Dextran-protamine polycation: An efficient nonviral and haemocompatible gene delivery system

Colloids and Surfaces B: Biointerfaces 81 (2010) 195–205

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Dextran-protamine polycation: An efficient nonviral and haemocompatible gene delivery system Jane Joy Thomas, M.R. Rekha, Chandra P. Sharma ∗ Division of Biosurface Technology, Biomedical Technology Wing, Sree Chitra Tirunal Institute for Medical Sciences & Technology, Trivandrum, Kerala, India

a r t i c l e

i n f o

Article history: Received 15 March 2010 Received in revised form 3 July 2010 Accepted 5 July 2010 Available online 13 July 2010 Keywords: Dextran Protamine ctDNA Nanoplex Haemocompatible Gene delivery

a b s t r a c t Despite the remarkable progress in the field of gene therapy with viral vectors, nonviral vectors have attracted great interests due to their unique properties. Imparting desired characteristics to nonviral gene delivery systems requires the development of cationic polymers. The purpose of this work was to design a cationic derivative (Dex-P) of dextran using protamine in order to assert target specific cellular binding. Our objective was to elucidate the potential use of Dex-P as a haemocompatible, nontoxic and efficient nonviral candidate for gene therapy. Nanoplexes were prepared with calf thymus DNA and Dex-P. Derivatization was confirmed by FTIR, gel permeation chromatography and TNBS assay. Dynamic light scattering and TEM studies determined the size and morphology of the nanoplex. The buffering behaviour was assessed by acid base titration. Complexation stability was evaluated using agarose gel electrophoresis and EtBr displacement assay. The protection of ctDNA from nuclear digestion and the effect of plasma components towards stability of the nanoplexes were also analyzed. Various haemocompatible studies were performed to check haemolysis, aggregation, clotting time, and complement activation. Transfection and cytotoxicity experiments were performed in vitro. The nanosize, spherical shape and stability of nanoplexes were affirmed. Various experiments conducted confirmed Dex-P to be nontoxic and haemocompatible. Transfection experiments revealed the capability of Dex-P to facilitate high gene expression and cellular uptake in HepG2 cells. With the improved physicochemical, biological and transfection properties, Dex-P seems to be a promising gene delivery system. © 2010 Elsevier B.V. All rights reserved.

1. Introduction For the past few decades, gene therapy has become important as a therapeutic entity for human diseases. The aim of gene therapy lies in the designing of delivery systems that can introduce exogenous genetic materials encoding a therapeutic protein or a specific virus antigen into target cells efficiently [1]. Significant progress has taken place in the development of viral gene delivery systems using retrovirus and adenovirus. Viral vectors can be employed for gene therapy because of their high gene transfer capability. Various disadvantages like long term risks, mutagenesis, toxic response, limit of plasmid size to be delivered, oncogenic effects and possible recombination with wild type virus have stimulated the development of nonviral gene delivery systems [2]. Nonviral vector mediated gene therapy is currently one of the most attractive strategies used due to their biocompatibility, biodegradability, minimal cytotoxicity, lack of pathogenicity and low immunogenicity. The development of nonviral vectors consisting of dendrimers, liposomes and cationic polymers are currently

∗ Corresponding author. Tel.: +91 471 2520214; fax: +91 471 2341814. E-mail address: [email protected] (C.P. Sharma). 0927-7765/$ – see front matter © 2010 Elsevier B.V. All rights reserved. doi:10.1016/j.colsurfb.2010.07.015

being focussed [3]. Cationic polymers have been useful in neutralizing the anionic nature of DNA thus efficiently condensing DNA to allow its easy cellular entry [4]. Cationic polymers represent an interesting class of vectors in plasmid DNA delivery formulations. Various cationic polymers including synthetic amino acid polymers like poly (l-lysine) and polyethyleneimine (PEI), natural DNA binding proteins like histones, carbohydrate based polymers like chitosan and various dextran derivatives have been reported [5]. Cationic polymers have nowadays reached the efficiencies of viral vectors due to their special abilities. They can perform multi tasks like condensation of DNA for easy cell migration, protection of DNA from degradation, shield against undesired interactions, enhancement of cell binding and cellular uptake. In reality, a polymer is unable to perform all the required tasks. Cationic polymers also face several drawbacks like lack of specificity, inhibition by serum components, toxicity and nonbiodegradability. Such obstacles have been overcome to a certain extent by designing a gene vector where dextran is catonized with protamine enabling the modified polymer to multi task. Dextrans have been proven to be efficient nonviral vectors [6]. Dextran is a biodegradable polysaccharide consisting of 1,6 linked d-glucopyranose residue [7]. Dextran exhibits various attractive properties like its remarkable degree of biocompatibility,

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biodegradability, availability, ease of use and improved transfection efficiency with reduced toxicity. Dextran has also been found to be used as a plasma extender, for ophthalmic use and for intrauterine examinations [8]. The major drawback of dextran limiting its usage in delivery systems is its high polarity which excludes its transcellular passage and its susceptibility to enzymatic digestion in human body [9]. Dextran-PEI conjugations having increased cytotoxicity and high transfection efficiency have also been reported [10]. In view of its numerous unique functional characteristics, dextran continues to find important applications in the field of gene therapy. As reported earlier by this group, dextran–glycidyltrimethylammonium chloride conjugate in complexation with DNA led to the formulation of a potential nonviral and haemocompatible gene delivery system [11]. Surface modification was found to be of great importance in the development of a gene carrier. This can be achieved with cationic electrolytes like chitosan, PEI or protamine [12]. Reports on synthesis of dextran based polycations using oligoamines have been investigated. The major drawback faced by many of them includes decrease in transfection efficiency [13]. Oligoamines have been found to possess similar characters as that of cell penetrating peptides. Its efficiency to deliver genes and proteins into cells has also been investigated [14]. Our work involves the construction of a new dextran derivative designated Dex-P where dextran is modified by protamine according to its specific functional and biological needs. Protamine is a positively charged polypeptide prepared from the sperm of salmon or herring. It is a cationic peptide having arginine residues [15]. It has been ascertained to act as a nuclear localizing signal [16]. The uniqueness of protamine which signifies its role in the development of a gene delivery system is its possibility to facilitate the intracellular release of nucleic acid [17]. Its special properties have bestowed it a significant role in medical applications as a carrier for injectable insulin, an antagonist for heparin, as an antibacterial ingredient in food products, for modification of various polymers [18] and in antisense delivery system [19]. Protamine was seen to neutralize the action of heparin through acid base interactions. Even though it acted as a heparin antagonist, excess of the protein prolonged the activated clotting time [20]. Protamine of low molecular weight was found to be less toxic and safer for clinical use [21]. With this information we hypothesized that the conjugation of protamine with dextan may lead to nuclear targeting which is a property lacking in the case of quaternary trimethyl ammonium groups leading to better dextran based gene delivery vector. The main aim of this study was to design a polycation that should be nontoxic, haemocompatible, biodegradable, able to form stable complex with DNA, protect DNA from degradation and deliver the genetic material to target cells. Our motivation focussed on the elucidation of the potential use of Dex-P as an efficient, safe and a promising candidate for gene therapy. The physicochemical characteristics, morphology, cytotoxicity, haemocompatibility, stability in serum and transfection efficiency were evaluated. The transfection efficiency and safety of the polymer/DNA complex was analyzed under in vitro conditions.

salt from calf thymus (ctDNA) was from Worthington Biochemical Corp. disuccinidylcarbamate (DSC) and 4-methylaminopyridine (DMAP) was from Fluka, USA. YOYO iodide and Hoechst 33342 was from Invitrogen. Fetal bovine serum (FBS) was from GIBCO (USA). All other reagents were of analytical grade from Merck, India. 3. Methods 3.1. Cationization of dextran To 10 ml of dimethylsulphoxide (DMSO), 500 mg of dextran was added and allowed to dissolve. Disuccinidylcarbamate (DSC) (6 mM) and 4-methylaminopyridine (DMAP) (6 mM) was added to the dextran solution respectively. The reaction mixture was continuously stirred for 6 h at room temperature. For the cationization reaction, 200 mg of protamine was added to the above solution and then subjected to continuous stirring for 24 h at 4 ◦ C. The reaction solution was precipitated with an excess of acetone. The precipitation obtained was washed twice with methanol. The product was then recovered by extensive dialysis against double distilled water. The obtained dextran derivative defined as Dex-P was finally stored at 4 ◦ C. 3.2. Fourier transform infrared spectroscopy The test samples of dextran and Dex-P were subjected to FTIR in a dry powdered form. The FTIR spectra of the samples was detected and compared using Nicolet 5700 Spectrophotometer. 3.3. Gel permeation chromatography Gel permeation chromatography of Dex-P was performed based on size exclusion using Sephadex G100-120 column. The column was calibrated using three different dextran molecular weight standards (Sigma–Aldrich Chemicals Co, USA). 3.4. TNBS assay Test samples containing dextran, protamine and Dex-P was prepared at a concentration of 1 mg/ml of which 200 ␮l of the samples were used for the assay. To the samples, 200 ␮l each of 4% sodium bicarbonate and 0.1% of 2,4,6-trinitrobenzenesulfonic acid (TNBS) was added and incubated for 2 h at 37 ◦ C. Later, 200 ␮l of 1 N hydrochloride (HCl) was added. The absorbance was read at 355 nm against blank prepared with water taken as sample and all the above. 3.5. Formulation of nanoplexes Nanoplexes of various weight ratios ranging from 0.25:1 to 7:1 were formulated by the addition of varying concentrations of Dex-P to a constant amount of ctDNA (10 ␮g) and diluted to a total volume of 100 ␮l with saline. The test nanoplexes were then vortexed and incubated for 30 min at room temperature before use.

2. Materials and methods

3.6. Particle size and zeta potential determination

2.1. Materials

The nanoplexes were characterized by the determination of their size and zeta potential. The particle size distribution was analyzed by the dynamic laser light scattering measurement using Zetasizer Nano ZS (Malvern Instruments Ltd., UK) at a temperature of 25 ◦ C. The complexes were prepared in saline with Dex-P/DNA ratios ranging from 1:1 to 4:1. The surface charge of the complexes of varying ratios was also evaluated using the Zetasizer Nano ZS (Malvern Instruments Ltd., UK) at 25 ◦ C. The Smoluchowsky approximation was used to check the zeta potential.

Dextran (MW 35,600 Da), protamine chloride, sodium hydroxide, Ethidium bromide (EtBr), 3-(4,5-dimethylthialzol-2-yl)-2,5diphenyl tetrazolium bromide (MTT), Dulbecco’s modified Eagle’s medium (DMEM), trypsin, ethylenediaminetetraacetic acid (EDTA), branched PEI (MW 25,000 Da), Sephadex G100-120 and DNase I was purchased from Sigma–Aldrich Chemicals Co, USA. pGL3 control DNA was from Promega, USA. Deoxyribonucleic acid sodium

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3.7. Transmission electron microscopy The morphology of Dex-P/DNA nanoplexes was observed by transmission electron microscopy. The test sample was prepared by placing a drop of the sample onto a formvar coated grid and allowed to dry at room temperature. The samples were then visualized. 3.8. Acid base titration assay The buffering capacity of Dex-P was evaluated by acid base titration over a pH range of 10–5. Test solutions consisting of different concentrations of Dex-P was first adjusted to pH 10 with 1 N sodium hydroxide. The solution was then titrated with 20 ␮l aliquots of 0.1 N HCl to a pH of 5. The mixtures of different pH values obtained were recorded during the titration. 3.9. Gel retardation studies The ability of Dex-P to condense ctDNA was confirmed by the gel retardation assay. Nanoplexes at desired ratios were loaded onto a 0.8% agarose gel containing ethidium bromide in Tris–acetate–EDTA (TAE) buffer solution. Electrophoresis was performed for 30 min at 100 V in a Bio-rad electrophoresis system (Bio-rad laboratories CA, USA). The DNA bands were visualized and the gel was photographed with using MultiImageTM Light Cabinet (Alpha Innotech Corp., San Leandro, CA, USA). 3.10. Blood compatibility studies Whole blood was collected from a healthy volunteer and anticoagulated with sodium citrate (ratio of blood to anticoagulant taken was 9:1). Isolation of blood components were done as describe elsewhere [11]. 3.10.1. Haemolysis and blood cell aggregation The erythrocytes were washed thrice with saline before use. Nanoplexes of varying ratios were mixed with RBCs at 1:1 ratio and then incubated for 2 h at 37 ◦ C. The supernatant was spinned off at 1500 × g for 5 min. Haemoglobin release was monitored spectrophotometrically at 399 nm. Triton X-100 and 0.9% NaCl were taken as the positive and negative control respectively. Erythrocytes and leukocyte aggregation studies were done as per the protocol reported previously [11]. Varying ratios of DexP/DNA complexes were incubated with 100 ␮l of erythrocyte suspensions and 100 ␮l leukocyte suspensions respectively. The nanoplexes were added and the mixture was incubated at 37 ◦ C for 1 h. Nanoplexes of different ratios were also mixed with 100 ␮l of platelets isolated from whole blood and kept for incubation at 37 ◦ C for 1 h. Aggregation was examined through phase contrast microscope (Leica DMI 3000B, Germany) at a magnification of 40×. 3.10.2. In vitro whole blood clotting time Nanoplexes were added to 100 ␮l of citrated whole blood at ratio 1:1 (v/v) and incubated for 5 min at room temperature. This was followed by the addition of 25 ␮l of 50 mM calcium chloride solution. The time from the addition of calcium chloride to the first visible sign of clot formation was recorded as the whole blood clotting time. Incubation of blood with 0.9% NaCl was taken as the control. 3.10.3. Complement activation The activation of the complement system was determined by recording the concentration of the complement protein C3 (mg/100 ml) present in serum by the turbidity assay. The test samples included PEI, protamine and Dex-P taken in three different concentrations (25 ␮g, 50 ␮g, 100 ␮g). The samples were incubated in 100 ␮l of serum for 1 h at 37 ◦ C and then mixed with the reagents

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provided. The intensity of turbidity was determined by measuring the absorbance at 340 nm and the concentration of C3 was thus calculated. The turbidity assay was performed using the C3 assay kit according to the protocol provided by the manufacturers. 3.11. EtBr displacement assay The extent of compaction of ctDNA by Dex-P was affirmed by the EtBr displacement assay as described elsewhere [11]. The fluorescence intensity of the complexes was measured at an excitation at 510 nm and emission at 590 nm using an automated microplate reader (Finstruments Micro plate Reader USA). The results were annotated as relative fluorescence intensity. 3.12. Cytotoxicity The influence of Dex-P on cell proliferation and viability was assayed by the 3-(4,5-dimethylthialzol-2-yl)-2,5-diphenyl tetrazolium bromide (MTT) assay. Mouse fibroblast cell lines (L929 cell lines) were seeded in multi well tissue culture plates and cultured at 37 ◦ C, 5% CO2 atmosphere for 24 h. The cells were then incubated with Dex-P of varying concentrations along with positive and negative controls in fresh DMEM medium containing 10% FBS at 37 ◦ C for 24 h under 5% CO2 atmosphere. Cells incubated with medium were taken as the negative control. After 24 h, samples were removed, MTT reagent (0.2 mg/ml) was added to each well and the cells were further incubated for 4 h. Later, the reagent was removed and DMSO was added to dissolve the formazan crystals thus formed. Plates were incubated for 5 min. The results were expressed as the mean % cell viability relative to untreated cells by measuring the absorbance of the samples at 640 nm using an automated microplate reader (Finstruments Micro plate Reader USA). 3.13. In vitro transfection studies HepG2 cell lines (human hepatocellular liver carcinoma cell lines) were used to evaluate the transfection of pGL3 plasmid mediated by Dex-P. Before transfection trials, cells were plated in multi well plates in DMEM medium containing 10% FBS and incubated overnight. Nanoplexes were prepared from ratios 2:1 to 4:1 using Dex-P and pGL3 plasmid. The complexes were added to the cells and incubated at 37 ◦ C for 24 h under 5% CO2 atmosphere. The medium was replaced later with fresh medium and the cells were cultured for another 48 h. The cells were washed with normal saline and the cell lysates were harvested after permeabilization with lysis buffer for 5 min. Luciferase assay reagent was added to the supernatant collected. The luciferase activity expressed as RLU (relative light units)/mg cellular protein was recorded by a luminometer (Chamaleon, Hidex). Total protein was measured using bicinchonic acid (BCA) protein assay (Pierce, USA). 3.14. Plasmid trafficking studies The capability of Dex-P as a gene carrier was assured by the observation of pGL3 plasmid trafficking. Plasmid DNA was tagged with a high affinity intercalating fluorescent labeling dye, YOYO iodide by incubation for 1 h. Nanoplexes were prepared with DexP and YOYO tagged plasmid at ratio 3:1 and then incubated with HepG2 cells for 4 h at 37 ◦ C in DMEM with 10% FBS. The entry of the complexes into the nucleus was assessed with the help of nuclear staining by Hoechst 33342 on incubation for 0.5 h. The cells were then washed with phosphate buffer saline (PBS) and fixed in 4% formaldehyde. Plasmid trafficking was viewed and photographed using fluorescence microscope (Leica DM IRB, Germany).

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Fig. 1. FTIR spectra of dextran (A) and of Dex-P (B).

4. Results 4.1. Cationization of dextran To increase the delivery of complexes into the cells, dextran was modified with protamine to obtain a water soluble dextran derivative (Dex-P) containing amino groups. Synthesis of cationized dextran proceeded in two steps. In the first step, DSC and DMAP were used to activate the hydroxyl groups present on dextran. In the following step, the activated hydroxyl group of dextran was conjugated to the amino group of protamine at different ratios. The optimized ratio of dextran to protamine being 2.5 was then used for further studies.

chromatography analysis revealed the increased molecular weight of modified dextran, Dex-P, as 39,810 Da. Fig. 2 depicts the comparison of percentage of free amino groups of dextran, Dex-P and protamine evaluated by the reaction with TNBS. Dextran showed negligible percentage of amino groups whereas protamine contained higher percentage of free amino groups. Dex-P on reaction with TNBS provided an interme-

4.2. Characterisation of Dex-P FTIR studies were performed to get an insight into the characterization of Dex-P. The spectra of dextran and Dex-P were compared as in Fig. 1. The spectrum of Dex-P depicted strong amide absorbance as new peaks at 1559.0 cm−1 , 1541.0 cm−1 and 1457.7 cm−1 . A shift in the characteristic peak of hydroxyl group of dextran occurred from 3264 cm−1 to 3199 cm−1 . The molecular weight of the native dextran is 35,600 Da and the gel permeation

Fig. 2. Percentage profile of free amino groups present in dextran, protamine and Dex-P evaluated on reaction with TNBS.

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Fig. 4. Acid base titration curves of Dex-P of different concentrations (40–100 ␮g/ml). The solution was titrated with 0.01 N HCl. The titration curve of PEI (1 mg/ml) is depicted as reference.

ity versus pH profile of Dex-P on comparison with PEI taken as the reference polymer. The slope of Dex-P obtained in the pH range of 10–5 of pH titration curves was found to be similar from pH 10 to 6.5 at all concentrations taken. Buffering capacity can be controlled with the concentration of Dex-P. Beyond the point of addition of 500 ␮l of HCL, pH curve of Dex-P decreased rapidly than PEI. The pH curve of Dex-P at concentration 40 ␮g was similar to that of PEI. Fig. 3. Transmission electron micrograph of Dex-P/DNA nanoplex formulated.

4.6. Gel retardation studies

diate absorbance proving the success of protamine substitution by the presence of free amino groups.

The strength of DNA binding by Dex-P was assessed by the complete retardation of the migration of DNA in agarose gel. Upon increase in the ratio of Dex-P/DNA complexes, migration of DNA towards anode was observed to be retarded as shown in Fig. 5. No release of DNA from the complexes indicated the strong interaction between Dex-P and DNA. At ratio 2:1 and above, ctDNA was found to be totally retained in the complex which was observed by the inhibition of DNA migration in agarose gel electrophoresis (Lanes 9–13).

4.3. Polyplex formation—zeta potential and particle size determination Constant amount of ctDNA was complexed with varying concentrations of Dex-P to obtain complexes of various weight ratios ranging from 0.25:1 to 7:1. Complete condensation of ctDNA by Dex-P resulted in the formulation of small and compact nanoplexes of nanometer size. Zeta potential determination provides information on the surface charge and stability of the nanoplexes. The zeta potential value for dextran and naked DNA was evaluated to be −2.97 mV and −14.7 mV respectively. The negative zeta potential was provided by the presence of the hydroxyl groups of dextran and the phosphate groups of ctDNA respectively. The surface charge of Dex-P/DNA complexes ranging from 35.9 mV to 39.6 mV increased according to the increase in the weight ratio of Dex-P to ctDNA (Table 1). The nanoplexes exhibited positive charge due to the presence of amino groups of protamine. Particle size was measured based on dynamic light scattering performed at 25 ◦ C. As shown in Table 1, Dex-P complexed ctDNA to a mean hydrodynamic diameter ranging from 7.741 nm to 6.481 nm according to the polymer/DNA weight ratio. No significant differences were detected in the size distribution profile obtained along with their polydispersity indices.

4.7. In vitro plasma stability The stability of Dex-P/DNA complexes in the presence of human plasma was evaluated on agarose gel as shown in Fig. 6. DNA bands obtained both in the presence and absence of plasma was found to be similar. Experiments clearly showed that the complexes are stable in the presence of plasma. No detectable DNA migrations from

4.4. Transmission electron microscopy The size exclusion barrier of Dex-P was checked using transmission electron microscopy. The diameter of the nanoplex was observed to be significantly similar to the hydrodynamic diameter measured by dynamic light scattering as depicted in Fig. 3. 4.5. Acid base titration studies Buffering capability of Dex-P was studied by the acid base titration of the polymer in the pH range of 10–5. PEI showed a buffering capability over a wide pH range. Fig. 4 shows the buffering capac-

Fig. 5. Agarose gel electrophoresis of Dex-P/DNA complexes at different weight ratios detected after 30 min of incubation compared with naked DNA. Lane 1 indicates calf thymus DNA. Lanes 2–13 indicates the polymer/DNA complexes at various ratios (0.25, 0.5, 0.75, 1, 1.25 1.5, 1.75, 2, 3, 4, 5, 6 and 7).

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Table 1 Zeta potential (mV), hydrodynamic diameter (nm) and polydispersity index of Dex-P–ctDNA complexes formulated at different weight ratios. Samples (polymer:DNA weight ratios)

Zeta potential (mV) ±sd

Dex-P:DNA 1:1 Dex-P:DNA 2:1 Dex-P:DNA 3:1 Dex-P:DNA 4:1 Dex-P:DNA 5:1 Dex-P:DNA 6:1

35.9 36.2 37.5 37.7 38.9 39.6

± ± ± ± ± ±

1.80 0.97 1.25 1.09 0.79 0.63

Average diameter (nm) ±sd 6.780 6.481 7.741 6.928 6.843 6.998

± ± ± ± ± ±

0.55 1.25 0.28 1.27 1.39 2.30

Polydispersity index (PDi) ±sd 0.174 0.203 0.216 0.232 0.217 0.222

± ± ± ± ± ±

0.049 0.039 0.037 0.028 0.015 0.003

Dex-P/DNA complexes were observed in the presence of plasma (Lanes 2–8).

4.8. DNase I sensitivity assay Resistance of ctDNA to DNase I degradation was analyzed on agarose gel. Fig. 7 shows the capability of Dex-P/DNA complexes to protect DNA from nuclease degradation. Intact DNA was observed to be totally degraded (Lane 1). Excellent protection ability of DexP/DNA complexes at higher ratios was assessed by the fluorescence seen in the wells which was due to the strong interaction between the polymer and DNA. (Lanes 2–8).

4.9. Interaction of Dex-P with plasma proteins The extent of binding of Dex-P with plasma proteins was studied by performing native polyacrylamide gel electrophoresis (PAGE). Fig. 8 shows the comparative study of Dex-P with dextran and PEI. The absence of protein bands proved the capability of PEI to bind to plasma proteins (Lane 2). The interaction of dextran with plasma proteins were less pronounced (Lane 1). No significant binding of plasma proteins to Dex-P was detected by native PAGE due to unfavourable electrostatic interactions (Lanes 4–6). Dex-P confirmed its safety as a gene carrier.

Fig. 6. Stability of Dex-P/DNA complexes after 30 min incubation with plasma at room temperature. Lane 1 indicates naked calf thymus DNA. Lanes 2–8 indicates the polymer/DNA complexes at different weight ratios (1–7).

Fig. 7. Electrophoretic mobility data of Dex-P/DNA complexes following DNase I digestion. Lane 1: enzyme treated naked DNA, Lanes 2–8: enzyme treated DexP/DNA complexes formulated at different weight ratios (1–7).

Fig. 8. Native PAGE analysis of Dex-P, PEI and dextran with plasma after 20 min incubation at room temperature. Lane 3 indicates plasma proteins. Lanes 4–6 indicates the Dex-P and plasma. Lane 2 indicates PEI and plasma. Lane 1 indicates dextran and plasma.

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Fig. 9. Microscopic view of erythrocytes after incubation with Dex-P/DNA nanoplexes at ratio 2:1 and 4:1 (A and B), 0.9% NaCl (C) and PEI (D) for 30 min respectively.

4.10. Haemolysis At physiological pH, Dex-P/DNA complexes at varying ratios showed negligible haemolytic activity as shown in Table 2. The nanoplexes showed 0.0240% of % haemolysis at a lower ratio of 1:1 and 0.7454% at a higher ratio of 7:1. 4.11. RBC aggregation The effect of Dex-P/DNA complexes on RBCs was investigated by performing an erythrocyte aggregation assay in vitro. Very strong aggregation of erythrocytes was observed on incubation of Table 2 Percentage of haemolysis of Dex-P/DNA complexes of different weight ratios measured after incubation with erythrocytes for 30 min. Triton X-100 and 0.9% NaCl was taken as the positive and negative control respectively. Samples

(%) Haemolysis

Standard deviation

Dex-P:DNA 1:1 Dex-P:DNA 2:1 Dex-P:DNA 3:1 Dex-P:DNA 4:1 Dex-P:DNA 5:1 Dex-P:DNA 6:1 Dex-P:DNA 7:1 Dex-P (100 ␮g) Triton X-100 Saline

0.0240 0.0961 0.1563 0.1923 0.2404 0.4568 0.7454 0.5242 100 0

0.0007 0.0154 0.0042 0.0754 0.0077 0.0139 0.0269 0.0875 – –

Nanoplexes of varying ratios were mixed with RBCs at 1:1 ratio and then incubated for 2 h at 37 ◦ C. The supernatant was spinned off at 1500 × g for 5 min. haemoglobin release was monitored spectrophotometrically at 399 nm. Triton X-100 and 0.9% NaCl were taken as the positive and negative control respectively.

PEI/ctDNA complex at ratio 2:1. No RBC aggregation was observed in the Dex-P/DNA nanoplex group of ratio 2:1 and 4:1 as shown in Fig. 9. 4.12. WBC aggregation The studies showed no aggregation evidence for ratio 2:1 and 4:1 of Dex-P/DNA complexes incubated with WBC for 30 min (data not shown). PEI/ctDNA complex at ratio 2:1 caused WBC aggregation whereas Dex-P proved the ability to inhibit WBC aggregation. 4.13. Platelet aggregation Platelets suspended with Dex-P and ctDNA complexes of ratios 2:1 and 4:1 did not aggregate (data not shown). PEI/DNA complex of ratio 2:1 aggregated platelets. Dex-P/DNA complexes either inhibited platelet aggregation or had no demonstrable aggregation effect. 4.14. In vitro clotting time Protamine presence provided the shortest clotting time of 70.5 s. The plasma solutions incubated with Dex-P/DNA complexes of various ratios on recalcification gave a clotting time ranging from 85 s to 116.5 s as shown in Table 3. The upper limit of the clotting time as detected using the negative control was 132 s. 4.15. Complement activation The effect on Dex-P on the complement system was assessed and compared with PEI by the turbidity assay as depicted in Fig. 10.

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Table 3 In vitro clotting time of dextran, protamine and Dex-P/DNA complexes of various ratios in seconds with standard deviation. 0.9% NaCl is taken as the negative control. Samples

In vitro clotting time (s)

0.9% NaCl (saline) Dextran Protamine Dex-P/DNA complex ratio 1:1 Dex-P/DNA complex ratio 2:1 Dex-P/DNA complex ratio 3:1 Dex-P/DNA complex ratio 4:1 Dex-P/DNA complex ratio 5:1

132.0 135.5 70.5 85.0 115.5 116.5 95.0 98.5

± ± ± ± ± ± ± ±

2.829 0.707 0.707 1.414 0.707 0.707 1.414 0.707

Fig. 12. Cytotoxicity of HepG2 cells after incubation of dextran, Dex-P and PEI for 24 h. Cytotoxicity was evaluated by the MTT assay and expressed as % cell viability.

toxicity by Dex-P was observed on L929 cell lines. Dex-P at lower concentration exhibited substantially lower cellular toxicity. Triton X-100 and 10% Phenol was taken as negative and positive control respectively. Dex-P provided more than 80% viable cells even at a higher concentration of 100 ␮g in comparison with untreated cells. Dextran showed similar results as Dex-P whereas PEI enhanced cellular toxicity. 4.18. In vitro transfection studies Fig. 10. Complement activation by PEI, Dex-P and protamine on incubation with plasma at 37 ◦ C for 1 h. Samples were tested at three different concentrations. Concentration of C3 was expressed as mg/100 ml of plasma.

PEI was found to strongly activate the complement system. Dex-P of various concentrations did not activate C3 and was similar to the extent of the C3 activation by protamine. 4.16. Dye intercalation assay The dye intercalation assay was performed to prove the stable complexation of Dex-P with ctDNA. EtBr alone showed a % fluorescence of 20.24% whereas the % fluorescence of EtBr bound with DNA was taken as 100%. As shown in Fig. 11, the successive addition of Dex-P to DNA resulted in gradual decrease of the fluorescence intensity monitored from ratio 2:1 to 3.8:1. 4.17. Effect of Dex-P on cell viability MTT assay was used to determine the cytotoxicity of Dex-P at varying concentrations. As shown in Fig. 12, no significant cyto-

The pattern of transfection in HepG2 cells by various weight ratios of Dex-P complexed with pGL3 plasmid DNA ranging from 2:1 to 4:1 was shown in Fig. 13. The highest level of gene expression was detected at lower ratio of Dex-P/DNA complexes. The transfection efficiency of Dex-P was compared with branched PEI, 25 kDa, used as control. As the ratio of the complex went higher, the efficiency of transfection reduced due to the reduction in electrostatic interaction after a certain ratio. 4.19. Plasmid trafficking studies The evidence for the uptake of nanoplexes by HepG2 cells were revealed by fluorescence microscopy. After 1 h incubation of nanoplexes with the cells, nanoplexes were observed in the region of cytosol beneath the cell membrane. A 4 h incubation micrograph depicted the presence of nanoplexes in the nuclear region stained blue with Hoechst 33342 (Fig. 14). 5. Discussion Various classes of cationic polymers efficient to be used as gene carriers have been designed [5]. The major drawback faced by various polymers is cytotoxicity. Dextran, a natural biocompatible polymer, possesses many favourable properties. Several studies

Fig. 11. The decrease in the relative fluorescence intensity was detected with the increase in the concentration of Dex-P. Complexes of Dex-P and ctDNA were taken in different ratios (2:1–3.8:1). The fluorescence intensity of EtBr and EtBr complexed with ctDNA was also detected.

Fig. 13. Luciferase expression in HepG2 cells transfected by Dex-P/DNA complexes and PEI/DNA complexes taken at three different ratios (2:1–7:1).

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Fig. 14. Internalization of Dex-P/DNA nanoplex in HepG2 cells at two different incubation times, 1 h (A) and 4 h (B). The ratio of the nanoplex taken was 2:1.

on gene delivery systems have been performed using cationized dextran [13]. The objective of this study was to prepare a haemocompatible, nontoxic and water soluble dextran derivative. One of the main barriers towards high transfection efficiency is the disruption of complexes made by anionic polymers and DNA. The rationale for selecting protamine for this study was to cationize dextran so as to improve DNA complexation and its entry into mammalian cells. Dex-P was prepared by the attachment of amino groups of protamine to the dextran chain. Polyplexes of positive surface charges were formed by exceeding the number of positive charges of Dex-P than that of the negative charges present in DNA. Complexation in different weight ratios were made possible by the electrostatic interaction between constant ctDNA concentration and varying Dex-P concentration. The encounter of these polyplexes enabled their interaction with the negatively charged cellular membrane and in their uptake via endocytosis. The absorption bands characteristic of dextran and Dex-P was compared by FTIR to confirm the derivatization of dextran with protamine. Attention was focussed on characterizing the occurrence of new peaks at 559.0 cm−1 , 1541.0 cm−1 and 1457.7 cm−1 which was due to the presence of amide bond. The reaction of dextran with protamine led to the decrease in the peak characteristic for hydroxyl group and to the appearance of new peaks characteristic to amino groups. The molecular weight of Dex-P was determined to be 39,810 Da by gel permeation chromatography. The TNBS assay helped to determine the presence of free amino groups in a polymer [22]. The absorbance of dextran showed the lack of amino groups in the polymer chain. Protamine showed a high absorbance due to the availability of free amino groups. Dex-P gave an intermediate absorbance reading affirming the cationization of dextran using the amino groups of protamine. This ascertained the derivatization of dextran by protamine. Investigation of zeta potential is an important part of characterization. It has a substantial influence on the interaction of the polymer with DNA, the stability of nanoplexes and its entry into the cell [23]. Zeta potential of Dex-P/ctDNA complexes of various ratios was evaluated to be positive. The higher the polymer concentration, the more was the zeta potential value detected. The positive charges obtained by Dex-P have been advantageous for the bioadhesion of the nanoplex to the negatively charged biological membranes. Cationization of dextran also helped to reduce charge–charge repulsion at the surface of the biological membranes and form stable complexes with gene expression systems. Particle size determination is an important parameter that controls their uptake by target cells [24]. Cationic polymers have the advantage to formulate nanoplexes. The evaluation of particle size showed the condensation of Dex-P/ctDNA nanoplexes to a diame-

ter range of 6–7 nm. Nanometer size helps in the mimicking of cell organelles which thus helps in the transport of therapeutic genes from the cytoplasm to the nucleus. Particle size can be influenced by many factors including the concentration of DNA [25]. The proton binding capacity of a polymer correlates with transfection efficiency [26]. Cationic polymers induce facilitated endosomal escape. Proton sponge effect promotes the endosomal escape of polymer/DNA complexes by the endosomal acidification process based on the capacity of the polymer to bind protons. An increase in the swelling of endocytic vesicles causes the escape of the complexes into the cytoplasm [27]. Polyethyleneimine (PEI) is considered as the best vector due to its proton sponge effect based on the presence of high amount of amine functions [28]. Dex-P showed considerable buffering capacity in the pH range from 7 to 6. This ascertained the polymer a buffering capability built due to the cationic behaviour extended to dextran by protamine. The proton sponge effect played a major role in the escape of Dex-P/ctDNA nanoplexes from the lysozyme thus avoiding its degradation by the enzyme. The decrease in the titration curves of Dex-P when compared to PEI may be due to the decrease in the number of amino groups which may be protonated. The buffering capacity of Dex-P can be controlled with the concentration of the polymer. Cationic polymers having primary amines can strongly interact with ctDNA to form nanoplexes [29]. The ability of Dex-P to interact with DNA, sensitivity of the nanoplexes to DNase I enzyme and the stability of nanoplexes in plasma was studied by observing the electrophoretic mobility behaviour of free DNA and DNA released by Dex-P/DNA complexes through agarose gel electrophoresis studies. Reduction in the electrophoretic mobility of DNA relates to the ability of Dex-P to interact strongly with DNA. Dex-P/ctDNA nanoplexes at ratios less than 2:1 had lower DNA binding capability. As the concentration of Dex-P increased, the ctDNA gradually lost its mobility due to the increase in the shielding effect of the polymer. The electrostatic interaction between the polymer and ctDNA caused the retardation of DNA in the well [30]. Naked DNA was used as a control. The determination of the stability of nanoplexes in plasma supports the prediction of its possibility for in vivo use. The increase in the binding ability of DNA can be correlated with the increase in the charge of Dex-P/ctDNA complex [31]. Dex-P was capable of protecting DNA from dissociation by serum. This was confirmed by the detection of similar DNA bands both in the presence and absence of plasma. Retardation of ctDNA in the well affirmed the stability of Dex-P. Plasma showed no effect on Dex-P/DNA complexes due to the strong shielding effect of Dex-P over DNA. The major obstacle during the formulation of a gene carrier is the exposure of DNA to degradation by serum DNase I [32]. In the intracellular environment, endosomes fuse with lysosomes

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thus exposing DNA of the nanoplexes to the lysosome enzymes. A capable gene vector provides protection to DNA against nucleases. This was assessed by the DNase protection assay. Naked DNA was detected to be completely degraded. DNA was found to be protected from DNase effect in the Dex-P/DNA complexes by the intense fluorescence seen in the well when compared to the naked DNA digested with DNase I where no band was seen. The entrapment of DNA by Dex-P was affirmed by the detection of intense fluorescence whereas diffuse fluorescence indicated the leaching of small amounts of DNA from the nanoplexes. Cationic polymer can result in nonspecific interactions with plasma proteins. Native PAGE was performed to check the binding extent of Dex-P with plasma proteins. No observable absence of plasma protein bands was detected. Dex-P showed better stability in comparison with PEI as the presence of PEI led to the absence of protein bands confirming its interaction with plasma proteins. One of the most important properties required for a gene delivery system is haemocompatibility. The biological environment first encountered by complexes after administration into the body is blood. Stability in blood is an important criterion for the development of gene carriers in in vivo applications. Haemolysis study was performed to quantify the damaging property of polymers on the cells membranes of blood cells. It is expressed as a percentage of haemoglobin release. Varying polymers like PEI exhibited high haemolysis [33]. To improve gene transfer, Dex-P/ctDNA complexes should be stable and inert in blood. The haemocompatibility of Dex-P was supported by the absence of its lytic activity. DexP showed no detectable RBC membrane disruptive activity thus proving itself to be nonhaemolytic. In in vivo studies, cationic polymers can interact nonspecifically with blood components. A potential carrier should be capable of minimizing the aggregation of blood cells. The basis of aggregation lies in the electrostatic interactions between the anionic cell membrane and cationic polymers. Various aggregation assays were performed in vitro to investigate the effect of Dex-P on erythrocytes, leukocytes and platelets. Blood rheology mainly depends on RBC aggregation [34]. No aggregation was detected in the case of RBC aggregation studies with Dex-P. Use of cationic polymers as gene vectors is often hampered due to imperfect haemocompatibility. No evidence of aggregation was observed on incubation of Dex-P with WBCs. Platelet aggregation is the most widely investigated functional responses of clinical relevance. Dex-P proved its uniqueness in haemocompatibility by finalizing the aggregation studies with platelets. Platelets suspended in Dex-P did not aggregate due to the compatible behaviour of the polymer. Protamine was discovered to have the shortest clotting time. According to Mochizuki et al., excess protamine weakens the clot structure and thus increases activated clotting time after the reversal of heparin effect [20]. Modification of dextran with protamine provided a clotting time similar to that obtained with normal saline. This proved the efficiency of Dex-P as a biocompatible gene carrier. When different ratios of Dex-P/ctDNA complexes came in contact with the recalcified blood solution, the clotting time showed very few variations. An important criterion to be studied while designing a gene vector is the activation of complement system induced by the carrier. Complement activation mainly depends on the donor’s complement reactivity and type of polymer surface. Amino groups have the potential to activate the complement system [35]. The effect of Dex-P of different concentrations was examined on complement activation using turbidity assay. The intensity of turbidity caused by the immuno complexes formed by C3 in sample and anti C3 antibody is measured to find the concentration of C3 present in the sample. This provides information on the activation of the complement system. PEI was found to strongly activate the complement

system. Dex-P, on the other hand, did not activate C3 and was similar to the extent of the C3 activation by protamine. The stability of condensation of DNA with Dex-P was characterized by EtBr displacement assay. EtBr intercalates with the minor groove of DNA helix fluorescing the nucleic acids [36]. The pattern of decreased fluorescence intensity observed was due to the exclusion of EtBr molecules from DNA upon addition of Dex-P. As the concentration of Dex-P increased, the polymer began to replace EtBr in its accessibility towards DNA by strong interactions. The lack of DNA accessibility by EtBr is shown by the significant reduction of the fluorescence signals with increase in Dex-P/DNA ratio. An important prerequisite for gene vectors is that they should not provoke toxic effects on healthy cells and tissues. The major cause of cytotoxicity was the interaction of the cationic polymer with either the plasma membrane or cell components [37]. A good indication of cell injury is the reduction in cellular metabolic activity. Viability of cells were assessed on L929 cells by reduction of MTT following exposure to various concentrations of Dex-P. Modification of dextran with protamine reduced polymer cytotoxicity. As the concentration of Dex-P decreased, it significantly reduced the cytotoxic potential of the polymer. This may be due to the counterbalance of the negative charge of DNA by the positive charge of Dex-P which minimizes the interactions with cell membrane. The effect of Dex-P can be minimized by keeping the polymer induced cytotoxicity minimal. A major requirement in transfection is the release of DNA by the vector after cell entry. Several studies have been performed on cationic polymers and their modification so as to increase transfection efficiency while keeping cytotoxicity manageable [38]. This provides Dex-P the capability to deliver DNA to its target site without enhancing cellular toxicity. The interpretation of gene transport experiments mainly rely on the entry of DNA into mammalian cells. Transfection was performed in HepG2 cells using Dex-P and pGL3 plasmid DNA complexed in different weight ratios. On comparison with branched PEI, 25 kDa, Dex-P showed a considerable high level of gene expression. The strong complexation of Dex-P with plasmid DNA provided the polymer favourable transfection efficiency at a lower ratio. As the ratio increased, transfection efficiency reduced due to the decrease in the shielding effect of polymer over DNA. The efficiency of transfection was found to be enhanced in the presence of serum. In this study, in the presence of 10% FBS, the transfection capability of Dex-P increased for all ratios investigated. Transfecting cells in the presence of serum have several advantages like less time consuming and easier transfections, low experimental cost and avoidance of cell’s deprivation of serum [39]. The major aim of gene carriers is to deliver DNA to its target site in the nucleus. The major role of polymer/DNA complexes is to unpack the plasmid which when translocated into the nucleus leads to the production of the desired protein by the gene expression system. The assessment of the fluorescence of YOYO tagged plasmid/Dex-P nanoplexes assured the uptake of the nanoplexes rather than their adsorption onto the cell surface. The localization of the plasmid seen within the nucleus correlates to its transfection efficiency. This study also signified the dependence of nanoplex uptake on incubation time of nanoplexes with HepG2 cells. As the incubation time increased, the plasmid which was seen distributed in the cytoplasm was found to enter the nucleus. This proved the nuclear entry of the nanoplex. Acknowledgements We express our sincere thanks to the director, SCTIMST and the head, BMT Wing for the facilities provided. Authors are thankful to the financial support from FADDS, DST, New Delhi. The authors would also like to thank Dr. H.K. Varma for the FTIR facility and Dr. Annie John for the TEM facility.

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References [1] [2] [3] [4] [5] [6] [7] [8] [9] [10] [11] [12] [13] [14] [15] [16] [17] [18] [19]

S. Choi, K.D. Lee, J. Control. Release 131 (2008) 70. Y. Tsuchiya, T. Ishii, Y. Okahata, T. Sato, J. Bioact. Compat. Polym. 21 (2006) 519. H. Eliyahu, Y. Barenholz, A.J. Domb, Molecules 10 (2005) 34. B. Vroman, I. Ferreira, C. Jerome, R. Jerome, V. Preat, Int. J. Pharm. 344 (2007) 88. S.C. De Smedt, J. Demeester, W.E. Hennink, Pharm. Res. 17 (2000) 113. W.C. Tseng, C.M. Jong, Biomacromolecules 4 (2003) 1277. M. Yalpani, P.O. Hedman, Crit. Rev. Biotechnol. 3 (1985) 375. H. Hosseinkhani, T. Aoyama, O. Ogawa, Y. Tabata, J. Control. Release 88 (2003) 297. Y. Zhang, C.Y. Won, C.C. Chu, J. Polym. Sci. Polym. Chem. 38 (2000) 2392. W.C. Tseng, C.H. Tang, T.Y. Fang, J. Gene Med. 6 (2004) 895. J.J. Thomas, M.R. Rekha, C.P. Sharma, Int. J. Pharm. 389 (2010) 195. M. Singh, G. Ott, J. Kazzaz, et al., Pharm. Res. 18 (2001) 1476. T. Azzam, H. Eliyahu, L. Shapira, M. Linial, Y. Barenholz, A.J. Domb, J. Med. Chem. 45 (2002) 1817. D.J. Mitchell, D.T. Kim, L. Steinman, C.G. Fathman, J.B. Rothbard, J. Pept. Res. 56 (2000) 318. J.D. Lewis, N. Saperas, Y. Song, M.J. Zamora, M.J. Chiva, Proc. Natl. Acad. Sci. 101 (2004) 4148. F.L. Sorgi, S. Bhattacharya, L. Huang, Gene Ther. 4 (1997) 961. S. Futaki, W. Ohashi, T. Suzuki, et al., Bioconjug. Chem. 12 (2001) 1005. J.M. Martínez Gómez, N. Csaba, S. Fischer, et al., J. Control. Release 130 (2008) 161. D. Lochmann, J. Weyermann, C. Georgens, R. Prassl, A. Zimmer, Eur. J. Pharm. Biopharm. 59 (2005) 419.

205

[20] T. Mochizuki, P.J. Olson, F. Szlam, J.G. Ramsay, J.H. Levy, Anesth. Analg. 87 (1998) 781. [21] J.F. Liang, V.C. Yang, Y. Vaynshteyna, Biochem. Biophys. Res. Commun. 336 (2005) 653. [22] A.F.S. Habeeb, Anal. Biochem. 14 (1966) 328. [23] A. Berthold, K. Cremer, J. Kreuter, J. Control. Release 39 (1996) 17. [24] M. Ruponen, S. Yla-Herttuala, A. Urtti, Biochim. Biophys. Acta 1415 (1999) 331. [25] R. Kircheis, L. Wightman, E. Wagner, Adv. Drug Deliv. Rev. 53 (2001) 341. [26] W.T. Godbey, K.K. Wu, G.J. Hirasaki, A.G. Mikos, Gene Ther. 6 (1999) 1380. [27] J.P. Behr, Chimia 51 (1997) 34. [28] M. Hashida, R.I. Mahato, K. Kawabata, T. Miyao, M. Nishikawa, Y. Takakura, J. Control. Release 41 (1996) 91. [29] A.M. Funhoff, C.F. van Nostrum, G.A. Koning, N.M.E. SchuurmansNieuwenbroek, D.J.A. Crommelin, W.E. Hennink, Biomacromolecules 5 (2004) 32. [30] M.A. Wolfert, E.H. Schacht, V. Toncheva, K. Ulbrich, O. Nazarova, L.W. Seymour, Hum. Gene Ther. 7 (1996) 2123. [31] S. Danielsen, K.M. Vårum, B.T. Stokke, Biomacromolecules 5 (2004) 928. [32] A. Akinc, M. Thomas, A.M. Klibanov, R. Langer, J. Gene Med. 7 (2004) 657. [33] L. Aravindan, K.A. Bicknell, G. Brooks, V.V. Khutoryanskiy, A.C. Williams, Int. J. Pharm. 378 (2009) 201. [34] H. Demiroglu, Blood 89 (1997) 4236. [35] M. Toda, T. Kitazawa, I. Hirata, Y. Hirano, H. Iwata, Biomaterials 29 (2008) 407. [36] M.J. Warning, J. Mol. Biol. 13 (1965) 269. [37] D. Fischer, Y.X. Li, B. Ahlemeyer, J. Krieglstein, T. Kissel, Biomaterials 24 (2003) 1121. [38] M.L. Forrest, J.T. Koerber, D.W. Pack, Bioconjug. Chem. 14 (2003) 934. [39] E. Brunette, R. Stribling, R. Debs, Nucleic. Acids Res. 20 (1992) 1151.