A highly conserved G-rich consensus sequence in hepatitis ...small molecules specifically targeting G4 can stabilize this structure to reduc e RNA replication and inhibit protein translation

R E S EARCH ART I C L E

CHEM ICAL B IOLOGY

1College of Chemistry and Molecular Sciences, Key Laboratory of Biomedical Polymersof Ministry of Education, Wuhan University, Wuhan 430072, Hubei, China. 2State KeyLaboratory of Virology and Hubei Province Key Laboratory of Allergy and Immunology andDepartment of Immunology, School of Medicine,WuhanUniversity, Wuhan 430071, Hubei,China. 3Institute of Hydrobiology, Chinese Academy of Sciences, Wuhan 430072, Hubei,China. 4Wuhan Institute of Physics andMathematics, Chinese Academy of Sciences, Wuhan430071, Hubei, China. 5College of Life Sciences, Wuhan University, Wuhan 430072, Hubei,China.*These authors contributed equally to this work.†Corresponding author. E-mail: [email protected] (X.Z.); [email protected](X.-L.Z.); [email protected] (T.T.)

Wang et al. Sci. Adv. 2016; 2 : e1501535 1 April 2016

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10.1126/sciadv.1501535

A highly conserved G-rich consensus sequencein hepatitis C virus core gene represents a newanti–hepatitis C target
Shao-Ru Wang,1* Yuan-Qin Min,2* Jia-Qi Wang,1 Chao-Xing Liu,1 Bo-Shi Fu,1 Fan Wu,1 Ling-Yu Wu,1 Zhi-Xian Qiao,3
Yan-Yan Song,1 Guo-Hua Xu,4 Zhi-Guo Wu,5 Gai Huang,5 Nan-Fang Peng,5 Rong Huang,1 Wu-Xiang Mao,1

Shuang Peng,1 Yu-Qi Chen,1 Ying Zhu,5 Tian Tian,1† Xiao-Lian Zhang,2† Xiang Zhou1†

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G-quadruplex (G4) is one of the most important secondary structures in nucleic acids. Until recently, G4 RNAs havenot been reported in any ribovirus, such as the hepatitis C virus. Our bioinformatics analysis reveals highly con-served guanine-rich consensus sequences within the core gene of hepatitis C despite the high genetic variability ofthis ribovirus; we further show using various methods that such consensus sequences can fold into unimolecular G4RNA structures, both in vitro and under physiological conditions. Furthermore, we provide direct evidences thatsmall molecules specifically targeting G4 can stabilize this structure to reduce RNA replication and inhibit proteintranslation of intracellular hepatitis C. Ultimately, the stabilization of G4 RNA in the genome of hepatitis C repre-sents a promising new strategy for anti–hepatitis C drug development.

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INTRODUCTION

Viruses rapidly mutate, and RNA viruses (particularly riboviruses,rather than retroviruses) have higher genetic variation than DNAviruses (1). Hepatitis C virus (HCV) infection is estimated to affect2.8% of the population worldwide (2). Recent medical studies havefocused on the development of small molecules targeting viral en-zymes (3, 4). However, resistance rapidly emerges in HCV patientswho are treated with specific inhibitors targeting nonstructural proteins,reflecting the error-prone replication by NS5B, a viral RNA-dependentRNA polymerase (RdRp) (5). Hence, the development of compoundstargeting highly conserved regions or structural motifs in the HCV ge-nome would have the most potential to protect against not only thecurrent virus but also the varied ones (6, 7). Moreover, the use ofsmall-molecule RNA binders as therapeutics is an area of intense in-terest at the interface of chemistry and biology (8, 9).

G-quadruplex (G4) contains stacked planar G-quartets, which arestabilized through Hoogsteen hydrogen bonding (fig. S1) (10–12). G4RNAs have been identified in telomeric transcripts (13) and 5′ or 3′ un-translated regions (UTRs) ofmRNAs (14–16). Recently, G4 RNAs havebeen associatedwith some viruses (17) such asHIV (18), herpes simplexvirus (HSV) (19), and Epstein-Barr virus (EBV) (20). BRACO-19, aG4-binding ligand, has been demonstrated to be an active anti-HIVagent (21). However, for HSV and EBV, G4 RNAs only form in nascentRNA transcripts during transcription, and HIV only exists in the latentform of proviral DNA upon entry into human cells. To the best of ourknowledge, the roles of G4 RNA in these studies are confined to theregulation of transient gene expression at the translational level. Until

recently, there have been no reports concerning the presence of G4structure in riboviruses. Because RNA replication and translation occuron the same RNA of positive-sense single-stranded RNA (ssRNA)viruses, we suggest that HCV G4 RNA (if present) play a more directrole in regulating both of these processes.

Here, we provide the first evidence that a highly conserved guanine-rich (G-rich) sequence is present in theHCV core (C) gene. Using var-iousmethods, we demonstrate the high potential of HCVC consensussequences to formG4 RNAs. Furthermore, our biological findings pro-vide direct evidences that, because of the actions of G4 ligands in bindingto and stabilizingG4RNA in the specificG-rich regionof theCgene, theycan reduce RNA replication and inhibit protein translation in HCV. Ul-timately, the highly conserved nature ofHCVG-rich RNAmight repre-sent a challenging new target for anti-HCV drug development.

RESULTS

Bioinformatics analysis to reveal HCV G-rich sequencesOn the basis of a genomic variance analysis, HCV was classified intomultiple genotypes (1 to 7) with varied subtypes (22). We retrieved77 complete genomic sequences of all available genotypes (GenBankaccession numbers in table S1) and conducted bioinformatics anal-ysis to assess the level of sequence conservation (23). As shown in fig. S2,a low ratio (30.89%) of conserved nucleotide sites across the wholegenome was observed, whereas much higher conservation (46.77%)was identified within the nucleotide sites of the C gene. Thus, we areparticularly interested in the development of a new antiviral strategytargeting the HCV C gene.

We next performed multiple sequence alignments of the C geneacross these 77 HCV genomes (figs. S3 to S5). In this example, a G-richconsensus sequence harboring four G-tracts was observed in the centralregion of the C gene, between positions +259 and +285. A high proportion(47.37%) of the nucleotide sites is conserved across such aG-rich region,which is highly characteristic of sequences with high potential to formG4 structures. HCV subtypes 1a and 1b are commonly observed in

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East Asia andNorth America (24), prompting significant interest in thestudy of these specific subtypes within the field. Hence, we aligned 1056partial coding sequences (cds) of theCgene forHCVsubtype 1a and 1025cds for HCV subtype 1b (tables S2 and S3), retrieved from the HCVdatabase. WebLogo was used to generate a graphical representation ofthealigned sequences (25) betweenpositions+267and+285.As illustratedinFig. 1,RNA1a(GGGCUGCGGGUGGGCGGGA,735/1056) andRNA1b(GGGCAUGGGGUGGGCAGGA, 656/1025) were among the sequencesmost frequently observed to display great potential for G4 formation;therefore, these sequenceswere selected as targets for the following studies.

G4 RNAs evidenced through gel electrophoresis and 1Hnuclear magnetic resonanceNative polyacrylamide gel electrophoresis (PAGE) was performedusing fluorescently labeled RNAs (RNA1a-FAM and RNA1b-FAMin Table 1) to monitor structure compaction, as G4 RNA should mi-grate faster than ssRNA (26). As shown in Fig. 2A, the bandcorresponding to target RNA migrated significantly faster than itsG4-mutated counterpart, indicating the formation of a stable uni-molecular structure. Compared with RNA1b-FAM, RNA1a-FAM mi-grated significantly faster, indicating a more compact structure forRNA1a. As there are only two Gs located in the fourth tract ofRNA1b, the structure of this RNA is expected to be less stable thanthat of RNA1a, which contains three stacked G-quartets.

To further confirm the G4 structures, we conducted 1H nuclearmagnetic resonance (NMR) analysis using chemically synthesizedRNAs (RNA1a and RNA1b) (13, 27). Imino proton peaks withinthe 10.0- to 11.5-ppm region are highly characteristic of the Hoogs-teen hydrogen bonds of G-quartets (28). Accordingly, the 1H NMRspectrum of RNA1a revealed well-resolved imino peaks within thisregion (red spectrum in Fig. 2B and fig. S6), indicating the formationof G4 structures. The 1H NMR spectrum of RNA1b demonstratedbroad imino peaks within the same range (red spectrum in Fig. 2C andfig. S7), suggesting inhomogeneous G4 RNAs. Because there were onlytwo G bases at the fourth G track, a two–G-tetrad motif was the mostprobable structure for RNA1b. Hence, RNA1b could potentially beinvolved inmultiple alternative foldingpatterns (29). Furthermore, the effectsof antisense oligonucleotides (ASOs), includingAS-RNA1a andAS-RNA1b(Table 1), were examined. When RNA1a was probed with AS-RNA1a,additional peaks appeared at the lower field of 11.5 to 13.5 ppm (bluespectrum in Fig. 2B), suggesting the partial conversion of G4 into double-strandedRNAs (dsRNAs) (20). A similar phenomenonwas alsoobservedfor RNA1b and AS-RNA1b (blue spectrum in Fig. 2C).

In a long RNA structural context, the folding of a specific structuremight be influenced by the neighboring sequences (30, 31). MfoldWeb server is used for folding analysis of the primary sequence of the


HCV C gene (32). The results indicated that RNA1a and RNA1b arelocated in a very structured (dsRNA) region (blue zone in figs. S8 andS9). Here, longer RNAs (RNA1a Long and RNA1b Long in table S4)containing the sequence in the green area (positions +264 to +311; figs.S8 and S9) were synthesized. Although most of the peaks are centered inthe “ds region,” the NMR results showed that the short motifs, includingRNA1a and RNA1b, were capable of forming G4 structures (redspectrum in fig. S10). Moreover, G-A mutations largely disrupted theG4 formation (blue spectrum in fig. S10).

Highly stable parallel G4 RNAs of the target HCV sequencesTo confirm the formation ofG4 structures in the targetHCVsequences,we performed circular dichroism (CD) analysis. As shown in fig. S11,the recorded spectra were consistent with parallel G4 structures (33).For RNA1a, the high stability of G4 RNAwas confirmed through smallvariations of CD spectra observed at elevated temperatures (fig. S12A),whereas the spectra of RNA1b showed more variation within the samerange (fig. S12B), indicating a less stable G4 RNA structure.

Next, we measured the spectra of these sequences in the presenceof NaCl or LiCl. As shown in fig. S13, the spectra of the sequences weresimilar to those recorded in KCl solution, indicating parallel G4s.

Furthermore, CD spectroscopy of RNA1a revealed a minimumpeak at 210 nm in the presence of AS-RNA1a (fig. S14A), whichwas highly reminiscent of the spectral signature associated withthe A-form of dsRNA. Moreover, increasing the level of AS-RNA1apromoted a shift from 264 to 275 nm, indicating a structural switchalleviated from G4. When RNA1b was titrated with less AS-RNA1b,the minimum at 210 nm could be evidently observed (fig. S14B), con-firming the increased G4 stability of RNA1a compared with RNA1b.

Fig. 1. Graphical representations of G-rich sequences consensus in the HCV genome. (A and B) A total of 1056 partial cds of the C gene for (A)subtype 1a and 1025 partial cds of the C gene for (B) subtype 1b were retrieved from the National Center for Biotechnology Information Web site(www.ncbi.nlm.nih.gov) and the HCV database (www.hcv.lanl.gov/) and aligned using WebLogo software.

Table 1. Sequences of some oligomers used in our studies.

Oligomer
Sequence (from 5′ to 3′)
RNA1a
5′-GGGCUGCGGGUGGGCGGGA-3′
RNA1b
5′-GGGCAUGGGGUGGGCAGGA-3′
RNA1a-FAM
5′-FAM-AGGGCUGCGGGUGGGCGGGA-3′
RNA1a–Mut-F
5′-FAM-AGAGCUGCGAGUGAGCGAGA-3′
RNA1b-FAM
5′-FAM-AGGGCAUGGGGUGGGCAGGA-3′
RNA1b–Mut-F
5′-FAM-AGAGCAUGGAGUGAGCAAGA-3′
AS-RNA1a
5′-UCCCGCCCACCCGCAGCCC-3′
AS-RNA1b
5′-UCCUGCCCACCCCAUGCCC-3′
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To quantitatively determine the G4 stabilities of target RNA, weconducted a thermal CD study (34). Because the unfolding process forRNA1a in the presence of a physiological concentration of K+ (100mM)could not be accomplished (fig. S15), thermal studies were initially con-ducted using subphysiological levels of K+. Figure S16A shows thethermal profiles of RNA1a in the presence of different salts. The melt-ing temperature (Tm) for RNA1a was dependent on the K+ concentra-tions. The alkali metal ion dependence for the stabilization of foldedRNA1a, on the basis of the Tm, was in the order K+ > Na+ > Li+ (fig.S16A and table S5), which is highly characteristic of G4 structures (35).Consistent with CD spectroscopy studies at variable temperatures, theTm of RNA1b was lower than that of RNA1a (fig. S16B), suggesting aless stable structure. An evident multiphasic stage was characterized forRNA1b (fig. S16B), further supporting the notion of the inhomogeneousG4 conformations of this RNA.

In addition, the Tm for RNA1a or RNA1b was independent of thestrand concentration, consistent with intramolecular G4 formation (fig.S17) (36). The melting curve analysis also revealed hysteresis in thecooling-versus-heating ramps (fig. S18), demonstrating slow foldingkinetics for target RNAs (20).

Stabilization of G4 RNAs through G4 ligandsRecently, a number of representative G4 DNA ligands, such as PDP (37)(structure in Fig. 3A) andTMPyP4 (38) (fig. S19), have been developed. Asdemonstrated in Fig. 3 (B and C) and fig. S20, the binding of G4 ligands totarget RNA resulted in a significant temperature shift in the meltingcurve, indicating evident G4 RNA stabilization at physiological ionicstrength. To further confirm that G4 ligands act specifically on HCVG4 RNAs, we incubated these molecules with the G4-mutated versionof target RNAs. No significant temperature shift in the melting curvewas observed (fig. S21). We showed that PDP increased the G4 stabilityof target RNA and therefore inhibited the opening of the G4 motifthrough ASO treatment, using a fluorescence resonance energy transfer(FRET) kinetic assay (figs. S22 and S23) (20).

G4 ligands block RNA-dependent RNA synthesisNext, we examined whether ligand-mediated G4 stabilization can reg-ulate RNA replication. In the HCV life cycle, the viral RNA is repli-cated by RdRp NS5B (39). For the simulation of this process, an RNA


stop assay is designed (Fig. 4A) (40, 41) using 3Dpol, which is an RdRpwith a primer-dependent mechanism (42, 43). The HCV G-richsegments (red part in Fig. 4A) have been incorporated into the 5′ endsof RNA templates, and the 5′ FAM-labeled primer p15 (green part inFig. 4A) is designed to target the 3′ end of templates (blue part in Fig.4A). Upon the addition of enzyme and nucleotide triphosphate(NTP), p15 is extended along the complementary template RNAs.When no G4 ligand is present, a full extension can be achieved (lefthalf in Fig. 4A). In contrast, if the binding of G4 ligands occurred, theextension would be stopped at the G4 site (right half in Fig. 4A).

As shown in lane 3 of Fig. 4B, fully extended products were ob-served along template 1a-G4, and few stopped products were observed(due to G4 formation) in the absence of G4 ligand. However, whenincreasing amounts of G4 ligand were incubated with template 1a-G4,the template-directed primer extension was gradually inhibited at theG4 site (lanes 4 to 7 in Fig. 4B). On the contrary, the site-specific ter-mination event was not characterized for G4-mutated template uponthe addition of G4 ligand (lanes 10 and 11 in Fig. 4B). The G4 site–specific blockade in RNA synthesis was also detected when template1b-G4 was treated with G4 ligands (fig. S24). Consistent with theresults of the melting studies, ligand PDP exhibited superior G4 sta-bilization and polymerization inhibition.

Inhibition of the full-length C gene expression through G4RNA stabilizationTo investigate the effects of G4 RNA stabilization on the expression of afull-length HCV C gene, we performed Western blot analysis. Plasmid24480 (pMO29) carrying the genotype 1bC gene (fig. S25) and the con-served backbone [pcDNA3.1(+)/pEV 204-Hind III] was used (44). Asdemonstrated in Fig. 4C, both PDP and TMPyP4 evidently inhibitedthe expression level of theHCVCgene. To better understand themech-anism behind the inhibition of HCV C gene expression through G4ligands, we examined TMPyP2 (structure in fig. S19), a positional iso-mer of TMPyP4 with low affinity for G4DNA (38). As demonstratedin fig. S26, TMPyP2 showed much less G4 RNA-stabilizing activitytoward RNA1b, contained in pMO29. Comparisons of TMPyP2 (lane4 in Fig. 4C) and TMPyP4 (lane 1 in Fig. 4C) on the basis of HCV Cgene expression were consistent with the above results. To demonstratethat the observed effects are specific for G4 sequence, we prepared a

Fig. 2. Synthetic HCV G-rich sequences form G4 RNAs. (A) Formation of compact G4 RNAs characterized on the basis of species that move morerapidly than the G4-mutated oligonucleotides. Lane 1, RNA1a–Mut-F; lane 2, RNA1a-FAM; lane 3, RNA1b-FAM; lane 4, RNA1b–Mut-F. (B) G4structures of RNA1a evidenced by 1H NMR. The chemical shift of Hoogsteen imino peaks in the range of 10.0 to 11.5 ppm was partially suppressedthrough AS-RNA1a (100 mM) in favor of Watson-Crick imino peaks. The expanded imino proton signals were analyzed by TopSpin 2.0 software.(C) G4 structures of RNA1b evidenced using 1H NMR.

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relevant control plasmid with a G4-mutated sequence. The followingresults (fig. S27) indicate that both PDP and TMPyP4 did not inhibitthe translation of the G4-mutated C gene. Together, these results sug-gest that because of the actions of G4 ligand in binding to and stabilizingG4RNA in the indicated region of the C gene (fig. S25), it is a promisingcandidate for further study.

Repression of enhanced green fluorescent proteinexpression through G4 RNA stabilizationNext, an enhanced green fluorescent protein (EGFP) reporter gene sys-temwas built to examine whether ligand-mediated G4 stabilization canregulate protein expression.We constructed a variety of pEGFP-C1 de-rivatives (45) after cloning the 21-bp sequences, containing either wild-type HCVG-rich sequences or the G-to-Amutant sequences. PlasmidsGFP-1a core G4, GFP-1b core G4, GFP-1a core Mut, and GFP-1b coreMut were prepared, and the insert was placed immediately downstreamof the human cytomegalovirus immediate early promoter (1 to 589) andupstream of the EGFP-cds present on the parental plasmid (figs. S28Aand S29A).We next performed a confocal fluorescence assay. Upon ad-


dition of G4 ligand, the expression of EGFP inGFP-1a core G4 or GFP-1bcore G4 was significantly inhibited compared with the DMSO treatment(fig. S28B), whereas the expression of EGFP in G4-mutated plasmids orthe empty vector was not influenced under the same conditions (fig.S29B). Consistent with the previous results, TMPyP2 did not inhibit theexpression of EGFP (fig. S28B). Together, the results suggest that G4ligands inhibit reporter gene expression by targeting HCVG-rich RNAs.

To quantitatively evaluate the effects of G4 ligands on EGFP ex-pression, we performed flow cytometry, and the results showed thatEGFPwas expressedmuch less efficiently inGFP-1a coreG4– orGFP-1bcore G4–transfected cells following G4 ligand treatment compared withtheDMSOcontrol (fig. S28C). In contrast,G4 ligands had amuch less pro-nounced effect on EGFP expression in G4-mutated plasmids. Consistentwith previous results, PDP showed better inhibition of EGFP expression.

G4 ligands inhibit HCV replication in cellsNext, we investigated whether G4 ligands can inhibit HCV infection.Currently, the most commonly used infectious HCV culture system isbased on JFH1 (Japanese fulminant hepatitis 1, genotype 2a) (46),

Fig. 3. G4 DNA ligand can stabilize target HCV G4 RNAs. (A) The structure of compound PDP. (B and C) Melting profiles of RNA1a (8.0 mM) orRNA1b (8.0 mM) were recorded in 10 mM tri-HCl buffer (pH 7.0) (100 mM KCl), in the absence or presence of PDP (8.0 mM).

Fig. 4. G4 ligands inhibit RNA-dependent RNA synthesis and HCV C gene expression through G4 RNA stabilization. (A) Schematic repre-sentation of the RNA stop assay. The artificial RNA template containing HCV G-rich or G4-mutated sequences was used. (B) The extended RNAs oftemplate 1a-G4 (lanes 2 to 7) or template 1a-G4 mut (lanes 8 to 11) analyzed through denaturing PAGE. The arrows indicate the positions of the full-length product, G4-pausing product, and free primer. The fully extended product and the template RNA formed a stable duplex, which was notdenatured and moved much slower than the corresponding ssRNA. Lane 1, RNA markers (p15, m17, m19, m21, and m39-1a in table S4); lanes 2 and3, no enzyme or no PDP control; lanes 4 to 7, G4 ligand–dependent inhibition; lanes 8 and 9, no enzyme or no PDP control; lanes 10 and 11,treatment with the G4 ligand. (C) Western blot analysis showing the suppression of HCV C gene expression through G4 ligands. The values indicatethe percentage of densitometry of the HCV Core protein relative to b-actin. DMSO, dimethyl sulfoxide; nt, nucleotides.

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which undergoes efficient replication in Huh-7 cells and other cell lines(47). We analyzed the sequence of the C gene in the JFH1 genome andalso identified the G4-forming sequence around the same region (fig.S30). A similar sequence alignment of the C gene for subtype 2a wasfurther performed (table S6 and figs. S31 to S33), and a conserved G4motif was also identified (fig. S34). The 1H NMR spectrum of RNA2ademonstrated broad imino peaks within the 10.0- to 11.5-ppm range(red spectrum in fig. S35A), providing direct evidence for G4 formation.G-A mutations in RNA2a resulted in the disappearance of the Hoogs-teen imino proton resonances (blue spectrum in fig. S35A), and the G4structure was stable (S35B). Hence, JFH1 is a suitable model for testingthe anti-HCV strategy. Here, JFH1-infected Huh-7.5.1 cells were trea-ted with different G4 ligands. Quantitative real-time polymerase chainreaction (RT-qPCR) was performed as previously described (48), andHCV RNA levels were determined relative to the transcription ofglyceraldehyde-3-phosphate dehydrogenase (GAPDH) in host cells.Because interferon-a (IFN-a) is best known as an effective treatmentfor HCV infection, it is used as a positive control in our studies examiningthe anti-HCV effects of G4 ligands. As shown in fig. S36, viral RNA levelsin JFH1-infected cells were markedly decreased by G4 ligands in adose-dependent manner, with measured median inhibitory concentra-tions of 1.2 and 3.2 mM for PDP and TMPyP4, respectively.

Next, we further evaluated the antiviral activity of G4 ligandsagainst two different intergenotypic HCV chimeras. The H77/JFH1chimeric genome contained the genotype 1a Con1 sequence fromthe C gene (sequence in fig. S37) to the NS2 region of H77 (genotype1a) and the nonstructural region of JFH1 (49), and the Con1/JFH1 chi-meric virus contained the genotype 1b Con1 sequence from the C gene(sequence in fig. S38) to the first 99 nucleotides of the NS2 gene and therest of the sequence from JFH1 (50, 51). The RT-qPCR results revealedthat PDP and TMPyP4 display adequate activity against HCV H77/JFH1 or Con1/JFH1 infection and evidently inhibit the viral RNA levelsat the C gene (figs. S39A and Fig. 5A). Not surprisingly, bothG4 ligandsalso effectively inhibit the viral RNA levels at 5′UTR in living cells (figs.S39B and Fig. 5B).

Moreover, Western blot analysis was performed to determine theCore protein levels of H77/JFH1- or Con1/JFH1-infected Huh-7.5.1cells using the commercial anti–HCV Core antibody (1a or 1b)(52). As shown in Fig. 5C and fig. S40, the level of HCV Core proteinexpression in cultured cells was significantly decreased by G4 ligandsin a dose-dependent manner. The nearly complete absence of the HCVCore protein was observed in infected cells treated with 2.5 mM PDP,indicating total viral inhibition. Consistent with these results, TMPyP4treatment significantly reduced the level of the HCV Core protein to10 mM, whereas a higher concentration of TMPyP2 did not inhibitHCV infection. Together, these results suggest that PDP is a more ef-fective agent for the inhibition of HCV Core protein expression.

Next, we also investigated the antiviral effects of G4 ligands on JFH1virus. The NS3 acts as a serine protease/helicase, which is an importantcomponent in the HCV replication complex. Therefore, the amount ofNS3 is a good indicator of the level of HCV activity in cells (53) andwas selected as a target in this assay. As demonstrated in Fig. 5D, G4ligands significantly inhibited the level of JFH1 virus and decreased theexpression of HCV NS3 protein in infected cells, and total viral inhibi-tion could be achieved through 2.5 mM PDP treatment. These resultsconfirm that PDP is a better antiviral agent against genotype 2a HCV.

During HCV replication, the positive-sense RNA genome is usedas a template to produce a negative-strand RNA intermediate (HCV−


RNA) (39). To further demonstrate that targeting G4 inhibits HCVreplication in vivo, we evaluated the effects of G4 ligands on the levelof HCV− RNA using Tth-based RT-qPCR (54) in JFH1-infectedHuh-7.5.1 cells. Our results clearly showed that G4 ligands signifi-cantly inhibit HCV− RNA levels in living cells in a dose-dependentmanner (fig. S41).

G4 mutations in the HCV C gene disrupt G4ligand–virus interactionsTo further demonstrate the existence of G4 RNAs in the HCV ge-nome under physiological conditions, we used a pull-down strategy(37, 55). Because the introduction of a biotin affinity tag on PDPhas been successfully applied in the selective enrichment of G4 DNAs(37), the same molecule (structure in fig. S42) was used in this assay.As demonstrated in fig. S43, biotin modification on PDP did not im-pair the stabilization of HCV G4 RNAs. Xba I restriction digestion ofthe plasmid pJ6/JFH1 at the 3′ end of the HCV complementary DNA(cDNA) was performed (56), and the linearized plasmid was tran-scribed in vitro to generate full-length HCV genomic RNA. Moreover,a pJ6/JFH1–G4-Mut plasmid containing a G4-mutated sequence in theC gene was prepared using overlapping extension PCR (OE-PCR).The transcribed J6/JFH1 or J6/JFH1–G4-Mut RNA was incubatedin the presence or absence of biotin-PDP and sonicated to shear thegenomic RNA. Subsequently, hydrophilic streptavidin-coatedmagnetic beads were used to capture the desired G4 RNAs (schemein fig. S44). As shown in fig. S45, an evident peak at approximately264 nm, representing parallel G4 RNAs, was observed for recoveredwild-type viral RNA (green line), whereas almost no signals were ob-served for G4-mutated viral RNA (blue line) and the control samplewithout using biotin-PDP (black line).

We next performed RT-qPCR to determine the abundance of theHCV G4 motif in RNA samples before and after pull-down manipu-lation (57), using G4-fwd and G4-rev (sequences in table S4). The nor-malized results revealed an approximately 28-fold enrichment (DDCT =4.82), whereas the control assay that did not use biotin-PDP did notdisplay any enrichment.

Next, we separately delivered the transcribed J6/JFH1 and J6/JFH1–G4-Mut RNA into Huh-7.5.1 cells through electroporation. The cellswere infected and treated with various G4 ligands, and the replicated viralRNA levels were determined relative to the host cell GAPDHmRNA. Asdemonstrated in Fig. 6A, much more attenuated inhibitions were ob-served for the J6/JFH1–G4-Mut virus. Western blot analysis furtherindicated that G4 ligands strongly inhibited Core protein expressionof the wild-type, but not of the G4-mutated, J6/JFH1 (Fig. 6B). Theseresults provided direct and solid evidence that G4 RNA in the HCV Cgene represents a cellular target for typical G4 ligands such as PDP.

To further confirm whether the G4 motif is a molecular target ofG4 ligands, we evaluated the activity of these molecules toward a dif-ferent natural virus without a G4-forming sequence. Influenza A virus,a negative-sense ssRNA virus, was examined (58). As expected, G4ligands were much less effective on this virus strain (fig. S46).

Previous investigations demonstrated that ASOs can bind to G4RNA and affect specific mRNA translation (20, 59). Here, ASOs com-plementary to G4 were delivered into different HCV-infected cells.The results clearly demonstrated that such ASOs can destabilize theHCV G4s and show an opposite effect on RNA replication whenusing G4 ligands (fig. S47). As expected, the stimulatory effect on viralreplication was significantly alleviated using mutant ASOs. The effect

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observed for G4-mutated virus was nearly abolished in the presence ofASO targeting the same nucleotide position.

Full HCV genome forms G4 at the targeted site in cellsBecause G4 structures have been visualized in human cells usingantibody-based fluorescence imaging (11, 60, 61), we applied the samestrategy to verify whether the target sequence truly formed G4 in cells.


The well-developed G4 binding antibody BG4 (11) was used in theseexperiments. We first transfected FAM-labeled target RNAs into Huh-7.5.1 cells (fig. S48). After fixation and permeabilization, amplified redfluorescence indicating G4 formation was generated (the second columnof images) according to a previously described method (60). Yellowregions indicating colocalization (indicated by white arrows in therightmost column of images) were detected in the cytoplasm of cells.

Fig. 5. G4 ligands suppress intracellular HCV replication. (A) RT-qPCR was used to determine the amount of HCV RNA in HCV Con1/JFH1-infected Huh-7.5.1 cells treated in triplicate with the G4 ligands or control (DMSO or IFN-a). IFN-a was used at 150 ng/ml. The values observedwere normalized to GAPDH. All data are presented as the means ± SEM from three independent experiments. The error bars reflect the SD. G4ligand groups versus DMSO group, *P < 0.05. The primers were designed to target the C gene of Con1/JFH1 RNA. (B) RT-qPCR was performed, andthe primers were designed to target the 5′UTR of Con1/JFH1 RNA. (C) Western blot analysis showed the suppression of intracellular HCV replication.A commercial anti–HCV Core 1b antibody was used, and the values indicate the percentage of densitometry of the target HCV protein relative tob-actin. (D) Western blot analysis was performed, and a commercial anti–HCV nonstructural protein 3 (NS3) antibody was used for detection.

Fig. 6. G4-disruptive mutations in the HCV C gene inhibit G4 ligand–virus interactions. (A) RT-qPCR was performed. The primers were designed totarget the C gene of J6/JFH1 virus. All data are presented as the means ± SEM from three independent experiments. The error bars reflect the SD.(B) Western blot analysis was performed. The values indicate the percentage of densitometry of the target HCV NS3 protein relative to b-actin. Lane 1, noHCV control; lanes 2 to 7, J6/JFH1–G4-Mut virus; lanes 8 to 13, J6/JFH1 virus.

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Moreover, the colocalization was inhibited after the ASO (targetingG4 site) treatment (the second, fourth, and sixth rows of images).We did not observe the colocalization (bottom row of images) usingthe G4-mutated sequence (RNA1b–Mut-F). The results providedstrong evidence that BG4 binds to intracellular G4 structures formedin target RNAs.

To confirm that the target G4 motif was targeted by the G4 ligandin living cells, we performed further tricolor confocal microscopy (fig.S49). The signals for biotinylated G4 ligand (biotin-PDP) were devel-oped using allophycocyanin (APC)–conjugated streptavidin (the mid-most column of images). As shown in the merged channel (therightmost column of images), the overlapping regions of red, green,and blue generated white images (indicated by white arrows), indicat-ing the binding of biotin-PDP, BG4, and target G4 RNA in cells. Inaddition, the ASO (targeting G4 site) treatment inhibited the observedcolocalization (the second, fourth, and sixth rows of images). Hence,target G4 RNAs were directly targeted by the G4 ligand.

To further verify whether the full HCV genome forms G4 at thetarget site in cells, we performed intracellular tracking of G4 in HCV(fig. S50). The ASO complementary to the adjacent sequence upstreamof the target G4 site (probe1a-FAM, probe1b-FAM, or probe2a-FAM intable S4) was used as a probe to visualize HCV in cells (the leftmostcolumn of images). The binding of the probe, BG4, and biotin-PDPis indicated as overlapping white regions (indicated by white arrowsin the rightmost column of images). As expected, the colocalizationwas disrupted after treatment with the ASO complementary to thetarget G4 site (the second, fourth, and sixth rows of images). More-over, no overlapping white region was observed in the G4-mutatedvirus (bottom row of images). These results provided direct evidencethat G4 ligand binds to the target HCV G4 site under physiologicalconditions.

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DISCUSSION

Despite the development of combined treatmentwith IFN and ribavirinas an antiviral therapy for HCV (62), IFN is poorly tolerated by manypeople and has limited availability in many countries (63). Moreover,combination therapy has significant side effects (64). Sofosbuvir-basedtherapy is a new and very effective treatment for HCV infection, butis extremely expensive (65). Hence, there is an urgent need for betteranti-HCV regimens with lower costs. Here, we conducted explicit bio-informatics analysis of the HCV genomes of all available genotypes.Remarkably, we first identified a highly conserved G-rich region intheHCVCgene of all available genotypes.Hence, this region representsa good candidate for the formation of G4 structures.

In a collection of critical studies, G4 DNA has been treated as aregulatorymotif (66–68), making it a potentially important target fordrug development (69, 70). Recently, a G4-based DNA aptamer wasused to inhibit HIV-1 replication ex vivo (71). In contrast to G4 DNAs,G4 RNAs are much less well characterized. G4 RNA structures exhibitmore stability than do G4 DNA versions (16), and a previous study hascharacterizedG4RNA in the 5′UTR of theNRAS proto-oncogene (35),whose expression level was modulated through this element. Remark-ably, it has been reported that EBV-encoded nuclear antigen 1 mRNAtranslation could be regulated through stabilization/destabilization ofG4 RNAs (20). These findings indicate that G4 RNA within virally en-coded transcripts might play an important role in translational control


and immune evasion.However, the reportedG4RNAswere all producedduring DNA transcription.

Here, we report the first evidence for the presence of G4 structurein HCV. Because patients with HCV genotype 1 responded poorly tothe current therapy (72), we aligned more than 1000 sequences of theC gene for subtype 1a or 1b from the HCV database and generatedtwo representative sequences with high potential to form stable G4RNAs. Various assays have been used to confirm the formation ofG4 RNA. A 1H NMR study revealed far fewer imino peaks in theG4 region for the longer RNA (fig. S10). This analysis, together withthe secondary structure prediction (figs. S8 and S9), may suggest acompetition between stable stem-loop structures and the proposedG4 structures. Hence, we propose that a target genome sequence doesnot form a very stable G4 structure under native conditions but ratheris induced to refold into a G4 by the presence of a G4 ligand. Furtherstudies confirmed the strong interactions between HCV G4 RNAs andG4 ligands in the presence of potassium at physiological concentra-tions. Although it has been reported that TMPyP4 might destabilize,rather than stabilize, G4 RNAs (73), this compound acts specificallyon target HCV G4 RNAs and shows a much weaker binding to theG4-mutated RNAs.

It has been demonstrated that stable G4 RNA motifs in 5′UTRsmight block the translation initiation of proteins in mammalian cells(35). A previous study also demonstrated the suppression of gene ex-pression through G4 RNA formation in several open reading framesencoded within the Escherichia coli genome (74). However, in thisstudy, G4 RNAs only inhibited the translation of specific proteins.During the life cycle of HCV, the positive-sense RNA genome is rep-licated through an RNA intermediate (39). The RNA stop assay in thepresent study revealed that this specific G4 RNA formation intemplate RNA led to the inhibition of RNA-dependent RNA syn-thesis, suggesting the direct inhibition of viral genomic replicationthrough G4 ligands. Moreover, the expression of EGFP containingHCV G-rich RNAs decreased significantly after treatment with G4 li-gands, whereas constructs containing G4-disruptive mutations (G toA) were relatively insensitive to G4 ligands. In conjunction with thefull-length C gene expression assay, we propose that the expressionlevel of the HCV C gene could be down-regulated through selectiveG4 RNA formation.

On the basis of these new findings, we further evaluated theantiviral activities of G4 ligands. Here, we first revealed that G4 ligandsinhibit intracellular HCV replication (genotype 1a, 1b, or 2a) at bothRNA and protein levels. We also confirmed that the specific cel-lular target of active G4 ligands was HCV G-rich RNA, using aninactive analog as a negative control. The results demonstrated thatTMPyP4 efficiently inhibited the expression of a full-length HCV Cgene on an engineered plasmid. TMPyP4 further suppressed intra-cellular HCV (genotype 1a, 1b, or 2a) levels in infected Huh-7.5.1 cells.In contrast to TMPyP4, much alleviated effects in all these studies couldbe characterized for TMPyP2. Because TMPyP2 is a bad ligand for thebinding and stabilization of G4 DNAs (38), these results suggested thatthe action mode of TMPyP4 in the inhibition of HCV replication mightbe mediated through binding to target HCV G4 RNA.

To further validate the cellular target for G4 ligand, we mutated theG4-forming sequence between positions +267 and +279 in the J6/JFH1C gene. The enrichment of G4 RNA in the pull-down extracts ofJ6/JFH1 was further demonstrated through CD and RT-qPCR deter-mination of RNA recovered from beads. In contrast, almost no RNA

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was recovered from the beads for J6/JFH1–G4-Mut RNA. Together,these results indicate that G4 RNA on the HCV C gene could be se-lectively recognized and targeted by the G4 ligand. Because the G4-mutated virus survived and reproduced in Huh-7.5.1 cells, this mutantvirus could be an ideal model for examining the on-target specificityof the G4 ligand. Comparative analysis of the antiviral activities of theG4 ligandwas performed throughRT-qPCRandWestern blot analyses,and the results showed a significant decrease toward the G4-mutatedvirus. We hypothesized that G4 RNA formation in the HCV C genecould lead to viral mRNA release from ribosomes due to the steric hin-drance of the stable structure.

We further examined the G4 structures in HCV-infected Huh-7.5.1 cells using indirect immunofluorescence microscopy, and clearoverlapping regions of BG4, HCV probe, and biotin-PDP immuno-fluorescence indicated G4 formation in the full HCV genome underphysiological conditions. We also characterized the competition be-tween ASO and BG4 (also biotin-PDP) for binding to the target G4site. For G4-mutated virus, almost no binding between the probe andBG4 (or biotin-PDP) was observed in the cytoplasm of cells. Together,these findings suggest that HCV G4 RNA in the C gene could serve asa new cellular target for anti-HCV drug development.


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MATERIALS AND METHODS

Experimental designThere is an urgent need for better anti-HCV regimens with lowercosts. Here, our bioinformatics analysis reveals a highly conservedG-rich consensus sequence within the HCV C gene, despite the highgenetic variability of riboviruses; we further show using variousmethods that the HCV C consensus sequences can fold into uni-molecular G4 RNA structures both in vitro and under physiologicalconditions. We then demonstrate that G4 ligands effectively inhibitintracellular HCV replication (genotype 1a, 1b, or 2a) and further val-idate the specific cellular target of G4 ligands to be HCV G-rich RNA.Ultimately, the stabilization of G4 RNA in the HCV genome repre-sents a promising new strategy for anti-HCV drug development.

SynthesisCompound PDP and biotin-PDP were synthesized according to a pre-viously described procedure (37).

Bioinformatics analysisSequence conservation analysis was conducted using Molecular Evo-lutionary Genetics Analysis (MEGA) software (version 6.0; available atwww.megasoftware.net). The multiple sequence alignment was alsocarried out with MEGA 6.0 software. HCV sequences were retrievedfrom the HCV database (www.hcv.lanl.gov/), and the graphical repre-sentation of the aligned sequences was generated using the WebLogosoftware program (version 2.8.2; http://weblogo.berkeley.edu/).

Gel electrophoresisSee the Supplementary Materials for a more detailed protocol.

NMR spectroscopy1H NMR spectra were recorded at 298 K using an 800-MHz BrukerAvance DRX-800 spectrometer equipped with a cryogenic triple-resonance inverse automatic tuning and matching probe. Water sup-


pression was achieved using the excitation sculpting method. TheRNA samples were dissolved in 10 mM phosphate-buffered saline(PBS) (pH 7.0) containing 100 mM KCl and 10% D2O at a final con-centration of 0.5 mM. The samples were annealed after heating at 90°Cfor 4.0 min and slowly cooled to 25°C. All experiments were performedusing a diffusion time (TD) of 100 ms, an eddy current recovery time(TE) of 50 ms, and a relaxation delay (TR) of 2 s.

CD studiesSee the Supplementary Materials for a more detailed protocol.

CD melting studiesG4 ligands were dissolved as a 10 mM stock solution in DMSO. Seethe Supplementary Materials for a more detailed protocol.

FRET kinetic assayDual-labeled G4 RNA probe (RNA1a-dual or RNA1b-dual)containing a donor fluorophore (FAM) and an acceptor fluorophore(TAMRA) was used. The probe samples (at a final concentration of200 nM) were prepared in 10 mM tris-HCl buffer (pH 7.0) (100 mMKCl) containing different amounts of PDP, and equilibrated at 4°Covernight. For the FRET kinetic assays, the ASO sample (ASO-1aor ASO-1b at a final concentration of 2.0 mM) was mixed at t = 0.Fluorescence detection was conducted at 25°C in kinetics mode.The same LS55 spectrometer was used with a 1-cm path length cell.The excitation and emission wavelengths were set to 494 and 580 nm,respectively.

RNA stop assay3Dpol was a gift from P. Gong (Wuhan Institute of Virology, ChineseAcademy of Sciences, Wuhan, China). The assay was performed as de-scribed previously (40). P15 (300 nM) and template RNA (600 nM)were annealed in a reaction buffer [50 mM Hepes, 20 mM NaCl,1.0 mM KCl, 5 mM MgCl2, and 4 mM DTT (pH 7.0) at 25°C] afterheating at 85°C for 2.0 min and slowly cooled to 4°C. Subsequently,NTPs (final concentration, 200 mM) and 3Dpol (0.02 mg/reaction) wereadded, and the reactions were performed at 33°C for 20 min. The re-actions were stopped after adding an equal volume of stop buffer (95%formamide) and heated at 90°C for 4.0 min. The products were loadedand separated on 20% denaturing polyacrylamide gels. Finally, the gelswere scanned by a Pharos FX Molecular Imager (Bio-Rad) operated influorescence mode.

Cell linesHuman embryonic kidney (HEK) 293 cells and Huh-7.5.1 cells werecultured under standard culture conditions.

EGFP expression repression through HCV G4RNA stabilizationHCV G-rich sequences were cloned into the pEGFP-C1 vector byPCR with the following primer sets. The forward primer was designedto include a pendant 5′ segment comprising an Nhe I cleavage site andan HCV G-rich sequence, along with the 3′ region perfectly matchingthe N terminus of EGFP in frame. The reverse primer was designed tohave an Hind III site at the 5′ end and a 3′ region, perfectly matchingthe C terminus of EGFP. Plasmid pEGFP-C1 (Clontech) was used asPCR template. These two primers (C1-1a-F and C1-R) were used toamplify the 758-bp fragment. The fragment was digested with Nhe I

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and Hind III, and subcloned into the same restriction sites of pEGFP-C1 vector to generate the plasmid construct pEGFP–C1-1a. pEGFP–C1-1b was prepared using the forward primer (C1-1b-F), togetherwith the same reverse primer (C1-R). The primer pair sets (C1-1aMut-F, C1-R and C1-1bMut-F, C1-R) were used to generatepEGFP–C1-1aMut and pEGFP–C1-1bMut. All constructs were con-firmed through sequencing. HEK293 cells were transfected with theplasmid using Lipofectamine 2000 (Invitrogen) at a 1:2 ratio (∼500 ngof plasmid/1 ml of reagent) and were subsequently treated for 48 hourswith different amounts of freshly dissolved G4 ligands. Subsequently, thecells were washed with PBS and fixed using 4% paraformaldehyde at25°C for 15 min. The cells were washed three times with PBS for 5.0 mineach and permeabilized with 0.5% Triton X-100 at 4°C for 10 min.Coverslips with the cells were then mounted using 50% glycerol in PBSand observed using a laser scanning confocal microscope (Leica TCSSP2). The excitation laser used for EGFP was 488 nm; the emissionof green fluorescence was collected using the 505- to 530-nm band-passfilter. A laser diode (excitation, 405 nm) and a band-pass filter (420 to480 nm) were used to record the fluorescence of Hoechst 33258 (blue).Flow cytometry was conducted on a flow cytometer (Coulter XL). Sta-tistical analysis was performed using EXPO32 v1.2 Analysis software(Beckman Coulter).

Construction of a plasmid carrying the G4-mutated C genepcDNA3.1-N-Flag vector (75) was a gift from H. Wang (WuhanInstitute of Virology, Chinese Academy of Sciences). The G4-mutatedC gene was chemically synthesized at Invitrogen, and the sequence inthe G-rich region (positions +265 to +279) was mutated (GAGGGCT-TGGGGTGG to GAAGGCTTAGTATTT). Two primers (1b coreG4-Mut F and 1b core G4-Mut R) were used to amplify the 595-bp frag-ment. The fragment was digested with Bam HI and Not I and sub-cloned into the same restriction sites of pcDNA3.1-N-Flag vector togenerate the plasmid construct 1b core G4-Mut.

Western blot analysis to detect HCV C gene expressionFor this assay, total proteins were extracted using a radioimmunopre-cipitation assay lysis buffer and quantitated using bicinchoninic acidprotein assay. The lysates were mixed with an equal volume of 2×loading buffer and boiled for 5 min. Then, 50 mg of proteins wassubjected to electrophoresis on 10% SDS-PAGE gels (1.0 mm thick ×15 cm long), along with 15 ml of Kaleidoscope protein standards(Bio-Rad), and transferred to polyvinylidene difluoride membranes(Millipore). After blocking with 5.0% bovine serum albumin in tris-buffered saline and Tween 20, target proteins on membranes wereprobed at 37°C for 1 hour with anti–HCV Core 1b antibody (1:1000dilution; Abcam). b-Actin was used as a loading control for analysisand was probed by monoclonal mouse anti-actin antibody (1:1000 di-lution; Abcam). After the final wash, the signal was developed withcorresponding horseradish peroxidase–conjugated secondary antibody(1:5000; Proteintech). Finally, protein bands were visualized on an x-rayfilm using enhanced chemiluminescence reagents (Millipore).

Antiviral activity assayGenotype 2a JFH1 HCVcc (cell culture–derived HCV) stock was gen-erated as previously described (46, 53). Genotype 1a/2a chimera (H77/JFH1 chimera) was a gift from Z. Qi (Department of Microbiology,SecondMilitaryMedical University, Shanghai, China), and genotype1b/2a chimera (Con1/JFH1 chimera) HCVcc stock (50) was a gift from


X. Chen (Wuhan Institute of Virology, Chinese Academy of Sciences).The infectious titers were determined after immunofluorescencestaining againstHCVNS5A (anti-NS5Apolyclonal antibody, a gift fromJ. Zhong, Unit of Viral Hepatitis, Institut Pasteur of Shanghai, China).Daclatasvir and IFN-a were used as positive controls. The cells werecultured in complete Dulbecco’s minimum essential medium (DMEM)and passaged every 3 to 5 days. The corresponding supernatants ofthe HCV-containing cell culture were collected and filtered through a0.45-mm filter membrane (Thomas Scientific), and the filtrates wereconcentrated with a Vivaspin (100-kD cutoff; Millipore). The concen-trated virus-containing mediumwas loaded onto a 20% sucrose cushionin an ultracentrifuge tube, and the cell culture–adapted virus particleswere purified using ultracentrifugation (4°C for 4 hours) at 28,000 rpm(SW48 rotor; BeckmanCoulter). The resulting pellets were resuspendedin 0.5 ml of complete medium, and naïve Huh-7.5.1 cells were infectedwith the diluted viruses at 3.0 multiplicity of infection for 4 hours. Afterinfection, the cellswerewashedwithPBS to removeunbound virus. Freshcomplete DMEM was added, and HCV-infected cells were treated for72 hours with different amounts of freshly dissolved G4 ligands. ForRT-qPCR, cells were lysed with TRIzol reagent (Invitrogen), and totalRNAwas extracted and purified according to themanufacturer’s instruc-tions. ThepurifiedRNAsweredissolved indiethyl pyrocarbonate–treatedwater and spectrophotometrically quantitated. Reverse transcriptionof RNA to synthesize cDNA was performed with a ReverTra Ace-a-Kit(TOYOBO). RT-qPCR was conducted in optical tubes in a 96-well mi-crotitre plate (Applied Biosystems) with an ABI PRISM 7700 SequenceDetector Systems (Applied Biosystems). SYBR Green Realtime PCRMaster Mix-Plus Kit (TOYOBO) was used, and fluorescence signalswere generated and recorded during each PCR cycle. HCV RNA levelsat the C gene and 5′UTR were quantitated, and the GAPDH mRNAof host cells was used as a control. The following primer sets were usedin this study: JFH1Core-F/R,H77Core-F/Core-R,Con1Core-F/Core-R,5′UTR-F/R, andGAPDH-F/R (table S4).Western blot analysis was con-ducted using the above procedure described for HCV C gene expres-sion, and the target proteins were probed with anti–HCV Core 1a or1b antibody (1:1000 dilution; Abcam) or anti-NS3 antibody (1:1000 di-lution; Abcam).

Construction of G4-mutated plasmids and G4-mutated virusTo construct a target plasmid containing a G4-mutated HCV C genewith multiple point mutations, we used a starting plasmid vectorpJ6/JFH1 (a gift from Z. Qi) (56). The multiple point mutationsat nucleotides 267, 269, 275, 278, and 279, respectively (GAGG-GACTCGGC TGG was changed to GAAGTACTCGTCTAC), wereintroduced after amplifying DNA fragments containing the mutations.Point mutations were created through OE-PCR of the region betweenthe unique restriction sites (Eco RI/Kpn I) of pJ6/JFH1 template DNA,and two primer pairs [forward primer in upstream region (J6 up F),reverse primer in upstream region (J6 up R); forward primer indownstream region (J6 down F), reverse primer in downstream region(J6 down R)] were used. The target fragment was digested with Eco RIand Kpn I and subcloned into the same restriction sites of the pJ6/JFH1vector to generate the plasmid construct pJ6/JFH1–G4-Mut, which wasfurther confirmed by sequencing.

In vitro transcription and activity assayIn vitro transcription reactions were performed according to the man-ufacturer’s instructions in the MEGAscript T7 Transcription Kit

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(Invitrogen) in a 30-ml reaction containing 3.0 ml of 10× reaction buffer,11.0 ml of nuclease-free water, 1.0 ml of Xba I–linearized pJ6/JFH1 DNAor pJ6/JFH1–G4-MutDNA (1.0 mg/ml), 3.0 ml of adenosine triphosphatesolution, 3.0 ml of cytidine triphosphate solution, 3.0 ml of guanosinetriphosphate solution, 3.0 ml of uridine triphosphate solution, and 3.0 mlof enzyme mix. In vitro transcription reactions were incubated at 37°Cfor 6 hours. The RNA transcripts were purified through spin-columnchromatography according to the manufacturer’s instructions (PureYieldRNA Midiprep System, Promega Corporation). The recovered RNAswere analyzed for purity and concentration with the NanoDrop ND-2000spectrophotometer. Delivery of in vitro–transcribed viral RNA intoHuh-7.5.1 cells (3 × 106) was performed through electroporation, andthe activity assay was performed accordingly. The primer sets JFH1Core-F/R and GAPDH-F/R were used in this study.

Pull-down assayIn vitro–transcribed RNAs (400 mg) were incubated in 10 mM tris-HCl (pH 7.0) buffer containing 100 mM KCl in the presence or ab-sence of 5.0 mM biotin-PDP and then sheared with the SB-5200 DTDsonicator (300 W; Ningbo Scientz Biotechnology) for 2 hours at highpower with a pulse of 30 s on/30 s off, to an average of 100 bp. Fourhundred microliters of each sonicated RNA sample was incubatedwith 20 ml of hydrophilic streptavidin magnetic beads (S1420S, NewEngland Biolaboratories) for 1 hour at 37°C. The separated magneticbeads were subsequently incubated with 10mMEDTA and 95% form-amide [2.5 ml of 0.2MEDTA (pH8.0) and 47.5 ml of formamide] at 90°Cfor 5 min. The eluted RNAs were purified through spin-column chro-matography and redissolved in 10 mM tris-HCl buffer (pH 7.0) con-taining 100 mM KCl. CD experiments were performed using a quartzcell with a 1.0-cm path length.

RT-qPCR validation of G4-specific enrichmentInput reverse-transcribed genomic DNA or reverse-transcribed G4-enriched DNA was diluted to 1 ng/ml, and 1 ml was used in triplicate20-ml qPCRs containing 1× LightCycler 480 SYBR Green I Master(Roche Diagnostics), 0.5 mM forward and reverse primers, and water.The reactions were run on a Roche LightCycler 480 II under stan-dard cycling conditions. The G4 locus-associated primers includedG4-fwd and G4-rev, and the fold enrichment was calculated as 2−dCT,where dCt = Ct (G4 enriched) − Ct (input).

ASO treatment enhances viral replicationHCV-infected Huh-7.5.1 cells (5 × 105) were transfected with either a19-mer ASO (AS-RNA1a, AS-RNA1b, AS-RNA2a, or AS-G4mut2a),a 19-mer mutant ASO (AS-mutant1a, AS-mutant1b, or AS-mutant2a),or a random 19-mer oligonucleotide (oligo control), each at a final con-centration of 100 nM, using Oligofectamine reagent (Invitrogen)according to the manufacturer’s protocol. At 48 hours after transfec-tion, the cells were lysed with TRIzol reagent, and total RNA wasdetermined through RT-qPCR. In these studies, the HCV RNA levelswere determined relative to the transcription of GAPDH in host cells.The primer sets (UTR and GAPDH) were the same as those forantiviral activity assay. Values for the “ASO” or “mutant ASO” groupswere calculated relative to the “oligo control” value.

Fluorescent imaging to visualize HCV G4 RNAs in cellsThe expression and purification of the G4-specific antibody BG4were performed according to the previously described protocol (11).


Huh-7.5.1 cells were transfected with FAM-labeled oligomers or full-length HCV genomes using Oligofectamine reagent (Invitrogen) andwere treated with or without 2.5 mMbiotin-PDP for 4 hours. The dem-onstration of full-length HCV genomes inside cells was confirmedthrough fluorescence in situ hybridization using a FAM-labeled ASOcomplementary to a sequence upstream of the target G4 site. Cells onglass coverslips were washed with a culture medium and PBS, fixed, per-meabilized, and blocked following a previous protocol (60). Specifically,cells were permeabilized in 0.02% Triton X-100/PBS buffer for 15 minat 4°C (76). Subsequently, immunofluorescence was measured. Briefly,the cells were incubated at 37°C with BG4 (4 mg/ml), followed by anti-Flag antibody (1:500 dilution; no. 2368, Cell Signaling Technology) andanti-rabbit rhodamine-conjugated antibody (1:500 dilution; R-6394, LifeTechnologies). APC-conjugated streptavidin (1:500 dilution; S868, LifeTechnologies) was used to demonstrate biotin-PDP. The excitationlaser used for the FAM group was 488 nm. Rhodamine was excitedwith a 568-nmdiode laser and detected using a 590-nm long-pass filter.APCwas excited at 639 nmand detectedwith a 647-nm long-pass filter.

Statistical analysisThe experimental data were analyzed with Student’s t test using theSPSS 17.0 software. *P < 0.05 was considered statistically significant.

SUPPLEMENTARY MATERIALSSupplementary material for this article is available at http://advances.sciencemag.org/cgi/content/full/2/4/e1501535/DC1Materials and MethodsFig. S1. Structural illustration of a G-quartet.Fig. S2. Conservation analysis of HCV genomes.Fig. S3. Premade sequence alignment in the central part of the HCV C gene, between positions+253 and +294.Fig. S4. Premade sequence alignment in the central part of the HCV C gene, between positions+253 and +294.Fig. S5. Premade sequence alignment in the central part of the HCV C gene, between positions+253 and +294.Fig. S6. Expansion of the 1H NMR spectra of RNA1a.Fig. S7. Expansion of the 1H NMR spectra of RNA1b.Fig. S8. Prediction of the RNA secondary structure of the C gene (subtype 1a) using free-energy minimization.Fig. S9. Prediction of the RNA secondary structure of the C gene (subtype 1b) using free-energy minimization.Fig. S10. G4 formation in a long structural context evidenced by 1H NMR.Fig. S11. Synthetic HCV G-rich sequences form parallel G4 RNAs.Fig. S12. G4 structure of RNA1a is more stable than that of RNA1b.Fig. S13. G4 RNAs are characterized in the presence of alkali metal ions (K+, Na+, or Li+).Fig. S14. HCV G4 RNA structures are destabilized through the ASO.Fig. S15. CD melting curves of HCV G-rich RNAs.Fig. S16. Influence of different alkali metal ions on the thermal stabilities of HCV G4 RNAs.Fig. S17. Analysis of concentration-independent melting curves of target HCV RNAs.Fig. S18. CD melting studies of target HCV RNAs.Fig. S19. Structures of TMPyP4 and TMPyP2.Fig. S20. G4 ligand stabilizes target HCV G4 RNAs.Fig. S21. Little interaction is observed between the G4 ligand and G4-mutated RNAs.Fig. S22. Schematic depiction of the inhibition of FRET through the binding between PDP andG4 RNA.Fig. S23. PDP binds to target G4 RNA and inhibits the trap by the corresponding ASO.Fig. S24. G4 ligand inhibits RNA-dependent RNA synthesis through G4 RNA stabilization.Fig. S25. Map of the plasmid 24480 (pMO29) and a sequenced portion of this plasmid forverification.Fig. S26. TMPyP2 does not stabilize G4 RNA for RNA1b.Fig. S27. G4 ligands do not suppress the expression of the HCV C gene containing a G4-mutated sequence.Fig. S28. G4 ligands repress the in vitro expression of EGFP through G4 RNA stabilization.

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Fig. S29. G4 ligands do not repress the in vitro expression of EGFP in empty vector or G4-mutated plasmids.Fig. S30. Sequence of the C gene for HCV JFH1 virus.Fig. S31. Premade sequence alignment in the central part of the HCV C gene (subtype 2a),between positions +253 and +296.Fig. S32. Premade sequence alignment in the central part of the HCV C gene (subtype 2a),between positions +253 and +296.Fig. S33. Premade sequence alignment in the central part of the HCV C gene (subtype 2a),between positions +253 and +296.Fig. S34. Graphical representation of G-rich consensus sequences in genotype 2a HCV genomes.Fig. S35. G4 RNA structure of RNA2a evidenced in different studies.Fig. S36. G4 ligands inhibit intracellular HCV JFH1 replication.Fig. S37. Sequence of the C gene for HCV H77.Fig. S38. Sequence of the C gene for HCV Con1.

Fig. S39. G4 ligands suppress intracellular HCV H77/JFH1 replication.

Fig. S40. Western blot analysis shows suppression of intracellular HCV H77/JFH1 replicationthrough G4 ligands.

Fig. S41. Detection of HCV− RNA using Tth-based RT-qPCR.

Fig. S42. Structure of biotin-PDP.

Fig. S43. Biotin modification on PDP does not impair the stabilization of HCV G4 RNAs.

Fig. S44. Pull down of G4 RNA through biotin-PDP.

Fig. S45. HCV G4 RNAs evidenced by a selective G4 affinity probe.

Fig. S46. G4 ligands do not inhibit influenza A virus without a G-rich region.

Fig. S47. ASOs destabilize the HCV G4s and show stimulatory effects on viral replication.

Fig. S48. Target HCV sequences form intracellular G4 structures.

Fig. S49. Target HCV sequences form intracellular G4 structures probed by the G4 ligand.

Fig. S50. G4 ligand binds to the target G4 site in the full HCV genome under physiologicalconditions.

Table S1. GenBank accession numbers of the HCV genomes analyzed.

Table S2. List of 1056 sequenced partial cds of the C genes for subtype 1a.

Table S3. List of 1025 sequenced partial cds of the C genes for subtype 1b.

Table S4. Sequences of oligomers used in our studies.

Table S5. Calculated Tm of synthetic HCV G-rich sequences in the presence of different alkalimetal ions.

Table S6. List of 143 sequenced partial cds of the C genes for subtype 2a.

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Acknowledgments: We thank the instrument and equipment sharing foundation of WuhanUniversity. Funding: X.Z., X.-L.Z., and T.T. thank the National Basic Research Program of China(973 Program) (grants 2012CB720600, 2012CB720603, and 2012CB720604), the National ScienceFoundation of China (grants 91213302, 81373256, 21432008, 31370197, 31221061, 81025008,21572173, and 21372182), and the Hubei Province’s Outstanding Medical Academic LeaderProgram. Author contributions: T.T. and S.-R.W. performed the bioinformatics and biophysicalanalyses. Y.-Q.M. and J.-Q.W. performed molecular cloning, RT-qPCR, Western blot analysis, andantiviral assays. C.-X.L. and W.-X.M. performed the synthesis. B.-S.F. and G.-H.X. performed theNMR assay. Z.-X.Q., L.-Y.W., Z.-G.W., and G.H. performed the pull-down assay. Y.-Q.C., F.W., Y.-Y.S.,S.P., and R.H. performed the ASO test. N.-F.P. and Y.Z. performed the influenza A virus test. X.Z., X.-L.Z.,and T.T. analyzed the data, discussed the results, and wrote the paper. Competing interests: Theauthors declare that they have no competing interests.Data andmaterials availability: Additionalexperimental details are included in the Supplementary Materials. All data needed to evaluate theconclusions in the paper are present in the paper and/or the Supplementary Materials. Additionaldata related to this paper may be requested from the authors.

Submitted 29 October 2015Accepted 18 February 2016Published 1 April 201610.1126/sciadv.1501535

Citation: S.-R Wang, Y.-Q. Min, J.-Q. Wang, C.-X. Liu, B.-S. Fu, F. Wu, L.-Y. Wu, Z.-X. Qiao, Y.-Y. Song,G.-H. Xu, Z.-G. Wu, G. Huang, N.-F. Peng, R. Huang, W.-X. Mao, S. Peng, Y.-Q. Chen, Y. Zhu, T. Tian,X.-L. Zhang, X. Zhou, A highly conserved G-rich consensus sequence in hepatitis C virus coregene represents a new anti–hepatitis C target. Sci. Adv. 2, e1501535 (2016).

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hepatitis C target−new anti A highly conserved G-rich consensus sequence in hepatitis C virus core gene represents a

Tian Tian, Xiao-Lian Zhang and Xiang ZhouGuo-Hua Xu, Zhi-Guo Wu, Gai Huang, Nan-Fang Peng, Rong Huang, Wu-Xiang Mao, Shuang Peng, Yu-Qi Chen, Ying Zhu, Shao-Ru Wang, Yuan-Qin Min, Jia-Qi Wang, Chao-Xing Liu, Bo-Shi Fu, Fan Wu, Ling-Yu Wu, Zhi-Xian Qiao, Yan-Yan Song,

DOI: 10.1126/sciadv.1501535 (4), e1501535.2Sci Adv

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A highly conserved G-rich consensus sequence in hepatitis ...small molecules specifically targeting G4 can stabilize this structure to reduc e RNA replication and inhibit protein translation