Advanced Drug Delivery Reviews - diyhpl

Advanced Drug Delivery Reviews 61 (2009) 746–759

Contents lists available at ScienceDirect

Advanced Drug Delivery Reviews

j ourna l homepage: www.e lsev ie r.com/ locate /addr

siRNA vs. shRNA: Similarities and differences☆

Donald D. Rao a, John S. Vorhies a, Neil Senzer a,b,c,d, John Nemunaitis a,b,c,d,⁎a Gradalis, Inc., Dallas, TX, USAb Mary Crowley Cancer Research Centers, Dallas, TX, USAc Texas Oncology PA, USAd Baylor Sammons Cancer Center, Dallas, TX, USA

☆ This review is part of the Advanced Drug Delivery ReTherapeutic Application of RNA-mediated Gene Regulat⁎ Corresponding author.1700 Pacific, Suite 1100, Dallas

658 1964; fax: +1 214 658 1992.E-mail addresses: [email protected] (D.D. Rao),

(J.S. Vorhies), [email protected] (N. Senzer), jn(J. Nemunaitis).

0169-409X/$ – see front matter © 2009 Elsevier B.V. Adoi:10.1016/j.addr.2009.04.004

a b s t r a c t
a r t i c l e i n f o
Article history:Received 23 January 2009Accepted 13 April 2009Available online 20 April 2009

Keywords:RNA interferenceBi-functionalCancerPersonalized

RNA interference (RNAi) is a natural process through which expression of a targeted gene can be knockeddown with high specificity and selectivity. Using available technology and bioinformatics investigators willsoon be able to identify relevant bio molecular tumor network hubs as potential key targets for knockdownapproaches. Methods of mediating the RNAi effect involve small interfering RNA (siRNA), short hairpin RNA(shRNA) and bi-functional shRNA. The simplicity of siRNA manufacturing and transient nature of the effectper dose are optimally suited for certain medical disorders (i.e. viral injections). However, using theendogenous processing machinery, optimized shRNA constructs allow for high potency and sustainableeffects using low copy numbers resulting in less off-target effects, particularly if embedded in a miRNAscaffold. Bi-functional design may further enhance potency and safety of RNAi-based therapeutics. Remainingchallenges include tumor selective delivery vehicles and more complete evaluation of the scope and scale ofoff-target effects. This review will compare siRNA, shRNA and bi-functional shRNA.

© 2009 Elsevier B.V. All rights reserved.

Contents

1. Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 7462. Targeted cancer gene therapy and RNA interference. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 747

2.1. Personalized approach for cancer therapy . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 7472.2. RNA interference for cancer. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 747

3. Small interfering RNA (siRNA) and short hairpin RNA (shRNA) . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 7483.1. siRNA . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 7483.2. shRNA. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 7503.3. Bi-functional shRNA . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 7513.4. Summary of si/sh/bi . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 752

4. Delivery . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 7535. Off-target effects . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 753

5.1. Specific off-target effects . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 7535.2. Nonspecific off-target effects . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 754

6. The future outlook . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 7557. Conclusions . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 755Acknowledgements . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 756References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 756

views theme issue on “Towardsion”., Texas 75201, USA. Tel.:+1214

[email protected]@marycrowley.org

ll rights reserved.

1. Introduction

Cancer is a disease of genes, whether based on aberrant changes insequence or expression (epigenomics). The constellation of geneticand epigenetic abnormalities characterizing cancer cells present newand more specific targets for cancer treatment and, hopefully,prevention. Over the last decade, the mapping of the human genome,

mailto:[email protected]




http://dx.doi.org/10.1016/j.addr.2009.04.004

http://www.sciencedirect.com/science/journal/0169409X

747D.D. Rao et al. / Advanced Drug Delivery Reviews 61 (2009) 746–759

along with improved understanding of signal transduction and thepathways responsible for tumor survival, have been transformingtherapeutic oncology from a more promiscuously targeted che-motherapeutic approach towards a highly selective targeted ther-apeusis. Indeed, in 1997, antibody-based Herceptin (trastuzumab)became the first targeted therapy for breast cancer, specifically forHER2-positive metastatic breast cancer. In 2001, the small moleculeGleevec (imatinib mesylate), became the first approved kinaseinhibitor for cancer targeting bcr-abl in chronic myeloid leukemia(CML), and it has since been approved for the treatment ofgastrointestinal stromal tumors (GIST) targeting c-kit. Over the lastfew years, several other targeted cancer therapies have beenapproved. Targeted therapeutics directed against amplified genesand/or over-expressed proteins in malignant cells have proven to bepowerful tools for cancer treatment. A growing understanding and useof proteomic, genetic, and pharmacogenomic tools are actualizing along desired concept of personalized cancer therapy. Geneticabnormalities of each patient's tumor can be analyzed through avariety of established means to quantitatively determine both geneand protein over- and under-expression. Moreover, functional path-ways can be determined and integrated within the cancer networkallowing for the identification of key molecular relays enablingexperimental testing of target specific therapeutics. Such informationcan potentially allow medical care takers to prioritize, if not yetoptimize, treatment for cancer patients, and to uncover surrogatebiomarkers for prognosis, prediction, and therapy assessment.

The recent discovery of RNA interference (RNAi), a natural processthroughwhich the expression of a targeted gene can be knocked downwith high specificity and selectivity, presents an invaluable tool forpersonalized cancer therapy. Target specific RNAi agents have thepotential to selectively knockdown key abnormally over- or constitu-tively expressed molecular targets that are essential for the survival ofeach patient's tumor for effective personalized cancer treatment.Conceptually, target specific RNAi agents can also be applied incombination with immune modulating agents or small molecules toimprove the efficacy of cancer treatment. Like other new therapeuticparadigms, there are a multitude of issues which need to be addressedin order for us to translate RNA interference technology (siRNA,shRNA, bi-functional RNA) from laboratory to bedside and fromconcept to reality. These issues include comparison of each of the RNAitechnologies with respect to effective delivery, possible off-targeteffects and the pharmacokinetics and pharmacodynamics. This reviewwill focus on several issues currently confronting clinical developmentof RNAi therapeutics. We will discuss the role of RNAi technology inpersonalized cancer gene therapy and address clinical considerationsof appropriate RNAi-based therapeutic agents for cancer.

2. Targeted cancer gene therapy and RNA interference

2.1. Personalized approach for cancer therapy

Most human tumors manifest gene expression patterns that differnot only from their normal counterparts but, to a lesser extent, evenfrom each other based on both intrinsic gene modifications andmodulated cancer cell–matrix interactions. Such variability in geneticpatterns found between histologically identical tumors arising indifferent patients may well explain the widely divergent responses tothe standard treatment regimens most often prescribed for a particulartumor type. This is supported by recent studies demonstrating thatcertain patterns of genetic expression (i.e. expression signatures)identified in tumor samples from patients with breast cancer are notonly strongly correlated with prognosis [1–4] but can actually be sub-classified into differing prognostic categories [5]. The presence offunctional redundancy in a robust, predominantly scale-free networksuch as cancer “buffers” the effect of any single gene/targetmodificationon themalignant process, with rare exception (e.g., CML). The hierarchy

of cancer scale-free networks does not have a threshold for single targetdisintegration insofar as random pathway component failure predomi-nantly affects targets with lowconnectivitywithin the network, therebyhaving limited functional impact. However, highly connected informa-tion-transfer targets do allow for “attack vulnerability.” In other words,the disordered circuitry characteristic ofmalignancy results in a change,such that the otherwise robust oncogenic process can become, almostparadoxically, more highly dependent on a specific rewired pathway(i.e., “pathway addiction”). Conceptually, knockout of those “rewired”tumor specific oncogenic pathways should produce a lethal effect oncancer cells yet not significantly perturb normal cell functionality.Genetic diversity of cancer, pathway addiction and targeted therapy arenot the subject of this review and have been extensively reviewedelsewhere [6–8]; here, we discuss how RNAi-based therapeutics can bebest applied in light of these mechanisms.

For example, we harvested tumor and normal cells from the lymphnodes of a melanoma cancer patient for molecular profiling bymicroarray and proteomic analysis [9]. Expression profiles formalignant versus normal tissue were compared at the mRNA andprotein levels. The goal was to identify a group of gene and proteindoublets differentially over-expressed in that individual's malignancy.Fig. 1 is a comparative analysis of protein expression profile by thetwo-dimensional difference gel electrophoresis (2D-DIGE) analysis.2D-DIGE can very effectively identify several over-expressed proteinsin the patient's tumor. Correlated DNA/RNA over-expression can thenbe confirmed with microarray data. The resulting data is furtheranalyzed by a modeling and simulation computational systemdeveloped by our team specifically for clinical application including,but not limited to, gene set enrichment analysis and networkinference modeling platforms (Fig. 2). Grouped gene expressionpatterns that are highly correlated with the pathway phenotype in anindividual patient allow target genes to be selected, and prioritizedbased on connectivity and vector-driven criteria developed analyticnetwork algorithms. Once this individual “cancer fingerprint” iscreated, it serves as the template for the design, synthesis, andsubsequent validation of individualized therapeutic RNAi moleculeswith knockdown activity against these “high-degree hub” genes for apersonalized and targeted cancer gene therapy.

2.2. RNA interference for cancer

The concept of antisense oligodeoxynucleotides as modulators ofgene expression and their application in targeted cancer gene therapywas developedmore than 25 years ago (for review, [10]). By processesstill unknown, the antisense nucleic acid (ASNA) strand and themRNAtarget come into proximity leading to the destruction of the mRNAtarget either by endogenous nucleases, such as RNase H [11,12] thatare recruited into the mRNA–ASNA duplex or by intrinsic enzymaticactivity engineered into the ASNA sequence, as is the case withribozymes [13,14] and DNAzymes [15,16]. The discovery of anevolutionarily conserved gene silencing mechanism whereby smallsequences of extrinsic dsRNA or intrinsic microRNA inhibit comple-mentary post-transcriptional mRNA (siRNA) or suppress translation(miRNA), respectively, ignited strong hope that the natural genesilencing process would be specific and robust. The silencing processoccurs following interaction of the RNA effector precursors with theRNase III enzymes Drosha (for miRNA) and Dicer (for miRNA andsiRNA) and subsequent formation of the RNA-interfering silencingcomplex (RISC) [17]. Endonucleolytic cleavage of the target mRNAoccurs at a single site ~10 nucleotides from the 5′ end of the guide(antisense) siRNA sequence [18,19]. RNAi offers several advantagesover antisense and ribozyme approaches, including ease of synthesis[20] and greater activity [18,21–24].

Preclinical studies confirm that RNAi techniques can be used tosilence cancer-related targets [25–35]. In vivo studies have also shownfavorable outcomes by RNAi targeting of components critical for

Fig. 1. Analysis of the 2-D DIGE images by DeCyder Software and mass spectrometry protein identification. The upper panel shows on the left the protein expression pattern of anormal lymph node from patient RW following 2-D gel electrophoresis. In themiddle of the upper panel the protein expression pattern of amalignant lymph node from patient RW isshown and to the right is an overlaid imagewith proteins from the normal lymph node labeled with Cy3 (green) and proteins from the malignant lymph node labeled with Cy5 (red).Circles indicate protein spots with significant expression level changes. The upper right panel shows the fold-of-change distribution curve and the level of change for the spots ofinterest. Themiddle panel shows the 3D view of one protein spot change between the normal andmalignant lymph nodes. Mass spectrometry (lower panel) subsequently (followingrobotic spot picking) identified this protein as RACK1.

748 D.D. Rao et al. / Advanced Drug Delivery Reviews 61 (2009) 746–759

tumor cell growth [26,36–39], metastasis [40–42], angiogenesis[43,44], and chemoresistance [45–47].

3. Small interfering RNA (siRNA) and short hairpin RNA (shRNA)

The applications of RNAi can be mediated through two types ofmolecules; the chemically synthesized double-stranded small interfer-ing RNA (siRNA) or vector based short hairpin RNA (shRNA). EffectiveRNAi was initially demonstrated by the application of synthetic siRNA[48]; later, siRNA produced in vitro by T7 RNA polymerase was found tobe active and it was soon demonstrated that active siRNA consists of ahairpin structure can be transcribed in cells from an RNA polymerase IIIpromoter on a plasmid construct [49,50]. Although siRNA and shRNAcan be applied to achieve similar functional outcomes, siRNA and shRNAare intrinsically different molecules. Therefore, the molecular mechan-isms of action, the RNA interference pathways, the off-target effects andthe applications can also be different.

3.1. siRNA

Fluorescent labeled siRNA has been used to trace the fate ofdelivered siRNA. A fluorescent label can either be tagged onto the 5′

end or the 3′ end of the siRNA. Using a fluorescence resonance energytransfer (FRET)-based visualization method, the intact siRNA can beobserved to be translocated into the nucleus within 15 min of thedelivery and then disseminated into the cytoplasmwithin the next 4 hboth in intact and dissociated form [51]. The initial accumulation ofsiRNA in the nuclei is similar to observations made on the behavior ofantisense olignucleotides [52]. The translocation of antisense oligo-nucleotide into the nuclei was not dependent on either the ATP pool ortemperature and thus may not involve the active import transportsystem of the nuclear pore [52]; it is not clear whether siRNAtranslocate into the nuclei using the same mechanism as antisenseolignucleotides. Using HeLa cells and targeting 7SK snRNA which isexclusively located in the nucleus, Robb showed the efficiency ofsiRNA mediated silencing to be greater than that effected by antisense7SK DNA [53]. Berezhna et al. observed nuclear localization of siRNAtargeted against small nuclear RNA (snRNA) and cytoplasmiclocalization of siRNA targeted against viral mRNA suggesting selectivelocalization and compartmentalization of siRNA based on its intendedtarget [54]. Ago1 and Ago2 containing RISC were found both in thecytoplasm and nucleus [55–57]. A recent study using fluorescencecorrelation spectroscopy and fluorescence cross-correlation spectro-scopy (FCS/FCCS) to correlate the presence of siRNA with Ago2

Fig. 2. (A) Nearest neighbor protein–protein (first order) interactions of the 6 prioritized proteins in VisualCell. Second order interactions for Stathmin1 (SDCBP) (B) and RACK1(GNB2L1) (C).


protein, indicated a shuttling of RISC between nucleus and cytoplasm[58]. Importin 8 (Imp8) binds to all Ago proteins in a Ran-dependentmanner, but independently of RNA [59]. Knockdown of Imp8 results ina shift of Ago2 from the nucleus to the cytoplasmwithout affecting thetotal quantity of Ago2. However, although Imp8 is not required fortarget mRNA cleavage it is necessary for Ago2 binding to miRNAtargets. In Caenorhabditis elegans, the argonaute protein NRDE-3 isessential for binding nuclear RNAs and appears to interact withcytoplasmic siRNAs generated by RNA-dependent RNA polymerase(RdRP) followed by redistribution to the nucleus [60]. The nuclearRISC (nRISC) is a complex that is 20× smaller in size than the cyto-plasmic RISC (cRISC). The nucleus may be the check point controllingdistribution as either the nuclear acting siRNA or the cytoplasmicacting siRNA.

Dynamically, siRNA steadily increases its accumulation in cells for4 h before plateau [61]. The steady-state nuclear distribution of siRNAwas mainly found in the nucleolus region and was excluded from thenucleoplasm [62]. The cytoplasmic distribution of siRNA appears to bein the perinuclear region forming a ring-like pattern around thenucleus [63]. The nucleolus and perinuclear regions are possibly themain site for RNAi. However, Ohrt et al. labeled siRNAwith fluorescentdye at the 3′ end of either strand of siRNA and did not findaccumulation of siRNA at the perinuclear region, but rather evenlydistributed throughout the cytoplasm [62]. This discrepancy may bethe result of the fluorescent tagging process. At 48 h post injection, themajority of siRNA appears to have been degraded with only 1%fluorescence remaining in the cell. The spatial and temporal distribu-tion of siRNA within the cell is in accord with the observed kinetics of

siRNA mediated RNA interference activity which peaks around 24 hpost delivery and diminishes within 48 h.

The life-cycle of siRNA inside transfected cells is diagrammaticallyillustrated in Fig. 3. In Drosophila, double-stranded RNA-bindingproteins (dsRBPs), such as R2D2 and Loquacious (Loqs), function intandem with Dicer (Dcr) enzymes in RNA interference (RNAi) [64–66]. Dcr-1/Loqs and Dcr-2/R2D2 complexes generate microRNAs(miRNAs) and small interfering RNAs (siRNAs), respectively. Thus,Loqs and R2D2 represent two distinct functional modes for dsRBPs inthe RNAi pathways [67]. In mammalian cells, only one Dicer gene hasthus far been identified [68]. Human Dicer is an integral component ofthe RNA interference pathway. Dicer processes pre-microRNA anddouble-strand RNA (dsRNA) to mature miRNA and siRNA, respec-tively, and transfers the processed products to the RISC [69,70]. Sincethere is only one Dicer in the human, the RNA-interfering pathway forsiRNA and for miRNA may not be as compartmentalized as forDrosophila. Dicer is a multi-domain RNase III-related endonucleaseresponsible for processing dsRNA to siRNAs [71]. Dicer preferentiallybinds to the 5′ phosphate of 2 nt 3′ overhang and cleaves dsRNAs into21 to 22 nucleotide siRNAs [72,73]. Mammalian Dicer interacts withthe double-stranded Tat–RNA-binding protein (TRBP) or PACT (PKRactivating protein) to mediate RNA interference and miRNA proces-sing. TRBP and PACT are structurally related but exert oppositeregulatory activities on RNA-dependent protein kinase (PKR). Knock-down of both TRBP and PACT in cultured cells leads to significantinhibition of gene silencing mediated by short hairpin RNA but not bysiRNA, suggesting that TRBP and PACT function primarily at the step ofsiRNA production [74]. Human TRBP and PACT directly interact with

Fig. 3. Schematic of the siRNA mediated RNA interference pathway. After entry into the cytoplasm, siRNA is either loaded onto RISC directly or utilize a Dicer mediated process. AfterRISC loading, the passenger strand departs, thereby commencing the RNA interference process via target mRNA cleavage and degradation. siRNA loaded RISCs are also found to beassociated with nucleolus region and maybe shuttled in and out of nucleus through an yet unidentified process.


each other and associate with Dicer to stimulate the cleavage ofdouble-stranded or short hairpin RNA to siRNA [74]. Dicer knockout EScells can effectively load processed siRNA onto RISC and carry out RNAinterference as efficiently as Dicer+ ES cells [68]. So, it appears that inmammalian cells, a perfectly processed siRNA can be effectivelyloaded onto RISC for RNAi without the help of the TRBP/PACT/Dicercomplex. The TRBP/PACT/Dicer complex, however, is required toprocess either shRNA or long dsRNA to appropriate size and form fortheir loading onto RISC.

Duplex siRNA in association with holo-RISC, composed of at leastAgo-2, Dicer and TRBP, is identified as the RISC loading complex (RLC)[75]. In the RLC, the two strands of the duplex are separated, resulting inthe departure of the passenger strand [76–78]. The passenger strand iscleaved by the RNase-H like activity of Ago-2, provided there arethermodynamically favorable conditions for passenger strand depar-ture. This is referred to as the cleavage-dependent pathway [79]. There isalso a cleavage-independent by-pass pathway, in which the passengerstrand with mismatches is induced to unwind and depart by an ATPdependent helicase activity [76,79,80]. The RISC with single-strandedguide strand siRNA is then able to execute multiple rounds of RNAinterference. ATP is not required for shRNA processing, RISC assembly,cleavage-dependent pathway, or multiple rounds of target-RNAcleavage [81–83]. Single-stranded siRNA (containing 5′-phosphates)and pre-miRNA can be loaded on RISC, but not duplex siRNA [84].

3.2. shRNA

shRNAs, as opposed to siRNAs, are synthesized in the nucleus ofcells, further processed and transported to the cytoplasm, and thenincorporated into the RISC for activity [85]. The life-cycle of shRNAinside of transfected cells is diagrammatically illustrated in Fig. 4. Tobe effective, the shRNA are designed to follow the rules predicated bythe specifics of the cellular machinery and are presumably processedsimilar to the microRNA maturation pathways. Thus, studies on thesynthesis and maturation of miRNAs have provided the groundwork

for the synthesis of shRNA [86], particularly the miR-30 based shRNAs[87].

shRNA can be transcribed by either RNA polymerase II or IIIthrough RNA polymerase II or III promoters on the expression cassette.The primary transcript generated from RNA polymerase II promotercontains a hairpin like stem-loop structure that is processed in thenucleus by a complex containing the RNase III enzyme Drosha andthe double-stranded RNA-binding domain protein DGCR8 [88]. Thecomplex measures the hairpin and allows precise processing ofthe long primary transcripts into individual shRNAs with a 2 nt 3′overhang [89]. The processed primary transcript is the pre-shRNAmolecule. It is transported to the cytoplasm by exportin 5, a Ran-GTP-dependent mechanism [90,91]. In the cytoplasm the pre-shRNA isloaded onto another RNase III complex containing the RNase IIIenzyme Dicer and TRBP/PACT where the loop of the hairpin isprocessed off to form a double-stranded siRNAwith 2 nt 3′ overhangs[92–94]. The Dicer containing complex then coordinates loading ontothe Ago2 protein containing RISC as described earlier for siRNA. Pre-shRNA has been found to be part of the RLC; thus, pre-shRNA maypotentially directly associate with RLC rather than through a two stepsprocess via a different Dicer/TRBP/PACT complex [95].

After loading onto RLC and passenger strand departure; both siRNAand shRNA in the RISC, in principle, should behave the same. Theargonaute family of proteins is the major component of RISC [96,97].Within the Argonaute family of proteins, only Ago2 contains theendonuclease activity necessary to cleave and release the passengerstrand of the double-stranded stem [76,77,79]. The remaining threemembers of Argonaute family, Ago1, Ago3 and Ago4, which do nothave identifiable endonuclease activity, are also assembled into RISCand presumably function through a cleavage-independent manner.Thus, RISC can be further classified as cleavage-dependent andcleavage-independent [79].

The argonaute family of proteins in RISCs are not only involved in theloading of siRNA or miRNA, but also implicated in both transcriptional(targeting heterochromatin) and post-transcriptional gene silencing.Ago protein complexes loaded with passenger strandless siRNA or

Fig. 4. Schematic of the shRNA mediated RNA interference pathway. After delivery of the shRNA expression vector into the cytoplasm, the vector needs to be transported into thenucleus for transcription. The primary transcripts (pre-shRNA) follow a similar route as discovered for the primary transcripts of microRNA. The primary transcripts are processed bythe Drosha/DGCR8 complex and form pre shRNAs. Pre-shRNAs are transported to the cytoplasm via exportin 5, to be loaded onto the Dicer/TRBP/PACT complex where they arefurther processed to mature shRNA. Mature shRNA in the Dicer/TRBP/PACT complex are associated with Argonaute protein containing RISC and provide RNA interference functioneither through mRNA cleavage and degradation, or through translational suppression via p-bodies.


miRNA seeks out complementary target sites in mRNAs, whereendonucleolytically active Ago-2 cleaves mRNA to initiate mRNAdegradation [98,99]. Other Ago protein containing complexes withoutendonucleolytic activity predominantly bind to partially complemen-tary target sites located at the 3′ UTR for translation repression throughmRNA sequestration in processing bodies (p-bodies) [100–102]. Thedetailed mechanism of mRNA sequestration in p-bodies and laterrelease fromp-bodies is still a debated issue; deadenylation of the targetmRNAwhich leads to destabilization of the mRNAwas also observed tooccur in p-bodies [103,104]. Coimmunoprecipitation experimentsdetermined that RISCs are also strongly associated with polyribosomesor the small subunit ribosomes [95] and Ago-2 (actually identified aselF2c2), strongly suggesting that RISC surveillance is compartmenta-lized with translational machinery of the cell. Details of the mechanisminvolving mRNA scanning and target mRNA identification are stilllargely unknown. Whatever the scanning or surveillance mechanismmay be, once the target mRNA is identified, the target mRNA is eithercleaved or conformationally changed following which both types ofstructures are routed to the p-body for either sequestration ordegradation [103,104]. The active siRNA or miRNA loaded complex isthen released for additional rounds of gene silencing activity.

3.3. Bi-functional shRNA

There is, however, a third unique RNAi option in developmentcalled bi-functional shRNA. shRNA can potentially be manipulated totake advantage of the gene silencing machinery within the cells toimprove its efficiency and durability of action. Conceptually, targetedshRNAs can be designed so as to effectively load shRNA onto both thecleavage-dependent and the cleavage-independent RISCs. This differ-ential processing is mediated by two pathways primarily dependenton strand complementarity and/or access to RNase-H cleavage and,presumably, for final target effect, on interaction with Imp8.Simultaneous expression of both types of shRNAs (i.e. the bi-

functional shRNA) in cells should achieve a higher level of efficacy,greater durability compared to siRNA, and a more rapid onset of geneexpression silencing (the rate dependent on mRNA turnover andprotein kinetics) compared to shRNA as illustrated on Fig. 5. The “bi-functional” shRNA, by virtue of loading onto multiple types of RISCs, isthus able to simultaneously induce degradation of target mRNA andalso inhibit translation through mRNA sequestration. This bi-func-tional design should be, in principal, much more efficient for tworeasons; first, the bi-functional will promote loading the guide strandonto at least two types of RISCs to increase activity; second, by loadingonto both cleavage-dependent RISC and cleavage-independent RISC,target mRNA can be silenced both through mRNA degradation andtranslational inhibition or sequestration.

Thedesignof the bi-functional shRNAexpressionunit consists of twostem-loop shRNA structures; one stem-loop structure composed of fullymatched passenger and guide strand for cleavage-dependent RISCloading, the second stem-loop structure composed of mis-matchedpassenger strand (at the positions 9–12) for cleavage-independent RISCloading.

There are several experimental observations that support thisapproach. In Drosophila, Ago1 preferentially binds to miRNAs thathave been excised from imperfectly paired hairpin precursors, whereasthose miRNAs that have near-perfectly paired hairpin precursors arebound by Ago2 [105–108]. In HEK293 cells transfected with tagged-Agoproteins, coimmunoprecipitation found similar sets of about 600transcripts to be bound to Ago1, 2, 3 or 4 [95], suggest all fourmammalian Ago protein containing RISCs are involved in the RNAifunction. Insofar as most mRNA have multiple miRNA target sites (withdistance constraints) at their 3′ UTR, the miRNA mediated RNAi systemappears to be redundant for the targeted mRNAs allowing forcooperative downregulation to ensure target mRNA knockdown. Thebi-functional shRNA approachmimics the natural process by mediatingtarget mRNA knockdown through multiple RNAi pathways andcomplexes.

Fig. 5. Schematic of the bi-functional shRNA concept. The bi-functional concept is to design two shRNAs for each targetedmRNA; onewith perfect match, one with mismatches at thecentral location (bases 9–12). The purpose of the bi-functional design is to promote loading ofmature shRNAs onto both cleavage-dependent and cleavage-independent RISCs, so thatthe expression of target mRNA can be more effectively and efficiently shut down both through target mRNA degradation and through translational repression.


In C. elegans, structural features of small RNA precursors determineArgonaute loading [109]. Recently, Azuma-Mukai et al. observedmiRNAs associated with hAgo-2 and hAgo-3 have some overlaps;however, some are discriminately loaded onto hAgo-2 or hAgo-3[110]. Further work is needed to resolve the specificity of miRNAloading onto different Ago containing RISCs. Although most miRNAtarget sites have been identified to be located at the 3′-UTR region,recent systemic identification of mRNAs recruited to hAgo-2 haveidentified many mRNAs with target sites located at the coding regionand some at the 5′-UTR [111]. hAgo-2 could initiate the target mRNAdegradation with its slicing activity in the coding region. Tay et al.recently found many naturally occurring miRNA targets are located inthe coding region of embryonic regulated genes to modulateembryonic stem cell differentiation [112], further support thatmiRNA can act through mRNA regions other than 3′-UTR.

3.4. Summary of si/sh/bi

In summary, exogenously introduced siRNA with appropriatelength (19–21 nt) and 2 nt 3′ overhang, can be loaded onto RISC forRNAi function without interacting with either Dicer, TRBP or PACT;however, the loading process is10× less efficient than shRNA.Increasing the length of the siRNA duplex to 29–30 nt with a 2 nt 3′overhang only at one end of the duplex (specifically antisense [113])appears to improve efficacy [114]. If so, it is possibly becauseincreasing the length of an siRNA duplex with an unprocessed endforces directionality as a result of imposed thermodynamic instabilitydetermining the guide strand motif and thereby enhancing itsassociation with Dicer/TRBP/PACT complex for more efficient loadingonto RLC [114]. shRNA, on the other hand, assimilates into theendogenous miRNA pathway and in so doing is significantly moreefficient [115–117]. Additionally, fluorescent tagged siRNA tracingindicated high degradation and turnover of exogenously introducedsiRNA. Less than 1% of the introduced duplex remains in the cell 48 hafter administration. shRNA can be continuously synthesized by thehost cell, therefore, its effect should be much more durable.Concentrations necessary for effective knockdown are usually in the

low nM range for most siRNAs, while less than 5 copies of shRNAintegrated in the host genome is sufficient to provide continual geneknockdown effect (Cleary M, personal communication). The higherdose required for siRNA can further contribute to the off-target effectsto be discussed later.

Understanding the mechanism and the dynamics of siRNA andshRNA at the cellular and molecular level greatly enhanced the effortin developing therapeutic siRNAs for various diseases. As a result,modifications can be made to improve the efficacy and stability ofRNAi agents. Chemically synthesized siRNA is easier to modifythrough chemistry; however, bulk manufacturing of complex struc-tures such as modified siRNA is more expensive. Vector based shRNArelies on the host machinery for expression, but, on the other hand, ismore difficult to modify. Modification can only be achieved throughmanipulating expression strategy (e.g. bi-functional shRNA), rede-signing shRNA structure, or by varying promoter regulation. TheshRNA expression units can be incorporated into varieties of plasmidsand viral vectors for delivery and integration. In addition, vector basedshRNA expression can also be regulated or induced [118–120].

Numerous siRNAs have been demonstrated to be effective for in-vivo tumor growth modulations via intratumoral, ex-vivo, or systemicroutes of application (For review see [121–123]. Vector based shRNAhas, likewise, demonstrated in-vivo effectiveness (For review see[121,123,124]. In-vivo studies employ a variety of delivery methodsand may not ensure equivalency of strand biasing; therefore, it is hardto perform direct comparison between siRNA and shRNA. McAnuff etal., using a luciferase expression system, compared the potency ofsiRNA versus shRNA mediated knockdown in vivo; they found thatsiRNA and shRNA are equivalent in potency at 10 mg dose; however,on a molar basis, the shRNA was 250 fold more effective than thesiRNA [115]. In an effort to assess the potential of RNAi as a therapeuticfor hepatitis C (HCV), siRNANS5B was targeted against the non-structural protein 5B viral polymerase coding region fused withluciferase gene [125]. Luciferase expression in vivo was reduced by75%. Using a cognate shRNANS5B produced a 92.8% average inhibitionover 3 experiments. McCleary et al. incubated shRNA, directed againstfirefly luciferase and containing 29 mer stems and 2-nt 3′ overhangs,


with recombinant human Dicer [116]. The resulting 22-nt shRNAproducts, with predictable guide strands, were compared to identi-cally targeted 21 mer siRNA in HeLa cells. More effective inhibitionwas seenwith the shRNA. In another study of the feasibility of RNAi fortreatment of HCV, 19 and 25 bp shRNAs were compared with 19 and25 bp siRNA directed against the HCV IRES (internal ribosomal entrysite) using a luciferase reporter in the AVA5 cell line with stableexpression of the HCV subgenomic genotype 1b replicon [117]. The19 bp shRNAs were more potent than either the 19- or 25-bp siRNAs.

A recent paper evaluated the levels of Dicer and Drosha in cell linesand in tumor samples from patients with ovarian cancer [126]. Thedistribution of Dicer mRNA levels was bimodal and 60% of specimenshad decreased Dicer mRNA. Furthermore, low Dicer levels were foundto be a predictor of reduced disease-specific survival in multivariateanalysis. Of particular interest was the finding that, compared to siRNAmediated silencing of the galectin-3 gene, poor silencing was achievedwith shRNA in ovarian cancer cell lines with low versus high Dicerexpression. These data will need to be confirmed and evaluatedfurther. Although there is a global downregulation of miRNAexpression in cancer [127], whether this is, in large part, due to lowDicer expression or to shortened 3′ UTRs with fewer miRNA-bindingsites [128] in highly proliferating tumors with modulated feedbackmechanisms is not known. miRNA functionality has been confirmed inthe three tumor types (ovary, breast, and lung) evaluated in this studyraising questions regarding both qualitative and quantitative issues.As the authors note, there are data correlating high Dicer expressionwith poor prognostic features in other tumor types [129,130]. In theirretrospective evaluation of lung cancer, the role of let-7 as both aregulator of Dicer and ras and their feedback networks could not beassessed [131,132]. The Dicer levels in the tumors used for functionalassay of gene silencing are not given nor is there confirmation of theequivalency of strand biasing between the siRNA and shRNAconstructs.

RNAi therapeutics have been shown to be well tolerated innumerous animal models allowing for transition into the clinic. Atleast 10 RNAi-based drugs are currently in early phase clinical trials[133], two of which are cancer related; one targeted against the M2subunit of ribonucleotide reductase (RRM2) [134] and the othertargeted against tenascin-C [135]. Animal studies with siRNAinhibitors for RRM2 show efficacy [134] and safety in non-humanprimates [136]. shRNA for the treatment of hepatitis B was alsoapproved for clinical trial by FDA.

Both siRNA and shRNA have their respective advantages anddisadvantages from the mechanistic point of view. However, safety ofthis new therapeutic paradigm is of the utmost importance. Althoughno significant adverse event involving initial RNAi-based clinical trialshas been reported, there are concerns over the potential off-targeteffect of RNAi-based agents which will be discussed separately below.

4. Delivery

Efficacy of an RNAi cancer therapeutic is limited by the quantity ofthe oligomer that effectively enters the tumor cells. In the clinicalsetting this is primarily dependent on the method of delivery. An idealdelivery vehicle must be able to selectively and differentially targettumors versus normal tissue, homogeneously distribute through thetumor mass and penetrate the tumor cells following systemicadministration. If cell entry is mediated by endocytosis the deliveryvehicle must negotiate endosomal/lysosomal escape and, in the caseof shRNA, the payload must penetrate the nuclear membrane as well.Viral vectors are popular for laboratory delivery of shRNA because oftheir high transfection efficiency and effective integration of exogen-ous DNA, but they have been losing support in recent years because ofconcerns over safety and immunogenicity [137,138]. Non-viral poly-meric delivery systems, in particular those with biodegradablecomponents, have much better safety profiles than their viral

counterparts though their transfection efficiency is generally lower[139–141]. Non-viral vehicles for delivery of siRNA and shRNA aretypically cationic preparations. Their positive charge facilitatescomplexation with negatively charged nucleic acids and also bindingto the negatively charged glycocalyx on external cell membranespromoting endocytosis. Both tumor targeting and cell entry can beenhanced by decoration or complexation of the vehicle with targetingmoieties, such as monoclonal antibodies, peptides, small moleculeligands, and aptamers to recognize cell surface markers [124,142].Once endocytosed, the vehicle's positive charge facilitates early escapefrom the endosome [143,144]. Though the positive charge of thesevehicles improves their transfection efficiency, it is also associatedwith increased toxicity [140,145].

Awide variety of potential vehicles are being developed to addressthe different issues associated with the delivery of shRNA and siRNA.There are three major classes of non-viral delivery vehicle systems:synthetic polymers, natural/biodegradable polymers, and lipids;many of the vehicles that are showing promise are actually hybridsof these classes. For instance, there is a cyclodextrin-based cationicpolymer which has been used successfully to deliver siRNA targeted toRRM2 in various in vivo cancer models [134,146]. This preparation iscurrently in Phase I clinical trials. Lipid based nanoparticles areshowing potential for the delivery of shRNA and siRNA [147]. ProtivaBiotherapeutics and Alnaylam have developed nanoparticles com-posed of a lipid–PEG conjugate that is capable of encapsulating andprotecting nucleic acids for the purpose of systemic delivery. Thesestable nucleic acid lipid particles (SNALPs) were used in the firstsuccessful administration of siRNAs to a non-human primate[148,149]. Silence Therapeutics has developed a lipid-based deliveryvehicle specifically designed for siRNA delivery to endothelial cells.This vehicle, called AtuPLEX, contains a mix of cationic and fusogeniclipids [150,151]. This vehicle has been used effectively to knockdownprotein kinase N3 in murine prostate and pancreatic cancer models,inhibiting cancer progression [152,153]. More detailed discussions ofdelivery vehicles for shRNA [124,154] and siRNA [141,155–157] as wellas general discussions of organ and tissue specific RNAi delivery[138,158,159] may be found elsewhere.

The magnitude of cytokine induction associated with in vivodelivery of siRNA has been noted to vary widely based on the deliveryvehicle used [160]. The well recognized conundrum in cationic non-viral nucleic delivery is that transfection efficiency usually correlateswith toxicity [161,162]. Effective strategies being pursued to break thiscorrelation include molecular modifications to shield positive chargeand the use of biodegradable polymers [154,163].

5. Off-target effects

Despite initial results showing excellent specificity in RNAimediated gene silencing, over the past 5 years many studies haveshown that there are multiple specific and nonspecific mechanismsthrough which siRNA and shRNA can cause effects other than theintended mRNA suppression. Unintended effects on gene expressionmediated by RNAi are termed “off-target effects.” Specific off-targeteffects are mediated by partial sequence complementarity of the RNAiconstruct to mRNAs other than the intended target. Nonspecific off-target effects include a wide variety of immune and toxicity relatedeffects that are intrinsic to the RNAi construct itself or its deliveryvehicle. The following provides a brief review to the mechanismssurrounding off-target effects and addresses strategies that are beingdeveloped to minimize those effects.

5.1. Specific off-target effects

Expression profiling experiments have shown that partial sequencecomplementarity in the passenger or guide strands of the RNAiconstruct can produce off-target gene suppression. The first group to


definitively demonstrate this used an unbiased genome-wide micro-array profile to search for downregulated mRNA immediately aftertransfecting a variety of different siRNA constructs directed to differentgenes into HeLa cells. This study demonstrated that few off-target geneswere regulated in common. Furthermore, the off-target expressionpatterns observed for each individual siRNA were consistent, withrepeated runs revealing the same off-target expression profile. It wasalso noted that the number and identity of the off-target transcriptswasunrelated to the ability of the siRNA to silence the target gene [164]. Ithas subsequently been shown that both siRNA and shRNA constructswith complementarity in the “seed region” can produce the same off-target expressionprofiles, even across cell lines, independent of deliverymethod [165].

Even limited sequence siRNA:mRNA complementarity with theintended target, which is usually less than optimal, can produce off-target suppression. Off-target silencing effects have been demon-strated in transcripts with complementarity as low as 7 nucleotideswith the guide siRNA strand [166]. Due to a variety of known andunknown mechanisms, not all transcripts with this level of similarityare silenced. The location of the region of complementarity within theRNAi construct and the mRNA transcript are important predictors ofpotential for suppression. Complementarity within nucleotides 2–7 atthe 5′ end of either the siRNA passenger or guide strands has beenshown to be a key determinant in directing off-target effects [167].This region of the construct is reminiscent of the “seed” region withinmiRNA, i.e., a heptameric sequence beginning at the first or secondposition from the 5′ end of the miRNA that is complementary to sitesin the mRNA 3′-UTRs, which guide silencing in endogenous RNAi.Indeed, RNAi off-targeting seems to be mechanistically related to anmiRNA-like effect in that complementarity between the miRNA seedhexamer and the 3′UTR of the off-targeted mRNA produce effectivesuppression of gene expression [103,168,169].

Various in vitro and in silico methods are either available or underdevelopment for analysis of the off-target effects of a given RNAiconstruct. Screening can be accomplished effectively using mRNAexpression data from transfected cells. Expression profiling for off-target screening must be temporally controlled to ensure observationof primary changes in mRNA levels and not secondary changes as aresult of downregulation of target protein expression. Results can thenbe correlated to qRT-PCR data and to protein expression data throughWestern blots. Methods for microarray-based off-target screening arereviewed in detail elsewhere [169,170].

Computational methods are also being developed so that RNAiconstructs may be more effectively screened prior to in vitro testing.Global complementarity search algorithms such as BLASTn andSmith–Waterman have been shown to be poor measures of off-targetpotential in an RNAi construct [168]. Methods for in silico screening ofRNAi constructs must be optimized to consider seed complementarityin the constructs as well as 3′ UTR complementarity in thetranscriptome. Several groups are working to develop algorithmsand some are freely available online [171–174].

The specific off-target effects of a given construct can bemitigated byseveral methods. siRNAs with an asymmetric unilateral 2-nt-overhangon the antisense strand have greater potency than conventional siRNAsas well as reduced off-target effects due to preferential strand selection[113]. Sequence based modifications designed to reduce specific off-target effects are likely to benefit both siRNA and shRNA approaches.Single and double base mismatches between an RNAi construct and itstarget transcript are often tolerated without reducing the potency ofsuppression [168,175–177]. This allows for the optimization of theconstruct sequence to minimize complementarity with 3′UTRs ofunintended targets.

Although the vector-driven shRNA approach to RNAi does notpermit specific chemicalmodification of the silencing construct, siRNAoligomers can be chemically modified in order to reduce direct off-target effects. These modifications must be carefully applied as

reductions in off-target effects are sometimes accompanied bydeceases in the overall potency of suppression. [178–180].

Chemical modifications can also be asymmetrically applied to thepassenger strand of the siRNA construct in order to specifically inhibit itsparticipation in silencing. Annealing of the guide strand to a passengerstrand composed of two segments has been shown to reduce off-targeteffects mediated by passenger strand complementarity [181]. Theaddition of other chemical moieties to one or both strands can alsolimit specific off-target effects. The 5′-phosphorylation status of thesiRNA strands has been shown to be a determinant of which strand isinvolved in silencing, and thus replacement of the 5′ phosphate by of ansiRNA strand with a methyl group has been shown to reduce itsparticipation in silencing [182].

Chemical modifications to nucleotides within the seed region ofthe passenger or guide strand can reduce unintended entry of siRNAconstructs into the endogenous miRNA gene silencing pathway byinhibiting the interaction of the RISC complex with mRNA. It has beenshown that 2′-O-methyl ribosyl substitution at position 2 in the siRNAguide strand can reduce off-target silencing of transcripts withcomplementarity to the seed region of the siRNA guide strand [165].Asymmetric replacement of seed region nucleotides with DNA baseshas also been shown to reduce off-targeting as a result of seed regioncomplementarity within the passenger strand [183].

Recent in vitro studies have shown that shRNA produces fewer off-target effects than siRNA. In one study shRNA and siRNA of the samecore sequence directed towards TP53 were applied to HCT-116 coloncarcinoma cells in concentrations necessary to achieve comparablelevels of target knockdown. Microarray profiling demonstrated amuch higher degree of up- and downregulation of off-targettranscripts in the siRNA transfected cells (M. Mehaffey, T. Ward, andM. Cleary, in prep.). It has been suggested that these differences arisefrom the fact that shRNA is transcribed in the nucleus and is thereforesubject to endogenous processing and regulatory mechanisms.Additionally, siRNA is more susceptible to degradation in thecytoplasm, which may also lead to off-target silencing [184].

5.2. Nonspecific off-target effects

Nonspecific off-target effects are those unintended perturbationsin gene expression not resulting from the direct interaction of an RNAiconstruct with an mRNA transcript. Included in this category areinterferon (IFN) and other immune mediated responses to exogenousRNAi, cellular toxicities due to the nucleotide construct, and effectsrelated to the delivery vehicle. There is strong reason to believe thatshRNA and siRNAwould have very different profiles of nonspecific off-target effects because of the mechanistic differences between the twoapproaches.

Introduction of dsRNA longer than 29–30 bp into mammalian cellsresults in a potent induction of the innate immune system via PKR,similar to the mammalian cell defense mechanism against viralinfection [185]. Activation of the innate immune response by receptorssensitive to exogenous nucleic acids leads to global degradation ofmRNA and thus broad inhibition of translation as well as globalupregulation of IFN-stimulated gene expression. Although shortersiRNA constructs have been shown to avoid receptor activation andwere initially considered to be non-immunogenic [18], subsequent invitro data have shown that the introduction of both synthetic siRNAoligomers and shRNA can induce a partial interferon response [186–188] some of which are sequence dependent (e.g. GU-dependent, 5′-UGUGU-3′ and GU-independent, 5′-GUCCUUCAA-3′) and, therefore,avoidable. Activation of the innate immune system in the case ofexogenous RNAi is likely mediated through several cytoplasmic andendosomal mechanisms attuned to recognize exogenous nucleic acidsfrom infectious agents. The relevant mediators of nucleic acid-stimulated immunoactivation at the level of the endosome are Toll-like receptors (TLRs) 7 and 8 (typically activated by ssRNA), TLR 9 (via


unmethylated CpG activation) and TLR 3 (via dsRNA) [189–193].Immune activation by nucleic acids at the cytoplasmic level ismediated through RNA sensing receptors such as RIG-I and MDA-5[114,194]. Mechanisms surrounding immune activation by RNAi arereviewed more thoroughly elsewhere [160,195].

Recently, naked siRNA has been shown to activate TLR-3 on thesurface of vascular endothelial cells and trigger the release of IFN-γand IL-12 that mediate nonspecific anti-angiogenic effects in-vivo[196]. Immune stimulation often is largely responsible for theobserved therapeutic effects of siRNA rather than the direct targetedeffect [197]. Misinterpreting the therapeutic effect of siRNA needs tobe carefully monitored.

Immune activation at the endosomal level is more readily avoidableby shRNA constructs because the construct is presented on a DNAplasmid obviating dsRNA activation of TLR3.However, TLR 9, as noted, isactivated by unmethylated CpG motifs, which are typically found inbacterial DNA [198,199]. Careful shRNA-encoding plasmid design,avoiding unmethylated CpG motifs, can effectively attenuate if noteliminate TLR 9 mediated endosomal immunoactivation [200].

There is also reason to believe that shRNA is less likely to induce aninflammatory response through cytoplasmic dsRNA receptors in vivobecause shRNA is spliced by endogenous mechanisms. It has beensuggested that the 5′ ends of the endogenous-dicer spliced RNAoligomers are less immunogenic than the 5′ ends of exogenous siRNAoligomers [114,194]. This has been supported in vitro in an experimentthat compared liposome delivered siRNAs versus Pol III promoter—expressed shRNAs of the same sequence in primary CD34+ progenitor—derived hematopoietic cells. In this study it was found that siRNAinduced IFN-alpha and type I IFN genes, while the shRNA of the samesequence did not induce an immune response [201].

Sequence modifications can be made to shRNA or siRNA in order toreduce immunogenicity. It has been shown that endosomal immu-noactivation by siRNA through TLR7 and TLR8 can be sequencedependant [189,190,202]. Some simple sequence modifications, suchas the introduction of G:Umismatches into the sequence also seem tolower the IFN response in vitro [203,204]. One recent study showedthat amarked reduction in the expression of the interferon-stimulatedgene oligoadenylate synthetase 1 (Oas1) could be achieved bymodifying the shRNA to contain features of the naturally occurringmicroRNA-30 (miR-30) precursor [205].

As in the case of specific off-target effects, chemical modification tosiRNA oligomers can make them less immunostimulatory, howeverthese modifications must be fine-tuned so as not to negatively affectthe potency of intended target silencing. Suppression of the TLRmediated immune response has been achieved by substituting the 2′-hydroxyl uridines in the construct with 2′-fluoro, 2′-deoxy, or 2′-O-methyl uridines [206–208], the products of which do retain target-silencing potency while reducing immunogenicity.

Usage of endogenous processing systems gives shRNA an advan-tage over siRNA in terms of its propensity for induction of IFN but itsover-saturation of these systems has been shown to have otherconsequences that are more easily avoided by siRNA. In a key in vivostudy of the safety effects of long term expression of shRNA in thelivers of adult mice, a type 8 adeno-associated virus with a Pol IIIpromoter was used to drive expression of 49 different shRNAs ofdifferent lengths and sequences directed against six targets. 36 of theconstructs tested resulted in a dose dependant liver injury that wasdetermined to be associated with the downregulation of criticalendogenous miRNAs. The degree of miRNA downregulation wasrelated neither to the shRNA sequence nor to the degree ofdownregulation of the target mRNA [209]. Subsequent transfectionstudies suggested the degree of miRNA downregulation to result froma competitive bottleneck in shared miRNA/shRNA processing, mostlikely at the level of exportin-5 (the nuclear membrane export proteinused to transfer pre-miRNAs into the cytoplasm) and the RISCcomponent, Argonaute-2 [210]. A similar in vivo study of hepatic

toxicity of siRNAs in mice and hamsters showed that systemicintroduction of synthetic siRNAs does not result in the suppressionof endogenous miRNA levels. In this study siRNAs targeting twohepatocyte-specific genes (apolipoprotein B and factor VII) and ascramble control were administered to mice and one siRNA targetingthe hepatocyte-expressed gene Scap was administered to hamsters.Robust suppression of target genes was achieved in all cases. Nochanges in levels of the hepatocyte endogenously expressed miR-122were observed in the mice or the hamsters and no changes in thebroadly expressed miRNAs, miR-16 and let-7a, were observed in themice [211].

Though siRNA likely does not compete with endogenous miRNAfor processing proteins, care must be taken when using shRNA as aneffector of RNAi in order to minimize the potential for damagemediated by over-saturation of exportin-5. In one in vitro study, over-expression of the exportin-5 protein has been shown to eliminate thenuclear export bottleneck and allow cells to tolerate higher dosages ofshRNA without toxicity [212]. Another study showed that an adeno-associated virus construct using a Pol-II promoter was able to achievestable target gene suppression at high shRNA doses for over 1 yearafter the initial dosing [213]. In order to minimize the risk of toxicityfrom over-saturation of miRNA professing systems, data to datesuggest both selective promoter integration (e.g. pol II versus pol III)and limiting the dosage of shRNA so as to stay below the threshold ofcompetitive inhibition of the endogenous miRNA biogenesismachinery.

6. The future outlook

The ability to precisely and differentially target functionally bio-relevant molecular signals in patient's cancers will establish a newparadigm in cancer management; one which focuses on defining theuniqueness of each patient's tumor and tumor-host processes andinteractions following rational target prioritization using computa-tional systems biology algorithms. This, then, would allow forexploitation of the “attack vulnerability” of the rewired cancernetwork by deconstructing essential hubs and linkages, multiplytargeting and eliminating them. Both siRNA and shRNA effectors areattractive opportunities. The capability of potentiating activity using abi-functional design may further enhance safety and efficacy. Thesimplicity of siRNA manufacturing and the transient nature of theeffect per dose may be optimal for certain medical disorders in whichhigh vector doses are required, e.g. some of the viral infections,however, by using the endogenous processing machinery, optimizedshRNA constructs allow for high potency sustainable effects usinglow-copy numbers resulting in less off-target effects (particularly ifembedded in an miRNA scaffold) thereby ensuring greater safety.Though shRNA seems ideal for cancer-related therapeutic develop-ment, new technology such as bi-functional RNA interference mayprovide an even greater opportunity for enhancement in potency aswell as heightening safety thereby increasing the opportunities formultiple target therapy. This, of course, is contingent on optimizationof delivery and minimization of off-target effect which will need to beestablished through early clinical testing.

7. Conclusions

Our understanding and application of RNAi has dramaticallyadvanced over the last 5 years. Despite limitations in developingeffective delivery vehicles and concerns regarding potential off-targetactivity, clinical development has been initiated. As the science of thisfledgling technology advances, it is evident that issues such as targetselection, effector potency, delivery vehicle design, and off-targeteffects will continue to be addressed and resolved. Bi-functional RNAiproducts are evolving components in this transition to clinicallyeffective and safer therapeutics.


Acknowledgements

The authors would like to acknowledge Brenda Marr and SusanMill for their competent and knowledgeable assistance in thepreparation of this manuscript, and M. Cleary for providing the databefore publication.

References

[1] V.G. Kaklamani, W.J. Gradishar, Gene expression in breast cancer, Curr. Treatm.Opt. Oncol. 7 (2006) 123–128.

[2] L. Marchionni, R.F. Wilson, S.S. Marinopoulos, A.C. Wolff, G. Parmigiani, E.B. Bass,S.N. Goodman, Impact of gene expression profiling tests on breast canceroutcomes, Evid. Rep./Technol. Assess. (Full Rep.) (2007) 1–105.

[3] J.K. Habermann, J. Doering, S. Hautaniemi, U.J. Roblick, N.K. Bundgen, D. Nicorici, U.Kronenwett, S. Rathnagiriswaran, R.K.Mettu,Y.Ma, S. Kruger,H.P. Bruch,G.Auer, N.L.Guo, T. Ried, The gene expression signature of genomic instability in breast cancer isan independent predictor of clinical outcome, Int. J. Cancer. 124 (7) (2009 Apr 1)1552–1564.

[4] P. Wirapati, C. Sotiriou, S. Kunkel, P. Farmer, S. Pradervand, B. Haibe-Kains, C.Desmedt, M. Ignatiadis, T. Sengstag, F. Schutz, D.R. Goldstein, M. Piccart, M.Delorenzi, Meta-analysis of gene expression profiles in breast cancer: toward aunified understanding of breast cancer subtyping and prognosis signatures,Breast Cancer Res. 10 (2008) R65.

[5] T. Sorlie, R. Tibshirani, J. Parker, T. Hastie, J.S. Marron, A. Nobel, S. Deng, H. Johnsen,R. Pesich, S. Geisler, J. Demeter, C.M. Perou, P.E. Lonning, P.O. Brown, A.L. Borresen-Dale, D. Botstein, Repeated observation of breast tumor subtypes in independentgene expression data sets, Proc. Natl. Acad. Sci. U. S. A. 100 (2003) 8418–8423.

[6] M. Davies, B. Hennessy, G.B. Mills, Point mutations of protein kinases andindividualised cancer therapy, Expert Opin. Pharmacother. 7 (2006) 2243–2261.

[7] A.G. Letai, Diagnosing and exploiting cancer's addiction to blocks in apoptosis,Nat. Rev., Cancer 8 (2008) 121–132.

[8] O.I. Olopade, T.A. Grushko, R. Nanda, D. Huo, Advances in breast cancer: pathwaysto personalized medicine, Clin. Cancer Res. 14 (2008) 7988–7999.

[9] J. Nemunaitis, N. Senzer, I. Khalil, Y. Shen, P. Kumar, A. Tong, J. Kuhn, J. Lamont, M.Nemunaitis, D. Rao, Y.A. Zhang, Y. Zhou, J. Vorhies, P. Maples, C. Hill, D. Shanahan,Proof concept for clinical justification of network mapping for personalizedcancer therapeutics, Cancer Gene Ther. 14 (2007) 686–695.

[10] C.A. Stein, J.S. Cohen, Oligodeoxynucleotides as inhibitors of gene expression: areview, Cancer Res. 48 (1988) 2659–2668.

[11] E. Zamaratski, P.I. Pradeepkumar, J. Chattopadhyaya, A critical survey of thestructure–function of the antisense oligo/RNA heteroduplex as substrate forRNase H, J. Biochem. Biophys. Methods 48 (2001) 189–208.

[12] S.T. Crooke, Molecular mechanisms of action of antisense drugs, Biochim.Biophys. Acta 1489 (1999) 31–44.

[13] D. Castanotto, M. Scherr, J.J. Rossi, Intracellular expression and function ofantisense catalytic RNAs, Methods Enzymol. 313 (2000) 401–420.

[14] J.J. Rossi, Ribozymes in the nucleolus, Science 285 (1999) 1685.[15] S.W. Santoro, G.F. Joyce, A general purpose RNA-cleaving DNA enzyme, Proc. Natl.

Acad. Sci. U. S. A. 94 (1997) 4262–4266.[16] Y. Wu, L. Yu, R. McMahon, J.J. Rossi, S.J. Forman, D.S. Snyder, Inhibition of bcr-abl

oncogene expression by novel deoxyribozymes (DNAzymes), Hum. Gene Ther. 10(1999) 2847–2857.

[17] T.E. Ichim,M. Li, H. Qian, I.A. Popov, K. Rycerz, X. Zheng, D.White, R. Zhong,W.P.Min,RNA interference: a potent tool for gene-specific therapeutics, Am. J. Transplant.4 (2004) 1227–1236.

[18] S.M. Elbashir, J. Harborth, W. Lendeckel, A. Yalcin, K. Weber, T. Tuschl, Duplexes of21-nucleotide RNAs mediate RNA interference in cultured mammalian cells,Nature 411 (2001) 494–498.

[19] J. Liu, M.A. Carmell, F.V. Rivas, C.G. Marsden, J.M. Thomson, J.J. Song, S.M.Hammond, L. Joshua-Tor, G.J. Hannon, Argonaute2 is the catalytic engine ofmammalian RNAi, Science 305 (2004) 1437–1441.

[20] Z. Paroo, D.R. Corey, Challenges for RNAi in vivo, Trends Biotechnol. 22 (2004)390–394.

[21] Y. Aoki, D.P. Cioca, H. Oidaira, J. Kamiya, K. Kiyosawa, RNA interference may bemore potent than antisense RNA in human cancer cell lines, Clin. Exp. Pharmacol.Physiol. 30 (2003) 96–102.

[22] B.L. Bass, RNA interference. The short answer, Nature 411 (2001) 428–429.[23] S. Coma, V. Noe, C. Lavarino, J. Adan, M. Rivas, M. Lopez-Matas, R. Pagan, F.

Mitjans, S. Vilaro, J. Piulats, C.J. Ciudad, Use of siRNAs and antisense oligonucleo-tides against survivin RNA to inhibit steps leading to tumor angiogenesis,Oligonucleotides 14 (2004) 100–113.

[24] M. Miyagishi, M. Hayashi, K. Taira, Comparison of the suppressive effects ofantisense oligonucleotides and siRNAs directed against the same targets inmammalian cells, Antisense Nucleic Acid Drug Dev. 13 (2003) 1–7.

[25] M. Scherr, K. Battmer, T.Winkler,O.Heidenreich,A.Ganser,M. Eder, Specific inhibitionof bcr-abl gene expression by small interfering RNA, Blood 101 (2003) 1566–1569.

[26] T.R. Brummelkamp, R. Bernards, R. Agami, Stable suppression of tumorigenicityby virus-mediated RNA interference, Cancer Cells 2 (2002) 243–247.

[27] L.A. Martinez, I. Naguibneva, H. Lehrmann, A. Vervisch, T. Tchenio, G. Lozano, A.Harel-Bellan, Synthetic small inhibiting RNAs: efficient tools to inactivate oncogenicmutations and restore p53 pathways, Proc. Natl. Acad. Sci. U. S. A. 99 (2002)14,849–14,854.

[28] M. Yoshinouchi, T. Yamada, M. Kizaki, J. Fen, T. Koseki, Y. Ikeda, T. Nishihara, K.Yamato, In vitro and in vivo growth suppression of human papillomavirus 16-positive cervical cancer cells by E6 siRNA, Mol. Ther. 8 (2003) 762–768.

[29] A. Choudhury, J. Charo, S.K. Parapuram, R.C. Hunt, D.M. Hunt, B. Seliger, R.Kiessling, Small interfering RNA (siRNA) inhibits the expression of the Her2/neugene, upregulates HLA class I and induces apoptosis of Her2/neu positive tumorcell lines, Int. J. Cancer 108 (2004) 71–77.

[30] G. Yang, K.Q. Cai, J.A. Thompson-Lanza, R.C. Bast Jr., J. Liu, Inhibition of breast andovarian tumor growth through multiple signaling pathways by using retrovirus-mediated small interfering RNA against Her-2/neu gene expression, J. Biol. Chem.279 (2004) 4339–4345.

[31] B. Farrow, P. Rychahou, C. Murillo, K.L. O'Connor, T. Iwamura, B.M. Evers,Inhibition of pancreatic cancer cell growth and induction of apoptosis with noveltherapies directed against protein kinase A, Surgery 134 (2003) 197–205.

[32] E. Yague, C.F. Higgins, S. Raguz, Complete reversal of multidrug resistance by stableexpressionof small interferingRNAs targetingMDR1,GeneTher.11 (2004)1170–1174.

[33] B.A. Kosciolek, K. Kalantidis, M. Tabler, P.T. Rowley, Inhibition of telomerase activityin human cancer cells by RNA interference, Mol. Cancer Ther. 2 (2003) 209–216.

[34] D.P. Cioca, Y. Aoki, K. Kiyosawa, RNA interference is a functional pathway withtherapeutic potential in human myeloid leukemia cell lines, Cancer Gene Ther.10 (2003) 125–133.

[35] H. Kawasaki, K. Taira, Short hairpin type of dsRNAs that are controlled by tRNA(Val)promoter significantly induce RNAi-mediated gene silencing in the cytoplasm ofhuman cells, Nucleic Acids Res. 31 (2003) 700–707.

[36] K. Li, S.Y. Lin, F.C. Brunicardi, P. Seu, Use of RNA interference to target cyclinE-overexpressing hepatocellular carcinoma, Cancer Res. 63 (2003) 3593–3597.

[37] U.N. Verma, R.M. Surabhi, A. Schmaltieg, C. Becerra, R.B. Gaynor, Small interferingRNAs directed against beta-catenin inhibit the in vitro and in vivo growth of coloncancer cells, Clin. Cancer Res. 9 (2003) 1291–1300.

[38] S. Aharinejad, P. Paulus, M. Sioud, M. Hofmann, K. Zins, R. Schafer, E.R. Stanley, D.Abraham, Colony-stimulating factor-1 blockade by antisense oligonucleotides andsmall interfering RNAs suppresses growth of humanmammary tumor xenografts inmice, Cancer Res. 64 (2004) 5378–5384.

[39] H. Uchida, T. Tanaka, K. Sasaki, K. Kato, H. Dehari, Y. Ito, M. Kobune, M. Miyagishi,K. Taira, H. Tahara, H. Hamada, Adenovirus-mediated transfer of siRNA againstsurvivin induced apoptosis and attenuated tumor cell growth in vitro and in vivo,Mol. Ther. 10 (2004) 162–171.

[40] A.J. Salisbury, V.M. Macaulay, Development of molecular agents for IGF receptortargeting, Horm. Metab. Res. 35 (2003) 843–849.

[41] M.S. Duxbury, H. Ito, M.J. Zinner, S.W. Ashley, E.E. Whang, Focal adhesion kinasegene silencing promotes anoikis and suppresses metastasis of human pancreaticadenocarcinoma cells, Surgery 135 (2004) 555–562.

[42] M.S. Duxbury, E. Matros, H. Ito, M.J. Zinner, S.W. Ashley, E.E. Whang, SystemicsiRNA-mediated gene silencing: a new approach to targeted therapy of cancer,Ann. Surg. 240 (2004) 667–674 [discussion 675–666].

[43] S. Filleur, A. Courtin, S. Ait-Si-Ali, J. Guglielmi, C. Merle, A. Harel-Bellan, P.Clezardin, F. Cabon, SiRNA-mediated inhibition of vascular endothelial growthfactor severely limits tumor resistance to antiangiogenic thrombospondin-1 andslows tumor vascularization and growth, Cancer Res. 63 (2003) 3919–3922.

[44] Y. Takei, K. Kadomatsu, Y. Yuzawa, S. Matsuo, T. Muramatsu, A small interferingRNA targeting vascular endothelial growth factor as cancer therapeutics, CancerRes. 64 (2004) 3365–3370.

[45] S.S. Lakka, C.S. Gondi, N. Yanamandra, W.C. Olivero, D.H. Dinh, M. Gujrati, J.S. Rao,Inhibition of cathepsin B andMMP-9 gene expression in glioblastoma cell line viaRNA interference reduces tumor cell invasion, tumor growth and angiogenesis,Oncogene 23 (2004) 4681–4689.

[46] A. Singh, S. Boldin-Adamsky, R.K. Thimmulappa, S.K. Rath, H. Ashush, J. Coulter, A.Blackford, S.N. Goodman, F. Bunz, W.H. Watson, E. Gabrielson, E. Feinstein, S.Biswal, RNAi-mediated silencing of nuclear factor erythroid-2-related factor 2gene expression in non-small cell lung cancer inhibits tumor growth andincreases efficacy of chemotherapy, Cancer Res. 68 (2008) 7975–7984.

[47] S. Nakahira, S. Nakamori, M. Tsujie, Y. Takahashi, J. Okami, S. Yoshioka, M.Yamasaki, S. Marubashi, I. Takemasa, A. Miyamoto, Y. Takeda, H. Nagano, K. Dono,K. Umeshita, M. Sakon, M. Monden, Involvement of ribonucleotide reductase M1subunit overexpression in gemcitabine resistance of human pancreatic cancer,Int. J. Cancer 120 (2007) 1355–1363.

[48] A. Fire, S. Xu, M.K. Montgomery, S.A. Kostas, S.E. Driver, C.C. Mello, Potent andspecific genetic interference by double-stranded RNA in Caenorhabditis elegans,Nature 391 (1998) 806–811.

[49] J.Y. Yu, S.L. DeRuiter, D.L. Turner, RNA interference by expression of short-interferingRNAs and hairpin RNAs in mammalian cells, Proc. Natl. Acad. Sci. U. S. A. 99 (2002)6047–6052.

[50] M. Miyagishi, K. Taira, U6 promoter-driven siRNAs with four uridine 3′ overhangsefficiently suppress targeted gene expression in mammalian cells, Nat.Biotechnol. 20 (2002) 497–500.

[51] A. Jarve, J. Muller, I.H. Kim, K. Rohr, C. MacLean, G. Fricker, U. Massing, F. Eberle, A.Dalpke, R. Fischer, M.F. Trendelenburg, M. Helm, Surveillance of siRNA integrityby FRET imaging, Nucleic Acids Res. 35 (2007) e124.

[52] J.P. Leonetti, N. Mechti, G. Degols, C. Gagnor, B. Lebleu, Intracellular distribution ofmicroinjected antisense oligonucleotides, Proc. Natl. Acad. Sci. U. S. A. 88 (1991)2702–2706.

[53] G.B. Robb, K.M. Brown, J. Khurana, T.M. Rana, Specific and potent RNAi in thenucleus of human cells, Nat. Struct. Mol. Biol. 12 (2005) 133–137.

[54] S.Y. Berezhna, L. Supekova, F. Supek, P.G. Schultz, A.A. Deniz, siRNA in human cellsselectively localizes to target RNA sites, Proc. Natl. Acad. Sci. U. S. A. 103 (2006)7682–7687.


[55] B.A. Janowski, K.E. Huffman, J.C. Schwartz, R. Ram, R. Nordsell, D.S. Shames, J.D.Minna, D.R. Corey, Involvement of AGO1 and AGO2 inmammalian transcriptionalsilencing, Nat. Struct. Mol. Biol. 13 (2006) 787–792.

[56] D.H. Kim, L.M. Villeneuve,K.V.Morris, J.J. Rossi, Argonaute-1 directs siRNA-mediatedtranscriptional gene silencing in human cells, Nat. Struct. Mol. Biol. 13 (2006)793–797.

[57] S. Rudel, A. Flatley, L. Weinmann, E. Kremmer, G. Meister, A multifunctionalhuman Argonaute2-specific monoclonal antibody, RNA 14 (2008) 1244–1253.

[58] T. Ohrt, J. Mutze, W. Staroske, L. Weinmann, J. Hock, K. Crell, G. Meister, P. Schwille,Fluorescence correlation spectroscopy and fluorescence cross-correlation spectro-scopy reveal the cytoplasmic origination of loaded nuclear RISC in vivo in humancells, Nucleic Acids Res. 36 (2008) 6439–6449.

[59] L. Weinmann, J. Hock, T. Ivacevic, T. Ohrt, J. Mutze, P. Schwille, E. Kremmer, V.Benes, H. Urlaub, G. Meister, Importin 8 is a gene silencing factor that targetsargonaute proteins to distinct mRNAs, Cell 136 (2009) 496–507.

[60] S. Guang, A.F. Bochner, D.M. Pavelec, K.B. Burkhart, S. Harding, J. Lachowiec, S.Kennedy, An Argonaute transports siRNAs from the cytoplasm to the nucleus,Science 321 (2008) 537–541.

[61] A. Grunweller, C. Gillen, V.A. Erdmann, J. Kurreck, Cellular uptake and localizationof a Cy3-labeled siRNA specific for the serine/threonine kinase Pim-1,Oligonucleotides 13 (2003) 345–352.

[62] T. Ohrt, D. Merkle, K. Birkenfeld, C.J. Echeverri, P. Schwille, In situ fluorescenceanalysis demonstrates active siRNA exclusion from the nucleus by exportin 5,Nucleic Acids Res. 34 (2006) 1369–1380.

[63] Y.L. Chiu, A. Ali, C.Y. Chu, H. Cao, T.M. Rana, Visualizing a correlation betweensiRNA localization, cellular uptake, and RNAi in living cells, Chem. Biol. 11 (2004)1165–1175.

[64] K. Forstemann, Y. Tomari, T. Du, V.V. Vagin, A.M. Denli, D.P. Bratu, C. Klattenhoff,W.E. Theurkauf, P.D. Zamore, Normal microRNA maturation and germ-line stemcell maintenance requires Loquacious, a double-stranded RNA-binding domainprotein, PLoS. Biol. 3 (2005) e236.

[65] F. Jiang, X. Ye, X. Liu, L. Fincher, D. McKearin, Q. Liu, Dicer-1 and R3D1-L catalyzemicroRNA maturation in Drosophila, Genes Dev. 19 (2005) 1674–1679.

[66] K. Saito, A. Ishizuka, H. Siomi, M.C. Siomi, Processing of pre-microRNAs by theDicer-1-Loquacious complex in Drosophila cells, PLoS. Biol. 3 (2005) e235.

[67] X. Liu, J.K. Park, F. Jiang, Y. Liu, D. McKearin, Q. Liu, Dicer-1, but not Loquacious, iscritical for assembly of miRNA-induced silencing complexes, RNA 13 (2007)2324–2329.

[68] E.P. Murchison, J.F. Partridge, O.H. Tam, S. Cheloufi, G.J. Hannon, Characterization ofDicer-deficientmurine embryonic stemcells, Proc. Natl. Acad. Sci. U. S. A.102 (2005)12,135–12,140.

[69] E. Bernstein, A.A. Caudy, S.M. Hammond, G.J. Hannon, Role for a bidentateribonuclease in the initiation step of RNA interference, Nature 409 (2001) 363–366.

[70] P. Provost, D. Dishart, J. Doucet, D. Frendewey, B. Samuelsson, O. Radmark,Ribonuclease activity and RNA binding of recombinant human Dicer, EMBO J.21 (2002) 5864–5874.

[71] S.M. Hammond, A.A. Caudy, G.J. Hannon, Post-transcriptional gene silencing bydouble-stranded RNA, Nat. Rev., Genet. 2 (2001) 110–119.

[72] M.A. Carmell, G.J. Hannon, RNase III enzymes and the initiation of gene silencing,Nat. Struct. Mol. Biol. 11 (2004) 214–218.

[73] I.J. Macrae, K. Zhou, F. Li, A. Repic, A.N. Brooks, W.Z. Cande, P.D. Adams, J.A.Doudna, Structural basis for double-stranded RNA processing by Dicer, Science311 (2006) 195–198.

[74] K.H. Kok, M.H. Ng, Y.P. Ching, D.Y. Jin, Human TRBP and PACT directly interact witheach other and associate with dicer to facilitate the production of smallinterfering RNA, J. Biol. Chem. 282 (2007) 17,649–17,657.

[75] G.B. Robb, T.M. Rana, RNA helicase A interacts with RISC in human cells andfunctions in RISC loading, Mol. Cell 26 (2007) 523–537.

[76] C. Matranga, Y. Tomari, C. Shin, D.P. Bartel, P.D. Zamore, Passenger–strandcleavage facilitates assembly of siRNA into Ago2-containing RNAi enzymecomplexes, Cell 123 (2005) 607–620.

[77] T.A. Rand, S. Petersen, F. Du, X. Wang, Argonaute2 cleaves the anti-guide strand ofsiRNA during RISC activation, Cell 123 (2005) 621–629.

[78] P.J. Leuschner, S.L. Ameres, S. Kueng, J. Martinez, Cleavage of the siRNA passengerstrand during RISC assembly in human cells, EMBO Rep. 7 (2006) 314–320.

[79] J.B. Preall, E.J. Sontheimer, RNAi: RISC gets loaded, Cell 123 (2005) 543–545.[80] B. Haley, P.D. Zamore, Kinetic analysis of the RNAi enzyme complex, Nat. Struct.

Mol. Biol. 11 (2004) 599–606.[81] R.I. Gregory, T.P. Chendrimada, N. Cooch, R. Shiekhattar, Human RISC couples

microRNA biogenesis and posttranscriptional gene silencing, Cell 123 (2005)631–640.

[82] E. Maniataki, Z. Mourelatos, A human, ATP-independent, RISC assembly machinefueled by pre-miRNA, Genes Dev. 19 (2005) 2979–2990.

[83] R.E. Collins, X. Cheng, Structural and biochemical advances in mammalian RNAi,J. Cell. Biochem. 99 (2006) 1251–1266.

[84] R.L. Boudreau, A.M. Monteys, B.L. Davidson, Minimizing variables among hairpin-based RNAi vectors reveals the potency of shRNAs, RNA 14 (2008) 1834–1844.

[85] B.R. Cullen, RNAi the natural way, Nat. Genet. 37 (2005) 1163–1165.[86] Y. Zeng, R. Yi, B.R. Cullen, Recognition and cleavage of primary microRNA

precursors by the nuclear processing enzyme Drosha, EMBO J. 24 (2005)138–148.

[87] J.M. Silva, M.Z. Li, K. Chang, W. Ge, M.C. Golding, R.J. Rickles, D. Siolas, G. Hu, P.J.Paddison, M.R. Schlabach, N. Sheth, J. Bradshaw, J. Burchard, A. Kulkarni, G. Cavet,R. Sachidanandam,W.R. McCombie, M.A. Cleary, S.J. Elledge, G.J. Hannon, Second-generation shRNA libraries covering the mouse and human genomes, Nat. Genet.37 (2005) 1281–1288.

[88] Y. Lee, C. Ahn, J. Han, H. Choi, J. Kim, J. Yim, J. Lee, P. Provost, O. Radmark, S. Kim, V.N.Kim, Thenuclear RNase III Drosha initiatesmicroRNAprocessing,Nature 425 (2003)415–419.

[89] H. Zhang, F.A. Kolb, V. Brondani, E. Billy, W. Filipowicz, Human Dicerpreferentially cleaves dsRNAs at their termini without a requirement for ATP,EMBO J. 21 (2002) 5875–5885.

[90] Y. Lee, K. Jeon, J.T. Lee, S. Kim, V.N. Kim, MicroRNA maturation: stepwiseprocessing and subcellular localization, EMBO J. 21 (2002) 4663–4670.

[91] B.R. Cullen, Transcription and processing of human microRNA precursors, Mol.Cell 16 (2004) 861–865.

[92] R. Yi, Y. Qin, I.G. Macara, B.R. Cullen, Exportin-5 mediates the nuclear export ofpre-microRNAs and short hairpin RNAs, Genes Dev. 17 (2003) 3011–3016.

[93] E. Lund, S. Guttinger, A. Calado, J.E. Dahlberg, U. Kutay, Nuclear export ofmicroRNA precursors, Science 303 (2004) 95–98.

[94] Y.S. Lee, K. Nakahara, J.W. Pham, K. Kim, Z. He, E.J. Sontheimer, R.W. Carthew,Distinct roles for Drosophila Dicer-1 and Dicer-2 in the siRNA/miRNA silencingpathways, Cell 117 (2004) 69–81.

[95] M. Landthaler, D.Gaidatzis, A. Rothballer, P.Y. Chen, S.J. Soll, L. Dinic, T. Ojo,M.Hafner,M. Zavolan, T. Tuschl, Molecular characterization of human Argonaute-containingribonucleoprotein complexes and their bound target mRNAs, RNA 14 (2008)2580–2596.

[96] S.M. Hammond, S. Boettcher, A.A. Caudy, R. Kobayashi, G.J. Hannon, Argonaute2, alink between genetic and biochemical analyses of RNAi, Science 293 (2001)1146–1150.

[97] E.J. Sontheimer, R.W. Carthew, Molecular biology. Argonaute journeys into theheart of RISC, Science 305 (2004) 1409–1410.

[98] G. Hutvagner, P.D. Zamore, A microRNA in a multiple-turnover RNAi enzymecomplex, Science 297 (2002) 2056–2060.

[99] S. Yekta, I.H. Shih, D.P. Bartel, MicroRNA-directed cleavage of HOXB8 mRNA,Science 304 (2004) 594–596.

[100] D.T. Humphreys, B.J. Westman, D.I. Martin, T. Preiss, MicroRNAs control translationinitiation by inhibiting eukaryotic initiation factor 4E/cap and poly(A) tail function,Proc. Natl. Acad. Sci. U. S. A. 102 (2005) 16,961–16,966.

[101] R.S. Pillai, S.N. Bhattacharyya, C.G. Artus, T. Zoller, N. Cougot, E. Basyuk, E.Bertrand, W. Filipowicz, Inhibition of translational initiation by Let-7 MicroRNAin human cells, Science 309 (2005) 1573–1576.

[102] R. Thermann, M.W. Hentze, Drosophila miR2 induces pseudo-polysomes andinhibits translation initiation, Nature 447 (2007) 875–878.

[103] M.A. Valencia-Sanchez, J. Liu, G.J. Hannon, R. Parker, Control of translation andmRNA degradation by miRNAs and siRNAs, Genes Dev. 20 (2006) 515–524.

[104] R. Parker, U. Sheth, P bodies and the control of mRNA translation and degradation,Mol. Cell 25 (2007) 635–646.

[105] K. Okamura, A. Ishizuka, H. Siomi, M.C. Siomi, Distinct roles for Argonauteproteins in small RNA-directed RNA cleavage pathways, Genes Dev. 18 (2004)1655–1666.

[106] K. Miyoshi, H. Tsukumo, T. Nagami, H. Siomi, M.C. Siomi, Slicer function ofDrosophila Argonautes and its involvement inRISC formation, Genes Dev.19 (2005)2837–2848.

[107] Y. Tomari, T. Du, P.D. Zamore, Sorting of Drosophila small silencing RNAs, Cell130 (2007) 299–308.

[108] K. Forstemann, M.D. Horwich, L. Wee, Y. Tomari, P.D. Zamore, DrosophilamicroRNAs are sorted into functionally distinct argonaute complexes afterproduction by dicer-1, Cell 130 (2007) 287–297.

[109] F.A. Steiner, S.W. Hoogstrate, K.L. Okihara, K.L. Thijssen, R.F. Ketting, R.H. Plasterk,T. Sijen, Structural features of small RNA precursors determine Argonaute loadingin Caenorhabditis elegans, Nat. Struct. Mol. Biol. 14 (2007) 927–933.

[110] A. Azuma-Mukai, H. Oguri, T. Mituyama, Z.R. Qian, K. Asai, H. Siomi, M.C. Siomi,Characterization of endogenous human Argonautes and their miRNA partners inRNA silencing, Proc. Natl. Acad. Sci. U. S. A. 105 (2008) 7964–7969.

[111] D.G. Hendrickson, D.J. Hogan, D. Herschlag, J.E. Ferrell, P.O. Brown, Systematicidentification of mRNAs recruited to argonaute 2 by specific microRNAs andcorresponding changes in transcript abundance, PLoS ONE 3 (2008) e2126.

[112] Y. Tay, J. Zhang,A.M. Thomson, B. Lim, I. Rigoutsos,MicroRNAs toNanog,Oct4andSox2coding regions modulate embryonic stem cell differentiation, Nature 455 (2008)1124–1128.

[113] M. Sano, M. Sierant, M. Miyagishi, M. Nakanishi, Y. Takagi, S. Sutou, Effect ofasymmetric terminal structures of short RNA duplexes on the RNA interferenceactivity and strand selection, Nucleic Acids Res. 36 (2008) 5812–5821.

[114] D.H. Kim,M.A. Behlke, S.D. Rose,M.S. Chang, S. Choi, J.J. Rossi, Synthetic dsRNADicersubstrates enhance RNAi potency and efficacy, Nat. Biotechnol. 23 (2005) 222–226.

[115] M.A. McAnuff, G.R. Rettig, K.G. Rice, Potency of siRNA versus shRNA mediatedknockdown in vivo, J. Pharm. Sci. 96 (2007) 2922–2930.

[116] D. Siolas, C. Lerner, J. Burchard, W. Ge, P.S. Linsley, P.J. Paddison, G.J. Hannon, M.A.Cleary, Synthetic shRNAsaspotentRNAi triggers, Nat. Biotechnol. 23 (2005)227–231.

[117] A.V. Vlassov, B. Korba, K. Farrar, S. Mukerjee, A.A. Seyhan, H. Ilves, R.L. Kaspar, D.Leake, S.A. Kazakov, B.H. Johnston, shRNAs targeting hepatitis C: effects ofsequence and structural features, and comparision with siRNA, Oligonucleotides17 (2007) 223–236.

[118] M. Gossen, H. Bujard, Tight control of gene expression in mammalian cells bytetracycline-responsive promoters, Proc.Natl. Acad. Sci. U. S.A. 89 (1992)5547–5551.

[119] S.Gupta, R.A. Schoer, J.E. Egan, G.J.Hannon, V.Mittal, Inducible, reversible, and stableRNA interference in mammalian cells, Proc. Natl. Acad. Sci. U. S. A. 101 (2004)1927–1932.

[120] R.A. Dickins, M.T. Hemann, J.T. Zilfou, D.R. Simpson, I. Ibarra, G.J. Hannon, S.W.Lowe, Probing tumor phenotypes using stable and regulated synthetic microRNAprecursors, Nat. Genet. 37 (2005) 1289–1295.


[121] A.W. Tong, Y.A. Zhang, J. Nemunaitis, Small interfering RNA for experimentalcancer therapy, Curr. Opin. Mol. Ther. 7 (2005) 114–124.

[122] A. Aigner, Cellular delivery in vivo of siRNA-based therapeutics, Curr. Pharm. Des.14 (2008) 3603–3619.

[123] A.W. Tong, C.M. Jay, N. Senzer, P.B. Maples, J. Nemunaitis, Systemic therapeutic genedelivery for cancer: crafting Paris' arrow, Curr. Gene Ther. 9 (1) (2009 Feb) 45–60.

[124] J.S. Vorhies, J. Nemunaitis, Nonviral delivery vehicles for use in short hairpin RNA-based cancer therapies, Expert Rev. Anticancer Ther. 7 (2007) 373–382.

[125] A.P. McCaffrey, L. Meuse, T.T. Pham, D.S. Conklin, G.J. Hannon, M.A. Kay, RNAinterference in adult mice, Nature 418 (2002) 38–39.

[126] W.M.Merritt, Y.G. Lin, L.Y. Han, A.A. Kamat,W.A. Spannuth, R. Schmandt, D. Urbauer,L.A. Pennacchio, J.F. Cheng, A.M. Nick, M.T. Deavers, A. Mourad-Zeidan, H. Wang, P.Mueller,M.E. Lenburg, J.W.Gray, S.Mok,M.J. Birrer, G. Lopez-Berestein,R.L. Coleman,M. Bar-Eli, A.K. Sood, Dicer, Drosha, and outcomes inpatientswith ovarian cancer, N.Engl. J. Med. 359 (2008) 2641–2650.

[127] J. Lu, G. Getz, E.A.Miska, E. Alvarez-Saavedra, J. Lamb, D. Peck, A. Sweet-Cordero, B.L.Ebert, R.H. Mak, A.A. Ferrando, J.R. Downing, T. Jacks, H.R. Horvitz, T.R. Golub,MicroRNA expression profiles classify human cancers, Nature 435 (2005) 834–838.

[128] R. Sandberg, J.R. Neilson, A. Sarma, P.A. Sharp, C.B. Burge, Proliferating cellsexpress mRNAs with shortened 3′ untranslated regions and fewer microRNAtarget sites, Science 320 (2008) 1643–1647.

[129] S. Chiosea, E. Jelezcova, U. Chandran, M. Acquafondata, T. McHale, R.W. Sobol, R.Dhir, Up-regulation of dicer, a component of theMicroRNAmachinery, in prostateadenocarcinoma, Am. J. Pathol. 169 (2006) 1812–1820.

[130] N. Sugito, H. Ishiguro, Y. Kuwabara, M. Kimura, A. Mitsui, H. Kurehara, T. Ando, R.Mori, N. Takashima,R.Ogawa,Y. Fujii, RNASEN regulates cell proliferation andaffectssurvival in esophageal cancer patients, Clin. Cancer Res. 12 (2006) 7322–7328.

[131] S. Tokumaru, M. Suzuki, H. Yamada, M. Nagino, T. Takahashi, Let-7 regulates Dicerexpression and constitutes a negative feedback loop, Carcinogenesis 29 (2008)2073–2077.

[132] J.J. Forman, A. Legesse-Miller, H.A. Coller, A search for conserved sequences incoding regions reveals that the let-7 microRNA targets Dicer within its codingsequence, Proc. Natl. Acad. Sci. U. S. A. 105 (2008) 14,879–14,884.

[133] Breakdown of RNAi-Based Drugs in the Clinic, RNAi News, vol. 6, 2008.[134] J.D. Heidel, J.Y. Liu, Y. Yen, B. Zhou, B.S. Heale, J.J. Rossi, D.W. Bartlett, M.E. Davis,

Potent siRNA inhibitors of ribonucleotide reductase subunit RRM2 reduce cellproliferation in vitro and in vivo, Clin. Cancer Res. 13 (2007) 2207–2215.

[135] R. Zukiel, S. Nowak, E. Wyszko, K. Rolle, I. Gawronska, M.Z. Barciszewska, J.Barciszewski, Suppression of human brain tumor with interference RNA specificfor tenascin-C, Cancer Biol. Ther. 5 (2006) 1002–1007.

[136] J.D. Heidel, Z. Yu, J.Y. Liu, S.M. Rele, Y. Liang, R.K. Zeidan, D.J. Kornbrust, M.E. Davis,Administration in non-human primates of escalating intravenous doses oftargeted nanoparticles containing ribonucleotide reductase subunit M2 siRNA,Proc. Natl. Acad. Sci. U. S. A. 104 (2007) 5715–5721.

[137] E. Check, A tragic setback, Nature 420 (2002) 116–118.[138] T. Nguyen, E.M. Menocal, J. Harborth, J.H. Fruehauf, RNAi therapeutics: an update

on delivery, Curr. Opin. Mol. Ther. 10 (2008) 158–167.[139] J. Luten, C.F. van Nostrum, S.C. De Smedt, W.E. Hennink, Biodegradable polymers as

non-viral carriers for plasmid DNA delivery, J. Control. Release 126 (2008) 97–110.[140] S. Zhang, B. Zhao, H. Jiang, B. Wang, B. Ma, Cationic lipids and polymers mediated

vectors for delivery of siRNA, J. Control. Release 123 (2007) 1–10.[141] A. Aigner, Nonviral in vivo delivery of therapeutic small interfering RNAs, Curr.

Opin. Mol. Ther. 9 (2007) 345–352.[142] J.A. Hughes, G.A. Rao, Targeted polymers for gene delivery, Expert Opin. Drug

Deliv. 2 (2005) 145–157.[143] W.T. Godbey, M.A. Barry, P. Saggau, K.K. Wu, A.G. Mikos, Poly(ethylenimine)-

mediated transfection: a new paradigm for gene delivery, J. Biomed. Mater. Res.51 (2000) 321–328.

[144] M. Thomas, A.M. Klibanov, Enhancing polyethylenimine's delivery of plasmidDNA intomammalian cells, Proc. Natl. Acad. Sci. U. S. A. 99 (2002) 14,640–14,645.

[145] Y. Kim, M. Tewari, J.D. Pajerowski, S. Cai, S. Sen, J. Williams, S. Sirsi, G. Lutz, D.E.Discher, Polymersome delivery of siRNA and antisense oligonucleotides, J. Control.Release 134 (2) (2009 Mar 4) 132–140.

[146] J.D. Heidel, Linear cyclodextrin-containing polymers and their use as deliveryagents, Expert Opin. Drug Deliv. 3 (2006) 641–646.

[147] J.N. Moreira, A. Santos, V. Moura, M.C. Pedroso de Lima, S. Simoes, Non-viral lipid-based nanoparticles for targeted cancer systemic gene silencing, J. Nanosci.Nanotechnol. 8 (2008) 2187–2204.

[148] N. Blow, Small RNAs: delivering the future, Nature 450 (2007) 1117–1120.[149] T.S. Zimmermann, A.C. Lee, A. Akinc, B. Bramlage, D. Bumcrot, M.N. Fedoruk, J.

Harborth, J.A. Heyes, L.B. Jeffs, M. John, A.D. Judge, K. Lam, K. McClintock, L.V.Nechev, L.R. Palmer, T. Racie, I. Rohl, S. Seiffert, S. Shanmugam, V. Sood, J.Soutschek, I. Toudjarska, A.J. Wheat, E. Yaworski, W. Zedalis, V. Koteliansky, M.Manoharan, H.P. Vornlocher, I. MacLachlan, RNAi-mediated gene silencing innon-human primates, Nature 441 (2006) 111–114.

[150] A. Santel, M. Aleku, O. Keil, J. Endruschat, V. Esche, B. Durieux, K. Loffler, M.Fechtner, T. Rohl, G. Fisch, S. Dames, W. Arnold, K. Giese, A. Klippel, J. Kaufmann,RNA interference in the mouse vascular endothelium by systemic administrationof siRNA-lipoplexes for cancer therapy, Gene Ther. 13 (2006) 1360–1370.

[151] A. Santel, M. Aleku, O. Keil, J. Endruschat, V. Esche, G. Fisch, S. Dames, K. Loffler, M.Fechtner, W. Arnold, K. Giese, A. Klippel, J. Kaufmann, A novel siRNA-lipoplextechnology for RNA interference in the mouse vascular endothelium, Gene Ther.13 (2006) 1222–1234.

[152] M. Aleku, G. Fisch, K. Mopert, O. Keil, W. Arnold, J. Kaufmann, A. Santel,Intracellular localization of lipoplexed siRNA in vascular endothelial cells ofdifferent mouse tissues, Microvasc. Res. 76 (2008) 31–41.

[153] M.Aleku, P. Schulz, O.Keil, A. Santel, U. Schaeper, B.Dieckhoff,O. Janke, J. Endruschat,B.Durieux,N.Roder, K. Loffler, C. Lange,M. Fechtner, K.Mopert,G. Fisch, S.Dames,W.Arnold, K. Jochims, K. Giese, B. Wiedenmann, A. Scholz, J. Kaufmann, Atu027, aliposomal small interfering RNA formulation targeting protein kinase N3, inhibitscancer progression, Cancer Res. 68 (2008) 9788–9798.

[154] J.S. Vorhies, J.J. Nemunaitis, Synthetic vs. natural/biodegradable polymers fordelivery of shRNA-based cancer therapies, Methods Mol. Biol. 480 (2009) 11–29.

[155] P. Kumar, H.S. Ban, S.S. Kim, H. Wu, T. Pearson, D.L. Greiner, A. Laouar, J. Yao, V.Haridas, K. Habiro, Y.G. Yang, J.H. Jeong, K.Y. Lee, Y.H. Kim, S.W. Kim,M. Peipp, G.H.Fey, N. Manjunath, L.D. Shultz, S.K. Lee, P. Shankar, T cell-specific siRNA deliverysuppresses HIV-1 infection in humanized mice, Cell 134 (2008 Dec 30) 577–586.

[156] K. Gao, L. Huang, Nonviralmethods for siRNADelivery,Mol. Pharmacol. 2008Dec 30.[157] A. Akinc, A. Zumbuehl, M. Goldberg, E.S. Leshchiner, V. Busini, N. Hossain, S.A.

Bacallado, D.N. Nguyen, J. Fuller, R. Alvarez, A. Borodovsky, T. Borland, R. Constien,A. de Fougerolles, J.R. Dorkin, K. Narayanannair Jayaprakash, M. Jayaraman, M.John, V. Koteliansky, M. Manoharan, L. Nechev, J. Qin, T. Racie, D. Raitcheva, K.G.Rajeev, D.W. Sah, J. Soutschek, I. Toudjarska, H.P. Vornlocher, T.S. Zimmermann, R.Langer, D.G. Anderson, A combinatorial library of lipid-like materials for deliveryof RNAi therapeutics, Nat. Biotechnol. 26 (2008) 561–569.

[158] S. Ghatak, V.C. Hascall, F.G. Berger, M.M. Penas, C. Davis, E. Jabari, X. He, J.S. Norris,Y. Dang, R.R. Markwald, S. Misra, Tissue-specific shRNA delivery: a novelapproach for gene therapy in cancer, Connect. Tissue Res. 49 (2008) 265–269.

[159] P. Kumar, H. Wu, J.L. McBride, K.E. Jung, M.H. Kim, B.L. Davidson, S.K. Lee, P.Shankar, N. Manjunath, Transvascular delivery of small interfering RNA to thecentral nervous system, Nature 448 (2007) 39–43.

[160] A. Judge, I. MacLachlan, Overcoming the innate immune response to smallinterfering RNA, Hum. Gene Ther. 19 (2008) 111–124.

[161] H. Lv, S. Zhang, B. Wang, S. Cui, J. Yan, Toxicity of cationic lipids and cationicpolymers in gene delivery, J. Control. Release 114 (2006) 100–109.

[162] T.G. Park, J.H. Jeong, S.W. Kim, Current status of polymeric gene delivery systems,Adv. Drug Deliv. Rev. 58 (2006) 467–486.

[163] M. Breunig, U. Lungwitz, R. Liebl, A. Goepferich, Breaking up the correlation betweenefficacyand toxicity fornonviral genedelivery, Proc.Natl.Acad. Sci. U. S. A.104 (2007)14,454–14,459.

[164] A.L. Jackson, S.R. Bartz, J. Schelter, S.V. Kobayashi, J. Burchard, M. Mao, B. Li, G.Cavet, P.S. Linsley, Expression profiling reveals off-target gene regulation by RNAi,Nat. Biotechnol. 21 (2003) 635–637.

[165] A.L. Jackson, J. Burchard, J. Schelter, B.N. Chau, M. Cleary, L. Lim, P.S. Linsley,Widespread siRNA “off-target” transcript silencing mediated by seed regionsequence complementarity, RNA 12 (2006) 1179–1187.

[166] X. Lin, X. Ruan, M.G. Anderson, J.A. McDowell, P.E. Kroeger, S.W. Fesik, Y. Shen,siRNA-mediated off-target gene silencing triggered by a 7 nt complementation,Nucleic Acids Res. 33 (2005) 4527–4535.

[167] A.L. Jackson, J. Burchard, D. Leake, A. Reynolds, J. Schelter, J. Guo, J.M. Johnson, L.Lim, J. Karpilow, K. Nichols, W. Marshall, A. Khvorova, P.S. Linsley, Position-specific chemical modification of siRNAs reduces “off-target” transcript silencing,RNA 12 (2006) 1197–1205.

[168] A. Birmingham, E.M. Anderson, A. Reynolds, D. Ilsley-Tyree, D. Leake, Y. Fedorov, S.Baskerville, E. Maksimova, K. Robinson, J. Karpilow, W.S. Marshall, A. Khvorova, 3′UTR seedmatches, but not overall identity, are associatedwith RNAi off-targets, Nat.Methods 3 (2006) 199–204.

[169] E. Anderson, Q. Boese, A. Khvorova, J. Karpilow, Identifying siRNA-induced off-targets by microarray analysis, Methods Mol. Biol. 442 (2008) 45–63.

[170] S. van Dongen, C. Abreu-Goodger, A.J. Enright, Detecting microRNA binding andsiRNA off-target effects from expression data, Nat. Methods 5 (2008) 1023–1025.

[171] S. Qiu, C.M. Adema, T. Lane, A computational study of off-target effects of RNAinterference, Nucleic Acids Res. 33 (2005) 1834–1847.

[172] Y.K. Park, S.M. Park, Y.C. Choi, D. Lee, M. Won, Y.J. Kim, AsiDesigner: exon-basedsiRNA design server considering alternative splicing, Nucleic Acids Res. 36 (2008)W97–103.

[173] T. Yamada, S. Morishita, Accelerated off-target search algorithm for siRNA,Bioinformatics 21 (2005) 1316–1324.

[174] Y. Naito, T. Yamada, T. Matsumiya, K. Ui-Tei, K. Saigo, S. Morishita, dsCheck: highlysensitive off-target search software for double-stranded RNA-mediated RNAinterference, Nucleic Acids Res. 33 (2005) W589–591.

[175] Q. Du, H. Thonberg, J. Wang, C. Wahlestedt, Z. Liang, A systematic analysis of thesilencing effects of an active siRNA at all single-nucleotide mismatched targetsites, Nucleic Acids Res. 33 (2005) 1671–1677.

[176] D.S. Schwarz, H. Ding, L. Kennington, J.T. Moore, J. Schelter, J. Burchard, P.S. Linsley,N. Aronin, Z. Xu, P.D. Zamore, Designing siRNA that distinguish between genesthat differ by a single nucleotide, PLoS Genet. 2 (2006) e140.

[177] C. Dahlgren, H.Y. Zhang, Q. Du, M. Grahn, G. Norstedt, C. Wahlestedt, Z. Liang,Analysis of siRNA specificity on targets with double-nucleotide mismatches,Nucleic Acids Res. 36 (2008) e53.

[178] J.K. Watts, G.F. Deleavey, M.J. Damha, Chemically modified siRNA: tools andapplications, Drug Discov. Today 13 (2008) 842–855.

[179] O. Snove Jr., J.J. Rossi, Chemical modifications rescue off-target effects of RNAi,ACS Chem. Biol. 1 (2006) 274–276.

[180] M.A. Behlke, Chemical modification of siRNAs for in vivo use, Oligonucleotides18 (2008) 305–319.

[181] J.B. Bramsen, M.B. Laursen, C.K. Damgaard, S.W. Lena, B.R. Babu, J. Wengel, J. Kjems,Improved silencing properties using small internally segmented interfering RNAs,Nucleic Acids Res. 35 (2007) 5886–5897.

[182] P.Y. Chen, L. Weinmann, D. Gaidatzis, Y. Pei, M. Zavolan, T. Tuschl, G. Meister,Strand-specific 5′-O-methylation of siRNA duplexes controls guide strandselection and targeting specificity, RNA 14 (2008) 263–274.


[183] K. Ui-Tei, Y. Naito, S. Zenno, K. Nishi, K. Yamato, F. Takahashi, A. Juni, K. Saigo,Functional dissection of siRNA sequence by systematic DNA substitution:modifiedsiRNAwith a DNA seed arm is a powerful tool for mammalian gene silencing withsignificantly reduced off-target effect, Nucleic Acids Res. 36 (2008) 2136–2151.

[184] D. Rao, N. Senzer, M.A. Cleary, J. Nemunaitis, Comparative assessment of siRNAand shRNA off target effects: what is slowing clinical development. Cancer GeneTherapy (2009).

[185] D. Bumcrot, M. Manoharan, V. Koteliansky, D.W. Sah, RNAi therapeutics: apotential new class of pharmaceutical drugs, Nat. Chem. Biol. 2 (2006) 711–719.

[186] A.J. Bridge, S. Pebernard, A. Ducraux, A.L. Nicoulaz, R. Iggo, Induction of aninterferon response by RNAi vectors in mammalian cells, Nat. Genet. 34 (2003)263–264.

[187] C.A. Sledz, M. Holko, M.J. de Veer, R.H. Silverman, B.R. Williams, Activation of theinterferon system by short-interfering RNAs, Nat. Cell Biol. 5 (2003) 834–839.

[188] K. Kariko, P. Bhuyan, J. Capodici, D. Weissman, Small interfering RNAs mediatesequence-independent gene suppression and induce immune activation bysignaling through toll-like receptor 3, J. Immunol. 172 (2004) 6545–6549.

[189] V. Hornung, M. Guenthner-Biller, C. Bourquin, A. Ablasser, M. Schlee, S. Uematsu,A. Noronha, M. Manoharan, S. Akira, A. de Fougerolles, S. Endres, G. Hartmann,Sequence-specific potent induction of IFN-alpha by short interfering RNA inplasmacytoid dendritic cells through TLR7, Nat. Med. 11 (2005) 263–270.

[190] A.D. Judge, V. Sood, J.R. Shaw, D. Fang, K. McClintock, I. MacLachlan, Sequence-dependent stimulation of the mammalian innate immune response by syntheticsiRNA, Nat. Biotechnol. 23 (2005) 457–462.

[191] S.S. Diebold, T. Kaisho, H. Hemmi, S. Akira, C. Reis e Sousa, Innate antiviralresponses by means of TLR7-mediated recognition of single-stranded RNA,Science 303 (2004) 1529–1531.

[192] F. Heil, H. Hemmi, H. Hochrein, F. Ampenberger, C. Kirschning, S. Akira, G. Lipford,H. Wagner, S. Bauer, Species-specific recognition of single-stranded RNA via toll-like receptor 7 and 8, Science 303 (2004) 1526–1529.

[193] M. Karin, Y. Yamamoto, Q.M. Wang, The IKK NF-kappa B system: a treasure trovefor drug development, Nat. Rev., Drug Discov. 3 (2004) 17–26.

[194] J.T. Marques, T. Devosse, D. Wang, M. Zamanian-Daryoush, P. Serbinowski, R.Hartmann, T. Fujita, M.A. Behlke, B.R. Williams, A structural basis fordiscriminating between self and nonself double-stranded RNAs in mammaliancells, Nat. Biotechnol. 24 (2006) 559–565.

[195] M. Sioud, RNA interference and innate immunity, Adv. Drug Deliv. Rev. 59 (2007)153–163.

[196] M.E. Kleinman, K. Yamada, A. Takeda, V. Chandrasekaran, M. Nozaki, J.Z. Baffi, R.J.Albuquerque, S. Yamasaki, M. Itaya, Y. Pan, B. Appukuttan, D. Gibbs, Z. Yang, K.Kariko, B.K. Ambati, T.A. Wilgus, L.A. DiPietro, E. Sakurai, K. Zhang, J.R. Smith, E.W.Taylor, J. Ambati, Sequence- and target-independent angiogenesis suppression bysiRNA via TLR3, Nature 452 (2008) 591–597.

[197] M. Robbins, A. Judge, E. Ambegia, C. Choi, E. Yaworski, L. Palmer, K. McClintock, I.Maclachlan, Misinterpreting the therapeutic effects of siRNA caused by immunestimulation, Hum. Gene Ther. 19 (10) (2008 Oct) 991–999.

[198] S. Bauer, C.J. Kirschning, H. Hacker, V. Redecke, S. Hausmann, S. Akira, H. Wagner,G.B. Lipford, Human TLR9 confers responsiveness to bacterial DNA via species-specific CpGmotif recognition, Proc. Natl. Acad. Sci. U. S. A. 98 (2001) 9237–9242.

[199] H. Hemmi, O. Takeuchi, T. Kawai, T. Kaisho, S. Sato, H. Sanjo, M. Matsumoto, K.Hoshino, H. Wagner, K. Takeda, S. Akira, A toll-like receptor recognizes bacterialDNA, Nature 408 (2000) 740–745.

[200] H.K. de Wolf, N. Johansson, A.T. Thong, C.J. Snel, E. Mastrobattista, W.E. Hennink,G. Storm, Plasmid CpG depletion improves degree and duration of tumor geneexpression after intravenous administration of polyplexes, Pharm. Res. 25 (2008)1654–1662.

[201] M.A. Robbins, M. Li, I. Leung, H. Li, D.V. Boyer, Y. Song, M.A. Behlke, J.J. Rossi, Stableexpression of shRNAs in human CD34+ progenitor cells can avoid induction ofinterferon responses to siRNAs in vitro, Nat. Biotechnol. 24 (2006) 566–571.

[202] M. Sioud, Induction of inflammatory cytokines and interferon responses bydouble-stranded and single-stranded siRNAs is sequence-dependent andrequires endosomal localization, J. Mol. Biol. 348 (2005) 1079–1090.

[203] H. Akashi, M. Miyagishi, T. Yokota, T. Watanabe, T. Hino, K. Nishina, M. Kohara, K.Taira, Escape from the interferon response associated with RNA interference usingvectors that encode long modified hairpin-RNA, Mol. Biosyst. 1 (2005) 382–390.

[204] T. Watanabe, M. Sudoh, M. Miyagishi, H. Akashi, M. Arai, K. Inoue, K. Taira, M.Yoshiba, M. Kohara, Intracellular-diced dsRNA has enhanced efficacy for silencingHCV RNA and overcomes variation in the viral genotype, Gene Ther. 13 (2006)883–892.

[205] M. Bauer, N. Kinkl, A. Meixner, E. Kremmer, M. Riemenschneider, H. Forstl, T.Gasser, M. Ueffing, Prevention of interferon-stimulated gene expression usingmicroRNA-designed hairpins, Gene Ther. 16 (2009) 142–147.

[206] M. Sioud, Single-stranded small interfering RNA are more immunostimulatorythan their double-stranded counterparts: a central role for 2′-hydroxyl uridinesin immune responses, Eur. J. Immunol. 36 (2006) 1222–1230.

[207] J.M. Layzer, A.P. McCaffrey, A.K. Tanner, Z. Huang, M.A. Kay, B.A. Sullenger, In vivoactivity of nuclease-resistant siRNAs, RNA 10 (2004) 766–771.

[208] K. Kariko, M. Buckstein, H. Ni, D. Weissman, Suppression of RNA recognition byToll-like receptors: the impact of nucleoside modification and the evolutionaryorigin of RNA, Immunity 23 (2005) 165–175.

[209] D. Grimm, K.L. Streetz, C.L. Jopling, T.A. Storm, K. Pandey, C.R. Davis, P. Marion, F.Salazar, M.A. Kay, Fatality in mice due to oversaturation of cellular microRNA/shorthairpin RNA pathways, Nature 441 (2006) 537–541.

[210] D. Grimm, M.A. Kay, Therapeutic application of RNAi: is mRNA targeting finallyready for prime time? J. Clin. Invest. 117 (2007) 3633–3641.

[211] M. John, R. Constien, A. Akinc, M. Goldberg, Y.A. Moon, M. Spranger, P. Hadwiger, J.Soutschek, H.P. Vornlocher, M. Manoharan, M. Stoffel, R. Langer, D.G. Anderson, J.D.Horton, V. Koteliansky, D. Bumcrot, Effective RNAi-mediated gene silencingwithoutinterruption of the endogenous microRNA pathway, Nature 449 (2007) 745–747.

[212] R. Yi, B.P. Doehle, Y. Qin, I.G. Macara, B.R. Cullen, Overexpression of exportin 5enhances RNA interference mediated by short hairpin RNAs and microRNAs, RNA11 (2005) 220–226.

[213] J.C. Giering, D. Grimm, T.A. Storm, M.A. Kay, Expression of shRNA from a tissue-specific pol II promoter is aneffectiveand safeRNAi therapeutic,Mol. Ther.16 (2008)1630–1636.

Advanced Drug Delivery Reviews - diyhpl

Documents

Transcript of Advanced Drug Delivery Reviews - diyhpl