G-quadruplex (G4) is one of the most important secondary structures in nucleic acids. Until recently, G4 RNAs have not been reported in any ribovirus, such as the hepatitis C virus. Our bioinformatics analysis reveals highly conserved guanine-rich consensus sequences within the core gene of hepatitis C despite the high genetic variability of this ribovirus; we further show using various methods that such consensus sequences can fold into unimolecular G4 RNA structures, both in vitro and under physiological conditions. Furthermore, we provide direct evidences that small molecules specifically targeting G4 can stabilize this structure to reduce RNA replication and inhibit protein translation of intracellular hepatitis C. Ultimately, the stabilization of G4 RNA in the genome of hepatitis C represents a promising new strategy for anti–hepatitis C drug development.

Keywords

Here, we provide the first evidence that a highly conserved guanine-rich (G-rich) sequence is present in the HCV core (C) gene. Using various methods, we demonstrate the high potential of HCV C consensus sequences to form G4 RNAs. Furthermore, our biological findings provide direct evidences that, because of the actions of G4 ligands in binding to and stabilizing G4 RNA in the specific G-rich region of the C gene, they can reduce RNA replication and inhibit protein translation in HCV. Ultimately, the highly conserved nature of HCV G-rich RNA might represent a challenging new target for anti-HCV drug development.

G-quadruplex (G4) contains stacked planar G-quartets, which are stabilized through Hoogsteen hydrogen bonding (fig. S1) ( 10 – 12 ). G4 RNAs have been identified in telomeric transcripts ( 13 ) and 5′ or 3′ untranslated regions (UTRs) of mRNAs ( 14 – 16 ). Recently, G4 RNAs have been associated with some viruses ( 17 ) such as HIV ( 18 ), herpes simplex virus (HSV) ( 19 ), and Epstein-Barr virus (EBV) ( 20 ). BRACO-19, a G4-binding ligand, has been demonstrated to be an active anti-HIV agent ( 21 ). However, for HSV and EBV, G4 RNAs only form in nascent RNA transcripts during transcription, and HIV only exists in the latent form of proviral DNA upon entry into human cells. To the best of our knowledge, the roles of G4 RNA in these studies are confined to the regulation of transient gene expression at the translational level. Until recently, there have been no reports concerning the presence of G4 structure in riboviruses. Because RNA replication and translation occur on the same RNA of positive-sense single-stranded RNA (ssRNA) viruses, we suggest that HCV G4 RNA (if present) play a more direct role in regulating both of these processes.

Viruses rapidly mutate, and RNA viruses (particularly riboviruses, rather than retroviruses) have higher genetic variation than DNA viruses ( 1 ). Hepatitis C virus (HCV) infection is estimated to affect 2.8% of the population worldwide ( 2 ). Recent medical studies have focused on the development of small molecules targeting viral enzymes ( 3 , 4 ). However, resistance rapidly emerges in HCV patients who are treated with specific inhibitors targeting nonstructural proteins, reflecting the error-prone replication by NS5B, a viral RNA-dependent RNA polymerase (RdRp) ( 5 ). Hence, the development of compounds targeting highly conserved regions or structural motifs in the HCV genome would have the most potential to protect against not only the current virus but also the varied ones ( 6 , 7 ). Moreover, the use of small-molecule RNA binders as therapeutics is an area of intense interest at the interface of chemistry and biology ( 8 , 9 ).

RESULTS

Bioinformatics analysis to reveal HCV G-rich sequences On the basis of a genomic variance analysis, HCV was classified into multiple genotypes (1 to 7) with varied subtypes (22). We retrieved 77 complete genomic sequences of all available genotypes (GenBank accession numbers in table S1) and conducted bioinformatics analysis to assess the level of sequence conservation (23). As shown in fig. S2, a low ratio (30.89%) of conserved nucleotide sites across the whole genome was observed, whereas much higher conservation (46.77%) was identified within the nucleotide sites of the C gene. Thus, we are particularly interested in the development of a new antiviral strategy targeting the HCV C gene. We next performed multiple sequence alignments of the C gene across these 77 HCV genomes (figs. S3 to S5). In this example, a G-rich consensus sequence harboring four G-tracts was observed in the central region of the C gene, between positions +259 and +285. A high proportion (47.37%) of the nucleotide sites is conserved across such a G-rich region, which is highly characteristic of sequences with high potential to form G4 structures. HCV subtypes 1a and 1b are commonly observed in East Asia and North America (24), prompting significant interest in the study of these specific subtypes within the field. Hence, we aligned 1056 partial coding sequences (cds) of the C gene for HCV subtype 1a and 1025 cds for HCV subtype 1b (tables S2 and S3), retrieved from the HCV database. WebLogo was used to generate a graphical representation of the aligned sequences (25) between positions +267 and +285. As illustrated in Fig. 1, RNA1a (GGGCUGCGGGUGGGCGGGA, 735/1056) and RNA1b (GGGCAUGGGGUGGGCAGGA, 656/1025) were among the sequences most frequently observed to display great potential for G4 formation; therefore, these sequences were selected as targets for the following studies. 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.

G4 RNAs evidenced through gel electrophoresis and 1H nuclear magnetic resonance Native polyacrylamide gel electrophoresis (PAGE) was performed using fluorescently labeled RNAs (RNA1a-FAM and RNA1b-FAM in Table 1) to monitor structure compaction, as G4 RNA should migrate faster than ssRNA (26). As shown in Fig. 2A, the band corresponding to target RNA migrated significantly faster than its G4-mutated counterpart, indicating the formation of a stable unimolecular structure. Compared with RNA1b-FAM, RNA1a-FAM migrated significantly faster, indicating a more compact structure for RNA1a. As there are only two Gs located in the fourth tract of RNA1b, the structure of this RNA is expected to be less stable than that of RNA1a, which contains three stacked G-quartets. Table 1 Sequences of some oligomers used in our studies. View this table: Fig. 2 Synthetic HCV G-rich sequences form G4 RNAs. (A) Formation of compact G4 RNAs characterized on the basis of species that move more rapidly than the G4-mutated oligonucleotides. Lane 1, RNA1a–Mut-F; lane 2, RNA1a-FAM; lane 3, RNA1b-FAM; lane 4, RNA1b–Mut-F. (B) G4 structures 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 suppressed through AS-RNA1a (100 μM) 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. To further confirm the G4 structures, we conducted 1H nuclear magnetic resonance (NMR) analysis using chemically synthesized RNAs (RNA1a and RNA1b) (13, 27). Imino proton peaks within the 10.0- to 11.5-ppm region are highly characteristic of the Hoogsteen hydrogen bonds of G-quartets (28). Accordingly, the 1H NMR spectrum of RNA1a revealed well-resolved imino peaks within this region (red spectrum in Fig. 2B and fig. S6), indicating the formation of G4 structures. The 1H NMR spectrum of RNA1b demonstrated broad imino peaks within the same range (red spectrum in Fig. 2C and fig. S7), suggesting inhomogeneous G4 RNAs. Because there were only two G bases at the fourth G track, a two–G-tetrad motif was the most probable structure for RNA1b. Hence, RNA1b could potentially be involved in multiple alternative folding patterns (29). Furthermore, the effects of antisense oligonucleotides (ASOs), including AS-RNA1a and AS-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 (blue spectrum in Fig. 2B), suggesting the partial conversion of G4 into double-stranded RNAs (dsRNAs) (20). A similar phenomenon was also observed for RNA1b and AS-RNA1b (blue spectrum in Fig. 2C). In a long RNA structural context, the folding of a specific structure might be influenced by the neighboring sequences (30, 31). Mfold Web server is used for folding analysis of the primary sequence of the HCV C gene (32). The results indicated that RNA1a and RNA1b are located in a very structured (dsRNA) region (blue zone in figs. S8 and S9). 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 in the “ds region,” the NMR results showed that the short motifs, including RNA1a and RNA1b, were capable of forming G4 structures (red spectrum in fig. S10). Moreover, G-A mutations largely disrupted the G4 formation (blue spectrum in fig. S10).

Highly stable parallel G4 RNAs of the target HCV sequences To confirm the formation of G4 structures in the target HCV sequences, 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 RNA was confirmed through small variations of CD spectra observed at elevated temperatures (fig. S12A), whereas the spectra of RNA1b showed more variation within the same range (fig. S12B), indicating a less stable G4 RNA structure. Next, we measured the spectra of these sequences in the presence of NaCl or LiCl. As shown in fig. S13, the spectra of the sequences were similar to those recorded in KCl solution, indicating parallel G4s. Furthermore, CD spectroscopy of RNA1a revealed a minimum peak at 210 nm in the presence of AS-RNA1a (fig. S14A), which was highly reminiscent of the spectral signature associated with the A-form of dsRNA. Moreover, increasing the level of AS-RNA1a promoted a shift from 264 to 275 nm, indicating a structural switch alleviated from G4. When RNA1b was titrated with less AS-RNA1b, the minimum at 210 nm could be evidently observed (fig. S14B), confirming the increased G4 stability of RNA1a compared with RNA1b. To quantitatively determine the G4 stabilities of target RNA, we conducted a thermal CD study (34). Because the unfolding process for RNA1a in the presence of a physiological concentration of K+ (100 mM) could not be accomplished (fig. S15), thermal studies were initially conducted using subphysiological levels of K+. Figure S16A shows the thermal profiles of RNA1a in the presence of different salts. The melting temperature (T m ) for RNA1a was dependent on the K+ concentrations. The alkali metal ion dependence for the stabilization of folded RNA1a, on the basis of the T m , 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, the T m of RNA1b was lower than that of RNA1a (fig. S16B), suggesting a less stable structure. An evident multiphasic stage was characterized for RNA1b (fig. S16B), further supporting the notion of the inhomogeneous G4 conformations of this RNA. In addition, the T m for RNA1a or RNA1b was independent of the strand concentration, consistent with intramolecular G4 formation (fig. S17) (36). The melting curve analysis also revealed hysteresis in the cooling-versus-heating ramps (fig. S18), demonstrating slow folding kinetics for target RNAs (20).

Stabilization of G4 RNAs through G4 ligands Recently, a number of representative G4 DNA ligands, such as PDP (37) (structure in Fig. 3A) and TMPyP4 (38) (fig. S19), have been developed. As demonstrated in Fig. 3 (B and C) and fig. S20, the binding of G4 ligands to target RNA resulted in a significant temperature shift in the melting curve, indicating evident G4 RNA stabilization at physiological ionic strength. To further confirm that G4 ligands act specifically on HCV G4 RNAs, we incubated these molecules with the G4-mutated version of target RNAs. No significant temperature shift in the melting curve was observed (fig. S21). We showed that PDP increased the G4 stability of target RNA and therefore inhibited the opening of the G4 motif through ASO treatment, using a fluorescence resonance energy transfer (FRET) kinetic assay (figs. S22 and S23) (20). 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 μM) or RNA1b (8.0 μM) were recorded in 10 mM tri-HCl buffer (pH 7.0) (100 mM KCl), in the absence or presence of PDP (8.0 μM).

G4 ligands block RNA-dependent RNA synthesis Next, we examined whether ligand-mediated G4 stabilization can regulate RNA replication. In the HCV life cycle, the viral RNA is replicated 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 RdRp with a primer-dependent mechanism (42, 43). The HCV G-rich segments (red part in Fig. 4A) have been incorporated into the 5′ ends of RNA templates, and the 5′ FAM-labeled primer p15 (green part in Fig. 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 (left half in Fig. 4A). In contrast, if the binding of G4 ligands occurred, the extension would be stopped at the G4 site (right half in Fig. 4A). Fig. 4 G4 ligands inhibit RNA-dependent RNA synthesis and HCV C gene expression through G4 RNA stabilization. (A) Schematic representation of the RNA stop assay. The artificial RNA template containing HCV G-rich or G4-mutated sequences was used. (B) The extended RNAs of template 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 not denatured and moved much slower than the corresponding ssRNA. Lane 1, RNA markers (p15, m17, m19, m21, and m39-1a in table S4); lanes 2 and 3, 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 indicate the percentage of densitometry of the HCV Core protein relative to β-actin. DMSO, dimethyl sulfoxide; nt, nucleotides. As shown in lane 3 of Fig. 4B, fully extended products were observed along template 1a-G4, and few stopped products were observed (due to G4 formation) in the absence of G4 ligand. However, when increasing amounts of G4 ligand were incubated with template 1a-G4, the template-directed primer extension was gradually inhibited at the G4 site (lanes 4 to 7 in Fig. 4B). On the contrary, the site-specific termination event was not characterized for G4-mutated template upon the addition of G4 ligand (lanes 10 and 11 in Fig. 4B). The G4 site–specific blockade in RNA synthesis was also detected when template 1b-G4 was treated with G4 ligands (fig. S24). Consistent with the results of the melting studies, ligand PDP exhibited superior G4 stabilization and polymerization inhibition.

Inhibition of the full-length C gene expression through G4 RNA stabilization To investigate the effects of G4 RNA stabilization on the expression of a full-length HCV C gene, we performed Western blot analysis. Plasmid 24480 (pMO29) carrying the genotype 1b C gene (fig. S25) and the conserved backbone [pcDNA3.1(+)/pEV 204-Hind III] was used (44). As demonstrated in Fig. 4C, both PDP and TMPyP4 evidently inhibited the expression level of the HCV C gene. To better understand the mechanism behind the inhibition of HCV C gene expression through G4 ligands, we examined TMPyP2 (structure in fig. S19), a positional isomer of TMPyP4 with low affinity for G4 DNA (38). As demonstrated in fig. S26, TMPyP2 showed much less G4 RNA-stabilizing activity toward RNA1b, contained in pMO29. Comparisons of TMPyP2 (lane 4 in Fig. 4C) and TMPyP4 (lane 1 in Fig. 4C) on the basis of HCV C gene expression were consistent with the above results. To demonstrate that the observed effects are specific for G4 sequence, we prepared a relevant control plasmid with a G4-mutated sequence. The following results (fig. S27) indicate that both PDP and TMPyP4 did not inhibit the translation of the G4-mutated C gene. Together, these results suggest that because of the actions of G4 ligand in binding to and stabilizing G4 RNA in the indicated region of the C gene (fig. S25), it is a promising candidate for further study.

Repression of enhanced green fluorescent protein expression through G4 RNA stabilization Next, an enhanced green fluorescent protein (EGFP) reporter gene system was built to examine whether ligand-mediated G4 stabilization can regulate protein expression. We constructed a variety of pEGFP-C1 derivatives (45) after cloning the 21-bp sequences, containing either wild-type HCV G-rich sequences or the G-to-A mutant sequences. Plasmids GFP-1a core G4, GFP-1b core G4, GFP-1a core Mut, and GFP-1b core Mut were prepared, and the insert was placed immediately downstream of the human cytomegalovirus immediate early promoter (1 to 589) and upstream of the EGFP-cds present on the parental plasmid (figs. S28A and S29A). We next performed a confocal fluorescence assay. Upon addition of G4 ligand, the expression of EGFP in GFP-1a core G4 or GFP-1b core G4 was significantly inhibited compared with the DMSO treatment (fig. S28B), whereas the expression of EGFP in G4-mutated plasmids or the empty vector was not influenced under the same conditions (fig. S29B). Consistent with the previous results, TMPyP2 did not inhibit the expression of EGFP (fig. S28B). Together, the results suggest that G4 ligands inhibit reporter gene expression by targeting HCV G-rich RNAs. To quantitatively evaluate the effects of G4 ligands on EGFP expression, we performed flow cytometry, and the results showed that EGFP was expressed much less efficiently in GFP-1a core G4– or GFP-1b core G4–transfected cells following G4 ligand treatment compared with the DMSO control (fig. S28C). In contrast, G4 ligands had a much less pronounced effect on EGFP expression in G4-mutated plasmids. Consistent with previous results, PDP showed better inhibition of EGFP expression.

G4 ligands inhibit HCV replication in cells Next, we investigated whether G4 ligands can inhibit HCV infection. Currently, the most commonly used infectious HCV culture system is based on JFH1 (Japanese fulminant hepatitis 1, genotype 2a) (46), 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 and also identified the G4-forming sequence around the same region (fig. S30). A similar sequence alignment of the C gene for subtype 2a was further performed (table S6 and figs. S31 to S33), and a conserved G4 motif was also identified (fig. S34). The 1H NMR spectrum of RNA2a demonstrated 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 Hoogsteen imino proton resonances (blue spectrum in fig. S35A), and the G4 structure was stable (S35B). Hence, JFH1 is a suitable model for testing the anti-HCV strategy. Here, JFH1-infected Huh-7.5.1 cells were treated with different G4 ligands. Quantitative real-time polymerase chain reaction (RT-qPCR) was performed as previously described (48), and HCV RNA levels were determined relative to the transcription of glyceraldehyde-3-phosphate dehydrogenase (GAPDH) in host cells. Because interferon-α (IFN-α) is best known as an effective treatment for HCV infection, it is used as a positive control in our studies examining the anti-HCV effects of G4 ligands. As shown in fig. S36, viral RNA levels in JFH1-infected cells were markedly decreased by G4 ligands in a dose-dependent manner, with measured median inhibitory concentrations of 1.2 and 3.2 μM for PDP and TMPyP4, respectively. Next, we further evaluated the antiviral activity of G4 ligands against two different intergenotypic HCV chimeras. The H77/JFH1 chimeric genome contained the genotype 1a Con1 sequence from the C gene (sequence in fig. S37) to the NS2 region of H77 (genotype 1a) and the nonstructural region of JFH1 (49), and the Con1/JFH1 chimeric 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 the rest of the sequence from JFH1 (50, 51). The RT-qPCR results revealed that PDP and TMPyP4 display adequate activity against HCV H77/JFH1 or Con1/JFH1 infection and evidently inhibit the viral RNA levels at the C gene (figs. S39A and Fig. 5A). Not surprisingly, both G4 ligands also effectively inhibit the viral RNA levels at 5′UTR in living cells (figs. S39B and Fig. 5B). 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-α). IFN-α was used at 150 ng/ml. The values observed were normalized to GAPDH. All data are presented as the means ± SEM from three independent experiments. The error bars reflect the SD. G4 ligand 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, and the 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 to β-actin. (D) Western blot analysis was performed, and a commercial anti–HCV nonstructural protein 3 (NS3) antibody was used for detection. Moreover, Western blot analysis was performed to determine the Core protein levels of H77/JFH1- or Con1/JFH1-infected Huh-7.5.1 cells using the commercial anti–HCV Core antibody (1a or 1b) (52). As shown in Fig. 5C and fig. S40, the level of HCV Core protein expression in cultured cells was significantly decreased by G4 ligands in a dose-dependent manner. The nearly complete absence of the HCV Core protein was observed in infected cells treated with 2.5 μM PDP, indicating total viral inhibition. Consistent with these results, TMPyP4 treatment significantly reduced the level of the HCV Core protein to 10 μM, whereas a higher concentration of TMPyP2 did not inhibit HCV infection. Together, these results suggest that PDP is a more effective agent for the inhibition of HCV Core protein expression. Next, we also investigated the antiviral effects of G4 ligands on JFH1 virus. The NS3 acts as a serine protease/helicase, which is an important component in the HCV replication complex. Therefore, the amount of NS3 is a good indicator of the level of HCV activity in cells (53) and was selected as a target in this assay. As demonstrated in Fig. 5D, G4 ligands significantly inhibited the level of JFH1 virus and decreased the expression of HCV NS3 protein in infected cells, and total viral inhibition could be achieved through 2.5 μM PDP treatment. These results confirm that PDP is a better antiviral agent against genotype 2a HCV. During HCV replication, the positive-sense RNA genome is used as a template to produce a negative-strand RNA intermediate (HCV− RNA) (39). To further demonstrate that targeting G4 inhibits HCV replication in vivo, we evaluated the effects of G4 ligands on the level of HCV− RNA using Tth-based RT-qPCR (54) in JFH1-infected Huh-7.5.1 cells. Our results clearly showed that G4 ligands significantly inhibit HCV− RNA levels in living cells in a dose-dependent manner (fig. S41).

G4 mutations in the HCV C gene disrupt G4 ligand–virus interactions To further demonstrate the existence of G4 RNAs in the HCV genome under physiological conditions, we used a pull-down strategy (37, 55). Because the introduction of a biotin affinity tag on PDP has 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 impair the stabilization of HCV G4 RNAs. Xba I restriction digestion of the plasmid pJ6/JFH1 at the 3′ end of the HCV complementary DNA (cDNA) was performed (56), and the linearized plasmid was transcribed in vitro to generate full-length HCV genomic RNA. Moreover, a pJ6/JFH1–G4-Mut plasmid containing a G4-mutated sequence in the C gene was prepared using overlapping extension PCR (OE-PCR). The transcribed J6/JFH1 or J6/JFH1–G4-Mut RNA was incubated in the presence or absence of biotin-PDP and sonicated to shear the genomic RNA. Subsequently, hydrophilic streptavidin-coated magnetic beads were used to capture the desired G4 RNAs (scheme in fig. S44). As shown in fig. S45, an evident peak at approximately 264 nm, representing parallel G4 RNAs, was observed for recovered wild-type viral RNA (green line), whereas almost no signals were observed for G4-mutated viral RNA (blue line) and the control sample without using biotin-PDP (black line). We next performed RT-qPCR to determine the abundance of the HCV G4 motif in RNA samples before and after pull-down manipulation (57), using G4-fwd and G4-rev (sequences in table S4). The normalized results revealed an approximately 28-fold enrichment (ΔΔC T = 4.82), whereas the control assay that did not use biotin-PDP did not display 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 cells were infected and treated with various G4 ligands, and the replicated viral RNA levels were determined relative to the host cell GAPDH mRNA. As demonstrated in Fig. 6A, much more attenuated inhibitions were observed for the J6/JFH1–G4-Mut virus. Western blot analysis further indicated that G4 ligands strongly inhibited Core protein expression of the wild-type, but not of the G4-mutated, J6/JFH1 (Fig. 6B). These results provided direct and solid evidence that G4 RNA in the HCV C gene represents a cellular target for typical G4 ligands such as PDP. Fig. 6 G4-disruptive mutations in the HCV C gene inhibit G4 ligand–virus interactions. (A) RT-qPCR was performed. The primers were designed to target 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 β-actin. Lane 1, no HCV control; lanes 2 to 7, J6/JFH1–G4-Mut virus; lanes 8 to 13, J6/JFH1 virus. To further confirm whether the G4 motif is a molecular target of G4 ligands, we evaluated the activity of these molecules toward a different natural virus without a G4-forming sequence. Influenza A virus, a negative-sense ssRNA virus, was examined (58). As expected, G4 ligands were much less effective on this virus strain (fig. S46). Previous investigations demonstrated that ASOs can bind to G4 RNA and affect specific mRNA translation (20, 59). Here, ASOs complementary to G4 were delivered into different HCV-infected cells. The results clearly demonstrated that such ASOs can destabilize the HCV G4s and show an opposite effect on RNA replication when using G4 ligands (fig. S47). As expected, the stimulatory effect on viral replication was significantly alleviated using mutant ASOs. The effect observed for G4-mutated virus was nearly abolished in the presence of ASO targeting the same nucleotide position.