Enhanced Transfection Efficiency and Reduced Cytotoxicity of Novel

Apr 18, 2013 - The present study reports the development, characterization, and evaluation of novel polyelectrolytes stabilized lipoplexes as a nonvir...
0 downloads 9 Views 532KB Size
Download PDF
Article pubs.acs.org/molecularpharmaceutics

Enhanced Transfection Efficiency and Reduced Cytotoxicity of Novel Lipid−Polymer Hybrid Nanoplexes Sanyog Jain,*,† Sandeep Kumar,†,‡ Ashish Kumar Agrawal,† Kaushik Thanki,† and Uttam Chand Banerjee‡ †

Centre for Pharmaceutical Nanotechnology, Department of Pharmaceutics and ‡Department of Pharmaceutical Technology (Biotechnology), National Institute of Pharmaceutical Education and Research (NIPER), Sector 67, S.A.S. Nagar (Mohali), Punjab-160062, India ABSTRACT: The present study reports the development, characterization, and evaluation of novel polyelectrolytes stabilized lipoplexes as a nonviral vector for gene delivery. In order to achieve the advantage of both DOTAP (1,2-dioleoyl-3-trimethylammonium propane) and PEI (high transfection efficiency) a system was hypothesized in which DOTAP/phosphatidyl choline (PC) lipoplexes were electrostatically coated with anionic poly(acrylic acid) (PAA) and cationic polyethylenimine (PEI) alternatively to finally shape a robust structure PEIPAA-DOTAP/PC-lipoplexes (nanoplexes). The nanoplexes were found to have size of 242.6 ± 9.4 nm and zeta potential of +23.1 ± 1.5 mV. Following development nanoplexes were evaluated for cellular uptake, nuclear colocalization, transfection efficiency, and cellular toxicity in MCF-7, HeLa, and HEK-293 cell lines. In support of our hypothesis nanoplexes exhibited higher uptake and nuclear colocalization in comparison with DOTAP/PC, DOTAP/DOPE lipoplexes, and PEI polyplexes. Nanoplexes also exhibited 50−80, 11−12, 6−7, and 5−6 fold higher transfection efficiency in comparison with DOTAP/PC-lipoplexes, DOTAP/DOPE-lipoplexes, PEI-polyplexes, and lipofectamine, respectively, and significantly lower toxicity in comparison with DOTAP/PC, DOTAP/DOPE lipoplexes, PEI polyplexes, and commercial lipofectamine. KEYWORDS: gene delivery, lipoplexes, nanoplexes, PEI polyplexes, DOTAP, transfection efficiency, cytotoxicity

1. INTRODUCTION Malfunctioning of proteins encoded by mutated genes result in diverse genetic disorders including cystic fibrosis, hemophilia, sickle cell anemia, Tay−Sachs disease, and Niemann−Pick disease.1 Such genetic defects may be corrected by site-specific replacement of disease-causing gene with healthy gene, leading to expression of therapeutic proteins. Although viral vectors including retroviruses, adenoviruses, and adeno-associated viruses are known for their intrinsic ability to deliver genetic cargoes to target cells with high transfection efficiency, their clinical applications are often restricted by severe toxicity that causes even patient death in certain clinical situations.2 Additional limitations pertinent to nonviral vectors include mutational insertion risks, limited DNA loading efficiency, difficulties in attachment of targeting ligand to virus surface, large-scale production, storage, stability issues, and above all, high cost of production.3−5 To prevail over these limitations, nonviral vectors with lower immunogenicity and toxicity have been developed. The most commonly used nonviral vectors include cationic liposomes, cationic polymers, and dendrimers, which electrostatically interact with the negatively charged phosphate backbone of the DNA to form lipoplexes or polyplexes.6−8 In contrast to viral vectors, the journey of nonviral vectors from their site of administration to pathological target is full of © 2013 American Chemical Society

obstacles. At every stage of the journey, the vector is exposed at the risk of destabilization by plasma proteins and degradation by nucleases. Even after reaching the target cells, the delivery vector has to overcome two major membrane barriers: the plasma membrane and, finally, the nuclear envelope. Subsequently, the transfection efficiency of nonviral vectors is usually lower than their viral analogues.9−11 Over the last two decades, cationic liposomes have been recognized as prospective candidates for nonviral gene delivery because of their easy scalability, low immunogenicity, high DNA loading efficiency, and innate ability to ferry the cargo to their appropriate targets. Unfortunately, cationic liposomes too exhibit certain limitations, which include low plasma circulation time due to nonspecific sequestration by macrophages of the mononuclear phagocytic system (MPS), nonspecific interaction with plasma proteins leading to poor transfection efficiency, lack of endolysosomal escape ensuing degradation of DNA, and high cytotoxicity due to its polycationic nature. A number of strategies have been used to overcome the limitations associated with cationic lipoplexes. These include: prolongation of plasma circulation time via PEGylation, masking of excess Received: January 19, 2013 Accepted: April 18, 2013 Published: April 18, 2013 2416

dx.doi.org/10.1021/mp400036w | Mol. Pharmaceutics 2013, 10, 2416−2425

Molecular Pharmaceutics

Article

positive charges by shielding with polysaccharides,12−14 attaining site-specificity by attachment of active targeting ligand(s),15 and improvement of transfection efficiency by incorporation of helper lipids in formulation along with conventional lipids.16 Although these approaches have been reported to improve the transfection efficiency of lipoplexes to some extent, the delivery of loaded DNA into nucleus still remains a major hurdle. In the present work, we hypothesized that the coating of conventional 1,2-dioleoyl-3-trimethylammonium propane (DOTAP)/phosphatidyl choline (PC)-lipoplexes with alternative layer of polyelectrolytes will improve the transfection efficiency by protecting the DNA against deactivation/ degradation against plasma proteins and deoxyribonucleases while circumventing the drawbacks associated with conventional liposomes. In line with that hypothesis, conventional DOTAP/PC-lipoplexes were electrostatically coated with the anionic polyelectrolyte, poly(acrylic acid) (PAA), followed by a second layer of cationic poly(ethylene imine) (PEI). Layer-bylayer deposition of polyelectrolytes resulted in the formation of a robust structure called “Nanoplexes”.17,18 In this study, PEI was selected as a formulation component based on its unique ability to undergo endolysosomal escape via proton sponge effect19,20 and subsequent nuclear entry through formation of nuclear pore complexes (NPC).15,21 The in vitro transfection efficiency and cytotoxicity of the synthesized nanoplexes compared to other lipoplexes, namely, PEI-polyplexes and lipofectamine were evaluated on three cancer cell lines of different origins: human breast adenocarcinoma (MCF-7), cervical carcinoma (HeLa), and embryonic kidney (HEK-293) cell lines. To the best of our knowledge, no previous report has embarked on the feasibility of using polyelectrolytes coated lipoplexes as a potential nonviral vector for gene delivery.

(DH5a), was obtained from Addgene. The pDNA was extracted using maxiprep columns by strictly following the manufacturer protocol. The purity and concentration of pDNA was determined by measuring the absorbance ratio at 260/280 nm using UV spectrophotometer and agarose gel electrophoresis.14 The absorbance ratio (1.87) at 260/280 and agarose gel electrophoresis confirmed the purity of pDNA. Purified pDNA was resuspended in sterile Tris buffer and frozen in aliquots at a concentration of 0.25 mg/mL. 2.3. Formulation of Cationic Lipoplexes and PEI Polyplexes. The cationic liposomes were prepared by the thin film hydration method.17,22,23 Briefly, DOTAP:PC (1:1 molar ratio) was dissolved in chloroform:methanol mixture (9:1 v/v) following solvent removal under reduced pressure in a rotavapor (Buchi, Switzerland) at 45 °C. The resultant thin film, formed on the walls of a round bottomed flask, was subsequently hydrated with water for 2 h and further subjected to probe sonication (Cole Parmar Instrument Company, USA) for 30 s at amplitude 80 (3 cycles with 1 min pulse off) which resulted in formation of liposomes. These liposomes were further incubated with pDNA at room temperature for 30 min to finally form the lipoplexes.24 DOTAP/DOPE lipoplexes were formed following the same procedure except the use of DOPE instead of PC. PEI polyplexes were formed by simple incubation of the PEI solution with pDNA at room temperature for 30 min.12 2.4. Gel Retardation Assay. Complexation efficiency between liposomes and pDNA was evaluated using gel retardation assay. Briefly, pDNA was incubated with increasing concentration of lipids for 30 min, following which the lipoplexes were mixed with loading buffer containing a tracking dye (xylene cyanol) and loaded into individual wells of 0.8% agarose gel and electrophoresed at 100 V for 45 min in TAE buffer (40 mM Tris-HCl, 1% (v/v) acetic acid, 1 mM EDTA). Finally, resolved bands were stained with ethidium bromide and visualized under a UV transilluminator.12,14 2.5. Formulation and Characterization of Polyelectrolyte Coated Lipoplexes (Nanoplexes). Cationic lipoplexes were taken as the template, and the coating of anionic PAA was implemented over this cationic template. Coating optimization was done at different concentrations (0.01−0.07 μg/μL) of PAA by simply mixing the lipoplex dispersion with equal volume of PAA solution by vortexing for 30 s followed by incubation at room temperature for 1 h. The coating was confirmed by charge reversal (from positive to negative) and size of PAA coated lipoplexes. These PAA coated lipoplexes were further taken as a template for coating of oppositely charged PEI in next step. Coating optimization was done at different concentrations (0.01− 0.07 μg/μL) of PEI following the above procedure. The coating confirmation was validated by charge reversal (from negative to positive) and size of nanoplexes. The excess of polyelectrolytes after each coating step were removed by previously reported method using highspeed centrifugation (15 000 rpm for 20 min) and a dialysis bag (70 kD, Sigma Aldrich, St. Louis, MO).17 Following the formation, nanoplexes were characterized for size and zeta potential. The particle size was measured by dynamic light scattering (DLS) (Nano ZS, Malvern, UK) while zeta potential was estimated on the basis of electrophoretic mobility under an electric field using same instrument. The surface morphology of the optimized polyplexes was determined using scanning electron microscopy (SEM) and Atomic force microscopy (AFM). In SEM analysis polyplexes were placed on a glass

2. MATERIALS AND METHODS 2.1. Materials. Branched poly(ethylene imine) MW 25 000 (PEI25000), poly(acrylic acid) MW 5000 (PAA5000), minimum essential medium (MEM), fetal bovine serum (FBS), antibioticantimycotic solution, 3-(4,5-dimethylthiazol-2-yl)-2,5-diphenyltetrazolium bromide (MTT), Tris, bromophenol blue (BPB), ethidium bromide (EtBr), sodium dodecyl sulfate (SDS), Triton X-100, ethylenediaminetetraacetic acid (EDTA), xylene cyanol (XC), rhodamine isothiocynate (RITC), and 4′,6diamidino-2-phenylindole (DAPI) were purchased from SigmaAldrich Chemical Co. (St. Louis, MO, USA). 1,2-Dioleoyloxy3-trimethylammoniumpropanchloride (DOTAP), 1,2-dioleoylsn-glycero-3-phosphoethanolamine (DOPE), and phosphatidyl choline (PC) were obtained as a generous gift sample from lipoid. The plasmid purification kit and lipofectamine were purchased from Qiagen (France). Human breast adenocarcinoma (MCF-7), human cervix carcinoma (HeLa), and HEK293 (human embryonic kidney-293) cell lines were obtained from the cell repository facility of National Centre for Cell Sciences (NCCS), Pune, India. DNase I was purchased from Fermentas Molecular Biology Tools. Tissue culture plates were procured from Nunc, Roskilde, Denmark. Deionized (Milli-Q) and Millipore filtered (pore diameter 0.22 μm) water was used throughout the experiments. All other chemicals and reagents were of analytical grade and procured from local suppliers. 2.2. Preparation and Extraction of Plasmid DNA (pDNA). The plasmid (pEGFP-N3) encoded for enhanced green fluorescence protein, by inserting gene to cytomegalovirus (CMV) and propagated in competent Escherichia coli 2417

dx.doi.org/10.1021/mp400036w | Mol. Pharmaceutics 2013, 10, 2416−2425

Molecular Pharmaceutics

Article

PEI (10 mg/mL) was added to RITC solution, and the reaction mixture was stirred in dark for 6 h to yield RITC-PEI. Following overnight incubation, MCF-7, HeLa, and HEK293 cells were allowed to incubate with RITC incorporated complexes for 4 h. After 4 h of incubation cells were thoroughly washed with PBS to remove the noninternalized complexes. Cells were fixed by adding 4% paraformaldehyde (PFA) in PBS (pH 7.4) for 10 min and thoroughly washed with PBS and subjected to Triton X-100 (0.2% in PBS) for 2 min for permeabilization. Subsequently, cells nuclei were counterstained with DAPI (1 μg/mL in PBS) for 1 min followed by thorough washing with PBS.30 The cells were visualized in CLSM (Olympus FV1000, Japan) at excitation/emission maxima ∼358/461 for DAPI and ∼560/580 maxima for RITC, respectively. Nuclear colocalization was studied using scatter plot analysis. 2.8.3. In Vitro Transfection. MCF-7, HeLa, and HEK-293 cells (1 × 105 cells/well) in six well plates were incubated with different complexes (lipoplexes, polyplexes, and nanoplexes; each containing electrostatically complexed DNA at N/P ratio 10) and commercially available nonviral vectors lipofectamine for 4 h in MEM supplemented with 10% (v/v) fetal bovine serum (FBS). Cells were washed three times with PBS (pH 7.4) and further incubated up to 48 h in fresh MEM supplemented with 10% FBS. After incubation, cells were washed three times with PBS (pH 7.4) and observed under CLSM (Olympus FV1000, Japan), using He−Ne green laser at an excitation wavelength of 488 nm and emission wavelength of 509 nm. Images were captured at 10X magnification.14 To quantify the protein expression level, cells were then washed with 50 μL of PBS and incubated with 100 μL of lysis buffer (10 mM Tris HCl, pH 7.4, 0.5% SDS, 0.5% Triton X100, 1 mM EDTA). The cell lysates were centrifuged to pelletize the cellular debris. The total protein concentration in cell lysate from each well was estimated by Bradford method, while EGFP content in per mg of protein was determined by measuring fluorescent intensity. A 2 μL of cell lysate was loaded on a Nanodrop spectrofluorimeter (NanoDrop 3300, Thermo Fisher Scientific, USA), and EGFP expression was estimated fluorimetrically at excitation and emission wavelengths at 488 and 509 nm, respectively. EGFP expression of nanoplexes was also measured against the blank cells (without pDNA).14 2.8.4. Cytotoxicity Study. Cytotoxicity of formulated lipoplexes was evaluated using MTT assay12 which is based on the principal of reduction of tetrazolium dye, MTT, to its insoluble formazan giving a purple color due to the activity of cellular enzymes. The attached cells in 96 well plates were incubated with 100 μL of developed lipoplexes dispersion for 4 h. Cells were washed three times with PBS (pH 7.4) and further incubated up to 48 h in fresh MEM supplemented with 10% FBS. Subsequently, the medium was aspirated, and 100 μL of MTT (0.5 mg/1.0 mL in PBS)29 was added to each well. After 3 h, the supernatant was aspirated, and the formazan crystals were suspended in 200 μL of DMSO followed by gentle shaking of plates for one minute and absorbance measurement at 540 nm in ELISA plate reader (Biotek Inc., USA). All of the experiments were performed in quadruplet. Untreated cells were taken as control with 100% viability, and relative cell viability (%) was calculated using the following formula:

coverslip, previously adhered to metallic stub by a biadhesive tape. Subsequently, the samples were vacuum coated with gold and visualized under SEM (Hitachi, S-3400N, Japan).25 In AFM analysis a drop of polyplexes dispersion was placed on the silicon wafer and air-dried. Scanning was carried out at nominal force constant at 0.1 N/m with a cantilever of length 325 μm and width 26 μm. Images were obtained by displaying the amplitude and height signals of cantilever in the trace and retrace direction, respectively.26 2.6. DNase Protection Assay. Protecting efficacy of developed complexes was evaluated by DNase I protection assay. Briefly, developed complexes at the desired weight ratio (complexes/DNA = 10) were incubated at 37 °C for half an hour in buffer solution (10 × 10−3 M Tris-Cl, 150 × 10−3 M NaCl, 1 × 10−3 M MgCl2, pH 7.4) containing 10 μL DNase I (1000 units/mL). Enzyme reaction was initiated by adding 50 μL of Mg2+ solution (50 × 10−3 M). The absorbance change at 260 nm was measured by spectrophometer at the interval of 2 min.27 2.7. Serum Stability. In vitro serum stability of the developed complexes was determined by measuring change in physicochemical characteristics and ethidium bromide (EtBr) intercalation assay. 2.7.1. Physicochemical Characterization. Briefly, complexes (1 mL) containing 10 μg of pDNA were incubated at 37 °C for 4 h with an equal volume of minimum essential medium (MEM) supplemented with 10% fetal bovine serum (FBS), following which changes in size and zeta potential were measured.12 2.7.2. EtBr Intercalation Assay. EtBr intercalation assay was performed according to reported method with slight modification.28 Briefly, different complexes (1 mL) containing 10 μg pDNA were incubated at 37 °C up to 4 h with equal volume of MEM supplemented with 10% fetal bovine serum (FBS). Aliquots were withdrawn at different time points (5, 30, 60, 120, and 240 min) and mixed with EtBr solution in water to achieve the concentration (0.2 μg/mL) of EtBr in final mixture. Fluorescent intensity of the resultant mixture was determined at 516 and 618 nm excitation and emission wavelengths, respectively. The fluorescence intensity of nanocarriers was calculated as the fraction of maximum fluorescence, obtained when EtBr was added to free plasmid. 2.8. Cell Uptake, Colocalization, In Vitro Transfection, and Cytotoxicity Studies. 2.8.1. Cell Culture. MCF-7 (Human breast adenocarcinoma), HeLa (human cervix adenocarcinoma), and HEK-293 (human embryonic kidney293) cells were grown in (MEM) supplemented with 10% (v/ v) fetal bovine serum (FBS), 100 units/mL penicillin, and 100 μg/mL streptomycin at 37 °C in a 5% CO2 and 95% air humidified atmosphere. Confluent cultures were harvested by trypsinization and cells counted and suitably diluted to obtain 1 × 105 cells/well followed by overnight incubation for cell attachment.22,29 2.8.2. Cell Uptake and Nuclear Colocalization. Cellular uptake and nuclear colocalization of various complexes was visualized through confocal light scanning microscopy (CLSM). For the purpose RITC encapsulated DOTAP/PC and DOTAP/DOPE lipoplexes were formed by taking RITC (0.5 mg/mL) in hydration medium while RITC incorporated PEI-polyplexes were formed by first synthesizing RITC-PEI conjugate subsequently forming RITC-PEI-polyplexes. RITCPEI was synthesized by dissolving RITC (0.5 mg/mL) in water simultaneously adjusting the pH to 8 and stirring for 10 min.

relative cell viability = 2418

absorbance(sample) × 100 absorbance(control)

dx.doi.org/10.1021/mp400036w | Mol. Pharmaceutics 2013, 10, 2416−2425

Molecular Pharmaceutics

Article

2.9. Statistical Analysis. All results have been demonstrated as mean ± standard deviation (SD). Statistical analysis was performed using Graph Pad Prism 6 using one-way analysis of variance (ANOVA) followed by Tukey-Kramer multiple comparison test. p < 0.05 was considered as statistically significant.

ratios. Smaller size and negative zeta potential at 0.5 and 1 N/P ratios might be the result of insufficient amount of DOTAP (N counterpart of the complex) to achieve complete complexation. A sharp increase in particle size at N/P ratio (2.5) could be attributed to aggregation as a result of charge neutralization. A decrease in particle size and increase in zeta potential from 2.5 to 10 could be ascribed to the formation of compact structure due to strong electrostatic attraction and increased concentration of DOTAP, respectively. 3.2. Gel Retardation Assay. The free DNA moves along the electric field due to presence of negative charge. The complexation of DNA with cationic lipid results in charge neutralization and ultimately loss of electrophoretic mobility. The loss of electrophoretic mobility clearly indicates the efficiency of complexation of negatively charged DNA with cationic lipid. The N/P ratio (10) resulted in a complete loss of electrophoretic mobility (Figure 2). The efficient complexation of anionic pDNA with cationic lipid at the N/P ratio (10) indicated the absence of free pDNA at this ratio. Therefore, the N/P ratio (10) was chosen for the development of DOTAP/DOPE-lipoplexes and PEI-polyplexes. 3.3. Formulation of Nanoplexes. The effect of different concentration of PAA on quality attributes of PAA-DOTAP/ PC-lipoplexes is shown in Figure 3. A significant (p < 0.05)

3. RESULTS AND DISCUSSION 3.1. Preparation and Characterization of Lipoplexes. Thin film hydration method was used for the preparation of cationic liposomes. The selection of mole ratio (1:1 w/w mole ratio) of lipids, chloroform:methanol mixture (9:1 v/v), hydration time (2 h), and sonication time (for 30 s at amplitude 80 (3 cycles with 1 min pulse off) was based on our previous experience in formation of cationic liposomes.22,23 Lipoplexes were formed by electrostatic attraction between positively charged quaternary amine groups of DOTAP with negatively charged phosphate groups of DNA. Because the lipoplex formation was based on electrostatic attraction, hence optimized N/P ratio (ratios of moles of the amine groups of cationic lipid to those of the phosphate groups of DNA) can play a crucial role in complexation efficiency. By keeping this point in mind, a different concentration of lipid was incubated with a constant concentration of pDNA. A change in N/P ratio significantly (p < 0.05) affected the particle size and zeta potential of the lipoplexes as evident by Figure 1.

Figure 1. Effect of N/P ratio on particle size and zeta potential. Values are expressed as mean ± SD (n = 6).

Figure 3. Effect of PAA coating on particle size and zeta potential. Values are expressed as mean ± SD (n = 6).

A significant (p < 0.01) increase (from N/P ratio 1 to 2.5) followed by decrease (from N/P ratio 2.5 to 10) in particle size was observed while the difference was insignificant (p > 0.05) at lower (0.5−1) and higher (10−12.5) N/P ratio. At lower N/P ratios (0.5 and 1) zeta potential was negative while shifting of zeta potential toward positive side was observed at higher N/P

reduction in particle size and zeta potential was observed as the PAA concentration was increased from 0.01 to 0.05 μg/μL although the difference was insignificant (p > 0.05) at higher concentrations (0.05−0.07 μg/μL). Aggregation at lower

Figure 2. Electrophoretic mobility of DNA at different N/P ratios. 2419

dx.doi.org/10.1021/mp400036w | Mol. Pharmaceutics 2013, 10, 2416−2425

Molecular Pharmaceutics

Article

A continuous increase in absorbance up to 10 min was observed in case of naked DNA which could be attributed to cleavage of DNA to nucleotide in presence of DNase, while almost constant values of absorbance were observed in case of all lipoplexes which could be attributed to the electrostatic interactions of cationic amine group in carrier moiety with negatively charged phosphate group of DNA which ultimately protected it against DNase activity. Although the protection efficiency of all lipoplexes formulations was significantly (p < 0.001) higher in comparison with naked DNA yet, the difference was insignificant (p > 0.05) among different lipoplexes, polyplexes, and nanoplexes. This revealed that DNA interacted and complexed with all of the cationic formulations with similar efficiency. 3.5. Serum Stability of Developed Complexes. 3.5.1. Physicochemical Characterization. Changes in physicochemical characteristics were taken as a preliminary quality control tool to determine the stability of developed formulation in presence of serum. The changes observed in physicochemical characteristics of different formulations are shown in Table 2. A significant increase (p < 0.001) in size and switch of zeta potential from positive to negative side was observed in case of conventional lipoplex formulations which could be ascribed to the adsorption of negatively charged proteins over the positively charged systems. Our findings are in agreement with previous reports in which high surface charge density has been reported to cause higher adsorption of plasma proteins.31 A small increase in size was also observed in case of developed nanoplexes, but the changes were insignificant in comparison with other lipoplexes and polyplexes. A drop in zeta potential was also recorded, but still it was maintained toward positive side. The stability of developed nanoplexes might be the consequence of lower tendency of protein adsorption over the nanoplexes due to the strong electrostatic attraction already present between alternative layers of polyelectrolytes. 3.5.2. EtBr Intercalation Assay. EtBr interaction assay is based on the principle of intercalation of EtBr between the base pairs of DNA double helix which results in emission of intense fluorescence signal at 618 nm when excited at 516 nm. Presence of negatively charged serum proteins such as heparin and albumin can displace the DNA from its complexes.24 This free DNA can be intercalated by EtBr which results in change in fluorescent intensity. The fluorescent intensity observed in case of different complexes is shown in Figure 7. Except nanoplexes all of the complexes were found somewhat unstable as evident by significantly higher (p < 0.001) fluorescence observed in case of lipoplexes and polyplexes. Higher fluorescence observed in case of conventional complexes clearly revealed the poor stability of these complexes in the presence of serum which ultimately led to a dissociation of the complex and accessibility of free DNA to intercalate with EtBr. Also the dissociation of complex and release of DNA was found to be a slower and time-dependent process. The dissociation of complexes and release of DNA could be attributed to the interaction of lipidic vectors with negatively charged serum proteins. It is worthy to note that DOTAP/DOPE-lipoplexes were found less stable in comparison to DOTAP/PC-lipoplexes as evident by significantly higher (p < 0.05) fluorescence observed at the end of 4 h. The poor stability of DOTAP/DOPE-lipoplexes might be the result of fluid nature of the liposomes. DOPE contains a relatively small headgroup with two bulky acyl chains which inherently provide an inverted cone shape which is not favorable for

concentration could be attributed to an insufficient amount of PAA which resulted in aggregation rather than uniform coating with PAA. A shifting of zeta potential toward the negative side was observed in response to a higher concentration of PAA which might be due to the coating of PAA over the positively charged lipoplexes. By considering PAA (0.05 μg/μL) as optimized, the system was subsequently optimized for PEI coating. The effect of different concentrations of PEI on quality attributes of lipoplexes is shown in Figure 4.

Figure 4. Effect of PEI coating on particle size and zeta potential. Values are expressed as mean ± SD (n = 6).

A significant (p < 0.05) decrease in particle size was observed from 0.01 to 0.05 μg/μL of PEI while the difference was insignificant (p > 0.05) at higher concentrations. Large aggregates observed at lower concentrations could be due to the same reason as an insufficient amount of PEI which resulted in aggregation. At higher PEI concentrations the zeta potential shifted to the positive side and can be attributed to deposition of positively charged PEI over the negatively charged PAADOTAP/PC-lipoplexes. PEI (0.05 μg/μL) was finally selected as it resulted in lowest particle size and sufficient charge reversal from negative to positive. The quality attributes of different lipoplexes are shown in Table 1. SEM and AFM analysis demonstrated formation of almost spherical particles. A good correlation was observed between the results of DLS and SEM/ AFM analysis (Figure 5A and B). Table 1. Quality Attributes of Optimized Lipoplexesa formulation DOTAP/PC liposome (without DNA) DOTAP/PC lipoplexes DOTAP/DOPE lipoplexes PEI polyplexes nanoplexes a

size (nm) 114.5 167.1 191.8 185.4 242.6

± ± ± ± ±

9.2 14.6 13.5 12.3 9.4

zeta potential (mV) 56.8 29.4 27.6 36.2 23.0

± ± ± ± ±

1.7 1.3 2.7 1.6 1.5

Values are expressed as mean ± SD (n = 6).

3.4. DNase Protection Assay. An efficient gene delivery system should offer protection to associated plasmid DNA against nucleases present in serum and in cytoplasm during its journey from site of administration to its pathological target site. DNase I is a versatile enzyme that nonspecifically cleaves DNA to release 5′-phosphorylated nucleotides. The cleavage of DNA results in an increase of absorbance at 260 nm due to the free nucleotide. The change in absorbance value at different time points for different formulations is shown in Figure 6. 2420

dx.doi.org/10.1021/mp400036w | Mol. Pharmaceutics 2013, 10, 2416−2425

Molecular Pharmaceutics

Article

Figure 5. Surface morphology analysis of nanoplexes formed at a N/P ratio of 10 using (A) SEM and (B) AFM.

Figure 6. DNase protection assay of lipoplex.

Figure 7. Stability of different complexes in presence of serum determined using the EtBr intercalation assay. Values are expressed as mean ± SD (n = 6).

bilayer formation under physiological conditions. Although, bilayer can be formed in association with other lipids, yet the formed bilayer structure can undergo destabilization upon change in pH and ionic strength.32 Our findings are in union with previous reports.28 PEI-polyplexes were somewhat stable in comparison to lipoplexes which could be attributed to strong electrostatic interaction of DNA with polymer. Nanoplexes were found almost stable as no significant change in fluorescent intensity was observed up to 4 h which might be the consequence of protective barrier of alternative layer of polyelectrolytes which sheltered direct exposure of DNA to negatively charged fractions of serum responsible for dissociation and release of DNA from the complex. 3.6. Cell Uptake and Nuclear Colocalization. The cellular uptake and subcellular translocation of nanoplexes inside their target cells was studied using confocal microscopy. Representative confocal images of nanoplex-incubated MCF-7, HeLa, and HEK-293 cells are shown in Figure 8. Cell nuclei were stained with DAPI (blue fluorescence), while nanoplexes are labeled with RITC (red fluorescence). As evident from the images, all formulations showed appreciable internalization in

all the three cell lines. It seems that the positive charges associated with the formulations facilitated their binding and internalization through interaction with negatively charged phospholipids on cell membranes and/or passive diffusion. Our next step was to elucidate whether the nanoplexes developed in course of our study were able to deliver the loaded DNA into the site of transfection, that is, nucleus. Subsequently, nuclear colocalization studies were performed by analyzing the degree of overlap between DAPI stained nuclei and RITC labeled nanoplexes. The colocalization in the entire field of view was assessed through scatter plot analysis. The extent of colocalization between RITC labeled nanoplexes and DAPI stained nuclei was measured in terms of Pearson’s correlation coefficient (r), which was calculated using instrument software. A colocalization coefficient greater than or equal to 0.5 (r ≥ 0.5) is usually taken as an indicator of good colocalization. In the present case, the highest nuclear colocalization was exhibited by the nanoplexes (r > 0.5), followed by PEIpolyplexes and lipoplexes. The observed trend can be

Table 2. Physicochemical Characteristics of Different Formulations after Exposure to Seruma size (nm) formulation DOTAP/PC-lipoplexes DOTAP/DOPE-lipoplexes PEI-polyplexes nanoplexes a

zeta potential (mV)

before 167.1 191.8 185.4 242.6

± ± ± ±

14.6 13.5 12.3 9.4

after 472.8 689.3 527.3 289.2

± ± ± ±

before 27.2 31.4 38.6 11.9

29.4 27.6 36.2 23.0

± ± ± ±

1.3 2.7 1.6 1.5

after −21.3 −24.4 −26.1 12.5

± ± ± ±

1.6 1.2 2.3 2.2

Values are expressed as mean ± SD (n = 6). 2421

dx.doi.org/10.1021/mp400036w | Mol. Pharmaceutics 2013, 10, 2416−2425

Molecular Pharmaceutics

Article

Figure 8. (A) Confocal microscopic images of MCF-7, HeLa, and HEK-293 cell lines treated with RITC-complexes. The white line represents scale bar of 10 nm. The left, middle, and right panel of each incubation type represents RITC fluorescence, an overlay of RITC and DAPI fluorescence, and scatter plot analysis for the entire field of view. The horizontal and vertical axes of each scatter plot represents the values of pixels in channel 2 (ch2) and channel 1 (ch1), respectively.

Figure 9. CLSM images of MCF-7, HeLa, and HEK-293 treated with DOTAP/PC-lipoplexes, DOTAP/DOPE-lipoplexes, PEI-polyplexes, nanoplexes, and lipofectamine.

rationalized if we concentrate on the stability profile of individual formulation in presence of serum. As evident from Table 2, all formulations other than nanoplexes displayed negative charges in the presence of serum. Thus, among all of the investigated complexes, only nanoplexes are supposed to retain positive charges in the intracellular milieu, which facilitate their rapid endolysosomal escape via proton sponge effect and subsequent colocalization in the nucleus. 3.7. Transfection Efficiency. The gene transfection efficiency of nanocarriers largely depends on the cell type and uptake mechanism. The cell uptake of naked DNA is restricted due to its large size, hydrophilic nature, and highly negatively

charged surface. These limitations can be overcome by complexing DNA with an oppositely charged carrier, which not only renders the complex positive but also aids in their internalization through adsorptive endocytosis.33 Figure 10 presents the comparative transfection efficiency of different formulations in different cell lines. In line with the results of nuclear colocalization experiments, the highest transfection was exhibited by the nanoplexes (Figure 9). The DOTAP/DOPE based formulation exhibited higher transfection than their DOTAP/PC analogues; albeit the latter was found to be less stable in serum. The observed anomaly may be attributed to the pH sensitivity of DOPE, which results in inversion of lipid 2422

dx.doi.org/10.1021/mp400036w | Mol. Pharmaceutics 2013, 10, 2416−2425

Molecular Pharmaceutics

Article

2,3-bis(dioleyloxy)-1-propaniminium pentahydrochloride, DOSPA) with five potentially charged amine groups in contrast to DOTAP, a monovalent lipid which makes the lipofectamine comparable to PEI but superior to DOTAP. The results are in concordance with previous reports. 38 The transfection efficiency of nanoplexes in all cell lines was significantly higher (P < 0.001) than any other formulation. The transfection efficiency of nanoplexes was 50−80, 11−12, 6−7, and 5−6 fold higher as compared to DOTAP/PC-lipoplexes, DOTAP/ DOPE-lipoplexes, PEI-polyplexes, and lipofectamine, respectively. The highest transfection observed in case of nanoplexes could be attributed to additional protection provided by alternative coating barrier of polyelectrolytes along with endolysosomal escape due to proton sponge effect. 3.8. Cytotoxicity Assay. High transfection efficiency and low cytotoxicity are the primary requirements of an ideal gene delivery vector. Most of the conventional gene delivery vectors suffer from high cytotoxicity due to excessive cationic charge density, which depends upon the nature of the polycation, that is, the primary, secondary, tertiary, or quaternary amine group as well as the molecular weight of cationic polymer/lipid. Molecules with high charge density and molecular weight have been reported to exhibit high cytotoxicity.8,39 The percentage cell viability observed in case of different carriers on different cell lines is shown in Figure 11. Among all of the complexes, PEI-polyplexes exerted the highest cytotoxicity which might be a consequence of high surface charge due to multivalent cationic nature of the polymer. For the same reason, lipofectamine also exhibited significantly higher toxicity than nanoplexes. Even after executing the highest transfection efficiency, nanoplexes induced the lowest cytotoxicity (almost 90% cell viability) in all cell lines. The observed results could be ascribed to the charge neutralization of PEI with PAA.8 Moreover, the PEI concentration used in nanoplexes (0.05 μg/μL) was much lower in comparison to PEI-polyplexes (10 μg/μL), which could also be attributed to reduced toxicity of nanoplexes.

Figure 10. Transfection efficiency of DOTAP/PC-lipoplexes, DOTAP/DOPE-lipoplexes, PEI-polyplexes, nanoplexes, and lipofectamine (*** p < 0.001). Values are expressed as mean ± SD (n = 4).

bilayer structure to hexagonal-II structure that can fuse and destabilize the membrane while facilitating endolysosomal escape.34,35 Among other formulations, DOTAP/DOPE-lipoplexes showed lower transfection efficiency than the PEIpolyplexes. The observed effects may be attributed to the wellreported proton sponge effect. Following lysosomal compartmentalization, the secondary and tertiary amines of PEI get protonated in the acidic endosomal environment, which prevented pH reduction of endosomes. The increased cationic charge density of PEI further assists the influx of chloride and water leading to osmotically triggered vesicle burst and subsequent release of DNA from endosome without lysosomal degradation.36,37 As compared to DOTAP/DOPE lipoplexes, lipofectamine showed significantly higher (p < 0.05) transfection; albeit the efficiency was comparable (p > 0.05) with PEI polyplexes. Lipofectamine contains a multivalent cationic lipid ((+)-N,N-dimethyl-N-[2-(sperminecarboxamido)ethyl]-

Figure 11. Percentage cell viability of MCF-7, HeLa, and HEK-293 cell lines after 48 incubation with different carriers (a; in comparison with PEI polyplexes, b; in comparison with lipofectamine, c; in comparison with DOTAP/DOPE lipoplexes) (*** p < 0.001, **p < 0.01, * p < 0.05). Values are expressed as mean ± SD (n = 4). 2423

dx.doi.org/10.1021/mp400036w | Mol. Pharmaceutics 2013, 10, 2416−2425

Molecular Pharmaceutics

Article

constructs mediated tumor gene therapy. ACS Nano 2009, 3, 1493− 1505. (15) Godbey, W.; Wu, K. K.; Mikos, A. G. Tracking the intracellular path of poly (ethylenimine)/DNA complexes for gene delivery. Proc. Natl. Acad. Sci. U.S.A. 1999, 96, 5177−5181. (16) Zhang, S.; Xu, Y.; Wang, B.; Qiao, W.; Liu, D.; Li, Z. Cationic compounds used in lipoplexes and polyplexes for gene delivery. J. Controlled Release 2004, 100, 165−180. (17) Jain, P.; Jain, S.; Prasad, K. N.; Jain, S. K.; Vyas, S. P. Polyelectrolyte coated multilayered liposomes (nanocapsules) for the treatment of Helicobacter pylori infection. Mol. Pharmaceutics 2009, 6, 593−603. (18) Ge, L.; Möhwald, H.; Li, J. Polymer-stabilized phospholipid vesicles formed on polyelectrolyte multilayer capsules. Biochem. Biophys. Res. Commun. 2003, 303, 653−659. (19) Mansoori, G. A.; Brandenburg, K. S.; Shakeri-Zadeh, A. A comparative study of two folate-conjugated gold nanoparticles for cancer nanotechnology applications. Cancers 2010, 2, 1911−1928. (20) Guo, W.; Lee, R. J. Receptor-targeted gene delivery via folateconjugated polyethylenimine. AAPS J. 1999, 1, 20−26. (21) Garcia L, B. M.; Duzgunes, N.; Tros de Ilarduya, C. Serumresistant lipopolyplexes for gene delivery to liver tumour cells. Eur. J. Pharm. Biopharm. 2007, 67, 58−66. (22) Jain, S.; Kumar, D.; Swarnakar, N. K.; Thanki, K. Polyelectrolyte stabilized multilayered liposomes for oral delivery of paclitaxel. Biomaterials 2012, 33, 6758−6768. (23) Jain, S.; Patil, S. R.; Swarnakar, N. K.; Agrawal, A. K. Oral delivery of doxorubicin using novel polyelectrolytes stabilized liposomes (layersomes). Mol. Pharmaceutics 2012, 9, 2626−2635. (24) Xu, Y.; Szoka, F. C., Jr. Mechanism of DNA release from cationic liposome/DNA complexes used in cell transfection. Biochemistry 1996, 35, 5616−5623. (25) Agrawal, A. K.; Gupta, P. N.; Khanna, A.; Sharma, R. K.; Chandrawanshi, H. K.; Gupta, N.; Patil, U. K.; Yadav, S. K. Development and characterization of in situ gel system for nasal insulin delivery. Pharmazie 2010, 65, 188−93. (26) Jain, A. K.; Swarnakar, N. K.; Das, M.; Godugu, C.; Singh, R. P.; Rao, P. R.; Jain, S. Augmented anticancer efficacy of doxorubicinloaded polymeric nanoparticles after oral administration in a breast cancer induced animal model. Mol. Pharmaceutics 2011, 8, 1140−51. (27) Hwa Kim, S.; Hoon Jeong, J.; Joe, C. O.; Gwan Park, T. Folate receptor mediated intracellular protein delivery using PLL-PEG-FOL conjugate. J. Controlled Release 2005, 103, 625−634. (28) Li, S.; Tseng, W. C.; Stolz, D. B.; Wu, S. P.; Watkins, S. C.; Huang, L. Dynamic changes in the characteristics of cationic lipidic vectors after exposure to mouse serum: implications for intravenous lipofection. Gene Ther. 1999, 6, 585−994. (29) Singh, R. P.; Das, M.; Thakare, V.; Jain, S. Functionalization density dependent toxicity of oxidized multiwalled carbon nanotubes in a murine macrophage cell line. Chem. Res. Toxicol. 2012, 25, 2127− 37. (30) Kasturi, S. P.; Sachaphibulkij, K.; Roy, K. Covalent conjugation of polyethyleneimine on biodegradable microparticles for delivery of plasmid DNA vaccines. Biomaterials 2005, 26, 6375−6385. (31) Gessner, A.; Lieske, A.; Paulke, B.; Muller, R. Influence of surface charge density on protein adsorption on polymeric nanoparticles: analysis by two-dimensional electrophoresis. Eur. J. Pharm. Biopharm. 2002, 54, 165−70. (32) Litzinger, D. C.; Huang, L. Phosphatidylethanolamine liposomes: drug delivery, gene transfer and immunodiagnostic applications. Biochim. Biophys. Acta 1992, 1113, 201−227. (33) Wiethoff, C. M.; Middaugh, C. R. Barriers to nonviral gene delivery. J. Pharm. Sci. 2002, 92, 203−217. (34) Chesnoy, S.; Huang, L. Structure and function of lipid-DNA complexes for gene delivery. Annu. Rev. Biophys. Biomol. Struct. 2000, 29, 27−47. (35) Safinya, C. R. Structures of lipid−DNA complexes: supramolecular assembly and gene delivery. Curr. Opin. Struct. Biol. 2001, 11, 440−448.

4. CONCLUSION The proposed nanoplexes were found to have great potential in the development of lipid−polymer based hybrid novel gene delivery vehicle with improved transfection efficiency and reduced cytotoxicity in comparison with lipoplexes and polyplexes. Also the ease of preparation and scalability can fortify the research in the area of polyelectrolytes and their use to design effective nonviral vector in gene delivery. Encouraged with our findings, we are at the stage of animal experimentation which would be reported in due course.

■

AUTHOR INFORMATION

Corresponding Author

*Telephone: 0172-2292055. Fax: 0172-2214692. E-mail: [email protected]; sanyogjain@rediffmail.com. Notes

The authors declare no competing financial interest.

■

ACKNOWLEDGMENTS

■

REFERENCES

Authors are thankful to Indian National Science Academy (INSA), Government of India, New Delhi, for providing financial assistance and Department of Biotechnology (DBT) for providing fellowship to S.K. Authors are also thankful to Director, NIPER for providing necessary infrastructure facilities.

(1) Friedmann, T.; Roblin, R. Gene therapy for human genetic disease? Science 1972, 175, 949−955. (2) Cavazzana-Calvo, M.; Thrasher, A.; Mavilio, F. The future of gene therapy. Nature 2004, 427, 779−781. (3) Zhang, X.; Godbey, W. T. Viral vectors for gene delivery in tissue engineering. Adv. Drug Delivery Rev. 2006, 58, 515−534. (4) Buchschacher, J. G. L. Safety considerations associated with development and clinical application of lentiviral vector systems for gene transfer. Curr. Genomics 2004, 5, 19−35. (5) Mintzer, M. A. S.; E., E. Nonviral vectors for gene delivery. Chem. Rev. 2009, 109, 259−302. (6) Dufes, C.; Uchegbu, I. F.; Schatzlein, A. G. Dendrimers in gene delivery. Adv. Drug Delivery Rev. 2005, 57, 2177−2202. (7) Felgner, P. L.; Ringold, G. M. Cationic liposome-mediated transfection. Nature 1989, 337, 387−388. (8) Lv, H.; Zhang, S.; Wang, B.; Cui, S.; Yan, J. Toxicity of cationic lipids and cationic polymers in gene delivery. J. Controlled Release 2006, 114, 100−109. (9) Brown, M. D.; Schatzlein, A. G.; Uchegbu, I. F. Gene delivery with synthetic (non viral) carriers. Int. J. Pharmaceutics 2001, 229, 1− 21. (10) Luo, D.; Saltzman, W. M. Synthetic DNA delivery systems. Nat. Biotechnol. 2000, 18, 33−37. (11) 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. (12) Pathak, A.; Swami, A.; Patnaik, S.; Jain, S.; Chuttani, K.; Mishra, A. K.; Vyas, S. P.; Kumar, P.; Gupta, K. C. Efficient tumor targeting by polysaccharide decked polyethylenimine based nanocomposites. J. Biomed. Nanotechnol. 2009, 5, 264−277. (13) Jiang, D.; Salem, A. K. Optimized dextran-polyethylenimine conjugates are efficient non-viral vectors with reduced cytotoxicity when used in serum containing environments. Int. J. Pharmaceutics 2012, 427, 71−9. (14) Pathak, A.; Kumar, P.; Chuttani, K.; Jain, S.; Mishra, A. K.; Vyas, S. P.; Gupta, K. C. Gene expression, biodistribution, and pharmacoscintigraphic evaluation of chondroitin sulfate-PEI nano2424

dx.doi.org/10.1021/mp400036w | Mol. Pharmaceutics 2013, 10, 2416−2425

Molecular Pharmaceutics

Article

(36) Sonawane, N. D.; Szoka, F. C.; Verkman, A. Chloride accumulation and swelling in endosomes enhances DNA transfer by polyamine-DNA polyplexes. J. Biol. Chem. 2003, 278, 44826−44831. (37) Wagner, E. Polymers for siRNA Delivery: Inspired by Viruses to be Targeted, Dynamic, and Precise. Acc. Chem. Res. 2012, 45, 1005− 1013. (38) Uchida, E.; Mizuguchi, H.; Ishii-Watabe, A.; Hayakawa, T. Comparison of the efficiency and safety of non-viral vector-mediated gene transfer into a wide range of human cells. Biol. Pharm. Bull. 2002, 25, 891−897. (39) Choksakulnimitr, S.; Masuda, S.; Tokuda, H.; Takakura, Y.; Hashida, M. In vitro cytotoxicity of macromolecules in different cell culture systems. J. Controlled Release 1995, 34, 233−241.

2425

dx.doi.org/10.1021/mp400036w | Mol. Pharmaceutics 2013, 10, 2416−2425