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Hollow and degradable polyelectrolyte nanocapsules for protein drug delivery
Acta Biomaterialia 6 (2010) 210–217
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Hollow and degradable polyelectrolyte nanocapsules for protein drug delivery Shujun Shu a, Chunyang Sun a, Xinge Zhang a,*, Zhongming Wu b, Zhen Wang a, Chaoxing Li a,* a b
Key Laboratory of Functional Polymer Materials Ministry of Education, Institute of Polymer Chemistry, Nankai University, Tianjin 300071, People’s Republic of China Metabolic Diseases Hospital, Tianjin Medical University, Tianjin 300070, People’s Republic of China
a r t i c l e
i n f o
Article history: Received 2 March 2009 Received in revised form 18 May 2009 Accepted 11 June 2009 Available online 14 June 2009 Keywords: Degradable Polyelectrolyte capsules Protein Polysaccharides Layer-by-layer technique
a b s t r a c t Biodegradable hollow capsules encapsulating protein drugs were prepared via layer-by-layer assembly of water-soluble chitosan and dextran sulfate on protein-entrapping amino-functionalized silica particles and the subsequent removal of the silica. In order to enhance the encapsulated efficiency and decrease its burst release, we designed this new system to fulfill these two goals. Bovine serum albumin (BSA), which was used as model protein, was entrapped in the nanocapsules. This system demonstrated a good capacity for the encapsulation and loading of BSA. The burst release was decreased to less than 10% in phosphate-buffered saline within 2 h. No significant conformation change was noted from the released BSA in comparison with native BSA by using circular dichroism spectroscopy. Cell viability study suggested that the nanocapsules had good biocompatibility. The drug release kinetics mechanism is Fickian diffusion. These kinds of novel composite nanocapsules may offer a promising delivery system for watersoluble proteins and peptides. Ó 2009 Acta Materialia Inc. Published by Elsevier Ltd. All rights reserved.
1. Introduction An oral route is the most convenient and comfortable means of administering protein drugs. However, peptides and proteins are administered through the non-parenteral route, which has poor absorption efficiency for patients. The bioavailability of peptide after oral administration is low due to its instability under low pH circumstances and poor absorption of proteins in the gastrointestinal (GI) tract. One possible way to improve the GI uptake of peptides is to encapsulate them in nanocapsules that can protect the peptide from being degraded in the GI tract and facilitate their transportation into systemic circulation [1,2]. In general, drug loading in micro/nanoparticulate systems can be done at one of two stages: (i) during the preparation of particles (incorporation) or (ii) after the formation of particles (adsorption by physical interaction). In this system, the drug is physically embedded into the matrix or adsorbed onto the surface [3]. These two methods of drug release readily lead to an initial burst release effect. Various methods of loading have been developed to improve the encapsulation efficiency, but no research has been found that focuses on both increasing the encapsulation efficiency and decreasing the burst release effect. Microencapsulation is a promising technique for biomedical applications. However, much research has provided evidence that particles (<500 nm) can cross membranes of epithelial cells * Corresponding authors. Tel.: +86 22 2350 1645; fax: +86 22 2350 5598. E-mail addresses: [email protected] (X. Zhang), [email protected] (C. Li).
through endocytosis while larger particles (>5 lm) would be taken up via the lymphatic system [4,5]. Furthermore, the nanosized system stays in circulation for longer and therefore greatly extends the macromolecular biological activity compared to microparticles [6]. Another drawback of microcapsules is that they can be easily broken and collapsed. Numerous folds and creases observed are attributed to drying of the shells. The shells are also flattened and some spreading is noticed [7]. Chitosan (CS), a weak cationic polysaccharide produced by deacetylation of the natural polymer chitin, has many useful biological properties, such as biocompatibility, biodegradability and bioactivity. Most commercially available CS has a quite large molecular weight (mol. wt.) and needs to be dissolved in an acetic acid solution at a pH value of approximately 4.0. However, there are potential applications of CS in which a low mol. wt. would be essential. Given a low mol. wt., the polycationic characteristic of water-soluble chitosan (WSC) can be used together with a good solubility at a pH value close to physiological ranges. Loading of peptide or protein drugs at physiological pH ranges may maintain their bioactivity from decreasing [8]. WSC can improve the transportation of hydrophilic drugs across intestinal epithelium and it is believed that WSC can increase the permeability of epithelial tissues by disrupting intercellular tight junctions [9,10]. On this basis, WSC was used to prepare the nanocapsules. In this study, hollow polyelectrolyte nanocapsules prepared by layer-by-layer adsorption of polymer on nanosized sacrificial template particles hold immense potential for oral protein drug delivery. The capsules can be engineered with controlled sizes, composition and functionality, and can be loaded with model
1742-7061/$ - see front matter Ó 2009 Acta Materialia Inc. Published by Elsevier Ltd. All rights reserved. doi:10.1016/j.actbio.2009.06.020
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therapeutic such as insulin and low mol. wt. drugs. However, the encapsulated efficiency was so low following burst release [11]. In order to overcome these problems, we designed a new system to enhance the encapsulated efficiency and decrease its burst release. Bovine serum albumin (BSA), a protein which was used as model, was entrapped in the nanocapsules by both physical interaction and electrostatic attraction. After the required numbers of polyelectrolyte layers were deposited, the silica core was removed by exposure to an aqueous solution of HF/NH4F at pH 5.4. Otherwise, in this way, we can prevent the burst release by changing the membrane thickness and permeability of the nanocapsule. This result can also be avoided by washing nanocapsules with a proper solvent, which may lead to low encapsulation efficiency [12,13]. Herein, we report a polycation-free encapsulation method to obtain high concentrations of uncomplexed BSA chains confined within monodisperse, degradable nanocapsules. The encapsulation method exploits amino functionalized silica (SiO2–NH2) particles with mean size about 166 nm to adsorb BSA, followed by the assembly of WSC and dextran sulfate (DS). Removal of the template particles produces degradable and hollow nanocapsules filled with BSA [14].
2. Experimental 2.1. Materials WSC with a mol. wt. of 6 kDa and high mol. wt. chitosan (CS; mol. wt. = 50 kDa) were purchased from Yuhuan Ocean Biochemical Co. Ltd. (Zhejiang, China) and the degree of deacetylation was 93%. High mol. wt. dextran sulfate (500 kDa) was obtained from Amresco Company (USA). Tetraethoxysilane (TEOS) was purchased from Ke MiOu chemical Co. Ltd. (Tianjin, China). 3-Aminopropyltriethoxysilane (APTES, 99%) was obtained from Acros (USA). BSA, with a mol. wt. of 66.7 kDa, was purchased from Beijing Jun Yao WeiYe Biotechnology Co. Ltd. Coomassie brilliant blue G-250 was obtained from Fluka (USA). Fluorescein isothiocyanate (FITC) was purchased from Tianjin LianXing Biotechnology Co. Ltd. (Tianjin, China). All other reagents were of analytical grade without further purification before used. 2.2. Preparation of amino-functionalized silica nanospheres Amino-functionalized silica nanospheres (SiO2–NH2) were prepared using the modified Stöber method [15]. Briefly, 4 ml of tetraethoxysilane (TEOS) was added to a mixture of 3.3 ml ammonium hydroxide and 47 ml of ethanol with stirring; the reaction was continued for 24 h. The resulting silica colloidal dispersion was functionalized with APTES by quickly adding 0.3 ml of APTES with vigorous stirring. The mixture was stirred overnight. The nanosphere was purified by centrifugation and redispersion in ethanol. The former procedure was repeated three times. Meanwhile, bare silica nanospheres (SiO2) were obtained under the same condition without addition of APTES [16].
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[17], the amino groups of WSC (pKa = 6.5) are protonated with positive charge [18,19]. To form hollow capsules, the silica core was dissolved by treatment with HF/NH4F solution (2:8 M, pH 5.4) at 20 °C for 5 min, followed by multiple centrifugation–washing cycles. The washing cycles were repeated as necessary until the pH of the capsule suspension became identical to the pH of the washing buffer solution [20,21]. The schematic representation of the encapsulation of BSA within polyelectrolyte nanocapsules is shown in Scheme 1. 2.4. Characterization of nanospheres and nanocapsules Particle size was measured by photon correlation spectroscopy (PCS) at 25 °C with a detection angle of 90° and zeta potential was measured by ZetaPlus Measurement (Brookhaven, USA). Zeta potential is measured by applying an electric field across the dispersion. Particles within the dispersion with a zeta potential will migrate toward the electrode of opposite charge with a velocity proportional to the magnitude of the zeta potential. The velocity is measured using the technique of laser Doppler anemometry (LDA). The morphologies and approximate sizes of nanospheres, nanocapsules and the composite nanospheres were studied with a scanning electron microscope (X650 Hitachi) and a transmission electron microscope (Philips, Tecnai G2 T20ST). The Fourier transform infrared spectra of the nanocapsule samples were used to identify the removal of silica core. The nanocapsules were lyophilized by the freeze-dryer system (Flexi-Dry) to obtain dried capsules. These gained nanocapsules were mixed with KBr and pressed to a plate for further measurement. 2.5. Determination of BSA encapsulation efficiency and loading capacity of the nanocapsules Bound and unbound BSA was separated by ultracentrifugation of the nanosuspension at 38,000 rpm at 4 °C for 30 min (Optima LE-80k Ultracentrifuge, Beckman). The amount of free BSA in the clear supernatant was measured by a Bradfold protein assay using a UV spectrometer at 595 nm (Shimadzu UV 2550, Japan) [22]. The BSA encapsulation efficiency (EE) and loading capacity (LC) of the nanocapsules were calculated using the following equation:
total protein free protein 100% total protein total protein free protein LC% ¼ 100% nanocapsules weight
EE% ¼
All measurements were performed in triplicate and averaged.
2.3. Preparation of nanocapsules through layer-by-layer assembly A suspension of the SiO2–NH3+ particles (0.25 wt.%) was combined with a BSA solution and allowed to interact for 15 min, after which time the suspension was charged with WSC to a final concentration of 1.0 mg ml1. The WSC/DS multilayer build-up was initiated by exposing the particles with preadsorbed BSA (pI = 4.6) to a WSC solution at pH 5.4. At this pH value, DS shows negative sulfate groups (pKa < 1)
Scheme 1. Schematic representation of the encapsulation of BSA within polyelectrolyte capsules.
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2.6. Confocal laser scanning microscope visualization (CLSM) Fluorescent FITC-labeled BSA trapped in WSC/DS nanocapsules were prepared for the CLSM study (Olympus, 1 51). Synthesis of the FITC-labeled BSA was based on the reaction between the isothiocyanate group of FITC and the primary amino groups of BSA as reported in the literature [23]. Briefly, 100 mg of FITC in 150 ml of dehydrated methanol were added to 100 ml of 1% BSA solution. After 3 h of reaction in the dark at ambient conditions, FITC-labeled BSA was dialyzed at 4 °C to remove the unconjugated FITC. No fluorescence was detected in the supernatant. Then the dialyzed solution was freeze-dried for 24 h. The drug loaded WSC/DS nanocapsules were prepared as the procedure described above. Subsequently, the fluorescence images were observed using an argon laser (excitation at 488 nm, emission collected at a range of 510–540 nm). 2.7. Cell viability Cell viability was evaluated by using NIH 3T3 cell line. The cell line was cultured in Dulbecco’s modified eagles medium (DMEM) in a humidified atmosphere (5% CO2/95% O2). The cells were seeded into 96-well plates at 10,000 cells per well. The plates were then returned to the incubator and the cells were allowed to grow to confluence for 24 h. Various nanoparticles were dissolved in deionized water. After being filtered by paper filter, the resulting solution was diluted with culture medium to give a final range of concentrations from 0.1 to 1.5 mg ml1. Then the media in the wells were replaced with the pre-prepared culture medium–sample mixture (200 ll). The plates were then returned to the incubator and maintained in 5% CO2 at 37 °C for 48 h. Each sample was tested in six replicates per plate. After incubation culture medium and 20 ll of MTT solutions were used to replace the mixture in each well. The plates were then returned to the incubator and incubated for a further 4 h in 5% CO2 at 37 °C. Then, the culture medium and MTT were removed. DMSO (150 ll) was then added to each well to dissolve the formazane crystals. The plate was placed in 5% CO2 at 37 °C for 10 min and for 15 min at 6 °C before measurement. The optical density was read on a microplate reader at 490 nm. Cell viability was determined as a percentage of the negative control (untreated cells). 2.8. In vitro release studies The obtained hollow nanocapsules were redispersed in PBS solution after freeze-drying. The in vitro release profiles of BSA from nanocapsules were determined as follows: the BSA-loaded nanocapsules separated from 9 ml suspension was placed into test tubes with 6 ml of HCl solution of pH 1.4, and incubated at 37 °C under stirring at 60 rpm. After 2 h, the nanocapsules were separated and placed into PBS solution of pH 7.4. At appropriate intervals samples were ultracentrifuged, and 1 ml of the supernatant was replaced by fresh medium. The amount of BSA released from the microcapsules was evaluated by the Bradfold method [22]. The calibration curve was made using non-loaded BSA microcapsules as correction. All release tests were run in triplicate, and the error bar in the plot was the standard deviation. 2.9. Kinetics of drug release To determine the drug release mechanism and compare the release profile differences among nanocapsules, the amount of drug released vs. time was used. We analyzed the release data with the following mathematical models:
Mt =M 1 ¼ k1 t n ; where Mt/M1 is the fractional amount of the drug released at time t, n is a diffusion exponent which indicates the release mechanism, and k1 is a characteristic constant of the system. From the slope and intercept of the plot of log(Mt/M1) vs. log t, kinetic parameters n and k1 were calculated [24]. For comparison purposes, the data was also subjected to the following equation, which may be considered a simple, Higuchi equation:
Mt =M 1 ¼ k2 t 0:5 : This equation, for release data dependent on the square root of time, would give a straight-line release profile, with k2 presented as a root time dissolution rate constant. 2.10. Circular dichroism (CD) measurements CD spectroscopy (Jascow 715 spectropolarimeter) was used to measure the conformational change of the released BSA with respect to the native one. Solution of the native BSA or the released BSA was diluted to 0.1 mg ml1 and scanned over the wavelength range 190–260 nm, using 1 mm quartz cylindrical cell [25]. 2.11. Statistical analysis Analysis of variance was used to test the statistical significance of release rate and amount released for the samples. Comparison between two groups was analyzed by one-tailed Student’s t-test and multiple group comparison was performed by one-way analysis of variance using statistical software (SPSS). All data are presented as mean values with standard deviations indicated (mean ± SD). Differences were considered to be statistically significant when the p values were less than 0.05. 3. Results and discussion 3.1. Characterization of nanospheres and nanocapsules 3.1.1. Morphology The silica particles before and after being coated were characterized with scanning electron micrographs (SEM) and transmission electron micrographs (TEM). As shown in Fig. 1a, the silica nanospheres were found to be spherical, smooth and non-aggregated and the mean size was about 140 nm, which was smaller than what was measured in liquid state by photon correlation spectroscopy (PCS) (see Table 1). From Fig. 1b, small, smooth and spherical nanospheres coated with WSC and DS through layer-by-layer assembly technique became rough nanospheres. The diameter of the shells is about 210 ± 10 nm, which is larger than that of the silica particles. The increase in diameter is ascribed to a combination of WSC/DS film on the silica particles by the electrostatic interaction. Due to the different mean sizes between solid state and liquid state of silica, the nanocapsules have a cavity before removal of silica core in solid state. The results suggest that sequential WSC/DS nanocapsules were formed on the silica core. It can be seen in Fig. 1c that we obtained hollow capsules after removal of the silica core from the WSC/DS polymer capsules by HF/NH4F buffer solution. The result suggests that stable capsule films were constructed by the LBL assembly and these films were not damaged by HF etching. The hollow capsules maintained their spherical shape in the dry state. With respect to microcapsules, the numerous folds and creases of microcapsules observed are attributed to the large size and unstable construction of the shells. The shells are also flattened and some spreading is seen. However, the spherical structure of nanocapsules is retained.
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Fig. 1. The morphologies of nanoparticulate systems: (a) SEM of amino-functionalized silica spheres; (b) TEM of silica spheres coated by degradable polymer; and (c) TEM of hollow nanocapsules removed the silica core.
Table 1 Mean size and zeta potential of nanoparticles. Sample
Mean size (nm)
pI
Zeta potential (mV)
Si–NH2 Si–NH2@BSA Si–NH2@BSA/WSC Si–NH2@(WSC/DS)3 Si–NH2@(WSC/DS)5
166.5 ± 10.2 170.6 ± 5.6 167.6 ± 4.5 185.8 ± 10.5 217.6 ± 3.9
0.236 0.319 0.215 0.421 0.217
+22.21 ± 0.83 57.13 ± 1.34 +25.8 ± 2.17 62.30 ± 2.23 63.07 ± 1.29
3.1.2. Mean size and zeta potential measurement Fig. 2 demonstrates that the zeta potential was a function of the number of polyelectrolyte layers coating the silica particles. The
Fig. 2. The potential change of polyelectrolyte nanocapsules with polyelectrolyte multilayers, measured at pH 5.4. Odd layers are WSC and even layers are DS from the second layer.
zeta potential alternated between positive and negative values, indicating the successful recharging of the particle with the adsorbed polyelectrolyte multilayer upon each layer deposition. The alternating adsorption of the oppositely charged polyelectrolyte on silica nanospheres was confirmed by improved dispersivity and zeta potential measurements [11]. The magnitudes of the zeta potential give an indication of the potential stability of nanoparticle system. All the nanoparticles have negative or positive charge: they will repel each other and there is dispersion stability. This system can be stable for more than 2 months without aggregation (data not shown).
Fig. 3. FT-IR spectra of the WSC/DS, Si–NH2@WSC/DS and core removed WSC/DS nanocapsules.
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3.1.3. Fourier transform infrared (FTIR) spectroscopy From Fig. 3, from FTIR spectroscopy, we can confirm that the silica core was completely removed from SiO2–NH2/WSC/DS nanoparticles. The strong absorption of Si–O at 1100 cm1 can be clearly observed. However, when the core is removed, the absorption peak disappears, accompanied by the removal of the core. This confirms that the silica core has been completely removed.
3.3. BSA encapsulation efficiency and loading capacity of the nanocapsule
The confocal microscopy visualization was aimed to determine the localization of BSA on the surface of or inside nanocapsules. Fig. 4a and b shows the same nanocapsules but with different luminance. From Fig. 4a, the morphology of the nanocapsules can be clearly observed. As shown in Fig. 4b, fluorescent images were observed inside the nanocapsules, suggesting that the FITC-labeled BSA has been trapped into the nanocapsules. Almost every nanocapsule has a fluorescent point. The result suggests that the model drug BSA has been encapsulated inside the nanocapsules.
Encapsulation efficiency and loading capacity of nanocapsules are shown in Fig. 5a, which clearly indicates that encapsulation efficiency of these particles was affected by initial concentration of the BSA used. As the initial concentration of BSA was increasing, encapsulation efficiency decreased slowly while loading capacity increased. As shown in Fig. 5b, amino-functionalized silica has higher encapsulation efficiency than bare silica, because aminofunctionalized silica adsorbed the BSA by electrostatic interaction, while the bare silica mainly by physical adsorption through many hydrogen bonds. A possible explanation for the very high encapsulation efficiency can be proposed which is based on the role of electrostatic effect. The isoelectric point of BSA is 4.6, and the BSA molecules are negatively charged in pH 5.4 solution. As a result, strong electrostatic interactions are established between BSA and amino-functionalized silica particles, resulting in an enhanced encapsulation efficiency.
Fig. 4. Confocal laser scanning microscope of FITC-labeled BSA entrapped in the nanocapsules: (a) differential interferometer micrograph: particles shown are nanocapsules; and (b) fluorescence micrograph: almost every nanocapsules show FITC fluorescence.
Fig. 5. Effect on BSA encapsulation efficiency and loading capacity: (a) BSA encapsulation efficiency and loading capacity of nanocapsules under different initial concentrations of BSA; (b) BSA encapsulation efficiency of nanocapsules used by amino-functionalized and non-functionalized silica core.
3.2. Confocal laser scanning microscope visualization
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3.4. Biocompatibility evaluation Cell viability was used to evaluate the biocompatibility of the nanocapsules. In order to further evaluate the role of WSC, the CS/DS and SiO2–NH2@(WSC/DS)5 nanocomplexes were evaluated as control. The cells were exposed to SiO2–NH2@(WSC/DS)5, WSC/DS nanocapsules, and CS/DS nanoparticulate dispersions with various concentration and incubated for 48 h (Fig. 6). As shown in Fig. 6, WSC/DS nanocapsule system has a better biocompatibility than others. Obviously, with the SiO2–NH2@(WSC/DS)5 concentrations increasing, the cell viability decreased sharply. One possible explanation for this can be ascribed to the cell toxicity of silane coupling agent at high concentrations. The CS/DS has lower cell viability than WSC/DS. It may be due to chitosan, which, having a high molecular weight, needs to be dissolved in an acetic acid solution at the pH value of approximately 4.0. At this circumstance, the bioactivity of cells is easily damaged. The results suggested that WSC/DS may be a good carrier in biocompatibility for drug delivery and have potential for in vivo use. 3.5. In vitro release studies and kinetics of drug release The release of BSA from nanocapsules was performed in the buffer solution of pH 1.4 and 7.4, respectively, which was simulation of the pH of condition in the GI tract. From Fig. 7, at pH 1.4 (simulating the pH in the stomach), the initial burst release was decreased to lower than 10%. Similarly, at pH 7.4 (simulating the pH at the bloodstream), in the first 2 h, more than 60% of BSA was released. Finally, it is noted that the nanoparticles present pH-dependent release pattern, which can not only protect protein drug from losing in acid environment but also control drug release in GI tract [23]. At pH 1.4, the amino groups in WSC are protonated and the sulfate groups in DS are ionized (pKa < 1). The strong electrostatic attractions between WSC and DS restrain the swelling of nanocapsules, so the rate of drug release is slow. At pH 7.4, most of the amino groups of WSC are in the –NH2 form and most of the sulfate groups of DS are in the –SO3 form. Therefore, the electrostatic attractions were weakened in nanocapsules. The amount of released BSA increased due to the swelling of nanocapsule enhanced. Based on the above results, the WSC/DS nanocapsules may be a suitable vehicle for oral drug administration. The release profile of the encapsulated protein of the hollow nanocapsules can be controlled by altering the layer of the hollow
Fig. 6. Viability of cells after incubation as a function of nanoparticle concentration by MTT assay, at 37 °C for 48 h.
Fig. 7. In vitro release of BSA from nanocapsules at different pH values (WSC/DS was prepared by mixing negatively charged DS and positively charged WSC with dropping method, LBL WSC/DS was prepared through layer-by-layer method).
capsules and can be sustained for a long period (P < 0.05). From Fig. 8a, we can conclude that by altering the thickness of the nano-
Fig. 8. (a) The profile of BSA release from nanocapsules with different layer thickness; (b) the profile of BSA release profile from nanocapsules with different surface charge.
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Table 2 Drug release kinetic data for WSC/DS nanocapsules obtained from fitting drug release experimental data to Higuchi and Ritger–Peppas equation (n, diffusion exponent; k1 and k2, kinetic constants; R2, correlation coefficient). Sample
Release medium
WSC/DS (mixed) WSC/DS (mixed) (WSC/DS)4 (LBL) (WSC/DS)4 (LBL)
pH pH pH pH
1.4 7.4 1.4 7.4
Higuchi model
Ridger–Peppas model R2
n
k1
R2
1.1820 1.4644 0.7549 3.4464
0.9391 0.9591 0.8061 0.7353
0.1484 0.1701 0.1164
37.3760 39.8375 32.0306
0.9907 0.9900 0.9060 0.8970
capsules membrane we are able to control protein drugs release. It was found that the layer-by-layer assembly nanocapsules with the different outmost surface had similar release profile. When the outmost surface of nanocapsules is WSC, it released as fast as the one whose outmost surface is DS (Fig. 8b). A possible explanation for this phenomenon can be accounted for the role of electrostatic interaction. At pH 7.4, the amino groups of WSC are in the –NH2 form and DS are negatively charged. There is little electrostatic interaction between WSC and DS, and the nanocapsules readily swell and release BSA. So, when drugs are released, there is no relationship to surface charge [25]. To investigate more precisely the mechanism of the protein drug release from nanocapsules, the results were analyzed according to the Ritger–Peppas equation and Higuchi equation. Moreover, according to the Ritger–Peppas equation, R2 was higher than 0.90 and n was lower than 0.5 in all cases (Table 2). The fact that we have achieved drug release kinetics is due to Fickian diffusion. These Fickian behaviours may suggest that BSA released from nanocapsules is controlled by a drug diffusion process. The drug release from particulate systems by diffusion involves three steps. Firstly, water penetrates into particulate system, which causes swelling of the nanocapsules; secondly, the conversion of glassy polymer into rubbery matrix takes place; while the third step is the diffusion of drug from the swollen rubbery matrix. Hence, the release is slowly initial. 3.6. CD measurements CD spectroscopy was used to examine the conformation and self-association of BSA. There are three common secondary structures in BSA, namely a-helices, b-sheets, and turns. The native BSA has two extreme valleys at 208 and 222 nm [26]. Since a-helices are one of the elements of secondary structure, the quantitative analysis of the structural change of BSA could be evaluated by the content of a-helix preserved. The a-helix content of protein is estimated according to the following equation [27]:
%a-helix content ¼
hmrd 4000 33000 4000
where hmrd is the mean molar ellipticity per residue at 208 nm (deg cm2 dmol1). Usually the raw data from the experiment is expressed in terms of hd (the ellipticity in the unit of mdeg). However, it can be converted to mean molar ellipticity per residue, using the following equation [28]:
hmrd ¼
Transport mechanism
k1
hd M ; 10CLN
where M is the BSA mol. wt. (Da), C is the BSA concentration (mg ml1), L is the sample cell path length (cm), and N is the number of amino residue. As indicated by the circular dichroic spectra (Fig. 9), the calculated percentage of a-helix in the native BSA and the released BSA from WSC/DS are 58% and 57%, respectively. In other words, no significant conformation change was noted for the released BSA by using HF/NH4F solution treatment as compared with the native BSA. However, the BSA that was released from the
Diffusion Diffusion Diffusion Diffusion
controlled controlled controlled controlled
Fig. 9. Circular dichroic (CD) spectra of BSA: (A) native BSA; (B) BSA released from WSC/DS nanocapsules in pH 7.4 PBS solution; (C) BSA released from CS/DS nanocapsules in pH 7.4 solution.
CS/DS nanocapsules has obviously changed their conformation. The calculated percentage of a-helix is 37%, which may be due to the fact that CS should be dissolved in an acetic acid solution at a pH value of approximately 4.0 in the preparation of CS/DS nanocapsule process [29]. In this case, the bioactivities of protein drugs are easily damaged. 4. Conclusions In this study, we have developed a method to obtain monodisperse and degradable polyelectrolyte capsules filled with bovine serum albumin as oral protein delivery carriers. Protein-encapsulating hollow capsules were successfully prepared by a combination of layer-by-layer assembly of oppositely charged biodegradable polyelectrolytes and protein-entrapping aminofunctionalized silica templates. The system demonstrated a good capacity for the encapsulation and loading of BSA. The burst release was decreased to lower than 10% in saline within 2 h. HF/ NH4F solution do not damage the nanocapsules and the bioactivity of BSA. It can provide sustained release of entrapped protein for extended periods of time. WSC/DS nanocapsule has better biocompatibility than high molecular chitosan/dextran sulfate. Hence, this kind of novel composite nanocapsules may be a promising delivery system for ionic protein and peptide drugs. Acknowledgements This work was supported in part by the National Natural Science Foundation of China (Grant No. 20804021), the National Science Foundation of Tianjin (Grant No. 08JCYBJC00300), and the PhD Programs Foundation for New Teachers of Ministry of Education of China (Grant No. 200800551030) and the Programs Foundation for Tianjin Bureau of Public Health (Grant No. 2007kZ047).
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[14] Gao CY, Moya S, Donath E, Möhwald H. Melamine formaldehyde core decomposition as the key step controlling capsule integrity: optimizing the polyelectrolyte capsule fabrication. Macromol Chem Phys 2002;203:953–60. [15] Stöber W, Fink A, Bohn EJ. Controlled growth of monodisperse silica spheres in the micron size range. J Colloid Interface Sci 1968;26:62–9. [16] An YQ, Chen M, Xue QJ, Liu WM. Preparation and self-assembly of carboxylic acid-functionalized silica. J Colloid Interface Sci 2007;311:507–13. [17] Sarmento B, Ribeiro A, Verga F, Ferreira D. Development and characterization of new insulin containing polysaccharide nanoparticles. Colloids Surf B: Biointerf 2006;53:193–202. [18] Itoh Y, Matsusaki M, Kida T, Akashi M. Preparation of biodegradable hollow nanoparticles by silica template method. Chem Lett 2004;33:1552–3. [19] Qiao SZ, Zhang HY, Zhou XF, Budihartono S, Lu GQ. Bovine serum albumin adsorption in large pore amine functionalized mesoporous silica. Stud Surf Sci Catal 2007;165:425–8. [20] Zelikin AN, Li Q, Caruso F. Degradable polyelectrolyte capsules filled with oligonucleotide sequences. Angew Chem Int Ed 2006;45:7743–5. [21] Mahalingam V, Onclin S, Peter M, Ravoo BJ, Huskens J, Rein-Houdt DB. Directed self-assembly of functionalized silica nanoparticles on molecular printboards through multivalent supramolecular interactions. Langmuir 2004;20:11756–62. [22] Bradford MM. A rapid and sensitive method for the quantitation of microgram quantities of protein utilizing the principle of protein–dye binding. Anal Biochem 1976;72:248–54. [23] Lin YH, Mi FL, Chen CT, Chang WC, Peng SF, Liang HF, et al. Preparation and characterization of nanoparticles shelled with chitosan for oral insulin delivery. Biomacromolecules 2007;8:146–52. [24] Peppas NA. A model of dissolution-controlled solute release from porous drugdelivery polymeric systems. Pharm Acta Helv 1985;60:110–1. [25] Serizawa T, Yamaguchi M, Akashi M. Enzymatic hydrolysis of a layer-by-layer assembly prepared from chitosan and dextran sulfate. Macromolecules 2002;35:8656–8. [26] Peng ZG, Hidajat K, Uddin MS. Adsorption and desorption of lysozyme on nano-sized magnetic particles and its conformational changes. Colloids Surf B: Biointerf 2004;35:169–74. [27] Greenfield N, Fasman GD. Computed circular dichroism spectra for the evaluation of protein conformation. Biochemistry 1969;8:4108–16. [28] Xu YM, Du YM, Huang RH, Gao LP. Preparation and modification of N-(2hydroxyl) propyl-3-trimethyl ammonium chitosan chloride nanoparticle as a protein carrier. Biomaterials 2003;24:5015–22. [29] Lin YH, Chung CK, Chen CT, Liang HF, Chen SC, Sung HW. Preparation of nanoparticles composed of chitosan/poly-c-glutamic acid and evaluation of their permeability through caco-2 cells. Biomacromolecules 2005;6:1104–12.
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Acta Biomaterialia journal homepage: www.elsevier.com/locate/actabiomat
Hollow and degradable polyelectrolyte nanocapsules for protein drug delivery Shujun Shu a, Chunyang Sun a, Xinge Zhang a,*, Zhongming Wu b, Zhen Wang a, Chaoxing Li a,* a b
Key Laboratory of Functional Polymer Materials Ministry of Education, Institute of Polymer Chemistry, Nankai University, Tianjin 300071, People’s Republic of China Metabolic Diseases Hospital, Tianjin Medical University, Tianjin 300070, People’s Republic of China
a r t i c l e
i n f o
Article history: Received 2 March 2009 Received in revised form 18 May 2009 Accepted 11 June 2009 Available online 14 June 2009 Keywords: Degradable Polyelectrolyte capsules Protein Polysaccharides Layer-by-layer technique
a b s t r a c t Biodegradable hollow capsules encapsulating protein drugs were prepared via layer-by-layer assembly of water-soluble chitosan and dextran sulfate on protein-entrapping amino-functionalized silica particles and the subsequent removal of the silica. In order to enhance the encapsulated efficiency and decrease its burst release, we designed this new system to fulfill these two goals. Bovine serum albumin (BSA), which was used as model protein, was entrapped in the nanocapsules. This system demonstrated a good capacity for the encapsulation and loading of BSA. The burst release was decreased to less than 10% in phosphate-buffered saline within 2 h. No significant conformation change was noted from the released BSA in comparison with native BSA by using circular dichroism spectroscopy. Cell viability study suggested that the nanocapsules had good biocompatibility. The drug release kinetics mechanism is Fickian diffusion. These kinds of novel composite nanocapsules may offer a promising delivery system for watersoluble proteins and peptides. Ó 2009 Acta Materialia Inc. Published by Elsevier Ltd. All rights reserved.
1. Introduction An oral route is the most convenient and comfortable means of administering protein drugs. However, peptides and proteins are administered through the non-parenteral route, which has poor absorption efficiency for patients. The bioavailability of peptide after oral administration is low due to its instability under low pH circumstances and poor absorption of proteins in the gastrointestinal (GI) tract. One possible way to improve the GI uptake of peptides is to encapsulate them in nanocapsules that can protect the peptide from being degraded in the GI tract and facilitate their transportation into systemic circulation [1,2]. In general, drug loading in micro/nanoparticulate systems can be done at one of two stages: (i) during the preparation of particles (incorporation) or (ii) after the formation of particles (adsorption by physical interaction). In this system, the drug is physically embedded into the matrix or adsorbed onto the surface [3]. These two methods of drug release readily lead to an initial burst release effect. Various methods of loading have been developed to improve the encapsulation efficiency, but no research has been found that focuses on both increasing the encapsulation efficiency and decreasing the burst release effect. Microencapsulation is a promising technique for biomedical applications. However, much research has provided evidence that particles (<500 nm) can cross membranes of epithelial cells * Corresponding authors. Tel.: +86 22 2350 1645; fax: +86 22 2350 5598. E-mail addresses: [email protected] (X. Zhang), [email protected] (C. Li).
through endocytosis while larger particles (>5 lm) would be taken up via the lymphatic system [4,5]. Furthermore, the nanosized system stays in circulation for longer and therefore greatly extends the macromolecular biological activity compared to microparticles [6]. Another drawback of microcapsules is that they can be easily broken and collapsed. Numerous folds and creases observed are attributed to drying of the shells. The shells are also flattened and some spreading is noticed [7]. Chitosan (CS), a weak cationic polysaccharide produced by deacetylation of the natural polymer chitin, has many useful biological properties, such as biocompatibility, biodegradability and bioactivity. Most commercially available CS has a quite large molecular weight (mol. wt.) and needs to be dissolved in an acetic acid solution at a pH value of approximately 4.0. However, there are potential applications of CS in which a low mol. wt. would be essential. Given a low mol. wt., the polycationic characteristic of water-soluble chitosan (WSC) can be used together with a good solubility at a pH value close to physiological ranges. Loading of peptide or protein drugs at physiological pH ranges may maintain their bioactivity from decreasing [8]. WSC can improve the transportation of hydrophilic drugs across intestinal epithelium and it is believed that WSC can increase the permeability of epithelial tissues by disrupting intercellular tight junctions [9,10]. On this basis, WSC was used to prepare the nanocapsules. In this study, hollow polyelectrolyte nanocapsules prepared by layer-by-layer adsorption of polymer on nanosized sacrificial template particles hold immense potential for oral protein drug delivery. The capsules can be engineered with controlled sizes, composition and functionality, and can be loaded with model
1742-7061/$ - see front matter Ó 2009 Acta Materialia Inc. Published by Elsevier Ltd. All rights reserved. doi:10.1016/j.actbio.2009.06.020
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therapeutic such as insulin and low mol. wt. drugs. However, the encapsulated efficiency was so low following burst release [11]. In order to overcome these problems, we designed a new system to enhance the encapsulated efficiency and decrease its burst release. Bovine serum albumin (BSA), a protein which was used as model, was entrapped in the nanocapsules by both physical interaction and electrostatic attraction. After the required numbers of polyelectrolyte layers were deposited, the silica core was removed by exposure to an aqueous solution of HF/NH4F at pH 5.4. Otherwise, in this way, we can prevent the burst release by changing the membrane thickness and permeability of the nanocapsule. This result can also be avoided by washing nanocapsules with a proper solvent, which may lead to low encapsulation efficiency [12,13]. Herein, we report a polycation-free encapsulation method to obtain high concentrations of uncomplexed BSA chains confined within monodisperse, degradable nanocapsules. The encapsulation method exploits amino functionalized silica (SiO2–NH2) particles with mean size about 166 nm to adsorb BSA, followed by the assembly of WSC and dextran sulfate (DS). Removal of the template particles produces degradable and hollow nanocapsules filled with BSA [14].
2. Experimental 2.1. Materials WSC with a mol. wt. of 6 kDa and high mol. wt. chitosan (CS; mol. wt. = 50 kDa) were purchased from Yuhuan Ocean Biochemical Co. Ltd. (Zhejiang, China) and the degree of deacetylation was 93%. High mol. wt. dextran sulfate (500 kDa) was obtained from Amresco Company (USA). Tetraethoxysilane (TEOS) was purchased from Ke MiOu chemical Co. Ltd. (Tianjin, China). 3-Aminopropyltriethoxysilane (APTES, 99%) was obtained from Acros (USA). BSA, with a mol. wt. of 66.7 kDa, was purchased from Beijing Jun Yao WeiYe Biotechnology Co. Ltd. Coomassie brilliant blue G-250 was obtained from Fluka (USA). Fluorescein isothiocyanate (FITC) was purchased from Tianjin LianXing Biotechnology Co. Ltd. (Tianjin, China). All other reagents were of analytical grade without further purification before used. 2.2. Preparation of amino-functionalized silica nanospheres Amino-functionalized silica nanospheres (SiO2–NH2) were prepared using the modified Stöber method [15]. Briefly, 4 ml of tetraethoxysilane (TEOS) was added to a mixture of 3.3 ml ammonium hydroxide and 47 ml of ethanol with stirring; the reaction was continued for 24 h. The resulting silica colloidal dispersion was functionalized with APTES by quickly adding 0.3 ml of APTES with vigorous stirring. The mixture was stirred overnight. The nanosphere was purified by centrifugation and redispersion in ethanol. The former procedure was repeated three times. Meanwhile, bare silica nanospheres (SiO2) were obtained under the same condition without addition of APTES [16].
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[17], the amino groups of WSC (pKa = 6.5) are protonated with positive charge [18,19]. To form hollow capsules, the silica core was dissolved by treatment with HF/NH4F solution (2:8 M, pH 5.4) at 20 °C for 5 min, followed by multiple centrifugation–washing cycles. The washing cycles were repeated as necessary until the pH of the capsule suspension became identical to the pH of the washing buffer solution [20,21]. The schematic representation of the encapsulation of BSA within polyelectrolyte nanocapsules is shown in Scheme 1. 2.4. Characterization of nanospheres and nanocapsules Particle size was measured by photon correlation spectroscopy (PCS) at 25 °C with a detection angle of 90° and zeta potential was measured by ZetaPlus Measurement (Brookhaven, USA). Zeta potential is measured by applying an electric field across the dispersion. Particles within the dispersion with a zeta potential will migrate toward the electrode of opposite charge with a velocity proportional to the magnitude of the zeta potential. The velocity is measured using the technique of laser Doppler anemometry (LDA). The morphologies and approximate sizes of nanospheres, nanocapsules and the composite nanospheres were studied with a scanning electron microscope (X650 Hitachi) and a transmission electron microscope (Philips, Tecnai G2 T20ST). The Fourier transform infrared spectra of the nanocapsule samples were used to identify the removal of silica core. The nanocapsules were lyophilized by the freeze-dryer system (Flexi-Dry) to obtain dried capsules. These gained nanocapsules were mixed with KBr and pressed to a plate for further measurement. 2.5. Determination of BSA encapsulation efficiency and loading capacity of the nanocapsules Bound and unbound BSA was separated by ultracentrifugation of the nanosuspension at 38,000 rpm at 4 °C for 30 min (Optima LE-80k Ultracentrifuge, Beckman). The amount of free BSA in the clear supernatant was measured by a Bradfold protein assay using a UV spectrometer at 595 nm (Shimadzu UV 2550, Japan) [22]. The BSA encapsulation efficiency (EE) and loading capacity (LC) of the nanocapsules were calculated using the following equation:
total protein free protein 100% total protein total protein free protein LC% ¼ 100% nanocapsules weight
EE% ¼
All measurements were performed in triplicate and averaged.
2.3. Preparation of nanocapsules through layer-by-layer assembly A suspension of the SiO2–NH3+ particles (0.25 wt.%) was combined with a BSA solution and allowed to interact for 15 min, after which time the suspension was charged with WSC to a final concentration of 1.0 mg ml1. The WSC/DS multilayer build-up was initiated by exposing the particles with preadsorbed BSA (pI = 4.6) to a WSC solution at pH 5.4. At this pH value, DS shows negative sulfate groups (pKa < 1)
Scheme 1. Schematic representation of the encapsulation of BSA within polyelectrolyte capsules.
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2.6. Confocal laser scanning microscope visualization (CLSM) Fluorescent FITC-labeled BSA trapped in WSC/DS nanocapsules were prepared for the CLSM study (Olympus, 1 51). Synthesis of the FITC-labeled BSA was based on the reaction between the isothiocyanate group of FITC and the primary amino groups of BSA as reported in the literature [23]. Briefly, 100 mg of FITC in 150 ml of dehydrated methanol were added to 100 ml of 1% BSA solution. After 3 h of reaction in the dark at ambient conditions, FITC-labeled BSA was dialyzed at 4 °C to remove the unconjugated FITC. No fluorescence was detected in the supernatant. Then the dialyzed solution was freeze-dried for 24 h. The drug loaded WSC/DS nanocapsules were prepared as the procedure described above. Subsequently, the fluorescence images were observed using an argon laser (excitation at 488 nm, emission collected at a range of 510–540 nm). 2.7. Cell viability Cell viability was evaluated by using NIH 3T3 cell line. The cell line was cultured in Dulbecco’s modified eagles medium (DMEM) in a humidified atmosphere (5% CO2/95% O2). The cells were seeded into 96-well plates at 10,000 cells per well. The plates were then returned to the incubator and the cells were allowed to grow to confluence for 24 h. Various nanoparticles were dissolved in deionized water. After being filtered by paper filter, the resulting solution was diluted with culture medium to give a final range of concentrations from 0.1 to 1.5 mg ml1. Then the media in the wells were replaced with the pre-prepared culture medium–sample mixture (200 ll). The plates were then returned to the incubator and maintained in 5% CO2 at 37 °C for 48 h. Each sample was tested in six replicates per plate. After incubation culture medium and 20 ll of MTT solutions were used to replace the mixture in each well. The plates were then returned to the incubator and incubated for a further 4 h in 5% CO2 at 37 °C. Then, the culture medium and MTT were removed. DMSO (150 ll) was then added to each well to dissolve the formazane crystals. The plate was placed in 5% CO2 at 37 °C for 10 min and for 15 min at 6 °C before measurement. The optical density was read on a microplate reader at 490 nm. Cell viability was determined as a percentage of the negative control (untreated cells). 2.8. In vitro release studies The obtained hollow nanocapsules were redispersed in PBS solution after freeze-drying. The in vitro release profiles of BSA from nanocapsules were determined as follows: the BSA-loaded nanocapsules separated from 9 ml suspension was placed into test tubes with 6 ml of HCl solution of pH 1.4, and incubated at 37 °C under stirring at 60 rpm. After 2 h, the nanocapsules were separated and placed into PBS solution of pH 7.4. At appropriate intervals samples were ultracentrifuged, and 1 ml of the supernatant was replaced by fresh medium. The amount of BSA released from the microcapsules was evaluated by the Bradfold method [22]. The calibration curve was made using non-loaded BSA microcapsules as correction. All release tests were run in triplicate, and the error bar in the plot was the standard deviation. 2.9. Kinetics of drug release To determine the drug release mechanism and compare the release profile differences among nanocapsules, the amount of drug released vs. time was used. We analyzed the release data with the following mathematical models:
Mt =M 1 ¼ k1 t n ; where Mt/M1 is the fractional amount of the drug released at time t, n is a diffusion exponent which indicates the release mechanism, and k1 is a characteristic constant of the system. From the slope and intercept of the plot of log(Mt/M1) vs. log t, kinetic parameters n and k1 were calculated [24]. For comparison purposes, the data was also subjected to the following equation, which may be considered a simple, Higuchi equation:
Mt =M 1 ¼ k2 t 0:5 : This equation, for release data dependent on the square root of time, would give a straight-line release profile, with k2 presented as a root time dissolution rate constant. 2.10. Circular dichroism (CD) measurements CD spectroscopy (Jascow 715 spectropolarimeter) was used to measure the conformational change of the released BSA with respect to the native one. Solution of the native BSA or the released BSA was diluted to 0.1 mg ml1 and scanned over the wavelength range 190–260 nm, using 1 mm quartz cylindrical cell [25]. 2.11. Statistical analysis Analysis of variance was used to test the statistical significance of release rate and amount released for the samples. Comparison between two groups was analyzed by one-tailed Student’s t-test and multiple group comparison was performed by one-way analysis of variance using statistical software (SPSS). All data are presented as mean values with standard deviations indicated (mean ± SD). Differences were considered to be statistically significant when the p values were less than 0.05. 3. Results and discussion 3.1. Characterization of nanospheres and nanocapsules 3.1.1. Morphology The silica particles before and after being coated were characterized with scanning electron micrographs (SEM) and transmission electron micrographs (TEM). As shown in Fig. 1a, the silica nanospheres were found to be spherical, smooth and non-aggregated and the mean size was about 140 nm, which was smaller than what was measured in liquid state by photon correlation spectroscopy (PCS) (see Table 1). From Fig. 1b, small, smooth and spherical nanospheres coated with WSC and DS through layer-by-layer assembly technique became rough nanospheres. The diameter of the shells is about 210 ± 10 nm, which is larger than that of the silica particles. The increase in diameter is ascribed to a combination of WSC/DS film on the silica particles by the electrostatic interaction. Due to the different mean sizes between solid state and liquid state of silica, the nanocapsules have a cavity before removal of silica core in solid state. The results suggest that sequential WSC/DS nanocapsules were formed on the silica core. It can be seen in Fig. 1c that we obtained hollow capsules after removal of the silica core from the WSC/DS polymer capsules by HF/NH4F buffer solution. The result suggests that stable capsule films were constructed by the LBL assembly and these films were not damaged by HF etching. The hollow capsules maintained their spherical shape in the dry state. With respect to microcapsules, the numerous folds and creases of microcapsules observed are attributed to the large size and unstable construction of the shells. The shells are also flattened and some spreading is seen. However, the spherical structure of nanocapsules is retained.
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Fig. 1. The morphologies of nanoparticulate systems: (a) SEM of amino-functionalized silica spheres; (b) TEM of silica spheres coated by degradable polymer; and (c) TEM of hollow nanocapsules removed the silica core.
Table 1 Mean size and zeta potential of nanoparticles. Sample
Mean size (nm)
pI
Zeta potential (mV)
Si–NH2 Si–NH2@BSA Si–NH2@BSA/WSC Si–NH2@(WSC/DS)3 Si–NH2@(WSC/DS)5
166.5 ± 10.2 170.6 ± 5.6 167.6 ± 4.5 185.8 ± 10.5 217.6 ± 3.9
0.236 0.319 0.215 0.421 0.217
+22.21 ± 0.83 57.13 ± 1.34 +25.8 ± 2.17 62.30 ± 2.23 63.07 ± 1.29
3.1.2. Mean size and zeta potential measurement Fig. 2 demonstrates that the zeta potential was a function of the number of polyelectrolyte layers coating the silica particles. The
Fig. 2. The potential change of polyelectrolyte nanocapsules with polyelectrolyte multilayers, measured at pH 5.4. Odd layers are WSC and even layers are DS from the second layer.
zeta potential alternated between positive and negative values, indicating the successful recharging of the particle with the adsorbed polyelectrolyte multilayer upon each layer deposition. The alternating adsorption of the oppositely charged polyelectrolyte on silica nanospheres was confirmed by improved dispersivity and zeta potential measurements [11]. The magnitudes of the zeta potential give an indication of the potential stability of nanoparticle system. All the nanoparticles have negative or positive charge: they will repel each other and there is dispersion stability. This system can be stable for more than 2 months without aggregation (data not shown).
Fig. 3. FT-IR spectra of the WSC/DS, Si–NH2@WSC/DS and core removed WSC/DS nanocapsules.
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3.1.3. Fourier transform infrared (FTIR) spectroscopy From Fig. 3, from FTIR spectroscopy, we can confirm that the silica core was completely removed from SiO2–NH2/WSC/DS nanoparticles. The strong absorption of Si–O at 1100 cm1 can be clearly observed. However, when the core is removed, the absorption peak disappears, accompanied by the removal of the core. This confirms that the silica core has been completely removed.
3.3. BSA encapsulation efficiency and loading capacity of the nanocapsule
The confocal microscopy visualization was aimed to determine the localization of BSA on the surface of or inside nanocapsules. Fig. 4a and b shows the same nanocapsules but with different luminance. From Fig. 4a, the morphology of the nanocapsules can be clearly observed. As shown in Fig. 4b, fluorescent images were observed inside the nanocapsules, suggesting that the FITC-labeled BSA has been trapped into the nanocapsules. Almost every nanocapsule has a fluorescent point. The result suggests that the model drug BSA has been encapsulated inside the nanocapsules.
Encapsulation efficiency and loading capacity of nanocapsules are shown in Fig. 5a, which clearly indicates that encapsulation efficiency of these particles was affected by initial concentration of the BSA used. As the initial concentration of BSA was increasing, encapsulation efficiency decreased slowly while loading capacity increased. As shown in Fig. 5b, amino-functionalized silica has higher encapsulation efficiency than bare silica, because aminofunctionalized silica adsorbed the BSA by electrostatic interaction, while the bare silica mainly by physical adsorption through many hydrogen bonds. A possible explanation for the very high encapsulation efficiency can be proposed which is based on the role of electrostatic effect. The isoelectric point of BSA is 4.6, and the BSA molecules are negatively charged in pH 5.4 solution. As a result, strong electrostatic interactions are established between BSA and amino-functionalized silica particles, resulting in an enhanced encapsulation efficiency.
Fig. 4. Confocal laser scanning microscope of FITC-labeled BSA entrapped in the nanocapsules: (a) differential interferometer micrograph: particles shown are nanocapsules; and (b) fluorescence micrograph: almost every nanocapsules show FITC fluorescence.
Fig. 5. Effect on BSA encapsulation efficiency and loading capacity: (a) BSA encapsulation efficiency and loading capacity of nanocapsules under different initial concentrations of BSA; (b) BSA encapsulation efficiency of nanocapsules used by amino-functionalized and non-functionalized silica core.
3.2. Confocal laser scanning microscope visualization
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3.4. Biocompatibility evaluation Cell viability was used to evaluate the biocompatibility of the nanocapsules. In order to further evaluate the role of WSC, the CS/DS and SiO2–NH2@(WSC/DS)5 nanocomplexes were evaluated as control. The cells were exposed to SiO2–NH2@(WSC/DS)5, WSC/DS nanocapsules, and CS/DS nanoparticulate dispersions with various concentration and incubated for 48 h (Fig. 6). As shown in Fig. 6, WSC/DS nanocapsule system has a better biocompatibility than others. Obviously, with the SiO2–NH2@(WSC/DS)5 concentrations increasing, the cell viability decreased sharply. One possible explanation for this can be ascribed to the cell toxicity of silane coupling agent at high concentrations. The CS/DS has lower cell viability than WSC/DS. It may be due to chitosan, which, having a high molecular weight, needs to be dissolved in an acetic acid solution at the pH value of approximately 4.0. At this circumstance, the bioactivity of cells is easily damaged. The results suggested that WSC/DS may be a good carrier in biocompatibility for drug delivery and have potential for in vivo use. 3.5. In vitro release studies and kinetics of drug release The release of BSA from nanocapsules was performed in the buffer solution of pH 1.4 and 7.4, respectively, which was simulation of the pH of condition in the GI tract. From Fig. 7, at pH 1.4 (simulating the pH in the stomach), the initial burst release was decreased to lower than 10%. Similarly, at pH 7.4 (simulating the pH at the bloodstream), in the first 2 h, more than 60% of BSA was released. Finally, it is noted that the nanoparticles present pH-dependent release pattern, which can not only protect protein drug from losing in acid environment but also control drug release in GI tract [23]. At pH 1.4, the amino groups in WSC are protonated and the sulfate groups in DS are ionized (pKa < 1). The strong electrostatic attractions between WSC and DS restrain the swelling of nanocapsules, so the rate of drug release is slow. At pH 7.4, most of the amino groups of WSC are in the –NH2 form and most of the sulfate groups of DS are in the –SO3 form. Therefore, the electrostatic attractions were weakened in nanocapsules. The amount of released BSA increased due to the swelling of nanocapsule enhanced. Based on the above results, the WSC/DS nanocapsules may be a suitable vehicle for oral drug administration. The release profile of the encapsulated protein of the hollow nanocapsules can be controlled by altering the layer of the hollow
Fig. 6. Viability of cells after incubation as a function of nanoparticle concentration by MTT assay, at 37 °C for 48 h.
Fig. 7. In vitro release of BSA from nanocapsules at different pH values (WSC/DS was prepared by mixing negatively charged DS and positively charged WSC with dropping method, LBL WSC/DS was prepared through layer-by-layer method).
capsules and can be sustained for a long period (P < 0.05). From Fig. 8a, we can conclude that by altering the thickness of the nano-
Fig. 8. (a) The profile of BSA release from nanocapsules with different layer thickness; (b) the profile of BSA release profile from nanocapsules with different surface charge.
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Table 2 Drug release kinetic data for WSC/DS nanocapsules obtained from fitting drug release experimental data to Higuchi and Ritger–Peppas equation (n, diffusion exponent; k1 and k2, kinetic constants; R2, correlation coefficient). Sample
Release medium
WSC/DS (mixed) WSC/DS (mixed) (WSC/DS)4 (LBL) (WSC/DS)4 (LBL)
pH pH pH pH
1.4 7.4 1.4 7.4
Higuchi model
Ridger–Peppas model R2
n
k1
R2
1.1820 1.4644 0.7549 3.4464
0.9391 0.9591 0.8061 0.7353
0.1484 0.1701 0.1164
37.3760 39.8375 32.0306
0.9907 0.9900 0.9060 0.8970
capsules membrane we are able to control protein drugs release. It was found that the layer-by-layer assembly nanocapsules with the different outmost surface had similar release profile. When the outmost surface of nanocapsules is WSC, it released as fast as the one whose outmost surface is DS (Fig. 8b). A possible explanation for this phenomenon can be accounted for the role of electrostatic interaction. At pH 7.4, the amino groups of WSC are in the –NH2 form and DS are negatively charged. There is little electrostatic interaction between WSC and DS, and the nanocapsules readily swell and release BSA. So, when drugs are released, there is no relationship to surface charge [25]. To investigate more precisely the mechanism of the protein drug release from nanocapsules, the results were analyzed according to the Ritger–Peppas equation and Higuchi equation. Moreover, according to the Ritger–Peppas equation, R2 was higher than 0.90 and n was lower than 0.5 in all cases (Table 2). The fact that we have achieved drug release kinetics is due to Fickian diffusion. These Fickian behaviours may suggest that BSA released from nanocapsules is controlled by a drug diffusion process. The drug release from particulate systems by diffusion involves three steps. Firstly, water penetrates into particulate system, which causes swelling of the nanocapsules; secondly, the conversion of glassy polymer into rubbery matrix takes place; while the third step is the diffusion of drug from the swollen rubbery matrix. Hence, the release is slowly initial. 3.6. CD measurements CD spectroscopy was used to examine the conformation and self-association of BSA. There are three common secondary structures in BSA, namely a-helices, b-sheets, and turns. The native BSA has two extreme valleys at 208 and 222 nm [26]. Since a-helices are one of the elements of secondary structure, the quantitative analysis of the structural change of BSA could be evaluated by the content of a-helix preserved. The a-helix content of protein is estimated according to the following equation [27]:
%a-helix content ¼
hmrd 4000 33000 4000
where hmrd is the mean molar ellipticity per residue at 208 nm (deg cm2 dmol1). Usually the raw data from the experiment is expressed in terms of hd (the ellipticity in the unit of mdeg). However, it can be converted to mean molar ellipticity per residue, using the following equation [28]:
hmrd ¼
Transport mechanism
k1
hd M ; 10CLN
where M is the BSA mol. wt. (Da), C is the BSA concentration (mg ml1), L is the sample cell path length (cm), and N is the number of amino residue. As indicated by the circular dichroic spectra (Fig. 9), the calculated percentage of a-helix in the native BSA and the released BSA from WSC/DS are 58% and 57%, respectively. In other words, no significant conformation change was noted for the released BSA by using HF/NH4F solution treatment as compared with the native BSA. However, the BSA that was released from the
Diffusion Diffusion Diffusion Diffusion
controlled controlled controlled controlled
Fig. 9. Circular dichroic (CD) spectra of BSA: (A) native BSA; (B) BSA released from WSC/DS nanocapsules in pH 7.4 PBS solution; (C) BSA released from CS/DS nanocapsules in pH 7.4 solution.
CS/DS nanocapsules has obviously changed their conformation. The calculated percentage of a-helix is 37%, which may be due to the fact that CS should be dissolved in an acetic acid solution at a pH value of approximately 4.0 in the preparation of CS/DS nanocapsule process [29]. In this case, the bioactivities of protein drugs are easily damaged. 4. Conclusions In this study, we have developed a method to obtain monodisperse and degradable polyelectrolyte capsules filled with bovine serum albumin as oral protein delivery carriers. Protein-encapsulating hollow capsules were successfully prepared by a combination of layer-by-layer assembly of oppositely charged biodegradable polyelectrolytes and protein-entrapping aminofunctionalized silica templates. The system demonstrated a good capacity for the encapsulation and loading of BSA. The burst release was decreased to lower than 10% in saline within 2 h. HF/ NH4F solution do not damage the nanocapsules and the bioactivity of BSA. It can provide sustained release of entrapped protein for extended periods of time. WSC/DS nanocapsule has better biocompatibility than high molecular chitosan/dextran sulfate. Hence, this kind of novel composite nanocapsules may be a promising delivery system for ionic protein and peptide drugs. Acknowledgements This work was supported in part by the National Natural Science Foundation of China (Grant No. 20804021), the National Science Foundation of Tianjin (Grant No. 08JCYBJC00300), and the PhD Programs Foundation for New Teachers of Ministry of Education of China (Grant No. 200800551030) and the Programs Foundation for Tianjin Bureau of Public Health (Grant No. 2007kZ047).
S. Shu et al. / Acta Biomaterialia 6 (2010) 210–217
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