siRNA nanoparticle formulation and gene silencing

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Biomaterials 28 (2007) 1280–1288 www.elsevier.com/locate/biomaterials

The influence of polymeric properties on chitosan/siRNA nanoparticle formulation and gene silencing Xiudong Liua,b, Kenneth A. Howarda,b,, Mingdong Donga, Morten Ø. Andersena,b, Ulrik L. Rahbeka,b, Mads G. Johnsenc, Ole C. Hansenc, Flemming Besenbachera, Jørgen Kjemsa,b a

Interdisciplinary Nanoscience Center (iNANO), University of Aarhus, DK-8000, Aarhus C, Denmark b Department of Molecular Biology, University of Aarhus, DK-8000, Aarhus C, Denmark c Bioneer A/S, Denmark Received 28 July 2006; accepted 4 November 2006

Abstract We have previously introduced the use of the biomaterial chitosan to form chitosan/siRNA nanoparticles for gene silencing protocols. This present study shows that the physicochemical properties (size, zeta potential, morphology and complex stability) and in vitro gene silencing of chitosan/siRNA nanoparticles are strongly dependent on chitosan molecular weight (Mw) and degree of deacetylation (DD). High Mw and DD chitosan resulted in the formation of discrete stable nanoparticles 200 nm in size. Chitosan/siRNA formulations (N:P 50) prepared with low Mw (10 kDa) showed almost no knockdown of endogenous enhanced green fluorescent protein (EGFP) in H1299 human lung carcinoma cells, whereas those prepared from higher Mw (64.8–170 kDa) and DD (80%) showed greater gene silencing ranging between 45% and 65%. The highest gene silencing efficiency (80%) was achieved using chitosan/siRNA nanoparticles at N:P 150 using higher Mw (114 and 170 kDa) and DD (84%) that correlated with formation of stable nanoparticles of 200 nm. In conclusion, this work confirms the application of chitosan as a non-viral carrier for siRNA and the importance of polymeric properties for the optimisation of gene silencing using chitosan/siRNA nanoparticles. r 2006 Elsevier Ltd. All rights reserved. Keywords: RNA interference; Chitosan; siRNA; Nanoparticle; Delivery; Gene silencing

1. Introduction RNA interference (RNAi) has been discovered as a conserved mechanism in higher eukaryotic cells to eliminate harmful or unwanted genes, and thereby contribute to the defence from viral infection and generation of microRNA that regulate gene expression in many cellular pathways [1]. In the RNAi process, double-stranded small interfering RNA (siRNA) of 21–23 nucleotides, endogenously produced or exogenously introduced, associates with a nucleic acid-protein complex called RNA-induced silencing complex (RISC). One of the strands is selected and used to target a specific sequence in a messenger RNA Corresponding author. Department of Molecular Biology, iNANO, University of Aarhus, Building 1130. Fax: +45 8942 3690. E-mail address: [email protected] (K.A. Howard).

0142-9612/$ - see front matter r 2006 Elsevier Ltd. All rights reserved. doi:10.1016/j.biomaterials.2006.11.004

(mRNA) leading to its degradation. Hence, the synthesis of the protein encoded by the mRNA is prevented [2]. Since the first report of the use of RNAi as a tool for gene silencing in Caenorhabditis elegans [3], RNAi-mediated gene silencing (or knockdown) has also been proven effective in mammalian cells [4]. This has led to the use of RNAi technology for the study of functional genomics and the therapy of human diseases including viral infection and cancer [5,6]. The challenge, however, is to overcome extracellular and intracellular barriers to achieve efficient target cell delivery of siRNA. Previous studies have shown the difficulty for nucleic acids such as siRNA and DNA to circulate in the bloodstream, pass across cellular membranes, escape from endosomal-lysosomal compartments and reach the nucleus (in the case of RNA-expressing plasmids) [7]. Recently, viral and non-viral carrier systems have been developed to

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increase the delivery of siRNA. The use of viral vectors, such as retrovirus and adenovirus, to deliver siRNAs has shown effective gene silencing in vitro and in vivo [8,9]; however, immunogenicity and safety concerns render viralmediated delivery of siRNA less attractive [5,10]. Non-viral delivery systems using cationic liposome and polycationbased carriers such as polyethylenimine (PEI), have been used for in vivo siRNA delivery and gene silencing after intravenous or intranasal administration. These systems, however, exhibit in vivo toxicity [11–14]. Much effort, therefore, has been dedicated to the development of efficient carrier materials that are non-toxic, biocompatible and biodegradable. Chitosan, a naturally occurring cationic polysaccharide, has been widely used in drug delivery systems, especially for DNA-mediated gene therapy [15–17]. Chitosan has been shown to be biocompatible, non-inflammatory, nontoxic and biodegradable [18–20]. The protonated amine groups allow transport across cellular membranes [21,22] and subsequent endocytosis into cells [23]. Moreover, the positively charged amines (under slightly acidic conditions) allow electrostatic interaction with phosphate bearing nucleic acids to form polyelectrolyte complexes. Numerous studies on DNA delivery with chitosan as a carrier biomaterial have shown effective expression of reporter genes in vitro and in vivo [24–28], promoting chitosan as an attractive candidate for siRNA delivery. We have previously introduced a chitosan/siRNA nanoparticle system showing gene silencing in vitro and in vivo [29]. Here, we investigate the effects of chitosan properties such as molecular weight (Mw) and degree of deacetylation (DD) and the N:P ratio (theoretical charge ratio between amino groups of chitosan and phosphate groups of siRNA), on physicochemical properties (size, zeta potential, morphology and complex stability) of chitosan/siRNA nanoparticles and concomitant gene silencing efficiency. 2. Materials and methods 2.1. Materials The chitosan samples used in this study were all prepared from the Chitopharms product provided by Cognis Deutschland GmbH & Co. (Du¨sseldorf, Germany). This product is obtained by heterogeneous de-Nacetylation of shrimp chitin resulting in a disperse product with Mw spanning from 50 to 1000 kDa as determined by Gel Permeation Chromatography (GPC) and DD 80–95% determined by nuclear magnetic resonance (NMR) spectroscopy. The Chitopharms product was treated with nitrous acid [30] for depolymerisation into samples of lower Mw as presented in Table 1. The Mw of all sample preparations was verified by GPC using dextran standards (Polymer Standard Service, USA) as molecular weight references [31]. A high Mw sample was reacetylated by treatment with acetic acid anhydride [32] in order to obtain a DD close to 60%. The DD of all samples was verified by first derivative ultraviolet spectrophotometry [33,34]. siRNA-EGFP duplex (21 bp), sense sequence: 50 -GACGUAAACGG CCACAAGUUC-30 , antisense sequence: 30 -CGCUGCAUUUGCCGGU GUUCA-50 ) and four-base mismatch siRNA-EGFP (21 bp), sense sequence: 50 -GACUUAGACUGACACAAGUUC-30 , antisense sequence:

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Table 1 Chitosan samples with different Mw and DD Sample

C9–95a

C12–77

C65–78

C114–84

C170–84

C173–54

Mw, kDa DD, %

8.9 95

11.9 77

64.8 78

114.2 84

170 84

173 54

a Numbers indicate Mw (molecular weight, kDa) and DD (degree of deacetylation, %).

30 -CGCUGAAUCUGACUGUGUUCA-50 ) were obtained from Dharmacon. SYBRs Gold Nucleic Acid Gel Stain was purchased from Molecular Probes Inc. (Eugene, USA). Poly (L-aspartic) acid (PAA) and the other chemical reagents were supplied by Sigma-Aldrich (Poole, Dorset, UK). Human lung cancer cell line (H1299) expressing enhanced green fluorescent protein (EGFP) (half-life 2 h), kindly provided by Dr. Anne Chauchereau (CNRS, Villejuif, France), was used to test the gene silencing efficiency and cytotoxicity of chitosan/siRNA nanoparticles. RPMI 1640 culture medium, +GlutaMAXTMI, Penicillin/Streptomycin (P/S), G418 selection factor, Trypsin-EDTA (1  ), fetal bovine serum (FBS) and 10  TBE buffer (pH 8.470.10) were purchased from Invitrogen Corporation (Carlsbad, USA). CellTiter 96s AQueous One Solution Cell Proliferation Assay (MTS) was purchased from Promega Corporation (Madison, USA). Commercial transfection reagent, TransIT-TKO was obtained from Mirus Corporation (Madison, WI, USA).

2.2. Formulation of chitosan/siRNA nanoparticles Chitosan was dissolved in 0.2 M sodium acetate buffer and then adjusted to pH 5.5 at a final concentration of 10 mg/mL (stock solution). The concentration of siRNA-EGFP was 20 mM in RNAse-free water following the instruction of Dharmacon. 20 mL siRNA-EGFP was used, if not specifically indicated, and added to chitosan solution under stirring condition to allow nanoparticle formation. Various concentrations of chitosans (working range 10–500 mg/ml) were used to complex with siRNA at different N:P ratios. The nanoparticles were characterised and used without further treatment.

2.3. Size and surface charge determination of chitosan/siRNA nanoparticles The size of the nanoparticles was determined by photon correlation spectroscopy (PCS) and zeta potential by laser doppler velocimetry (LDV) at 25 1C using a Zetasizer Nano ZS (Malvern Instruments, Malvern, UK) The size and zeta potential of nanoparticles formulated at different parameters are presented as the mean values of three measurements7SD (standard deviation).

2.4. Morphology observation of chitosan/siRNA nanoparticles by AFM AFM imaging was performed with a commercial Digital Instruments Nanoscope IIIa MultiMode SPM (Veeco Instruments, Santa Barbara, CA) under ambient conditions. Samples were prepared by depositing a 1 ml aliquot of chitosan/siRNA nanoparticles solution on freshly cleaved mica surfaces and allowed to air dry. Standard V-shaped silicon tips (Ultrasharp cantilevers, NSC11, MikroMasch Germany) were used with a resonance frequency of 45 kHz, a spring constant of 1.5 N/m and a tip radius ofo10.0 nm. AFM images were obtained in tapping mode at 12 Hz scan rates. The AFM images were flattened using the DI software algorithm (excluding the particles from the flattened area) and then analyzed automatically by using commercial Scanning Probe Image Processor (SPIPTM) software (Image Metrology ApS) [35].

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2.5. Evaluation of the stability of chitosan/siRNA nanoparticles using native polyacrylamide gel electrophoresis (PAGE) Nanoparticle stability and siRNA integrity was investigated using a displacement assay in which the polyanion (poly (L-aspartic acid) PAA) competes and displaces the anionic siRNA from interaction with the polycationic chitosan. Samples were incubated with or without PAA (5 mg/ml) at a 1:4 volumetric ratio (PAA: complex) at 37 1C for 30 min. Samples were then analysed by electrophoresis using a 10% polyacrylamide gel (50 mM Tris–Borate, pH 7.9, 1 mM EDTA) at 150–230 V for 2 h, stained with SYBR Gold nucleic acid stain and visualised using a UV illuminator.

2.6. Gene silencing in EGFP expressed human cell line H1299 green cells were plated on 24-well plates (105 cells/well) in RPMI media (containing 10% FBS, 1% penicillin/streptomycin, and G418 selection factor) 24 h prior to transfection. The media was removed and replaced with 250 ml serum-free media and the chitosan/siRNA nanoparticles or TransIT-TKO/siRNA formulations added at 50 nM siRNA per well. After 4 h, the media was replaced with 0.5 ml fresh media containing 10% FBS. The cells were left for 44 h and then removed using a standard trypsin protocol and resuspended in PBS containing 1% paraformaldehyde. The EGFP cell fluorescence was measured using a Becton Dickenson FACSCalibur flow cytometer. A histogram plot with log green fluorescence intensity on the x-axis and cell number on the y-axis is used to define median fluorescence intensity of the main cell population defined by scatter properties (FSC, SSC, not shown). The Geomean was taken for the measure of fluorescence intensity.

2.7. Evaluation of chitosan/siRNA nanoparticle cytotoxicity Cellular cytotoxicity of chitosan/siRNA nanoparticles in H1299 green cells was determined using a tetrazolium-based viability assay. H1299 green cells in RPMI 1640+GlutaMAXTMI culture media supplemented with 10% FBS, 1% penicillin/streptomycin and G418 selection factor, were seeded at a density of 10,000 cells/well in a 96-well plate in a total well volume of 100 ml, 24 h prior to assay. The media was removed and chitosan/siRNA nanoparticles and TranslT-TKO added in triplicate and incubated for 4 h in serum-free medium (100 ml/well) after which the medium was replaced with medium containing 10% serum (100 ml/well). After 44 h, 20 ml of CellTitre 96 Aqueous proliferation assay solution was added to the plate and left for 3 h before absorbance measured at 490 nm using a 96-well plate reader (mQuant, Bio-Tek Instruments, Inc. USA).

3. Results and discussion 3.1. Effect of chitosan Mw and DD on size, zeta potential and morphology of chitosan/siRNA nanoparticles All chitosan species in this report are denoted as C Mw-DD where Mw and DD denote the molecular weight and the deacetylation degree, respectively. The Mw correlates with the physical size of the chitosan molecules, the high Mw forms are longer and more flexible molecules whereas the lower Mw forms are shorter and have stiffer molecular chains. The DD value signifies the percentage of deacetylated primary amine groups along the molecular chain, which subsequently determines the positive charge density when chitosan is dissolved in acidic conditions. Higher DD results in increased positive charge enabling a

greater siRNA binding capacity. Mw and DD are, therefore, important parameters for the interaction of chitosan with polyanionic species. Chitosan/siRNA nanoparticles were formulated at N:P 50 using different chitosan samples (Mw range 8.9–173 kDa and DD from 54% to 95% refer to Table 1) and the size and zeta potential of nanoparticles were studied (Fig. 1). Nanoparticles prepared using chitosan C9–95 (low Mw and high DD) measured 3500 nm and decreased to 500 nm with chitosan C12–77 (low Mw and medium DD). All other chitosan samples (C65–78, C114–84, C170–84, and C173–54) resulted in particles with a size of approximately 200 nm (Fig. 1A). The zeta potential for all formulations was in the range of 10–20 mV, suggesting a net positive surface charge due to excess chitosan (Fig. 1B). The charge increased slightly with Mw, with the exception of C173–54 which was reduced due to the relative low DD value. The formation of particles and the relative size was also determined by AFM (Fig. 2A–L). The large complexes formed by chitosan C9–95 were typified by rod- or circleshape self-assembly and aggregation (Fig. 2A and 2D). This might be due to shorter and stiffer chitosan chains that restrict inter-winding and further incorporation of siRNA molecules required for compact particle formation. Chitosan C12–77 formed nanoparticles and open structures but with clear signs of aggregation (500 nm) (Fig. 2B and E). Chitosan samples with both higher Mw and DD (C65–78, C114–84 and C170–84) have sufficient chain length and charge density to complex and condense siRNA into discrete nanoparticles (Fig. 2C and F, G and J, H and K). In this situation, the chitosan molecules (Mw 64.8–170 kDa), 5–10 times the length of the siRNA (Mw of 13.36 kDa), can recruit siRNA through electrostatic force along the same or adjacent chitosan molecules resulting in inter-chain connection. This may lead to further self-assembly and the formation of compact structures. In contrast to PCS, a subpopulation of nanoparticles in the 100 nm size range were evident using AFM for chitosan samples with high Mw and DD (Fig. 2J–L). This size discrepancy probably reflects the different methods used. PCS was performed on nanoparticles in a fully hydrated state in solution, whereas, AFM studies on samples dried to a mica surface. In addition, the PCS measurement presents an average size range whereas, AFM allows subpopulations to be visualised. For this reason, the use of different but complementary methods allows an overall evaluation to be made of both size and morphology. Chitosan sample C173–54 with lower DD but higher Mw can form nanoparticles (Fig. 2I and L) that suggests molecules with relatively low charge density can form complexes with siRNA if the chain length is sufficiently long. Moreover, the hydrophobic interactions or hydrogen bonds between the sugar residues of chitosan and the specific structure of the organic bases of the nucleotide may further contribute to formation of nanoparticles.

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A 5000 4500 4000 3500

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Fig. 1. The effect of chitosan Mw and DD on (A) size determined by photon correlation spectroscopy and (B) zeta potential determined by laser doppler velocimetry of chitosan/siRNA formulations at N:P 50. (Error bars represent7SD.)

3.2. Effect of chitosan Mw and DD on the complex stability of chitosan/siRNA nanoparticles The influence of chitosan Mw and DD on particle stability was examined by observing the electrophoretic

migration behaviour in the absence or presence of PAA (Fig. 3). In general, high complex stability correlated with the discrete nano-size particle morphology shown with higher Mw and DD. When chitosan samples of C9–95 and C12–77 were used to form complex at N:P 50, the

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Fig. 2. AFM images of chitosan/siRNA formulations formed with different chitosan samples at N:P 50 on mica surface. (A, D) C9–95/GFP; (B, E) C12–77/GFP; (C, F) C65–78/GFP; (G, J) C114–84/GFP; (H,K) C170–84/GFP; (I, L) C173–54/GFP. Scale bar, 500 nm (A–C and G–I); scale bar, 100 nm (D–F and J–L).

migration behaviour was almost the same as that of siRNA control (Fig. 3, Lanes 1–3) displaying no retardation effect commonly exhibited by polyelectrolyte complexes. This suggests that chitosan with lower Mw of 10 kDa (even with DD as high as 95%) cannot complex and compact siRNA into stable particles, which is in contrast to DNA plasmids where it has been reported that chitosan with 24-mer (approximately 4.7 kDa) can form stable chitosan/ DNA nanoparticles [26]. A possible explanation could be that longer DNA strands may be able to compensate for the shorter chitosan chains in the assembly process. Minimal retardation effect on the migration of siRNA was observed when using C173–54 (Fig. 3, Lane 7), which suggests the complexes formed with low deacetylated chitosan have less charge interaction and are consequently unstable. Chitosan with greater charge density (C65–78, C114–84 and C170–84) showed retarded siRNA migration

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siRNA Fig. 3. Evaluation of the effect of chitosan Mw and DD on stability of chitosan/siRNA nanoparticles at N:P 50 using native polyacrylamide gel electrophoresis (PAGE). Lane 1: siRNA-GFP; Lane 2: C9–95/GFP; Lane 3: C12–77/GFP; Lane 4: C65–78/GFP; Lane 5: C114-84/GFP; Lane 6: C170–84/GFP; Lane 7: C173–54/GFP; Lanes 8: C9–95/GFP+poly (L-aspartic acid) (PAA); Lanes 9: C12–77/GFP+PAA; Lanes 10: C65–78/GFP+PAA; Lanes 11: C114–84/GFP+PAA; Lanes 12: C170–84/GFP+PAA; Lanes 13: C173–54/GFP+PAA.

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(Fig. 3, Lanes 4–6) verifying the necessity of high charge for complex stability. In contrast, chitosan/DNA nanoparticles formed using chitosans of comparable Mw and DD show complete DNA retardation even at lower N:P ratios [36] indicating the stronger interactive forces of large polyanionic species in comparison to siRNA. Nanoparticle stability is necessary for extracellular siRNA protection; however, disassembly is needed to allow RNA-mediated gene silencing through interaction with intracellular components such as RISC. It has been reported that stable chitosan/DNA complexes are beneficial for the protection of DNA in the cellular endosomallysosomal compartments, but restricts the release of DNA to the nucleus that results in low gene expression [25,37]. This emphasises the importance of a sufficient balance between protection and release for siRNA biological functionality. Using a polyanion displacement method, siRNA was rapidly displaced from the complexes upon addition of polyanionic PAA (Fig. 3, Lanes 8–13). The demonstration of weak electrostatic interaction between chitosan and siRNA could be potentially exploited under intracellular conditions to enable siRNA release for increased siRNA activity. Importantly, the displaced siRNAs were structurally intact (in comparison to naked siRNA control) that confirmed the integrity of siRNA was maintained after nanoparticle formation and disassembly. 3.3. In vitro gene silencing and cytotoxicity of chitosan/ siRNA nanoparticles The stably expressing EGFP cell line (H1299 green cells) was used to investigate the gene silencing efficiency of the various chitosan/siRNA formulations. When compared to the non-transfected cells (negative control), chitosan/ siRNA formulations (N:P 50) prepared with low Mw (C9–95, C12–77) or low DD (C173–54) showed low or no EGFP knockdown, whereas those prepared from higher Mw or DD showed greater gene silencing efficiencies of 45% (C65–78), 54% (C170–84) and 65% (C114–84). In comparison, the commercial TransIT-TKO (positive control) resulted in 85% EGFP knockdown (Fig. 4A). A cytotoxicity assay performed using the same chitosan formulations showed a 20–40% reduction in cell viability comparable with the commercial TransIT-TKO (Fig. 4B). The level of EGFP knockdown increased at higher N:P ratios (50 and 150) in comparison to low N:P (2 and 10) formulations (Fig. 4C). Nanoparticles formed at N:P 150 showed the greatest level (80%) of EGFP knockdown. The total chitosan used at high N:P ratios does not completely participate in the formation of the nanoparticle, as demonstrated by excess chitosan found in the filtrate when centrifugal concentration was performed (data not shown). In our studies, increased nanoparticle stability, shown by a decrease in siRNA migration in nanoparticles formed at high N:P (Fig. 5) demonstrates the role of excess chitosan in particle stability. We propose that a proportion

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of chitosan more likely loosely associates with nanoparticles at high N:P and contributes to the improved stability and increased gene silencing. Moreover, removal of excess polycation prior to transfection resulted in virtually no cellular knockdown (data not shown). This suggests the requirement of stabilised particles for cellular entry or a possible role of excess chitosan in cellular permeation. Based on the findings shown in Fig. 4C, nanoparticles formed at N:P 150 were selected to further investigate the influence of chitosan Mw and DD on cellular EGFP knockdown. When high Mw chitosan (C114–84 and C170–84) was used gene silencing was increased to 80% that was comparable with commercial TransIT-TKO (Fig. 4D). In contrast, low levels of knockdown were again achieved with low Mw and DD chitosans. The specificity of knockdown was confirmed using mismatch siRNA formulations (containing 4 nucleotides substitutions as control to EGFP) although a significant nonspecific knockdown with mismatch siRNA was observed. It has been reported that perfect complementary of base pairs between siRNA and targeted mRNA leads to the degradation of mRNA, while several mismatched base may result in the silencing via translation pathway [38]. Based on the above report and the fact that we measure protein (EGFP) expression, it is reasonable that nanoparticles formed by complexing mismatch siRNA might show some silencing efficiency (although still less than specific siRNA, Fig. 4D). The observation that the ability of mismatch siRNA to knockdown EGFP expression is dependent on the formulation (chitosan Mw and DD) is not surprising since the activity follows closely the trend observed for the matching siRNA, namely that highest knockdown with both mismatch and specific siRNA was observed with formulations using higher chitosan Mw and DD (C65–78, C114–84, C170–84, Fig. 4D). This may reflect increased particle stability shown with these formulations. Overall, a slight increase in cytotoxicity was found with formulations at N:P 150 compared to N:P 50 that may reflect the increased level of free excess chitosan in the system. Cytotoxicity levels, however, were no higher than those observed for the commercial reagent TransIT-TKO (Fig. 4E). The biological knockdown results are fully consistent with the described physicochemical properties of chitosan/ siRNA nanoparticles that suggest a correlation between the stability of the nanoparticles and gene silencing efficiency in vitro. Chitosan samples with high Mw of 114 and 170 kDa and DD over 80% are highly suitable for formulation of siRNA carrier systems. These formulations provide an efficient balance between appropriate protection and release, resulting in high in vitro gene silencing efficiencies equivalent to what can be achieved with existing commercial reagents. As discussed earlier, chitosan samples with high Mw and DD have longer molecular chains and higher charge density which enable efficient siRNA interaction and the formation of discrete nanoparticles. Nanoparticles formed with C114–84 and C170–84 have

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Fig. 4. Flow analysis of average GFP expression showing the gene silencing efficiency and cytotoxicity of chitosan/siRNA nanoparticles. (A) In vitro transfection using nanoparticles prepared with 6 chitosan samples of different Mw and DD at N:P 50; (B) cytotoxicity assay of nanoparticles prepared with 6 chitosan samples of different Mw and DD at N:P 50; (C) in vitro transfection using nanoparticles prepared at different N:P ratio (D) in vitro transfection using nanoparticles prepared with 6 chitosan samples of different Mw and DD at N:P 150; (E) cytotoxicity assay of nanoparticles prepared with 6 chitosan samples of different Mw and DD at N:P 150. Non-transfected cells (Control), nanoparticles prepared with commercial transfection reagent, TransITTKOs, is included as a positive control. (Error bars represent7SD.)

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siRNA Fig. 5. Effect of N:P ratio on the complex stability of nanoparticles formulated with chitosan (C170–84) studied by native PAGE. Lane 1: siRNA-GFP; Lane 2: N:P 2; Lane 3: N:P 10; Lane 4: N:P 50; Lane 5: N:P 150.

small size, higher surface charge and higher complex stability, which should enable them to become more easily endocytosed into cells and withstand endosomal-lysosomal conditions prior to cargo release resulting in higher gene silencing efficiency. With the increase of N:P ratio to 150, the complex stability of nanoparticles was increased as indicated by the slower mobility of siRNA complexed with C170–84 (Fig. 5). The increasing stability could be attributed to increased levels of loosely bound chitosan at higher N:P. The excess chitosan loosely bound to the outer surface of nanoparticles should promote binding and uptake across anionic cell surfaces whilst providing subsequent protection against siRNA breakdown within endosomal compartments. In addition, exploitation of the endosomolytic property of chitosan [39,40] should improve cytoplasmic localisation prior to interaction with RISC.

4. Conclusion Interpolyelectrolyte complexes between chitosan and siRNA were used to form nanoparticles for siRNA delivery and gene silencing applications. Physicochemical properties such as size, zeta potential and complex stability of the nanoparticles were shown to be highly dependent on the structural parameters Mw and DD of the chitosan polymer. It was found that chitosan/siRNA nanoparticles formed using high Mw (114 and 170 kDa) and DD (84%) chitosan formed at N:P 150 were the most stable and exhibited the highest (80%) in vitro gene knockdown, which was comparable to the best we have observed using other commercial reagents. This work demonstrates the application of chitosan as a non-viral carrier for siRNA and the pivotal role of polymeric properties in the optimisation of gene silencing protocols.

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Acknowledgements The authors thank Claus Bus, Rita Rosendahl Hansen and Tonnie Holm Nygaard for excellent technical assistance. We also thank Dr. Anne Chauchereau (CNRS, Villejuif, France) for providing the EGFP cell line and Dr. Alexander Schmitz for help with flow cytometry. The work was supported by the Danish Technical Research Council, the Danish Strategic Research Council, the Danish Cancer Society and the Sixth Research Framework Programme of the European Union, Project RIGHT (LSHB-CT-2004-005276).

References [1] Plasterk RH. RNA silencing: the genome’s immune system. Science 2002;296:1263–5. [2] Milhavet O, Gary DS, Mattson MP. RNA interference in biology and medicine. Pharmacol Rev 2003;55(4):629–48. [3] Fire A, Xu S, Montgomery MK, Kostas SA, Driver SE, Mello CC. Potent and specific genetic interference by double-stranded RNA in Caenorhabditis elegans. Nature 1998;391:806–11. [4] Elbashir SM, Harborth J, Lendeckel W, Yalcin A, Weber K, Tuschl T. Duplexes of 21-nucleotide RNAs mediate RNA interference in cultured mammalian cells. Nature 2001;411:494–8. [5] Hannon GJ, Rossi JJ. Unlocking the potential of the human genome with RNA interference. Nature 2004;431:371–8. [6] Dorsett Y, Tuschl T. siRNAs: applications in functional genomics and potential as therapeutics. Nat Rev Drug Discov 2004;3:318–29. [7] Pouton CW, Seymour LW. Key issues in non-viral gene delivery. Adv Drug Deliver Rev 2001;46:187–203. [8] Xia HB, Mao QW, Paulson HL, Davidson BL. siRNA-mediated gene silencing in vitro and in vivo. Nat Biotechnol 2002;20:1006–10. [9] Barton GM, Medzhitov R. Retroviral delivery of small interfering RNA into primary cells. PNAS 2002;99(23):14943–5. [10] Lundstrom K. Latest development in viral vectors for gene therapy. Trends Biotechnol 2003;21(3):117–22. [11] Soutschek J, Akinc A, Bramlage B, Charisse K, ZConstien R, Donoghue M, et al. Therapeutic silencing of an endogenous gene by systemic administration of modified siRNAs. Nature 2004;432:173–8. [12] Ichim TE, Li M, Qian H, Popov IA, Rycerz K, Zheng XF, White D, Zhong R, Min WP. RNA interference: a potent tool for gene-specific therapeutics. Am J Transplant 2004;4:1227–36. [13] Schiffelers RM, Ansari A, Xu J, Zhou Q, Tang QQ, Storm G, et al. Cancer siRNA therapy by tumor selective delivery with ligandtargeted sterically stabilized nanoparticle. Nucleic Acids Res 2004; 32(19):e149. [14] Bitko V, Musiyenko A, Shulyayeva O, Barik S. Inhibition of respiratory viruses by nasally administered siRNA. Nat Med 2005; 11(1):50–5. [15] Borchard G. Chitosans for gene delivery. Adv Drug Deliver Rev 2001;52:145–50. [16] Liu WG, Yao KD. Chitosan and its derivatives—a promising nonviral vector for gene transfection. J Control Release 2002;83:1–11. [17] Mansouri S, Lavigne P, Corsi K, Benderdour M, Beaumont E, Fernandes JC. Chitsoan-DNA nanoparticles as non-viral vectors in gene therapy: strategies to improve transfection efficacy. Eur J Pharm Biopharm 2004;57:1–8. [18] Richardson SCW, Kolbe HVJ, Duncan R. Potential of low molecular mass chitosan as a DNA delivery system: biocompatibility, body distribution and ability to complex and protect DNA. Int J Pharm 1999;178:231–43. [19] Corsi K, Chellat F, Yahia L, Fernandes JC. Mesenchymal stem cells, MG63 and HEK 293 transfection using chitosan-DNA nanoparticles. Biomaterials 2003;24:1255–64.

ARTICLE IN PRESS 1288

X. Liu et al. / Biomaterials 28 (2007) 1280–1288

[20] Chandy T, Sharma CP. Chitosan—as a biomaterial. Biomater Artif Cell Artif Organ 1990;18:1–24. [21] Illum L, Farraj NF, Davis SS. Chitosan as a novel nasal delivery system for peptide drugs. Pharm Res 1994;11:1186–9. [22] Verhoef JC, Junginger HE, Thanou M. Chitosan and its derivatives as intestinal absorption enhancers. Adv Drug Deliver Rev 2001; 50:S91–S101. [23] Carren˜o-Go´mez B, Duncan R. Evaluation of the biological properties of soluble chitosan and chitosan microspheres. Int J Pharm 1997;148:231–40. [24] Mao HQ, Roy K, Truong-Le VL, Janes KA, Lin KY, Wang Y, et al. Chitosan-DNA nanoparticles as gene carriers: synthesis, characterization and transfection efficiency. J Control Release 2001;70: 399–421. [25] Li XW, Lee DKL, Chan ASC, Alpar HO. Sustained expression in mammalian cells with DNA complexed with chitosan nanoparticles. Biochim Biophys Acta 2003;1630:7–18. [26] Ko¨ping-Ho¨gga˚rd M, Va˚rum KM, Issa M, Danielsen S, Christensen BE, Stokke BT, Artursson P. Improved chitosan-mediated gene delivery based on easily dissociated chitosan polyplexes of highly defined chitosan oligomers. Gene Ther 2004;11:1441–52. [27] Kiang T, Wen J, Lim HW, Leong KW. The effect of the degree of chitosan deacetylation on the efficiency of gene transfection. Biomaterials 2004;25:5293–301. [28] Liu WG, Sun SJ, Cao ZQ, Zhang X, Yao KD, Lu WW, et al. An investigation on the physicochemical properties of chitosan/DNA polyelectrolyte complexes. Biomaterials 2005;26:2705–11. [29] Howard KA, Rahbek UL, Liu XD, Damgaard CK, Glud SZ, Anderson MØ, et al. RNA interference in vitro and in vivo using a chitosan/siRNA nanoparticle system. Mol Ther 2006;14:476–84.

[30] Va˚rum KM, Ottøy MH, Smidsrød O. Water-solubility of partially N-acetylated chitosan as a function of pH: effect of chemical composition and depolymerization. Carbohydr Polym 1994;25: 65–70. [31] Ottøy MH, Va˚rum KH, Christensen BE, Anthonsen MW, Smidsrød O. Preparative and analytical size-exclusion chromatography of chitosans. Carbohydr Polym 1996;31:253–61. [32] Hirano S, Ohe Y, Ono H. Selective N-acetylation of chitosan. Carbohydr Res 1976;47:315–20. [33] Muzzarelli RAA, Rocchetti R. Determination of the degree of acetylation of chitosans by first derivative ultraviolet spectrophotometry. Carbohydr Polym 1985;5:461–72. [34] Khan TA, Peh KK, Ch’ng HS. Reporting degree of deacetylation values of chitosan: the influence of analytical methods. J Pharm Pharm Sci 2002;5:205–12. [35] www.imgemet.com. [36] Sato T, Ishii T, Okahata Y. In vitro gene delivery mediated by chitosan. Effect of pH, serum, and molecular mass of chitosan on the transfection efficiency. Biomaterials 2001;22:2075–80. [37] MacLaughlin FC, Mumper RJ, Wang JJ, Tagliaferri JM, Gill I, Hinchcliffe M, et al. Chitosan and depolymerized chitosan oligomers as condensing carriers for in vivo plasmid delivery. J Control Release 1998;56:259–72. [38] Doench J, Petersen C, Sharp P. siRNAs can function as miRNAs. Genes Dev 2003;17:438–42. [39] Huang M, Fong CW, Khor E, Lim LY. Transfection efficiency of chitosan vectors: effect of polymer molecular weight and degree of deacetylation. J Control Release 2005;106:391–406. [40] Ishii T, Okahata Y, Sato T. Mechanism of cell transfection with plasmid/chitosan complexes. Biochim Biophys Acta 2001;1514:51–64.