Improving siRNA Bio-Distribution and Minimizing Side Effects

Current Drug Metabolism, 2011, 12, 11-23 11

1389-2002/11 $58.00+.00 © 2011 Bentham Science Publishers Ltd.

Improving siRNA Bio-Distribution and Minimizing Side Effects

Bruna Scaggiante1, Barbara Dapas1, Rossella Farra2, Mario Grassi3, Gabriele Pozzato2, Carlo Giansante2, Nicola Fiotti2 and Gabriele Grassi1,2,*

1Department of Life Sciences, University of Trieste;

2Department of Clinical, Morphological and Technological Sciences, University

Hospital of Cattinara; 3Department of Chemical Engineering, University of Trieste, Italy

Abstract: The RNA interference (RNAi) is a biological process by which a double stranded RNA (dsRNA also called small interfering RNA - siRNA) triggers the sequence-dependent degradation of a target RNA within the cellular environment. Thus siRNAs can be used to combat the expression of deleterious gene(s) causing disease or to destroy invading pathogen RNAs. Despite their enormous therapeu-tic potential, the use of siRNA as drugs presents two major problems: the difficulties to identify optimal delivery systems and the possible induction of different unwanted side effects. In this review, after presenting an overview about the mechanisms ruling the process of RNAi, we focus the attention on the description of the strategies developed to optimise systemic siRNA delivery; in this sense, considera-tions about the attempts to improve siRNA stability in the biological environment, the development of synthetic vectors for siRNA deliv-ery, the siRNA bio-distribution and pharmacokinetics together with the selection of siRNA targeted delivery systems, are discussed. Since in the optimisation of the siRNA delivery systems the minimization of siRNA side effects should not be neglected, in the last part of the review we consider the problems related to the possible induction of siRNA mediated side effects focusing on the so called mi-croRNA like off-targeting.

Keywords: siRNA, siRNA delivery, bio-distribution, miRNA, off-targeting, RNAi interference.

1. INTRODUCTION

The RNA interference (RNAi) is commonly known as the proc-ess by which a double stranded RNA (dsRNA) triggers the se-quence-dependent degradation of a target RNA within the cellular environment. This results in either the inhibition of cellular gene expression or the inhibition of the expression of deleterious foreign genes such as viral genes. The first evidence of RNAi came in 1990 in plants [1] and the phenomenon was subsequently properly de-scribed in 1998 when Fire et al. [2] observed that the introduction of a double stranded RNA into C. elegans resulted in the expression inhibition of an homologous gene. This revolutionary biological discovery, for which Fire and Mello got the Nobel price in 2006, further increased its scientific relevance when Elbashir et al. [3] reported the possibility to induce RNAi also in human cells by us-ing short double stranded RNA molecules (small interfering RNAs – siRNAs). This observation opened the possibility to use siRNAs as therapeutic agents for human diseases. The interest of the scien-tific community for RNAi is well demonstrated by the explosion of the publication number from 1998 up till now (from few papers in 1998 till about 6000 papers in 2009).

Slightly after the demonstration of the potential therapeutic value of RNAi inducing molecules, it became evident that a major problem to be solved before the therapeutic use of siRNA was the choice of the appropriate delivery system which can guarantee ade-quate bio-distribution, i.e. the siRNA release in the proper amount to the wanted organ/tissue. A second problem to be addressed deals with the possible induction of unwanted effects triggered by the siRNAs [4], a fact which can obviously impair their therapeutic potential.

In this review we present an overview about the general mecha-nisms ruling RNAi together with ongoing applications (section 2), we stress the relevance of the selection of the appropriate delivery system able to achieve adequate bio-distribution and possibly a cell targeted delivery (section 3) and finally we discuss the problems related to the possible induction of side effects (section 4) in par-ticular focussing on the so called microRNA like off-targeting.

*Address correspondence to this author at the Department of Clinical, Mor-phological and Technological Sciences, University Hospital of Cattinara, Strada di Fiume 447, 34100 Trieste, Italy; Tel: +39-040-3996227; Fax: +39-040-3994593; E-mail: [email protected]

2. RNAi: MECHANISMS OF ACTION

The RNA interference (RNAi) pathway Fig. (1) represents a conserved biological mechanism through which eukaryotic cells control gene expression [5]. RNAi is triggered by non coding short double stranded RNA molecules termed microRNAs (miRNAs) and small interfering RNAs (siRNAs). Different hundreds of miRNA have been described in rodents and most of these have been also detected in humans. To date the exact biological functions of many miRNAs are unknown; however evidences of their implications in the regulation of pivotal cellular processes such as apoptosis and cell proliferation are rising. In addition, the observation that most of the vertebrate miRNAs are expressed during development [6] clearly points towards a relevant role in developmental control. In contrast to miRNA which are of endogenous origin, siRNAs can be also of exogenous origin. The siRNAs can indeed be generated either from invasive nucleic acids such as viruses and transposones or are the products of different endogenous sources including pseu-dogene-derived transcripts [7], in both cases their function is to preserve the host genome integrity.

The miRNA pathway starts with the transcription of a long precursor defined primary miRNA (pri-miRNA) [8] Fig. (1) which is subsequently processed (pre-miRNA) in the nucleus by a cellular enzyme called, in animal, Drosha [9]. Following processing, the pre-miRNA is exported from the nucleus to the cytoplasm by means of Exportin 5 (Exp5). Once in the cytoplasm, the pre-miRNA is processed by the DICER enzyme [7] to generate an ap-proximately 22 bp long RNA duplexes bearing 2 nt 3’ overhangs [10]. The product of the processing is then loaded onto RISC (RNA-induced silencing complexes) whose best known component is the Argonaute (Ago2) protein [7, 11, 12]. Within the two miRNA strands, the “sense” or “passenger strand” is discarded while the other, named “antisense” or “guide strand”, is the one selected to function as mature miRNA in the RISC complex Fig. (1, 2). Strand selection is dictated by the relative thermodynamic stabilities of the two duplex ends as reported in section 4. The loading of the guide strand onto RISC is a complex process whose mechanism is still under investigation [11, 13-15]. Once loaded, the guide strand binds to the target RNA most often inducing translational repression. For effective miRNA mediated translational repression, a contiguous and perfect base pairing of the first 2-8 nucleotides, from the 5’ end of the guide strand with the target, is required Fig. (2). This region has been defined as the “seed region” [16, 17] and

12 Current Drug Metabolism, 2011, Vol. 12, No. 1 Scaggiante et al.

Fig. (1). miRNA and siRNA pathways miRNA (left) and siRNA (right) pathway are reported; the guide strand is indicated in bold.

with few exceptions, it binds to sequences found in the 3’UTR of mRNA molecules, often in multiple copies [18-21]. Finally, al-though less frequently than the translational inhibition, miRNA may also induce mRNA degradation [22].

In addition to the miRNAs, also small interfering RNAs (siR-NAs), can enter the RNAi pathway Fig. (1). Although most of them are represented by synthetic molecules [3], naturally occurring molecules termed endogenous siRNA (esiRNA) have been de-scribed [7, 23, 24]. Structurally similar to miRNAs, synthetic siR-NAs (also termed short hairpin RNA (shRNA) when transcribed from an expression vector [25, 26]) induce the target RNA degrada-tion via a perfect complementarity between the guide strand and the target RNA using the same cellular enzymes used by miRNA. Only a single mismatch between the guide strand and the target RNA could profoundly impair target RNA destruction [27, 28].

Despite the theoretic possibility to generate siRNAs directed against any region of a given target RNA, it is now clear that not all possible synthetic siRNAs can efficiently induce the silencing of the target RNA. Thus, approaches have been defined to select siR-

NAs for which RISC uptake/activation and target recogni-tion/cleavage are optimal [29, 30]. Among the different parameters considered, siRNA duplex stability and sequences characteristics have been found to be of relevance. In this sense, it has been ob-served that functional siRNAs display in the overall nucleotide sequence a low-to-medium G+C content (30% -52% [29, 31]). It is possible that too low G+C content may destabilize siRNA duplexes and reduce target mRNA binding; in contrast, too high G+C content may impair RISC loading and/or cleavage-product release. Interest-ingly, although the overall duplex stability is important, the centre of the duplex (positions 9–14 on the guide strand) needs to have low stability [31-33] Fig. (2A). Additional sequences requirements have been also identified in efficient siRNAs [30]. Finally, not only single nucleotides can determine siRNA efficiency, also siRNA [29, 31, 34] secondary structure is relevant as well as the targeted RNA region [35] secondary structure. These technical problems have not impeded the identification of effective siRNAs, some of which have been tested in different clinical trials as reported in Table 1.

siRNA, Bio-Distribution and Off-Targeting Current Drug Metabolism, 2011, Vol. 12, No. 1 13

Fig. (2). Details of the siRNA/miRNA structures.A) In grey the seed region (SR) is reported: to induce expression inhibition via the off-targeting mechanism, a perfect complementarity between the SR (nucleotide position 2-7 or 2-8 of the antisense strand of the siRNA) and the 3’UTR, but not the 5’ UTR, of the tran-script is necessary. The dotted area indicates the siRNA region which needs to have low stability (positions 9–14 on the guide strand) for optimal siRNA func-tion. B) Possible role of extra SR in inducing off-targeting: the presence, on the target mRNA, of an adenosine opposite to siRNA base 1 and the presence of an adenosine or uridine opposite to siRNA base 9 enhanced mRNA repression. C) A plot of all possible 4096 hexamers (X-axis, nucleotides 2-7 of the siRNA guide strand, i.e. SR) versus the number of 3´ UTRs that contain a given hexamer (Y-axis) is presented. The approximate seed frequencies ranging from low, medium, and high-frequency seeds are indicated by circles (from Anderson [107] with permission). Table 1.

siRNA mRNA Target Disease Company

siRNA Bevasiranib against the Vascular endothelial growth factor (VEGF)

Wet age-related macular degeneration (AMD) Acuity Pharmaceuticals, Philadelphia, Pennsylvania

siRNA Sirna-027 against the Vascular endothelial growth factor receptor (VEGFR1)

Wet age-related macular degeneration (AMD) Merck’s Sirna Therapeutics (San Francisco,California)

siRNA PF-4523655 against the hypoxia-inducible gene, RTP801

Wet age-related macular degeneration (AMD) and Dia-betic Macular Degeneration (DME)

Quark Pharmaceuticals (Fremont, California)

siRNA QPI-1002 directed against the protein p53 Ischemia-reperfusion induced acute kidney injury (AKI) Quark Pharmaceuticals (Fremont, California)

siRNA ALN-RSV01 against the Respiratory syncytial virus (RSV),

Serious respiratory infectious disease in children, in elderly and in adults with compromised immune system

Alnylam Pharmaceuticals, Inc.

siRNA ALN-VSP, directed against kinesin spindle pro-tein (KSP) and VEGF,

Cancer proliferation and cancer angiogenesis Alnylam Pharmaceuticals, Inc.

NA CALAA-01 against the M2 subunit of the ribonu-cleotide reductase, an enzyme involved in DNA synthesis

Relapsed or refractory solid cancer Calando Pharmaceuticals

https://www.researchgate.net/publication/5482382_Identifying_siRNA-Induced_Off-Targets_by_Microarray_Analysis?el=1_x_8&enrichId=rgreq-d1ad5fef-31dc-4e2c-b9c2-47c5567f0f2b&enrichSource=Y292ZXJQYWdlOzQ5NzQ0MDg4O0FTOjk5Nzk0MTQzOTQwNjE2QDE0MDA4MDQxODYzOTc=


3. siRNA BIO-DISTRIBUTION

Before considering siRNA as novel therapeutic agents, the dif-ficulties to find delivery systems able to optimize bio-distribution and pharmacokinetic have to be overcome.

In vivo delivery strategies can be substantially classified into two groups based on the route of administration, i.e. localized or systemic [36]. Localized approaches, can be used for easily acces-sible tissues; in this case siRNA delivery occurs directly in the tar-get tissue, it allows the accumulation of the proper siRNA amount and, obviously, a substantial targeted release. An example of local delivery is represented by the direct siRNA release to keratinocytes which are able to efficiently up-take unmodified siRNA thus hold-ing promise for topically cutaneous gene therapy. Benefit for pa-tients affected by the autosomal dominant keratin disorder, pachyonychia congenita, has been demonstrated in clinical trials using siRNAs for one mutation in the gene for keratin 6a. For skin diseases, most of the rate limiting barriers to siRNA delivery have been overcome in mice by combining physically methods and liposomal RNA encapsulation [37].

Local delivery is very often not possible as most tissues are not easily accessible and require the development of systemic delivery approaches. Despite the obvious difference between local and sys-temic delivery systems, both of them have to overcome different physiological barriers. Any kind of in vivo delivery systems has to bypass several physiological barriers which can severely prevent siRNA to reach the target cells Fig. (3). First of all (1), for systemic applications, siRNAs have to come in contact with the blood nucle-ases which can degrade them when administered in the naked form. Then (2) siRNAs have to cross the vessel barrier and the extra-cellular matrix compartment. Once in proximity to the cell, siRNAs have to pass through the cellular membrane (3), a difficult step due to the electrostatic repulsion between the negatively charged phos-phate groups present in siRNA structure and the negatively charged surface of cellular membranes. Moreover, the hydrophilic nature of siRNAs does not allow an efficient passage through the inner hy-drophobic layer of cellular membranes. In the cellular environment (4), siRNAs are then susceptible to further degradation by cellular nucleases and have to face with the problem of cellular trafficking such as endosomal release and cytoplasmic transport [38, 39]. It is therefore clear that without the proper delivery system an irrelevant fraction of the administered siRNA can reach the desired cells. Whereas this is particularly true for systemic application, also local delivery approaches have to face with steps (3) and (4). In the next section a description about the proposed solutions to circumvent the physiological barrier is reported.

3.1. siRNA Chemical Modifications

As above mentioned once in blood siRNAs are exposed to the nucleases which most often have a 3’exonuclease activity but also an inter-nucleotide nuclease activity can take place. Chemical modifications of the siRNA structure have shown to significantly reduce nuclease digestion thus increasing siRNA half life; however the problem is that the modifications can negatively affect siRNA activity. It has been demonstrated that modifications in the sense strand are in general well tolerated, whereas in the antisense strand this depends on the position [40]. The relationship between the chemical modifications, the position within the strands and siRNA efficacy/toxicity needs to be further elucidated.

Four major types of modification of the siRNA nucleotides have been investigated: i) modifications involving the phosphate moiety, ii) modifications involving the sugar moiety, iii) deep modifications of the chemical structure and iiii) siRNA conjugation with different molecules [41].

Phosphorothioate modification (PS) involves the substitution of the non-bridge oxygen of the phosphate group with a sulphur atom Fig. (4); this modification is the most commonly used chemical

Fig. (3). Biological barriers for siRNA delivery. Following systemic ad-ministration, siRNAs come in contact with blood nucleases (1); then (2) they have to pass through the vessel barrier and the extra-cellular matrix compartment; in proximity to the cell (3) siRNAs need to cross the cellular membrane; once in the cell (4), they have to face with the problem of en-dosomal release and with nuclear membrane crossing, at least for the siR-NAs whit a nuclear target.

modification for both sense and antisense strands due to its easy synthesis. Preferentially PS are located in the siRNA extremities as their introduction in the siRNA centre negatively affect siRNA activity [42]. Notably, due to the tendency of PS to non specifically bind to protein, a certain toxicity has been described [43]. However, this phenomenon enhances the pharmacokinetic of siRNAs, delay-ing kidney excretion. In addition to PS, the insertion of a boran group - BH3 – in the place of the native free hydroxylic moiety of the phosphoric group has been developed. When present in nucleo-tides at the end of the chain, this modification confers to siRNA increased activity in comparison to the respective unmodified nu-cleotides [44], unfortunately, however, this modification presents some difficulties with regard to the chemical synthesis.

Among sugar modifications, position 2’ on the sugar moiety has been extensively modified resulting in increased siRNA stability and protection from immune system activation. Etherification of the sugar 2’–OH was made using several chemical groups such as the 2’-O-methylation of nucleotide Fig. (4) in both the sense and an-tisense strands. Whereas partial 2’-O-Me modification gives perfect functional siRNAs [45], full modification of the double strand may impair siRNA activity [46]. Another possible modification of 2'OH involves the use of a fluorine atom (2’F modification), tolerated at the site of Ago2 cleavage. Partial modification is tolerated in both


siRNA strands obtaining functional derivatives [47]. Some authors have also observed that full fluorinated siRNAs have good activity [48]. Other ribose modifications consist, for example, in the change of the oxygen in the ring with sulphur (4’S-RNA). The observed increased affinity of 4’S-RNA for the serum proteins determines an increased bio-distribution. Optimal results in terms of activity were obtained with modifications introduced in terminal siRNA position [49] but not in the inner positions [50].

Fig. (4). Scheme of the experimental oligonucleotide modifications. Chemi-cal structures of a non-modified nucleotide (A) and some typical modifica-tions in the phosphate portion (B), in the sugar portions (C) and a deep modification of the chemical structure.

Deep modifications of the chemical structure involves funda-mental substitutions in oligonucleotide backbone; this is the case for peptide nucleic acid (PNA) [51] Fig. (4), for phosphorodiami-date morpholino oligonucleotide (PMO) where ribose is replaced by morpholino [52], for locked nucleic acid (LNA) that contains a methylene bridge connecting the 2 -oxygen of the ribose with the 4 -carbon [53] and for nucleic acid containing modified units with 2 -deoxy-2 -fluoro- -D-arabinonucleotide (FANA) that are suitable for sense-strand [54]. Whereas all these modifications in-crease nuclease stability, caution has to be put in the amount of modified nucleotides introduced as excessive modification may negatively alter siRNA functions.

Finally, the conjugation of different molecules to siRNAs (oligo-conjugation) represents another strategy to improve nuclease stability; moreover, this approach has been shown to facilitate siRNA cellular uptake and intracellular delivery [55]. Oligo-conjugation consists of the chemical conjugation of various mole-cules, such as cholesterol and derivatives, peptides, polyethylene glycol to the siRNA [56-58], whereas these molecules can be added at 3’ and 5’ end of each strand, terminal antisense strand modifica-tions should be carefully evaluated due to the possibility to affect RISC activity.

Together, the above reported examples indicate the consider-able number of strategies used to improve siRNA stability. So far, in our opinion, the ideal modification which can be indicated for all experimental set up does not exist; in contrast, the optimal modifi-

cation(s) has to be selected on the basis of the biological variables such as the target cells and tissue as well as the delivery strategy, i.e. systemic or localized.

3.2. Synthetic Vector Mediated Delivery of siRNA

Whereas siRNA chemical modifications can significantly im-prove stability, they may not properly bypass the other biological barriers such as the crossing of the cellular membrane. For this reason chemical modified siRNAs are often delivered via synthetic vectors. These are a heterogeneous group of vectors, typically in the nano-micro scale range, each with its peculiarity with regard to the delivery properties. The most popular synthetic vectors are certainly liposomes, vesicles containing an internal aqueous compartment separated from the external environment by a bi-layer membrane constituted by amphiphilic lipids. While polar heads are oriented towards the external and internal environments, a-polar tails form a hydrophobic environment inside the membrane. Liposomes used for siRNA delivery are cationic lipids formed either by a single type of cationic amphiphilic molecules or, more commonly, by a combi-nation of cationic amphiphilic molecules and a neutral lipid [59, 60]. Liposome/siRNA complexes are very heterogeneous and dy-namic, varying in size and shape depending upon the molar ratio of liposomes to nucleic acid molecule (most important changes in structure taking place when the positive/negative charge ratio is around 1) [59, 61]. It has been proposed that liposomes can neutral-ize the negative charge of siRNAs to induce a cooperative collapse of the siRNA structure. In virtue of the conspicuous reduction of the exposed surface area, collapsed siRNAs could then be effi-ciently encapsulated by lipid [62]. Upon encapsulation, siRNAs not only are protected from degradation, they can easily cross the nega-tively charged cell membrane and the hydrophobic component of the cellular membrane thus reaching the cytoplasm. Endocytosis is the leading mechanism ruling liposome-siRNA complex entry into cells [63, 64]. An initial association to the cell surface is followed by complex entry and storing inside endosomes. Here, retained complexes destabilise endosome membrane according to a flip-flop mechanism involving endosome membrane anionic lipids. This destabilisation allows siRNA escaping from the endosome to reach the cytoplasm. In case endosome escape does not occur efficiently, siRNAs are substantially sequestered and cannot exert the biologi-cal function; thus, proper and efficient endosome escape has to be accurately evaluated for any specific application to ensure success-ful siRNA activity. Beside this aspect, liposomes are commonly used for their general ability to cross cellular membrane and for the low immunogenicity responses [65].

A variation of the liposome delivery system is represented by echogenic liposomes [66]. These are liposomes incorporating a gas presents in the internal aqueous environment or in the lipidic bi-layer. The essential advantage of containing a gas relies in the echogenic sensitivity of liposomes. When exposed to appropriate ultrasounds, the gas contained in the echogenic liposomes under-goes compression, expansion and vibration (cavitation phenome-non); this in turn determines liposome stretching till rupture with consequent siRNA delivery. Basically, the advantages of using echogenic liposomes rely on i) the possibility of realizing a targeted delivery (ultrasound can be focused), ii) an easy liposome visualisa-tion by means of diagnostic ultrasound and iii) on the possibility of improving siRNA cellular internalisation as ultrasound cavitation effects can provoke the formation of transient pores in the cellular membrane enhancing siRNA income [67].

In addition to the above reported synthetic vectors, many others are being developed. Among these mesoporous silica nanoparticles, polymers, dendrimers and cyclodextrins represent interesting alter-native under active investigation [41]. Among synthetic polymers, polyethylenimine (PEI) is widely used due to its strong proton sponge effect that increases the endosomal escape of siRNA, a phe-nomenon useful to prevent the toll-like receptor-mediated immu-


nological response that starts in endosome of macrophages [68]. However, synthetic polymer-based delivery systems, such as PEI, can cause toxicity because they are poorly degradable. Thus de-gradable polymers have been developed and a clinical trial for solid tumours targeted delivery of siRNA started using a cyclodextrin polymer-based nanoparticles (www.clinicaltrials.gov NTC0068-9065).

Finally it should be mentioned that a complete different ap-proach from the synthetic-vector delivery is represented by viral vectors. This approach implies the introduction of the nucleic acid materials coding for the siRNA into the backbone of a viral vector which in turn carries the siRNA into the cell [25]. Once in the cells, the siRNA is continuously expressed from the viral backbone by means of appropriate promoters [26]. The main advantage of this approach is a more prolonged presence of the siRNA within the cellular environment; among the main disadvantage is the safe con-cern about the use of viral-derived vectors in humans.

3.3. Evaluation of siRNA Bio-Distribution and Pharmacokinet-ics

The first evidences of siRNA gene silencing in vivo was dem-onstrated by systemic administration in mice of naked molecule [69]. In general, the distribution behavior of naked siRNAs follow-ing i.v. injection is comparable to other types of oligonucleotide: rapid elimination from the blood, hepatic uptake, degradation by nucleases and urinary excretion. In particular, the major accumula-tion is found in the kidney accounting for about the 30% of the quote injected and in liver where parenchymal and non-parenchymal cells take up the siRNA by non-specific mechanisms [40]. Although most studies dealing with the bio-distribution and pharmacokinetics of nucleic acid molecules have been performed for antisense oligodeoxynucleotides, siRNA delivery has also re-ceived recent attention. For example, Merkel et al. [70] studied the siRNA bio-distribution by using the single photon emission com-puter tomography (SPECT) in mice. siRNAs were delivered either as naked molecules or conjugated with polyethylenimine (PEI), a widely used polymer [71] which has the property to efficiently escape endosomes according to a “proton sponge” effect [72]. After the injection of 35 μg of siRNA i.v. to the tail vein, bio-distribution was evaluated. PEI conjugated siRNA had a rapid uptake and ac-cumulation in the liver and kidneys with a half life in the blood of 168.5 s. The naked siRNA showed an initial uptake in the liver and kidneys but no accumulation; differently from the PEI conjugate siRNA, an increased half life in blood of 692.8 s and rapid excre-tion into the bladder was observed. Together, these data indicate that PEI is potentially suitable for siRNA delivery and that it favors a more rapid blood clearance with siRNA accumulation in kidneys and liver.

PEI combined with PEG has also been used for siRNA delivery [73]. Following i.v. injection of radioactively labeled PEI(-PEG) complexed siRNAs, it has been observed uptake in the liver, spleen, lung and kidney; however marked differences in the pharmacoki-netics and bio-distribution occurred depending on the PEGylation pattern as well as on the PEI/siRNA ratio. These variables are also relevant with regard to the induction of erythrocyte aggregation and hemorrhage. Furthermore, this study indicates that whereas siRNA uptake in liver and spleen, but not in lung, is dependent on macro-phage activity, in the kidney PEI(-PEG)/siRNA uptake is mostly passive and reflects the total stability of the complexes.

In another study [74] it was compared the bio-distribution of naked siRNAs containing either phosphodiester linkages in both strands (PO/PO) or one strand with PO and the other one with phosphorothioate (PS) linkages. Both siRNA types, labeled by 125I, and administered i.v. in mice, accumulated predominantly in the liver and kidneys with lower amounts detected in the lung, spleen and heart. No dramatic differences were observed between the amounts of PO/PO and PO/PS siRNA in the liver and kidneys,

suggesting a minor role for PS linkages in determining the siRNA distribution. Notably, within 4 hours from the delivery, higher blood levels of the PO/PS siRNA were detected compared to the PO/PO siRNA, a fact probably reflecting the increased serum sta-bility and protein binding determined by PS linkages; however, from 24 hours till the ending point of the experiment (72 hours) the blood concentrations were comparable for the two types of siRNAs. Finally, no significant differences in terms of bio-distribution were observed between the i.v. and intra-peritoneal administration for the PO/PO siRNA suggesting that the route of administration is not relevant for siRNA bio-distribution, at least in the experimental set up considered.

Whereas the liver and kidneys seem to be the major target or-gans following siRNA systemic administration, a report from van de Water et al. [75] indicates the kidneys but not the liver as target organ for PO/PO siRNA. The reason for this discrepancy may de-pend on the radio-labeling method (111Indium labeling) used which may have determined the predominant kidney uptake in van de Water work compared to the other works above mentioned.

With the aim to further improve siRNA bio-distribution, siRNA conjugation with lipid molecules has been investigated. Soutschek et al. [76] tested the efficacy and bio-distribution of a partially PS containing siRNA conjugated with cholesterol (chol-siRNA). Com-pared to the naked siRNA, the chol-siRNA displayed an increased blood permanence following i.v administration most likely due to an efficient binding with serum albumin; moreover the chol-siRNA had a broader tissue distribution (liver, heart, kidney, adipose tissue and lung). Notably, the cholesterol conjugation did not alter the siRNA activity as shown by the fact that the siRNA target (apolipo-protein B mRNA) was efficiently silenced in the liver/jejunum with consequent reduction of the apolipoprotein B protein in the blood. A variation of the chol-siRNA consists of the conjugation of the siRNA with a -tocopherol (vitamin E) molecule [77]. This siRNA, containing modifications with PS linkages and sugar 2 -O-methylation on both the sense and the antisense strands, was cova-lently bound to -tocopherol ( -toco-siRNA); this choice was based on the fact that -tocopherol has its own physiological transport pathway to most of the organs. Notably, the -toco-siRNA was designed to be cleaved by DICER in order to produce a mature form lacking the -tocopherol molecule. The -toco-siRNA, di-rected against apolipoprotein B mRNA, efficiently reached the liver and silenced its target. Notably, compared to the chol-siRNA above reported, the -toco-siRNA was more effective as only 2mg/Kg were necessary to efficiently silence the target vs. the 50-100 mg/Kg for the chol-siRNA. Both for the chol-siRNA and -toco-siRNA no major systemic side effects were detected. It should be pointed out that siRNA conjugation with lipids is not limited to cholesterol and -tocopherol, it also includes other lipids such as bile acids and long-chain fatty acids which, based on their peculiar chemical structure, can preferentially direct siRNA to defined or-gans [78]. Moreover, very recently it has been developed a delivery strategy using lipid-like compounds based on epoxide chemistry [79]: this approach allows to significantly scale down the siRNA doses to about 0.01 mg⁄Kg in mice and 0.03 mg⁄Kg in non-human primates thus significantly improving the delivery efficiency.

Albeit ideally, for the safe and successful in vivo delivery of siRNA specific and strong gene silencing, biocompatibility, biode-gradability, absence of immunogenicity and escape from the rapid hepatic and renal clearance of the molecules should be achieved. Together, the data above reported indicate the significant advance-ments reached in this field and constitute a solid platform on which building more sophisticated delivery systems characterized by im-proved tissue specificity. So far, indeed, the liver and kidneys are easily reached by siRNAs but other organs are not. To try to get a more targeted delivery of siRNAs, different strategies have been developed as described in the following section.


3.4. siRNA Targeted Delivery

An interesting strategy for minimizing siRNA delivery to by-stander tissues, which may be negatively affected by the siRNA, is based on the use of agents that recognize specific molecules on the surface of distinct cell populations [80, 81]. In this sense, oligo-conjugation represents a strategy to improve the delivery specific-ity: it is in principle possible to link to the siRNA cell-specific ligand such as aptamers, antibodies, sugar molecules, vitamins, and hormones [82-86] to confer cell specificity to siRNA delivery sys-tems. Specific interaction between a ligand and its cellular mem-brane receptor can enhance siRNA cellular uptake by means of receptor-mediated endocytosis. In addition to conjugate the target-ing molecule to the siRNA, it is also possible to link the targeting molecule to the siRNA delivery system.

In the field of cancer specific delivery, the use of Arg-Gly-Asp (RGD) peptide has been investigated. RGD peptide binds to trans-membrane integrins, over-expressed in many types of cancer, thus preferentially promoting the entry of its cargo into cancer cells. The RGD sequence has been successfully combined to the surface of different siRNA delivery systems resulting in a preferential target-ing of the cancer cells leaving normal cells relatively unaffected [87]. Another and more complex approach involving the use of targeting peptides is represented by nanogels [88]. The method is based on core/shell hydro-gel nanoparticle (nanogel): the nanogel is composed by a porous hydro-gel core able to host therapeutic molecules and by a porous hydro-gel shell that displays the appro-priate ligation sites for the attachment of targeting ligands. This delivery system was loaded with a siRNA directed against the epi-dermal growth factor receptor (EGFR) which is over-expressed in most epithelial ovarian cancers; EGFR and its ligand, epidermal growth factor (EGF), regulate the progression of ovarian cancer. As targeting ligand, the YSA peptide mimicking the ligand ephrin-A1, which binds to the erythropoietin-producing hepatocellular (Eph) A2 receptor, was used. The delivery of nanogel-loaded EGFR siRNA to EphA2 positive cells resulted in the loss of EGFR expres-sion; notably, in EphA2 negative cells, EGFR levels were not af-fected.

Another popular target molecule in cancer specific drug deliv-ery is represented by the folate receptor. Folic acid, which is in-volved in the DNA biosynthesis, is required in high amount in the case of accelerated cell growth; this is the reason why many cancer cells over-express folate receptors thus having higher amount of the folate receptor on the cell surface compared to non cancer cells. Taking advantage of this biological feature it has been though to conjugate folic acid onto the surface of liposomal and polymeric siRNA carriers [89]. As a result of this strategy it was possible to observe that, for example, folic acid-conjugated polyethylenimine had enhanced gene silencing via receptor-mediated endocytosis [90].

Very recently a clinical trial aimed at the targeted delivery of siRNA to patients with solid cancers refractory to standard-of-care therapies has begun [91]. The siRNA used is directed against the mRNA of M2 subunit of ribonucleotide reductase, an established anti-cancer target. The delivery strategy is based on the use of gold nanoparticles delivered by i.v. infusion; the particles are coated by a cyclodextrin-based polymer, a hydrophilic polymer (polyethylene glycol used to promote nanoparticle stability in biological fluids) and by the human transferrin protein (hTf) targeting ligand. The hTf ligand should bind the Tf receptors (hTfR) on the surface of the cancer cells which are known to over-express this receptor. Thus in principle cancer cells but not normal cells should predominantly uptake the particles carrying the therapeutic siRNA. Notably, these nanoparticles have been shown to be well tolerated in multi-dosing studies in non-human primates. The data so far available indicate the presence, in tumour tissue biopsies, of intracellular-localized nanoparticles in amounts that correlate with dose levels of the nanoparticles administered; in addition there is evidence of the

reduction in both the siRNA target mRNA and the respective pro-tein when compared to pre-treated sample. These and other data [92] encourage further testing of this approach whose therapeutic potential will be better clarified by additional future data.

As an alternative to peptide mediated targeting, the use of ap-tamers have been explored. Aptamers are short stretches of single-stranded DNA or RNA molecules potentially able to recognize with high affinity any given molecular target, such as proteins, carbohy-drates, metal ions and small chemicals, thus modulating biological functions in an agonistic or antagonistic fashion [41]. Attractive features of aptamers are that they are small molecules easy to syn-thesise and which can be much more easily internalised into the cells than bigger peptides. Applications of aptamers in siRNA de-livery include, among others, the use of aptamer targeted to a signa-ture surface molecule of prostate cancer. The aptamer portion of the aptamer-siRNA chimera mediates the binding to the prostate spe-cific membrane antigen (PSMA), a cell-surface receptor over-expressed in prostate cancer cells while the siRNA targets the ex-pression of survival genes [93, 94]. Whereas PSMA expressing cells internalise the RNA, thus resulting in depletion of the siRNA targets, in cells non expressing PSMA, the chimera neither binds nor affect PSMA expression. This chimera can effectively and spe-cifically inhibit tumour growth and mediate tumour regression in a xenograft model of prostate cancer.

Finally, a substantial different approach to introduce the mole-cule(s) of interest in a given target cells is represented by vertical silicon nanowires [95]. The method is based on the ability of the silicon nanowires to penetrate the cell’s membrane and subse-quently release surface-bound molecules directly into the cytosol, without the need for chemical modification or viral packaging. By this method, it is possible to obtain localized and efficient delivery of bio-molecules, including siRNA, into immortalized and primary mammalian cells. One of the potential advantages of this approach is represented by the fact that not only siRNA but also other thera-peutic molecules such as DNAs, peptides, and proteins can be re-leased alone or in combination. This method is particularly suitable for the introduction of bioactive molecules into cells explanted from the donor (ex-vivo approaches) in a high-throughput fashion; less evident is, at the moment, the use for a systemic delivery.

4. siRNA: UNDESIRABLE SIDE EFFECTS

The optimisation of the delivery system is not the only require-ment to be satisfied for the establishment of an effective therapeutic strategy based on the use of siRNAs. Indeed, once the siRNA has safely reached the target cells, care should be put to consider and minimize the possible siRNA induced side effects. In this regards, a number of works have indicated that siRNAs can trigger non-specific effects which can be grouped into three categories. The first includes the effects caused by the activation of the so called “interferon response” (IFN-response), the second is represented by the “saturation effects” and the third is referred to as the “off-targeting effect”. The first two types of side effects will not be fur-ther described in this review (for details see [96]). In contrast, a description of the off-targeting effect is presented in the following section.

4.1. siRNA microRNA Like Off-Targeting: Past and Present

The off-targeting effect can be defined as the possibility of certain siRNA/shRNAs to interfere, in addition to the target RNA, with the expression of different other transcripts. This phenomenon is based on an imperfect hybridisation between the guide strand of the siRNA/shRNAs to transcripts different from the target. This behaviour, clearly resembling that of miRNAs and for this reason defined as microRNA like off-targeting, was not expected soon after siRNA discovery. This convincement was based on the obser-vation that only a single mismatch between the guide strand and the target RNA could profoundly impair target RNA destruction [27,


97]. In addition, the limited amount of control (i.e. the evaluation of the expression levels of few unrelated genes) included in siRNA experiments did not facilitate the detection of the off-targeting ef-fect. Only more recently with the introduction of microarray and proteomics analysis in siRNA experiments which allowed studying siRNA/shRNA effects on global gene expression, the off-targeting effects could be documented and studied.

The first evidence of the off-targeting effect came in 2003 by Jackson et al. [4], just two years after the demonstration [3] of the siRNA therapeutic potential. Jackson et al. found that every siRNA can induce a unique and reproducible expression pattern, suggestive of a siRNA-specific rather than a target specific effect. The effect tends to persist also at low siRNA concentration and involves genes which present a sequence similarity of approximately 11 nucleo-tides encompassing the 3’end of siRNA sense strand. Subsequent experimentation [17, 98] revealed that, to induce expression inhibi-tion via the off-targeting mechanism, a perfect complementarity between nucleotide position 2-7 or 2-8 of the antisense strand of the siRNA (the so called “seed region” – SR, Fig. (2A) and the 3’UTR, but not the 5’ UTR, of the transcript is necessary. However, Lin et al. [16] showed that SRs are necessary but not sufficient to trigger the off-targeting. Thus, the only search for SRs may not necessarily predict the risk of off-targeting for a given gene. In this regards, Nielsen et al. [21] reported that the presence, on the target mRNA, of an adenosine opposite to siRNA base 1 Fig. (2B) and the pres-ence of an adenosine or uridine opposite to siRNA base 9 enhanced mRNA repression. Moreover, it was observed that defined se-quence in the 50 bases 5’ and 3’ of the SR as well as increased AU content in the 3’ of the SR were each independently associated with an increase in mRNA down regulation.

More recently a paper by Anderson et al.l [99] pointed back to the predominant role of SR compared to the extra SR sequences, on the extent of the off-targeting. The authors, analysing the frequen-cies of all possible (4096) hexamers (nucleotides 2-7 of the siRNA guide strand, i.e. SR) in the 3ÙTR of the trascriptome, found a not uniform frequency of the different hexamers within the 3ÙTR regions. It was thus possible to identify hexamers present in a low, medium or high number in the 3ÙTRs Fig. (3C). The authors could show that siRNAs bearing SR complementary to high fre-quency hexamers induced higher off-targeting than siRNAs with SR complementary to low frequency hexamers thus strongly point-ing towards the relevant role of SR in triggering the off- targeting.

Very recently, Burchard et al. [100] addressed other relevant aspects of the siRNA off-targeting, i.e. its reproducibility between in vitro and in vivo tests and between animal models and human. In this work it was shown a good correlation between the off targeting effects observed in vitro and in vivo; in contrast, a specie-dependent off targeting effect probably depending on the cross-species differ-ences in the 3ÚTR composition was observed. A relevant conse-quence of this finding is that experiments in mice cells cannot be used to accurately predict the off-targeting effect in human cells. To add further complexity to the off-targeting effect, it should be pointed out that this phenomenon can be triggered also by I) siR-NAs directed against gene promoter region [101] and II) by both sense and antisense strand [4]. This last phenomenon may occur when the sense strand is preferentially up-taken by RISC. The dif-ferential affinity of the two siRNA strands for RISC has been ele-gantly addressed by Schwarz [102]. The authors dissected the mechanisms of this phenomenon using an in vitro system where a siRNA was directed against either the sense or antisense sequence of the firefly luciferase mRNA Fig. (5A). The siRNA duplex in-duced more efficient degradation of the sense target than of the antisense target Fig. (5B); this effect was explained by the fact that the antisense strand was up-taken by RISC more efficiently that the sense strand. Notably, testing the ability of each single strand of the siRNA against the respective target mRNA, both siRNA strands showed comparable efficacy in target degradation Fig. (5C), al-

though at a much lower degree than the double stranded siRNA. This supports the concept that the difference observed in the effi-ciency of target degradation by the duplex did not depend on each single strand per se but rather on the ability of each strand to be assembled onto RISC. The authors could also show that the siRNA strand whose 5’ end is more weakly bound to the complementary strand (i.e. has higher A-U content) is more efficiently incorporated into RISC. Thus, to minimize the risk of a sense strand-induced off-targeting, it is advisable to design an siRNA duplex with the only antisense strand able to form complex with RISC, i.e. with an an-tisense containing weak bound at the 5’, a feature which is also linked to siRNA efficacy [32].

Together the above data clearly indicate that siRNA-induced off-targeting is reminiscent of the expression regulation exerted by miRNAs where a partial complementarity between the target RNA and the miRNA is necessary to modulate gene expression. Moreo-ver, since miRNAs affect gene expression via a combination of inhibition of translation and mRNA cleavage, it cannot be excluded that the same occurs also for the siRNA mediated off-targeting. This leads to the consideration that for a comprehensive evaluation of the siRNA off-targeting effect, a global evaluation of both the transcript and the protein levels would be necessary.

4.2. Genuine siRNA Off-Targeting

Given the demonstrated possible occurrence of off-targeting in siRNA-based experiments, it is obvious that the identification and minimization of this effect is of utmost relevance. Obviously, off-targeting occurrence in tumour tissues is not relevant for therapeutic purposes, unless either it favours tumour cell survival or it affects the surrounding normal tissue; in contrast, this phenomenon has to be prevented when siRNAs are used to control non tumour patholo-gies where a gene expression defect needs to be corrected with no cell vitality impairment.

Before studying any possible genuine off-targeting effect, it is however necessary to minimize gene expression variations due to other variables such as the delivery system used and the cell cultur-ing conditions. With regard to the delivery methods, the most commonly used are those lipid and viral-vector mediated. For other systems based on the use of different compounds such as polymers, dendrimers, silica nanoparticles, cyclodextrin and polyplexes [41] there is no extensive knowledge about their effects on global gene expression and therefore they will not be considered here.

The availability of several different lipid-based transfection reagents for siRNA allows the selection of the most suited for the specific cell line. We typically perform the selection by testing different amounts of a given lipid, different ratio lipid/siRNA, dif-ferent transfection times and different transfection medium [103-105]. In parallel to the above mentioned tests, we also evaluate the transfection efficiency and siRNA cellular distribution by using FITC-labelled siRNAs. In this way we can select the transfection lipid with the highest possible transfection efficiency and the lowest possible impact on cell viability. The optimisation of the transfec-tion conditions is very important as, for example, too high lipid concentration may alter gene expression in different unpredictable ways [106, 107]. Additionally, the analysis time is relevant. For example, unspecific lipid-mediated effects on gene transcription were considerable 24-48 hours post transduction of siRNAs into mouse liver but rapidly diminished in the extent at 72 hours [100].

Other siRNA delivery systems such as electroporation and vec-tor-based systems are not free from the possible induction of unspe-cific gene expression. Although in some cases electroporation re-sulted in fewer non-specific gene expression compared to lipid-mediated transfection [108], this technique does not eliminate the risk of non-specific gene expression. Finally, viral vectors-based delivery systems [25] have been shown in some cases to perturb the processing of native RNAi [109] and thus their use may bring to confounding results.


With regard to culturing conditions, the second variable to be optimised to identify genuine off-targeting, cell density is a particu-lar critical parameter. It is known that lipid mediated transfection at low cell density induces more unspecific toxicity than in the pres-ence of higher cell density [107]. In our experience (unpublished observation) we observed that not only cell density but also the total amount of cell is relevant. A low plating density may be plagued by increased toxicity when the cells are cultured in 96 well compared to the same plating density but in larger plates (6 wells or bigger plates). In principle the ideal density would resemble the conflu-ence: however under this condition the transfection efficiency by lipid mediated systems may be considerably reduced. Thus a com-promise between transfection efficiency and the impact on cell viability should be found by testing different cell density and total cell number.

The choice of the cell type is another relevant variable. If possi-ble it would be desirable to start off-targeting investigation with cells characterized by an easy growth, transfectability and manipu-lation. In this regard it has been observed [107] that, with few ex-ceptions, the off-target effect observed in one cell type is consis-tently observed among unrelated cell types. Thus, after studying and minimizing the off-targeting in a convenient cell type, it is possible to switch to the cell type of interest. This approach can be followed only among cells of the same species as relevant species-variability in the off-targeting effect has been reported [100].

4.2.1. Strategies to Minimize siRNA Induced Off-Targeting

So far different strategies have been proposed to minimize siRNA off-targeting: basically all of them deal with the proper planning of siRNA experiments and with the proper siRNA design.

With regard to the siRNA experiment planning, it has been observed that, at least in part, siRNA off-targeting is dose depend-

ent [4]. Therefore it is in general recommendable to scale down as much as possible the concentration of siRNA, when delivered by lipids, to reduce the off-targeting. An additional strategy to scale down the off-targeting consists of the chemical modifications of the synthetic siRNA. Jackson et al. [106] have shown that the substitu-tion by 2’-O-methyl group at position 2 in the siRNA guide strand, significantly reduced off-targeting without negatively affecting target transcript silencing. Chen et al. also reported that it is possi-ble to reduce the off-targeting of the siRNA sense (passenger) strand by 5’-O-methylation of the strand itself [110].

In addition to the introduction of chemical modifications, the off-targeting effect may be controlled by properly choosing the siRNA sequence. In this regard it has been suggested (see also the above section 3) that, to minimize the risk of a sense strand-induced off-targeting, it is advisable to design a siRNA duplex with the only antisense strand able to form complex with RISC, i.e. an antisense containing weak bound at the 5’[32]. Additionally, as the number of off-targets generated by an siRNA depends on the frequency at which the SR sequence is present in the 3’UTR trascriptome, it is advisable to generate siRNA with the antisense strand containing low frequency SR [99]. Finally, design rules useful to select siR-NAs with high specificity and low risk of generating the off-targeting induction have been proposed [111] (http://informatics-eskitis.griffith.edu.au/SpecificityServer).

4.2.2. Experimental Approaches to Detect siRNA Off-Targeting

and Specificity

Minimizing the likelihood of off-targeting and any other unspe-cific effects is very important; despite this, experimental controls have to be planned to study these aspects.

Microarray is particularly indicated to detect the specific and unwanted effects, such as off-targeting. Obviously, the off-targeting

Fig. (5). Scheme of the experimental procedure by Schwarz et al. ref 102. A) A siRNA was directed against either the sense or antisense sequence of the fire-fly luciferase mRNA. B) The siRNA duplex induced a more efficient degradation of the sense target than of the antisense target. C) Both siRNA strands showed comparable efficacy in the respective target degradation, although at a much lower degree than the double stranded siRNA.


evidenced by microarray is restricted to the unwanted gene regula-tion which occurs at the mRNA level; a comprehensive evaluation of the off-targeting may come from a global proteomic analysis, an approach so far technically not easy to perform. In contrast, a more contained proteomic analysis limited to some proteins interacting with the target protein, are technically easier to perform, still being reasonably informative. For example, while studying the effects of the depletion of the transcription factor E2F1 [104], we evaluated also the effects on the protein levels of some of its transcriptional regulated proteins such as cyclin E1 and E2. A higher, although still limited, amount of protein interactions can be evaluated by a re-cently developed approach. This strategy is based on an algorithm able to integrate the siRNA effects with protein-protein interaction; by this approach it is possible to discriminate at the protein level siRNA off-targeting from genuine effects [112].

In addition to the above reported experimental procedures, the possibility to distinguish between off-targeting and specific siRNA effects is based on: I) the demonstration of the target mRNA and protein reduction in comparison to one or more unrelated mRNAs/ proteins; II) the execution of titration experiments to minimize the siRNA amount used, still keeping a good transfection efficiency; III) the comparison between the effetcs of the specific siRNA/ shRNA with those of control siRNA/shRNA either directed against an irrelevant gene or bearing 3 or more mismatches in the critical seed region. To further distinguish between off-targeting and spe-cific siRNA effects other checks may be performed [4, 104, 113-116]. Finally, it has to be excluded the possibility that the siRNA triggers the unspecific expression of genes involved in the activa-tion of the interferon response [117], thus generating a non specific phenotype. This aspect can be simply accomplished by quantitative RT-PCR technique, in addition to microarry test.

CONCLUSIONS

Since the demonstration of the possibility to induce RNAi in human cells, it became clear the therapeutic potential of siRNAs. In this sense, the explosion in the number of published papers well documents the interest of the scientific community in this field. However, some problems must be solved before the therapeutic use of siRNAs: the first deals with the identification of appropriate delivery systems which must guarantee adequate bio-distribution and the second concerns the possible induction of side effects which can impair the siRNA therapeutic potential. Despite not hav-ing completely overcome these problems, the works performed so far to select adequate delivery systems and to minimize the possible side effects, have significantly shortened the distance between siR-NAs and therapeutic applications; we thus believe that these mole-cules have the power to become in the next future a valid alterna-tive to the commonly used drugs for which the therapeutic potential is far from being optimal.

ACKNOWLEDGEMENTS

This work was in part supported by the “Fondazione Cassa di Risparmio of Trieste”, by the “Fondazione Benefica Kathleen Foreman Casali of Trieste” and by Italian Ministry of University and Research, project P.R.I.N. (2008HCAJ9T).

REFERENCES

[1] Napoli, C.; Lemieux, C.; Jorgensen, R. Introduction of a chimeric chalcone synthase gene into petunia results in reversible co-suppression of homologous genes in trans. Plant Cell, 1990, 2(4), 279-289.

[2] Fire, A.; Xu, S.; Montgomery, M.K.; Kostas, S.A.; Driver, S.E. Mello, C.C. Potent and specific genetic interference by double-stranded RNA in Caenorhabditis elegans. Nature, 1998, 391(6669), 806-811.

[3] Elbashir, S.M.; Harborth, J.; Lendeckel, W.; Yalcin, A.; Weber, K. Tuschl, T. Duplexes of 21-nucleotide RNAs mediate RNA interfer-

ence in cultured mammalian cells. Nature, 2001, 411(6836), 494-498.

[4] Jackson, A.L.; Bartz, S.R.; Schelter, J.; Kobayashi, S.V.; Burchard, J.; Mao, M.; Li, B.; Cavet, G. Linsley, P.S. Expression profiling re-veals off-target gene regulation by RNAi. Nat. Biotechnol., 2003, 21(6), 635-637.

[5] Bartel, D.P. MicroRNAs: genomics, biogenesis, mechanism, and function. Cell, 2004, 116(2), 281-297.

[6] Lagos-Quintana, M.; Rauhut, R.; Yalcin, A.; Meyer, J.; Lendeckel, W. Tuschl, T. Identification of tissue-specific microRNAs from mouse. Curr. Biol., 2002, 12(9), 735-739.

[7] Carthew, R.W.; Sontheimer, E.J. Origins and Mechanisms of miR-NAs and siRNAs. Cell, 2009, 136(4), 642-655.

[8] Lee, Y.; Jeon, K.; Lee, J.T.; Kim, S. Kim, V.N. MicroRNA matura-tion: stepwise processing and subcellular localization. EMBO J., 2002, 21(17), 4663-4670.

[9] Kim, K.; Lee, Y.S. Carthew, R.W. Conversion of pre-RISC to holo-RISC by Ago2 during assembly of RNAi complexes. RNA, 2007, 13(1), 22-29.

[10] Bernstein, E.; Caudy, A.A.; Hammond, S.M. Hannon, G.J. Role for a bidentate ribonuclease in the initiation step of RNA interference. Nature, 2001, 409(6818), 363-366.

[11] Peters, L.; Meister, G. Argonaute proteins: mediators of RNA silencing. Mol. Cell, 2007, 26(5), 611-623.

[12] Tolia, N.H.; Joshua-Tor, L. Slicer and the argonautes. Nat. Chem.

Biol., 2007, 3(1), 36-43. [13] Bushati, N.; Cohen, S.M. microRNA functions. Annu. Rev. Cell.

Dev. Biol., 2007, 23, 175-205. [14] Rana, T.M. Illuminating the silence: understanding the structure

and function of small RNAs. Nat. Rev.Mol. Cell. Biol., 2007, 8(1), 23-36.

[15] Du, T.; Zamore, P.D. MicroPrimer: the biogenesis and function of microRNA. Development, 2005, 132(21), 4645-4652.

[16] Lin, X.; Ruan, X.; Anderson, M.G.; McDowell, J.A.; Kroeger, P.E.; Fesik, S.W. Shen, Y. siRNA-mediated off-target gene silencing triggered by a 7 nt complementation. Nucleic Acids Res., 2005, 33(14), 4527-4535.

[17] Jackson, A.L.; Burchard, J.; Schelter, J.; Chau, B.N.; Cleary, M.; Lim, L. Linsley, P.S. Widespread siRNA "off-target" transcript si-lencing mediated by seed region sequence complementarity. RNA, 2006, 12(7), 1179-1187.

[18] Doench, J.G.; Sharp, P.A. Specificity of microRNA target selection in translational repression. Genes. Dev., 2004, 18(5), 504-511.

[19] Lewis, B.P.; Burge, C.B. Bartel, D.P. Conserved seed pairing, often flanked by adenosines, indicates that thousands of human genes are microRNA targets. Cell, 2005, 120(1), 15-20.

[20] Grimson, A.; Farh, K.K.; Johnston, W.K.; Garrett-Engele, P.; Lim, L.P. Bartel, D.P. MicroRNA targeting specificity in mammals: de-terminants beyond seed pairing. Mol. Cell, 2007, 27(1), 91-105.

[21] Nielsen, C.B.; Shomron, N.; Sandberg, R.; Hornstein, E.; Kitzman, J. Burge, C.B. Determinants of targeting by endogenous and ex-ogenous microRNAs and siRNAs. RNA, 2007, 13(11), 1894-1910.

[22] Newbury, S.F.; Muhlemann, O. Stoecklin, G. Turnover in the Alps: an mRNA perspective. Workshops on mechanisms and regulation of mRNA turnover. EMBO Rep., 2006, 7(2), 143-148.

[23] Watanabe, T.; Totoki, Y.; Toyoda, A.; Kaneda, M.; Kuramochi-Miyagawa, S.; Obata, Y.; Chiba, H.; Kohara, Y.; Kono, T.; Na-kano, T.; Surani, M.A.; Sakaki, Y. Sasaki, H. Endogenous siRNAs from naturally formed dsRNAs regulate transcripts in mouse oo-cytes. Nature, 2008, 453(7194), 539-543.

[24] Tam, O.H.; Aravin, A.A.; Stein, P.; Girard, A.; Murchison, E.P.; Cheloufi, S.; Hodges, E.; Anger, M.; Sachidanandam, R.; Schultz, R.M. Hannon, G.J. Pseudogene-derived small interfering RNAs regulate gene expression in mouse oocytes. Nature, 2008, 453(7194), 534-538.

[25] Kurreck, J. RNA interference: from basic research to therapeutic applications. Angew. Chem. Int. Ed. Engl., 2009, 48(8), 1378-1398.

[26] Zentilin, L.; Giacca, M. In vivo transfer and expression of genes coding for short interfering RNAs. Curr. Pharm. Biotechnol., 2004, 5(4), 341-347.

[27] Brummelkamp, T.R.; Bernards, R. Agami, R. A system for stable expression of short interfering RNAs in mammalian cells. Science, 2002, 296(5567), 550-553.


[28] Fedorov, Y.; Anderson, E.M.; Birmingham, A.; Reynolds, A.; Karpilow, J.; Robinson, K.; Leake, D.; Marshall, W.S. Khvorova, A. Off-target effects by siRNA can induce toxic phenotype. RNA, 2006, 12(7), 1188-1196.

[29] Reynolds, A.; Leake, D.; Boese, Q.; Scaringe, S.; Marshall, W.S. Khvorova, A. Rational siRNA design for RNA interference. Nat.

Biotechnol., 2004, 22(3), 326-330. [30] Pei, Y.; Tuschl, T. On the art of identifying effective and specific

siRNAs. Nat. Methods, 2006, 3(9), 670-676. [31] Chalk, A.M.; Wahlestedt, C. Sonnhammer, E.L. Improved and

automated prediction of effective siRNA. Biochem. Biophys. Res.

Commun., 2004, 319(1), 264-274. [32] Khvorova, A.; Reynolds, A. Jayasena, S.D. Functional siRNAs and

miRNAs exhibit strand bias. Cell, 2003, 115(2), 209-216. [33] Taxman, D.J.; Livingstone, L.R.; Zhang, J.; Conti, B.J.; Iocca,

H.A.; Williams, K.L.; Lich, J.D.; Ting, J.P. Reed, W. Criteria for effective design, construction, and gene knockdown by shRNA vectors. BMC. Biotechnol., 2006, 6, 7.

[34] Patzel, V.; Rutz, S.; Dietrich, I.; Koberle, C.; Scheffold, A. Kauf-mann, S.H. Design of siRNAs producing unstructured guide-RNAs results in improved RNA interference efficiency. Nat. Biotechnol., 2005, 23(11), 1440-1444.

[35] Shao, Y.; Chan, C.Y.; Maliyekkel, A.; Lawrence, C.E.; Roninson, I.B. Ding, Y. Effect of target secondary structure on RNAi effi-ciency. RNA, 2007, 13(10), 1631-1640.

[36] Juliano, R.; Bauman, J.; Kang, H. Ming, X. Biological barriers to therapy with antisense and siRNA oligonucleotides. Mol. Pharm., 2009, 6(3), 686-695.

[37] Geusens, B.; Sanders, N.; Prow, T.; Van Gele, M. Lambert, J. Cutaneous short-interfering RNA therapy. Expert. Opin. Drug. De-

liv., 2009, 6(12), 1333-1349. [38] Kaneda, Y. Gene therapy: a battle against biological barriers. Curr.

Mol. Med., 2001, 1(4), 493-499. [39] Brown, M.D.; Schatzlein, A.G. Uchegbu, I.F. Gene delivery with

synthetic (non viral) carriers. Int. J. Pharm., 2001, 229(1-2), 1-21. [40] Higuchi, Y.; Kawakami, S. Hashida, M. Strategies for in vivo de-

livery of siRNAs: recent progress. Bio. Drugs, 2010, 24(3), 195-205.

[41] Grassi, M.; Cavallaro, G.; Scirè, S.; Scaggiante, B.; Daps, B.; Farra, R.; Baiz, D.; Giansante, C.; Guarnieri, G.; Perin, D. Grassi, G. Current strategies to improve the efficacy and the delivery of nucleic acid based drugs. Curr. Signal Transduc. Therapy, 2010, 5(2), 92-120.

[42] Schwarz, D.S.; Tomari, Y. Zamore, P.D. The RNA-induced silenc-ing complex is a Mg2+-dependent endonuclease. Curr. Biol., 2004, 14(9), 787-791.

[43] Krieg, A.M.; Stein, C.A. Phosphorothioate oligodeoxynucleotides: antisense or anti-protein? Antisense Res. Dev., 1995, 5(4), 241.

[44] Hall, A.H.; Wan, J.; Shaughnessy, E.E.; Ramsay, S.B. Alexander, K.A. RNA interference using boranophosphate siRNAs: structure-activity relationships. Nucleic Acids Res., 2004, 32(20), 5991-6000.

[45] Choung, S.; Kim, Y.J.; Kim, S.; Park, H.O. Choi, Y.C. Chemical modification of siRNAs to improve serum stability without loss of efficacy. Biochem. Biophys. Res. Commun., 2006, 342(3), 919-927.

[46] Braasch, D.A.; Jensen, S.; Liu, Y.; Kaur, K.; Arar, K.; White, M.A. Corey, D.R. RNA interference in mammalian cells by chemically-modified RNA. Biochemistry, 2003, 42(26), 7967-7975.

[47] Chiu, Y.L.; Rana, T.M. siRNA function in RNAi: a chemical modi-fication analysis. RNA, 2003, 9(9), 1034-1048.

[48] Blidner, R.A.; Hammer, R.P.; Lopez, M.J.; Robinson, S.O. Monroe, W.T. Fully 2'-deoxy-2'-fluoro substituted nucleic acids in-duce RNA interference in mammalian cell culture. Chem. Biol.

Drug Des., 2007, 70(2), 113-122. [49] Hoshika, S.; Minakawa, N.; Kamiya, H.; Harashima, H. Matsuda,

A. RNA interference induced by siRNAs modified with 4'-thioribonucleosides in cultured mammalian cells. FEBS Lett., 2005, 579(14), 3115-3118.

[50] Dande, P.; Prakash, T.P.; Sioufi, N.; Gaus, H.; Jarres, R.; Berdeja, A.; Swayze, E.E.; Griffey, R.H. Bhat, B. Improving RNA interfer-ence in mammalian cells by 4'-thio-modified small interfering RNA (siRNA): effect on siRNA activity and nuclease stability when used in combination with 2'-O-alkyl modifications. J. Med. Chem., 2006, 49(5), 1624-1634.

[51] Nielsen P, Egholm M, Berg R, Buchardt O. Peptide nucleic acids (PNA) : oligonucleotide analogs with a polyamide backbone. In: Crooke S, Lebleu B, eds. Antisense Research and Applications. Boca Rato, FL: CRC Press, 1993; 363-73.

[52] Summerton, J.; Weller, D. Morpholino antisense oligomers: design, preparation, and properties. Antisense Nucleic Acid Drug Dev., 1997, 7(3), 187-195.

[53] Mook, O.R.; Baas, F.; de Wissel, M.B. Fluiter, K. Evaluation of locked nucleic acid-modified small interfering RNA in vitro and in

vivo. Mol. Cancer Ther., 2007, 6(3), 833-843. [54] Dowler, T.; Bergeron, D.; Tedeschi, A.L.; Paquet, L.; Ferrari, N.

Damha, M.J. Improvements in siRNA properties mediated by 2'-deoxy-2'-fluoro-beta-D-arabinonucleic acid (FANA). Nucleic Acids

Res., 2006, 34(6), 1669-1675. [55] Jeong, J.H.; Mok, H.; Oh, Y.K. Park, T.G. siRNA conjugate deliv-

ery systems. Bioconjug. Chem., 2009, 20(1), 5-14. [56] Muratovska, A.; Eccles, M.R. Conjugate for efficient delivery of

short interfering RNA (siRNA) into mammalian cells. FEBS Lett., 2004, 558(1-3), 63-68.

[57] Lorenz, C.; Hadwiger, P.; John, M.; Vornlocher, H.P. Unverzagt, C. Steroid and lipid conjugates of siRNAs to enhance cellular up-take and gene silencing in liver cells. Bioorg.Med. Chem. Lett., 2004, 14(19), 4975-4977.

[58] Kim, S.H.; Jeong, J.H.; Lee, S.H.; Kim, S.W. Park, T.G. PEG conjugated VEGF siRNA for anti-angiogenic gene therapy. J. Con-

trol. Release, 2006, 116(2), 123-129. [59] Miller, A.D. Cationic Liposomes for Gene Therapy. Angew. Chem.

Int. Ed., 1998, 37, 1768-1785. [60] Huang, S.L. Liposomes in ultrasonic drug and gene delivery. Adv.

Drug Deliv. Rev., 2008, 60(10), 1167-1176. [61] Grassi, G.; Farra, R.; Noro, E.; Voinovivh, D.; Lapasin, R.; Dapas,

B.; Alpar, O.; Zennaro, C.; Carraro, M.; Giansante, C.; Guarnieri, G.; Pascotto, A.; Reihmers, B. Grassi, M. Charactherization of nu-cleid acid molecule/liposome complexes and rheological effects on pluronic/alginate matrices. J. Drug Deliv. Sci. Technol., 2007, 17(5), 325-331.

[62] Gershon, H.; Ghirlando, R.; Guttman, S.B. Minsky, A. Mode of formation and structural features of DNA-cationic liposome com-plexes used for transfection. Biochemistry, 1993, 32(28), 7143-7151.

[63] Friend, D.S.; Papahadjopoulos, D. Debs, R.J. Endocytosis and intracellular processing accompanying transfection mediated by cationic liposomes. Biochim. Biophys. Acta., 1996, 1278(1), 41-50.

[64] Mislick, K.A.; Baldeschwieler, J.D. Evidence for the role of pro-teoglycans in cation-mediated gene transfer. Proc. Natl. Acad. Sci.

U S A, 1996, 93(22), 12349-12354. [65] Karmali, P.P.; Chaudhuri, A. Cationic liposomes as non-viral carri-

ers of gene medicines: resolved issues, open questions, and future promises. Med. Res. Rev., 2007, 27(5), 696-722.

[66] Huang, S.L. Liposomes in ultrasonic drug and gene delivery. Adv.

Drug Deliv. Rev., 2008, 60 (10), 1167-1176. [67] Nyborg, W.L. Ultrasound, contrast agents and biological cells; a

simplified model for their interaction during in vitro experiments. Ultrasound Med. Biol., 2006, 32(10), 1557-1568.

[68] Saito, Y.; Higuchi, Y.; Kawakami, S.; Yamashita, F. Hashida, M. Immunostimulatory characteristics induced by linear polyethyle-neimine-plasmid DNA complexes in cultured macrophages. Hum.

Gene Ther., 2009, 20(2), 137-145. [69] Song, E.; Lee, S.K.; Wang, J.; Ince, N.; Ouyang, N.; Min, J.; Chen,

J.; Shankar, P. Lieberman, J. RNA interference targeting Fas pro-tects mice from fulminant hepatitis. Nat. Med., 2003, 9(3), 347-351.

[70] Merkel, O.M.; Librizzi, D.; Pfestroff, A.; Schurrat, T.; Behe, M. Kissel, T. In vivo SPECT and real-time gamma camera imaging of biodistribution and pharmacokinetics of siRNA delivery using an optimized radiolabeling and purification procedure. Bioconjug.

Chem., 2009, 20(1), 174-182. [71] Park, T.G.; Jeong, J.H. Kim, S.W. Current status of polymeric gene

delivery systems. Adv. Drug Deliv. Rev., 2006, 58(4), 467-486. [72] York, A.W.; Kirkland, S.E. McCormick, C.L. Advances in the

synthesis of amphiphilic block copolymers via RAFT polymeriza-tion: stimuli-responsive drug and gene delivery. Adv. Drug Deliv.

Rev., 2008, 60(9), 1018-1036.


[73] Malek, A.; Merkel, O.; Fink, L.; Czubayko, F.; Kissel, T. Aigner, A. In vivo pharmacokinetics, tissue distribution and underlying mechanisms of various PEI(-PEG)/siRNA complexes. Toxicol.

Appl. Pharmacol., 2009, 236(1), 97-108. [74] Braasch, D.A.; Paroo, Z.; Constantinescu, A.; Ren, G.; Oz, O.K.;

Mason, R.P. Corey, D.R. Biodistribution of phosphodiester and phosphorothioate siRNA. Bioorg. Med. Chem.Lett., 2004, 14(5), 1139-1143.

[75] van de Water, F.M.; Boerman, O.C.; Wouterse, A.C.; Peters, J.G.; Russel, F.G. Masereeuw, R. Intravenously administered short inter-fering RNA accumulates in the kidney and selectively suppresses gene function in renal proximal tubules. Drug Metab. Dispos., 2006, 34(8), 1393-1397.

[76] Soutschek, J.; Akinc, A.; Bramlage, B.; Charisse, K.; Constien, R.; Donoghue, M.; Elbashir, S.; Geick, A.; Hadwiger, P.; Harborth, J.; John, M.; Kesavan, V.; Lavine, G.; Pandey, R.K.; Racie, T.; Ra-jeev, K.G.; Rohl, I.; Toudjarska, I.; Wang, G.; Wuschko, S.; Bum-crot, D.; Koteliansky, V.; Limmer, S.; Manoharan, M. Vornlocher, H.P. Therapeutic silencing of an endogenous gene by systemic ad-ministration of modified siRNAs. Nature, 2004, 432(7014), 173-178.

[77] Nishina, K.; Unno, T.; Uno, Y.; Kubodera, T.; Kanouchi, T.; Mi-zusawa, H. Yokota, T. Efficient in vivo delivery of siRNA to the liver by conjugation of alpha-tocopherol. Mol. Ther., 2008, 16(4), 734-740.

[78] Wolfrum, C.; Shi, S.; Jayaprakash, K.N.; Jayaraman, M.; Wang, G.; Pandey, R.K.; Rajeev, K.G.; Nakayama, T.; Charrise, K.; Ndungo, E.M.; Zimmermann, T.; Koteliansky, V.; Manoharan, M. Stoffel, M. Mechanisms and optimization of in vivo delivery of lipophilic siRNAs. Nat. Biotechnol., 2007, 25(10), 1149-1157.

[79] Love, K.T.; Mahon, K.P.; Levins, C.G.; Whitehead, K.A.; Querbes, W.; Dorkin, J.R.; Qin, J.; Cantley, W.; Qin, L.L.; Racie, T.; Frank-Kamenetsky, M.; Yip, K.N.; Alvarez, R.; Sah, D.W.; de Fougerolles, A.; Fitzgerald, K.; Koteliansky, V.; Akinc, A.; Langer, R. Anderson, D.G. Lipid-like materials for low-dose, in vivo gene silencing. Proc. Natl. Acad. Sci. U S A, 2010, 107(5), 1864-1869.

[80] Manjunath, N.; Dykxhoorn, D.M. Advances in synthetic siRNA delivery. Discov. Med., 2010, 9(48), 418-430.

[81] Shim, M.S.; Kwon, Y.J. Efficient and targeted delivery of siRNA in vivo. FEBS J., 2010, 277(23), 4814-4827.

[82] Jeong, J.H.; Mok, H.; Oh, Y.K. Park, T.G. siRNA conjugate deliv-ery systems. Bioconjug. Chem., 2009, 20(1), 5-14.

[83] Wu, G.; Yang, W.; Barth, R.F.; Kawabata, S.; Swindall, M.; Ban-dyopadhyaya, A.K.; Tjarks, W.; Khorsandi, B.; Blue, T.E.; Ferketich, A.K.; Yang, M.; Christoforidis, G.A.; Sferra, T.J.; Binns, P.J.; Riley, K.J.; Ciesielski, M.J. Fenstermaker, R.A. Molecular targeting and treatment of an epidermal growth factor receptor-positive glioma using boronated cetuximab. Clin. Cancer. Res., 2007, 13(4), 1260-1268.

[84] Ikeda, Y.; Taira, K. Ligand-targeted delivery of therapeutic siRNA. Pharm. Res., 2006, 23(8), 1631-1640.

[85] Cesarone, G.; Edupuganti, O.P.; Chen, C.P. Wickstrom, E. Insulin receptor substrate 1 knockdown in human MCF7 ER+ breast can-cer cells by nuclease-resistant IRS1 siRNA conjugated to a disul-fide-bridged D-peptide analogue of insulin-like growth factor 1. Bioconjug. Chem., 2007, 18(6), 1831-1840.

[86] Chu, T.C.; Twu, K.Y.; Ellington, A.D. Levy, M. Aptamer mediated siRNA delivery. Nucleic Acids Res., 2006, 34(10), e73.

[87] Schiffelers, R.M.; Ansari, A.; Xu, J.; Zhou, Q.; Tang, Q.; Storm, G.; Molema, G.; Lu, P.Y.; Scaria, P.V. Woodle, M.C. Cancer siRNA therapy by tumor selective delivery with ligand-targeted sterically stabilized nanoparticle. Nucleic Acids Res., 2004, 32(19), e149.

[88] Dickerson, E.B.; Blackburn, W.H.; Smith, M.H.; Kapa, L.B.; Lyon, L.A. McDonald, J.F. Chemosensitization of cancer cells by siRNA using targeted nanogel delivery. BMC Cancer, 2010, 10, 10.

[89] Gosselin, M.A.; Lee, R.J. Folate receptor-targeted liposomes as vectors for therapeutic agents. Biotechnol. Annu. Rev., 2002, 8, 103-131.

[90] Kim, S.H.; Mok, H.; Jeong, J.H.; Kim, S.W. Park, T.G. Compara-tive evaluation of target-specific GFP gene silencing efficiencies for antisense ODN, synthetic siRNA, and siRNA plasmid com-plexed with PEI-PEG-FOL conjugate. Bioconjug. Chem., 2006, 17(1), 241-244.

[91] Davis, M.E.; Zuckerman, J.E.; Choi, C.H.; Seligson, D.; Tolcher, A.; Alabi, C.A.; Yen, Y.; Heidel, J.D. Ribas, A. Evidence of RNAi in humans from systemically administered siRNA via targeted nanoparticles. Nature, 2010, 464(7291), 1067-1070.

[92] Pirollo, K.F.; Chang, E.H. Targeted delivery of small interfering RNA: approaching effective cancer therapies. Cancer Res., 2008, 68(5), 1247-1250.

[93] McNamara, J.O.; Andrechek, E.R.; Wang, Y.; Viles, K.D.; Rempel, R.E.; Gilboa, E.; Sullenger, B.A. Giangrande, P.H. Cell type-specific delivery of siRNAs with aptamer-siRNA chimeras. Nat.

Biotechnol., 2006, 24(8), 1005-1015. [94] Chu, T.C.; Twu, K.Y.; Ellington, A.D. Levy, M. Aptamer mediated

siRNA delivery. Nucleic Acids Res., 2006, 34(10), e73. [95] Shalek, A.K.; Robinson, J.T.; Karp, E.S.; Lee, J.S.; Ahn, D.R.;

Yoon, M.H.; Sutton, A.; Jorgolli, M.; Gertner, R.S.; Gujral, T.S.; MacBeath, G.; Yang, E.G. Park, H. Vertical silicon nanowires as a universal platform for delivering biomolecules into living cells. Proc. Natl. Acad. Sci. U S A, 2010, 107(5), 1870-1875.

[96] Jackson, A.L.; Linsley, P.S. Recognizing and avoiding siRNA off-target effects for target identification and therapeutic application. Nat. Rev. Drug Discov., 2010, 9(1), 57-67.

[97] Schubert, S.; Grunert, H.P.; Zeichhardt, H.; Werk, D.; Erdmann, V.A. Kurreck, J. Maintaining inhibition: siRNA double expression vectors against coxsackieviral RNAs. J. Mol. Biol., 2005, 346(2), 457-465.

[98] Birmingham, A.; Anderson, E.M.; Reynolds, A.; Ilsley-Tyree, D.; Leake, D.; Fedorov, Y.; Baskerville, S.; Maksimova, E.; Robinson, K.; Karpilow, J.; Marshall, W.S. Khvorova, A. 3' UTR seed matches, but not overall identity, are associated with RNAi off-targets. Nat. Methods, 2006, 3(3), 199-204.

[99] Anderson, E.M.; Birmingham, A.; Baskerville, S.; Reynolds, A.; Maksimova, E.; Leake, D.; Fedorov, Y.; Karpilow, J. Khvorova, A. Experimental validation of the importance of seed complement fre-quency to siRNA specificity. RNA, 2008, 14(5), 853-861.

[100] Burchard, J.; Jackson, A.L.; Malkov, V.; Needham, R.H.; Tan, Y.; Bartz, S.R.; Dai, H.; Sachs, A.B. Linsley, P.S. MicroRNA-like off-target transcript regulation by siRNAs is species specific. RNA, 2009, 15(2), 308-315.

[101] Weinberg, M.S.; Barichievy, S.; Schaffer, L.; Han, J. Morris, K.V. An RNA targeted to the HIV-1 LTR promoter modulates indis-criminate off-target gene activation. Nucleic Acids Res., 2007, 35(21), 7303-7312.

[102] Schwarz, D.S.; Hutvagner, G.; Du, T.; Xu, Z.; Aronin, N. Zamore, P.D. Asymmetry in the assembly of the RNAi enzyme complex. Cell, 2003, 115(2), 199-208.

[103] Racchi, G.; Klingel, K.; Kandolf, R. Grassi, G. Targeting of prote-ase 2A genome by single and multiple siRNAs as a strategy to im-pair CVB3 life cycle in permissive HeLa cells. Methods Find Exp.

Clin. Pharmacol., 2009, 31(2), 63-70. [104] Dapas, B.; Farra, R.; Grassi, M.; Giansante, C.; Fiotti, N.; Uxa, L.;

Rainaldi, G.; Mercatanti, A.; Colombatti, A.; Spessotto, P.; La-covich, V.; Guarnieri, G. Grassi, G. Role of E2F1-cyclinE1-cyclinE2 circuit in human coronary smooth muscle cell prolifera-tion and therapeutic potential of its down regulation by siRNAs. Mol. Med., 2009, 15(9-10), 297-306.

[105] Farra, R.; Dapas, B.; Pozzato, G.; Giansante, C.; Heidenreich, O.; Uxa, L.; Zennaro, C.; Guarnieri, G. Grassi, G. Serum response fac-tor depletion affects the proliferation of the hepatocellular carci-noma cells HepG2 and JHH6. Biochimie., 2010, 92(5), 455-463.

[106] Jackson, A.L.; Burchard, J.; Leake, D.; Reynolds, A.; Schelter, J.; Guo, J.; Johnson, J.M.; Lim, L.; Karpilow, J.; Nichols, K.; Mar-shall, W.; Khvorova, A. Linsley, P.S. Position-specific chemical modification of siRNAs reduces "off-target" transcript silencing. RNA, 2006, 12(7), 1197-1205.

[107] Anderson, E.; Boese, Q.; Khvorova, A. Karpilow, J. Identifying siRNA-induced off-targets by microarray analysis. Methods Mol.

Biol., 2008, 442, 45-63. [108] Fedorov, Y.; King, A.; Anderson, E.; Karpilow, J.; Ilsley, D.; Mar-

shall, W. Khvorova, A. Different delivery methods-different ex-pression profiles. Nat. Methods., 2005, 2(4), 241.

[109] Grimm, D.; Streetz, K.L.; Jopling, C.L.; Storm, T.A.; Pandey, K.; Davis, C.R.; Marion, P.; Salazar, F. Kay, M.A. Fatality in mice due to oversaturation of cellular microRNA/short hairpin RNA path-ways. Nature, 2006, 441(7092), 537-541.


[110] Chen, P.Y.; Weinmann, L.; Gaidatzis, D.; Pei, Y.; Zavolan, M.; Tuschl, T. Meister, G. Strand-specific 5'-O-methylation of siRNA duplexes controls guide strand selection and targeting specificity. RNA, 2008, 14(2), 263-274.

[111] Chalk, A.M.; Sonnhammer, E.L. siRNA specificity searching in-corporating mismatch tolerance data. Bioinformatics, 2008, 24(10), 1316-1317.

[112] Tu, Z.; Argmann, C.; Wong, K.K.; Mitnaul, L.J.; Edwards, S.; Sach, I.C.; Zhu, J. Schadt, E.E. Integrating siRNA and protein-protein interaction data to identify an expanded insulin signaling network. Genome Res., 2009, 19(6), 1057-1067.

[113] Cullen, B.R. Enhancing and confirming the specificity of RNAi experiments. Nat. Methods, 2006, 3(9), 677-681.

[114] Kittler, R.; Pelletier, L.; Ma, C.; Poser, I.; Fischer, S.; Hyman, A.A. Buchholz, F. RNA interference rescue by bacterial artificial chro-mosome transgenesis in mammalian tissue culture cells. Proc. Natl.

Acad. Sci. U S A, 2005, 102(7), 2396-2401.

[115] Echeverri, C.J.; Beachy, P.A.; Baum, B.; Boutros, M.; Buchholz, F.; Chanda, S.K.; Downward, J.; Ellenberg, J.; Fraser, A.G.; Ha-cohen, N.; Hahn, W.C.; Jackson, A.L.; Kiger, A.; Linsley, P.S.; Lum, L.; Ma, Y.; Mathey-Prevot, B.; Root, D.E.; Sabatini, D.M.; Taipale, J.; Perrimon, N. Bernards, R. Minimizing the risk of re-porting false positives in large-scale RNAi screens. Nat. Methods, 2006, 3(10), 777-779.

[116] Kittler, R.; Surendranath, V.; Heninger, A.K.; Slabicki, M.; Theis, M.; Putz, G.; Franke, K.; Caldarelli, A.; Grabner, H.; Kozak, K.; Wagner, J.; Rees, E.; Korn, B.; Frenzel, C.; Sachse, C.; Sonnich-sen, B.; Guo, J.; Schelter, J.; Burchard, J.; Linsley, P.S.; Jackson, A.L.; Habermann, B. Buchholz, F. Genome-wide resources of en-doribonuclease-prepared short interfering RNAs for specific loss-of-function studies. Nat. Methods, 2007, 4(4), 337-344.

[117] de Veer, M.J.; Sledz, C.A. Williams, B.R. Detection of foreign RNA: implications for RNAi. Immunol. Cell Biol., 2005, 83(3), 224-228.

Received: October 27, 2010 Revised: January 04, 2011 Accepted: January 04, 2011

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Improving siRNA Bio-Distribution and Minimizing Side Effects