RNA interference (RNAi) is now an umbrella term referring to post-transcriptional gene silencing mediated by either degradation or translation arrest of target RNA. This process is initiated by double-stranded RNA with sequence homology driving specificity. The discovery that 21–23 nucleotide RNA duplexes (small-interfering RNAs, siRNAs) mediate RNAi in mammalian cells opened the door to the therapeutic use of siRNAs. While much work remains to optimize delivery and maintain specificity, the therapeutic advantages of siRNAs for treatment of viral infection, dominant disorders, cancer, and neurological disorders show great promise.
The phenomena of RNA interference (RNAi) was originally observed in transgenic plants where it was termed cosuppression and later recognized as a form of homology-dependent gene silencing.1 Work in Caenorhabditis elegans led to the discovery that double-stranded RNA (dsRNA) triggers RNAi.2 Injection of sense and antisense RNAs resulted in negligible decreases in targeted mRNA, whereas introduction of dsRNA resulted in effective and specific mRNA knockdown. The silencing effects of dsRNA in C. elegans are systemic and heritable and RNAi is now routinely used as a reverse genetics tool in C. elegans.
Processing and production of small-interfering RNAs (siRNAs)
While initial studies utilized introduction of exogenous dsRNA, it is now clear that higher eukaryotes contain a large number of genes that encode small RNAs referred to as micro-RNAs (miRNAs).3 Both miRNAs and exogenous dsRNAs mediate their effects at the RNA level, miRNAs by inhibiting translation and exogenous dsRNAs through degradation of target RNAs. The difference is due to the degree of complementarity between the siRNA and the target. Perfect complementarity induces degradation, whereas several mismatches lead to translation arrest.4 For miRNAs, nuclear transcripts are processed into shorter hairpin structures that are then transported to the cytoplasm for final processing and assembly of effector complexes that are similar for both exogenous dsRNA and miRNAs (Figure 1). In both cases, processing by the enzyme Dicer leads to 21–23 nucleotide (nt) siRNAs, which guide silencing of target mRNAs through a base-pairing-dependent mechanism.5, 6, 7, 8, 9 Dicer cleaves long dsRNAs approximately every 22 nt yielding small duplexes containing 2–3 nt overhangs, 5′ phosphates, and free 3′ hydroxyl-termini.10 After processing, siRNAs are incorporated into a multiprotein complex termed the RNA-induced silencing complex (RISC).6 Selection of the strand incorporated into RISC is biased towards the strand with the lowest duplex stability at its 5′-end.11 Once paired, target RNA cleavage occurs near the center of the target–siRNA duplex.12
In invertebrates, siRNAs have shown great specificity and efficacy. As vertebrates exhibit a cell death response to long dsRNAs due to induction of an interferon response and activation of the kinase PKR, it was thought that RNAi might have limited utility in vertebrates.13 However, the use of small dsRNAs apparently sidesteps most problems allowing specific target cleavage.14 With the important caveat that two recent papers have reported activation of an interferon response even with short dsRNA,15, 16 the efficiency and specificity of siRNA-mediated target degradation reported in most studies opens the door to its possible use against human disease. Here, we focus on several obvious classes of disease where degradation of specific mRNA targets would have therapeutic utility.
siRNA targeting in dominant disorders
For disorders that exert their effects either via a dominant-negative or gain-of-function mechanism, the ability to specifically eliminate mutant transcripts would be highly beneficial. The major obstacle is to successfully distinguish and degrade mutant RNAs leaving wild-type transcripts unaffected. There are several targeting strategies that can be utilized.
Direct mutation targeting
The simplest siRNA design takes advantage of sequence differences between wild-type and mutant RNAs (Figure 2a). This strategy has been used in a cellular model of frontotemporal dementia with parkinsonism linked to chromosome 17 (FTDP-17).17 FTDP-17 is mediated by dominant mutations in Tau that lead to a buildup of neurofibrillary tangles and neuronal degeneration. Four different FTDP-17 missense mutations were targeted using 21–24 nt siRNAs with the mutant nucleotide located at one of four central positions within the siRNA. Only one of the four siRNAs (corresponding to the V337M mutation) efficiently degraded mutant Tau. Unfortunately, this siRNA also led to considerable degradation of wild-type Tau. To increase specificity, siRNAs were designed that contained a second mismatch either one nucleotide upstream or three nucleotides downstream of the V337M mutation. The introduction of a second mismatch just upstream of the V337M mutation resulted in specific degradation of mutant Tau without any apparent loss of wild-type protein.17
Small deletions can also be targeted in this manner. DYT1 dystonia is an incompletely penetrant, dominant disorder in which the overwhelming majority of patients inherit a 3 nt deletion that eliminates a glutamic acid residue in the torsinA protein.18 Using siRNAs that placed the 3 nt sequence difference near the central portion of the siRNA resulted in allele-specific degradation. The efficiency and specificity of these results were also obtained in the simulated heterozygous state in which both wild-type and mutant torsinA were cotransfected with siRNAs.18 Diseases such as DYT1 dystonia, where a common mutation is present in greater than 50% of the disease population, represent the ideal situation because the design and optimization of a single siRNA would apply to many patients. In contrast, patient- or family-specific therapy and specialized optimization would be required for diseases caused by variable mutations.
Indirect mutation targeting
In cases where mutant alleles are too similar to wild type, an indirect targeting approach might succeed (Figure 2b). One example would include polyglutamine-repeat expansions. Since smaller numbers of repeats are found in normal alleles, one might expect difficulty generating a specific siRNA. This was in fact the case when Miller et al17 designed siRNAs targeting the CAG repeat or an immediately adjacent region in spinocerebellar ataxia type 3 (SCA3), a dominant polyglutamine disorder resulting in aberrant accumulation of ataxin-3. To more selectively target the mutant SCA3 allele, a SNP (G to C transversion) was utilized immediately downstream of the CAG repeat. Chromosomes containing the C allele are in linkage disequilibrium with disease-causing expansions in the majority of patients. siRNAs were designed to target this region with the SNP located at position 10 of the siRNA, allowing selective degradation of mutant transcripts. Shifting the SNP to position 7, however, led to degradation of both wild-type and mutant SCA3 transcripts. Interestingly, specificity could also be achieved with the SNP in the seventh position by introducing a second mismatch such that wild-type SCA3 contained two mismatches while the C variant (and therefore mutant expansion) had only one. The use of a SNP in linkage disequilibrium may actually be a more useful strategy than directly targeting specific mutations, since only one siRNA design would be necessary to target a larger percentage of the disease population.
Targeting aberrant splicing isoforms
Disease-associated splicing isoforms19, 20 can be classified into two categories: intron retention (inclusion of specific introns or activation of cryptic splice sites within introns) or exon skipping (complete skipping or activation of cryptic splice sites within exons). Both groups can be degraded using sequence-specific siRNAs.
Exon-specific siRNA was tested by targeting the variable exons of the Drosophila Dscam exon 4 cluster in which only one of 12 variable exons are spliced into mRNA.21 Instead of using small, 21–23 nt RNA duplexes, ‘RNA triggers’ composed of dsRNAs corresponding to an entire exon sequence (in this case, either alternative exon 4.1 or 4.3) were used. Both sets of RNA triggers specifically and efficiently degraded their respective isoforms in Drosophila S2 cells. Sequence similarity between one or more of the variable exons led to some nonspecific degradation, which was eliminated by using shorter dsRNAs. Exon-specific siRNA has also been successful with more traditional approaches using siRNAs complementary to alternatively spliced isoforms (Figure 2c).22, 23
Aberrant splicing events can lead to the production of dominant-negative proteins. The majority of mutations that lead to isolated growth hormone deficiency (GHD) type II, a dominant form of GHD, affect splicing of exon 3 leading to the production of a smaller, 17.5 kDa isoform. The 17.5 kDa isoform exerts a dominant-negative effect on secretion of the biologically active 22 kDa growth hormone isoform. Since the 17.5 kDa isoform results from skipping of exon 3, it can be targeted with siRNAs directed against the unique exons 2–4 junction (Figure 2d). This targeting approach was used to degrade specifically transcripts encoding the 17.5 kDa isoform with no apparent effect on either the 22kDa isoform or another, smaller 20 kDa isoform.24 Thus, properly designed siRNAs can be used to specifically degrade aberrant or alternatively spliced mRNAs (Figure 2c and d).
siRNA and neurological disorders
Alzheimer's disease (AD) is a progressive neurodegenerative disorder whose pathological findings include neurofibrillary tangles and aggregation of β-amyloid peptides. One hypothesis as to the pathogenesis of AD involves increased production of β-amyloid peptides, particularly Aβ42, from amyloid precursor protein (APP) by two enzymes, a β-secretase and a γ-secretase. The β-secretase (BACE1) is upregulated in the brains of AD patients. Since mice lacking BACE1 do not generate β-amyloid peptides and have no obvious developmental abnormalities, it is an attractive target for siRNA therapeutics. siRNA-mediated knockdown of BACE1 decreased Aβ140 and Aβ142 secretion in mouse primary cortical neurons expressing recombinant APP and in primary cortical neurons carrying two human APP mutations (KM670/671NL).25 Also, knockdown of BACE1 reduced cell death and decreased secreted Aβ levels during oxidative stress, which induces BACE1 protein and its APP cleavage products. These findings are promising for the future of AD therapy, although more work is needed, especially in mouse models. The use of siRNAs in other neurological disorders has recently been reviewed.26
siRNA targeting of downstream effectors of disease
siRNAs could also play an important role in preventing side effects and in slowing or halting progression of disease. Previous studies of siRNA delivery via high-pressure tail vein injections demonstrated that the liver is the primary site of siRNA uptake.20, 27 Therefore, the liver and liver diseases such as viral hepatitis are an ideal system in which to study siRNA efficacy as a modifier of human disease (see Table 1 for viral studies). Fulminant hepatitis results from long-term liver injury via hepatic cell death from Fas-mediated apoptosis. Transgenic mice deficient in Fas exhibit decreased liver fibrosis and increased survival following exposure to various insults. siRNAs directed against Fas and delivered to the liver via hydrodynamic tail vein injection were shown to protect against fulminant hepatitis.28 Hydrodynamic transfection was also used to prevent acute liver failure (ALF), which results from activation of a caspase cascade triggering massive hepatocyte apoptosis.29 siRNAs targeted against caspase-8 protected mice from the development of ALF but in this case, a second delivery technique involving direct injection into the portal vein along with Lipiodol was also used. Portal vein injection successfully inhibited apoptosis in approximately 10–20% of hepatocytes compared to complete apoptosis in controls. In an accompanying set of experiments, liver damage was induced by delivery of vectors expressing FasL. Pretreatment via tail vein injection of caspase-8 siRNAs resulted in significant protection from FasL-mediated hepatocytic injury and improved overall survival of mice treated with activators of apoptosis. Importantly, the delivery of caspase-8 siRNAs after the induction of liver injury significantly improved the survival of ALF animals, demonstrating that siRNA therapeutics can be beneficial after the start of liver injury, the far more likely clinical scenario.
siRNA and cancer
Cancers often have upregulated or misappropriately expressed genes that lead to uncontrolled cell growth. One such target is the M-BCR/ABL fusion gene, the oncogenic product of the so-called Philadelphia chromosome, which leads to chronic myeloid leukemia (CML). A recent comparative study showed that siRNAs directed against BCR/ABL transcripts induced apoptosis roughly equivalent to STI 571, a small molecule drug currently in use in CML patients.30 The development of additional therapeutic options for CML beyond STI 571 is particularly important in light of documented resistance to STI 571.30
siRNAs have also been used to target K-RAS transcripts carrying the valine-112 oncogenic mutation (K-RASV112), which constitutively activates RAS leading to pancreatic and colon cancer.31 Knockdown of K-RASV112 resulted in specific degradation of K-RASV112 and inhibition of colony growth in soft agar. In addition, CAPAN-1 cells stably expressing siRNAs against K-RASV112 did not cause tumors when injected into nude mice, whereas control cells produced tumors. This work and others32 make it easy to imagine the numerous attractive siRNA targets in the fight against cancer. Several recent studies examining the use of siRNAs in cancer models are listed in Table 2.
Obstacles to effective siRNA therapeutics
For all siRNAs, target specificity needs to be validated. In general, it appears that specificity can be attained depending on the position and sequence of a given siRNA. To examine whether global gene expression patterns change in the presence and absence of siRNAs, microarrays have been used.33, 34 These experiments suggest that small numbers of nontarget transcripts with sequence homology as short as 9 nt may be affected. This demonstrates the critical importance of designing appropriate siRNAs and testing them in vivo. Fortunately, it appears that most nonspecific responses can be minimized or eliminated with small changes in siRNA sequences.
As one might expect, the exquisite specificity of siRNA-mediated degradation is both its greatest strength and its potential downfall as an effective anti-viral or chemotherapeutic agent. In fact, it is thought that the normal, endogenous RNAi pathway developed for protection against viruses in plants and fungi.1 Fittingly, siRNAs have been tested extensively against HIV-1, hepatitis, and other viruses (see Table 1).35, 36, 37, 38 Two recent reports have indicated that siRNA-resistant HIV strains can emerge in as little as 25 days.39, 40 Resistance has also been observed with other viral targets.41 To combat resistance, multiple sequences per target and multiple targets per viral genome will likely be necessary.
Another form of resistance is the fact that not all sequences can be targeted by siRNAs. This is likely due to a lack of accessibility of the RNA sequence, either hidden by RNA-binding proteins or by complex secondary structures. Lastly, cells may also develop resistance to RNAi through loss of genes essential for RISC complex formation or selection of suppressors that inhibit degradation. Cymbidium ringspot virus is resistant to RNAi via production of p19, a protein that inhibits RNAi by sequestering dsRNAs.42 Although these forms of resistance are largely hypothetical in humans, appropriate selective pressure could lead to similar problems.
Efficacy/length of effect
To be widely applicable in clinical settings, siRNAs must exert their effects over time. Multiple studies have indicated that degradation generally peaks 36–48 h after introduction and begins to wane around 96 h. This can be extended with repeated siRNA delivery and obviously depends on the rate of target turnover. Several modifications can be used to extend the life of the dsRNAs themselves, notably, 2′-O-methylation.43, 44, 45 In contrast, phosphorothioate backbones appear to be cytotoxic and 2′-O-allylation inhibits activity.45
A variety of strategies have been used to deliver dsRNAs to cells, either directly or by introduction of expression vectors. The advent of lentiviral, adeno-associated, and other retroviral, short hairpin vectors that produce siRNAs allows the use of traditional gene therapy delivery systems. As such, the clinical utility of siRNAs will depend, at least in part, on the development of safe and efficacious delivery systems. The majority of studies performed in mice have used high-pressure, large-volume tail vein injections that allows delivery of dsRNAs into multiple organs, notably the liver, spleen, lung, kidney, and pancreas.20, 46 However, except for limb delivery, this method is unlikely to be useful in humans. For in vivo delivery, cationic liposomes bearing siRNAs have been intravenously injected into mice,47 and electroporation has been used to deliver siRNAs and short hairpin RNAs (shRNAs) to postimplantation embryos and postnatal retinas.48, 49, 50, 51 Direct injection into the portal vein with lipiodol29 may be feasible in humans, although injection volumes were still quite large as a percentage of total blood volume. In addition, siRNAs have been successfully delivered via intranasal delivery to the lungs.52 One recent report examined direct application of siRNAs in the rat brain.53 Although siRNAs against the dopamine D1 receptor effectively degraded D1 transcipts in vitro, delivery with a mini-osmotic pump via a cannula into the caudate–putamen failed to induce RNAi in vivo, suggesting that the development of effective delivery systems may be the key barrier to siRNA therapeutics. Overall, unless direct delivery of dsRNA to target tissues proves feasible, many of the same delivery problems that plague traditional forms of gene therapy will still need to be overcome.
In conclusion, siRNAs have become not only an exciting new tool in molecular biology but also the next frontier in molecular medicine. Significant hurdles remain, most notably guaranteeing specificity and finding safe and efficacious delivery systems. While work is ongoing to solve these problems, the therapeutic promise of small RNAs remains great.
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This work was supported by the National Institutes of Health (DK035592 and GM62487). RCCR was supported by NIH 5T32 GM0347.
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Ryther, R., Flynt, A., Phillips, J. et al. siRNA therapeutics: big potential from small RNAs. Gene Ther 12, 5–11 (2005). https://doi.org/10.1038/sj.gt.3302356
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