Advanced Drug Delivery Reviews 61 (2009) 746–759 Contents lists available at ScienceDirect Advanced Drug Delivery Reviews j o u r n a l h o m e p a g e : w w w. e l s ev i e r. c o m / l o c a t e / a d d r 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, USA Mary Crowley Cancer Research Centers, Dallas, TX, USA c Texas Oncology PA, USA d Baylor Sammons Cancer Center, Dallas, TX, USA b a r t i c l e i n f o Article history: Received 23 January 2009 Accepted 13 April 2009 Available online 20 April 2009 Keywords: RNA interference Bi-functional Cancer Personalized a b s t r a c t RNA interference (RNAi) is a natural process through which expression of a targeted gene can be knocked down with high speciﬁcity and selectivity. Using available technology and bioinformatics investigators will soon be able to identify relevant bio molecular tumor network hubs as potential key targets for knockdown approaches. 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 effect per dose are optimally suited for certain medical disorders (i.e. viral injections). However, using the endogenous processing machinery, optimized shRNA constructs allow for high potency and sustainable effects using low copy numbers resulting in less off-target effects, particularly if embedded in a miRNA scaffold. Bi-functional design may further enhance potency and safety of RNAi-based therapeutics. Remaining challenges include tumor selective delivery vehicles and more complete evaluation of the scope and scale of off-target effects. This review will compare siRNA, shRNA and bi-functional shRNA. © 2009 Elsevier B.V. All rights reserved. Contents 1. 2. Introduction . . . . . . . . . . . . . . . . . . . . . . . . Targeted cancer gene therapy and RNA interference. . . . . . 2.1. Personalized approach for cancer therapy . . . . . . . 2.2. RNA interference for cancer. . . . . . . . . . . . . . 3. Small interfering RNA (siRNA) and short hairpin RNA (shRNA) 3.1. siRNA . . . . . . . . . . . . . . . . . . . . . . . . 3.2. shRNA. . . . . . . . . . . . . . . . . . . . . . . . 3.3. Bi-functional shRNA . . . . . . . . . . . . . . . . . 3.4. Summary of si/sh/bi . . . . . . . . . . . . . . . . . 4. Delivery . . . . . . . . . . . . . . . . . . . . . . . . . . 5. Off-target effects . . . . . . . . . . . . . . . . . . . . . . 5.1. Speciﬁc off-target effects . . . . . . . . . . . . . . . 5.2. Nonspeciﬁc off-target effects . . . . . . . . . . . . . 6. The future outlook . . . . . . . . . . . . . . . . . . . . . 7. Conclusions . . . . . . . . . . . . . . . . . . . . . . . . Acknowledgements . . . . . . . . . . . . . . . . . . . . . . . References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 746 747 747 747 748 748 750 751 752 753 753 753 754 755 755 756 756 1. Introduction ☆ This review is part of the Advanced Drug Delivery Reviews theme issue on “Towards Therapeutic Application of RNA-mediated Gene Regulation”. ⁎ Corresponding author. 1700 Paciﬁc, Suite 1100, Dallas, Texas 75201, USA. Tel.: +1 214 658 1964; fax: +1 214 658 1992. E-mail addresses: [email protected] (D.D. Rao), [email protected] (J.S. Vorhies), [email protected] (N. Senzer), [email protected] (J. Nemunaitis). 0169-409X/$ – see front matter © 2009 Elsevier B.V. All rights reserved. doi:10.1016/j.addr.2009.04.004 Cancer is a disease of genes, whether based on aberrant changes in sequence or expression (epigenomics). The constellation of genetic and epigenetic abnormalities characterizing cancer cells present new and more speciﬁc targets for cancer treatment and, hopefully, prevention. Over the last decade, the mapping of the human genome, D.D. Rao et al. / Advanced Drug Delivery Reviews 61 (2009) 746–759 along with improved understanding of signal transduction and the pathways responsible for tumor survival, have been transforming therapeutic oncology from a more promiscuously targeted chemotherapeutic approach towards a highly selective targeted therapeusis. Indeed, in 1997, antibody-based Herceptin (trastuzumab) became the ﬁrst targeted therapy for breast cancer, speciﬁcally for HER2-positive metastatic breast cancer. In 2001, the small molecule Gleevec (imatinib mesylate), became the ﬁrst approved kinase inhibitor for cancer targeting bcr-abl in chronic myeloid leukemia (CML), and it has since been approved for the treatment of gastrointestinal stromal tumors (GIST) targeting c-kit. Over the last few years, several other targeted cancer therapies have been approved. Targeted therapeutics directed against ampliﬁed genes and/or over-expressed proteins in malignant cells have proven to be powerful tools for cancer treatment. A growing understanding and use of proteomic, genetic, and pharmacogenomic tools are actualizing a long desired concept of personalized cancer therapy. Genetic abnormalities of each patient's tumor can be analyzed through a variety of established means to quantitatively determine both gene and protein over- and under-expression. Moreover, functional pathways can be determined and integrated within the cancer network allowing for the identiﬁcation of key molecular relays enabling experimental testing of target speciﬁc therapeutics. Such information can potentially allow medical care takers to prioritize, if not yet optimize, treatment for cancer patients, and to uncover surrogate biomarkers for prognosis, prediction, and therapy assessment. The recent discovery of RNA interference (RNAi), a natural process through which the expression of a targeted gene can be knocked down with high speciﬁcity and selectivity, presents an invaluable tool for personalized cancer therapy. Target speciﬁc RNAi agents have the potential to selectively knockdown key abnormally over- or constitutively expressed molecular targets that are essential for the survival of each patient's tumor for effective personalized cancer treatment. Conceptually, target speciﬁc RNAi agents can also be applied in combination with immune modulating agents or small molecules to improve the efﬁcacy of cancer treatment. Like other new therapeutic paradigms, there are a multitude of issues which need to be addressed in order for us to translate RNA interference technology (siRNA, shRNA, bi-functional RNA) from laboratory to bedside and from concept to reality. These issues include comparison of each of the RNAi technologies with respect to effective delivery, possible off-target effects and the pharmacokinetics and pharmacodynamics. This review will focus on several issues currently confronting clinical development of RNAi therapeutics. We will discuss the role of RNAi technology in personalized cancer gene therapy and address clinical considerations of 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 differ not only from their normal counterparts but, to a lesser extent, even from each other based on both intrinsic gene modiﬁcations and modulated cancer cell–matrix interactions. Such variability in genetic patterns found between histologically identical tumors arising in different patients may well explain the widely divergent responses to the standard treatment regimens most often prescribed for a particular tumor type. This is supported by recent studies demonstrating that certain patterns of genetic expression (i.e. expression signatures) identiﬁed in tumor samples from patients with breast cancer are not only strongly correlated with prognosis [1–4] but can actually be subclassiﬁed into differing prognostic categories . The presence of functional redundancy in a robust, predominantly scale-free network such as cancer “buffers” the effect of any single gene/target modiﬁcation on the malignant process, with rare exception (e.g., CML). The hierarchy 747 of cancer scale-free networks does not have a threshold for single target disintegration insofar as random pathway component failure predominantly affects targets with low connectivity within the network, thereby having limited functional impact. However, highly connected information-transfer targets do allow for “attack vulnerability.” In other words, the disordered circuitry characteristic of malignancy results in a change, such that the otherwise robust oncogenic process can become, almost paradoxically, 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 on cancer cells yet not signiﬁcantly perturb normal cell functionality. Genetic diversity of cancer, pathway addiction and targeted therapy are not the subject of this review and have been extensively reviewed elsewhere [6–8]; here, we discuss how RNAi-based therapeutics can be best applied in light of these mechanisms. For example, we harvested tumor and normal cells from the lymph nodes of a melanoma cancer patient for molecular proﬁling by microarray and proteomic analysis . Expression proﬁles for malignant versus normal tissue were compared at the mRNA and protein levels. The goal was to identify a group of gene and protein doublets differentially over-expressed in that individual's malignancy. Fig. 1 is a comparative analysis of protein expression proﬁle by the two-dimensional difference gel electrophoresis (2D-DIGE) analysis. 2D-DIGE can very effectively identify several over-expressed proteins in the patient's tumor. Correlated DNA/RNA over-expression can then be conﬁrmed with microarray data. The resulting data is further analyzed by a modeling and simulation computational system developed by our team speciﬁcally for clinical application including, but not limited to, gene set enrichment analysis and network inference modeling platforms (Fig. 2). Grouped gene expression patterns that are highly correlated with the pathway phenotype in an individual patient allow target genes to be selected, and prioritized based on connectivity and vector-driven criteria developed analytic network algorithms. Once this individual “cancer ﬁngerprint” is created, it serves as the template for the design, synthesis, and subsequent validation of individualized therapeutic RNAi molecules with knockdown activity against these “high-degree hub” genes for a personalized and targeted cancer gene therapy. 2.2. RNA interference for cancer The concept of antisense oligodeoxynucleotides as modulators of gene expression and their application in targeted cancer gene therapy was developed more than 25 years ago (for review, ). By processes still unknown, the antisense nucleic acid (ASNA) strand and the mRNA target come into proximity leading to the destruction of the mRNA target either by endogenous nucleases, such as RNase H [11,12] that are recruited into the mRNA–ASNA duplex or by intrinsic enzymatic activity engineered into the ASNA sequence, as is the case with ribozymes [13,14] and DNAzymes [15,16]. The discovery of an evolutionarily conserved gene silencing mechanism whereby small sequences of extrinsic dsRNA or intrinsic microRNA inhibit complementary post-transcriptional mRNA (siRNA) or suppress translation (miRNA), respectively, ignited strong hope that the natural gene silencing process would be speciﬁc and robust. The silencing process occurs following interaction of the RNA effector precursors with the RNase III enzymes Drosha (for miRNA) and Dicer (for miRNA and siRNA) and subsequent formation of the RNA-interfering silencing complex (RISC) . Endonucleolytic cleavage of the target mRNA occurs at a single site ~10 nucleotides from the 5′ end of the guide (antisense) siRNA sequence [18,19]. RNAi offers several advantages over antisense and ribozyme approaches, including ease of synthesis  and greater activity [18,21–24]. Preclinical studies conﬁrm that RNAi techniques can be used to silence cancer-related targets [25–35]. In vivo studies have also shown favorable outcomes by RNAi targeting of components critical for 748 D.D. Rao et al. / Advanced Drug Delivery Reviews 61 (2009) 746–759 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 a normal lymph node from patient RW following 2-D gel electrophoresis. In the middle of the upper panel the protein expression pattern of a malignant lymph node from patient RW is shown and to the right is an overlaid image with 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 of interest. The middle panel shows the 3D view of one protein spot change between the normal and malignant lymph nodes. Mass spectrometry (lower panel) subsequently (following robotic spot picking) identiﬁed this protein as RACK1. 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 of molecules; the chemically synthesized double-stranded small interfering RNA (siRNA) or vector based short hairpin RNA (shRNA). Effective RNAi was initially demonstrated by the application of synthetic siRNA ; later, siRNA produced in vitro by T7 RNA polymerase was found to be active and it was soon demonstrated that active siRNA consists of a hairpin structure can be transcribed in cells from an RNA polymerase III promoter on a plasmid construct [49,50]. Although siRNA and shRNA can be applied to achieve similar functional outcomes, siRNA and shRNA are intrinsically different molecules. Therefore, the molecular mechanisms of action, the RNA interference pathways, the off-target effects and the applications can also be different. 3.1. siRNA Fluorescent labeled siRNA has been used to trace the fate of delivered siRNA. A ﬂuorescent label can either be tagged onto the 5′ end or the 3′ end of the siRNA. Using a ﬂuorescence resonance energy transfer (FRET)-based visualization method, the intact siRNA can be observed to be translocated into the nucleus within 15 min of the delivery and then disseminated into the cytoplasm within the next 4 h both in intact and dissociated form . The initial accumulation of siRNA in the nuclei is similar to observations made on the behavior of antisense olignucleotides . The translocation of antisense oligonucleotide into the nuclei was not dependent on either the ATP pool or temperature and thus may not involve the active import transport system of the nuclear pore ; it is not clear whether siRNA translocate into the nuclei using the same mechanism as antisense olignucleotides. Using HeLa cells and targeting 7SK snRNA which is exclusively located in the nucleus, Robb showed the efﬁciency of siRNA mediated silencing to be greater than that effected by antisense 7SK DNA . Berezhna et al. observed nuclear localization of siRNA targeted against small nuclear RNA (snRNA) and cytoplasmic localization of siRNA targeted against viral mRNA suggesting selective localization and compartmentalization of siRNA based on its intended target . Ago1 and Ago2 containing RISC were found both in the cytoplasm and nucleus [55–57]. A recent study using ﬂuorescence correlation spectroscopy and ﬂuorescence cross-correlation spectroscopy (FCS/FCCS) to correlate the presence of siRNA with Ago2 D.D. Rao et al. / Advanced Drug Delivery Reviews 61 (2009) 746–759 749 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 . Importin 8 (Imp8) binds to all Ago proteins in a Ran-dependent manner, but independently of RNA . Knockdown of Imp8 results in a shift of Ago2 from the nucleus to the cytoplasm without affecting the total quantity of Ago2. However, although Imp8 is not required for target mRNA cleavage it is necessary for Ago2 binding to miRNA targets. In Caenorhabditis elegans, the argonaute protein NRDE-3 is essential for binding nuclear RNAs and appears to interact with cytoplasmic siRNAs generated by RNA-dependent RNA polymerase (RdRP) followed by redistribution to the nucleus . The nuclear RISC (nRISC) is a complex that is 20× smaller in size than the cytoplasmic RISC (cRISC). The nucleus may be the check point controlling distribution as either the nuclear acting siRNA or the cytoplasmic acting siRNA. Dynamically, siRNA steadily increases its accumulation in cells for 4 h before plateau . The steady-state nuclear distribution of siRNA was mainly found in the nucleolus region and was excluded from the nucleoplasm . The cytoplasmic distribution of siRNA appears to be in the perinuclear region forming a ring-like pattern around the nucleus . The nucleolus and perinuclear regions are possibly the main site for RNAi. However, Ohrt et al. labeled siRNA with ﬂuorescent dye at the 3′ end of either strand of siRNA and did not ﬁnd accumulation of siRNA at the perinuclear region, but rather evenly distributed throughout the cytoplasm . This discrepancy may be the result of the ﬂuorescent tagging process. At 48 h post injection, the majority of siRNA appears to have been degraded with only 1% ﬂuorescence remaining in the cell. The spatial and temporal distribution of siRNA within the cell is in accord with the observed kinetics of siRNA mediated RNA interference activity which peaks around 24 h post delivery and diminishes within 48 h. The life-cycle of siRNA inside transfected cells is diagrammatically illustrated in Fig. 3. In Drosophila, double-stranded RNA-binding proteins (dsRBPs), such as R2D2 and Loquacious (Loqs), function in tandem 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 in the RNAi pathways . In mammalian cells, only one Dicer gene has thus far been identiﬁed . Human Dicer is an integral component of the RNA interference pathway. Dicer processes pre-microRNA and double-strand RNA (dsRNA) to mature miRNA and siRNA, respectively, and transfers the processed products to the RISC [69,70]. Since there is only one Dicer in the human, the RNA-interfering pathway for siRNA and for miRNA may not be as compartmentalized as for Drosophila. Dicer is a multi-domain RNase III-related endonuclease responsible for processing dsRNA to siRNAs . Dicer preferentially binds to the 5′ phosphate of 2 nt 3′ overhang and cleaves dsRNAs into 21 to 22 nucleotide siRNAs [72,73]. Mammalian Dicer interacts with the double-stranded Tat–RNA-binding protein (TRBP) or PACT (PKR activating protein) to mediate RNA interference and miRNA processing. TRBP and PACT are structurally related but exert opposite regulatory activities on RNA-dependent protein kinase (PKR). Knockdown of both TRBP and PACT in cultured cells leads to signiﬁcant inhibition of gene silencing mediated by short hairpin RNA but not by siRNA, suggesting that TRBP and PACT function primarily at the step of siRNA production . Human TRBP and PACT directly interact with 750 D.D. Rao et al. / Advanced Drug Delivery Reviews 61 (2009) 746–759 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. After RISC loading, the passenger strand departs, thereby commencing the RNA interference process via target mRNA cleavage and degradation. siRNA loaded RISCs are also found to be associated 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 of double-stranded or short hairpin RNA to siRNA . Dicer knockout ES cells can effectively load processed siRNA onto RISC and carry out RNA interference as efﬁciently as Dicer+ ES cells . So, it appears that in mammalian cells, a perfectly processed siRNA can be effectively loaded onto RISC for RNAi without the help of the TRBP/PACT/Dicer complex. The TRBP/PACT/Dicer complex, however, is required to process either shRNA or long dsRNA to appropriate size and form for their loading onto RISC. Duplex siRNA in association with holo-RISC, composed of at least Ago-2, Dicer and TRBP, is identiﬁed as the RISC loading complex (RLC) . In the RLC, the two strands of the duplex are separated, resulting in the departure of the passenger strand [76–78]. The passenger strand is cleaved by the RNase-H like activity of Ago-2, provided there are thermodynamically favorable conditions for passenger strand departure. This is referred to as the cleavage-dependent pathway . There is also a cleavage-independent by-pass pathway, in which the passenger strand with mismatches is induced to unwind and depart by an ATP dependent helicase activity [76,79,80]. The RISC with single-stranded guide strand siRNA is then able to execute multiple rounds of RNA interference. ATP is not required for shRNA processing, RISC assembly, cleavage-dependent pathway, or multiple rounds of target-RNA cleavage [81–83]. Single-stranded siRNA (containing 5′-phosphates) and pre-miRNA can be loaded on RISC, but not duplex siRNA . 3.2. shRNA shRNAs, as opposed to siRNAs, are synthesized in the nucleus of cells, further processed and transported to the cytoplasm, and then incorporated into the RISC for activity . The life-cycle of shRNA inside of transfected cells is diagrammatically illustrated in Fig. 4. To be effective, the shRNA are designed to follow the rules predicated by the speciﬁcs of the cellular machinery and are presumably processed similar to the microRNA maturation pathways. Thus, studies on the synthesis and maturation of miRNAs have provided the groundwork for the synthesis of shRNA , particularly the miR-30 based shRNAs . shRNA can be transcribed by either RNA polymerase II or III through RNA polymerase II or III promoters on the expression cassette. The primary transcript generated from RNA polymerase II promoter contains a hairpin like stem-loop structure that is processed in the nucleus by a complex containing the RNase III enzyme Drosha and the double-stranded RNA-binding domain protein DGCR8 . The complex measures the hairpin and allows precise processing of the long primary transcripts into individual shRNAs with a 2 nt 3′ overhang . The processed primary transcript is the pre-shRNA molecule. It is transported to the cytoplasm by exportin 5, a Ran-GTPdependent mechanism [90,91]. In the cytoplasm the pre-shRNA is loaded onto another RNase III complex containing the RNase III enzyme Dicer and TRBP/PACT where the loop of the hairpin is processed off to form a double-stranded siRNA with 2 nt 3′ overhangs [92–94]. The Dicer containing complex then coordinates loading onto the Ago2 protein containing RISC as described earlier for siRNA. PreshRNA has been found to be part of the RLC; thus, pre-shRNA may potentially directly associate with RLC rather than through a two steps process via a different Dicer/TRBP/PACT complex . After loading onto RLC and passenger strand departure; both siRNA and shRNA in the RISC, in principle, should behave the same. The argonaute family of proteins is the major component of RISC [96,97]. Within the Argonaute family of proteins, only Ago2 contains the endonuclease activity necessary to cleave and release the passenger strand of the double-stranded stem [76,77,79]. The remaining three members of Argonaute family, Ago1, Ago3 and Ago4, which do not have identiﬁable endonuclease activity, are also assembled into RISC and presumably function through a cleavage-independent manner. Thus, RISC can be further classiﬁed as cleavage-dependent and cleavage-independent . The argonaute family of proteins in RISCs are not only involved in the loading 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 D.D. Rao et al. / Advanced Drug Delivery Reviews 61 (2009) 746–759 751 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 the nucleus for transcription. The primary transcripts (pre-shRNA) follow a similar route as discovered for the primary transcripts of microRNA. The primary transcripts are processed by the 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 are further processed to mature shRNA. Mature shRNA in the Dicer/TRBP/PACT complex are associated with Argonaute protein containing RISC and provide RNA interference function either through mRNA cleavage and degradation, or through translational suppression via p-bodies. miRNA seeks out complementary target sites in mRNAs, where endonucleolytically active Ago-2 cleaves mRNA to initiate mRNA degradation [98,99]. Other Ago protein containing complexes without endonucleolytic activity predominantly bind to partially complementary target sites located at the 3′ UTR for translation repression through mRNA sequestration in processing bodies (p-bodies) [100–102]. The detailed mechanism of mRNA sequestration in p-bodies and later release from p-bodies is still a debated issue; deadenylation of the target mRNA which leads to destabilization of the mRNA was also observed to occur in p-bodies [103,104]. Coimmunoprecipitation experiments determined that RISCs are also strongly associated with polyribosomes or the small subunit ribosomes  and Ago-2 (actually identiﬁed as elF2c2), strongly suggesting that RISC surveillance is compartmentalized with translational machinery of the cell. Details of the mechanism involving mRNA scanning and target mRNA identiﬁcation are still largely unknown. Whatever the scanning or surveillance mechanism may be, once the target mRNA is identiﬁed, the target mRNA is either cleaved or conformationally changed following which both types of structures are routed to the p-body for either sequestration or degradation [103,104]. The active siRNA or miRNA loaded complex is then released for additional rounds of gene silencing activity. 3.3. Bi-functional shRNA There is, however, a third unique RNAi option in development called bi-functional shRNA. shRNA can potentially be manipulated to take advantage of the gene silencing machinery within the cells to improve its efﬁciency and durability of action. Conceptually, targeted shRNAs can be designed so as to effectively load shRNA onto both the cleavage-dependent and the cleavage-independent RISCs. This differential processing is mediated by two pathways primarily dependent on 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 gene expression silencing (the rate dependent on mRNA turnover and protein kinetics) compared to shRNA as illustrated on Fig. 5. The “bifunctional” shRNA, by virtue of loading onto multiple types of RISCs, is thus able to simultaneously induce degradation of target mRNA and also inhibit translation through mRNA sequestration. This bi-functional design should be, in principal, much more efﬁcient for two reasons; ﬁrst, the bi-functional will promote loading the guide strand onto at least two types of RISCs to increase activity; second, by loading onto both cleavage-dependent RISC and cleavage-independent RISC, target mRNA can be silenced both through mRNA degradation and translational inhibition or sequestration. The design of the bi-functional shRNA expression unit consists of two stem-loop shRNA structures; one stem-loop structure composed of fully matched passenger and guide strand for cleavage-dependent RISC loading, the second stem-loop structure composed of mis-matched passenger strand (at the positions 9–12) for cleavage-independent RISC loading. There are several experimental observations that support this approach. In Drosophila, Ago1 preferentially binds to miRNAs that have been excised from imperfectly paired hairpin precursors, whereas those miRNAs that have near-perfectly paired hairpin precursors are bound by Ago2 [105–108]. In HEK293 cells transfected with tagged-Ago proteins, coimmunoprecipitation found similar sets of about 600 transcripts to be bound to Ago1, 2, 3 or 4 , suggest all four mammalian Ago protein containing RISCs are involved in the RNAi function. Insofar as most mRNA have multiple miRNA target sites (with distance constraints) at their 3′ UTR, the miRNA mediated RNAi system appears to be redundant for the targeted mRNAs allowing for cooperative downregulation to ensure target mRNA knockdown. The bi-functional shRNA approach mimics the natural process by mediating target mRNA knockdown through multiple RNAi pathways and complexes. 752 D.D. Rao et al. / Advanced Drug Delivery Reviews 61 (2009) 746–759 Fig. 5. Schematic of the bi-functional shRNA concept. The bi-functional concept is to design two shRNAs for each targeted mRNA; one with perfect match, one with mismatches at the central location (bases 9–12). The purpose of the bi-functional design is to promote loading of mature shRNAs onto both cleavage-dependent and cleavage-independent RISCs, so that the 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 determine Argonaute loading . Recently, Azuma-Mukai et al. observed miRNAs associated with hAgo-2 and hAgo-3 have some overlaps; however, some are discriminately loaded onto hAgo-2 or hAgo-3 . Further work is needed to resolve the speciﬁcity of miRNA loading onto different Ago containing RISCs. Although most miRNA target sites have been identiﬁed to be located at the 3′-UTR region, recent systemic identiﬁcation of mRNAs recruited to hAgo-2 have identiﬁed many mRNAs with target sites located at the coding region and some at the 5′-UTR . hAgo-2 could initiate the target mRNA degradation with its slicing activity in the coding region. Tay et al. recently found many naturally occurring miRNA targets are located in the coding region of embryonic regulated genes to modulate embryonic stem cell differentiation , further support that miRNA can act through mRNA regions other than 3′-UTR. 3.4. Summary of si/sh/bi In summary, exogenously introduced siRNA with appropriate length (19–21 nt) and 2 nt 3′ overhang, can be loaded onto RISC for RNAi 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 ) appears to improve efﬁcacy . If so, it is possibly because increasing the length of an siRNA duplex with an unprocessed end forces directionality as a result of imposed thermodynamic instability determining the guide strand motif and thereby enhancing its association with Dicer/TRBP/PACT complex for more efﬁcient loading onto RLC . shRNA, on the other hand, assimilates into the endogenous miRNA pathway and in so doing is signiﬁcantly more efﬁcient [115–117]. Additionally, ﬂuorescent tagged siRNA tracing indicated high degradation and turnover of exogenously introduced siRNA. Less than 1% of the introduced duplex remains in the cell 48 h after administration. shRNA can be continuously synthesized by the host 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 shRNA integrated in the host genome is sufﬁcient to provide continual gene knockdown effect (Cleary M, personal communication). The higher dose required for siRNA can further contribute to the off-target effects to be discussed later. Understanding the mechanism and the dynamics of siRNA and shRNA at the cellular and molecular level greatly enhanced the effort in developing therapeutic siRNAs for various diseases. As a result, modiﬁcations can be made to improve the efﬁcacy and stability of RNAi agents. Chemically synthesized siRNA is easier to modify through chemistry; however, bulk manufacturing of complex structures such as modiﬁed siRNA is more expensive. Vector based shRNA relies on the host machinery for expression, but, on the other hand, is more difﬁcult to modify. Modiﬁcation can only be achieved through manipulating expression strategy (e.g. bi-functional shRNA), redesigning shRNA structure, or by varying promoter regulation. The shRNA expression units can be incorporated into varieties of plasmids and viral vectors for delivery and integration. In addition, vector based shRNA expression can also be regulated or induced [118–120]. Numerous siRNAs have been demonstrated to be effective for invivo tumor growth modulations via intratumoral, ex-vivo, or systemic routes of application (For review see [121–123]. Vector based shRNA has, likewise, demonstrated in-vivo effectiveness (For review see [121,123,124]. In-vivo studies employ a variety of delivery methods and may not ensure equivalency of strand biasing; therefore, it is hard to perform direct comparison between siRNA and shRNA. McAnuff et al., using a luciferase expression system, compared the potency of siRNA versus shRNA mediated knockdown in vivo; they found that siRNA and shRNA are equivalent in potency at 10 mg dose; however, on a molar basis, the shRNA was 250 fold more effective than the siRNA . In an effort to assess the potential of RNAi as a therapeutic for hepatitis C (HCV), siRNANS5B was targeted against the nonstructural protein 5B viral polymerase coding region fused with luciferase gene . Luciferase expression in vivo was reduced by 75%. Using a cognate shRNANS5B produced a 92.8% average inhibition over 3 experiments. McCleary et al. incubated shRNA, directed against ﬁreﬂy luciferase and containing 29 mer stems and 2-nt 3′ overhangs, D.D. Rao et al. / Advanced Drug Delivery Reviews 61 (2009) 746–759 with recombinant human Dicer . The resulting 22-nt shRNA products, with predictable guide strands, were compared to identically targeted 21 mer siRNA in HeLa cells. More effective inhibition was seen with the shRNA. In another study of the feasibility of RNAi for treatment of HCV, 19 and 25 bp shRNAs were compared with 19 and 25 bp siRNA directed against the HCV IRES (internal ribosomal entry site) using a luciferase reporter in the AVA5 cell line with stable expression of the HCV subgenomic genotype 1b replicon . The 19 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 lines and in tumor samples from patients with ovarian cancer . The distribution of Dicer mRNA levels was bimodal and 60% of specimens had decreased Dicer mRNA. Furthermore, low Dicer levels were found to be a predictor of reduced disease-speciﬁc survival in multivariate analysis. Of particular interest was the ﬁnding that, compared to siRNA mediated silencing of the galectin-3 gene, poor silencing was achieved with shRNA in ovarian cancer cell lines with low versus high Dicer expression. These data will need to be conﬁrmed and evaluated further. Although there is a global downregulation of miRNA expression in cancer , whether this is, in large part, due to low Dicer expression or to shortened 3′ UTRs with fewer miRNA-binding sites  in highly proliferating tumors with modulated feedback mechanisms is not known. miRNA functionality has been conﬁrmed in the three tumor types (ovary, breast, and lung) evaluated in this study raising questions regarding both qualitative and quantitative issues. As the authors note, there are data correlating high Dicer expression with poor prognostic features in other tumor types [129,130]. In their retrospective evaluation of lung cancer, the role of let-7 as both a regulator of Dicer and ras and their feedback networks could not be assessed [131,132]. The Dicer levels in the tumors used for functional assay of gene silencing are not given nor is there conﬁrmation of the equivalency of strand biasing between the siRNA and shRNA constructs. RNAi therapeutics have been shown to be well tolerated in numerous animal models allowing for transition into the clinic. At least 10 RNAi-based drugs are currently in early phase clinical trials , two of which are cancer related; one targeted against the M2 subunit of ribonucleotide reductase (RRM2)  and the other targeted against tenascin-C . Animal studies with siRNA inhibitors for RRM2 show efﬁcacy  and safety in non-human primates . shRNA for the treatment of hepatitis B was also approved for clinical trial by FDA. Both siRNA and shRNA have their respective advantages and disadvantages from the mechanistic point of view. However, safety of this new therapeutic paradigm is of the utmost importance. Although no signiﬁcant adverse event involving initial RNAi-based clinical trials has been reported, there are concerns over the potential off-target effect of RNAi-based agents which will be discussed separately below. 4. Delivery Efﬁcacy of an RNAi cancer therapeutic is limited by the quantity of the oligomer that effectively enters the tumor cells. In the clinical setting this is primarily dependent on the method of delivery. An ideal delivery vehicle must be able to selectively and differentially target tumors versus normal tissue, homogeneously distribute through the tumor mass and penetrate the tumor cells following systemic administration. If cell entry is mediated by endocytosis the delivery vehicle must negotiate endosomal/lysosomal escape and, in the case of shRNA, the payload must penetrate the nuclear membrane as well. Viral vectors are popular for laboratory delivery of shRNA because of their high transfection efﬁciency and effective integration of exogenous DNA, but they have been losing support in recent years because of concerns over safety and immunogenicity [137,138]. Non-viral polymeric delivery systems, in particular those with biodegradable components, have much better safety proﬁles than their viral 753 counterparts though their transfection efﬁciency is generally lower [139–141]. Non-viral vehicles for delivery of siRNA and shRNA are typically cationic preparations. Their positive charge facilitates complexation with negatively charged nucleic acids and also binding to the negatively charged glycocalyx on external cell membranes promoting endocytosis. Both tumor targeting and cell entry can be enhanced by decoration or complexation of the vehicle with targeting moieties, such as monoclonal antibodies, peptides, small molecule ligands, and aptamers to recognize cell surface markers [124,142]. Once endocytosed, the vehicle's positive charge facilitates early escape from the endosome [143,144]. Though the positive charge of these vehicles improves their transfection efﬁciency, it is also associated with increased toxicity [140,145]. A wide variety of potential vehicles are being developed to address the 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 hybrids of these classes. For instance, there is a cyclodextrin-based cationic polymer which has been used successfully to deliver siRNA targeted to RRM2 in various in vivo cancer models [134,146]. This preparation is currently in Phase I clinical trials. Lipid based nanoparticles are showing potential for the delivery of shRNA and siRNA . Protiva Biotherapeutics and Alnaylam have developed nanoparticles composed of a lipid–PEG conjugate that is capable of encapsulating and protecting nucleic acids for the purpose of systemic delivery. These stable nucleic acid lipid particles (SNALPs) were used in the ﬁrst successful administration of siRNAs to a non-human primate [148,149]. Silence Therapeutics has developed a lipid-based delivery vehicle speciﬁcally designed for siRNA delivery to endothelial cells. This vehicle, called AtuPLEX, contains a mix of cationic and fusogenic lipids [150,151]. This vehicle has been used effectively to knockdown protein kinase N3 in murine prostate and pancreatic cancer models, inhibiting cancer progression [152,153]. More detailed discussions of delivery vehicles for shRNA [124,154] and siRNA [141,155–157] as well as general discussions of organ and tissue speciﬁc RNAi delivery [138,158,159] may be found elsewhere. The magnitude of cytokine induction associated with in vivo delivery of siRNA has been noted to vary widely based on the delivery vehicle used . The well recognized conundrum in cationic nonviral nucleic delivery is that transfection efﬁciency usually correlates with toxicity [161,162]. Effective strategies being pursued to break this correlation include molecular modiﬁcations to shield positive charge and the use of biodegradable polymers [154,163]. 5. Off-target effects Despite initial results showing excellent speciﬁcity in RNAi mediated gene silencing, over the past 5 years many studies have shown that there are multiple speciﬁc and nonspeciﬁc mechanisms through which siRNA and shRNA can cause effects other than the intended mRNA suppression. Unintended effects on gene expression mediated by RNAi are termed “off-target effects.” Speciﬁc off-target effects are mediated by partial sequence complementarity of the RNAi construct to mRNAs other than the intended target. Nonspeciﬁc offtarget effects include a wide variety of immune and toxicity related effects that are intrinsic to the RNAi construct itself or its delivery vehicle. The following provides a brief review to the mechanisms surrounding off-target effects and addresses strategies that are being developed to minimize those effects. 5.1. Speciﬁc off-target effects Expression proﬁling experiments have shown that partial sequence complementarity in the passenger or guide strands of the RNAi construct can produce off-target gene suppression. The ﬁrst group to 754 D.D. Rao et al. / Advanced Drug Delivery Reviews 61 (2009) 746–759 deﬁnitively demonstrate this used an unbiased genome-wide microarray proﬁle to search for downregulated mRNA immediately after transfecting a variety of different siRNA constructs directed to different genes into HeLa cells. This study demonstrated that few off-target genes were regulated in common. Furthermore, the off-target expression patterns observed for each individual siRNA were consistent, with repeated runs revealing the same off-target expression proﬁle. It was also noted that the number and identity of the off-target transcripts was unrelated to the ability of the siRNA to silence the target gene . It has subsequently been shown that both siRNA and shRNA constructs with complementarity in the “seed region” can produce the same offtarget expression proﬁles, even across cell lines, independent of delivery method . Even limited sequence siRNA:mRNA complementarity with the intended target, which is usually less than optimal, can produce offtarget suppression. Off-target silencing effects have been demonstrated in transcripts with complementarity as low as 7 nucleotides with the guide siRNA strand . Due to a variety of known and unknown mechanisms, not all transcripts with this level of similarity are silenced. The location of the region of complementarity within the RNAi construct and the mRNA transcript are important predictors of potential for suppression. Complementarity within nucleotides 2–7 at the 5′ end of either the siRNA passenger or guide strands has been shown to be a key determinant in directing off-target effects . This region of the construct is reminiscent of the “seed” region within miRNA, i.e., a heptameric sequence beginning at the ﬁrst or second position from the 5′ end of the miRNA that is complementary to sites in the mRNA 3′-UTRs, which guide silencing in endogenous RNAi. Indeed, RNAi off-targeting seems to be mechanistically related to an miRNA-like effect in that complementarity between the miRNA seed hexamer and the 3′UTR of the off-targeted mRNA produce effective suppression of gene expression [103,168,169]. Various in vitro and in silico methods are either available or under development for analysis of the off-target effects of a given RNAi construct. Screening can be accomplished effectively using mRNA expression data from transfected cells. Expression proﬁling for offtarget screening must be temporally controlled to ensure observation of primary changes in mRNA levels and not secondary changes as a result of downregulation of target protein expression. Results can then be correlated to qRT-PCR data and to protein expression data through Western blots. Methods for microarray-based off-target screening are reviewed in detail elsewhere [169,170]. Computational methods are also being developed so that RNAi constructs may be more effectively screened prior to in vitro testing. Global complementarity search algorithms such as BLASTn and Smith–Waterman have been shown to be poor measures of off-target potential in an RNAi construct . Methods for in silico screening of RNAi constructs must be optimized to consider seed complementarity in the constructs as well as 3′ UTR complementarity in the transcriptome. Several groups are working to develop algorithms and some are freely available online [171–174]. The speciﬁc off-target effects of a given construct can be mitigated by several methods. siRNAs with an asymmetric unilateral 2-nt-overhang on the antisense strand have greater potency than conventional siRNAs as well as reduced off-target effects due to preferential strand selection . Sequence based modiﬁcations designed to reduce speciﬁc offtarget effects are likely to beneﬁt both siRNA and shRNA approaches. Single and double base mismatches between an RNAi construct and its target transcript are often tolerated without reducing the potency of suppression [168,175–177]. This allows for the optimization of the construct sequence to minimize complementarity with 3′UTRs of unintended targets. Although the vector-driven shRNA approach to RNAi does not permit speciﬁc chemical modiﬁcation of the silencing construct, siRNA oligomers can be chemically modiﬁed in order to reduce direct offtarget effects. These modiﬁcations must be carefully applied as reductions in off-target effects are sometimes accompanied by deceases in the overall potency of suppression. [178–180]. Chemical modiﬁcations can also be asymmetrically applied to the passenger strand of the siRNA construct in order to speciﬁcally inhibit its participation in silencing. Annealing of the guide strand to a passenger strand composed of two segments has been shown to reduce off-target effects mediated by passenger strand complementarity . The addition of other chemical moieties to one or both strands can also limit speciﬁc off-target effects. The 5′-phosphorylation status of the siRNA strands has been shown to be a determinant of which strand is involved in silencing, and thus replacement of the 5′ phosphate by of an siRNA strand with a methyl group has been shown to reduce its participation in silencing . Chemical modiﬁcations to nucleotides within the seed region of the passenger or guide strand can reduce unintended entry of siRNA constructs into the endogenous miRNA gene silencing pathway by inhibiting the interaction of the RISC complex with mRNA. It has been shown that 2′-O-methyl ribosyl substitution at position 2 in the siRNA guide strand can reduce off-target silencing of transcripts with complementarity to the seed region of the siRNA guide strand . Asymmetric replacement of seed region nucleotides with DNA bases has also been shown to reduce off-targeting as a result of seed region complementarity within the passenger strand . Recent in vitro studies have shown that shRNA produces fewer offtarget effects than siRNA. In one study shRNA and siRNA of the same core sequence directed towards TP53 were applied to HCT-116 colon carcinoma cells in concentrations necessary to achieve comparable levels of target knockdown. Microarray proﬁling demonstrated a much higher degree of up- and downregulation of off-target transcripts in the siRNA transfected cells (M. Mehaffey, T. Ward, and M. Cleary, in prep.). It has been suggested that these differences arise from the fact that shRNA is transcribed in the nucleus and is therefore subject to endogenous processing and regulatory mechanisms. Additionally, siRNA is more susceptible to degradation in the cytoplasm, which may also lead to off-target silencing . 5.2. Nonspeciﬁc off-target effects Nonspeciﬁc off-target effects are those unintended perturbations in gene expression not resulting from the direct interaction of an RNAi construct with an mRNA transcript. Included in this category are interferon (IFN) and other immune mediated responses to exogenous RNAi, cellular toxicities due to the nucleotide construct, and effects related to the delivery vehicle. There is strong reason to believe that shRNA and siRNA would have very different proﬁles of nonspeciﬁc offtarget effects because of the mechanistic differences between the two approaches. Introduction of dsRNA longer than 29–30 bp into mammalian cells results in a potent induction of the innate immune system via PKR, similar to the mammalian cell defense mechanism against viral infection . Activation of the innate immune response by receptors sensitive to exogenous nucleic acids leads to global degradation of mRNA and thus broad inhibition of translation as well as global upregulation of IFN-stimulated gene expression. Although shorter siRNA constructs have been shown to avoid receptor activation and were initially considered to be non-immunogenic , subsequent in vitro data have shown that the introduction of both synthetic siRNA oligomers 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 of exogenous RNAi is likely mediated through several cytoplasmic and endosomal mechanisms attuned to recognize exogenous nucleic acids from infectious agents. The relevant mediators of nucleic acidstimulated immunoactivation at the level of the endosome are Tolllike receptors (TLRs) 7 and 8 (typically activated by ssRNA), TLR 9 (via D.D. Rao et al. / Advanced Drug Delivery Reviews 61 (2009) 746–759 unmethylated CpG activation) and TLR 3 (via dsRNA) [189–193]. Immune activation by nucleic acids at the cytoplasmic level is mediated through RNA sensing receptors such as RIG-I and MDA-5 [114,194]. Mechanisms surrounding immune activation by RNAi are reviewed more thoroughly elsewhere [160,195]. Recently, naked siRNA has been shown to activate TLR-3 on the surface of vascular endothelial cells and trigger the release of IFN-γ and IL-12 that mediate nonspeciﬁc anti-angiogenic effects in-vivo . Immune stimulation often is largely responsible for the observed therapeutic effects of siRNA rather than the direct targeted effect . Misinterpreting the therapeutic effect of siRNA needs to be carefully monitored. Immune activation at the endosomal level is more readily avoidable by shRNA constructs because the construct is presented on a DNA plasmid obviating dsRNA activation of TLR 3. However, TLR 9, as noted, is activated by unmethylated CpG motifs, which are typically found in bacterial DNA [198,199]. Careful shRNA-encoding plasmid design, avoiding unmethylated CpG motifs, can effectively attenuate if not eliminate TLR 9 mediated endosomal immunoactivation . There is also reason to believe that shRNA is less likely to induce an inﬂammatory response through cytoplasmic dsRNA receptors in vivo because shRNA is spliced by endogenous mechanisms. It has been suggested that the 5′ ends of the endogenous-dicer spliced RNA oligomers are less immunogenic than the 5′ ends of exogenous siRNA oligomers [114,194]. This has been supported in vitro in an experiment that 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 siRNA induced IFN-alpha and type I IFN genes, while the shRNA of the same sequence did not induce an immune response . Sequence modiﬁcations can be made to shRNA or siRNA in order to reduce immunogenicity. It has been shown that endosomal immunoactivation by siRNA through TLR7 and TLR8 can be sequence dependant [189,190,202]. Some simple sequence modiﬁcations, such as the introduction of G:U mismatches into the sequence also seem to lower the IFN response in vitro [203,204]. One recent study showed that a marked reduction in the expression of the interferon-stimulated gene oligoadenylate synthetase 1 (Oas1) could be achieved by modifying the shRNA to contain features of the naturally occurring microRNA-30 (miR-30) precursor . As in the case of speciﬁc off-target effects, chemical modiﬁcation to siRNA oligomers can make them less immunostimulatory, however these modiﬁcations must be ﬁne-tuned so as not to negatively affect the potency of intended target silencing. Suppression of the TLR mediated immune response has been achieved by substituting the 2′hydroxyl uridines in the construct with 2′-ﬂuoro, 2′-deoxy, or 2′-Omethyl uridines [206–208], the products of which do retain targetsilencing potency while reducing immunogenicity. Usage of endogenous processing systems gives shRNA an advantage over siRNA in terms of its propensity for induction of IFN but its over-saturation of these systems has been shown to have other consequences that are more easily avoided by siRNA. In a key in vivo study of the safety effects of long term expression of shRNA in the livers of adult mice, a type 8 adeno-associated virus with a Pol III promoter was used to drive expression of 49 different shRNAs of different lengths and sequences directed against six targets. 36 of the constructs tested resulted in a dose dependant liver injury that was determined to be associated with the downregulation of critical endogenous miRNAs. The degree of miRNA downregulation was related neither to the shRNA sequence nor to the degree of downregulation of the target mRNA . Subsequent transfection studies suggested the degree of miRNA downregulation to result from a competitive bottleneck in shared miRNA/shRNA processing, most likely at the level of exportin-5 (the nuclear membrane export protein used to transfer pre-miRNAs into the cytoplasm) and the RISC component, Argonaute-2 . A similar in vivo study of hepatic 755 toxicity of siRNAs in mice and hamsters showed that systemic introduction of synthetic siRNAs does not result in the suppression of endogenous miRNA levels. In this study siRNAs targeting two hepatocyte-speciﬁc genes (apolipoprotein B and factor VII) and a scramble control were administered to mice and one siRNA targeting the hepatocyte-expressed gene Scap was administered to hamsters. Robust suppression of target genes was achieved in all cases. No changes in levels of the hepatocyte endogenously expressed miR-122 were observed in the mice or the hamsters and no changes in the broadly expressed miRNAs, miR-16 and let-7a, were observed in the mice . Though siRNA likely does not compete with endogenous miRNA for processing proteins, care must be taken when using shRNA as an effector of RNAi in order to minimize the potential for damage mediated by over-saturation of exportin-5. In one in vitro study, overexpression of the exportin-5 protein has been shown to eliminate the nuclear export bottleneck and allow cells to tolerate higher dosages of shRNA without toxicity . Another study showed that an adenoassociated virus construct using a Pol-II promoter was able to achieve stable target gene suppression at high shRNA doses for over 1 year after the initial dosing . In order to minimize the risk of toxicity from over-saturation of miRNA professing systems, data to date suggest both selective promoter integration (e.g. pol II versus pol III) and limiting the dosage of shRNA so as to stay below the threshold of competitive inhibition of the endogenous miRNA biogenesis machinery. 6. The future outlook The ability to precisely and differentially target functionally biorelevant molecular signals in patient's cancers will establish a new paradigm in cancer management; one which focuses on deﬁning the uniqueness of each patient's tumor and tumor-host processes and interactions following rational target prioritization using computational systems biology algorithms. This, then, would allow for exploitation of the “attack vulnerability” of the rewired cancer network by deconstructing essential hubs and linkages, multiply targeting and eliminating them. Both siRNA and shRNA effectors are attractive opportunities. The capability of potentiating activity using a bi-functional design may further enhance safety and efﬁcacy. The simplicity of siRNA manufacturing and the transient nature of the effect per dose may be optimal for certain medical disorders in which high vector doses are required, e.g. some of the viral infections, however, by using the endogenous processing machinery, optimized shRNA constructs allow for high potency sustainable effects using low-copy numbers resulting in less off-target effects (particularly if embedded in an miRNA scaffold) thereby ensuring greater safety. Though shRNA seems ideal for cancer-related therapeutic development, new technology such as bi-functional RNA interference may provide an even greater opportunity for enhancement in potency as well as heightening safety thereby increasing the opportunities for multiple target therapy. This, of course, is contingent on optimization of delivery and minimization of off-target effect which will need to be established through early clinical testing. 7. Conclusions Our understanding and application of RNAi has dramatically advanced over the last 5 years. Despite limitations in developing effective delivery vehicles and concerns regarding potential off-target activity, clinical development has been initiated. As the science of this ﬂedgling technology advances, it is evident that issues such as target selection, effector potency, delivery vehicle design, and off-target effects will continue to be addressed and resolved. Bi-functional RNAi products are evolving components in this transition to clinically effective and safer therapeutics. 756 D.D. Rao et al. / Advanced Drug Delivery Reviews 61 (2009) 746–759 Acknowledgements The authors would like to acknowledge Brenda Marr and Susan Mill for their competent and knowledgeable assistance in the preparation of this manuscript, and M. Cleary for providing the data before publication. References  V.G. Kaklamani, W.J. Gradishar, Gene expression in breast cancer, Curr. Treatm. Opt. Oncol. 7 (2006) 123–128.  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 cancer outcomes, Evid. Rep./Technol. Assess. (Full Rep.) (2007) 1–105.  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 is an independent predictor of clinical outcome, Int. J. Cancer. 124 (7) (2009 Apr 1) 1552–1564.  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 a uniﬁed understanding of breast cancer subtyping and prognosis signatures, Breast Cancer Res. 10 (2008) R65.  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. BorresenDale, D. Botstein, Repeated observation of breast tumor subtypes in independent gene expression data sets, Proc. Natl. Acad. Sci. U. S. A. 100 (2003) 8418–8423.  M. Davies, B. Hennessy, G.B. Mills, Point mutations of protein kinases and individualised cancer therapy, Expert Opin. Pharmacother. 7 (2006) 2243–2261.  A.G. Letai, Diagnosing and exploiting cancer's addiction to blocks in apoptosis, Nat. Rev., Cancer 8 (2008) 121–132.  O.I. Olopade, T.A. Grushko, R. Nanda, D. Huo, Advances in breast cancer: pathways to personalized medicine, Clin. Cancer Res. 14 (2008) 7988–7999.  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 personalized cancer therapeutics, Cancer Gene Ther. 14 (2007) 686–695.  C.A. Stein, J.S. Cohen, Oligodeoxynucleotides as inhibitors of gene expression: a review, Cancer Res. 48 (1988) 2659–2668.  E. Zamaratski, P.I. Pradeepkumar, J. Chattopadhyaya, A critical survey of the structure–function of the antisense oligo/RNA heteroduplex as substrate for RNase H, J. Biochem. Biophys. Methods 48 (2001) 189–208.  S.T. Crooke, Molecular mechanisms of action of antisense drugs, Biochim. Biophys. Acta 1489 (1999) 31–44.  D. Castanotto, M. Scherr, J.J. Rossi, Intracellular expression and function of antisense catalytic RNAs, Methods Enzymol. 313 (2000) 401–420.  J.J. Rossi, Ribozymes in the nucleolus, Science 285 (1999) 1685.  S.W. Santoro, G.F. Joyce, A general purpose RNA-cleaving DNA enzyme, Proc. Natl. Acad. Sci. U. S. A. 94 (1997) 4262–4266.  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.  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.  S.M. Elbashir, J. Harborth, W. Lendeckel, A. Yalcin, K. Weber, T. Tuschl, Duplexes of 21-nucleotide RNAs mediate RNA interference in cultured mammalian cells, Nature 411 (2001) 494–498.  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 of mammalian RNAi, Science 305 (2004) 1437–1441.  Z. Paroo, D.R. Corey, Challenges for RNAi in vivo, Trends Biotechnol. 22 (2004) 390–394.  Y. Aoki, D.P. Cioca, H. Oidaira, J. Kamiya, K. Kiyosawa, RNA interference may be more potent than antisense RNA in human cancer cell lines, Clin. Exp. Pharmacol. Physiol. 30 (2003) 96–102.  B.L. Bass, RNA interference. The short answer, Nature 411 (2001) 428–429.  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 oligonucleotides against survivin RNA to inhibit steps leading to tumor angiogenesis, Oligonucleotides 14 (2004) 100–113.  M. Miyagishi, M. Hayashi, K. Taira, Comparison of the suppressive effects of antisense oligonucleotides and siRNAs directed against the same targets in mammalian cells, Antisense Nucleic Acid Drug Dev. 13 (2003) 1–7.  M. Scherr, K. Battmer, T. Winkler, O. Heidenreich, A. Ganser, M. Eder, Speciﬁc inhibition of bcr-abl gene expression by small interfering RNA, Blood 101 (2003) 1566–1569.  T.R. Brummelkamp, R. Bernards, R. Agami, Stable suppression of tumorigenicity by virus-mediated RNA interference, Cancer Cells 2 (2002) 243–247.  L.A. Martinez, I. Naguibneva, H. Lehrmann, A. Vervisch, T. Tchenio, G. Lozano, A. Harel-Bellan, Synthetic small inhibiting RNAs: efﬁcient tools to inactivate oncogenic mutations and restore p53 pathways, Proc. Natl. Acad. Sci. U. S. A. 99 (2002) 14,849–14,854.  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 16positive cervical cancer cells by E6 siRNA, Mol. Ther. 8 (2003) 762–768.  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/neu gene, upregulates HLA class I and induces apoptosis of Her2/neu positive tumor cell lines, Int. J. Cancer 108 (2004) 71–77.  G. Yang, K.Q. Cai, J.A. Thompson-Lanza, R.C. Bast Jr., J. Liu, Inhibition of breast and ovarian tumor growth through multiple signaling pathways by using retrovirusmediated small interfering RNA against Her-2/neu gene expression, J. Biol. Chem. 279 (2004) 4339–4345.  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 novel therapies directed against protein kinase A, Surgery 134 (2003) 197–205.  E. Yague, C.F. Higgins, S. Raguz, Complete reversal of multidrug resistance by stable expression of small interfering RNAs targeting MDR1, Gene Ther.11 (2004) 1170–1174.  B.A. Kosciolek, K. Kalantidis, M. Tabler, P.T. Rowley, Inhibition of telomerase activity in human cancer cells by RNA interference, Mol. Cancer Ther. 2 (2003) 209–216.  D.P. Cioca, Y. Aoki, K. Kiyosawa, RNA interference is a functional pathway with therapeutic potential in human myeloid leukemia cell lines, Cancer Gene Ther. 10 (2003) 125–133.  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 of human cells, Nucleic Acids Res. 31 (2003) 700–707.  K. Li, S.Y. Lin, F.C. Brunicardi, P. Seu, Use of RNA interference to target cyclin E-overexpressing hepatocellular carcinoma, Cancer Res. 63 (2003) 3593–3597.  U.N. Verma, R.M. Surabhi, A. Schmaltieg, C. Becerra, R.B. Gaynor, Small interfering RNAs directed against beta-catenin inhibit the in vitro and in vivo growth of colon cancer cells, Clin. Cancer Res. 9 (2003) 1291–1300.  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 and small interfering RNAs suppresses growth of human mammary tumor xenografts in mice, Cancer Res. 64 (2004) 5378–5384.  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 against survivin induced apoptosis and attenuated tumor cell growth in vitro and in vivo, Mol. Ther. 10 (2004) 162–171.  A.J. Salisbury, V.M. Macaulay, Development of molecular agents for IGF receptor targeting, Horm. Metab. Res. 35 (2003) 843–849.  M.S. Duxbury, H. Ito, M.J. Zinner, S.W. Ashley, E.E. Whang, Focal adhesion kinase gene silencing promotes anoikis and suppresses metastasis of human pancreatic adenocarcinoma cells, Surgery 135 (2004) 555–562.  M.S. Duxbury, E. Matros, H. Ito, M.J. Zinner, S.W. Ashley, E.E. Whang, Systemic siRNA-mediated gene silencing: a new approach to targeted therapy of cancer, Ann. Surg. 240 (2004) 667–674 [discussion 675–666].  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 growth factor severely limits tumor resistance to antiangiogenic thrombospondin-1 and slows tumor vascularization and growth, Cancer Res. 63 (2003) 3919–3922.  Y. Takei, K. Kadomatsu, Y. Yuzawa, S. Matsuo, T. Muramatsu, A small interfering RNA targeting vascular endothelial growth factor as cancer therapeutics, Cancer Res. 64 (2004) 3365–3370.  S.S. Lakka, C.S. Gondi, N. Yanamandra, W.C. Olivero, D.H. Dinh, M. Gujrati, J.S. Rao, Inhibition of cathepsin B and MMP-9 gene expression in glioblastoma cell line via RNA interference reduces tumor cell invasion, tumor growth and angiogenesis, Oncogene 23 (2004) 4681–4689.  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 2 gene expression in non-small cell lung cancer inhibits tumor growth and increases efﬁcacy of chemotherapy, Cancer Res. 68 (2008) 7975–7984.  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 M1 subunit overexpression in gemcitabine resistance of human pancreatic cancer, Int. J. Cancer 120 (2007) 1355–1363.  A. Fire, S. Xu, M.K. Montgomery, S.A. Kostas, S.E. Driver, C.C. Mello, Potent and speciﬁc genetic interference by double-stranded RNA in Caenorhabditis elegans, Nature 391 (1998) 806–811.  J.Y. Yu, S.L. DeRuiter, D.L. Turner, RNA interference by expression of short-interfering RNAs and hairpin RNAs in mammalian cells, Proc. Natl. Acad. Sci. U. S. A. 99 (2002) 6047–6052.  M. Miyagishi, K. Taira, U6 promoter-driven siRNAs with four uridine 3′ overhangs efﬁciently suppress targeted gene expression in mammalian cells, Nat. Biotechnol. 20 (2002) 497–500.  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 integrity by FRET imaging, Nucleic Acids Res. 35 (2007) e124.  J.P. Leonetti, N. Mechti, G. Degols, C. Gagnor, B. Lebleu, Intracellular distribution of microinjected antisense oligonucleotides, Proc. Natl. Acad. Sci. U. S. A. 88 (1991) 2702–2706.  G.B. Robb, K.M. Brown, J. Khurana, T.M. Rana, Speciﬁc and potent RNAi in the nucleus of human cells, Nat. Struct. Mol. Biol. 12 (2005) 133–137.  S.Y. Berezhna, L. Supekova, F. Supek, P.G. Schultz, A.A. Deniz, siRNA in human cells selectively localizes to target RNA sites, Proc. Natl. Acad. Sci. U. S. A. 103 (2006) 7682–7687. D.D. Rao et al. / Advanced Drug Delivery Reviews 61 (2009) 746–759  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 in mammalian transcriptional silencing, Nat. Struct. Mol. Biol. 13 (2006) 787–792.  D.H. Kim, L.M. Villeneuve, K.V. Morris, J.J. Rossi, Argonaute-1 directs siRNA-mediated transcriptional gene silencing in human cells, Nat. Struct. Mol. Biol. 13 (2006) 793–797.  S. Rudel, A. Flatley, L. Weinmann, E. Kremmer, G. Meister, A multifunctional human Argonaute2-speciﬁc monoclonal antibody, RNA 14 (2008) 1244–1253.  T. Ohrt, J. Mutze, W. Staroske, L. Weinmann, J. Hock, K. Crell, G. Meister, P. Schwille, Fluorescence correlation spectroscopy and ﬂuorescence cross-correlation spectroscopy reveal the cytoplasmic origination of loaded nuclear RISC in vivo in human cells, Nucleic Acids Res. 36 (2008) 6439–6449.  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 targets argonaute proteins to distinct mRNAs, Cell 136 (2009) 496–507.  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.  A. Grunweller, C. Gillen, V.A. Erdmann, J. Kurreck, Cellular uptake and localization of a Cy3-labeled siRNA speciﬁc for the serine/threonine kinase Pim-1, Oligonucleotides 13 (2003) 345–352.  T. Ohrt, D. Merkle, K. Birkenfeld, C.J. Echeverri, P. Schwille, In situ ﬂuorescence analysis demonstrates active siRNA exclusion from the nucleus by exportin 5, Nucleic Acids Res. 34 (2006) 1369–1380.  Y.L. Chiu, A. Ali, C.Y. Chu, H. Cao, T.M. Rana, Visualizing a correlation between siRNA localization, cellular uptake, and RNAi in living cells, Chem. Biol. 11 (2004) 1165–1175.  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 stem cell maintenance requires Loquacious, a double-stranded RNA-binding domain protein, PLoS. Biol. 3 (2005) e236.  F. Jiang, X. Ye, X. Liu, L. Fincher, D. McKearin, Q. Liu, Dicer-1 and R3D1-L catalyze microRNA maturation in Drosophila, Genes Dev. 19 (2005) 1674–1679.  K. Saito, A. Ishizuka, H. Siomi, M.C. Siomi, Processing of pre-microRNAs by the Dicer-1-Loquacious complex in Drosophila cells, PLoS. Biol. 3 (2005) e235.  X. Liu, J.K. Park, F. Jiang, Y. Liu, D. McKearin, Q. Liu, Dicer-1, but not Loquacious, is critical for assembly of miRNA-induced silencing complexes, RNA 13 (2007) 2324–2329.  E.P. Murchison, J.F. Partridge, O.H. Tam, S. Chelouﬁ, G.J. Hannon, Characterization of Dicer-deﬁcient murine embryonic stem cells, Proc. Natl. Acad. Sci. U. S. A. 102 (2005) 12,135–12,140.  E. Bernstein, A.A. Caudy, S.M. Hammond, G.J. Hannon, Role for a bidentate ribonuclease in the initiation step of RNA interference, Nature 409 (2001) 363–366.  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.  S.M. Hammond, A.A. Caudy, G.J. Hannon, Post-transcriptional gene silencing by double-stranded RNA, Nat. Rev., Genet. 2 (2001) 110–119.  M.A. Carmell, G.J. Hannon, RNase III enzymes and the initiation of gene silencing, Nat. Struct. Mol. Biol. 11 (2004) 214–218.  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, Science 311 (2006) 195–198.  K.H. Kok, M.H. Ng, Y.P. Ching, D.Y. Jin, Human TRBP and PACT directly interact with each other and associate with dicer to facilitate the production of small interfering RNA, J. Biol. Chem. 282 (2007) 17,649–17,657.  G.B. Robb, T.M. Rana, RNA helicase A interacts with RISC in human cells and functions in RISC loading, Mol. Cell 26 (2007) 523–537.  C. Matranga, Y. Tomari, C. Shin, D.P. Bartel, P.D. Zamore, Passenger–strand cleavage facilitates assembly of siRNA into Ago2-containing RNAi enzyme complexes, Cell 123 (2005) 607–620.  T.A. Rand, S. Petersen, F. Du, X. Wang, Argonaute2 cleaves the anti-guide strand of siRNA during RISC activation, Cell 123 (2005) 621–629.  P.J. Leuschner, S.L. Ameres, S. Kueng, J. Martinez, Cleavage of the siRNA passenger strand during RISC assembly in human cells, EMBO Rep. 7 (2006) 314–320.  J.B. Preall, E.J. Sontheimer, RNAi: RISC gets loaded, Cell 123 (2005) 543–545.  B. Haley, P.D. Zamore, Kinetic analysis of the RNAi enzyme complex, Nat. Struct. Mol. Biol. 11 (2004) 599–606.  R.I. Gregory, T.P. Chendrimada, N. Cooch, R. Shiekhattar, Human RISC couples microRNA biogenesis and posttranscriptional gene silencing, Cell 123 (2005) 631–640.  E. Maniataki, Z. Mourelatos, A human, ATP-independent, RISC assembly machine fueled by pre-miRNA, Genes Dev. 19 (2005) 2979–2990.  R.E. Collins, X. Cheng, Structural and biochemical advances in mammalian RNAi, J. Cell. Biochem. 99 (2006) 1251–1266.  R.L. Boudreau, A.M. Monteys, B.L. Davidson, Minimizing variables among hairpinbased RNAi vectors reveals the potency of shRNAs, RNA 14 (2008) 1834–1844.  B.R. Cullen, RNAi the natural way, Nat. Genet. 37 (2005) 1163–1165.  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.  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, Secondgeneration shRNA libraries covering the mouse and human genomes, Nat. Genet. 37 (2005) 1281–1288. 757  Y. Lee, C. Ahn, J. Han, H. Choi, J. Kim, J. Yim, J. Lee, P. Provost, O. Radmark, S. Kim, V.N. Kim, The nuclear RNase III Drosha initiates microRNA processing, Nature 425 (2003) 415–419.  H. Zhang, F.A. Kolb, V. Brondani, E. Billy, W. Filipowicz, Human Dicer preferentially cleaves dsRNAs at their termini without a requirement for ATP, EMBO J. 21 (2002) 5875–5885.  Y. Lee, K. Jeon, J.T. Lee, S. Kim, V.N. Kim, MicroRNA maturation: stepwise processing and subcellular localization, EMBO J. 21 (2002) 4663–4670.  B.R. Cullen, Transcription and processing of human microRNA precursors, Mol. Cell 16 (2004) 861–865.  R. Yi, Y. Qin, I.G. Macara, B.R. Cullen, Exportin-5 mediates the nuclear export of pre-microRNAs and short hairpin RNAs, Genes Dev. 17 (2003) 3011–3016.  E. Lund, S. Guttinger, A. Calado, J.E. Dahlberg, U. Kutay, Nuclear export of microRNA precursors, Science 303 (2004) 95–98.  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 silencing pathways, Cell 117 (2004) 69–81.  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-containing ribonucleoprotein complexes and their bound target mRNAs, RNA 14 (2008) 2580–2596.  S.M. Hammond, S. Boettcher, A.A. Caudy, R. Kobayashi, G.J. Hannon, Argonaute2, a link between genetic and biochemical analyses of RNAi, Science 293 (2001) 1146–1150.  E.J. Sontheimer, R.W. Carthew, Molecular biology. Argonaute journeys into the heart of RISC, Science 305 (2004) 1409–1410.  G. Hutvagner, P.D. Zamore, A microRNA in a multiple-turnover RNAi enzyme complex, Science 297 (2002) 2056–2060.  S. Yekta, I.H. Shih, D.P. Bartel, MicroRNA-directed cleavage of HOXB8 mRNA, Science 304 (2004) 594–596.  D.T. Humphreys, B.J. Westman, D.I. Martin, T. Preiss, MicroRNAs control translation initiation 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.  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 MicroRNA in human cells, Science 309 (2005) 1573–1576.  R. Thermann, M.W. Hentze, Drosophila miR2 induces pseudo-polysomes and inhibits translation initiation, Nature 447 (2007) 875–878.  M.A. Valencia-Sanchez, J. Liu, G.J. Hannon, R. Parker, Control of translation and mRNA degradation by miRNAs and siRNAs, Genes Dev. 20 (2006) 515–524.  R. Parker, U. Sheth, P bodies and the control of mRNA translation and degradation, Mol. Cell 25 (2007) 635–646.  K. Okamura, A. Ishizuka, H. Siomi, M.C. Siomi, Distinct roles for Argonaute proteins in small RNA-directed RNA cleavage pathways, Genes Dev. 18 (2004) 1655–1666.  K. Miyoshi, H. Tsukumo, T. Nagami, H. Siomi, M.C. Siomi, Slicer function of Drosophila Argonautes and its involvement in RISC formation, Genes Dev. 19 (2005) 2837–2848.  Y. Tomari, T. Du, P.D. Zamore, Sorting of Drosophila small silencing RNAs, Cell 130 (2007) 299–308.  K. Forstemann, M.D. Horwich, L. Wee, Y. Tomari, P.D. Zamore, Drosophila microRNAs are sorted into functionally distinct argonaute complexes after production by dicer-1, Cell 130 (2007) 287–297.  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 loading in Caenorhabditis elegans, Nat. Struct. Mol. Biol. 14 (2007) 927–933.  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 in RNA silencing, Proc. Natl. Acad. Sci. U. S. A. 105 (2008) 7964–7969.  D.G. Hendrickson, D.J. Hogan, D. Herschlag, J.E. Ferrell, P.O. Brown, Systematic identiﬁcation of mRNAs recruited to argonaute 2 by speciﬁc microRNAs and corresponding changes in transcript abundance, PLoS ONE 3 (2008) e2126.  Y. Tay, J. Zhang, A.M. Thomson, B. Lim, I. Rigoutsos, MicroRNAs to Nanog, Oct4 and Sox2 coding regions modulate embryonic stem cell differentiation, Nature 455 (2008) 1124–1128.  M. Sano, M. Sierant, M. Miyagishi, M. Nakanishi, Y. Takagi, S. Sutou, Effect of asymmetric terminal structures of short RNA duplexes on the RNA interference activity and strand selection, Nucleic Acids Res. 36 (2008) 5812–5821.  D.H. Kim, M.A. Behlke, S.D. Rose, M.S. Chang, S. Choi, J.J. Rossi, Synthetic dsRNA Dicer substrates enhance RNAi potency and efﬁcacy, Nat. Biotechnol. 23 (2005) 222–226.  M.A. McAnuff, G.R. Rettig, K.G. Rice, Potency of siRNA versus shRNA mediated knockdown in vivo, J. Pharm. Sci. 96 (2007) 2922–2930.  D. Siolas, C. Lerner, J. Burchard, W. Ge, P.S. Linsley, P.J. Paddison, G.J. Hannon, M.A. Cleary, Synthetic shRNAs as potent RNAi triggers, Nat. Biotechnol. 23 (2005) 227–231.  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 of sequence and structural features, and comparision with siRNA, Oligonucleotides 17 (2007) 223–236.  M. Gossen, H. Bujard, Tight control of gene expression in mammalian cells by tetracycline-responsive promoters, Proc. Natl. Acad. Sci. U. S. A. 89 (1992) 5547–5551.  S. Gupta, R.A. Schoer, J.E. Egan, G.J. Hannon, V. Mittal, Inducible, reversible, and stable RNA interference in mammalian cells, Proc. Natl. Acad. Sci. U. S. A. 101 (2004) 1927–1932.  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 microRNA precursors, Nat. Genet. 37 (2005) 1289–1295. 758 D.D. Rao et al. / Advanced Drug Delivery Reviews 61 (2009) 746–759  A.W. Tong, Y.A. Zhang, J. Nemunaitis, Small interfering RNA for experimental cancer therapy, Curr. Opin. Mol. Ther. 7 (2005) 114–124.  A. Aigner, Cellular delivery in vivo of siRNA-based therapeutics, Curr. Pharm. Des. 14 (2008) 3603–3619.  A.W. Tong, C.M. Jay, N. Senzer, P.B. Maples, J. Nemunaitis, Systemic therapeutic gene delivery for cancer: crafting Paris' arrow, Curr. Gene Ther. 9 (1) (2009 Feb) 45–60.  J.S. Vorhies, J. Nemunaitis, Nonviral delivery vehicles for use in short hairpin RNAbased cancer therapies, Expert Rev. Anticancer Ther. 7 (2007) 373–382.  A.P. McCaffrey, L. Meuse, T.T. Pham, D.S. Conklin, G.J. Hannon, M.A. Kay, RNA interference in adult mice, Nature 418 (2002) 38–39.  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 in patients with ovarian cancer, N. Engl. J. Med. 359 (2008) 2641–2650.  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.  R. Sandberg, J.R. Neilson, A. Sarma, P.A. Sharp, C.B. Burge, Proliferating cells express mRNAs with shortened 3′ untranslated regions and fewer microRNA target sites, Science 320 (2008) 1643–1647.  S. Chiosea, E. Jelezcova, U. Chandran, M. Acquafondata, T. McHale, R.W. Sobol, R. Dhir, Up-regulation of dicer, a component of the MicroRNA machinery, in prostate adenocarcinoma, Am. J. Pathol. 169 (2006) 1812–1820.  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 and affects survival in esophageal cancer patients, Clin. Cancer Res. 12 (2006) 7322–7328.  S. Tokumaru, M. Suzuki, H. Yamada, M. Nagino, T. Takahashi, Let-7 regulates Dicer expression and constitutes a negative feedback loop, Carcinogenesis 29 (2008) 2073–2077.  J.J. Forman, A. Legesse-Miller, H.A. Coller, A search for conserved sequences in coding regions reveals that the let-7 microRNA targets Dicer within its coding sequence, Proc. Natl. Acad. Sci. U. S. A. 105 (2008) 14,879–14,884.  Breakdown of RNAi-Based Drugs in the Clinic, RNAi News, vol. 6, 2008.  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 cell proliferation in vitro and in vivo, Clin. Cancer Res. 13 (2007) 2207–2215.  R. Zukiel, S. Nowak, E. Wyszko, K. Rolle, I. Gawronska, M.Z. Barciszewska, J. Barciszewski, Suppression of human brain tumor with interference RNA speciﬁc for tenascin-C, Cancer Biol. Ther. 5 (2006) 1002–1007.  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 of targeted nanoparticles containing ribonucleotide reductase subunit M2 siRNA, Proc. Natl. Acad. Sci. U. S. A. 104 (2007) 5715–5721.  E. Check, A tragic setback, Nature 420 (2002) 116–118.  T. Nguyen, E.M. Menocal, J. Harborth, J.H. Fruehauf, RNAi therapeutics: an update on delivery, Curr. Opin. Mol. Ther. 10 (2008) 158–167.  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.  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.  A. Aigner, Nonviral in vivo delivery of therapeutic small interfering RNAs, Curr. Opin. Mol. Ther. 9 (2007) 345–352.  J.A. Hughes, G.A. Rao, Targeted polymers for gene delivery, Expert Opin. Drug Deliv. 2 (2005) 145–157.  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.  M. Thomas, A.M. Klibanov, Enhancing polyethylenimine's delivery of plasmid DNA into mammalian cells, Proc. Natl. Acad. Sci. U. S. A. 99 (2002) 14,640–14,645.  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.  J.D. Heidel, Linear cyclodextrin-containing polymers and their use as delivery agents, Expert Opin. Drug Deliv. 3 (2006) 641–646.  J.N. Moreira, A. Santos, V. Moura, M.C. Pedroso de Lima, S. Simoes, Non-viral lipidbased nanoparticles for targeted cancer systemic gene silencing, J. Nanosci. Nanotechnol. 8 (2008) 2187–2204.  N. Blow, Small RNAs: delivering the future, Nature 450 (2007) 1117–1120.  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 in non-human primates, Nature 441 (2006) 111–114.  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 administration of siRNA-lipoplexes for cancer therapy, Gene Ther. 13 (2006) 1360–1370.  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-lipoplex technology for RNA interference in the mouse vascular endothelium, Gene Ther. 13 (2006) 1222–1234.  M. Aleku, G. Fisch, K. Mopert, O. Keil, W. Arnold, J. Kaufmann, A. Santel, Intracellular localization of lipoplexed siRNA in vascular endothelial cells of different mouse tissues, Microvasc. Res. 76 (2008) 31–41.  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, a liposomal small interfering RNA formulation targeting protein kinase N3, inhibits cancer progression, Cancer Res. 68 (2008) 9788–9798.  J.S. Vorhies, J.J. Nemunaitis, Synthetic vs. natural/biodegradable polymers for delivery of shRNA-based cancer therapies, Methods Mol. Biol. 480 (2009) 11–29.  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 delivery suppresses HIV-1 infection in humanized mice, Cell 134 (2008 Dec 30) 577–586.  K. Gao, L. Huang, Nonviral methods for siRNA Delivery, Mol. Pharmacol. 2008 Dec 30.  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 delivery of RNAi therapeutics, Nat. Biotechnol. 26 (2008) 561–569.  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 novel approach for gene therapy in cancer, Connect. Tissue Res. 49 (2008) 265–269.  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 the central nervous system, Nature 448 (2007) 39–43.  A. Judge, I. MacLachlan, Overcoming the innate immune response to small interfering RNA, Hum. Gene Ther. 19 (2008) 111–124.  H. Lv, S. Zhang, B. Wang, S. Cui, J. Yan, Toxicity of cationic lipids and cationic polymers in gene delivery, J. Control. Release 114 (2006) 100–109.  T.G. Park, J.H. Jeong, S.W. Kim, Current status of polymeric gene delivery systems, Adv. Drug Deliv. Rev. 58 (2006) 467–486.  M. Breunig, U. Lungwitz, R. Liebl, A. Goepferich, Breaking up the correlation between efﬁcacy and toxicity for nonviral gene delivery, Proc. Natl. Acad. Sci. U. S. A.104 (2007) 14,454–14,459.  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.  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 region sequence complementarity, RNA 12 (2006) 1179–1187.  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.  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, Positionspeciﬁc chemical modiﬁcation of siRNAs reduces “off-target” transcript silencing, RNA 12 (2006) 1197–1205.  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 seed matches, but not overall identity, are associated with RNAi off-targets, Nat. Methods 3 (2006) 199–204.  E. Anderson, Q. Boese, A. Khvorova, J. Karpilow, Identifying siRNA-induced offtargets by microarray analysis, Methods Mol. Biol. 442 (2008) 45–63.  S. van Dongen, C. Abreu-Goodger, A.J. Enright, Detecting microRNA binding and siRNA off-target effects from expression data, Nat. Methods 5 (2008) 1023–1025.  S. Qiu, C.M. Adema, T. Lane, A computational study of off-target effects of RNA interference, Nucleic Acids Res. 33 (2005) 1834–1847.  Y.K. Park, S.M. Park, Y.C. Choi, D. Lee, M. Won, Y.J. Kim, AsiDesigner: exon-based siRNA design server considering alternative splicing, Nucleic Acids Res. 36 (2008) W97–103.  T. Yamada, S. Morishita, Accelerated off-target search algorithm for siRNA, Bioinformatics 21 (2005) 1316–1324.  Y. Naito, T. Yamada, T. Matsumiya, K. Ui-Tei, K. Saigo, S. Morishita, dsCheck: highly sensitive off-target search software for double-stranded RNA-mediated RNA interference, Nucleic Acids Res. 33 (2005) W589–591.  Q. Du, H. Thonberg, J. Wang, C. Wahlestedt, Z. Liang, A systematic analysis of the silencing effects of an active siRNA at all single-nucleotide mismatched target sites, Nucleic Acids Res. 33 (2005) 1671–1677.  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 genes that differ by a single nucleotide, PLoS Genet. 2 (2006) e140.  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.  J.K. Watts, G.F. Deleavey, M.J. Damha, Chemically modiﬁed siRNA: tools and applications, Drug Discov. Today 13 (2008) 842–855.  O. Snove Jr., J.J. Rossi, Chemical modiﬁcations rescue off-target effects of RNAi, ACS Chem. Biol. 1 (2006) 274–276.  M.A. Behlke, Chemical modiﬁcation of siRNAs for in vivo use, Oligonucleotides 18 (2008) 305–319.  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.  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 strand selection and targeting speciﬁcity, RNA 14 (2008) 263–274. D.D. Rao et al. / Advanced Drug Delivery Reviews 61 (2009) 746–759  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: modiﬁed siRNA with a DNA seed arm is a powerful tool for mammalian gene silencing with signiﬁcantly reduced off-target effect, Nucleic Acids Res. 36 (2008) 2136–2151.  D. Rao, N. Senzer, M.A. Cleary, J. Nemunaitis, Comparative assessment of siRNA and shRNA off target effects: what is slowing clinical development. Cancer Gene Therapy (2009).  D. Bumcrot, M. Manoharan, V. Koteliansky, D.W. Sah, RNAi therapeutics: a potential new class of pharmaceutical drugs, Nat. Chem. Biol. 2 (2006) 711–719.  A.J. Bridge, S. Pebernard, A. Ducraux, A.L. Nicoulaz, R. Iggo, Induction of an interferon response by RNAi vectors in mammalian cells, Nat. Genet. 34 (2003) 263–264.  C.A. Sledz, M. Holko, M.J. de Veer, R.H. Silverman, B.R. Williams, Activation of the interferon system by short-interfering RNAs, Nat. Cell Biol. 5 (2003) 834–839.  K. Kariko, P. Bhuyan, J. Capodici, D. Weissman, Small interfering RNAs mediate sequence-independent gene suppression and induce immune activation by signaling through toll-like receptor 3, J. Immunol. 172 (2004) 6545–6549.  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 in plasmacytoid dendritic cells through TLR7, Nat. Med. 11 (2005) 263–270.  A.D. Judge, V. Sood, J.R. Shaw, D. Fang, K. McClintock, I. MacLachlan, Sequencedependent stimulation of the mammalian innate immune response by synthetic siRNA, Nat. Biotechnol. 23 (2005) 457–462.  S.S. Diebold, T. Kaisho, H. Hemmi, S. Akira, C. Reis e Sousa, Innate antiviral responses by means of TLR7-mediated recognition of single-stranded RNA, Science 303 (2004) 1529–1531.  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 tolllike receptor 7 and 8, Science 303 (2004) 1526–1529.  M. Karin, Y. Yamamoto, Q.M. Wang, The IKK NF-kappa B system: a treasure trove for drug development, Nat. Rev., Drug Discov. 3 (2004) 17–26.  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 for discriminating between self and nonself double-stranded RNAs in mammalian cells, Nat. Biotechnol. 24 (2006) 559–565.  M. Sioud, RNA interference and innate immunity, Adv. Drug Deliv. Rev. 59 (2007) 153–163.  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 by siRNA via TLR3, Nature 452 (2008) 591–597.  M. Robbins, A. Judge, E. Ambegia, C. Choi, E. Yaworski, L. Palmer, K. McClintock, I. Maclachlan, Misinterpreting the therapeutic effects of siRNA caused by immune stimulation, Hum. Gene Ther. 19 (10) (2008 Oct) 991–999.  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 speciesspeciﬁc CpG motif recognition, Proc. Natl. Acad. Sci. U. S. A. 98 (2001) 9237–9242. 759  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 bacterial DNA, Nature 408 (2000) 740–745.  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 gene expression after intravenous administration of polyplexes, Pharm. Res. 25 (2008) 1654–1662.  M.A. Robbins, M. Li, I. Leung, H. Li, D.V. Boyer, Y. Song, M.A. Behlke, J.J. Rossi, Stable expression of shRNAs in human CD34+ progenitor cells can avoid induction of interferon responses to siRNAs in vitro, Nat. Biotechnol. 24 (2006) 566–571.  M. Sioud, Induction of inﬂammatory cytokines and interferon responses by double-stranded and single-stranded siRNAs is sequence-dependent and requires endosomal localization, J. Mol. Biol. 348 (2005) 1079–1090.  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 using vectors that encode long modiﬁed hairpin-RNA, Mol. Biosyst. 1 (2005) 382–390.  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 silencing HCV RNA and overcomes variation in the viral genotype, Gene Ther. 13 (2006) 883–892.  M. Bauer, N. Kinkl, A. Meixner, E. Kremmer, M. Riemenschneider, H. Forstl, T. Gasser, M. Uefﬁng, Prevention of interferon-stimulated gene expression using microRNA-designed hairpins, Gene Ther. 16 (2009) 142–147.  M. Sioud, Single-stranded small interfering RNA are more immunostimulatory than their double-stranded counterparts: a central role for 2′-hydroxyl uridines in immune responses, Eur. J. Immunol. 36 (2006) 1222–1230.  J.M. Layzer, A.P. McCaffrey, A.K. Tanner, Z. Huang, M.A. Kay, B.A. Sullenger, In vivo activity of nuclease-resistant siRNAs, RNA 10 (2004) 766–771.  K. Kariko, M. Buckstein, H. Ni, D. Weissman, Suppression of RNA recognition by Toll-like receptors: the impact of nucleoside modiﬁcation and the evolutionary origin of RNA, Immunity 23 (2005) 165–175.  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/short hairpin RNA pathways, Nature 441 (2006) 537–541.  D. Grimm, M.A. Kay, Therapeutic application of RNAi: is mRNA targeting ﬁnally ready for prime time? J. Clin. Invest. 117 (2007) 3633–3641.  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 silencing without interruption of the endogenous microRNA pathway, Nature 449 (2007) 745–747.  R. Yi, B.P. Doehle, Y. Qin, I.G. Macara, B.R. Cullen, Overexpression of exportin 5 enhances RNA interference mediated by short hairpin RNAs and microRNAs, RNA 11 (2005) 220–226.  J.C. Giering, D. Grimm, T.A. Storm, M.A. Kay, Expression of shRNA from a tissuespeciﬁc pol II promoter is an effective and safe RNAi therapeutic, Mol. Ther.16 (2008) 1630–1636.
© Copyright 2017