Hexanoic Acid and Polyethylene Glycol Double Grafted Amphiphilic

Feb 5, 2014 - Hydrophilic Substitution Degree. Buddhadev Layek, Manas K. Haldar, Gitanjali Sharma, Lindsey Lipp, Sanku Mallik, and Jagdish Singh*...
0 downloads 0 Views 2MB Size
Download PDF
Article pubs.acs.org/molecularpharmaceutics

Hexanoic Acid and Polyethylene Glycol Double Grafted Amphiphilic Chitosan for Enhanced Gene Delivery: Influence of Hydrophobic and Hydrophilic Substitution Degree Buddhadev Layek, Manas K. Haldar, Gitanjali Sharma, Lindsey Lipp, Sanku Mallik, and Jagdish Singh* Department of Pharmaceutical Sciences, College of Pharmacy, Nursing, and Allied Sciences, North Dakota State University, Fargo, North Dakota 58105, United States S Supporting Information *

ABSTRACT: Gene therapy holds immense potential as a future therapeutic strategy for the treatment of numerous genetic diseases which are incurable to date. Nevertheless, safe and efficient gene delivery remains the most challenging aspects of gene therapy. To overcome this difficulty a series of hexanoic acid (HA) and monomethoxy poly(ethylene glycol) (mPEG) double grafted chitosan-based (HPC) nanomicelles were developed as nonviral gene carrier. HPC polymers with various HA and mPEG substitution degrees were synthesized, and their chemical structures were confirmed by 1H NMR spectroscopy. HPC nanomicelles exhibited excellent blood compatibility and cell viability, as demonstrated by in vitro hemolysis and MTT assay, respectively. The cationic HPC nanomicelles retained the plasmid DNA (pDNA) binding capacity of chitosan and formed stable HPC/pDNA polyplexes with diameters below 200 nm. Both hydrophobic and hydrophilic substitution resulted in suppressed nonspecific protein adsorption on HPC/pDNA polyplexes and increased pDNA dissociation. However, resistance against DNase I degradation was enhanced by HA conjugation while being inhibited by mPEG substitution. Amphiphilic modification resulted in 3−4.5-fold higher cellular uptake in human embryonic kidney 293 cells (HEK 293) mainly through clathrin-mediated pathway. The optimal HPC/pDNA polyplexes displayed 50-fold and 1.2-fold higher gene transfection compared to unmodified chitosan and Fugene, respectively, in HEK 293 cells. Moreover, both the cellular uptake and in vitro transfection study suggested a clear dependence of gene expression on the extent of HA and mPEG substitution. These findings demonstrate that amphiphilic HPC nanomicelles with the proper combination of HA and mPEG substitution could be used as a promising gene carrier for efficient gene therapy. KEYWORDS: chitosan, hexanoic acid, gene delivery, poly(ethylene glycol), transfection

■

INTRODUCTION Gene therapy has evolved as a promising therapeutic modality for the treatment of a wide range of diseases of both innate and acquired origin.1,2 The naked DNA is highly susceptible to nuclease degradation and displays poor cellular uptake, as well as low transfection efficiency.3,4 Therefore, the basic challenge for gene therapy is to design safe and effective carriers that assist efficient transfer of the therapeutic gene to targeted cells without degradation of the delivered gene. Recombinant viruses such as lentivirus, retrovirus, adenovirus, and adeno-associated viruses have been widely studied for this purpose and epitomize the vast majority of clinical trials.5 As of June 2012, there have been over 1843 gene therapy clinical trials worldwide, ∼68% of which have used various viral vectors.6 In spite of some limited success with recombinant viral vectors, there is still no FDA approved gene therapy products. Moreover, use of viral vectors in human has been associated with inflammation, immunogenicity, cancers, and even death in a few cases.7,8 Therefore, viral-based delivery systems need to be reevaluated with respect to their safety issues for human gene therapy. Nonviral © 2014 American Chemical Society

polymeric vectors have drawn increasing attention since their emergence, owing to their distinct advantages which include ease of synthesis, unrestricted gene loading capacity, low immunogenicity against the vector, and greater safety measures.9,10 In recent years, chitosan (CS) and CS derivatives have emerged as potential nonviral gene delivery vectors because of their excellent biocompatibility, biodegradability, negligible cytotoxicity, low immunogenicity, and high cationic charge density.11−13 Under acidic conditions, primary amines of CS remain protonated and can easily form nanoscale complexes (polyplexes) with negatively charged plasmid DNA (pDNA) through ionic interaction.14 These polyplexes have been reported to provide efficient protection of complexed pDNA from enzymatic degradation, facilitate delivery of complexed Received: Revised: Accepted: Published: 982

October 26, 2013 January 11, 2014 February 5, 2014 February 5, 2014 dx.doi.org/10.1021/mp400633r | Mol. Pharmaceutics 2014, 11, 982−994

Molecular Pharmaceutics

Article

Scheme 1. Synthetic Route of Hexanoic Acid and mPEG Double Grafted Amphiphilic Chitosan

surfaces and facilitate endocytosis of polymer/pDNA polyplexes.18 Furthermore, hydrophobic segments in CS assist dissociation of polymer/pDNA polyplexes to promote intracellular release of pDNA which otherwise would be tightly bound through ionic interactions between phosphate groups of pDNA and amino groups of CS.19 These favorable features of hydrophobic modification are reported to induce superior transfection efficiency for CS derivatives over other polymeric carriers using ionic interaction alone. With this purpose, CS has been tailored with numerous hydrophobic moieties such as alkyl,20 acyl,21 deoxycholic acid,22 and cholesterol.23 Meanwhile, the improved water solubility at physiological pH has been pursued by modifying CS with various hydrophilic molecules

pDNA into target cells via adsorption mediated endocytosis, and trigger endosomal escape.15 Despite these favorable physicochemical and biological characteristics, the gene transfection efficiency of unmodified CS is inadequate for clinical applications. Studies have reported that the low transfection efficiency of CS is attributed to its poor aqueous solubility in physiological solution, low cellular uptake, and strong ionic interactions between CS and pDNA, resulting in inefficient release of pDNA from CS/pDNA polyplexes.16,17 To overcome these challenges, various chemical modifications have been carried out on CS through easily modifiable hydroxyl and primary amino groups. Hydrophobic modifications of CS are expected to enhance its adsorption on cell 983

dx.doi.org/10.1021/mp400633r | Mol. Pharmaceutics 2014, 11, 982−994

Molecular Pharmaceutics

Article

Synthesis of mPEG Grafted N-Phthaloyl CS (mPEG-gCSPH). The synthesis of mPEG-g-CSPH was carried out in three steps (Scheme 1): (1) chlorination of CSPH, (2) activation of mPEG with NaH, and (3) grafting of activated mPEG to chlorinated CSPH. To prepare chlorinated CSPH, thionyl chloride (SOCl2, 0.2 mol/mol of CSPH) was added to a solution of CSPH (1.0 g) in pyridine (50 mL), and the mixture was stirred for 30 min at 80 °C under nitrogen. The reaction mixture was cooled to room temperature, precipitated in ice cold water, filtered, and vacuum-dried to obtain chlorinated CSPH. mPEG (80 mg, 0.05 mol/mol of sugar unit of CS) was dissolved in 30 mL of anhydrous THF and NaH (5.2 mg) was added in it, and the mixture was stirring under nitrogen at 60 °C. After 2 h, chlorinated CSPH (0.60 g) was added, and the mixture was stirred at same conditions for another 16 h. The resultant mixture was cooled to room temperature, precipitated in methanol, filtered, washed with dichloromethane (CH2Cl2), and dried under vacuum to get mPEG-g-CSPH.32 Synthesis of mPEG and Hexanoic Acid Double Grafted N-Phthaloyl CS (mPEG-g-CSPH-g-HA). The mPEG-g-CSPH (0.5 g) was dissolved in a mixture of DMF (15 mL) and pyridine (20 mL) and was stirred at 60 °C for 12 h to ensure total solubility. The solution was then cooled to 0− 5 °C, and a solution of hexanoyl chloride (HC) (55 mg, 0.25 mol/mol of sugar unit of CS) in chloroform (CHCl3) was added dropwise over a period of 30 min with constant stirring. The temperature was allowed to rise at room temperature, and the reaction mixture was stirred for another 6 h. Finally, the reaction mixture was dried by vacuum distillation followed by washing with CHCl3 and methanol to acquire mPEG-g-CSPHg-HA.33 Deprotection of N-Phthaloyl CS Derivatives. The mPEG-g-CSPH-g-HA (0.5 g) was dissolved in 25 mL of DMF. Hydrazine monohydrate (10 mL) and water (20 mL) were added to the mPEG-g-CSPH-HA solution, and the mixture was heated at 110 °C under nitrogen with constant stirring. After 15 h of stirring, the reaction mixture was cooled to room temperature, and the suspension was filtered.32 The solvents were evaporated under a reduced pressure. The residue was dissolved in deionized water and dialyzed (MWCO: 6000) against deionized water for 2 days followed by freeze-drying to get mPEG and HA double grafted CS (mPEG-g-CS-HA or HPC1). Similarly, HPC2, HPC3, and HPC4 were obtained when 110 mg (0.50 mol/mol of sugar unit of CS), 165 mg (0.75 mol/mol of sugar unit of CS), and 220 mg (1.0 mol/mol of sugar unit of CS) of HC were used, respectively. While the composition of HPC5 and HPC 6 were 0.75 mol of HC and 0.10 mol of mPEG/mol of sugar unit of CS and 0.75 mol of HC and 0.15 mol of mPEG/sugar unit of CS, respectively. 1 H NMR Spectroscopy. The 1H NMR analysis was performed on a Mercury Varian 400 MHz spectrometer at 25 °C. CS was dissolved in D2O with 1% DCl, and mPEG, HC, and HPC polymers were dissolved in deuterated dimethyl sulfoxide (DMSO-d6). Chemical shifts were recorded in parts per million (ppm) using the signal of TMS as the internal reference. Fourier Transform Infrared Spectroscopy (FTIR). FTIR spectra were recorded on a Thermo Nicolet Nexus 470 FT-IR spectrometer scanning from 4000 to 400 cm−1 at room temperature. CS, mPEG, HC, and HPC polymers were mixed

such as dextran,24 poly(ethylene glycol) (PEG),25,26 poly(vinyl pyrrolidone),27 and poly(β-malic acid).28 Furthermore, hydrophilic modifications of CS also alleviate serum opsonization and enhance intracellular pDNA release from the polyplexes.11 The distinct advantages of hydrophobic and hydrophilic modifications have encouraged the possibility of constructing welldefined amphiphilic CS derivatives with enhanced gene transfection efficiency. The objective of this study was to design, synthesize, and evaluate CS-based gene carriers that offer a solution for the aforementioned barriers in gene delivery. To accomplish our goal, hexanoic acid (HA) and monomethoxy poly(ethylene glycol) (mPEG) were synthesized. HA was selected as hydrophobic part of the system owing to its good safety profile and enhanced gene transfection efficiency as compared to other hydrophobic molecules.29 Similarly, mPEG was chosen as the hydrophilic unit due to its high water solubility, superior cell permeability, nonimmunogenicity, low cytotoxicity, and nonantigenicity.30 The most chemical modifications of CS would result in the reduction of the primary amine groups, which could alter its original physiochemical and biochemical properties, and interfere with the actual analysis of the impact exerted by the modifications alone. In this study, HPC polymers with different degrees of HA and mPEG substitutions were synthesized through chemoselective modification of the hydroxyl groups of CS and subjected them to form HPC/ pDNA polyplexes. The impacts of HA and mPEG substitutions were investigated in terms of critical micelle concentration (CMC), particle size, zeta potential, pDNA complexation and protection, serum adsorption inhibition, in vitro pDNA release rate, cellular uptake, and cell viability. The efficacy of HPC/ pDNA polyplexes was monitored by in vitro gene transfection efficiency in HEK 293 cells.

■

EXPERIMENTAL SECTION

Materials. Low molecular weight CS (MW ∼52 kDa, 90% deacetylated), hexanoyl chloride, monomethoxy poly(ethylene glycol) (MW 750 Da), phthalic anhydride, pyridine, sodium hydride (NaH), hydrazine monohydrate, fluorescein 5isothiocyanate (FITC), ethidium bromide, and 3-(4,5-dimethylthiazol-2-yl)-2,5-diphenyl-tetrazolium bromide (MTT) were purchased from Sigma-Aldrich (St. Louis, MO, USA). DNase I was obtained from Rockland Inc. (Gilbertsville, PA, USA). Plasmid DNA encoding green fluorescence protein (gWiz-GFP) and β-galactosidase (gWiz-βGal) were supplied from Aldevron LLC (Fargo, ND, USA). Human embryonic kidney (HEK 293) cell lines, Dulbecco’s modified Eagle’s medium (DMEM), and phosphate buffered saline (PBS) were purchased from American Type Culture Collection (ATCC, Rockville, MD, USA). Agarose and β-galactosidase enzyme assay kit were purchased from Promega (Madison, WI, USA). FuGENE was obtained from Roche Diagnostics (Indiapolis, IN, USA). All other chemicals were of analytical grade and used without further modification. N-Phthaloylation of CS. CS (3.0 g) was added to a solution of phthalic anhydride (8.0 g) in 50 mL of N,Ndimethylformamide (DMF), and the mixture was heated under nitrogen at 120 °C with constant stirring. After 14 h of reaction, the resultant product was allowed to cool to room temperature and poured into ice cold water.31 The precipitate was filtered, stirred with methanol overnight at room temperature, filtered, and dried under vacuum to get N-phthaloyl CS (CSPH). 984

dx.doi.org/10.1021/mp400633r | Mol. Pharmaceutics 2014, 11, 982−994

Molecular Pharmaceutics

Article

potential from the electrophoretic mobility. Further, the size and morphology of the polyplexes were studied using a DI3100 atomic force microscope (Veeco, Minnesota, USA). The sample (5 μL) of each polyplex was placed on a freshly cleaved untreated mica plate (grade V-4; 15 × 15 × 0.15 mm3) and allowed to dry at room temperature for 30 min. Consequently, the images were recorded in tapping mode using a pyramidal cantilever with a spring constant of 2.8 N/m and at a scan rate of 1 Hz. Gel Retardation Assay. The complex formation between pDNA and HPC polymers was evaluated by agarose gel electrophoresis experiments. To prepare polyplexes (15 μL total volume) at different weight ratios, 1 μg of pDNA was mixed with various concentrations of polymer solution and incubated at room temperature. After 30 min of incubation, 3 μL of gel loading dye (5 × ) was added into each polyplex, and then polyplexes were loaded on a 0.8% (w/v) agarose gel prestained with ethidium bromide (0.5 μg/mL). The electrophoresis was performed at 80 V for 80 min in 0.5 × TAE (TAE, Bio-Rad, CA, USA) buffer, and the resulting pDNA migration patterns were analyzed on a UV transilluminator at 254 nm. DNase I Protection. The resistance of HPC/pDNA polyplexes to pDNA hydrolysis by endonucleases was monitored by DNase I protection assay.28 The HPC/pDNA and CS/pDNA polyplexes containing 10 μg of pDNA at the weight ratio of 20 were incubated with 2.5 units of DNase I in a buffer solution (10 mM Tris-HCl, 2.5 mM MgCl2, and 0.5 mM CaCl2, pH 7.6) at 37 °C. The naked pDNA treated with DNase I was served as a positive control. A quantitative analysis of pDNA degradation was performed by measuring the variation in UV absorbance at 260 nm of each sample at 10 min interval up to 1 h. The absorbance at time zero served as a reference blank. Bovine Serum Albumin Challenge. One hundred microliters of HPC/pDNA and CS/pDNA polyplexes containing 10 μg of pDNA at the weight ratio of 20 were prepared and diluted with 400 μL of PBS (pH 7.4). The polyplexes were then mixed with 500 μL of BSA solution (pH 7.4) and incubated at 37 °C for 1 h. The final concentration of BSA was maintained at 0.2, 0.4, 0.6, 0.8, 1.0, 1.5, and 2.0 mg/mL. The changes in turbidity at 350 nm were monitored with Spectramax UV− visible spectrophotometer.35 In Vitro Release of pDNA. HPC/pDNA and CS/pDNA polyplexes, containing 20 μg of pDNA at a weight ratio of 20, were dispersed in 20 mL of PBS (pH 7.4). The polyplex suspension was incubated at 37 °C in a reciprocal shaking water bath at 50 rpm. At each sampling time, 0.5 mL of suspension was withdrawn and centrifuged at 30 000g, 4 °C for 30 min. The amount of released pDNA in the supernatant was quantified by fluorospectrophotometry. Cell Viability. The cytotoxicity of HPC polymers as well as HPC/pDNA polyplexes was evaluated by MTT assay.16 HEK 293 cells were seeded in a 96-well plate 24 h before the assay at a density of 1 × 104 cells/well in 150 μL of DMEM containing 10% FBS. The next day, different concentrations (100−1000 μg/mL) of polymers or polyplexes at various weight ratios (5, 10, 20, and 30) were added to the cells. After 48 h, media containing polymer or polyplex was removed, and 100 μL of serum-free media and 20 μL of MTT (5 mg/mL) solution were added to each well. Following 3 h of incubation, the MTT containing culture media was aspirated carefully, and the formazan crystals produced by live cells were dissolved in 150 μL of DMSO. Nontreated cells in DMEM containing 10% FBS

separately with KBr powder and pressed into pellets for the study. Critical Micelle Concentration (CMC). The CMC of HPC polymers was determined by fluorescence measurement using pyrene as a molecular probe.34 A series of polymer solutions containing 0.6 μM of pyrene was prepared at various concentrations (5−1000 μg/mL) in 20 mM sodium acetate buffer (pH 6.5). Emission spectra of polymer solutions were recorded from 360 to 450 nm at room temperature using a Fluoromax-4 spectrofluorometer (Horiba Jobin Yvon, NJ, USA). The excitation wavelength was fixed at 336 nm, and both the excitation and emission slit widths were set at 2 nm. The intensity ratios of the first peak (I1, 372 nm) to the third peak (I3, 395 nm) were analyzed for CMC determination. Blood Compatibility Study. The degree of blood compatibility of amphiphilic HPC polymers was estimated by performing in vitro hemolysis assay and morphological analysis of polymer-treated rat erythrocytes as described earlier.19 The erythrocytes were harvested from fresh citrated blood of Sprague−Dawley rats by centrifugation at 1500 rpm for 10 min, and the pellet was washed in PBS until the supernatant was clear. Then the erythrocytes were diluted in PBS to obtain a concentration of 5 × 109 cells/mL. The resultant cell suspension (100 μL) was incubated with 900 μL of the solution of HPC polymers at different concentrations (10−500 μg/mL) for 60 min at 37 °C. Following centrifugation at 1500 rpm for 10 min, the supernatant was analyzed for released hemoglobin at 540 nm using a SpectraMax M5 plate reader. Erythrocytes incubated with PBS and 1% (v/v) Triton X-100 was served as negative (0% hemolysis) and positive control (100% hemolysis), respectively. The percentage of hemolysis was calculated according to the following equation: hemolysis(%) = (A − A 0)/(A100 − A 0) × 100%

where A, A0, and A100 correspond to the absorbance of polymer sample, negative, and positive control, respectively. The morphology of the erythrocytes, incubated with the HPC polymers (500 μg/mL) as described above, was examined on an Olympus DP72 microscope at 40× magnification. Preparation of Polymer/pDNA Polyplexes. CS and HPC stock solutions (0.2−1 mg/mL) were prepared by dissolving the polymer in 20 mM sodium acetate buffer, pH 6.5, and then filtered through 0.2 μm cellulose acetate syringe filters (VWR International, IL, USA). The pDNA stock solution (200 μg/mL) was also prepared in 20 mM sodium acetate buffer, pH 6.5. The polymer/pDNA polyplexes were formulated by adding polymer solution to the pDNA solution at various weight ratios followed by gently vortexing for 10 s. The polyplexes were incubated for 30 min at room temperature before further assessment. Fugene/pDNA complexes were formulated with a pDNA (μg) to Fugene (μL) ratio of 3, according to manufacturer specifications. Particle Size, Zeta Potential, and Morphology. HPC/ pDNA and CS/pDNA polyplexes were prepared at various weight ratios in 20 mM sodium acetate buffer (pH 6.5) with a final pDNA concentration of 10 μg/mL. The average hydrodynamic diameters and zeta potentials of the polyplexes were measured using a Zetasizer Nano-ZS (Malvern Instruments, Malvern, UK) instrument equipped with a nominal 5 mW He−Ne laser operating at 633 nm at 25 °C and at a constant scattering angle of 90°. Before measurements polyplexes were incubated at room temperature for 30 min. The Smoluchowski equation was used to calculate zeta 985

dx.doi.org/10.1021/mp400633r | Mol. Pharmaceutics 2014, 11, 982−994

Molecular Pharmaceutics

Article

polymers to form micellar structure in aqueous environment. Furthermore, HA confers enhance cellular uptake of polymer/ pDNA polyplexes via adsorption-mediated endocytosis and facilitates intracellular release of complexed pDNA, while (3) mPEG offers stability to the polyplexes particularly in the presence of anionic plasma proteins in physiological conditions and also promotes intracellular pDNA release. The synthesis route of the HPC copolymers is shown in Scheme 1. The primary amine groups of CS were first protected with phthalic anhydride. Then, a fraction of 6-CH2OH groups on CS were converted to 6-CH2Cl to react with anionic mPEG to get mPEG-g-CSPH. In the subsequent step, HA was grafted on the remaining 6-CH2OH groups of CS. Finally, the primary amine groups were deprotected by hydrazine treatment. The 1H NMR spectroscopy was utilized to characterize the chemical structure of HPC polymers. In the 1H NMR spectrum of CS (Figure 1),

served as positive controls. The absorbance of each sample was measured at 570 nm using a microplate reader. The relative cell viability (%) was calculated using the following equation. cell viability(%) = (optical density of sample /optical density of control) × 100

Cellular Uptake of Polymer/pDNA Polyplexes. To evaluate cell uptake, HPC polymers and CS were covalently labeled with fluorescein 5-isothiocyanate (FITC) and were allowed to form stable polyplexes with pDNA at the weight ratio of 20. HEK 293 cells were seeded in 6-well plates at 2 × 105 cells/well 24 h prior to the assay. Then, the cells were incubated with the FITC-labeled polyplexes containing 4 μg of pDNA/well. Following 4 h of incubation at 37 °C, the uptake process was ceased by discarding polyplex containing culture media and rinsing the cells three times with PBS. After trypsinization, the percentage of FITC-positive cells was measured by flow cytometry analysis. In an effort to elucidate the mechanisms associated in cellular uptake, HEK 293 cells were treated with optimized concentrations of different well-known endocytosis inhibitors such as sodium azide (10 mM) to inhibit all energy-dependent endocytosis, chlorpromazine (10 μg/mL) to block clathrinmediated endocytosis, colchicine (100 μg/mL) to prevent caveolae formation, or amiloride (50 μg/mL) to inhibit macropinocytosis for 30 min at 37 °C before application of HPC/pDNA polyplexes.28,36 Following 4 h of uptake process, the percentage of FITC-positive cells was quantified by flow cytometry analysis. In Vitro Gene Transfection. The efficacy of HPC polymers as a nonviral gene delivery vector was evaluated by in vitro gene transfection assay. HEK 293 cells were seeded in 24-well plates at 1 × 105 cells/well in 0.5 mL of DMEM containing 10% FBS 24 h prior to transfection experiments. HPC/pDNA polyplexes (50 μL) prepared at different weight ratios equivalent to 1 μg of pDNA (gWiz-GFP or gWiz-βGal) were diluted with 450 μL of fresh culture media (DMEM with 10%FBS) and added to each well. After 4 h of incubation, the transfecting media was exchanged with fresh culture media and the cells were further incubated for 48 h. Naked pDNA and Fugene/pDNA complex were used as negative and positive control, respectively. Fluorescence activated cell sorting (FACS) analysis was performed to determine the percentage of GFP transfected cells. Transfected cells were also visualized and photographed via FV300 confocal laser scanning microscope (Olympus, NY, USA). To quantify the β-galactosidase expression, the culture media was removed, and the cells were washed with PBS. Following thorough lysis of the cells with 1× reporter lysis buffer, the β-galactosidase activity was quantified using a β-galactosidase assay kit (Promega, Madison, WI, USA) at 450 nm. The β-galactosidase activities were normalized against the protein concentration in the cell lysates. The protein concentration was determined using a micro BCA assay.

Figure 1. 1H NMR spectra of chitosan (CS) in D2O, mPEG, hexanoyl chloride (HC), and HPC1 in DMSO-d6.

the peak at 1.85 ppm was ascribed to the acetyl protons of Nacetyl glucosamine, and the peak at 2.9 ppm was attributed to H-2 proton of N-acetyl glucosamine or glucosamine residue. The multiple peaks between 3.4 and 4.0 ppm were assigned to the ring protons (H-3, 4, 5, 6, 6′) of CS. In comparison, HPC1 displayed the peak at 0.85 ppm, which was assigned to the methyl protons of HA. The peaks at 1.25, 1.5, and 2.2 ppm were assigned to the methylene protons of HA. The peak at 3.5 ppm was attributed to the methylene protons of mPEG. Thus, the 1H NMR spectrum confirmed the successful synthesis of HPC copolymers. The degree of substitution of (DS) HA in HPC was defined as the number of HA groups per 100 sugar units of CS and was calculated by comparing the ratio of methyl protons of HA (δ = 0.85) to H-2 proton of N-acetyl glucosamine or glucosamine residue of CS (δ = 2.9 ppm). The degree of mPEG substitution was determined by comparing the ratio of methylene protons of mPEG (δ = 3.5) to H-2 proton of CS (δ = 2.9 ppm). The results are presented in Table 1.

■

RESULTS AND DISCUSSION Chitosan (CS) has chemically active primary amino and hydroxyl groups that can be modified into desirable derivatives. In this report HA and mPEG double functionalized HPC polymers were designed to serve the following distinct purposes: (1) CS is used as polymeric backbone and provides cationic primary amine groups for electrostatic interaction with pDNA, and (2) HA grafting imparts amphiphilicity to the HCP 986

dx.doi.org/10.1021/mp400633r | Mol. Pharmaceutics 2014, 11, 982−994

Molecular Pharmaceutics

Article

CMC Measurement. The CMC is an important physicochemical parameter for amphiphilic polymers, indicating the micelle formation capability. The synthesized HPC polymers possessed self-organizing characteristics to form micellar structures in aqueous environment. The aggregation behavior of HPC polymers was examined by fluorometry with pyrene as a molecular probe. Figure S2 shows the alteration of emission intensity ratio for I1/I3 against the logarithmic of HPC concentration. With polymer concentration below the CMC, the I1/I3 ratio remained almost constant. As the HPC concentration increased to begin micelle formation, the incorporation of pyrene molecules into the micelles contributed to the increase of fluorescence intensity. The intensity of I3 of pyrene amplified at a significantly higher rate than that of the I1. Consequently, the value of the I1/I3 ratio decreased sharply, suggesting micelle formation. The CMC of HPC polymers as a function of HA and mPEG substitutions were shown in Table 1. It indicated that HPC with a higher degree of HA substitution possessed a lower CMC value, while the CMC of HPC with an equal degree of HA substitution, but a different PEGylation degree (3.9%, 7.6%, and 10.5%), had no significant (p > 0.05) difference. Blood Compatibility Study. Blood compatibility is one of the major criteria for the biological applications of cationic delivery systems.37 The nonspecific interactions of cationic polymers with blood components may results in substantial decrease in half-life, targeting ability of the polyplexes, and reproducibility of the therapeutics.38 The hemocompatibility of HPC polymers have been investigated by spectrophotometric measurement of released hemoglobin from erythrocytes after treatment with various concentrations of polymers. The hemolytic activity of PBS and 1% (v/v) Triton X-100 treated erythrocytes were set as 0% and 100%, respectively. Under the experimental conditions, HPC polymers exhibited no hemolytic effect, suggesting no detectable disruption of the erythrocyte membrane (Figure 3). The hemolysis rate of HPC polymers was less than 2%, a rate suggestive of good hemocompatibility. In general, formulation with an in vitro hemolysis rate less than 10% is considered to be nonhemolytic.39 The membrane-lytic potential of HPC polymers was further investigated by performing a detailed morphological analysis of the treated erythrocytes under light microscopy. As shown in Figure 4, no obvious erythrocyte aggregation was evident after

Table 1. Degree of Substitution (DS) and CMC of HPC Copolymers sample

DS (HA)

DS (mPEG)

CMC (μg/mL)

HPC1 HPC2 HPC3 HPC4 HPC5 HPC6

19.2 36.5 54.5 70.6 54.2 54.1

3.9 3.9 3.9 3.9 7.6 10.5

50 40 30 20 30 30

The chemical structures of HCP polymers were also confirmed by FTIR spectroscopy (Figure 2). The grafting of

Figure 2. FTIR spectra of CS, mPEG, HC, and HPC1.

mPEG to CS results in strong absorption peak at about 1075 cm−1 due to the increase of C−O−C bond in the FTIR spectrum of HPC1. The presence of sharp ester bands at 1726 cm−1 in the FTIR spectrum of HPC1 indicates the successful grafting of hexanoyl groups to CS. The peak at the 2912 cm−1 ascribes to the presence of (CH2) groups of CS and mPEG.

Figure 3. Hemolytic activity of HPC polymers at different concentrations. Hemolytic activity is normalized relative to a positive control 1% (v/v) Triton X-100 (set to 100%) and negative control PBS (set to 0%). Indicated values are mean ± SD (n = 4). (B) Photo images of Eppendorf tubes containing supernatant from rat erythrocytes exposed to different concentrations of HPC polymers. 987

dx.doi.org/10.1021/mp400633r | Mol. Pharmaceutics 2014, 11, 982−994

Molecular Pharmaceutics

Article

uniform HPC/pDNA polyplexes were formed at pH 6.5, which had particle diameters less than 200 nm with a polydispersity index of 0.08−0.23. The size of the polyplexes tended to decrease with the increasing weight ratio, suggesting the formation of condensed particles due to the presence of higher density of available protonated amines surrounding the polyplexes. The morphology of HPC/pDNA polyplexes was observed by AFM. As shown in Figure 6, the polyplexes were spherical in shape, and their sizes were ∼150 nm, which were in accordance with the results obtained by the DLS study. DNA Binding Ability. The pDNA binding ability of HPC polymers with various degrees of HA and mPEG substitution was monitored by agarose gel retardation assay using naked pDNA as a positive control. Figure 7 shows the electrophoretic mobility of pDNA in the presence of various amounts of HPC. The existence of partially dissociated pDNA bands in lane “b” indicates the formation of physically unstable complexes at the weight ratio of 0.5. Once the weight ratio increased to ≥5, the migration of pDNA was completely retarded for all polymer/ pDNA polyplexes investigated. These results clearly suggest that the degrees of HA and mPEG substitution did not affect the pDNA condensing capacity of HPC micelles. It is worthy to mention that we selectively conjugated HA and mPEG moieties to the −OH groups of C-6 position on CS via the ester bond. Consequently, the −NH2 groups on CS remained free, and the pDNA binding capacity at acidic pH was preserved. The results of gel retardation assay are in complete agreement with the zeta potential measurement data. Protection of pDNA against Nucleases. The structural integrity of the therapeutic gene is a prerequisite to ensure its desired function in vitro as well as in vivo.41 Thus, an efficient gene carrier must confer adequate protection to pDNA against nuclease degradation. The pDNA protection ability of HPC polymers was evaluated using DNase I as a model enzyme. The naked pDNA was susceptible to DNase I degradation as evident by rapid increase in absorbance at 260 nm. As demonstrated in Figure 8, the degradation rate of pDNA was reduced significantly when it was complexed with CS and HPC polymers at a weight ratio of 20. HPC-6 and HPC-1 displayed slightly lower pDNA protection ability than CS, while HPC5 and HPC2 provide similar extent of pDNA protection as that of CS. Interestingly, HPC3 and HPC4 polyplexes demonstrated slightly higher pDNA protection capacity than other polyplexes indicating higher degrees of HA substitution was favorable for DNase I protection. The higher pDNA protection observed in

Figure 4. Light microscopic images of rat erythrocytes exposed to HPC polymers (500 μg/mL), PBS (pH 7.4), and 1% (v/v) Triton X100. Images were captured at 40× magnification using Olympus DP72 microscope.

PBS or HPC treatment. Moreover, the erythrocytes in HPC polymer solution appeared as biconcave disc shaped normal cells with smooth surfaces, comparable with those in PBS. On the contrary, erythrocytes in 1% (v/v) Triton X-100 demonstrated complete disruption of the cell membrane. The hemolysis assay clearly shows that amphiphilic HPC polymers impose no adverse effects on erythrocyte membrane integrity as they exhibit negligible hemolytic activity equivalent to those of negative control. Characterization of HPC/pDNA Polyplexes. The particle size and surface charge of polyplexes control their cellular uptake.40 A positively charged polyplex could effectively interact with the anionic proteogylans of the cell membrane and thereby facilitate the adsorption mediated endocytosis of the polyplex. In general, polyplexes less than 200 nm in size are favorable for cellular uptake through endocytosis.40 The zeta potentials and mean hydrodynamic diameters of HPC/pDNA polyplexes at different weight ratios were determined by dynamic light scattering measurement. At the weight ratio of 1, HPC/pDNA polyplexes had negative potentials, indicating that the amount of HPC polymer was inadequate to completely condense pDNA. As the weight ratio increased, the zeta potentials of the polyplexes changed from negative to positive, and their surface potentials reached a plateau at or above weight ratio of 15 (Figure 5A). As depicted in Figure 5B, stable and

Figure 5. (A) Zeta potential and (B) average particle diameter measurements using dynamic light scattering as a function of the polymer/pDNA weight ratio. Polyplexes were prepared in 20 mM sodium acetate buffer at pH 6.5. Results are expressed as the mean ± SD (n = 6). 988

dx.doi.org/10.1021/mp400633r | Mol. Pharmaceutics 2014, 11, 982−994

Molecular Pharmaceutics

Article

Figure 6. Atomic force microscopy images of (A) HPC5 micelles and (B) HPC5/pDNA polyplexes (w/w = 20). Images were captured in tapping contact mode and at a scan rate of 1 Hz.

Figure 7. Analysis of HPC/pDNA complex formation at different weight ratios by agarose gel electrophoresis using 0.8% (w/v) agarose gel in 0.5× Tris-acetate-EDTA running buffer for 80 min.

Bovine Serum Albumin Challenge. The in vivo applications of cationic gene delivery vectors are restricted by their nonspecific interactions with anionic serum proteins. These proteins would adsorb on the surface of polyplexes inducing aggregation, initiate rapid clearance, and impede endocytosis.42 Thus, the stability of HPC/pDNA polyplexes in the presence of BSA was studied as a simple model. As shown in Figure 9, HPC/pDNA polyplexes were very stable up to a BSA concentration of 2 mg/mL, whereas CS/pDNA polyplexes showed significant turbidity at 350 nm. The superiority of HPC4/pDNA polyplexes over HPC1/pDNA, HPC2/pDNA, and HPC3/pDNA polyplexes indicated that higher degrees of HA substitution were favorable for stabilizing the polyplexes against the BSA challenge. These results can be explained by the greater extent of resistance offered by the increased hydrophobicity of the polyplexes at higher degrees of HA substitution which lower the nonspecific interaction between the polyplexes and anionic BSA. Compared to HPC3/pDNA polyplexes, the lower increase in absorbance at 350 nm for HPC6/pDNA polyplexes after BSA treatment suggested that higher mPEG substitution degrees provided superior stability from BSA induced aggregation owing to the formation of

Figure 8. DNase I protection assay measured by the variation in OD260 nm of HPC/pDNA polyplexes (w/w = 20) after DNase I addition for 1 h. Results are expressed as the mean ± SD (n = 4).

HPC3 and HPC4 could be attributed to the greater shielding of enzymatic degradation sites of pDNA by stronger hydrophobic interactions and reduced permeability to DNase I.20 989

dx.doi.org/10.1021/mp400633r | Mol. Pharmaceutics 2014, 11, 982−994

Molecular Pharmaceutics

Article

pDNA, and HPC6/pDNA polyplexes were increased to 18.2%, 22.8%, 25.0%, 27.5%, 31.9%, and 39.2%, respectively. The incorporation of hexanoyl side chain to CS backbone resulted in hydrophobicity-induced weakening of electrostatic interactions between HCP polymer and pDNA leading to accelerated pDNA release rate.20 Similarly, the grafting of mPEG chains on CS weakens the pDNA binding strength and makes polyplexes more kinetically labile than the ones formed with unmodified CS.45 In addition, a linear relationship of higher degrees of HA and mPEG substitution with enhanced pDNA release rate was observed. Cell Viability. The cytotoxicity of cationic polymers is considered as one of the limiting factor for their in vivo applications. Hence, in addition to blood compatibility, in vitro cytotoxicity of cationic polymers was evaluated by MTT assay. As depicted in Figure 11A, HPC polymers displayed no diminution in cell viability up to a concentration of 1 mg/mL, which was ∼25-fold higher than the maximum concentration applied in transfection experiments. Moreover, the degrees of HA and mPEG substitution exerted no significant impact on the cell viability of HPC polymers. The cytotoxicity of the HPC/pDNA polyplexes with different weight ratios varied from 5 to 30 was further assessed in HEK 293 cells by MTT assay. As shown in Figure 11B, HPC/pDNA polyplexes at different weight ratios did not alter the cell viability compared to the control. The cell viability results show great promise for the prospects of HPC polymers as gene delivery vectors in clinical applications. Cell Uptake. The cell internalization ability of HPC/pDNA polyplexes into HEK 293 cells was quantitatively determined by FACS analysis. To adequately track the cell uptake process, HPC polymers were chemically conjugated with FITC. As depicted in Figure 12A, CS/pDNA polyplexes mediated 20.7% of FITC positive cells, whereas the uptake percentages were improved by 3−4.5-fold for HPC/pDNA polyplexes. Increasing HA substitution from 19.2% to 54.5% enhanced the cell uptake from 61.2% to 92.5%, while a further increase in HA substitution to 70.6% resulted in the slight reduction in the uptake process. Since HPC polymers are comprised of cationic amine groups and hydrophobic HA moieties in their structure, both charge attraction and hydrophobic interactions facilitate the adsorption mediated endocytosis process. In addition, owing to the hydrophobicity of cell membranes, improved hydrophobicity of the polyplexes to a certain extent might be favorable for endocytosis. On the other hand, increase of mPEG substitution from 3.9% to 7.6% only slightly reduced the cellular internalization. However, a further increase in mPEG substitution (10.5%) seemed to disfavor uptake process, which could be attributable to the steric shielding of hydrophobic interactions between HA chains of polymer and cell membrane. Thus, the design of CS-based polymeric vector with proper degree of hydrophobic and hydrophilic substitutions is essential for enhanced cellular uptake of polyplexes. Endocytosis has been established as the principal mechanism involved in cellular uptake of nonviral gene delivery systems. Four pharmacologically distinct endocytic pathways have been recognized such as clathrin-mediated endocytosis, caveolaemediated endocytosis, macropinocytosis, and phagocytosis. Phagocytosis is typically restricted to specialized mammalian cells such as macrophages, neutrophils, monocytes, and dendritic cells, while all other pathways take place in all cells. To elucidate cell uptake pathways, HPC5/pDNA polyplexes at a weight ratio of 20 were chosen. As depicted in Figure 12B,

Figure 9. Colloidal stability of HPC/pDNA and CS/pDNA polyplexes in BSA. The alteration in turbidity was measured at 350 nm after incubation with increasing concentrations of BSA for 1 h at 37 °C. The polyplexes were prepared at the weight ratio of 20 with a final pDNA concentration of 100 μg/mL. Results are expressed as the mean ± SD (n = 4).

hydrophilic sheath on the surface of the polyplexes at physiological pH. In Vitro Release of pDNA. Once the polyplexes reach the cytoplasm, very strong interactions between pDNA and its vectors imposes difficulty in dissociation of the condensed pDNA and thereby impede gene transfection.43 Thus, the release profile of condensed pDNA plays a critical role to modulate the overall gene expression. Incorporation of hydrophilic or hydrophobic units to polymeric backbone has emerged as an efficient strategy to diminish polymer/pDNA binding strength.44 In vitro pDNA release profiles of HPC/ pDNA polyplexes at a weight ratio of 20 in pH 7.4 PBS are shown in Figure 10. The HPC/pDNA weight ratio of 20 was

Figure 10. Cumulative pDNA release profiles of HCP/pDNA and CS/pDNA polyplexes prepared at a weight ratio of 20 after exposing to PBS (pH 7.4) at 37 °C. Results are expressed as the mean ± SD (n = 4).

selected based on the pDNA binding, condensing (i.e., particle size), and protecting capability of the polyplexes at different weight ratios. After 48 h of incubation, compared to CS/pDNA polyplexes from which only 9.1% of complexed pDNA was released, the cumulative release percentages from HPC1/ pDNA, HPC2/pDNA, HPC3/pDNA, HPC4/pDNA, HPC5/ 990

dx.doi.org/10.1021/mp400633r | Mol. Pharmaceutics 2014, 11, 982−994

Molecular Pharmaceutics

Article

Figure 11. In vitro cytotoxicity of (A) HPC polymers and (B) HPC/pDNA polyplexes in HEK 293 cells. Indicated values are mean ± SD (n = 4).

Figure 12. (A) Cellular uptake of HCP/pDNA and CS/pDNA polyplexes in HEK 293 cells following incubation at 37 °C for 4 h. [*Significantly (p < 0.05) higher than CS]. (B) The effect of chemical inhibitors on the uptake of HCP5/pDNA polyplexes. All of the polyplexes were prepared at the weight ratio of 20. Results are expressed as mean ± SD (n = 4). [*Significantly (p < 0.05) lower than control].

Figure 13. In vitro transfection efficiency of CS and HCP polyplexes in HEK 293 cells at the weight ratio of 20. (A) Transfection at the cellular level using gWiz-GFP plasmid as measured by flow cytometry. (B) Transfection at protein level using gWiz-βGal plasmid as quantified by the βgalactosidase assay. Results are expressed as the mean ± SD (n = 4). [*Significantly (p < 0.05) higher than CS; †Significantly (p < 0.05) higher than Fugene].

inhibitor of the Na+/H+ exchange pump, demonstrated the irrelevance of macropinocytosis-mediated endocytosis in polyplex uptake. Similarly, the insignificant inhibition induced by colchicine ruled out the association of a caveolin-mediated endocytosis in cell internalization. The cellular uptake mechanism analysis was well-accorded with the size-dependent internalization theory, which stated that particles smaller than 200 nm are primarily taken up by the clathrin-mediated

pretreatment with sodium azide yielded striking depression in uptake by ∼82%. These data signified that an ATP/energydependent mechanism contributed to 82% of cellular internalization, while the remaining 15% might be originated from diffusion or physical adsorption. Significant uptake inhibition (61.4%, p < 0.001) exerted by chlorpromazine indicated the role of clathrin-mediated endocytosis in polyplex internalization. The negligible inhibition by amiloride, a selective 991

dx.doi.org/10.1021/mp400633r | Mol. Pharmaceutics 2014, 11, 982−994

Molecular Pharmaceutics

Article

Figure 14. Confocal microscopic images of (A) naked gWiz-GFP, (B) CS/gWiz-GFP, (C) Fugene/gWiz-GFP, (D) HPC1/gWiz-GFP, (E) HPC2/ gWiz-GFP, (F) HPC3/gWiz-GFP, (G) HPC4/gWiz-GFP, (H) HPC5/gWiz-GFP, and (I) HPC6/gWiz-GFP transfected HEK 293 cells after 48 h of transfection. Images were taken at 20× magnification.

pathway and internalization by this pathway was proven to be suitable for efficient gene transfection.46 In Vitro Transfection at Cell Level and Protein Level. To explore in vitro gene transfection capability of HPC polymers, transfection experiments were performed on HEK 293 cells using pGFP and pβ-Gal as reporter genes. HEK 293 cells have been extensively used as a standard model to evaluate gene transfection efficiency of nonviral vectors. To determine the optimal weight ratio of HPC/pDNA polyplexes for transfection, weight ratios ranging from 5 to 30 were investigated for initial screening, and the optimized weight ratio of 20 was used for all transfection studies (data not shown). After 48 h of transfection, the percentages of GFP transfected cells were quantitatively determined by FACS analysis. As illustrated in Figure 13A, HPC polymers effectively expressed GFP, whereas CS/pGFP showed only 7.4% of GFP positive cells even at the optimized weight ratio. Moreover, a strong correlation between degree of HA and mPEG substitution and transfection efficiency was observed. The gene transfection efficiency of HPC polymers increased significantly with the increasing HA substitution, attained the peak value at ∼54.5%, and then decreased at higher substitution of 70.6%. However, the gene transfection efficiency of HPC/ pGFP polyplexes only slightly increased when the degree of mPEG substitution increased from 3.6% to 7.6% followed by a sharp decrease of transfection efficiency at 10.5% of mPEG substitution. HPC5/pGFP polyplexes at the weight ratio of 20

were found to mediate the highest level of gene expression with 72.2% of GFP positive cells detected, which was 10-fold higher than CS/pGFP polyplexes and 1.2-fold higher as compared to positive control Fugene/pGFP complex. Gene transfection efficiency of HPC polymers at the protein level was evaluated using pβ-Gal, a plasmid encoding βgalactosidase protein. The amount of β-galactosidase activity was quantified spectrophotometrically with the help of β-gal assay kit, and data were expressed as the milliunit (mU) of βgalactosidase/mg of total cellular protein. As depicted in Figure 13B, HPC polymers exhibited a 27.5−50.4-fold improvement in gene transfection potency compared to unmodified CS. Moreover, the variation of β-galactosidase activity among different HPC/pβ-Gal polyplexes was in complete agreement with the results of GFP expression data. The transfection efficiency of HPC/pGFP polyplexes was also visualized by confocal laser scanning microscopy. As depicted in Figure 14, no GFP transfected cells were observed when the transfection was performed by naked pGFP. Only a few GFP positive cells were detected when CS was used as a gene carrier. However, highly intense GFP fluorescence was detected when the transfections were carried out by HPC polymers and Fugene. Furthermore, the fluorescence intensity of HPC5/pGFP appeared to be greater than that of all other formulations, including Fugene. Thus, the outcomes of confocal microscopic examination reinforce our conclusions derived 992

dx.doi.org/10.1021/mp400633r | Mol. Pharmaceutics 2014, 11, 982−994

Molecular Pharmaceutics

Article

from the quantitative assessments of GFP and β-gal expression data. In summary, the superior transfection efficiency of HPC polymers to CS is due to amphiphilic modification, which imparts enhanced stability of the polyplexes in the presence of serum protein, higher cellular internalization, and better unpacking of condensed pDNA. HPC polyplexes with higher degree of HA substitution were slightly superior in offering protection against DNase I degradation assay. Hence, a relatively higher amount of intact pDNA was available for subsequent cellular uptake and gene transfection. The cellular uptake of the HPC/pDNA polyplexes was increased with the higher HA substitution degree, reached peak value at 54.5%, and then slightly decreased at higher substitution of 70.6%. In addition, the increased HA substitution contributed better shielding of polyplexes from anionic serum protein BSA and promoted pDNA release rate from the polyplexes might be due to hydrophobicity-induced weakening of electrostatic interactions between HCP and pDNA. These above-mentioned favorable characteristics of HA grafting are responsible for enhanced transfection efficiency of HPC3/pDNA polyplexes over HPC1/pDNA, HPC2/pDNA, and HPC4/pDNA polyplexes. On the other hand, the higher degree of mPEG substitution provided stronger inhibition toward nonspecific serum proteins induced aggregation and improved pDNA release rate. However, higher mPEG substitution such as 10.5% inhibits cellular uptake of the polyplexes. These properties of mPEG substitution explained the superior transfection efficiency of HCP5/pDNA polyplexes over HCP3/pDNA and HCP6/pDNA polyplexes. Together with the enhanced safety profile due to excellent biocompatibility, HPC5 was proven as an efficient gene carrier which not only assists the cellular uptake of the loaded pDNA but also enhances the expression of transferred pDNA.

Notes

The authors declare no competing financial interest.

■

ACKNOWLEDGMENTS We would like to greatly acknowledge the financial support provided by NSF EPSCoR grant no EPS-0814442.

■

■

CONCLUSIONS In this report, a series of chitosan-based amphiphilic polymers was designed and synthesized as promising nonviral vectors for gene delivery. Our results confirmed that HPC polymers efficiently complexed pDNA into nanoscale cationic polyplexes and protected the pDNA from enzymatic degradation. Furthermore, our results support that pDNA delivery capacity of HPC polymers can be modulated by appropriate degree of hydrophobic and hydrophilic substitution. Using HEK 293 cells as a model cell type, we demonstrated that two HPC polymers (HPC3 and HPC5) induced significantly higher gene expression compared to Fugene. It is worth mentioning that the amphiphilic HPC polymers showed excellent blood compatibility and no cytotoxicity when utilized as transfection reagents. These findings highlight the potential of HPC nanomicelles as novel nonviral vectors for gene delivery.

■

ASSOCIATED CONTENT

S Supporting Information *

GPC analysis and alteration of emission intensity ratio for I1/I3 against the logarithmic of HPC concentration. This material is available free of charge via the Internet at http://pubs.acs.org.

■

REFERENCES

(1) McCormick, F. Cancer Gene Therapy: Fringe or Cutting Edge. Nat. Rev. Cancer 2001, 1, 130−141. (2) Opalinska, J. B.; Gewirtz, A. M. Nucleic-Acid Therapeutics: Basic Principles and Recent Applications. Nat. Rev. Drug Discovery 2002, 1, 503−514. (3) Al-Dosari, M. S.; Gao, X. Nonviral Gene Delivery: Principle, Limitations, and Recent Progress. AAPS J. 2009, 11, 671−681. (4) Kim, T.; Jiang, H.; Jere, D.; Park, I.; Cho, M.; Nah, J.; Choi, Y.; Akaike, T.; Cho, C. Chemical Modification of Chitosan as a Gene Carrier In Vitro and In Vivo. Prog. Polym. Sci. 2007, 32, 726−753. (5) Giacca, M.; Zacchigna, S. Virus-Mediated Gene Delivery for Human Gene Therapy. J. Controlled Release 2012, 161, 377−88. (6) Ginn, S. L.; Alexander, I. E.; Edelstein, M. L.; Abedi, M. R.; Wixon, J. Gene Therapy Clinical Trials Worldwide to 2012 − an Update. J. Gene Med. 2013, 15, 65−77. (7) Hollon, T. Researchers and Regulators Reflect on First Gene Therapy Death. Nat. Med. 2000, 6, 6. (8) Check, E. Gene Therapy Put on Hold as Third Child Develops Cancer. Nature 2005, 433, 561. (9) Jones, C. H.; Chen, C. K.; Ravikrishnan, A.; Rane, S.; Pfeifer, B. A. Overcoming Nonviral Gene Delivery Barriers: Perspective and Future. Mol. Pharmaceutics 2013, DOI: 10.1021/mp400467x. (10) Remaut, K.; Sanders, N. N.; De Geest, B. G.; Braeckmans, K.; Demeester, J.; De Smedt, S. C. Nucleic Acid Delivery: Where Material Sciences and Bio-Sciences Meet. Mater. Sci. Eng. R 2007, 58, 117−161. (11) Mao, S.; Sun, W.; Kissel, T. Chitosan-Based Formulations for Delivery of DNA and siRNA. Adv. Drug Delivery Rev. 2010, 62, 12−27. (12) Saranya, N.; Moorthi, A.; Saravanan, S.; Pandima Devi, M.; Selvamurugan, N. Chitosan and Its Derivatives for Gene Delivery. Int. J. Biol. Macromol. 2011, 48, 234−238. (13) Yan, J.; Du, Y. Z.; Chen, F. Y.; You, J.; Yuan, H.; Hu, F. Q. Effect of Proteins with Different Isoelectric Points on the Gene Transfection Efficiency Mediated by Stearic Acid Grafted Chitosan Oligosaccharide Micelles. Mol. Pharmaceutics 2013, 10, 2568−2577. (14) Chang, K. L.; Higuchi, Y.; Kawakami, S.; Yamashita, F.; Hashida, M. Efficient Gene Transfection by Histidine-Modified Chitosan through Enhancement of Endosomal Escape. Bioconjugate Chem. 2010, 21, 1087−1095. (15) Yang, X.; Yuan, X.; Cai, D.; Wang, S.; Zong, L. Low Molecular Weight Chitosan in DNA Vaccine Delivery via Mucosa. Int. J. Pharm. 2009, 375, 123−132. (16) Lu, B.; Wang, C. F.; Wu, D. Q.; Li, C.; Zhang, X. Z.; Zhuo, R. X. Chitosan Based Oligoamine Polymers: Synthesis, Characterization, and Gene Delivery. J. Controlled Release 2009, 137, 54−62. (17) Chan, P.; Kurisawa, M.; Chung, J. E.; Yang, Y. Y. Synthesis and Characterization of Chitosan-g-Poly (ethylene glycol)-Folate as a NonViral Carrier for Tumor-Targeted Gene Delivery. Biomaterials 2007, 28, 540−549. (18) Chiu, Y. L.; Ho, Y. C.; Chen, Y. M.; Peng, S. F.; Ke, C. J.; Chen, K. J.; Mi, F. L.; Sung, H. W. The Characteristics, Cellular Uptake and Intracellular Trafficking of Nanoparticles Made of HydrophobicallyModified Chitosan. J. Controlled Release 2010, 146, 152−159. (19) Layek, B.; Singh, J. Amino Acid Grafted Chitosan for High Performance Gene Delivery: Comparison of Amino Acid Hydrophobicity on Vector and Polyplex Characteristics. Biomacromolecules 2013, 14, 485−494. (20) Liu, W. G.; Zhang, X.; Sun, S. J.; Sun, G. J.; Yao, K. D.; Liang, D. C.; Guo, G.; Zhang, J. Y. N-Alkylated Chitosan as a Potential Nonviral Vector for Gene Transfection. Bioconjugate Chem. 2003, 14, 782−789.

AUTHOR INFORMATION

Corresponding Author

*Telephone: +1-701-231-7943; fax: +1-701-231-8333; e-mail: [email protected]. 993

dx.doi.org/10.1021/mp400633r | Mol. Pharmaceutics 2014, 11, 982−994

Molecular Pharmaceutics

Article

(21) Hu, F. Q.; Zhao, M. D.; Yuan, H.; You, J.; Du, Y. Z.; Zeng, S. A Novel Chitosan Oligosaccharide-Stearic Acid Micelles for Gene Delivery: Properties and In Vitro Transfection Studies. Int. J. Pharmaceutics 2006, 315, 158−166. (22) Lee, K. Y.; Kwon, I. C.; Jo, W. H.; Jeong, S. Y. Complex Formation Between Plasmid DNA and Self-Aggregates of Deoxycholic Acid-Modified Chitosan. Polymer 2005, 46, 8107−8112. (23) Son, S. H.; Chae, S. Y.; Choi, C. Y.; Kim, M. Y.; Ngugen, V. G.; Jang, M. K.; Nah, J. W.; Kweon, J. K. Preparation of a Hydrophobized Chitosan Oligosaccharide for Application as an Efficient Gene Carrier. Macromol. Res. 2004, 12, 573−580. (24) Park, I. K.; Park, Y. H.; Shin, B. A.; Choi, E. S.; Kim, Y. R.; Akaike, T.; Cho, C. S. Galactosylated Chitosan-Graft-Dextran as Hepatocyte-Targeting DNA Carrier. J. Controlled Release 2000, 9, 97− 108. (25) Jiang, X.; Dai, H.; Leong, K. W.; Goh, S. H.; Mao, H. Q.; Yang, Y. Y. Chitosan-g-PEG/DNA Complexes Deliver Gene to the Rat Liver via Intrabiliary and Intraportal Infusions. J. Gene Med. 2006, 8, 477− 487. (26) Zhang, Y.; Chen, J.; Zhang, Y.; Pan, Y.; Zhao, J.; Ren, L.; Liao, M.; Hu, Z.; Kong, L.; Wang, J. A Novel PEGylation of Chitosan Nanoparticles for Gene Delivery. Biotechnol. Appl. Biochem. 2007, 46, 197−204. (27) Park, I. K.; Ihm, J. E.; Park, Y. H.; Choi, Y. J.; Kim, S. I.; Kim, W. J.; Akaike, T.; Cho, C. S. Galactosylated Chitosan (GC)-GraftPoly(vinyl pyrrolidone) (PVP) as Hepatocyte-Targeting DNA Carrier: Preparation and Physicochemical Characterization of GC-Graft-PVP/ DNA Complex (1). J. Controlled Release 2003, 86, 349−359. (28) Wang, B.; He, C.; Tang, C.; Yin, C. Effects of Hydrophobic and Hydrophilic Modifications on Gene Delivery of Amphiphilic Chitosan Based Nanocarriers. Biomaterials 2011, 32, 4630−4638. (29) Layek, B.; Singh, J. N-hexanoyl, N-octanoyl and N-decanoyl Chitosans: Binding Affinity, Cell Uptake, and Transfection. Carbohydr. Polym. 2012, 89, 403−410. (30) Working, P. K.; Newman, M. S.; Johnson, J.; Cornacoff, J. B. Safety of Poly(ethylene glycol) and Poly(ethylene glycol) Derivatives. In Polyethylene Glycol Chemistry and Biological Applications; Harris, J. M., Zalipsky, S., Eds.; American Chemical Society: Washington, DC, 1997; p 45. (31) Kurita, K.; Ikeda, H.; Yoshida, Y.; Shimojoh, M.’; Harata, M. Chemoselective Protection of Amino Groups of Chitosan by Controlled Phthaloylation: Facile Preparation of a Precursor Useful for Chemical Modifications. Biomacromolecules 2002, 3, 1−4. (32) Malhotra, M.; Lane, C.; Tomaro-Duchesneau, C.; Saha, S.; Prakash, S. A Novel Method for Synthesizing PEGylated Chitosan Nanoparticles: Strategy, Preparation, and In Vitro Analysis. Int. J. Nanomed. 2011, 6, 485−494. (33) Zhang, J.; Chen, X. G.; Peng, W. B.; Liu, C. S. Uptake of OleoylChitosan Nanoparticles by A549 Cells. Nanomedicine 2008, 4, 208− 214. (34) Ye, Y. Q.; Yang, F. L.; Hu, F. Q.; Du, Y. Z.; Yuan, H.; Yu, H. Y. Core-Modified Chitosan-Based Polymeric Micelles for Controlled Release of Doxorubicin. Int. J. Pharmaceutics 2008, 352, 294−301. (35) Hashimoto, M.; Morimoto, M.; Saimoto, H.; Shigemasa, Y.; Sato, T. Lactosylated Chitosan for DNA Delivery into Hepatocytes: The Effect of Lactosylation on the Physicochemical Properties and Intracellular Trafficking of pDNA/Chitosan Complexes. Bioconjugate Chem. 2006, 17, 309−316. (36) Nam, H. Y.; Kwon, S. M.; Chung, H.; Lee, S. Y.; Kwon, S. H.; Jeon, H.; Kim, Y.; Park, J. H.; Kim, J.; Her, S.; Oh, Y. K.; Kwon, I. C.; Kim, K.; Jeong, S. Y. Cellular Uptake Mechanism and Intracellular Fate of Hydrophobically Modified Glycol Chitosan Nanoparticles. J. Controlled Release 2009, 135, 259−267. (37) Yang, J.; Liu, Y.; Wang, H.; Liu, L.; Wang, W.; Wang, C.; Wang, Q.; Liu, W. The Biocompatibility of Fatty Acid Modified DextranAgmatine Bioconjugate Gene Delivery Vector. Biomaterials 2012, 33, 604−613. (38) Kainthan, R. K.; Gnanamani, M.; Ganguli, M.; Ghosh, T.; Brooks, D. E.; Maiti, S.; Kizhakkedathu, J. N. Blood Compatibility of

Water Soluble Hyperbranched Polyglycerol-Based Multivalent Cationic Polymers and Their Interaction with DNA. Biomaterials 2006, 27, 5377−5390. (39) Amin, K.; Dannenfelser, R. M. In vitro hemolysis: guidance for the pharmaceutical scientist. J. Pharm. Sci. 2006, 95, 1173−1176. (40) Zauner, W.; Ogris, M.; Wagner, E. Polylysine-Based Transfection Systems Utilizing Receptor-Mediated Delivery. Adv. Drug Delivery Rev. 1998, 30, 97−113. (41) Katayose, S.; Kataoka, K. Remarkable Increase in Nuclease Resistance of Plasmid DNA through Supramolecular Assembly with Poly (ethylene glycol)-poly (l-lysine) Block Copolymer. J. Pharm. Sci. 1998, 87, 160−163. (42) Mao, H. Q.; Roy, K.; Troung-Le, V. L.; Janes, K. A.; Lin, K. Y.; Wang, Y.; August, J. T.; Leong, K. W. Chitosan-DNA Nanoparticles as Gene Carriers: Synthesis, Characterization and Transfection Efficiency. J. Controlled Release 2001, 70, 399−421. (43) Schaffer, D. V.; Fidelman, N. A.; Dan, N.; Lauffenburger, D. A. Vector Unpacking as a Potential Barrier for Receptor-Mediated Polyplex Gene Delivery. Biotechnol. Bioeng. 2000, 67, 598−606. (44) Pack, D. W.; Hoffman, A. S.; Pun, S.; Stayton, P. S. Design and Development of Polymers for Gene Delivery. Nat. Rev. Drug Discovery 2005, 4, 581−593. (45) Banaszczyk, M. G.; Lollo, C. P.; Kwoh, D. Y.; Phillips, A. T.; Amini, A.; Wu, D. P.; Mullen, P. M.; Coffin, C. C.; Brostoff, S. W.; Carlo, D. J. Poly-L-Lysine-Graft-PEG-Comb-Type Polycation Copolymers for Gene Delivery. J. Macromol. Sci., Pure Appl. Chem. 1999, 36, 1061−1084. (46) Douglas, K. L.; Piccirillo, C. A.; Tabrizian, M. Cell LineDependent Internalization Pathways and Intracellular Trafficking Determine Transfection Efficiency of Nanoparticle Vectors. Eur. J. Pharm. Biopharm. 2008, 68, 676−687.

994

dx.doi.org/10.1021/mp400633r | Mol. Pharmaceutics 2014, 11, 982−994