Size-Dependent Knockdown Potential of siRNA-Loaded Cationic

Oct 22, 2014 - Interestingly, primary cell uptake and colocalization studies with lysosomal compartments revealed that only these small sized nanogels...
1 downloads 9 Views 10MB Size
Download PDF
Article pubs.acs.org/Biomac

Size-Dependent Knockdown Potential of siRNA-Loaded Cationic Nanohydrogel Particles Lutz Nuhn,† Stephanie Tomcin,‡ Kanjiro Miyata,§ Volker Mailan̈ der,‡,∥ Katharina Landfester,‡ Kazunori Kataoka,§,⊥ and Rudolf Zentel*,† †

Institute of Organic Chemistry, Johannes Gutenberg-University, Duesbergweg 10-14, D-55099 Mainz, Germany Max-Planck-Institute for Polymer Research, Ackermannweg 10, D-55128 Mainz, Germany ∥ III. Medical Clinic (Hematology, Oncology and Pneumology), University Medical Center of the Johannes-Gutenberg University Mainz, Langenbeckstraße 1, D-55131 Mainz, Germany § Center for Disease Biology and Integrative Medicine, Graduate School of Medicine, and ⊥Department of Materials Engineering, Graduate School of Engineering, The University of Tokyo, Hongo 7-3-1, Bunkyo-ku, Tokyo 113-0033, Japan ‡

S Supporting Information *

ABSTRACT: To overcome the poor pharmacokinetic conditions of short double-stranded RNA molecules in RNA interference therapies, cationic nanohydrogel particles can be considered as alternative safe and stable carriers for oligonucleotide delivery. For understanding key parameters during this process, two different types of well-defined cationic nanohydrogel particles were synthesized, which provided nearly identical physicochemical properties with regards to their material composition and resulting siRNA loading characteristics. Yet, according to the manufacturing process using amphiphilic reactive ester block copolymers of pentafluorophenyl methacrylate (PFPMA) and tri(ethylene glycol)methyl ether methacrylate (MEO3MA) with similar compositions but different molecular weights, the resulting nanohydrogel particles differed in size after cross-linking with spermine (average diameter 40 vs 100 nm). This affected their knockdown potential significantly. Only the 40 nm sized cationic nanogel particles were able to generate moderate gene knockdown levels, which lasted, however, up to 3 days. Interestingly, primary cell uptake and colocalization studies with lysosomal compartments revealed that only these small sized nanogels were able to avoid acidic compartments of endolysosomal uptake pathways, which may contribute to their knockdown ability exclusively. To that respect, this sizedependent intracellular distribution behavior may be considered as an essential key parameter for tuning the knockdown potential of siRNA nanohydrogel particles, which may further contribute to the development of advanced siRNA carrier systems with improved knockdown potential.

■

INTRODUCTION RNA interference (RNAi)1,2 has opened alternative therapeutic pathways to treat pathogenic genes by silencing their protein expression,3,4 yet the pharmacokinetic conditions for short double-stranded RNA molecules to enter the cytoplasm as RNAi’s site of action are rather poor, especially after systemic application.5 Several delivery approaches have been developed to protect small interfering RNA (siRNA) from enzymatic degradation or rapid renal clearance and to promote cellular uptake of this highly negatively charged oligonucleotide into targeted cells: For example, small molecules like cationic lipids can successfully be used to formulate nanosized siRNA lipoplexes that mediate gene knockdown.6 Anderson et al., for instance, developed various libraries of lipid-like molecules to optimize knockdown efficiencies for both in vitro as well as in vivo applications.7−12 Generally, siRNA formulations using cationic lipids have so far mostly been applied in first clinical trials of nanotherapeutics with promising results.13 © 2014 American Chemical Society

Alternatively, cationic copolymers can facilitate siRNA delivery into the cytoplasm by electrostatic interaction with the oligonucleotides forming nanosized polyplexes, too.14−16 Among various multifunctional cationic copolymer systems a cyclodextrin-based siRNA polyplex system decorated with both adamantane-PEG and adamantane-PEG-transferrin could also be translated into first clinical trials providing first evidence of RNAi in humans,17,18 however, rapid disassembly was found at the glomerular basement membrane within the kidney recently.19 Thus, for siRNA carrying moieties, a high degree of stability under physiological conditions is of major importance, especially for systemic applications.20 Many polymeric delivery strategies originally developed for plasmid DNA (pDNA) in gene therapy can often not be adopted to siRNA delivery Received: August 6, 2014 Revised: October 1, 2014 Published: October 22, 2014 4111

dx.doi.org/10.1021/bm501148y | Biomacromolecules 2014, 15, 4111−4121

Biomacromolecules

Article

indicated. Oregon Green 488 cadaverine was obtained from Invitrogen. Pentafluorophenol was obtained from Fluorochem (Great Britain, U.K.). Heparin sodium salt from hog intestine was obtained from TCI Europe. Phosphate buffered saline (PBS) was obtained from Fisher BioReagents containing 137 mM NaCl, 11.9 mM phosphates, and 2.7 mM KCl with a pH at 7.4. 4-(2-Hydroxyethyl)-1piperazineethanesulfonic acid (HEPES) buffer was prepared at 10 mM and its pH was adjusted to 7.4 with 1 M NaOH. Dialysis was performed using Spectra/Por 3 membranes obtained from Carl Roth GmbH+Co.KG (Germany) molecular weight cut off 8000−10000 g/ mol. All siRNA sequences were purchased from Hokkaido System Science Co., Ltd. (Hokkaido, Japan) and used as follows:

without alterations, since the siRNA as anionic cargo is simply too small in size to stabilize the spontaneously self-assembled nano-object.21 The advantageous simplicity of both siRNA lipoplex as well as polyplex delivery systems is, therefore, diminished as siRNA is usually the only structure-promoting counterpart during the nanoparticle self-assembling process but cannot guarantee enough stability under physiological relevant conditions. Yet, attempts to introduce covalent cross-linkers after polyplex formulations have shown to increase stability and to improve efficacy of these carriers substantially.22−25 Alternatively, well-defined nanosized siRNA delivery systems that are predefined in size and shape but independent from their siRNA cargo can serve as ideal carriers with improved stability properties for in vivo applications. Nanohydrogel particles can provide all these properties and have, therefore, gained more attraction to siRNA delivery recently.26,27 As examples, De Smedt et al. developed a synthetic strategy for cationic nanogel carriers based on biodegradable dextran for cellular siRNA delivery,28,29 and Siegwart et al. synthesized libraries of cationic nanogel particles based on synthetic block copolymers, which were screened for siRNA in vivo transfection to liver hepatocytes successfully.30 To that respect, we have recently developed a novel synthetic route to access precise cationic nanohydrogel particles with adjustable sizes for cellular delivery of siRNA:31 By reversible addition−fragmentation chain transfer (RAFT) block copolymerization of pentafluorophenly methacrylate (PFPMA) and tri(ethylene glycol) methyl ether methacrylate) (MEO3MA) well-defined amphiphilic reactive ester block copolymers can be generated that self-assemble into nanosized micellar aggregates in polar-aprotic solvents (e.g., dimethyl sulfoxide). These precursor structures can permanently be locked-in by a crosslinking reaction of the hydrophobic pentafluorophenyl ester moieties inside the core with spermine. The resulting cationic nanohydrogel particles, now predefined in size, provide a hydrophilic, cross-linked cationic core for efficient siRNA complexation without changing the nanoparticle’s morphology. So far, we could demonstrate that these nanohydrogel particles facilitate uptake of siRNA into cells over time properly.31 Moreover, their stability properties could recently be investigated in human blood serum as well as in the bloodstream of mice.32 However, detailed investigation about the material’s cellular knockdown potential mediated by its payload has not been studied so far. In the present study two sets of well-defined cationic nanohydrogel particles were synthesized with diameters of about 40 and 100 nm, respectively. Both materials show similar physicochemical properties as well as loading efficiencies and release capabilities of siRNA, yet differ in their knockdown potential in luciferase expressing HeLa cells significantly. Only small siRNA loaded nanohydrogel particles with diameters of around 40 nm were able to induce gene silencing that lasts for about 3 days. Cell uptake studies provided preliminary information about those particles being able to avoid endolysosomal uptake pathways as less colocalization with acidic compartments was found over time. Therefore, according to previous studies,33,34 our results support that size-dependent uptake mechanism may contribute to the knockdown potential of siRNA nanocarriers.

■

(1) Firefly GL3 luciferase siRNA (luc-siRNA) sense strand: 5′CUUACGCUGAGUACUUCGAdTdT-3′; antisense strand: 3′-dTdTGAAUGCGACUCAUGAAGCU-5′. (2) Scramble luciferase (scr-siRNA) sense strand: 5′-UUCUCCGAACGUGUCACGUdTdT-3′; antisense strand: 3′-dTdTAAGAGGCUUGCACUGAGCA-5′. Instrumentation. All 1H-, 13C-, and 19F-NMR spectra were recorded on Bruker 300 and 400 MHz FT NMR spectrometers, respectively. Chemical shifts (δ) are given in ppm relative to TMS. Samples were prepared in deuterated solvents and their signals referenced to residual nondeuterated solvent signals. The polymers’ molecular weight was determined by size exclusion chromatography (SEC) in tetrahydrofuran (THF) as solvent and with the following parts: pump PU 1580, auto sampler AS1555, UV-detector UV 1575 (detection at 254 nm), RI-detector RI 1530 from JASCO, Germany. Columns were used from MZ-Analysentechnik, Germany: MZ-Gel SDplus 102 Å and MZ-Gel SDplus 106 Å. Calibration was done using polystyrene standards purchased from Polymer Standard Services, Germany. UV−vis spectra were recorded using a Jasco V-630 Spectrophotometer (1 cm × 1 cm quartz cell). Block Copolymer Syntheses. The syntheses of poly(tri(ethylene glycol) methyl ether methacrylate)-block-poly(pentafluorophenyl methacrylate) block copolymer P(MEO3MA)25-b-P(PFPMA)38 P1 (Mn = 14200 g/mol, PDI = 1.24) and poly(pentafluorophenyl methacrylate)-block- poly(tri(ethylene glycol) methyl ether methacrylate) block copolymer P(PFPMA)61-b-P(MEO3MA)40 P2 (Mn = 25100 g/mol, PDI = 1.23) were performed by RAFT polymerization as recently described.31 Analytical data can be found in the Supporting Information (for block copolymer P(MEO3MA)25-b-P(PFPMA)38 P1: Scheme S1 and Figures S1−S5; for block copolymer P(PFPMA)61-bP(MEO3MA)40 P2: Scheme S2 and Figures S6−S10). Synthesis of Cationic Nanohydrogel Particle NP1. P(MEO3MA)25-b-P(PFPMA)38 block copolymer P1 (30.0 mg; 2.11 μmol polymer or 80.3 μmol reactive ester) was transferred into a round-bottom flask equipped with a stir bar and anhydrous DMSO (3.0 mL). Supported by sonication for 1 h, the polymer could be dispersed in DMSO under a nitrogen atmosphere forming selfassembled micellar aggregates. For fluorescence labeling, Oregon Green cadaverine (16.0 μL of a 2.5 mg/mL solution in DMSO; 0.08 μmol) was added first, followed by triethylamine (56.0 μL; 404.0 μmol) and then spermine (8.1 mg; 40.1 μmol) for cross-linking. The flask containing the reaction mixture was immersed in an oil bath at 50 °C under vigorously stirring. After 64 h, a 19F-NMR sample (0.2 mL dissolved in 0.4 mL DMSO-d6) was taken showing complete reactive ester conversion (compare Supporting Information, Figure S11). Yet, to remove further traces of polymer bound pentafluorophenol below the NMR detection limit, excess of non-cross-linking methoxy triethylene glycol amine (13.1 mg; 80.3 μmol) was added and the reaction mixture was stirred for additional 25 h at 50 °C. To remove small molecular byproducts, the reaction mixture was purified by dialysis against Millipore water for several days (including water exchange twice every day) and subsequent lyophilization affording NP1 (19.7 mg, 83%) as a voluminous orange powder. Synthesis of Cationic Nanohydrogel Particle NP2. P(PFPMA)61-b-P(MEO3MA)40 block copolymer P2 (51.9 mg; 2.10 μmol polymer or 127 μmol reactive ester) was transferred into a round-bottom flask equipped with a stir bar and anhydrous DMSO

MATERIALS AND METHODS

Materials. All chemicals and solvents were purchased from Acros Organics or Sigma-Aldrich and used as received unless otherwise 4112

dx.doi.org/10.1021/bm501148y | Biomacromolecules 2014, 15, 4111−4121

Biomacromolecules

Article

(5.2 mL). Supported by sonication for 1 h the polymer could be dispersed in DMSO under nitrogen atmosphere forming selfassembled micellar aggregates. For fluorescence labeling, Oregon Green cadaverine (26.7 μL of a 2.5 mg/mL solution in DMSO; 0.13 μmol) was added first, followed by triethylamine (112 μL; 808.0 μmol), and then spermine (13.6 mg; 63.5 μmol) for cross-linking. The flask containing the reaction mixture was immersed in an oil bath at 50 °C under vigorously stirring. After 16 h, a 19F-NMR sample (0.2 mL dissolved in 0.4 mL DMSO-d6) was taken showing complete reactive ester conversion (compare Supporting Information, Figure S12). Yet, to remove further traces of polymer bound pentafluorophenol below the NMR detection limit, excess of non-cross-linking methoxy triethylene glycol amine (22.0 mg; 134 μmol) was added and the reaction mixture was stirred for additional 20 h at 50 °C. To remove small molecular byproducts, the reaction mixture was purified by dialysis against Millipore water for several days (including water exchange twice every day) and subsequent lyophilization affording NP2 (31.0 mg, 79%) as a voluminous orange powder. Cationic Nanohydrogel Particle Loading with siRNA. The lyophilized cationic nanohydrogel particles could easily be redispersed in water, PBS, or HEPES buffer at given concentrations supported by sonication. For loading with siRNA, the samples were mixed at the given concentrations with the corresponding weight-to-weight ratios of nanohydrogel particle to siRNA and incubated for 30 min at room temperature. To verify siRNA loading agarose gel electrophoresis was performed after each complexation. Nanohydrogel Particle Characterization. Dynamic light scattering (DLS) of the nanohydrogel particles alone or loaded with siRNA at the given weight-to-weight ratio (nanohydrogel particle to siRNA) was done using a ZetaSizer Nano ZS instrument (Malvern Instruments Ltd., Malvern, U.K.) equipped with a He−Ne laser (λ = 633 nm) as incident beam. Typically, 20 μL of each sample at 0.5 mg/ mL of nanohydrogel in HEPES was loaded into a Zen 2112 lowvolume cuvette and measured at 25 °C. The obtained data were analyzed by cumulant fitting for z-average mean diameter and PDI and CONTIN fitting method for particle diameter size distribution, respectively. For electron microscopy analysis, samples of the nanohydrogel particles were prepared at 0.02 mg/mL in water. Scanning electron microscopy (SEM) was performed after drying 5 μL of each sample on silica wafers. Images were recorded by a field emission microscope (LEO 1530 Gemini) working at an acceleration voltage of 0.2 kV. Transmission electron microcopy (TEM) was performed after dropcasting 4 μL of each the sample on a 300-mesh carbon-coated copper grid without any further staining. Images were recorded by a FEI Tecnai F20 transmission electron microscope working at 200 kV. Zeta potential analysis of the cationic nanohydrogel particles was done by Doppler electrophoresis using a ZetaSizer Nano ZS instrument (Malvern Instruments Ltd., Malvern, U.K.) equipped with a He−Ne laser (λ = 633 nm) as incident beam. For each cationic nanogel sample, 1 mL of a 0.1 mg/mL solution in Millipore H2O was loaded into a folded DTS1070 capillary cell and measured at 25 °C. The obtained data were analyzed by applying the obtained electrophoretic mobility to the Smoluchowski equation. Agarose Gel Electrophoresis. For agarose gel electrophoresis experiments, samples were usually prepared with 70 ng siRNA and nanohydrogel particles at given weight-to-weight ratios in PBS and incubated for 30 min for complete complexation. In case of competitive release studies with heparin, each sample was afterward mixed with an aliquot of heparin sodium salt from a stock solution (0.5 mg/mL) to adjust the given heparin concentration and incubated for another 30 min. Each sample was mixed with 6× loading buffer (30% glycerol, 0.25% bromophenol blue, and 0.25% xylene cyanol) prior to loading on a 0.5% agarose gel containing GelRed (Biotium). Electrophoresis was performed in TBE buffer (89 mM tris(hydroxymethyl)-aminomethane, 89 mM boric acid, and 2 mM Na2EDTA, pH 8) at 120 V for 30 min, and upon excitation at 365 nm, fluorescence was imaged by a conventional digital camera. Endogenous Luciferase Gene Silencing Assay. Human cervix adenocarcinoma cells (HeLa-Luc) stably expressing firefly luciferase

(Promega Corp.) were kept in Dulbecco’s Modified Eagle Medium (DMEM), supplemented with 10 vol % fetal calf serum (FCS), 100 units/mL penicillin, and 100 μg/mL streptomycin (all from Invitrogen). Cells were grown in a humidified incubator at 37 °C and 5% CO2. One day prior to the experiments, adherent cells were detached using 0.5% trypsin (Invitrogen) and seeded in 48-well plates at a density of 8500 cells per well. After readhesion overnight cells were incubated with nanohydrogel particles (alone or loaded with siRNA at a weight-to-weight ratio nanohydrogel particle to siRNA of 10:1) at the given concentrations (50, 100, 200, and 400 nM of siRNA or 7, 14, 28, and 56 mg/L of nanohydrogel particles, respectively) in cell culture media. After 48 h incubation, cells were washed with 0.2 mL of PBS and lysed with 0.2 mL of cell culture lysis buffer (Promega). The luciferase activity of the lysates was determined from the photoluminescence intensity using the Luciferase Assay System (Promega) and a luminometer Mithras LB 940 (Berthold Technologies). The relative luminescence (%) was calculated from the obtained luminescence intensity as a percentage to control wells without applying nanohydrogel particles (HEPES buffer only). The results are presented as mean and standard deviation obtained from four samples. Cell Viability Assay. After 48 h incubation of HeLa-Luc cells with nanohydrogel particles (alone or loaded with siRNA at a weight-toweight ratio nanohydrogel particle to siRNA of 10:1) at the given concentrations in cell culture media, their viability was analyzed using a Cell Counting Kit-8 (DOJINDO Laboratories, Kumamoto, Japan), following the manufacturer’s protocol. The absorbance at 450 nm in each well was measured using a microplate reader (Model 680, BIORAD). The cell viability in each well was calculated as a percentage to control wells without applying nanohydrogel particles (HEPES buffer only). The results are presented as mean and standard deviation obtained from four samples. Long-Term Luciferase Gene Silencing for Knockdown Kinetics. Human cervix adenocarcinoma cells stably expressing luciferase (HeLa-Luc) were seeded in 35 mm Petri dishes (25000 cells/dish) and allowed to attach for 24 h. After cell attachment, the medium was removed and replaced with 2 mL of DMEM medium (supplemented with FBS and antibiotics) containing 100 μM luciferin and siRNA loaded nanohydrogel particles (400 nM siRNA). For each analysis, control samples were prepared by the addition of media diluted with HEPES instead of nanogel solution. Samples were placed into a Kronos real-time photon countable incubator (Atto, Japan), and the luminescence intensity was measured periodically over an 80 h time period until complete cell confluence was reached, with the temperature and CO2 maintained at 37 °C and 5%, respectively. Relative luminescence was determined by dividing the average luminescence intensity of treated samples by the average luminescence intensity of control samples (for each n = 4). Confocal Laser Scanning Microscopy Studies. Human cervix adenocarcinoma cells (HeLa) were kept in Dulbecco’s modified Eagle medium (DMEM) without phenol Red, supplemented with 10 vol % fetal calf serum (FCS), 100 units/mL penicillin together with 100 μg/ mL streptomycin, and 1 vol % GlutaMAX (all from Invitrogen, Germany). Cells were grown in a humidified incubator at 37 °C and 5% CO2. One day prior to the experiments, adherent cells were detached using 0.5% trypsin (Invitrogen) and seeded in IBIDI μdish35mm,low (IBIDI, Germany) at a density of 8500 cells per cm2. After readhesion overnight cells were washed once with Dulbecco’s phosphate buffered saline (PBS, Invitrogen, Germany) before treatment. Cells were incubated with nanohydrogel particles at concentrations of 75 μg/mL in PBS for 0.5, 1, 2, 4, 8, 12, and 24 h. After each incubation time point cells were washed three times with PBS and covered with 1 mL of medium. Staining with LysoTracker Red (Invitrogen) was achieved by adding 0.04 μL of a 1 mM stock solution. After incubation for 30 min in a humidified incubator, cells were washed once with PBS, then covered with 1 mL medium and stained with 0.2 μL Cell Mask Deep Red (5 mg/mL, purchased from Invitrogen). Nanoparticles complexed with siRNA were treated as nanoparticles without siRNA (nanohydrogel particles were complexed with siRNA in PBS at 25:1 weight-to-weight ratio nanohydrogel 4113

dx.doi.org/10.1021/bm501148y | Biomacromolecules 2014, 15, 4111−4121

Biomacromolecules

Article

Figure 1. Synthesis of cationic nanohydrogel particles NP1 and NP2 based on cross-linking of self-assembled micellar aggregates of block copolymers P(MEO3MA)25-b-P(PFPMA)38 P1 and P(PFPMA)61-b-P(MEO3MA)40 P2 with spermine enabling complexation with siRNA.

Figure 2. Characterization of cationic nanohydrogel particles NP1 and NP2: (1) scanning electron microscopy (SEM) images; (2) dynamic light scattering (DLS) results for NP1 with hydrodynamic diameter Dh = 36.6 nm and PDI = 0.21 and for NP2 with hydrodynamic diameter Dh = 103.1 nm and PDI = 0.27; (3) transmission electron microscopy (TEM) images.

■

particle to siRNA prior to the cell culture experiments). To demonstrate intracellular uptake of nanohydrogel particle (loaded with or without siRNA) and potential accumulation in lysosomes, confocal laser scanning microscopy (CLSM) was applied directly after staining with CellMask Deep Red. Images were taken with Leica LAS AF Software on a Leica TCS SP5 II microscope equipped with five lasers (multiline argon laser with 458, 476, 488, 496, 514 nm, a 561 nm DPSS laser, a HeNe laser with 594 and 633 nm, and a 592 nm CW STED laser) with a HCX PL APO CS 63×/1.4−0.6 oil-immersion objective. Nanohydrogel particles labeled with Oregon Green were excited with a 488 nm argon laser and detected at 500−550 nm. To detect lysosomes stained with LysoTracker Red a 561 nm DPSS laser was used for excitation and emission was detected at 575−610 nm. The detection of the CellMask Deep Red labeled cell membranes occurred at 655−730 nm when excited at 633 nm with a HeNe laser. Images were taken with a pinhole size of 1 AE, a line average of 2, a resolution of 1024 × 1024 8-bit-pixels, and by using PMTs for detection. To avoid crosstalk a serial mode was applied for imaging. Image processing was done with Leica LAS AF software and ImageJ and quantitative analysis for nanogel uptake and lysosomal colocalization by calculation of the Pearson’s Correlation Coefficient (PCC) was done using Volocity software (PerkinElmer). Statistical Analysis. The p values were determined by the Student’s t test using a two-tailed distribution and two-sample unequal variance with the t test function of Microsoft Excel. The p values of less than 0.05 were considered as statistically significant.

RESULTS 1. Synthesis of Cationic Nanohydrogel Particles. Precise nanosized siRNA hydrogels are generally considered to provide improved stability properties for siRNA delivery.26,27 Our group recently developed a synthetic pathway to generate well-defined cationic nanohydrogel particles for siRNA delivery by cross-linking self-assembled precursor polymers synthesized via RAFT block copolymerization: Amphiphilic reactive ester block copolymers composed of pentafluorophenyl methacrylate (PFPMA) and tri(ethylene glycol) methyl ether methacrylate (MEO3MA) self-assemble into nm-sized micellar aggregates in polar protic solvents like dimethyl sulfoxide (DMSO). The fluorinated reactive ester moieties inside the core can react with the amine functionalities of spermine stoichiometrically resulting in permanently cross-linked nanohydrogel particles with cationic cores provided by residual amine functionalities of spermine. Interestingly, based on the composition and the molecular weight of the amphiphilic reactive ester precursor block copolymer the size and the morphology of the resulting nanohydrogel particle can be adjusted effectively.31 For the present study two sets of nanohydrogel particles were synthesized: For nanohydrogel particle NP1 an amphiphilic reactive ester block copolymer P(MEO3MA)25-bP(PFPMA)38 P1 was synthesized by RAFT block copolymerization with an average molecular weight of Mn = 14200 g/ 4114

dx.doi.org/10.1021/bm501148y | Biomacromolecules 2014, 15, 4111−4121

Biomacromolecules

Article

transmission electron microscopy images of the dried nanohydrogel particles NP1 on solid substrate mostly showed small spherical particles with sizes far below 100 nm, too. Because of the higher molecular weight of precursor polymer P2, on the other hand, the characterization data of NP2 revealed larger cross-linked spherical particles with diameters of about 100 nm. In this case, DLS measurements detected a hydrodynamic diameter of 103.1 nm (PDI = 0.27), and both electron microscopy techniques could visualize spherical particles with sizes of about 100 nm. Besides, zeta potential measurements for both particles were performed in water demonstrating successful transformation of the fluorinated inner core of the self-assembled precursor polymers into a strongly cationic network (compare Supporting Information, Figure S13). The measurements for the small sized nanohydrogel NP1 provided a zeta potential value of +44.4 ± 4.3 mV, while for the larger sized NP2 a zeta potential of +34.9 ± 4.1 mV was found, which lies in a comparable regime. To that respect, both highly cationic nanogels can be utilized for siRNA complexation. 2. Characterization of siRNA-Loaded Cationic Nanohydrogel Particles. SiRNA loading capacity was investigated for the two nanohydrogel particles carefully. To that respect, gel electrophoresis experiments were performed using a 0.5% agarose gel (containing GelRed as nucleic acid stain) in TBE buffer and applying 120 V for 30 min for nucleic acid separation. For each sample, 70 ng of siRNA were incubated with increasing amount of nanohydrogel particles (0.07, 0.70, 1.75, and 3.50 μg) at the given weight-to-weight ratios of nanohydrogel particle to siRNA in PBS for 30 min prior to loading onto the gel (Figure 3.1). For both nanohydrogel particles at the 1:1 weight-to-weight ratio siRNA was not complexed by the particle completely and, thus, able to penetrate into the gel like free siRNA. However, already at the 10:1 ratio, no free siRNA could be detected but, instead, a diffuse smearing close to the sample well suggests compensated siRNA loading into the nanogel. At higher loading ratios, this smearing seemed to be retained more inside the sample well showing identical siRNA complexation properties for both particles. Moreover, similar results could be obtained for both nanohydrogel particle samples complexed with different types of siRNA in HEPES buffer, which were used later on for transfection studies (either luc-siRNA for luciferase gene knockdown or scr-siRNA as control). In this sodium chloride free buffer already at a 7.5:1 weight-to-weight ratio siRNA to nanohydrogel particle sample NP1 or NP2 no free siRNA was visualized by agarose gel electrophoresis (compare Supporting Information, Figure S14.1 for nanohydrogel particle sample NP1 and Figure S14.2 for nanohydrogel particle sample NP2). Interestingly, in all those cases, the siRNA payload did not affect the nanoparticles’ sizes significantly, as DLS measurements showed again similar results for both nanohydrogel particle types loaded with either luc-siRNA for luciferase gene knockdown or scr-siRNA as control: While NP1 provided average hydrodynamic diameters of about 40 nm either with or without different siRNA, the sizes for NP2 seemed to retain again around 100 nm (compare Supporting Information, Figure S15 for NP1 and Figure S16 for NP2). These results are also summarized in Table 1. As previously shown, the siRNA loading process only influences the zeta potential of the cationic nanohydrogel particles by neutralizing the cationic charge of the nanoparticle.32 This could also be observed for the two nanohydrogel particle sample NP1 or NP2 within this study. Interestingly, a neutral zeta potential could generally be

mol and a dispersity of PDI = 1.24 (compare Supporting Information, Scheme S1). Successful block copolymerization, removal of the RAFT-derived dithiobenzoate end group as well as copolymer composition could be verified by size exclusion chromatography (SEC) as well as 1H, 13C, 19F NMR, and UV− vis spectroscopy (compare Supporting Information, Figures S1−S5). Similarly, the amphiphilic reactive ester block copolymer P(PFPMA)61-b-P(MEO3MA)40 P2 was generated by RAFT block copolymerization for the synthesis of nanohydrogel particle NP2 (compare Supporting Information, Scheme S2, as shown previously,31 the order of block copolymerization does not affect nanohydrogel particle synthesis). SEC in this case provided an average molecular weight of Mn = 25100 g/mol and a dispersity of PDI = 1.23, while the UV−vis spectra during end group removal as well as the 1H, 13 C, and 19F NMR spectra of the final block copolymer P2 showed no difference to the previous block copolymer P1 (compare Supporting Information, Figures S6−S10). Therefore, both polymers have similar molar block ratios of MEO3MA/PFPFMA = 4:6, yet their molecular weights differ by a factor of 1.8. Both reactive ester block copolymers can independently be used for nanohydrogel particle synthesis as described in Figure 1. To that respect, each precursor polymer was dispersed in dimethyl sulfoxide supported by sonication forming selfassembled micellar aggregates. For fluorescent labeling 0.001 equiv of Oregon Green cadaverine (related to 1.0 equiv of PFPMA reactive ester) was added to each solution prior to spermine addition for covalent cross-linking. Complete crosslinking was monitored by detecting quantitative release of pentafluorophenol from the polymer via 19F NMR spectroscopy (compare Supporting Information, Figure S11 for nanohydrogel particle NP1 and Figure S12 for nanohydrogel particle NP2). After purification, both types of particles could be redispersed again in Millipore water, HEPES, or PBS buffer supported by sonication easily. They were applied to three independent characterization methods, which, according to the utilized precursor polymer, provide corresponding results that are summarized in Figure 2 and Table 1: On the one hand, for nanohydrogel particle NP1 the small precursor polymer P1 was used affording cross-linked spherical particles with diameters of about 40 nm. Thus, dynamic light scattering data in aqueous solution provided a hydrodynamic diamter of 36.6 nm (PDI = 0.21), and both scanning electron microscopy images as well as Table 1. Characterization of siRNA-Loaded Cationic Nanohydrogel Particles sample NP1

NP2

used precursor polymer P1 P(MEO3MA)25b-P(PFPMA)38

P2 P(PFPMA)61b-P(MEO3MA)40

loaded witha

hydrodynamic diameterb (nm)

PDIb

lucsiRNA scrsiRNA alone lucsiRNA scrsiRNA alone

35.8

0.15

36.5

0.14

36.3 96.1

0.21 0.24

95.2

0.25

103.1

0.27

a Weight-to-weight loading ratio NP:siRNA = 10:1. bDetermined by dynamic light scattering of nanohydrogel samples at 0.5 mg/mL in 10 mM HEPES pH 7.4.

4115

dx.doi.org/10.1021/bm501148y | Biomacromolecules 2014, 15, 4111−4121

Biomacromolecules

Article

Figure 3. Agarose gel electrophoresis of siRNA loaded cationic nanohydrogel particles: (1) loading capacity of NP1 (left) and NP2 (right) mixed with siRNA at the given weight-to-weight ratios of nanohydrogel particle to siRNA in PBS; (2) heparin competition of siRNA-loaded NP1 (left) and NP2 (right) at the given heparin concentrations in PBS.

Figure 4. Cell viability of HeLa-Luc cells treated with siRNA loaded cationic nanohydrogel particles NP1 (left) and NP2 (right) at the given concentrations (weight-to-weight loading 10:1 nanohydrogel particle to siRNA) after incubation for 48 h (n = 4).

ratio 25:1 of nanogel to siRNA still complexed siRNA robustly at heparin concentrations up to 10 μg/mL in PBS. Only heparin levels of 50 μg/mL and higher were able to release siRNA from both particles completely at that nanogel to siRNA ratio, as demonstrated in Figure 3.2. However, these heparin levels are more than 30 times higher than physiological heparin concentrations found in human blood plasma.36 Thus, the two nanohydrogel particle samples both not only provide similar loading efficiencies but also release capabilities for the siRNA payload. 3. Cell Viability and Knockdown Potential of siRNALoaded Cationic Nanohydrogel Particles. The ability of both nanohydrogel particle samples NP1 and NP2 to deliver and release siRNA was determined in human cervix adenocarcinoma cells stably expressing luficerase (HeLa-Luc). For this experiment, anti-luciferase coding siRNA (luc-siRNA) was loaded into both nanogels in HEPES buffer at a weight-toweight ratio of 10:1 nanohydrogel to siRNA, as described and then applied to HeLa-Luc cells for 48 h. As control samples,

obtained independently form the applied siRNA sequence (lucsiRNA or scr-siRNA) loaded into the nanogel, as shown for NP2 in Figure S17 in the Supporting Information. As previously demonstrated, cationic nanohydrogel particles can provide sufficient stability for the siRNA as polyanionic cargo in high protein-rich solutions like human blood serum as well as the bloodstream of mice.32 Alternatively, a competition mechanism with polyanionic molecules can occur under physiological relevant conditions that promote release of the siRNA cargo and, thus, contribute to the knockdown potential of the carrier after cellular uptake. To that respect, we chose heparin as one of the most negatively charged glycosaminoglycan polysaccharides present in the extracellular matrix of many tissues on the cell surface35 and added it to siRNAcomplexed nanohydrogel particles at increasing concentration. In this case, PBS was chosen to mimic physiological ionic strength during the addition of heparin sodium salt. The release of siRNA was afterward detected by agarose gel electrophoresis as performed earlier. To that respect, we found that both nanohydrogel particles NP1 and NP2 at a weight-to-weight 4116

dx.doi.org/10.1021/bm501148y | Biomacromolecules 2014, 15, 4111−4121

Biomacromolecules

Article

Figure 5. Endogenous luciferase gene silencing in HeLa-Luc cells treated with siRNA loaded cationic nanohydrogel particles NP1 (left) and NP2 (right) at the given concentrations (weight-to-weight loading 10:1 nanohydrogel particle to siRNA) after incubation for 48 h (n = 4, *p < 0.05, and **p < 0.001; Student’s t test for luc-siRNA compared to scr-siRNA or NP only).

cells were incubated for 48 h with both nanogels alone as well as loaded with a scramble siRNA (scr-siRNA), too. Primarily, to exclude nonspecific gene knockdown caused by metabolic disorder, cell viability was tested for each sample prior to luciferase luminescence. The results for both particles are summarized in Figure 4. None of the samples at concentrations up to 400 nM siRNA or 56 mg/L nanohydrogel particle, which were applied to study their knockdown potential, were able to harm the metabolic state of the HeLa cells. In all cases, cell viability was detected around 100% relative to control samples treated with HEPES buffer only. This was also observed not only for both siRNA sequences (luc-siRNA and scr-siRNA) loaded into the nanogels, but also for the empty particles applied to the cells at the given concentrations. Different results, however, were obtained for the luciferase expression levels by measuring the cells’ relative luminescence. The results are summarized in Figure 5. As negative control samples, scr-siRNA loaded nanohydrogel particles or nanohydrogel particles alone did not reduce the expression of luciferase significantly. Thus, carrier related off-target effects can be excluded for the cationic nanohydrogel particles in this study. However, for luc-siRNA loaded nanohydrogel particles significant differences were observed for both particles: While for NP2 no knockdown potential was detected at all for siRNA concentrations up to 400 nM, reduced luciferase expression levels were monitored for NP1 already at 200 nM. The expression levels could be further lowered when cells were treated with 400 nM siRNA. To that respect, siRNA loaded nanohydrogel particle sample NP2 obviously does not provide any knockdown potential, while NP1 seems to promote sufficient delivery of its siRNA payload. To verify this knockdown potential for NP1 over time more precisely, luciferase gene silencing with NP1 was monitored real time for about 80 h (Figure 6). Cells were either incubated with NP1 loaded with luc-siRNA or scr-siRNA as negative control. Already after 12 h a reduced luciferase derived luminescence was detected for the luc-siRNA loaded nanohydrogel particles compared to the control samples. These results turned out to be more significant after 24 h and especially after 48 h, as measured by the knockdown assay before, too. Consequently, this elaborate long-term gene silencing study confirms again the knockdown potential for NP1. Although the values for reduced gene expression are rather moderate, they are nonetheless quite stable and

Figure 6. Long-term luciferase gene silencing in HeLa-Luc cells treated with one single dose of 400 nM siRNA loaded into cationic nanohydrogel particle NP1 at a weight-to-weight loading ratio of 10:1 nanohydrogel particle to siRNA (n = 4, *p < 0.005 at t = 24 h, and **p < 0.001 at t = 48 h; Student’s t test for luc-siRNA compared to scrsiRNA).

continuing over time, as they can last up to 80 h according to the results presented in Figure 6. 4. Cell Uptake and Lysosomal Colocalization of Cationic Nanohydrogel Particles. Based on these promising results, we were interested in some mechanistic insights leading to the exclusive knockdown potential of only the small sized nanohydrogel particle NP1, although their physicochemical properties are nearly identical to NP2, except for their size. Therefore, several cell uptake experiments were performed in human cervix adenocarcinoma cell lines (HeLa). Due to fluorescent labeling of the cationic nanohydrogel particle with Oregon Green, both nanogels NP1 and NP2 could be monitored during their cell uptake. For better visualization of the cell morphology, cell membranes were stained with CellMask Deep Red. Moreover, to get information about the nanoparticles’ intracellular distribution, cells were further stained with LysoTracker, a cationic fluorescent dye that preferentially accumulates in acidic compartments like late endosomes or lysosomes.37 In Figure 7 representative confocal laser scanning images of HeLa cells incubated with either NP1 or NP2 for 24 h were chosen. The fluorescent signal of the nanogel particles is presented in green, while the poststained cell membranes are visualized in blue and the lysosomal compartments are shown in red. Sufficient uptake inside all cells was found for both 4117

dx.doi.org/10.1021/bm501148y | Biomacromolecules 2014, 15, 4111−4121

Biomacromolecules

Article

Figure 7. Incubation of HeLa cells over 24 h with cationic nanohydrogel particles NP1 (left) and NP2 (right) at a particle concentration of 75 μg/ mL. All images are overlays of the fluorescent signals from Oregon Green labeled nanohydrogel particles (green), lysosomal compartments stained with LysoTracker (red), and cell membranes stained with CellMask Deep Red (blue). Areas showing colocalization of nanohydrogel particles with lysosomal compartments appear in yellow; scale bars = 25 μm.

lysosomal compartments (Figure 8, top). Additionally, quantitative colocalization analyses by Pearson’s Correlation Coefficient (PCC) provided again significantly less lysosomal colocalization for NP1 than for NP2, both at 12 and 24 h (compare Supporting Information, Figure S19). To conclude, the time-dependent uptake studies demonstrate that both cationic nanohydrogel particles are taken up by cells similarly, yet, intracellular colocalization studies with lysosomal compartments show that the 100 nm sized nanogel particles NP2 accumulate there already after 4 h, while most of the 40 nm sized nanogel particles NP1 seem to avoid these compartments even after incubation for 24 h. These studies were done with nanogel particles only, but no significant differences of the nanoparticles’ uptake and intracellular distribution performance could be found in further studies when siRNA was conjugated to the nanogels by complexation prior to incubation with cells. As an example, representative confocal laser scanning images of HeLa cells incubated with siRNA loaded nanogel particles NP1 and NP2 for 12 h can be found in the Supporting Information, Figures S20 and S21.

particle systems. Having a closer look at the intracellular distribution, one can easily see that all nanohydrogel particles NP2 mostly colocalize with the fluorescence of the lysosomal compartments, resulting in the yellow color visualized in Figure 7. For the cationic nanogel NP1, however, less colocalization between nanoparticle and lysosome was found. In many cells the fluorescent signal of the nanogel demonstrated in green in Figure 7 can be separated from the red fluorescence derived by the lysosomal compartments. Moreover, quantitative image analyses additionally provided a significantly lower Person’s Correlation Coefficient (PCC) as lysosomal colocalization value for NP1 than for NP2 at this time point (compare Supporting Information, Figure S19). Thus, the 40 nm sized nanohydrogel particles NP1 seem to avoid lysosomal accumulation, while the 100 nm sized cationic nanohydrogel particles NP2 are found there exclusively. To study this intracellular distribution behavior over time, additional kinetic uptake studies were performed. HeLa cells were incubated with the fluorescently labeled nanogels for 0.5, 1, 2, 4, 8, 12, or 24 h and imaged by fluorescence confocal laser scanning microscopy under identical setup conditions. Representative pictures are presented in Figure 8 for NP1 and NP2. The obtained images were analyzed quantitatively, revealing similar uptake efficiencies for both particles that increase over time (compare Supporting Information, Figure S18). Already after 0.5 h some nanoparticles were found inside the cells in both cases. Interestingly, while for NP1 some fluorescence is found in the interior of the cell, most of the nanogel particles NP2 are preferentially located in the cellular periphery. Usually, none of the particles’ green fluorescence is generally colocalized with the red fluorescence of the lysosomal compartments at this early time point. This was also the case for NP2 after 1 and 2 h; however, after 4 h, most of these particles were colocalized with the red fluorescence of the lysosomes, remaining there at further time points (8, 12, and 24 h), too (Figure 8, bottom). For NP1, though, less colocalization could be visualized at these time points inside the cells, since the fluorescent signal of this nanogel colored in green could again often be separated from the red fluorescence of the

■

DISCUSSION In this study, two different types of cationic nanohydrogel particles were generated composed of similar material and synthesized by an identical manufacturing process. They could serve as siRNA carriers for in vitro luciferase gene silencing studies. The nanogels’ physicochemical properties concerning loading with siRNA and their release ability by anionic competition seemed to be similar. Only their average size differed (40 nm vs 100 nm) according to the different molecular weights of the precursor polymers with a similar block ratio. This size difference obviously has significant influence on the knockdown potential for the siRNA carrier. Only the small sized nanogels with average diameters of about 40 nm were able to reduce the luciferase expression levels when loaded with anti-luciferase siRNA, while the 100 nm-sized nanogels did not affect the gene expression of luciferase at all. Cell uptake and lysosomal colocalization studies demonstrated that the siRNA loaded 40 nm-sized-nanohydrogel’s ability to 4118

dx.doi.org/10.1021/bm501148y | Biomacromolecules 2014, 15, 4111−4121

Biomacromolecules

Article

Figure 8. Incubation of HeLa cells with cationic nanohydrogel particles NP1 (top) and NP2 (bottom) (each 75 μg/mL) at given time points. All images are overlays of the fluorescent signals from Oregon Green labeled nanohydrogel particles (green), lysosomal compartments stained with LysoTracker (red) and cell membranes stained with CellMask Deep Red (blue). Areas showing colocalization of nanohydrogel particles with lysosomal compartments appear in yellow; scale bars = 25 μm.

subcellular distribution and transfection efficiency.48−51 Avoiding endosomal pathways that are directed to lysosomal compartments, which would result in oligonucleotide degradation, is generally considered to be the major bottleneck in siRNA delivery for efficient gene silencing. Recent studies by Gilleron et al. demonstrated that for effective lipid-based siRNA carriers only a minor fraction (1−2%) of the siRNA payload was released from the endosome.33 However, Sahay et al. showed that methods to retain siRNA inside endosomal compartments for longer time periods can further enhance their silencing potential effectively.34,52 Increased residence time of siRNA in nonlysosomal compartments may, therefore, contribute to a slow, controlled diffusion of the oligonucleotides from such reservoirs into the cytoplasm, followed by robust gene knockdown.

knockdown luciferase can be correlated with less colocalization with lysosomal compartments over time, whereas the 100 nmsized nanohydrogel particles accumulate in the lysosomes after 4 h incubation. Thus, the nanogel’s size may be an essential key parameter in intracellular distribution and trafficking after cellular uptake, and, what is of more importance, also for the effectiveness of the delivery of a susceptible cargo (e.g., siRNA). So far, previous studies could demonstrate for different nanosized material that their size is an important factor for their cellular uptake and subsequent subcellular trafficking, which mostly affects their biological response in different ways.38−43 Endocytotic pathways, like phagocytosis, marcopinocytosis, and clathrin- and caveolin-dependent or independent endocytosis have been discussed for many nanosized materials in literature.44−47 Especially for siRNA or pDNA carriers in gene delivery, these uptake mechanisms mostly affect their 4119

dx.doi.org/10.1021/bm501148y | Biomacromolecules 2014, 15, 4111−4121

Biomacromolecules

Article

competitive heparin replacement. Interestingly, the nanogels’ different sizes were not affected during the loading process with siRNA. These well-characterized carriers were applied to luciferase gene silencing assays. Interestingly, only the small sized siRNA loaded nanohydrogel particles with diameters of around 40 nm could induce moderate knockdown. Long-term luciferase gene silencing confirmed this knockdown potential lasting even for about 3 days. Cell uptake and colocalization studies with lysosomal compartments revealed that, in contrast to the 100 nm sized cationic nanohydrogel particles, especially the small-sized nanogels were able to avoid cellular endolysosomal uptake pathways, as they were found less accumulating in acidic compartments. In analogy to previously reported studies, we believe that this behavior might also explain their exclusive knockdown potential. Taking these aspects into account, we believe that sizedependent uptake and intracellular distribution mechanism may be essential key parameters for tuning the knockdown potential of siRNA nanocarriers.

To that respect, the 40 nm sized nanohydrogel particles presented in this manuscript might support this hypothesis. In contrast to the 100 nm sized nanohydrogel particles, they have found endocytotic pathways to avoid lysosomal accumulation and thus, contribute to the siRNA related knockdown potential even over longer time periods, as demonstrated especially in Figure 6. The gene silencing values are moderate (luciferase luminescence was reduced to about 50%), probably because of the high stability of the nanohydrogel as siRNA carrier system under physiological conditions, as demonstrated recently.32 To that respect, the covalently cross-linked nanogels may serve as siRNA reservoir, which continuously releases their payload, contributing to their preservative knockdown ability. We hypothesize that the small-sized NP1 siRNA carriers might have access to the 50 nm sized caveolae, in contrast to the larger sized NP2,53 and would preferentially be internalized by caveolin-mediated endocytosis. This is known to mostly avoid vesicle fusion with lysosomes,54 but leaving the cargo longer in endosomal compartments. In addition, it has been shown that the RNA-induced silencing complex (RISC) can also be found associated with the membranes of such endosomal multivesicular bodies.55,56 This process would bring together siRNA and RISC. Alternatively, stimuliresponsive cross-linked nanohydrogel particles may enhance the silencing characteristics of these carrier systems. They would allow more siRNA payload to get access to the RNAi machinery by exposing membrane disruptive elements for enhanced endosomal escape. Besides, this degradability potential may further promote in vivo applicabilities, where biodegradability is mandatory to avoid nanoparticle accumulation. To that respect, we were able to introduce a novel siRNA carrier system based on reductive degradable cationic nanohydrogel particles cross-linked by disulfide-modified spermine derivatives recently, whose siRNA delivery properties are currently under evaluation.57 We are aware that the results found for the 40 and 100 nm sized nanohydrogel particles are so far only preliminary and further detailed investigation, especially focusing on the uptake pathways of the cationic nanohydrogel particles and their subcellular trafficking, will have to be done to understand the mechanism for this behavior more in detail. Still, we believe that the size-dependence found for the presented cationic nanohydrogel particle systems may offer novel opportunities to adjust gene silencing ability and, moreover, contribute to the development of further advanced siRNA carrier systems with improved knockdown potential.

■

ASSOCIATED CONTENT

S Supporting Information *

Analytica data of the applied P(MEO3MA)-b-P(PFPMA) block copolymers and the resulting cationic nanohydrogel particles, agarose gel electrophresis, and dynamic light scattering experiments of the siRNA-loaded nanohydrogels in HEPES buffer, quantitative cellular uptake, and lysosomal colocalization data for each nanohydrogel particle in HeLa cells, confocal laser scanning microscopy images of HeLa cells treated with nanohydrogel particles loaded with and without siRNA after incubation for 12 h. This material is available free of charge via the Internet at http://pubs.acs.org.

■

AUTHOR INFORMATION

Corresponding Author

*E-mail: [email protected]. Fax: +49-6131-3924778. Tel.: +49-6131-3920361. Notes

The authors declare no competing financial interest.

■

ACKNOWLEDGMENTS The authors thank the Fonds der Chemischen Industrie (FCI), the Max Planck Graduate Center with the Johannes GutenbergUniversität Mainz (MPGC) and the DFG Sonderforschungsbereich SFB 1066 for financial support. Moreover, Annette Kelsch (Institute of Organic Chemistry, Johannes GutenbergUniversity Mainz) and Claudia Messerschmidt (Max Planck Institute for Polymer Research, Mainz) are gratefully acknowledged for the electron microscopy imaging.

■

CONCLUSION Nanogel particles can be utilized as safe and stable carriers for effective siRNA delivery. We have recently developed a new concept to synthesize cationic nanohydrogel particles for cellular delivery31 and could demonstrate that these carriers provide sufficient stability under physiologically relevant conditions like human blood serum or the blood circulation of mice.32 In this work, two types of well-defined cationic nanohydrogel particles of different size were synthesized for subsequent gene knockdown studies. According to the manufacturing process using two amphiphilic reactive ester block copolymers of similar composition but different molecular weight, nanogel particles could be generated with diameters of about 40 and 100 nm, respectively. These materials showed similar loading characteristics with siRNA as well as release capabilities by

■

REFERENCES

(1) Fire, A.; Xu, S.; Montgomery, M. K.; Kostas, S. A.; Driver, S. E.; Mello, C. C. Nature 1998, 391, 806−811. (2) Elbashir, S. M.; Harborth, J.; Lendeckel, W.; Yalcin, A.; Weber, K.; Tuschl, T. Nature 2001, 411, 494−498. (3) Dorsett, Y.; Tuschl, T. Nat. Rev. Drug Discovery 2004, 3, 318− 329. (4) Davidson, B. L.; McCray, P. B. Nat. Rev. Genet. 2011, 12, 329− 340. (5) Whitehead, K. A.; Langer, R.; Anderson, D. G. Nat. Rev. Drug Discovery 2009, 8, 129−138. (6) Tseng, Y.-C.; Mozumdar, S.; Huang, L. Adv. Drug Delivery Rev. 2009, 61, 721−731. 4120

dx.doi.org/10.1021/bm501148y | Biomacromolecules 2014, 15, 4111−4121

Biomacromolecules

Article

(7) Akinc, A.; Zumbuehl, A.; Goldberg, M.; Leshchiner, E. S.; Busini, V.; Hossain, N.; Bacallado, S. A.; Nguyen, D. N.; Fuller, J.; Alvarez, R.; Borodovsky, A.; Borland, T.; Constien, R.; de Fougerolles, A.; Dorkin, J. R.; Narayanannair Jayaprakash, K.; Jayaraman, M.; John, M.; Koteliansky, V.; Manoharan, M.; Nechev, L.; Qin, J.; Racie, T.; Raitcheva, D.; Rajeev, K. G.; Sah, D. W.; Soutschek, J.; Toudjarska, I.; Vornlocher, H. P.; Zimmermann, T. S.; Langer, R.; Anderson, D. G. Nat. Biotechnol. 2008, 26, 561−569. (8) 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. Proc. Natl. Acad. Sci. U.S.A. 2010, 107, 1864−1869. (9) Mahon, K. P.; Love, K. T.; Whitehead, K. A.; Qin, J.; Akinc, A.; Leshchiner, E.; Leshchiner, I.; Langer, R.; Anderson, D. G. Bioconjugate Chem. 2010, 21, 1448−1454. (10) Whitehead, K. A.; Sahay, G.; Li, G. Z.; Love, K. T.; Alabi, C. A.; Ma, M.; Zurenko, C.; Querbes, W.; Langer, R. S.; Anderson, D. G. Mol. Ther. 2011, 19, 1688−1694. (11) Zhang, Y.; Pelet, J. M.; Heller, D. a; Dong, Y.; Chen, D.; Gu, Z.; Joseph, B. J.; Wallas, J.; Anderson, D. G. Adv. Mater. 2013, 25, 4641− 4645. (12) Dong, Y.; Love, K. T.; Dorkin, J. R.; Sirirungruang, S.; Zhang, Y.; Chen, D.; Bogorad, R. L.; Yin, H.; Chen, Y.; Vegas, a. J.; Alabi, C. a.; Sahay, G.; Olejnik, K. T.; Wang, W.; Schroeder, A.; Lytton-Jean, a. K. R.; Siegwart, D. J.; Akinc, A.; Barnes, C.; Barros, S. a.; Carioto, M.; Fitzgerald, K.; Hettinger, J.; Kumar, V.; Novobrantseva, T. I.; Qin, J.; Querbes, W.; Koteliansky, V.; Langer, R.; Anderson, D. G. Proc. Natl. Acad. Sci. U.S.A. 2014, 111, 3955−3960. (13) Kanasty, R.; Dorkin, J. R.; Vegas, A.; Anderson, D. G. Nat. Mater. 2013, 12, 967−977. (14) Wagner, E. Acc. Chem. Res. 2012, 45, 1005−1013. (15) Kesharwani, P.; Gajbhiye, V.; Jain, N. K. Biomaterials 2012, 33, 7138−7150. (16) Falamarzian, A.; Xiong, X.-B.; Uludag, H.; Lavasanifar, A. J. Drug Delivery Sci. Technol. 2012, 22, 43−54. (17) Davis, M. E. Mol. Pharmaceutics 2009, 6, 659−668. (18) Davis, M. E.; Zuckerman, J. E.; Choi, C. H.; Seligson, D.; Tolcher, A.; Alabi, C. A.; Yen, Y.; Heidel, J. D.; Ribas, A. Nature 2010, 464, 1067−1070. (19) Zuckerman, J. E.; Choi, C. H. J.; Han, H.; Davis, M. E. Proc. Natl. Acad. Sci. U.S.A. 2012, 109, 3137−3142. (20) Ballarín-González, B.; Howard, K. A. Adv. Drug Delivery Rev. 2012, 64, 1717−1729. (21) Scholz, C.; Wagner, E. J. Controlled Release 2012, 161, 554−565. (22) Christie, R. J.; Miyata, K.; Matsumoto, Y.; Nomoto, T.; Menasco, D.; Lai, T. C.; Pennisi, M.; Osada, K.; Fukushima, S.; Nishiyama, N.; Yamasaki, Y.; Kataoka, K. Biomacromolecules 2011, 12, 3174−3185. (23) Schaffert, D.; Troiber, C.; Salcher, E. E.; Fröhlich, T.; Martin, I.; Badgujar, N.; Dohmen, C.; Edinger, D.; Kläger, R.; Maiwald, G.; Farkasova, K.; Seeber, S.; Jahn-Hofmann, K.; Hadwiger, P.; Wagner, E. Angew. Chem., Int. Ed. 2011, 50, 8986−8989. (24) Christie, R. J.; Matsumoto, Y.; Miyata, K.; Nomoto, T.; Fukushima, S.; Osada, K.; Halnaut, J.; Pittella, F.; Kim, H. J.; Nishiyama, N.; Kataoka, K. ACS Nano 2012, 6, 5174−5189. (25) Fröhlich, T.; Edinger, D.; Kläger, R.; Troiber, C.; Salcher, E.; Badgujar, N.; Martin, I.; Schaffert, D.; Cengizeroglu, A.; Hadwiger, P.; Vornlocher, H.-P.; Wagner, E. J. Controlled Release 2012, 160, 532− 541. (26) Smith, M. H.; Lyon, L. A. Acc. Chem. Res. 2012, 45, 985−993. (27) Ramos, J.; Forcada, J.; Hidalgo-Alvarez, R. Chem. Rev. 2013, 114, 367−428. (28) Naeye, B.; Raemdonck, K.; Remaut, K.; Sproat, B.; Demeester, J.; De Smedt, S. C. Eur. J. Pharm. Sci. 2010, 40, 342−351. (29) Raemdonck, K.; Naeye, B.; Buyens, K.; Vandenbroucke, R. E.; Høgset, A.; Demeester, J.; De Smedt, S. C. Adv. Funct. Mater. 2009, 19, 1406−1415.

(30) Siegwart, D. J.; Whitehead, K. A.; Nuhn, L.; Sahay, G.; Cheng, H.; Jiang, S.; Ma, M.; Lytton-Jean, A.; Vegas, A.; Fenton, P.; Levins, C. G.; Love, K. T.; Lee, H.; Cortez, C.; Collins, S. P.; Li, Y. F.; Jang, J.; Querbes, W.; Zurenko, C.; Novobrantseva, T.; Langer, R.; Anderson, D. G. Proc. Natl. Acad. Sci. U.S.A. 2011, 108, 12996−13001. (31) Nuhn, L.; Hirsch, M.; Krieg, B.; Koynov, K.; Fischer, K.; Schmidt, M.; Helm, M.; Zentel, R. ACS Nano 2012, 6, 2198−2214. (32) Nuhn, L.; Gietzen, S.; Mohr, K.; Fischer, K.; Toh, K.; Miyata, K.; Matsumoto, Y.; Kataoka, K.; Schmidt, M.; Zentel, R. Biomacromolecules 2014, 15, 1526−1533. (33) Gilleron, J.; Querbes, W.; Zeigerer, A.; Borodovsky, A.; Marsico, G.; Schubert, U.; Manygoats, K.; Seifert, S.; Andree, C.; Stöter, M.; Epstein-Barash, H.; Zhang, L.; Koteliansky, V.; Fitzgerald, K.; Fava, E.; Bickle, M.; Kalaidzidis, Y.; Akinc, A.; Maier, M.; Zerial, M. Nat. Biotechnol. 2013, 31, 638−646. (34) Sahay, G.; Querbes, W.; Alabi, C.; Eltoukhy, A.; Sarkar, S.; Zurenko, C.; Karagiannis, E.; Love, K.; Chen, D.; Zoncu, R.; Buganim, Y.; Schroeder, A.; Langer, R.; Anderson, D. G. Nat. Biotechnol. 2013, 31, 653−658. (35) Ruponen, M.; Rönkkö, S.; Honkakoski, P.; Pelkonen, J.; Tammi, M.; Urtti, A. J. Biol. Chem. 2001, 276, 33875−33880. (36) Engelberg, H.; Dudley, A. Circulation 1961, 23, 578−581. (37) Neun, B. W.; Stern, S. T. In Methods Mol. Biol.; Springer: New York, 2009. (38) Rejman, J.; Oberle, V.; Zuhorn, I. S.; Hoekstra, D. Biochem. J. 2004, 377, 159−169. (39) Lai, S. K.; Hida, K.; Man, S. T.; Chen, C.; Machamer, C.; Schroer, T. A.; Hanes, J. Biomaterials 2007, 28, 2876−2884. (40) Jiang, W.; Kim, B. Y.; Rutka, J. T.; Chan, W. C. Nat. Nanotechnol. 2008, 3, 145−150. (41) Cureton, D. K.; Massol, R. H.; Whelan, S. P. J.; Kirchhausen, T. PLoS Pathog. 2010, 6, e1001127. (42) Albanese, A.; Tang, P. S.; Chan, W. C. W. Annu. Rev. Biomed. Eng. 2012, 14, 1−16. (43) Lerch, S.; Dass, M.; Musyanovych, A.; Landfester, K.; Mailänder, V. Eur. J. Pharm. Biopharm. 2013, 84, 265−274. (44) Sahay, G.; Alakhova, D. Y.; Kabanov, A. V. J. Controlled Release 2010, 145, 182−195. (45) Zhao, F.; Zhao, Y.; Liu, Y.; Chang, X.; Chen, C.; Zhao, Y. Small 2011, 7, 182−195. (46) Canton, I.; Battaglia, G. Chem. Soc. Rev. 2012, 41, 2718−2739. (47) Duncan, R.; Richardson, S. C. W. Mol. Pharmaceutics 2012, 9, 2380−2402. (48) Juliano, R. L.; Ming, X.; Nakagawa, O. Bioconjugate Chem. 2012, 23, 147−157. (49) Midoux, P.; Breuzard, G.; Gomez, J. P.; Pichon, C. Curr. Gene Ther. 2008, 8, 335−352. (50) Khalil, I. A.; Kogure, K.; Akita, H.; Harashima, H. Pharmacol. Rev. 2006, 58, 32−45. (51) Medina-Kauwe, L. K.; Xie, J.; Hamm-Alvarez, S. Gene Ther. 2005, 12, 1734−1751. (52) Eltoukhy, A. A.; Sahay, G.; Cunningham, J. M.; Anderson, D. G. ACS Nano 2014, 8, 7905−7913. (53) Mayor, S.; Pagano, R. E. Nat. Rev. Mol. Cell Biol. 2007, 8, 603− 612. (54) Shin, J. S.; Abraham, S. N. Immunology 2001, 102, 2−7. (55) Lee, S. L.; Pressman, S.; Andress, A. O.; Kim, K.; White, J. L.; Cassidy, J. J.; Li, X.; Lim, D. H.; Cho, I. S.; Nakajara, K.; Preall, J. B.; Bellare, P.; Sontheimer, E. J.; Carthew, R. W. Nat. Cell Biol. 2009, 11, 1150−1156. (56) Siomi, H.; Siomi, M. C. Nat. Cell Biol. 2009, 11, 1049−1051. (57) Nuhn, L.; Braun, L.; Overhoff, I.; Kelsch, A.; Schaeffel, D.; Koynov, K.; Zentel, R. Macromol. Rapid Commun. 2014, DOI: 10.1002/marc.201400458.

4121

dx.doi.org/10.1021/bm501148y | Biomacromolecules 2014, 15, 4111−4121