides of the tetraloop stack onthe 5′ guanosine of the sheared G-A pair. This fold creates anarray of three major groove exposed Watson-Crick faces(G295, A296, and U297). In contrast, the central purine andadenosine in a GNRA tetraloop point towards the minor
Structures of two RNAdomains essential forhepatitis C virus internalribosome entry sitefunctionPeter J. Lukavsky1, Geoff A. Ot to1, Alissa M. Lancaster2,Peter Sarnow 2 and Joseph D. Puglisi1
1Department of Structural Biology and 2Department of Microbiology andImmunology, Stanford University School of Medicine, Stanford, California94305-5126, USA.
groove, and are stacked on the 3’adenosine of the sheared G-Apair (Fig. 2b).
Functional importance of HCV IRES domain IIIeThe sequence of the GAUA tetraloop is conserved among allHCV isolates, and among related pestiviridae IRES16. Our bio-chemical studies suggest that this loop is a point of direct con-tact with the 40S subunit. G295, which is exposed to the majorgroove, is strongly reactive to kethoxal in the free IRES and is
1106 nature structural biology • volume 7 number 12 • december 2000
protected from chemical modification upon40S binding (Fig. 1b). The importance of theIIIe tetraloop for IRES function was also sup-ported by translation in cultured human HeLacells. In vivo IRES activity of wild type andmutant HCV IRES were measured using a dualluciferase reporter assay. The second cistroncontains the firefly luciferase gene, which istranslated under IRES control. The size andintegrity of the dicistronic mRNAs were moni-tored by Northern analysis to examine the pos-sibility that firefly luciferase protein wassynthesized from functionally monocistronicfirefly luciferase-containing mRNAs, whichcould have been generated by splicing or bycryptic promoter elements located in the dis-cistronic genes. These Northern analysesrevealed that all dicistronic mRNAs remainedlargely intact (data not shown), arguing thatthe second cistrons were translated by internalinitiation mediated by the HCV IRES.
Using the dual luciferase reporter assay, westudied the effects of mutations of the GAUAtetraloop motif on IRES activity. Mutation ofthe major groove exposed base U297 to a cyto-sine residue causes a more than 50% decreasein IRES-mediated translation compared to wildtype (Table 1). Converting the GAUA tetraloopinto the GNRA tetraloop sequence by mutatingU297 to an adenine residue, causes the samedecrease in IRES activity. These data are consis-tent with a previous mutational study ofdomain IIIe, which showed that virtually anyalteration of the loop sequence caused a signifi-cant decrease in IRES activity11. The presenta-
tion of bases by the domain IIIe tetraloop may be required forthe IRES-40S subunit interaction.
Structure of HCV IRES domain IIIdThe domain IIId RNA forms a helical stem with noncanonicalpairings, followed by a hexanucleotide loop region. The overallstructure is well defined by the NMR data (heavy atom r.m.s.deviation = 1.61 Å; Fig. 3a). The –UUGGGU– hairpin loop ismore disordered than the other regions of the RNA; the heavy
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GGCGUCUGGUGAU
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UGGGCAG
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44
70 97
118 133
144 249
155
171
191
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206
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296
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360
II
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IIIb
IIIc
IIId
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}
}
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b
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296 A U 297
295 G A 298
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260 A259 U
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U G264 U U 269
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257 A 256 G
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c
Fig. 1 a, Sequence and secondary st ructure of HCVIRES RNA (nucleot ides 1–383 of HCV genotype 1b).Domains are numbered according to ref . 34Nucleot ides protected f rom kethoxal modif icat ionupon 40S subunit binding are indicated by f illed circles.Nucleot ides that show increases in DMS modif icat ionupon 40S subunit binding are indicated by open circles.b, Autoradiograph of kethoxal and DMS probing ofHCV IRES RNA domains IIId and IIIe in the absence (–),or presence (+) of 40S subunits. The K lane is a primerextension react ion using the unmodif ied HCV IRESRNA. The kethoxal (ket ) and DMS probing and primerextension react ions are performed as described inMethods. The lanes demarked U, G, C and A aredideoxy sequencing react ions. c, Sequence and sec-ondary st ructure of the HCV IRES domain IIIe and IIIdRNA oligonucleot ides used for the NMR st ructuralstudies. Numbering according to Fig. 1a. Nucleot ides,that were changed to improve t ranscript ion eff iciency,are out lined.
atom r.m.s. deviation for U264–U269 is 1.46 Å(Fig. 3c). The central internal loop, which is highlyconserved among HCV isolates, adopts the well-char-acterized loop E fold, and is very well defined by theNMR data (r.m.s. deviation of 0.28 Å; Fig. 3d). Fourconsecutive noncanonical base pairs are formed — asheared G256-A276 pair, a parallel A257-A275 pair, areverse Hoogsteen U259-A274 pair and anothersheared A260-G273 base pair (Fig. 3e). The reverseHoogsteen hydrogen bonding scheme is supported bythe observation of internucleotide 2JNN scalar cou-plings. The arrangement of the base pairs within theloop E motif creates a continous stack of fouradenines (A260 and A274–A276) with their Watson-Crick faces exposed to the minor groove (Fig. 3d).The internal loop is asymmetric, with the bulgedG258 positioned in the major groove, where it forms abase triple with the U259-A274 reverse Hoogsteenpair (Fig. 3e). The phosphodiester backbone undergoes a localreversion of direction at A257 and G258, such that a parallelhydrogen bonding arrangement between A257 and A275 canform. The inversion in backbone direction leads to an S turn inthe backbone between G256 to A259 (Fig. 3b), which is charac-teristic for loop E motifs. The unusual backbone geometrywithin the loop E motif results from non A-form values for tor-sion angles β for G258 and A274 (gauche+), γ for A274 (trans),and ε for A257 (gauche+) to allow for the triple formation andbackbone inversion. The structure also explains unusual 1H,13C, and 31P chemical shifts observed for several loop E reso-nances (chemical shifts are available on our web page:http://puglisi.stanford.edu/).
Comparison to other loop E motifsThe loop E motif is common in RNAs, with different sequencefamilies17. All loop E motifs contain a sheared G-A pair and theadjacent U-A pair. In prokaryotic 5S ribosomal RNA, a loop Emotif is observed with a symmetric internal loop: a G-G pairand a sheared G-A pair18. In eukaryotic 28S ribosomal RNA,the sarcin-ricin loop (SRL) contains a loop E motif that con-tains the A-A pair, bulged G, U-A pair and G-A pair19,20; ther.m.s. deviation between the crystal structure of SRL and theHCV IRES domain IIId loop E motif is 1.15 Å. The SRL has aflexible region adjacent to the loop E motif, whereas in domainIIId the loop E is bordered by a sheared G-A pair and Watson-Crick base pairs. Loop E motifs present rich hydrogen bondingpotential in both the minor and major groove for bothRNA–RNA and RNA–protein interactions.
Functional importance of domain IIIdThe sequence of the –UUGGGU– hairpin loop of domain IIIdis absolutely conserved among all HCV isolates16. The hairpinloop is separated from the loop E motif by a short helical stemconsisting of a G-U wobble pair flanked by two G-C Watson-
Crick base pairs, of which one closes the hairpin loop. On the5′ side of the loop, U264 stacks on top of the loop closing G-Cbase pair, whereas U269 on the 3′ side is bulged into solutionand disordered in the ensemble of NMR structures (Fig. 3c).This positions the ribose of G268 above the ribose of C270 ofthe loop-closing G-C base pair, which exposes the base of G268to the major groove and introduces an inversion in backbonedirection similar to the loop E motif with an S turn betweenG267 to C270 (Fig. 3b). G267 stacks below G266 in the minorgroove of the loop and U265 is located in the major groove,where it loosely stacks on the 5′ side residues and is more dis-ordered compared to the three guanosine residues. The six basepair spacing between the loop E and the hairpin loop backbonereversion places both S turns on the same side of the hairpinloop structure. This creates a unique backbone feature for thedomain IIId motif.
The domain IIId hairpin loop clearly plays an important rolein IRES–40S subunit interaction. In the chemical probing
Fig. 2 Structure of HCV domain IIIe stem loop. a, Stereo viewf rom the major groove of the heavy atom superposit ion of f inal20 st ructures of HCV IRES domain IIIe. Bases are colored in blueand ribose-phosphate backbone in gray. b, Single representat ivest ructures of the GNRA14 and the GAUA tet raloop of HCV IRESdomain IIIe. Base nit rogens are in blue and base oxygens in red.Phosphorus atoms and phosphate oxygens are shown explicit lyin yellow and red, respect ively.
1All dicist ronic vectors contain an addit ional mutat ion of the HCV AUGstart codon to CUG, which did not af fect t ranslat ional act ivity6.2Translat ional ef f iciency of mutant IRES RNAs is displayed as LucF/LucRrat io in reference to wild type IRES RNA, whose LucF/LucR rat io was set to100% act ivity. Mean values f rom four independent t ransfect ion experi-ments performed as duplicates are shown with standard errors. The sizeand integrity of the dicist ronic mRNAs were monitored by Northernanalysis (data not shown). All mRNAs displayed similar int racellular sta-bilit ies, conf irming that the second cist rons (LucF) were t ranslated byinternal init iat ion mediated by the IRES element .
nature structural biology • volume 7 number 12 • december 2000 1107
experiments discussed above, the Watson-Crick faces of G266,G267 and G268 were strongly protected from reaction withkethoxal in the IRES–40S subunit complex (Fig. 1b). In addi-tion, the N7 positions of G266 and G267 are protected frommethylation by dimethyl sulfate (DMS) in the IRES–40S com-plex, whereas the N7 of G268, which is exposed on the majorgroove side of the IIId loop, is highly reactive in the complex(data not shown). The three guanosines in the loop arerequired for full IRES activity in internal initiation. Mutationof the three loop guanosines to cytosines had been previouslyshown to be deleterious to IRES activity in vitro12. Based on ourstructural data, we mutated all three guanosine residues (G266-G268) to adenines preserving purine residues in those posi-tions in order to maintain the fold of domain IIId, which wastested by chemical probing (data not shown). These mutations,which did not alter the local IRES fold like the G266-268C
1108 nature structural biology • volume 7 number 12 • december 2000
mutations, decreased IRES–mediated translation by 50%(Table 1).
What is the role of the loop E motif in domain IIId?The ability to form a loop E fold is conserved among HCV iso-lates. An observed change is G256-A276 to a U-C pair in HCVisolate HCV-2b, which would lead to the SRL loop E motif.Our preliminary data indicate that mutations that disrupt boththe loop E motif and folding of domain IIId are deleterious toIRES function (data not shown). The loop E motif, with itsnarrowed major groove, distorted phosphodiester backbone,and stretch of noncanonical pairings, may be involved in RNAtertiary interactions within the IRES, or intermolecular inter-actions with the 40S subunit; the interactions with the ribo-some may be with protein or RNA components. The adenosineN1 positions of A274–A276 in the minor groove of the loop E
Fig. 3 Structure of HCV domain IIId stem loop. a, Stereo view of theheavy-atom superposit ion of f inal 25 st ructures of HCV IRES domainIIId. The color scheme is the same as in Fig. 2a. b, Single representa-t ive st ructure of the HCV IRES domain IIId, with both S turns high-lighted. Ribose O4′ atoms are shown in red, the inverted riboses inblue and the phosphate backbone in yellow. c, Major groove view of the heavy-atom superposit ion of hairpin loop nucleot idesG263–C270 of the f inal 25 st ructures of the HCV IRES domain IIId anda single representat ive st ructure. The color scheme is the same as inFigs 2a,b. Phosphorus atoms are shown in yellow. d, Minor grooveview of the heavy atom superposit ion of loop E nucleot idesC255–G261 and C272–G277 of the f inal 25 st ructures of the HCV IRESdomain IIId and a single representat ive st ructure omit t ing the f lank-ing G-C base pairs. The color scheme is the same as in Fig. 3c. e, Basepairing schemes found within the loop E mot if of HCV IRES domainIIId. The color scheme is the same as in Fig. 2a. Hydrogen bondsshown by dashed lines are observed in all 25 f inal NMR st ructures.
clear and heteronuclear methods opt imized forRNA st ructure determinat ion (RnaPack, Varian UserLibrary). In short , constant t ime HSQC, 3D HCCH-TOCSY, 3D HCCH-COSY, and 2D HCCH-RELAY experi-ments were used to assign sugar spin systems, whilethrough-backbone assignments were made w ithHCP and HP-COSY experiments23. Base exchange-able protons were assigned by correlat ion to non-exchangeable base protons using heteroTOCSYmethods. Int ranucleot ide H1′ t o base proton corre-lat ions were obtained using a 2D MQ–HCN experi-ment 24. NOE distance rest raint s f romnon–exchangeable protons were obtained f rom2D–NOESY experiments (100% D2O) w ith mixingt imes of 50, 150 and 250 ms. Exchangeable protonNOEs were determined using SS-NOESY25 or WATER-GATE-NOESY experiments (4% D20) w ith mixingt imes of 50 and 150ms. A 3D 13C-edited NOESY-HSQCexperiment (4% D2O) w ith a mixing t ime of 150mswas used to conf irm both nonexchangeable andexchangeable proton NOE assignments. Base pair-ing schemes for IIId were established using the HNN-COSY experiment 13. NOEs f rom exchangeableprotons were characterized as st rong (0–3.5 Å),medium (0–4.5 Å) or weak (0–6 Å), while NOEs f romnonexchangeable protons were characterized aseither st rong (0–3 Å), medium (0–4 Å), weak (0–5 Å)or very weak (0–6 Å). Dihedral t orsion rest raint swere obtained f rom DQF-COSY, 3D HMQC-TOCSY,HP-COSY and HCP experiments, as described26.Spect ra were analyzed w ith SPARKY27.
Structure calculat ion. St ructures were calculatedusing a simulated annealing protocol w ithin the X-PLOR 3.1 package28. The protocol f or st ructuralcalculat ions included two stages; simulated anneal-ing of start ing st ructures w ith random angles andrest rained molecular dynamics (rMD) ref inement . Atotal of 699 NOE distance rest raints, 6 NN hydrogen
bond distance rest raint s and 200 dihedral rest raint s for IIId and271 NOE distance rest raints and 88 dihedral rest raints for IIIe wereused. The NOE distance force constants were set t o 50 kcal mol-1
Å-2 and torsion angle force constants were varied f rom 5 to 50 kcalmol-1 rad-2 during calculat ions. No hydrogen bonding rest raint sother than experimentally measured ones were used in calcula-t ions.
A total of 100 start ing st ructures were generated and subjectedto a simulated annealing protocol. This consisted of 500 cycles ofenergy minimizat ion, f ollowed by rMD at 1,000 K w ith low valuesfor interatomic repulsion, and subsequent rMD w ith increasingvalues for interatomic repulsion while cooling to 300 K. A f inalminimizat ion step w ith 1,000 cycles was performed, which includ-ed a Lennard-Jones potent ial and no elect rostat ic t erms. The 100st ructures were then subjected to a ref inement procedure: 500steps of rest rained energy minimizat ion; rMD at 1,000 K whileincreasing the torsion angle force constant ; rMD while cooling to300 K and f inally 1,000 cycles of energy minimizat ion, whichincluded a Lennard-Jones potent ial, but no elect rostat ic t erms.The f inal st ructures (25 IIId or 20 IIIe) were chosen, which had thelowest t otal and rest raint violat ion energies, whereas non con-verged st ructures were at least one standard deviat ion higher intotal and rest raint violat ion energy (Table 2).
Chemical probing. 40S subunits were isolated f rom HeLa S3 cellpellets (Nat ional Cell Culture Center) by the puromycin method ofBlobel and Sabat ini29. HCV IRES RNA (nt 40–375) was generated byT7 RNA polymerase run-off t ranscript ion and purif ied by gel elec-t rophoresis12. Chemical modif icat ion w ith kethoxal and DMS wasperformed essent ially as described in Moazed and Noller30.React ions were performed with an excess of 40S subunits in 125 mMKOAc, 10 mM MgCl2, 30 mM HEPES-KOH (pH 7.0) and 0.5 mM spermidine. Sodium borohydride reduct ion and aniline-induced st rand scission of DMS modif ied IRES was also performed31.
motif are highly accessible to modification by DMS in theIRES–40S subunit complex (Fig. 1b). Therefore, protein orRNA interactions with the loop E motif likely occur on themajor groove side. Additional experiments are required todetermine the precise role of the loop E motif of domain IIId inIRES–40S subunit interaction.
The results presented here demonstrate that two surface-accessible stem loops in the HCV IRES are involved in complexformation with 40S ribosomal subunits. The novel structuresof both domain IIId and IIIe are suggestive of their involve-ment in IRES function, and suggest experiments for a molecu-lar level understanding of HCV IRES function. The workpresented here demonstrates the powerful ability of RNA NMRto provide local structural information to drive biochemicalstudies of a large RNA system.
MethodsSample preparat ion. RNA oligonucleot ides were prepared byt ranscript ion f rom DNA templates by phage T7 RNA polymeraseand purif ied using polyacrylamide gel elect rophoresis21.Unlabeled and 13C, 15N-labeled RNAs were prepared. Labelednucleoside t riphosphates were prepared in-house using publishedmethods22. RNAs were elect roeluted f rom the gel and subsequent -ly dialyzed against f inal buf fer (10 mM Na phosphate, pH 6.4,1 mM d12-EDTA, 4% D2O or 100% D2O). NMR samples were pre-pared in a Shigemi NMR tube (sample volume 250 µL) at RNA con-cent rat ions of 1.0–2.5 mM.
NM R spect ral analyses. NMR data were acquired at eit her 15 or25 °C on Varian Inova 500 MHz and 800 MHz NMR spect rometersequipped w ith t riple resonance x,y,z-axis gradient probes. 1H, 13C,15N, and 31P assignments were obtained using standard homonu-
Table 2 Structural statistics for domain IIIe and IIId RNA oligonucleotidesdomain IIIe <SA>1 domain IIId <SA>2
Total number of experimental rest raintsDistance rest raints 271 705Dihedral rest raints 88 200
Final distance and dihedral rest raint Violat ion energies (kcal mol-1) 15.9 ± 0.8 29.9 ± 0.7
R.m.s. deviat ion f rom experimental rest raintsDistance rest raints (Å)3 0.03 ± 0.001 0.03 ± 0.001Dihedral rest raints (°)3 0.95 ± 0. 0.043 0.98 ± 0.06
<SA> versus SA <SA> versus SAHeavy-atoms r.m.s. deviat ion (Å)
All IIIe RNA 0.89All IIId RNA 1.61
Heavy-atoms r.m.s. deviat ion (Å)IIIe loop (U294–G299) 0.19IIId loop E mot if (G256–A260, G273–A276) 0.28IIId hairpin loop (U264–U269) 1.46
1<SA> refers to the f inal 20 simulated annealing st ructures, SA to the average st ructureobtained by taking the average coordinates of the 20 simulated annealing st ructuresbest -f it ted to one another.2<SA> refers to the f inal 25 simulated annealing st ructures, SA to the average st ructureobtained by taking the average coordinates of the 25 simulated annealing st ructuresbest -f it ted to one another.3The f inal st ructures did not contain distance violat ions of >0.25 Å or dihedral violat ionsof >5°. Numbers in parentheses refer to number of rest raints.
nature structural biology • volume 7 number 12 • december 2000 1109
Primer extension, using a primer complementary to the HCV IRESopen reading f rame was used to detect sites of RNA base modif ica-t ion32. Body labeled products were separated on an 8% 7 M Urea,1X TBE acrylamide gel and detected by autoradiography.
Const ruct ion of dual luciferase reporter const ructs andt ranslat ion assays. Wild t ype1 and mutated HCV IRES elementswere subcloned into the intercist ronic region of a dicist ronicluciferase reporter const ruct , as described previously33.
DNA plasmids were t ransfected into HeLa cells using theFuGENE 6 t ransfect ion reagent (Boehringer Mannheim).Transfected Cells were harvested 24 h af ter t ransfect ion andluciferase act ivit ies were measured using the Dual LuciferaseReporter Assay System (Promega Biotech).
Northern analysis. Total RNA was harvested f rom t ransfectedHeLa cells 24 h af ter t ransfect ion using the Trizol reagent(Gibco/BRL). Polyadenosine-containing (polyA+) RNA was isolatedf rom the total RNA using the Oligotex mRNA Kit (Qiagen).Approximately 1 µg of polyA+ RNA was separated on a formalde-hyde-containing gel and t ransferred to a nit rocellulose mem-brane. Radiolabled probe was generated f rom a PCR productcorresponding to nucleot ides 648–1,280 of t he f iref ly lucif erasegene using the RadPrime Kit (Gibco/BRL) and hybridized to themembrane using Express-hyb solut ion (Clonetech).
Coordinates. The cooridinates have been deposited in theProtein Data Bank (accession code 1F84 for HCV domain IIId and1F85 for HCV domain IIIe).
AcknowledgmentsThe authors thank E. Lau for preparation of labeled nucleotides. Supported bygrants from the NIH, the Hutchinson Foundation and Eli Lilly, Inc., and apostdoctoral grant to P. J. L. from the Max Kade Foundation. The StanfordMagnetic Resonance Laboratory is supported by the Stanford University Schoolof Medicine.
1110 nature structural biology • volume 7 number 12 • december 2000
Correspondence should be addressed to J.D.P. email: [email protected]
Received 10 May, 2000; accepted 3 October, 2000.
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