Design of Synthetic shRNAs for Targeting Hepatitis C: A New Approach to Antiviral Therapeutics

  • Brian H. JohnstonEmail author
  • Qing Ge
Part of the RNA Technologies book series (RNATECHN)


Small hairpin RNAs (shRNAs) are widely used as gene silencing tools and typically consist of a duplex stem of 19–29 bp, a loop, and often a dinucleotide overhang at the 3′ end. Like siRNAs, shRNAs show promise as potential therapeutic agents due to their high level of specificity and potency, although effective delivery to target tissues remains a challenge. Algorithms used to predict siRNA performance are frequently used to design shRNAs as well. However, the differences between these two kinds of RNAi mediators indicate that the factors affecting target gene silencing will not be the same for siRNAs and shRNAs. Stem and loop lengths, structures of the termini, the identity of nucleotides adjacent to and near the loop, and the position of the guide (antisense) strand all affect the efficacy of shRNAs. In addition, shRNAs with 19-bp or shorter stem lengths are processed and function differently than those with longer stems. In this review, we describe studies of targeting the hepatitis C virus that have provided guidelines for an optimal design for short (19 bp) shRNAs (sshRNAs) that are highly potent, stable in biological fluids, and have minimal immunostimulatory properties.


Antivirals Hepatitis C RNAi shRNAs sshRNAs 

1 Introduction

While antibiotics have greatly reduced the mortality and morbidity associated with bacterial infections, the development of effective antivirals has proceeded much more slowly. Hepatitis C virus (HCV) infections remain a worldwide health problem, and no vaccine is currently available. While a number of small molecule inhibitors of virally encoded enzymes are showing promise in clinical trials and some have now been approved by the US FDA, cures of HCV generally require multiple agents with different mechanisms of action due to the development of viral resistance. Such viral escape results from the high error rate of the viral RNA-dependent RNA polymerase, which allows rapid exploration of sequence space, creating myriad quasispecies that may include mutations conferring resistance to any given drug. Small interfering RNAs (siRNAs) and their cousins small hairpin RNAs (shRNAs) have two advantages over small molecule drugs that suggest a potential solution to the viral resistance problem: first, they can potentially target conserved but “undruggable” sequences anywhere in the viral genome, including noncoding sequences, and second, multiple small siRNAs or shRNAs can be combined to foil the ability of the virus to escape through single-site mutations. Chronic HCV infection, which represents the most common form of this disease, is also attractive as a disease target for RNAi because the course of the infection is slow. Thus, there is plenty of time for the biology of RNA interference to take place, in contrast to some very rapid viral infections in which viral replication may outrun treatments that require some hours to take full effect.

In this review, we describe the development of a new class of RNA interference (RNAi) effectors, short shRNAs, or sshRNAs, as HCV drugs. sshRNAs have been designed to target the conserved internal ribosome entry site (IRES) element of the HCV genome, one of the most highly conserved regions of the viral genome. Their short length (around 40 nt) distinguishes them from more traditional shRNAs in both potency and mechanism of action.

2 Hepatitis C Virus

HCV, a single-stranded, positive-sense RNA virus, is the most common blood-borne RNA virus, with an estimated 4.1 million persons currently infected in the United States and 180 million worldwide. A total of 3–4 million people are newly infected each year (Armstrong et al. 2000; Alter et al. 1999; Shepard et al. 2005; Ray Kim 2002). Approximately 250,000 human immunodeficiency virus (HIV)-infected persons in the United States are coinfected with HCV ( HCV is recognized as a major cause of end-stage liver disease, such as liver cancer (with 1–5% of HCV cases leading to this outcome) and cirrhosis (10–20% of cases), and is the leading indication for liver transplantation in the Western world (Shepard et al. 2005). Chronic liver disease ranks as the tenth leading cause of death in the United States, and HCV is estimated to account for 40–60% of these cases. Mortality related to HCV infection (death from liver failure or hepatocellular carcinoma) is expected to increase over the next two decades (Deuffic-Burban et al. 2007).

There are six major genotypes of HCV. The standard treatment is a combination of pegylated interferon (IFN)-α and ribavirin, which is effective in 40–80% of patients, depending on the genotype. Efficacy has been around 50% in patients infected with HCV genotype 1, which comprises nearly 70% of cases in the western hemisphere, although the success rate is improving with the addition of newly approved protease inhibitors to the standard treatment. The response rate for HIV–HCV coinfected patients is lower, estimated at 30–40% (Torriani et al. 2004). Because interferon is associated with severe adverse effects, including flu-like symptoms, hematologic abnormalities, and depression, patient compliance has been poor with an estimated 30% of patients refusing treatment. Considering that the incidence of HCV is increasing worldwide and that within 10 years more deaths from HCV than HIV are predicted by the Centers for Disease Control, safer and more efficacious HCV drugs are urgently needed (Deuffic-Burban et al. 2004, 2006; Law et al. 2003; Salomon et al. 2002).

More than 40 HCV drug candidates are under development. They can be divided into those that target the virus directly [direct-acting antiviral (DAA) agents] and those that affect host targets (Ronn and Sandstrom 2008; Liu-Young and Kozal 2008; Beaulieu 2007). DAAs include small molecules as well as ribozymes, antisense oligonucleotides (ASOs), decoy RNAs, RNA aptamers, siRNAs, and shRNAs. Host-targeting agents include small molecules such as cyclophilin inhibitors, antifibrotic agents, antibodies, modified interferons with improved pharmacokinetics, and oligonucleotides complementary to the required host factor microRNA-122 (miR-122) (Lanford et al. 2010). As mentioned above, the high genetic diversity and rapid mutation and turnover rates of HCV (1010–1012 new particles produced per day with an error rate of 10−3–10−5 mutations per nucleotide per genomic replication) result in the rapid emergence of viral resistance with many single DAAs (Okamoto et al. 1992; Neumann et al. 1998; Ogata et al. 1991; Sarrazin and Zeuzem 2010). For instance, although clinical studies showed that the use of protease inhibitors in combination with pegylated interferon-α (peg-IFN) and ribavirin increased the rate of sustained viral response (SVR) by at least 20% compared with peg-IFN and ribavirin alone in patients infected with HCV genotype 1, the existence of viral variants with reduced susceptibility has been observed (Deuffic-Burban et al. 2004, 2009, 2012; Sarrazin and Zeuzem 2010; Robinson et al. 2011).

Of the oligonucleotide-based DAAs, siRNAs and shRNA generally have the best efficacy and potency in vitro as well as in animals (Ronn and Sandstrom 2008; McHutchison et al. 2006). In addition, the ability of RNAi to efficiently limit viral replication, and to target multiple genes and/or sequences simultaneously, makes this an attractive therapeutic approach for limiting the emergence of resistant mutants. Both siRNAs and vector-expressed shRNAs have been shown to significantly decrease HCV RNA replication and protein expression in cell culture as well as in animal systems (Watanabe et al. 2007; Randall and Rice 2004; Kapadia et al. 2003; Wilson et al. 2003; Yokota et al. 2003; Seo et al. 2003; Takigawa et al. 2004; Prabhu et al. 2005; Sen et al. 2003; Kanda et al. 2007). More recently, synthetic shRNAs have gained attention, as discussed below.

3 RNA Interference

RNA interference (RNAi) plays a central role in the regulation of eukaryotic gene expressions associated with various biological processes ranging from development to cell homeostasis. Diseases, particularly cancers, are often associated with the dysregulation of particular miRNAs (Shenouda and Alahari 2009).

The mechanism of RNAi is complex and can involve alternative pathways (Filipowicz 2005; Siomi and Siomi 2009). Double-stranded RNA (dsRNA) molecules are recognized and processed by one or more RNase III-family enzymes (Hammond et al. 2000; Zamore et al. 2000; Bernstein et al. 2001). Primary transcripts encoding microRNAs (pri-miRs) are processed in the nucleus by a “microprocessor” complex to individual hairpins before export to the cytoplasm. In the cytoplasm, dsRNAs longer than ~23 bp, including hairpins, are processed by Dicer into double-stranded siRNAs whose strands are each 21–23 nucleotides (nt) in length. Dicer contains a PAZ domain that binds specifically to the 3′ end of single-stranded RNA and two RNase III domains that possess the catalytic cleavage activity. The distance between the 3′-overhang-binding PAZ domain and the active site of the RNase III domains provides a molecular ruler corresponding to the length of an siRNA duplex (Jinek and Doudna 2009). The cleavage products of Dicer have characteristic termini, a monophosphate group at the 5′ end, and a two-nucleotide overhang with a 3′-hydroxyl at the 3′ end (MacRae and Doudna 2007). sRNAs, whether generated by dicing of a precursor RNA or introduced from outside the cell, are loaded into an RNA-induced silencing complex (RISC) containing a protein of the Argonaute (Ago) family (Elbashir et al. 2001; Lee et al. 2004; Pham et al. 2004; Tomari et al. 2004a, b; Jinek and Doudna 2009). Ago proteins contain PAZ, middle (MID), and PIWI domains. The binding of an siRNA to an Ago is aided by the presence of a 5′ phosphate group and a 3′ dinucleotide overhang at the termini. As with Dicer, the 3′ overhang binds to the PAZ domain of Ago. The 5′ phosphate group binds in a pocket at the interface between the MID domain and the PIWI domain (Jinek and Doudna 2009).

Of the four human Agos, Ago2 is thought to be the only one able to mediate cleavage of a target mRNA (Hammond et al. 2001; Okamura et al. 2004; Meister et al. 2004; Rand et al. 2004; Liu et al. 2004; Song et al. 2004; Rivas et al. 2005). To render the “guide” (antisense) strand of an siRNA available to pair with its target, the passenger strand must be removed. The selection of which strand is to be removed and which retained is thought to be largely governed by asymmetry in the thermodynamic profile of the siRNA duplex termini (Schwarz et al. 2003; Khvorova et al. 2003). After the RISC-loading complex (which contains the dsRNA binding protein TRBP along with Dicer and Ago2) loads an siRNA duplex into Ago2, Ago2 cleaves (or “slices”) the “passenger” (sense) strand at a position opposite 10 nt from the 5′-phosphate of the guide strand, facilitating the dissociation and/or degradation by C3PO (Ye et al. 2011) of the resulting passenger-strand fragments. The loss of the passenger strand from the complex produces the active RISC. The anchoring of the 5′ phosphate group into the binding pocket of Ago is essential, and the distance from it determines the position of the cleavage site in the passenger strand (this Ago2-mediated passenger-strand cleavage was also found in the processing of some microRNAs [miRNAs] (Diederichs and Haber 2007).

When passenger-strand slicing is blocked by chemical modification or by mismatches between the two strands of the siRNA, a slower, alternative pathway dissociates and destroys the passenger strand, possibly via an ATP-dependent helicase (RNA helicase A), yielding the active RISC (Matranga et al. 2005; Leuschner et al. 2006; Miyoshi et al. 2005; Kraynack and Baker 2006; Robb and Rana 2007; Rand et al. 2005). RISC uses the bound single-stranded RNA molecule as a guide to “search” the resident population of messenger RNAs (mRNAs) for complementary sequences, eventually cleaving these transcripts and thereby downregulating the expression of the targeted gene (Hammond et al. 2000; Dorsett and Tuschl 2004; Kim and Rossi 2008; Elbashir et al. 2001; Nykanen et al. 2001; Martinez et al. 2002). The currently favored model for target recognition and cleavage by Ago2 is as follows: The target binds to the seed region of the 5′ half of the guide sequence and then base pairing proceeds toward its 3′ end. This results in the 3′ end of the guide strand dissociating from the PAZ domain, leading to a conformational change that positions the active site of Ago2 at the cleavage site on the target (Filipowicz 2005; Tomari and Zamore 2005). The position of the scissile phosphate group of the target mRNA is similar to that in the passenger strand, i.e., 10 nt from the 5′-phosphate group of the guide strand. Mismatches at the 10th and 11th nucleotides prevent the slicing activity (Jinek and Doudna 2009).

The endogenous RNAi machinery has been exploited to advance a wide range of studies involving gene function analysis, pathway mapping, drug target validation, and host–pathogen interactions (Natt 2007; Dorsett and Tuschl 2004; Iorns et al. 2007). New understanding of how RNAi regulates gene expression is also leading to the rapid development of RNAi-based therapeutics, especially in the area of viral disease. For example, RNAi approaches targeting viral genes, including those of the HIV, influenza, and hepatitis A, B, and C viruses (Table 1), respiratory syncytial virus, polio virus, the SARS coronavirus, alphaviruses, and the Marburg and dengue fever viruses, have been shown to limit viral replication in cell culture and, in some cases, in animals (Rossi et al. 2007; Barik and Bitko 2006; Watanabe et al. 2007; Arbuthnot et al. 2007; Seyhan et al. 2007; Gitlin et al. 2002; Fowler et al. 2005; Haasnoot et al. 2003). A number of siRNA drug candidates are in clinical trials (Castanotto and Rossi 2009). Moreover, mechanistic studies of dsRNA/siRNA/miRNA processing by Dicer and Ago2 have also led to the development of algorithms for efficient target selection; chemical modification for improved specificity, functionality, and longevity; and new designs for RNAi triggers such as Dicer substrate, ssiRNA, etc., that benefit both basic and applied research (Bolcato-Bellemin et al. 2007; Collingwood et al. 2008).
Table 1

Hepatitis C-specific siRNAs and shRNAs

Target region in HCV


Biological effect



Hydrodynamic injection of si/shRNA and the reporter plasmid of NS5B-luciferase fusion

Reduced luciferase expression in mouse liver by 75% (siRNA) to 98% (expressed shRNA)

McCaffrey et al. (2002)


Selectable subgenomic HCV replicon cell culture

HCV RNA replication and protein expression were inhibited more than 30-fold

Kapadia et al. (2003)


HCV replicon cell culture

80-fold decrease in HCV RNA

Randall et al. (2003)

NS3-1, NS5B

HCV replicon cell culture

Effectively suppressed replication of the HCV replicon without suppressing host gene expression

Takigawa et al. (2004)


HCV subgenomic replicon cell culture

HCV RNA synthesis reduced by 90%

Wilson et al. (2003), Wilson and Richardson (2005)

E2, NS3, NS5B

Transient HCV1a replication model

Effectively suppressed HCV RNA replication and protein expression

Prabhu et al. (2005)

Core, E2

EGFP reporter in cell culture

Suppressed EGFP expression

Liu et al. (2006)

Core, NS3, NS4A, NS4B

Genomic HCV replicon cell culture and hydrodynamic injection mouse model

Effectively suppressed viral replication in a dose-dependent manner

Kim et al. (2006), Shin et al. (2009)


Cell culture-grown HCV genotype 1a

Effectively inhibited NS5A and core protein expression

Sen et al. (2003)

Stem–loop II of 5′ UTR

HCV subgenomic and full-length infectious replicon cell culture

Suppressed GFP expression and IRES mRNA in the case of six different HCV genotypes

Prabhu t al. (2006)

5′ UTR

HCV IRES-reporter and HCV subgenomic replicon cell culture

~80% suppression of HCV replication with concentrations of siRNA as low as 2.5 nM

Yokota et al. (2003)

5′ UTR

HCV subgenomic replicon with the luciferase gene

Suppressed the luciferase reporter expression

Seo et al. (2003)

5′ UTR

Cell culture-grown HCV genotype 2a, 1a, and 1b replicon system

Inhibited viral genome replication and infectivity titers

Kanda et al. (2007)

5′ UTR

HCV serum infected Huh-7 cells that supports genotype-4 replication

Suppressed HCV RNA by ~25-fold

Zekri et al. (2009)

5′ UTR

Cell culture-grown HCV and replicon system

Suppressed HCV replications

Ray and Kanda (2009)

5′ UTR

HCV replicon cell culture

Additive HCV inhibitory effects for combinations of ribozymes and siRNAs

Jarczak et al. (2005)

5′ UTR

HCV subgenomic replicon, HCV-luciferase reporter in cell culture

Effectively suppressed RNA replication in replicon; up to 98% knockdown of HCV-luciferase reporter

Ge et al. (2010a, b), Ilves et al. (2006), Vlassov et al. (2007)

5′ UTR

Hydrodynamic injection of HCV-luciferase reporter with shRNA in mice

Up to 99% knockdown of HCV-luciferase reporter in mouse liver

Wang et al. (2005)

5′ UTR

HCV-luciferase reporter expressed in mouse liver with lipid nanoparticle formulation

90% suppression of IRES-luciferase expression in mouse liver

Dallas, Ma et al. (submitted)

5′ UTR

HCV-infected chimeric uPA/SCID mice, lipid nanoparticle formulation

2.5 log10 viral load reduction

Dallas, Ma et al. (submitted)

5′ UTR NS3, NS4A, NS4B, NS5B

HCV subgenomic replicon cell culture

Suppressed HCV RNA and NS5B protein levels up to 75% with single siRNA and 90% with siRNA combinations

Korf et al. (2007)

4 Structure–Activity Relationships of shRNAs

RNAi effectors can be generated by chemical synthesis of siRNAs and sshRNAs, Dicer cleavage of longer synthetic dsRNAs and shRNAs, or processing of shRNAs and long dsRNAs transcribed from DNA or viral vectors (Dorsett and Tuschl 2004; Chang et al. 2006; Bernards et al. 2006; Fewell and Schmitt 2006; Vlassov et al. 2006; Amarzguioui et al. 2006; Ge et al. 2010a, b). The transcription of shRNAs or dsRNAs can be driven by Pol II promoters or alternatively by Pol III promoters such as the H1 promoter of RNase P or the U6 snRNA promoter. shRNA has received considerable attention due to its widespread use in DNA vector-based shRNA libraries for various loss-of-function screens, generation of cell lines or transgenic animals that express silencing triggers against targets of interest, and therapeutic approaches (Grimm et al. 2006; Li et al. 2005a, b).

4.1 Parameters Involved

An shRNA consists of largely paired antisense and sense sequences connected by a loop of unpaired nucleotides. A duplex stem of typically 24–29 bp, either fully paired or with miRNA-style internal mismatches or loops, is commonly used in vector-expressed shRNAs (Silva et al. 2005; Stegmeier et al. 2005; Boudreau et al. 2008). Although target site selection is critical to silencing activity, the structural design of the shRNA also plays a significant role. With appropriate design, shRNAs can be at least as active as siRNAs targeting the same site. Although the structure–activity relationship of siRNAs has been extensively examined and a 19-bp RNA duplex with 2-nucleotide overhangs at the 3′ ends of each strand is widely used, there have been only a handful of studies on the effect of hairpin structures on efficacy of target silencing. The factors involved in shRNA design are loop size and sequence, stem length, presence of internal mismatches or single-stranded overhangs, and whether it is expressed or synthetic and directly delivered. As with siRNAs, stem length determines whether the molecule is a substrate for Dicer. In the context of vector expression, fully matched shRNAs have been compared with shRNAs containing internal mismatches from sequence alterations in the passenger arm, and the fully matched shRNAs were found to be more potent (Li et al. 2007; Boudreau et al. 2008). The loop can be almost any size, from 2 to 10 or more nucleotides. Brummelkamp et al. found that an expressed 19-bp shRNA with a 9-nt loop provided better target knockdown than similar shRNAs with 5- or 7-nt loops (Brummelkamp et al. 2002). The efficacy seen with expressed shRNAs depends on expression level as well as design parameters (Hinton et al. 2008; Kawasaki et al. 2003; Zhou et al. 2009). To investigate design parameters alone, several groups have examined the impact of structural changes to synthetic hairpins and found that the lengths of both stems and loops can affect efficacy. Li et al. reported that in the context of 4-nt loops, shRNAs with 29-bp stems silenced target gene expression more efficiently than those with 19-bp stems, but 19-bp shRNAs with 9-nt loops outperformed shRNAs with longer stems, including 29-bp shRNAs with 4-nt loops (Li et al. 2007). Our group found that, in the context of 10-nt loops, 19-bp shRNAs were somewhat more potent than similar 19-bp and 25-bp siRNAs and 25-bp shRNAs were less potent than any of the 19-bp shRNAs or siRNAs tested (Vlassov et al. 2007).

4.2 Loop Position

The position of the antisense sequence within the hairpin also affects shRNA efficacy. shRNAs have often been designed with the sense sequence at the 5′ end of the hairpin (right-hand loop, R-type shRNAs) (Vlassov et al. 2007; Li et al. 2007) (Fig. 1). Harborth et al. reported that an shRNA with its antisense sequence at the 5′ end of the hairpin (left-hand loop, L-type shRNA) showed comparable silencing efficacy to an R shRNA if the stem length was 21–29 nt (Harborth et al. 2003). However, when the stem length was shortened to 19 bp (with a 4-nt loop), much less potency was found with R shRNA, whereas L shRNA retained a potency comparable to that of shRNAs with 21–29-bp stems. Similar results were obtained by another group with a CD8-specific shRNA (McManus et al. 2002). Together, these results suggested that 19-bp shRNAs may be processed differently from shRNAs of 24 bp or longer. A major difference is that, of the longer shRNAs, both R and L types are processed by Dicer to generate the same ~19-bp siRNAs. In contrast, 19-bp shRNAs were found not to be Dicer substrates (Siolas et al. 2005; McManus et al. 2002). To distinguish shRNAs with 19 or fewer base pairs from longer, Dicer substrate shRNAs, we have designated the former as short shRNAs or sshRNAs.
Fig. 1

Structure and activity of R- and L-sshRNAs. (a) General structures of R- and L-sshRNAs. Antisense and sense are relative to target sequence. The antisense strand normally becomes the guide strand in RISC. (b) Representative structures of L-sshRNAs, with sense and antisense strands of equal or unequal length. Adapted from Ge et al. (2010b)

4.3 Duplex Length Effects on Activity of sshRNAs

Shortening the sense sequence of a synthetic 19-bp L-sshRNA from its 3′ end to 17 or 16 nt while maintaining the length of the antisense arm at 19 nt significantly reduced gene silencing activity, suggesting that having duplex structure at the 5′ end of the antisense sequence is important(Fig. 2) (Ge et al. 2010b). However, the overall length of the duplex can be shorter than 19 bp. sshRNAs having 17- or 18-bp stems can be virtually as potent as similar 19-bp versions (Fig. 2). Further shortening of the stem to 16 nt in each strand (connected by UU, which might form a 15-bp stem with a GUUC loop) resulted in somewhat lower activity, and an sshRNA with 15 nt in each arm of the duplex had very little activity. This indicates that potent silencing activity requires a hairpin with a duplex length of at least 16 bp. Interestingly, an sshRNA with a stem of 16 bp consisting of a 19-nt antisense sequence connected directly to a 17-nt sense sequence showed similar potency compared to its parent molecule with 19-bp stem and UU linker.
Fig. 2

Comparison of the activities of L-sshRNAs differing in 3′ overhang and stem length. (a) Overhang comparison. L-sshRNAs with and without a 3′ overhang as well as an siRNA, si19-3, all targeting the same sequence were chemically synthesized, and their ability to inhibit HCV IRES-dependent luciferase expression was compared in 293FT cells. (b) Stem length comparison. L-sshRNAs (UU loop) against the same target sequence but with different stem lengths were compared for their inhibitory activity in 293FT cells. SG105 differs from SG142 in lacking a 3′ UU overhang. (c) Effect of using part of the antisense sequence as the loop. SG119 has a 19-nt antisense sequence directly linked with a 17-nt sense sequence, probably forming a 16-bp duplex and 4-nt loop (UGCA). (d) Stem length comparison with SG119 derivatives. SG131 and SG132 have UU to connect the 3′ end of the antisense and 5′ end of the sense strands. They were compared with SG119 for target knockdown in 293FT cells. si131 has two complementary strands 16 nt in length with UU overhangs at their 3′ ends. as/s values represent the nucleotide lengths of antisense and sense strands. Adapted from Ge et al. (2010b)

4.4 Mechanistic Differences of L- and R-sshRNAs

To further examine the structure–function relationships of sshRNAs and how they may differ from Dicer-substrate shRNAs, our group undertook an extensive structure–activity and mechanistic study of sshRNAs targeting three partially overlapping sequences within the internal ribosome entry site (IRES) of HCV (Ge et al. 2010a, b; Dallas et al. 2012). For two of these target sites, L-type sshRNAs showed significantly higher potency than R-type; for the third, there was little difference between the two hairpin types. Unlike R-sshRNAs, where loop sizes of 5 nt or greater were optimal (Vlassov et al. 2007; Li et al. 2007), the L-sshRNAs were more potent when the loop size was very small (1 or 2 nt) than when it was larger (5 or 10 nt) (Fig. 3). The IC50 of an L-sshRNA of 19 bp and a UU loop was slightly more potent than a corresponding siRNA targeting the same region. The loop sequence appeared not to affect the sshRNA activity since three different loop sequences, including one derived from the microRNA miR-23, gave the same results. However, nucleotides adjacent to the loop may affect the activity as they can affect the actual size of the loop. For example, a CG base pair next to a UU loop can be paired and result in an actual 2-nt loop (Jucker and Pardi 1995), whereas if that base pair is AU or UA, strain in a 2-nt loop may keep them unpaired, resulting in a 4-nt loop of AUUU or UUUA. Based on studies involving chemical modification, conditional dicer-knockout cells, and Ago immunoprecipitation, we concluded (Dallas et al. 2012) that L-sshRNAs having UU loops [which are naturally quite resistant to RNase cleavage (Ge et al. 2010a)] require passenger-arm cleavage to be maximally active, but the loops remain intact in the active RISC, with the guide strand breaking its base pairs with the truncated passenger strand to allow pairing with the target. Passenger-strand “slicing” is needed apparently to facilitate opening of the guide-passenger duplex. L-sshRNAs have their guide sequence at the 5′ side of the loop, so the 5′ end of the guide sequence is immediately available for phosphorylation and binding to the MID pocket of Ago2. R-sshRNAs, on the other hand, require a larger loop for full activity because the loop must be cleaved by some nuclease to create a terminal phosphate at the 5′ end of the guide strand (see Fig. 1). In this case, passenger-arm slicing is less important because the cleavage of the loop itself facilitates removal of the passenger strand in active RISC.
Fig. 3

Comparison of L-sshRNA activity with various loop structures and base pairs adjacent to the loop. (a) Loop length comparison, using L-sshRNAs against the same target region as sh68 and si19-3 but with various lengths and sequences of loops. sshRNAs were chemically synthesized, and their abilities to inhibit HCV IRES-dependent luciferase expression were compared in 293FT cells. (b) Loop-adjacent base pair comparison. sshRNAs against the same target region as si72 but with loops of 5 nt (SG72 and SG72L) and 2 nt (SG118 and SG103) and left- (SG72L and SG118) and right-loop (SG72 and SG103) orientation were compared for their inhibitory activity in 293FT cells. Loop sequences are underlined. Adapted from Ge et al. (2010b)

Whereas the presence of short flanking sequences such as a 3′-overhang generally enhance the efficiency of gene knockdown for R shRNAs (Siolas et al. 2005; Vlassov et al. 2007), the presence or absence of a 3′ overhang has relatively little effect on the silencing ability of L-sshRNAs (Fig. 2a). This appears to be because the loop is generally not cleaved in L-sshRNAs, and when the stem opens up, the loop is available to bind to the PAZ domain of Ago2 (Dallas et al. 2012).
Fig. 4

Effect of 2′-O-Me modifications in the stem and/or loop regions on activities of L-sshRNAs. Activity was determined by potency in suppressing the expression of an HCV IRES-fLuc reporter in 293FT cells, by cotransfecting them in triplicate with the reporter DNA. An unmodified siRNA specific for the same target, si19-3, was used as a positive control. (ac) Comparison of an unmodified sshRNA (SG105) and its derivatives containing 2′-O-Me modifications in the stem and/or loop. (d) Comparison of SG119 and its derivatives containing 2′-O-Me modifications in the stem and loop. Each construct is identified by sequence number and a shorthand notation for the region modified, e.g., as-2 means 2′-O-methylated at position 2 of the antisense strand, ss-alt means 2′-O-methylated at every other nucleotide of the sense strand except for the slicer site. Adapted from Ge et al. (2010a)

5 Monomeric Versus Oligomeric shRNAs

Synthetic shRNAs are usually simply dissolved in H2O or a buffer prior to in vitro or in vivo use. Whether these shRNAs form the expected hairpin structures in solution is frequently ignored. We have found that synthetic shRNAs, irrespective of stem length, loop size, or L vs. R loop orientation, can form dimers or even oligomers (Ge et al. 2010b). Dimerization of hairpin RNAs has been also documented in retroviral RNAs, tRNAs, and some artificial RNA hairpins (Sun et al. 2007). The propensity of hairpin RNAs to dimerize depends on their loop size, sequence, and concentration as well as how they are handled (Bernacchi et al. 2005; Liu et al. 2005). When shRNAs are heated to 95°C and quickly cooled in an ice bath (snap cooling), dimerization can be eliminated (Ge et al. 2010a). Interestingly, sshRNAs showed efficient target knockdown both before and after the heating/snap cooling procedure, even for some sshRNAs that were predominantly dimers before the treatment, suggesting that the dimers of certain sshRNAs are functional molecules that can be processed and utilized by the RNAi machinery with similar efficiency as the monomers. However, the presence of long duplex regions in dimers can provoke immune stimulation. The heating/cooling procedure greatly reduces this immune stimulation (Table 2).
Table 2

Heating and snap cooling reduces immune stimulation by unmodified sshRNAs

Heating/snap cooling






47.5 ± 0.4

5.0 ± 0.6

6,192 ± 2,422

319 ± 81


23.1 ± 2.7

4.6 ± 0.7

2,552 ± 73

195 ± 37


7.3 ± 1.8

2.6 ± 0.6

5.9 ± 2.8

5.4 ± 2.1

Three different unmodified sshRNAs (100 nM), either untreated or subjected to 95°C heating (4 min) and snap cooling, were transfected into MRC5 cells in triplicate. RNA was extracted from cells 24 h post-transfection, and IFN-β and TNF-α mRNAs were quantified by RT-PCR. The mean values and standard errors of the relative RNA levels (fold differences) of cytokine genes were calculated and normalized to levels of GAPDH. Adapted from Ge et al. (2010b).

6 Chemical Modification to Improve Pharmacological Properties

Like all RNAs, shRNAs are nonideal as drugs due to their susceptibility to degradation by nucleases and the tendency of some sequences and structural features to cause unwanted immune stimulation. In a study of the effects of 2′-O-methyl, 2′-deoxy (DNA), and phosphorothioate modifications at various positions in the stem, loop, and overhangs, we found that placing a 2′-O-Me on each nucleotide of the loop and alternate nucleotides of the passenger arm, but leaving an unmodified window of 4 nt at the slicer site, provided significantly greater stability in 10% human serum (Fig. 5) while abrogating induction of the innate immune system (Ge et al. 2010a). It can be seen from Table 3 that two sshRNAs of the same structure but different sequences can have very different immunostimulatory properties. Blunt sshRNAs are particularly efficient at inducing RIG-I. In each case, however, 2′-O-methylation renders them non-stimulatory, by blocking recognition by pattern recognition sensors such as RIG-I. These modifications had essentially no effect on potency, but placing modifications in most positions of the guide arm or the slicer site of the passenger arm reduced their efficacy. Phosphorothioate modifications were found to induce interferon-beta (IFN-β) and TNF-α in MRC-5 cells.
Fig. 5

Effect of 2′-O-Me modifications and loop size on serum stability of sshRNAs. sshRNAs were incubated with 10% human serum at 37°C for the times shown. Aliquots were analyzed by denaturing 12% PAGE. Adapted from Ge et al. (2010a)

Table 3

Effect of 2′-O-Me modification on immune stimulation by sshRNAs


SG142 (3′-overhang)

SG118 (3′-overhang)

T7 transcribed shRNA

Blunt-ended version of SG142





1.5 ± 0.2

1.3 ± 0.1

192 ± 7

2.2 ± 0.5

393 ± 35

7.3 ± 1.8

0.6 ± 0.1


2.8 ± 0.1

1.3 ± 0.1

21.5 ± 1.7

1.5 ± 0.2

83 ± 27

4.6 ± 1.4

0.9 ± 0.1


7.7 ± 0.4

1.5 ± 0.4

548 ± 92

1.8 ± 0.6

319 ± 49

5.9 ± 2.8

1.3 ± 0.2


4.3 ± 0.2

1.3 ± 0.1

67.6 ± 3.7

1.5 ± 0.1

95 ± 5

8.0 ± 0.9

1.1 ± 0.1


1.5 ± 0.4

7.0 ± 3.0

6.5 ± 0.8

2.6 ± 1.2

7.8 ± 4.0

1.2 ± 0.4

0.3 ± 0.04


1.4 ± 0.1

4.5 ± 1.7

5.3 ± 0.4


4.1 ± 1.1

2.1 ± 0.4

0.5 ± 0.1


28.4 ± 10.2

2.8 ± 1.5

18.9 ± 8.5

3.9 ± 1.6

141 ± 35

289 ± 73

13.5 ± 6.1


3.6 ± 0.4

1.2 ± 0.1

12.1 ± 2.2

1.2 ± 0.1

6.2 ± 2.2

4.3 ± 0.4

1.1 ± 0.2

Shown are mean values and standard errors of the mRNA levels (relative to an untreated control and normalized to GAPDH) of genes of interest. 100-nM sshRNAs with (+) and without (–) 2′-O-Me modifications on the loop and alternate nucleotides of the passenger strand were transfected into human MRC-5 cells in triplicate (without heating and snap cooling). SG142 and SG118 are 19-bp sshRNAs with UU loops and 3′-UU overhangs targeting different sequences on the HCV viral RNA. RNA was extracted from cells 24 h post-transfection, and quantitative RT-PCR was performed. Cells that received the transfection reagent Lipofectamine 2000, alone showed no change in levels of the tested genes. A T7-transcribed shRNA, used as positive control, was transfected into cells in equivalent amounts (on a mononucleotide basis). N.D., not done. Adapted from Ge et al. (2010a).

7 Summary: Design of Active sshRNAs

Synthetic sshRNAs can be highly potent RNAi effectors when properly designed. 19-bp R-sshRNAs require longer loops (at least 5–6 nt) and the presence of a 3′ dinucleotide overhang for maximal efficacy (Li et al. 2007; Vlassov et al. 2007; Siolas et al. 2005). In contrast, L-sshRNAs, at least the ones tested, can possess a loop as short as 1–2 nt or even a direct connection between the two strands. A UU loop is particularly effective. A 3′ overhang is not essential for many active L-sshRNAs, and both antisense and sense sequences can be either 18 or 19 nt. Modification of every other nucleotide on the passenger arm, except around the slicer site, is helpful for increasing stability against nucleases (e.g., those found in serum) and minimizing immune stimulation, particularly for blunt-ended sshRNAs. A heat–snap cooling step immediately prior to use is advisable to eliminate dimers.



This work was supported by National Institutes of Health grant numbers R44AI056611, R44AI074256, and R43AI074214 (B.H.J.).


  1. Alter MJ, Kruszon-Moran D, Nainan OV et al (1999) The prevalence of hepatitis C virus infection in the United States, 1988 through 1994. N Engl J Med 341:556–562PubMedCrossRefGoogle Scholar
  2. Amarzguioui M, Lundberg P, Cantin E et al (2006) Rational design and in vitro and in vivo delivery of Dicer substrate siRNA. Nat Protoc 1:508–517PubMedCrossRefGoogle Scholar
  3. Arbuthnot P, Longshaw V, Naidoo T et al (2007) Opportunities for treating chronic hepatitis B and C virus infection using RNA interference. J Viral Hepat 14:447–459PubMedCrossRefGoogle Scholar
  4. Armstrong GL, Alter MJ, McQuillan GM et al (2000) The past incidence of hepatitis C virus infection: implications for the future burden of chronic liver disease in the United States. Hepatology 31:777–782PubMedCrossRefGoogle Scholar
  5. Barik S, Bitko V (2006) Prospects of RNA interference therapy in respiratory viral diseases: update 2006. Expert Opin Biol Ther 6:1151–1160PubMedCrossRefGoogle Scholar
  6. Beaulieu PL (2007) Non-nucleoside inhibitors of the HCV NS5B polymerase: progress in the discovery and development of novel agents for the treatment of HCV infections. Curr Opin Investig Drugs 8:614–634PubMedGoogle Scholar
  7. Bernacchi S, Ennifar E, Toth K et al (2005) Mechanism of hairpin-duplex conversion for the HIV-1 dimerization initiation site. J Biol Chem 280:40112–40121PubMedCrossRefGoogle Scholar
  8. Bernards R, Brummelkamp TR, Beijersbergen RL (2006) shRNA libraries and their use in cancer genetics. Nat Methods 3:701–706PubMedCrossRefGoogle Scholar
  9. Bernstein E, Caudy AA, Hammond SM et al (2001) Role for a bidentate ribonuclease in the initiation step of RNA interference. Nature 409:363–366PubMedCrossRefGoogle Scholar
  10. Bolcato-Bellemin AL, Bonnet ME, Creusat G et al (2007) Sticky overhangs enhance siRNA-mediated gene silencing. Proc Natl Acad Sci USA 104:16050–16055PubMedCrossRefGoogle Scholar
  11. Boudreau RL, Monteys AM, Davidson BL (2008) Minimizing variables among hairpin-based RNAi vectors reveals the potency of shRNAs. RNA 14:1834–1844PubMedCrossRefGoogle Scholar
  12. Brummelkamp TR, Bernards R, Agami R (2002) A system for stable expression of short interfering RNAs in mammalian cells. Science 296:550–553PubMedCrossRefGoogle Scholar
  13. Castanotto D, Rossi JJ (2009) The promises and pitfalls of RNA-interference-based therapeutics. Nature 457:426–433PubMedCrossRefGoogle Scholar
  14. Chang K, Elledge SJ, Hannon GJ (2006) Lessons from nature: microRNA-based shRNA libraries. Nat Methods 3:707–714PubMedCrossRefGoogle Scholar
  15. Collingwood MA, Rose SD, Huang L et al (2008) Chemical modification patterns compatible with high potency dicer-substrate small interfering RNAs. Oligonucleotides 18:187–200PubMedCrossRefGoogle Scholar
  16. Dallas A, Ge Q, Ilves H et al (2012) Right- and Left-loop short shRNAs have distinct and unusual mechanisms of silencing. SubmittedGoogle Scholar
  17. Deuffic-Burban S, Wong JB, Valleron AJ et al (2004) Comparing the public health burden of chronic hepatitis C and HIV infection in France. J Hepatol 40:319–326PubMedCrossRefGoogle Scholar
  18. Deuffic-Burban S, Mohamed MK, Larouze B et al (2006) Expected increase in hepatitis C-related mortality in Egypt due to pre-2000 infections. J Hepatol 44:455–461PubMedCrossRefGoogle Scholar
  19. Deuffic-Burban S, Poynard T, Sulkowski MS et al (2007) Estimating the future health burden of chronic hepatitis C and human immunodeficiency virus infections in the United States. J Viral Hepat 14:107–115PubMedCrossRefGoogle Scholar
  20. Deuffic-Burban S, Abiteboul D, Lot F et al (2009) Costs and cost-effectiveness of different follow-up schedules for detection of occupational hepatitis C virus infection. Gut 58:105–110PubMedCrossRefGoogle Scholar
  21. Deuffic-Burban S, Mathurin P, Pol S et al (2012) Impact of hepatitis C triple therapy availability upon the number of patients to be treated and associated costs in France: a model-based analysis. Gut 61:290–296PubMedCrossRefGoogle Scholar
  22. Diederichs S, Haber DA (2007) Dual role for argonautes in microRNA processing and posttranscriptional regulation of microRNA expression. Cell 131:1097–1108PubMedCrossRefGoogle Scholar
  23. Dorsett Y, Tuschl T (2004) siRNAs: applications in functional genomics and potential as therapeutics. Nat Rev Drug Discov 3:318–329PubMedCrossRefGoogle Scholar
  24. Elbashir SM, Lendeckel W, Tuschl T (2001) RNA interference is mediated by 21- and 22-nucleotide RNAs. Genes Dev 1:188–200CrossRefGoogle Scholar
  25. Fewell GD, Schmitt K (2006) Vector-based RNAi approaches for stable, inducible and genome-wide screens. Drug Discov Today 11:975–982PubMedCrossRefGoogle Scholar
  26. Filipowicz W (2005) RNAi: the nuts and bolts of the RISC machine. Cell 122:17–20PubMedCrossRefGoogle Scholar
  27. Fowler T, Bamberg S, Moller P et al (2005) Inhibition of Marburg virus protein expression and viral release by RNA interference. J Gen Virol 86:1181–1188PubMedCrossRefGoogle Scholar
  28. Ge Q, Dallas A, Ilves H et al (2010a) Effects of chemical modification on the potency, serum stability, and immunostimulatory properties of short shRNAs. RNA 16:118–130PubMedCrossRefGoogle Scholar
  29. Ge Q, Ilves H, Dallas A et al (2010b) Minimal-length short hairpin RNAs: the relationship of structure and RNAi activity. RNA 16:106–117PubMedCrossRefGoogle Scholar
  30. Gitlin L, Karelsky S, Andino R (2002) Short interfering RNA confers intracellular antiviral immunity in human cells. Nature 418:430–434PubMedCrossRefGoogle Scholar
  31. Grimm D, Streetz KL, Jopling CL et al (2006) Fatality in mice due to oversaturation of cellular microRNA/short hairpin RNA pathways. Nature 441:537–541PubMedCrossRefGoogle Scholar
  32. Haasnoot PC, Cupac D, Berkhout B (2003) Inhibition of virus replication by RNA interference. J Biomed Sci 10:607–616PubMedCrossRefGoogle Scholar
  33. Hammond SM, Bernstein E, Beach D et al (2000) An RNA-directed nuclease mediates post-transcriptional gene silencing in Drosophila cells. Nature 404:293–296PubMedCrossRefGoogle Scholar
  34. Hammond SM, Boettcher S, Caudy AA et al (2001) Argonaute2, a link between genetic and biochemical analyses of RNAi. Science 293:1146–1150PubMedCrossRefGoogle Scholar
  35. Harborth J, Elbashir SM, Vandenburgh K et al (2003) Sequence, chemical, and structural variation of small interfering RNAs and short hairpin RNAs and the effect on mammalian gene silencing. Antisense Nucleic Acid Drug Dev 13:83–105PubMedCrossRefGoogle Scholar
  36. Hinton TM, Wise TG, Cottee PA et al (2008) Native microRNA loop sequences can improve short hairpin RNA processing for virus gene silencing in animal cells. JRNAi Gene Silencing 4:295–301Google Scholar
  37. Ilves H, Kaspar RL, Wang Q et al (2006) Inhibition of hepatitis C IRES-mediated gene expression by small hairpin RNAs in human hepatocytes and mice. Ann NY Acad Sci 1082:52–55PubMedCrossRefGoogle Scholar
  38. Iorns E, Lord CJ, Turner N et al (2007) Utilizing RNA interference to enhance cancer drug discovery. Nat Rev Drug Discov 6:556–568PubMedCrossRefGoogle Scholar
  39. Jarczak D, Korf M, Beger C et al (2005) Hairpin ribozymes in combination with siRNAs against highly conserved hepatitis C virus sequence inhibit RNA replication and protein translation from hepatitis C virus subgenomic replicons. FEBS J 272:5910–5922PubMedCrossRefGoogle Scholar
  40. Jinek M, Doudna JA (2009) A three-dimensional view of the molecular machinery of RNA interference. Nature 457:405–412PubMedCrossRefGoogle Scholar
  41. Jucker FM, Pardi A (1995) Solution structure of the CUUG hairpin loop: a novel RNA tetraloop motif. Biochemistry 34:14416–14427PubMedCrossRefGoogle Scholar
  42. Kanda T, Steele R, Ray R et al (2007) Small interfering RNA targeted to hepatitis C virus 5′-nontranslated region exerts potent antiviral effect. J Virol 81:669–676PubMedCrossRefGoogle Scholar
  43. Kapadia SB, Brideau-Andersen A, Chisari FV (2003) Interference of hepatitis C virus RNA replication by short interfering RNAs. Proc Natl Acad Sci USA 100:2014–2018PubMedCrossRefGoogle Scholar
  44. Kawasaki H, Suyama E, Iyo M et al (2003) siRNAs generated by recombinant human Dicer induce specific and significant but target site-independent gene silencing in human cells. Nucleic Acids Res 31:981–987PubMedCrossRefGoogle Scholar
  45. Khvorova A, Reynolds A, Jayasena SD (2003) Functional siRNAs and miRNAs exhibit strand bias. Cell 115:209–216PubMedCrossRefGoogle Scholar
  46. Kim D, Rossi J (2008) RNAi mechanisms and applications. Biotechniques 44:613–616PubMedCrossRefGoogle Scholar
  47. Kim M, Shin D, Kim SI et al (2006) Inhibition of hepatitis C virus gene expression by small interfering RNAs using a tri-cistronic full-length viral replicon and a transient mouse model. Virus Res 122:1–10PubMedCrossRefGoogle Scholar
  48. Korf M, Meyer A, Jarczak D et al (2007) Inhibition of HCV subgenomic replicons by siRNAs derived from plasmids with opposing U6 and H1 promoters. J Viral Hepat 14:122–132PubMedCrossRefGoogle Scholar
  49. Kraynack BA, Baker BF (2006) Small interfering RNAs containing full 2′-O-methylribonucleotide-modified sense strands display Argonaute2/eIF2C2-dependent activity. RNA 12:163–176PubMedCrossRefGoogle Scholar
  50. Lanford RE, Hildebrandt-Eriksen ES, Petri A et al (2010) Therapeutic silencing of microRNA-122 in primates with chronic hepatitis C virus infection. Science 327:198–201PubMedCrossRefGoogle Scholar
  51. Law MG, Dore GJ, Bath N et al (2003) Modelling hepatitis C virus incidence, prevalence and long-term sequelae in Australia, 2001. Int J Epidemiol 32:717–724PubMedCrossRefGoogle Scholar
  52. Lee YS, Nakahara K, Pham JW et al (2004) Distinct roles for Drosophila Dicer-1 and Dicer-2 in the siRNA/miRNA silencing pathways. Cell 117:69–81PubMedCrossRefGoogle Scholar
  53. Leuschner PJ, Ameres SL, Kueng S et al (2006) Cleavage of the siRNA passenger strand during RISC assembly in human cells. EMBO Rep 7:314–320PubMedCrossRefGoogle Scholar
  54. Li L, Lin X, Staver M et al (2005a) Evaluating hypoxia-inducible factor-1alpha as a cancer therapeutic target via inducible RNA interference in vivo. Cancer Res 65:7249–7258PubMedCrossRefGoogle Scholar
  55. Li MJ, Kim J, Li S et al (2005b) Long-term inhibition of HIV-1 infection in primary hematopoietic cells by lentiviral vector delivery of a triple combination of anti-HIV shRNA, anti-CCR5 ribozyme, and a nucleolar-localizing TAR decoy. Mol Ther 12:900–909PubMedCrossRefGoogle Scholar
  56. Li L, Lin X, Khvorova A et al (2007) Defining the optimal parameters for hairpin-based knockdown constructs. RNA 13:1765–1774PubMedCrossRefGoogle Scholar
  57. Liu J, Carmell MA, Rivas FV et al (2004) Argonaute2 is the catalytic engine of mammalian RNAi. Science 305:1437–1441PubMedCrossRefGoogle Scholar
  58. Liu HW, Cosa G, Landes CF et al (2005) Single-molecule FRET studies of important intermediates in the nucleocapsid-protein-chaperoned minus-strand transfer step in HIV-1 reverse transcription. Biophys J 89:3470–3479PubMedCrossRefGoogle Scholar
  59. Liu M, Ding H, Zhao P et al (2006) RNA interference effectively inhibits mRNA accumulation and protein expression of hepatitis C virus core and E2 genes in human cells. Biosci Biotechnol Biochem 70:2049–2055PubMedCrossRefGoogle Scholar
  60. Liu-Young G, Kozal MJ (2008) Review: hepatitis C protease and polymerase inhibitors in development. AIDS Patient Care STDS 22:449–457PubMedCrossRefGoogle Scholar
  61. MacRae IJ, Doudna JA (2007) Ribonuclease revisited: structural insights into ribonuclease III family enzymes. Curr Opin Struct Biol 17:138–145PubMedCrossRefGoogle Scholar
  62. Martinez J, Patkaniowska A, Urlaub H et al (2002) Single-stranded antisense siRNAs guide target RNA cleavage in RNAi. Cell 110:563PubMedCrossRefGoogle Scholar
  63. Matranga C, Tomari Y, Shin C et al (2005) Passenger-strand cleavage facilitates assembly of siRNA into Ago2-containing RNAi enzyme complexes. Cell 123:607–620PubMedCrossRefGoogle Scholar
  64. McCaffrey AP, Meuse L, Pham TT et al (2002) RNA interference in adult mice. Nature 418:38–39PubMedCrossRefGoogle Scholar
  65. McHutchison JG, Patel K, Pockros P et al (2006) A phase I trial of an antisense inhibitor of hepatitis C virus (ISIS 14803), administered to chronic hepatitis C patients. J Hepatol 44:88–96PubMedCrossRefGoogle Scholar
  66. McManus MT, Petersen CP, Haines BB et al (2002) Gene silencing using micro-RNA designed hairpins. RNA 8:842–850PubMedCrossRefGoogle Scholar
  67. Meister G, Landthaler M, Patkaniowska A et al (2004) Human Argonaute2 mediates RNA cleavage targeted by miRNAs and siRNAs. Mol Cell 15:185–197PubMedCrossRefGoogle Scholar
  68. Miyoshi K, Tsukumo H, Nagami T et al (2005) Slicer function of Drosophila Argonautes and its involvement in RISC formation. Genes Dev 19:2837–2848PubMedCrossRefGoogle Scholar
  69. Natt F (2007) siRNAs in drug discovery: target validation and beyond. Curr Opin Mol Ther 9:242–247PubMedGoogle Scholar
  70. Neumann AU, Lam NP, Dahari H et al (1998) Hepatitis C viral dynamics in vivo and the antiviral efficacy of interferon-alpha therapy. Science 282:103–107PubMedCrossRefGoogle Scholar
  71. Nykanen A, Haley B, Zamore PD (2001) ATP requirements and small interfering RNA structure in the RNA interference pathway. Cell 107:309–321PubMedCrossRefGoogle Scholar
  72. Ogata N, Alter HJ, Miller RH et al (1991) Nucleotide sequence and mutation rate of the H strain of hepatitis C virus. Proc Natl Acad Sci USA 88:3392–3396PubMedCrossRefGoogle Scholar
  73. Okamoto H, Kojima M, Okada S et al (1992) Genetic drift of hepatitis C virus during an 8.2-year infection in a chimpanzee: variability and stability. Virology 190:894–899PubMedCrossRefGoogle Scholar
  74. Okamura K, Ishizuka A, Siomi H et al (2004) Distinct roles for Argonaute proteins in small RNA-directed RNA cleavage pathways. Genes Dev 18:1655–1666PubMedCrossRefGoogle Scholar
  75. Pham JW, Pellino JL, Lee YS et al (2004) A Dicer-2-dependent 80s complex cleaves targeted mRNAs during RNAi in Drosophila. Cell 117:83–94PubMedCrossRefGoogle Scholar
  76. Prabhu R, Vittal P, Yin Q et al (2005) Small interfering RNA effectively inhibits protein expression and negative strand RNA synthesis from a full-length hepatitis C virus clone. J Med Virol 76:511–519PubMedCrossRefGoogle Scholar
  77. Prabhu R, Garry RF, Dash S (2006) Small interfering RNA targeted to stem-loop II of the 5′ untranslated region effectively inhibits expression of six HCV genotypes. Virol J 3:100PubMedCrossRefGoogle Scholar
  78. Rand TA, Ginalski K, Grishin NV et al (2004) Biochemical identification of Argonaute 2 as the sole protein required for RNA-induced silencing complex activity. Proc Natl Acad Sci USA 101:14385–14389PubMedCrossRefGoogle Scholar
  79. Rand TA, Petersen S, Du F et al (2005) Argonaute2 cleaves the anti-guide strand of siRNA during RISC activation. Cell 123:621–629PubMedCrossRefGoogle Scholar
  80. Randall G, Rice CM (2004) Interfering with hepatitis C virus RNA replication. Virus Res 102:19–25PubMedCrossRefGoogle Scholar
  81. Randall G, Grakoui A, Rice CM (2003) Clearance of replicating hepatitis C virus replicon RNAs in cell culture by small interfering RNAs. Proc Natl Acad Sci USA 100:235–240PubMedCrossRefGoogle Scholar
  82. Ray Kim W (2002) Global epidemiology and burden of hepatitis C. Microbes Infect 4:1219–1225PubMedCrossRefGoogle Scholar
  83. Ray RB, Kanda T (2009) Inhibition of HCV replication by small interfering RNA. Methods Mol Biol 510:251–262PubMedCrossRefGoogle Scholar
  84. Rivas FV, Tolia NH, Song JJ et al (2005) Purified Argonaute2 and an siRNA form recombinant human RISC. Nat Struct Mol Biol 12:340–349PubMedCrossRefGoogle Scholar
  85. Robb GB, Rana TM (2007) RNA helicase A interacts with RISC in human cells and functions in RISC loading. Mol Cell 26:523–537PubMedCrossRefGoogle Scholar
  86. Robinson M, Tian Y, Delaney WEt et al (2011) Preexisting drug-resistance mutations reveal unique barriers to resistance for distinct antivirals. Proc Natl Acad Sci USA 108:10290–10295PubMedCrossRefGoogle Scholar
  87. Ronn R, Sandstrom A (2008) New developments in the discovery of agents to treat hepatitis C. Curr Top Med Chem 8:533–562PubMedCrossRefGoogle Scholar
  88. Rossi JJ, June CH, Kohn DB (2007) Genetic therapies against HIV. Nat Biotechnol 25:1444–1454PubMedCrossRefGoogle Scholar
  89. Salomon JA, Weinstein MC, Hammitt JK et al (2002) Empirically calibrated model of hepatitis C virus infection in the United States. Am J Epidemiol 156:761–773PubMedCrossRefGoogle Scholar
  90. Sarrazin C, Zeuzem S (2010) Resistance to direct antiviral agents in patients with hepatitis C virus infection. Gastroenterology 138:447–462PubMedCrossRefGoogle Scholar
  91. Schwarz DS, Hutvagner G, Du T et al (2003) Asymmetry in the assembly of the RNAi enzyme complex. Cell 115:199–208PubMedCrossRefGoogle Scholar
  92. Sen A, Steele R, Ghosh AK et al (2003) Inhibition of hepatitis C virus protein expression by RNA interference. Virus Res 96:27–35PubMedCrossRefGoogle Scholar
  93. Seo MY, Abrignani S, Houghton M et al (2003) Small interfering RNA-mediated inhibition of hepatitis C virus replication in the human hepatoma cell line Huh-7. J Virol 77:810–812PubMedCrossRefGoogle Scholar
  94. Seyhan AA, Alizadeh BN, Lundstrom K et al (2007) RNA interference-mediated inhibition of Semliki Forest Virus replication in mammalian cells. Oligonucleotides 17:473–484PubMedCrossRefGoogle Scholar
  95. Shenouda SK, Alahari SK (2009) MicroRNA function in cancer: oncogene or a tumor suppressor? Cancer Metastasis Rev 28:369–378PubMedCrossRefGoogle Scholar
  96. Shepard CW, Finelli L, Alter MJ (2005) Global epidemiology of hepatitis C virus infection. Lancet Infect Dis 5:558–567PubMedCrossRefGoogle Scholar
  97. Shin D, Lee H, Kim SI et al (2009) Optimization of linear double-stranded RNA for the production of multiple siRNAs targeting hepatitis C virus. RNA 15:898–910PubMedCrossRefGoogle Scholar
  98. Silva JM, Li MZ, Chang K et al (2005) Second-generation shRNA libraries covering the mouse and human genomes. Nat Genet 37:1281–1288PubMedGoogle Scholar
  99. Siolas D, Lerner C, Burchard J et al (2005) Synthetic shRNAs as potent RNAi triggers. Nat Biotechnol 23:227–231PubMedCrossRefGoogle Scholar
  100. Siomi H, Siomi MC (2009) On the road to reading the RNA-interference code. Nature 457:396–404PubMedCrossRefGoogle Scholar
  101. Song JJ, Smith SK, Hannon GJ et al (2004) Crystal structure of Argonaute and its implications for RISC slicer activity. Science 305:1434–1437PubMedCrossRefGoogle Scholar
  102. Stegmeier F, Hu G, Rickles RJ et al (2005) A lentiviral microRNA-based system for single-copy polymerase II-regulated RNA interference in mammalian cells. Proc Natl Acad Sci USA 102:13212–13217PubMedCrossRefGoogle Scholar
  103. Sun X, Li JM, Wartell RM (2007) Conversion of stable RNA hairpin to a metastable dimer in frozen solution. RNA 13:2277–2286PubMedCrossRefGoogle Scholar
  104. Takigawa Y, Nagano-Fujii M, Deng L et al (2004) Suppression of hepatitis C virus replicon by RNA interference directed against the NS3 and NS5B regions of the viral genome. Microbiol Immunol 48:591–598PubMedGoogle Scholar
  105. Tomari Y, Zamore PD (2005) Perspective: machines for RNAi. Genes Dev 19:517–529PubMedCrossRefGoogle Scholar
  106. Tomari Y, Du T, Haley B et al (2004a) RISC assembly defects in the Drosophila RNAi mutant armitage. Cell 116:831–841PubMedCrossRefGoogle Scholar
  107. Tomari Y, Matranga C, Haley B et al (2004b) A protein sensor for siRNA asymmetry. Science 306:1377–1380PubMedCrossRefGoogle Scholar
  108. Torriani FJ, Rodriguez-Torres M, Rockstroh JK et al (2004) Peginterferon Alfa-2a plus ribavirin for chronic hepatitis C virus infection in HIV-infected patients. N Engl J Med 351:438–450PubMedCrossRefGoogle Scholar
  109. Vlassov AV, Ilves H, Johnston BH (2006) Inhibition of hepatitis C IRES-mediated gene expression by 8-17 deoxyribozymes in human tissue culture cells. Dokl Biochem Biophys 410:257–259PubMedCrossRefGoogle Scholar
  110. Vlassov AV, Korba B, Farrar K et al (2007) shRNAs targeting hepatitis C: effects of sequence and structural features, and comparison with siRNA. Oligonucleotides 17:223–236PubMedCrossRefGoogle Scholar
  111. Wang Q, Contag CH, Ilves H et al (2005) Small hairpin RNAs efficiently inhibit hepatitis C IRES-mediated gene expression in human tissue culture cells and a mouse model. Mol Ther 12:562–568PubMedCrossRefGoogle Scholar
  112. Watanabe T, Umehara T, Kohara M (2007) Therapeutic application of RNA interference for hepatitis C virus. Adv Drug Deliv Rev 59:1263–1276PubMedCrossRefGoogle Scholar
  113. Wilson JA, Richardson CD (2005) Hepatitis C virus replicons escape RNA interference induced by a short interfering RNA directed against the NS5b coding region. J Virol 79:7050–7058PubMedCrossRefGoogle Scholar
  114. Wilson JA, Jayasena S, Khvorova A et al (2003) RNA interference blocks gene expression and RNA synthesis from hepatitis C replicons propagated in human liver cells. Proc Natl Acad Sci USA 100:2783–2788PubMedCrossRefGoogle Scholar
  115. Ye X, Huang N, Liu Y et al (2011) Structure of C3PO and mechanism of human RISC activation. Nat Struct Mol Biol 18:650–657PubMedCrossRefGoogle Scholar
  116. Yokota T, Sakamoto N, Enomoto N et al (2003) Inhibition of intracellular hepatitis C virus replication by synthetic and vector-derived small interfering RNAs. EMBO Rep 4:602–608PubMedCrossRefGoogle Scholar
  117. Zamore PD, Tuschl T, Sharp PA et al (2000) RNAi: double-stranded RNA directs the ATP-dependent cleavage of mRNA at 21 to 23 nucleotide intervals. Cell 101:25–33PubMedCrossRefGoogle Scholar
  118. Zekri AR, Bahnassy AA, El-Din HM et al (2009) Consensus siRNA for inhibition of HCV genotype-4 replication. Virol J 6:13PubMedCrossRefGoogle Scholar
  119. Zhou D, Zhang J, Wang C et al (2009) A method for detecting and preventing negative RNA interference in preparation of lentiviral vectors for siRNA delivery. RNA 15:732–740PubMedCrossRefGoogle Scholar

Copyright information

© Springer-Verlag Berlin Heidelberg 2012

Authors and Affiliations

  1. 1.SomaGenics, Inc.Santa CruzUSA
  2. 2.Department of ImmunologyPeking University Health Science CenterBeijingPR China

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