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Encapsulation of ascorbyl palmitate in chitosan nanoparticles by oil-in-water emulsion and ionic gelation processes
Colloids and Surfaces B: Biointerfaces 76 (2010) 292–297
Contents lists available at ScienceDirect
Colloids and Surfaces B: Biointerfaces journal homepage: www.elsevier.com/locate/colsurfb
Encapsulation of ascorbyl palmitate in chitosan nanoparticles by oil-in-water emulsion and ionic gelation processes Rangrong Yoksan a,b,∗ , Jatesuda Jirawutthiwongchai b , Kridsada Arpo b a b
Department of Packaging and Materials Technology, Faculty of Agro-Industry, Kasetsart University, Bangkok 10900, Thailand Division of Physico-Chemical Processing Technology, Faculty of Agro-Industry, Kasetsart University, Bangkok 10900, Thailand
a r t i c l e
i n f o
Article history: Received 8 July 2009 Received in revised form 13 October 2009 Accepted 11 November 2009 Available online 18 November 2009 Keywords: Chitosan Ascorbyl palmitate Encapsulation Nanoparticles Emulsion Ionic cross-linking
a b s t r a c t The encapsulation of ascorbyl palmitate (AP) in chitosan particles was carried out by droplet formation via an oil-in-water emulsion, followed by droplet solidification via ionic gelation using sodium triphosphate pentabasic (TPP) as a cross-linking agent. The success of AP encapsulation was confirmed by FT-IR, UV–vis spectrophotometry, TGA, and XRD techniques. The obtained AP-loaded chitosan particles were spherical in shape with an average diameter of 30–100 nm as observed by SEM and TEM. Loading capacity (LC) and encapsulation efficiency (EE) of AP in the nanoparticles were about 8–20% and 39–77%, respectively, when the initial AP concentration was in the range of 25–150% (w/w) of chitosan. Augmentation of the initial AP concentration led to an increase of LC and a reduction of EE. The amount of AP released from the nanoparticles in ethanol and tris buffer (pH∼8.0) increased with increasing LC and decreasing TPP concentration. © 2009 Elsevier B.V. All rights reserved.
1. Introduction For decades, ascorbyl palmitate (AP), a fat-soluble ester form of vitamin C, has been used as a source of vitamin C and as an antioxidant for foods, pharmaceuticals, and cosmetics [1,2]. Although AP is more stable than ascorbic acid, the low chemical stability and water insolubility limit its utilizations. Encapsulation potentially can protect active molecules from degradation by direct exposure to severe environments, e.g., light, oxygen, chemicals, etc. In other words, encapsulation can reduce the loss of activity of the active compounds. An encapsulant, or shell, frequently plays an important role as a carrier for delivery of the molecules to the target organs. In addition, the shell performs a release mechanism to control the diffusion level of active molecules under specific conditions, resulting in prolonged activities of these molecules. A few materials such as lipids [3–6], poly(d,l-lactide) [7], and poly(d,l-lactide-co-glycolide) [7] have been used as encapsulants for AP in the forms of microemulsions [3], liposomes [3,5,8], solid lipid nanoparticles [3], nanostructured lipid carriers [4,6] and nanoparticles [7]. In addition, the nano-level encapsulation would likely enhance the bioavailability of lipophilic compounds, thus
∗ Corresponding author at: Department of Packaging and Materials Technology, Faculty of Agro-Industry, Kasetsart University, 50 Paholyothin Rd., Ladyao, Jatujak, Bangkok 10900, Thailand. Tel.: +66 2 5625097; fax: +66 2 5625092. E-mail address: [email protected] (R. Yoksan). 0927-7765/$ – see front matter © 2009 Elsevier B.V. All rights reserved. doi:10.1016/j.colsurfb.2009.11.007
increasing the degree to which those compounds become available to the target tissues. Chitosan, a natural copolymer of N-acetyl-d-glucosamine and dglucosamine units, is one of the polysaccharides potentially most suitable as carriers for a large number of active compounds [9–16]. Amino groups along chitosan backbones allow solubility in dilute organic acid solutions, ionic cross-linking, and chemical modification of the biopolymer to form gels, beads, films, particles, etc. In addition to its biodegradability, biocompatibility and non-toxicity, chitosan has received much attention in the development of microand nanoencapsulation systems [9–16]. The formation of chitosan particles by ionic gelation or polyionic coacervation has been reported for the delivery systems of various active molecules, e.g., proteins [10,15], hydrophilic and hydrophobic drugs [9,11,13,16], and vitamins [12,14,17]. Although vitamin C has been incorporated into chitosan–tripolyphosphate particles by spray-drying [12,14,17], the encapsulation of its fat-soluble derivative, AP, by chitosan has not been reported. Spray-drying is a convenient process to produce particles encapsulating active compounds; however, particles obtained are of micro-level size. In addition, the use of high temperature (e.g., 170 ◦ C) during the microparticle preparation might induce the degradation of the active compounds. The present work thus focuses on the fabrication of AP-loaded chitosan nanoparticles by a two-step process: emulsion formation and ionic gelation. This process is unique because it not only provides very tiny particles or nanoparticles, but also avoids the use of
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high temperature. We also clarified the successful encapsulation by Fourier transform infrared (FT-IR) and ultraviolet–visible (UV–vis) spectrophotometry, thermal gravimetry analysis (TGA), and X-ray diffraction (XRD) techniques, and determined the shape and size of the particles by scanning electron microscopy (SEM) and transmission electron microscopy (TEM). In addition, the release of AP from chitosan nanoparticles was investigated. 2. Materials and methods 2.1. Materials Chitosan (deacetylation degree of 0.95 and molecular weight of ∼700,000 Da) was purchased from the Seafresh Chitosan (Lab) Co., Ltd. (Bangkok, Thailand). Ascorbyl palmitate (AP), sodium triphosphate pentabasic (TPP), Tween 60 and Span 60 were supplied by Fluka Chemika (Buchs, Switzerland). Acetic acid was obtained from Merck (Darmstadt, Germany). Soybean oil (Jade Brand) was purchased from Lam Soon Public Co., Ltd. (Bangkok, Thailand). All chemicals were used as received without further purification. 2.2. Preparation of AP-loaded chitosan nanoparticles AP-loaded chitosan nanoparticles were prepared according to a method modified from the ones described by Ko et al. [9] and Ajun et al. [16]. However, soybean oil was used instead of CH2 Cl2 for preparation of the oil phase in order to avoid toxicity of the chemical residue. Briefly, aqueous and oil phase solutions were produced. Chitosan solution (1.5% (w/v)) was prepared by agitating chitosan in an aqueous acetic acid solution (1% (v/v)) at ambient temperature (ca. 25–28 ◦ C) overnight. Tween 60 (0.45 g) was subsequently added to the solution (40 mL) and stirred at ambient temperature until the mixture was homogeneous. Soybean oil (10 mL) and Span 60 (0.05 g) were mixed at 50 ◦ C for 2 h and then cooled to ambient temperature. AP was added to the oil mixture and agitated to achieve a homogeneous oil phase solution. The oil–AP fraction (10 mL) was gradually dropped into the aqueous chitosan solution (40 mL) during homogenization at a speed of 16,000 rpm for 2 min to obtain an oil-in-water emulsion. TPP solution (0.5% (w/v), 40 mL) was then slowly dropped into the agitated emulsion. Agitation was continuously performed for 30 min. The formed particles were collected by centrifugation at 10,000 × g for 15 min at 20 ◦ C, and subsequently washed several times with Tween 60 solution (0.1% (v/v)) and water. The particles were dried at ambient temperature under reduced pressure and stored in dry condition at 25 ◦ C. Weight ratios of chitosan to AP (CTS:AP) of 1:0, 1:0.25, 1:0.50, 1:1.00 and 1:1.50 were used for the present study. 2.3. Characterization of nanoparticles FT-IR spectra were obtained by using a Thermo Nicolet Nexus 670 spectrometer (Thermo Electron Corp., Madison WI, USA) with 32 scans at a resolution of 4 cm−1 over a wavenumber range of 4000–400 cm−1 . XRD patterns were recorded over a 2 range of 5–50◦ by a JEOL JDX-3530 (JEOL Ltd., Tokyo, Japan) with a step angle of 0.04 ◦ C/min. A Mettler-Toledo TGA/SDTA 851e thermogravimetric analyzer (Columbus OH, USA) was used, with a N2 flow rate of 60 mL/min and a heating rate of 10 ◦ C/min from 30 to 600 ◦ C. The Zaverage diameter of samples was determined at 20 ◦ C by a Malvern Zetasizer (model 3600, Malvern Instruments Ltd., Worcestershire, UK) equipped with a He–Ne laser operating at 4.0 mW and 633 nm with a fixed scattering angle of 90◦ . SEM analysis of the products was carried out using a JEOL JSM LV-5600 at an operating voltage of 15 kV. Transmission electron micrographs were observed by a
Fig. 1. FT-IR spectra of (a) chitosan flakes, (b) chitosan particles and (c) AP-loaded chitosan particles with CTS to AP weight ratio of 1:1.50.
JEOL JEM-1220 at an accelerate voltage of 80 kV. UV–vis absorption spectra were recorded on a Thermo Spectronic Helios Gamma spectrophotometer (Thermo Scientific, Waltham MA, USA) with a scan speed of 60 nm/min over a wavelength range of 200–400 nm (max = 247 nm). 2.4. Determination of loading capacity and encapsulation efficiency of AP The content of AP loaded in chitosan nanoparticles was determined by TGA/DTG (derivative thermal gravimetric) technique. The amount of loaded AP per 100 g of sample—loading capacity (LC) and the amount of loaded AP per 100 g of initial AP (in feed)—encapsulation efficiency (EE) were thus calculated from Eqs. (1) and (2), respectively: %LC = %EE =
weight of loaded AP weight of sample
weight of loaded AP weight of initial AP
× 100
(1)
× 100
(2)
2.5. Study on in vitro release of AP from chitosan nanoparticles Ethanol and tris buffer (pH ∼8.4) were used as model media for an in vitro AP release study. Wet samples (10 mg) and media (1.2 mL) were placed in a microtube and incubated at ambient temperature. At sampling time, the incubated mixture was centrifuged and 100 L of supernatant was collected. Evaluation of the amount of AP released was determined using a spectrophotometer at a wavelength of 247 nm. An equal volume of fresh media was then replaced in the mixture, and the same procedure was repeated for the subsequent sampling. These in vitro release studies were performed in triplicate for each of the samples. 3. Results and discussion 3.1. Characteristics of AP-loaded chitosan nanoparticles AP-loaded chitosan particles were prepared by a two-step process. The first step involved the formation of oil droplets (including AP) by an oil-in-water emulsion. The second step was the solidification of the formed droplets by an ionic gelation of chitosan, enveloping the oil droplets, with TPP. FT-IR spectra of the obtained particles are presented in Fig. 1. In general, chitosan flakes show characteristic peaks at 3382 (–OH and –NH2 stretching), 2886–2854 (–CH stretching), 1634 (amide I), 1565 (amide II), 1062 (C–O–C) and 887 (pyranose ring) (Fig. 1a). For chitosan particles, the peak of amide II (–NH2 bending)
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Fig. 2. UV–vis absorption spectra of supernatant obtained from immersion of different samples in ethanol (0.0083 mg/mL) at ambient temperature for 150 min: (a) chitosan particles and (b)–(e) AP-loaded chitosan particles with different CTS to AP weight ratios: (b) 1:0.25, (c) 1:0.50, (d) 1:1.00 and (e) 1:1.50.
shifted from 1565 to 1550 cm−1 , and new peaks appeared around 1230–1160 cm−1 (P–O and P O), implying the complex formation via electrostatic interaction between phosphoric groups and ammonium ions (Fig. 1b). This result was similar to ones reported by Xu and Du [10], and Bhumkar and Pokharkar [18]. In comparison with the FT-IR spectrum of chitosan particles, the addition of AP resulted in a markedly increased intensity of the peak at 1732 cm−1 , indicating an increase in the content of ester groups, which might come from AP molecules (Fig. 1c). The success of AP encapsulation was also confirmed by UV–vis spectrophotometry. Since ethanol is a good solvent for AP, it was used as a release medium. After dispersing chitosan and AP-loaded chitosan particles in ethanol at ambient temperature for 150 min, the supernatants were collected and characterized by a UV–vis spectrophotometer. Chitosan (CTS) particles showed no absorption bands at wavelengths ranging from 220 to 300 nm (Fig. 2a), whereas AP-loaded chitosan particles gave a maximum absorption peak at a wavelength of ∼246–248 nm (Fig. 2b–e), corresponding to that of AP (data not shown). This implied that some content of AP was released or diffused out from chitosan particles. In other words, the encapsulation of AP in chitosan particles was accomplished. The absorption intensity at ∼246–248 nm increased with increasing initial AP content (Fig. 2b–e). This reflected that the amount of AP released from chitosan particles depended on the initial AP content (see Section 3.4, below). TGA is a simple analytical technique to study the weight change of a material as a function of temperature. Fig. 3A shows that the weight of chitosan and AP-loaded chitosan particles decreases
with increasing the temperature from 100 to 600 ◦ C. The degree of weight loss (slope) was different for different temperature ranges and samples. AP possessed only one level of weight loss (Fig. 3Aa), while chitosan and AP-loaded chitosan particles showed two (Fig. 3Ab) and three levels of weight loss (Fig. 3Ac and d), respectively. These weight losses reflected the degradation of each component in the materials. The temperatures corresponding to the maximum slopes of each weight change stage are clearly observed when the first derivative of the TGA curve with respect to temperature, the so-called derivative thermogravimetry (DTG) thermogram, is plotted (Fig. 3B). The DTG thermogram thus reflects the rate of weight change, or the change in sample weight over time [d(w)/dt] on the y-axis. The temperatures which give the highest rate of weight loss at each stage (i.e., peaks in the DTG thermogram) are usually considered as degradation temperatures (Td ) of components in the material. From DTG thermograms, chitosan particles exhibited two-step degradation, at 235 and 377 ◦ C—which might be the Td of free chitosan and of chitosan cross-linked with TPP, respectively (Fig. 3Bb). By loading of AP, the particles showed new Td at 277 ◦ C (Fig. 3Bc and d), which corresponded to the Td of AP (Fig. 3Ba). The result suggested the successful loading of AP into chitosan nanoparticles. The weight loss at this temperature was then used to determine the content of loaded AP (see Section 3.3, below). The packing structures of the obtained particles were determined by XRD technique. Generally, chitosan shows two peaks at 2 of 11◦ and 20◦ (Fig. 4a). After ionic cross-linking with TPP, a shift of peak positions, reduction of peak intensity, and broadness of peaks were observed, reflecting the destruction of the native chitosan packing structure (Fig. 4b). In addition, a new peak at 2 of 24◦ appeared for chitosan particles. These phenomena might be due to a modification in the arrangement of molecules in the crystal lattice induced by ionic interaction [18]. For AP-loaded chitosan particles, the peak intensity at 2 of 12◦ decreased significantly, while the peak at 2 of ∼20◦ was markedly sharp (Fig. 4c). This implied that the incorporation of AP resulted in a change in the chitosan–TPP packing structure. 3.2. Shape and size of AP-loaded chitosan nanoparticles The diameters of chitosan and AP-loaded chitosan particles were determined by dynamic light scattering (DLS) technique. Fig. 5 shows that chitosan particles possessed an average diameter of ∼2.5 m. A submicron-level size was obtained for the AP-loaded chitosan particles, i.e., 250–930 nm. The mean diameter of particles decreased with increasing the initial AP content. However, the size measured by the DLS technique might be the hydrodynamic
Fig. 3. (A) TGA and (B) DTG thermograms of (a) AP, (b) chitosan particles and (c)–(d) AP-loaded chitosan particles with different CTS to AP weight ratios: (c) 1:1.00 and (d) 1:1.50.
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Fig. 4. XRD patterns of (a) chitosan, (b) chitosan particles; and (c) AP-loaded chitosan particles with CTS to AP weight ratio of 1:1.00.
diameters of aggregate particles and/or hydrated individual particles. The results reflected that the aggregation and/or swelling of AP-loaded chitosan particles (in water) were lower than those of chitosan particles. This might be a result of the hydrophobicity of AP molecules entrapped inside/on the particles. In order to investigate the detailed shape and morphological information of the individual particles, electron microscopy techniques were applied. Fig. 6a shows the aggregation of chitosan particles. The individual particles were spheres with an average size of 200–400 nm. For AP-loaded chitosan particles, the aggregation of particles was also visible, and those aggregates seemed to be melting and combining with each other (Fig. 6b). This might have resulted from some AP content surrounding the particle surface. However, rounded individual particles with diameters ranging from 60 to 100 nm were observed (Fig. 6c and d). In addition, TEM micrographs confirmed the spherical shape and nanosize structure of individual chitosan and AP-loaded chitosan particles, and the aggregation of those individual particles (Fig. 7). Chitosan particle sizes were in a range of 25–50 nm (Fig. 7a), while AP-loaded chitosan particles were 30–60 nm (Fig. 7b). 3.3. Loading capacity and encapsulation efficiency of AP Although spectrophotometry and high performance liquid chromatography (HPLC) are effective techniques to determine the
Fig. 6. SEM micrographs at 15 kV of (a) chitosan particles (10,000×) and (b)–(d) APloaded chitosan particles with different CTS to AP weight ratios: (b) 1:1.00 (1000×), (c) 1:1.00 (10,000×) and (d) 1:1.50 (10,000×).
content of guest molecules loaded in the particles, the complete destruction of particles and dissolution of guest molecules in the medium should be considered. In our present research, AP-loaded chitosan nanoparticles were successfully degraded by the method reported by Wang et al. [13]; however AP did not dissolve in an aqueous HCl solution. As a result, spectrophotometry and HPLC techniques were not useful. Information obtained from TGA thermograms was used to determine the content of AP loaded in chitosan nanoparticles. Fig. 3Ab shows that chitosan particles exhibit three-step weight loss at temperatures below 150 ◦ C (loss of moisture), 180–310 ◦ C (loss of chitosan), and above 310 ◦ C (loss of chitosan cross-linked with TPP). By comparison with the TGA thermogram of chitosan particles, AP-loaded chitosan particles showed four-step weight loss (Fig. 3Ac and d). This new range of weight loss appeared at temperatures ranging from 250 to 310 ◦ C, corresponding to the Td of AP. The percent weight loss at this temperature range was thus used to compute the amount of loaded AP (see Supplementary material). The loading capacity (LC) and encapsulation efficiency (EE) were then calculated using Equations (1) and (2), respectively, and tabulated in Table 1. The LC of AP was in a range of 8–20% when the initial AP varied from 25 to 150% (w/w) of chitosan. In addition, the LC increased with an increase of the initial AP content. This result was in agreement with the findings regarding the loading of ammonium glycyrrhizinate or BSA into chitosan-TPP nanoparticles, which have been reported by Wu et al. [11] and Xu and Du Table 1 Loading capacity and encapsulation efficiency of AP in AP-loaded chitosan nanoparticles. Sample
Fig. 5. Z-average diameter of chitosan particles and AP-loaded chitosan particles with different CTS to AP weight ratios. Data are the mean ± standard deviation (n = 10).
CTS:AP
TPP (%)
1:0.25 1:0.50 1:1.00 1:1.50
0.5 0.5 0.5 0.5
a b
LCa (%)
EEb (%)
8.46 8.45 13.87 19.78
76.67 68.78 43.27 38.91
LC = (weight of loaded AP/weight of sample) × 100. EE = (weight of loaded AP/weight of AP in feed) × 100.
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Fig. 7. TEM micrographs at 80 kV of (a) chitosan particles (100,000×) and (b) AP-loaded chitosan particles with CTS to AP weight ratio of 1:1.00 (100,000×).
[10], respectively. The EE was about 39–77%. This value decreased with an increase of the initial AP content, a result which could be due to the saturation of AP loading into chitosan nanoparticles. 3.4. In vitro release of AP from chitosan nanoparticles The release profiles of AP from AP-loaded chitosan nanoparticles were investigated in vitro at ambient condition for 5 h. The media used were ethanol and tris buffer (pH∼8.0) because AP dissolved well in both solvents. The solubility of AP was restricted in phosphate-buffered saline (PBS) (pH∼7.4) and citric acid/trisodium citrate buffers (pH∼3.0); as a result, the amount of AP could not be determined by a spectrophotometer. The amount of AP released at different times was measured by a spectrophotometer at a wavelength of ∼247 nm. The release profiles of AP in ethanol (Fig. 8) and tris buffer (Fig. 9) were similar. There were two stages of AP release, depending on the release rate (the slope of the release profile). For the initial stage (i.e., the first hour for ethanol and the first 20 min for tris buffer), the release rate
Fig. 8. In vitro release profiles of AP in ethanol at ambient temperature from APloaded chitosan particles prepared by using different TPP concentrations: (a)–(d) 0.5%, (e) 2% and (f) 4%; and different CTS to AP weight ratios: (a) 1:0.25, (b) 1:0.50, (c) 1:1.00 and (d)–(f) 1:1.50. Data are the mean ± standard deviation (n = 3).
was rapid, especially in the case of tris buffer (Fig. 9). The mechanism of AP release at this stage could be explained by the diffusion of AP localized at the particle surface, which might be involved with the concentration gradient. The diffusion of AP was enhanced at high pH due to the deprotonation of chitosan; as a result, the electrostatic interaction between ammonium ions on chitosan chains and phosphoric groups of TPP molecules weakened or disappeared, and AP was eventually released very quickly (Scheme 1). In the second stage, the release rate was relatively slow, i.e., almost zero (Figs. 8 and 9). This might be due to the inability of ethanol and tris buffer to break or destroy the nanoparticles, resulting in no additional release of AP at this stage. The release of AP in ethanol reached a plateau within 1 h, while reaching a plateau in tris buffer required a shorter time (20 min). This might be a result of the weakening of electrostatic interaction between the cationic material and anionic TPP, which caused both faster release and shorter release periods, as mentioned above. However, the amount of AP released in ethanol was higher than that in tris buffer, which might be due to the superior solubility of AP in ethanol.
Fig. 9. In vitro release profiles of AP in tris buffer at ambient temperature from AP-loaded chitosan nanoparticles prepared by using CTS to AP weight ratio of 1:1.50, and different TPP concentrations: (a) 0.5%, (b) 2% and (c) 4%. Data are the mean ± standard deviation (n = 3).
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Scheme 1. Schematic drawing representing the loss of cross-linked structure via electrostatic interaction between ammonium ions on chitosan chains and phosphoric groups of TPP molecules (a) due to the deprotonation of chitosan in tris buffer (pH ∼ 8) (b).
The amount of AP released in ethanol was influenced by the amount of AP entrapped. The high LC provided a fast release rate and a concomitantly high amount of released AP (Fig. 8a-d). This might be explained by a wider AP concentration gap between the polymeric particles and the release medium, which caused a higher diffusion rate [10]. The concentration of TPP also affected the amount of released AP. AP was more easily released when TPP concentration was low (Figs. 8d–f and 9). This could be a result of low density structure, which is in agreement with previous studies [9,14,19]. 4. Conclusion Ascorbyl palmitate (AP)-loaded chitosan nanoparticles were prepared by droplet formation via an oil-in-water emulsion, followed by droplet solidification via ionic gelation using tripolyphosphate (TPP) as a cross-linking agent. The obtained APloaded chitosan nanoparticles were spherical, with an average size of 60–100 nm as observed by SEM, and 30–60 nm by TEM. The loading capacity (LC) and encapsulation efficiency (EE) of AP in the nanoparticles was about 8–20% and 39–77%, respectively, when the weight ratio of AP to chitosan was in the range of 0.25–1.50. By increasing the weight ratio of AP to chitosan, LC increased, while EE decreased. The release of AP in ethanol or tris buffer (pH∼8.0) via the diffusion mechanism was completed within 1 h (ethanol) and 20 min (tris buffer). The amount of released AP increased with increasing LC and with decreasing TPP concentration. Acknowledgements This work has been supported by the Thailand Research Fund (IRPUS/I250B02002); the Faculty of Agro-Industry, Kasetsart Uni-
versity, Thailand (AI-01-50); and the International Laboratories Corp., Ltd., Thailand. Appendix A. Supplementary data Supplementary data associated with this article can be found, in the online version, at doi:10.1016/j.colsurfb.2009.11.007. References [1] E. DeRitter, Science 113 (1951) 628–631. [2] R. Austria, A. Semenzato, A. Bettero, J. Pharm. Biomed. Anal. 15 (1997) 795– 801. [3] J. Kristl, B. Volk, M.G. Perlin, M.S. Entjurc, P. Jurkovic, Eur. J. Pharm. Sci. 19 (2003) 181–189. [4] V. Teeranachaideekul, R.H. Muller, V.B. Junyaprasert, Int. J. Pharm. 340 (2007) 198–206. [5] S. Lee, J. Lee, Y.W. Choi, Biol. Pharm. Bull. 30 (2007) 393–396. [6] V. Teeranachaideekul, E. Souto, R. Müller, V.B. Junyaprasert, J. Microencapsul. 25 (2008) 111–120. [7] A. Tangsumranjit, Y. Pellequer, H. Lboutounne, Y.C. Guillaume, A. Lamprecht, J. Millet, J. Drug Del. Sci. Technol. 16 (2006) 161–163. [8] S. Lee, J. Lee, W.C. Young, Bull. Korean Chem. Soc. 28 (2007) 99–102. [9] J.A. Ko, H.J. Park, S.J. Hwang, J.B. Park, J.S. Lee, Int. J. Pharm. 249 (2002) 165– 174. [10] Y. Xu, Y. Du, Int. J. Pharm. 250 (2003) 215–226. [11] Y. Wu, W. Yang, C. Wang, J. Hu, S. Fu, Int. J. Pharm. 295 (2005) 235–245. [12] K.G. Desai, H.J. Park, J. Microencapsul. 22 (2005) 179–192. [13] L.Y. Wang, Y.H. Gu, Q.Z. Zhou, G.H. Ma, Y.H. Wan, Z.G. Su, Colloids Surf. B 50 (2006) 126–135. [14] K.G. Desai, H.J. Park, J. Microencapsul. 23 (2006) 91–103. [15] Q. Gan, T. Wang, Colloids Surf. B 59 (2007) 24–34. [16] W. Ajun, S. Yan, G. Li, L. Huili, Carbohydr. Polym. 75 (2009) 566–574. [17] K.G. Desai, C. Lui, H.J. Park, J. Microencapsul. 23 (2006) 79–90. [18] D.R. Bhumkarl, V.B. Pokharkar1, AAPS Pharm. Sci. Tech. 7 (2) (2006) E1–E6. [19] D.V. Ratnam, D.D. Ankola, V. Bhardwaj, D.K. Sahana, M.N.V. Ravi Kumar, J. Controlled Release 113 (2006) 189–207.
Contents lists available at ScienceDirect
Colloids and Surfaces B: Biointerfaces journal homepage: www.elsevier.com/locate/colsurfb
Encapsulation of ascorbyl palmitate in chitosan nanoparticles by oil-in-water emulsion and ionic gelation processes Rangrong Yoksan a,b,∗ , Jatesuda Jirawutthiwongchai b , Kridsada Arpo b a b
Department of Packaging and Materials Technology, Faculty of Agro-Industry, Kasetsart University, Bangkok 10900, Thailand Division of Physico-Chemical Processing Technology, Faculty of Agro-Industry, Kasetsart University, Bangkok 10900, Thailand
a r t i c l e
i n f o
Article history: Received 8 July 2009 Received in revised form 13 October 2009 Accepted 11 November 2009 Available online 18 November 2009 Keywords: Chitosan Ascorbyl palmitate Encapsulation Nanoparticles Emulsion Ionic cross-linking
a b s t r a c t The encapsulation of ascorbyl palmitate (AP) in chitosan particles was carried out by droplet formation via an oil-in-water emulsion, followed by droplet solidification via ionic gelation using sodium triphosphate pentabasic (TPP) as a cross-linking agent. The success of AP encapsulation was confirmed by FT-IR, UV–vis spectrophotometry, TGA, and XRD techniques. The obtained AP-loaded chitosan particles were spherical in shape with an average diameter of 30–100 nm as observed by SEM and TEM. Loading capacity (LC) and encapsulation efficiency (EE) of AP in the nanoparticles were about 8–20% and 39–77%, respectively, when the initial AP concentration was in the range of 25–150% (w/w) of chitosan. Augmentation of the initial AP concentration led to an increase of LC and a reduction of EE. The amount of AP released from the nanoparticles in ethanol and tris buffer (pH∼8.0) increased with increasing LC and decreasing TPP concentration. © 2009 Elsevier B.V. All rights reserved.
1. Introduction For decades, ascorbyl palmitate (AP), a fat-soluble ester form of vitamin C, has been used as a source of vitamin C and as an antioxidant for foods, pharmaceuticals, and cosmetics [1,2]. Although AP is more stable than ascorbic acid, the low chemical stability and water insolubility limit its utilizations. Encapsulation potentially can protect active molecules from degradation by direct exposure to severe environments, e.g., light, oxygen, chemicals, etc. In other words, encapsulation can reduce the loss of activity of the active compounds. An encapsulant, or shell, frequently plays an important role as a carrier for delivery of the molecules to the target organs. In addition, the shell performs a release mechanism to control the diffusion level of active molecules under specific conditions, resulting in prolonged activities of these molecules. A few materials such as lipids [3–6], poly(d,l-lactide) [7], and poly(d,l-lactide-co-glycolide) [7] have been used as encapsulants for AP in the forms of microemulsions [3], liposomes [3,5,8], solid lipid nanoparticles [3], nanostructured lipid carriers [4,6] and nanoparticles [7]. In addition, the nano-level encapsulation would likely enhance the bioavailability of lipophilic compounds, thus
∗ Corresponding author at: Department of Packaging and Materials Technology, Faculty of Agro-Industry, Kasetsart University, 50 Paholyothin Rd., Ladyao, Jatujak, Bangkok 10900, Thailand. Tel.: +66 2 5625097; fax: +66 2 5625092. E-mail address: [email protected] (R. Yoksan). 0927-7765/$ – see front matter © 2009 Elsevier B.V. All rights reserved. doi:10.1016/j.colsurfb.2009.11.007
increasing the degree to which those compounds become available to the target tissues. Chitosan, a natural copolymer of N-acetyl-d-glucosamine and dglucosamine units, is one of the polysaccharides potentially most suitable as carriers for a large number of active compounds [9–16]. Amino groups along chitosan backbones allow solubility in dilute organic acid solutions, ionic cross-linking, and chemical modification of the biopolymer to form gels, beads, films, particles, etc. In addition to its biodegradability, biocompatibility and non-toxicity, chitosan has received much attention in the development of microand nanoencapsulation systems [9–16]. The formation of chitosan particles by ionic gelation or polyionic coacervation has been reported for the delivery systems of various active molecules, e.g., proteins [10,15], hydrophilic and hydrophobic drugs [9,11,13,16], and vitamins [12,14,17]. Although vitamin C has been incorporated into chitosan–tripolyphosphate particles by spray-drying [12,14,17], the encapsulation of its fat-soluble derivative, AP, by chitosan has not been reported. Spray-drying is a convenient process to produce particles encapsulating active compounds; however, particles obtained are of micro-level size. In addition, the use of high temperature (e.g., 170 ◦ C) during the microparticle preparation might induce the degradation of the active compounds. The present work thus focuses on the fabrication of AP-loaded chitosan nanoparticles by a two-step process: emulsion formation and ionic gelation. This process is unique because it not only provides very tiny particles or nanoparticles, but also avoids the use of
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high temperature. We also clarified the successful encapsulation by Fourier transform infrared (FT-IR) and ultraviolet–visible (UV–vis) spectrophotometry, thermal gravimetry analysis (TGA), and X-ray diffraction (XRD) techniques, and determined the shape and size of the particles by scanning electron microscopy (SEM) and transmission electron microscopy (TEM). In addition, the release of AP from chitosan nanoparticles was investigated. 2. Materials and methods 2.1. Materials Chitosan (deacetylation degree of 0.95 and molecular weight of ∼700,000 Da) was purchased from the Seafresh Chitosan (Lab) Co., Ltd. (Bangkok, Thailand). Ascorbyl palmitate (AP), sodium triphosphate pentabasic (TPP), Tween 60 and Span 60 were supplied by Fluka Chemika (Buchs, Switzerland). Acetic acid was obtained from Merck (Darmstadt, Germany). Soybean oil (Jade Brand) was purchased from Lam Soon Public Co., Ltd. (Bangkok, Thailand). All chemicals were used as received without further purification. 2.2. Preparation of AP-loaded chitosan nanoparticles AP-loaded chitosan nanoparticles were prepared according to a method modified from the ones described by Ko et al. [9] and Ajun et al. [16]. However, soybean oil was used instead of CH2 Cl2 for preparation of the oil phase in order to avoid toxicity of the chemical residue. Briefly, aqueous and oil phase solutions were produced. Chitosan solution (1.5% (w/v)) was prepared by agitating chitosan in an aqueous acetic acid solution (1% (v/v)) at ambient temperature (ca. 25–28 ◦ C) overnight. Tween 60 (0.45 g) was subsequently added to the solution (40 mL) and stirred at ambient temperature until the mixture was homogeneous. Soybean oil (10 mL) and Span 60 (0.05 g) were mixed at 50 ◦ C for 2 h and then cooled to ambient temperature. AP was added to the oil mixture and agitated to achieve a homogeneous oil phase solution. The oil–AP fraction (10 mL) was gradually dropped into the aqueous chitosan solution (40 mL) during homogenization at a speed of 16,000 rpm for 2 min to obtain an oil-in-water emulsion. TPP solution (0.5% (w/v), 40 mL) was then slowly dropped into the agitated emulsion. Agitation was continuously performed for 30 min. The formed particles were collected by centrifugation at 10,000 × g for 15 min at 20 ◦ C, and subsequently washed several times with Tween 60 solution (0.1% (v/v)) and water. The particles were dried at ambient temperature under reduced pressure and stored in dry condition at 25 ◦ C. Weight ratios of chitosan to AP (CTS:AP) of 1:0, 1:0.25, 1:0.50, 1:1.00 and 1:1.50 were used for the present study. 2.3. Characterization of nanoparticles FT-IR spectra were obtained by using a Thermo Nicolet Nexus 670 spectrometer (Thermo Electron Corp., Madison WI, USA) with 32 scans at a resolution of 4 cm−1 over a wavenumber range of 4000–400 cm−1 . XRD patterns were recorded over a 2 range of 5–50◦ by a JEOL JDX-3530 (JEOL Ltd., Tokyo, Japan) with a step angle of 0.04 ◦ C/min. A Mettler-Toledo TGA/SDTA 851e thermogravimetric analyzer (Columbus OH, USA) was used, with a N2 flow rate of 60 mL/min and a heating rate of 10 ◦ C/min from 30 to 600 ◦ C. The Zaverage diameter of samples was determined at 20 ◦ C by a Malvern Zetasizer (model 3600, Malvern Instruments Ltd., Worcestershire, UK) equipped with a He–Ne laser operating at 4.0 mW and 633 nm with a fixed scattering angle of 90◦ . SEM analysis of the products was carried out using a JEOL JSM LV-5600 at an operating voltage of 15 kV. Transmission electron micrographs were observed by a
Fig. 1. FT-IR spectra of (a) chitosan flakes, (b) chitosan particles and (c) AP-loaded chitosan particles with CTS to AP weight ratio of 1:1.50.
JEOL JEM-1220 at an accelerate voltage of 80 kV. UV–vis absorption spectra were recorded on a Thermo Spectronic Helios Gamma spectrophotometer (Thermo Scientific, Waltham MA, USA) with a scan speed of 60 nm/min over a wavelength range of 200–400 nm (max = 247 nm). 2.4. Determination of loading capacity and encapsulation efficiency of AP The content of AP loaded in chitosan nanoparticles was determined by TGA/DTG (derivative thermal gravimetric) technique. The amount of loaded AP per 100 g of sample—loading capacity (LC) and the amount of loaded AP per 100 g of initial AP (in feed)—encapsulation efficiency (EE) were thus calculated from Eqs. (1) and (2), respectively: %LC = %EE =
weight of loaded AP weight of sample
weight of loaded AP weight of initial AP
× 100
(1)
× 100
(2)
2.5. Study on in vitro release of AP from chitosan nanoparticles Ethanol and tris buffer (pH ∼8.4) were used as model media for an in vitro AP release study. Wet samples (10 mg) and media (1.2 mL) were placed in a microtube and incubated at ambient temperature. At sampling time, the incubated mixture was centrifuged and 100 L of supernatant was collected. Evaluation of the amount of AP released was determined using a spectrophotometer at a wavelength of 247 nm. An equal volume of fresh media was then replaced in the mixture, and the same procedure was repeated for the subsequent sampling. These in vitro release studies were performed in triplicate for each of the samples. 3. Results and discussion 3.1. Characteristics of AP-loaded chitosan nanoparticles AP-loaded chitosan particles were prepared by a two-step process. The first step involved the formation of oil droplets (including AP) by an oil-in-water emulsion. The second step was the solidification of the formed droplets by an ionic gelation of chitosan, enveloping the oil droplets, with TPP. FT-IR spectra of the obtained particles are presented in Fig. 1. In general, chitosan flakes show characteristic peaks at 3382 (–OH and –NH2 stretching), 2886–2854 (–CH stretching), 1634 (amide I), 1565 (amide II), 1062 (C–O–C) and 887 (pyranose ring) (Fig. 1a). For chitosan particles, the peak of amide II (–NH2 bending)
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Fig. 2. UV–vis absorption spectra of supernatant obtained from immersion of different samples in ethanol (0.0083 mg/mL) at ambient temperature for 150 min: (a) chitosan particles and (b)–(e) AP-loaded chitosan particles with different CTS to AP weight ratios: (b) 1:0.25, (c) 1:0.50, (d) 1:1.00 and (e) 1:1.50.
shifted from 1565 to 1550 cm−1 , and new peaks appeared around 1230–1160 cm−1 (P–O and P O), implying the complex formation via electrostatic interaction between phosphoric groups and ammonium ions (Fig. 1b). This result was similar to ones reported by Xu and Du [10], and Bhumkar and Pokharkar [18]. In comparison with the FT-IR spectrum of chitosan particles, the addition of AP resulted in a markedly increased intensity of the peak at 1732 cm−1 , indicating an increase in the content of ester groups, which might come from AP molecules (Fig. 1c). The success of AP encapsulation was also confirmed by UV–vis spectrophotometry. Since ethanol is a good solvent for AP, it was used as a release medium. After dispersing chitosan and AP-loaded chitosan particles in ethanol at ambient temperature for 150 min, the supernatants were collected and characterized by a UV–vis spectrophotometer. Chitosan (CTS) particles showed no absorption bands at wavelengths ranging from 220 to 300 nm (Fig. 2a), whereas AP-loaded chitosan particles gave a maximum absorption peak at a wavelength of ∼246–248 nm (Fig. 2b–e), corresponding to that of AP (data not shown). This implied that some content of AP was released or diffused out from chitosan particles. In other words, the encapsulation of AP in chitosan particles was accomplished. The absorption intensity at ∼246–248 nm increased with increasing initial AP content (Fig. 2b–e). This reflected that the amount of AP released from chitosan particles depended on the initial AP content (see Section 3.4, below). TGA is a simple analytical technique to study the weight change of a material as a function of temperature. Fig. 3A shows that the weight of chitosan and AP-loaded chitosan particles decreases
with increasing the temperature from 100 to 600 ◦ C. The degree of weight loss (slope) was different for different temperature ranges and samples. AP possessed only one level of weight loss (Fig. 3Aa), while chitosan and AP-loaded chitosan particles showed two (Fig. 3Ab) and three levels of weight loss (Fig. 3Ac and d), respectively. These weight losses reflected the degradation of each component in the materials. The temperatures corresponding to the maximum slopes of each weight change stage are clearly observed when the first derivative of the TGA curve with respect to temperature, the so-called derivative thermogravimetry (DTG) thermogram, is plotted (Fig. 3B). The DTG thermogram thus reflects the rate of weight change, or the change in sample weight over time [d(w)/dt] on the y-axis. The temperatures which give the highest rate of weight loss at each stage (i.e., peaks in the DTG thermogram) are usually considered as degradation temperatures (Td ) of components in the material. From DTG thermograms, chitosan particles exhibited two-step degradation, at 235 and 377 ◦ C—which might be the Td of free chitosan and of chitosan cross-linked with TPP, respectively (Fig. 3Bb). By loading of AP, the particles showed new Td at 277 ◦ C (Fig. 3Bc and d), which corresponded to the Td of AP (Fig. 3Ba). The result suggested the successful loading of AP into chitosan nanoparticles. The weight loss at this temperature was then used to determine the content of loaded AP (see Section 3.3, below). The packing structures of the obtained particles were determined by XRD technique. Generally, chitosan shows two peaks at 2 of 11◦ and 20◦ (Fig. 4a). After ionic cross-linking with TPP, a shift of peak positions, reduction of peak intensity, and broadness of peaks were observed, reflecting the destruction of the native chitosan packing structure (Fig. 4b). In addition, a new peak at 2 of 24◦ appeared for chitosan particles. These phenomena might be due to a modification in the arrangement of molecules in the crystal lattice induced by ionic interaction [18]. For AP-loaded chitosan particles, the peak intensity at 2 of 12◦ decreased significantly, while the peak at 2 of ∼20◦ was markedly sharp (Fig. 4c). This implied that the incorporation of AP resulted in a change in the chitosan–TPP packing structure. 3.2. Shape and size of AP-loaded chitosan nanoparticles The diameters of chitosan and AP-loaded chitosan particles were determined by dynamic light scattering (DLS) technique. Fig. 5 shows that chitosan particles possessed an average diameter of ∼2.5 m. A submicron-level size was obtained for the AP-loaded chitosan particles, i.e., 250–930 nm. The mean diameter of particles decreased with increasing the initial AP content. However, the size measured by the DLS technique might be the hydrodynamic
Fig. 3. (A) TGA and (B) DTG thermograms of (a) AP, (b) chitosan particles and (c)–(d) AP-loaded chitosan particles with different CTS to AP weight ratios: (c) 1:1.00 and (d) 1:1.50.
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Fig. 4. XRD patterns of (a) chitosan, (b) chitosan particles; and (c) AP-loaded chitosan particles with CTS to AP weight ratio of 1:1.00.
diameters of aggregate particles and/or hydrated individual particles. The results reflected that the aggregation and/or swelling of AP-loaded chitosan particles (in water) were lower than those of chitosan particles. This might be a result of the hydrophobicity of AP molecules entrapped inside/on the particles. In order to investigate the detailed shape and morphological information of the individual particles, electron microscopy techniques were applied. Fig. 6a shows the aggregation of chitosan particles. The individual particles were spheres with an average size of 200–400 nm. For AP-loaded chitosan particles, the aggregation of particles was also visible, and those aggregates seemed to be melting and combining with each other (Fig. 6b). This might have resulted from some AP content surrounding the particle surface. However, rounded individual particles with diameters ranging from 60 to 100 nm were observed (Fig. 6c and d). In addition, TEM micrographs confirmed the spherical shape and nanosize structure of individual chitosan and AP-loaded chitosan particles, and the aggregation of those individual particles (Fig. 7). Chitosan particle sizes were in a range of 25–50 nm (Fig. 7a), while AP-loaded chitosan particles were 30–60 nm (Fig. 7b). 3.3. Loading capacity and encapsulation efficiency of AP Although spectrophotometry and high performance liquid chromatography (HPLC) are effective techniques to determine the
Fig. 6. SEM micrographs at 15 kV of (a) chitosan particles (10,000×) and (b)–(d) APloaded chitosan particles with different CTS to AP weight ratios: (b) 1:1.00 (1000×), (c) 1:1.00 (10,000×) and (d) 1:1.50 (10,000×).
content of guest molecules loaded in the particles, the complete destruction of particles and dissolution of guest molecules in the medium should be considered. In our present research, AP-loaded chitosan nanoparticles were successfully degraded by the method reported by Wang et al. [13]; however AP did not dissolve in an aqueous HCl solution. As a result, spectrophotometry and HPLC techniques were not useful. Information obtained from TGA thermograms was used to determine the content of AP loaded in chitosan nanoparticles. Fig. 3Ab shows that chitosan particles exhibit three-step weight loss at temperatures below 150 ◦ C (loss of moisture), 180–310 ◦ C (loss of chitosan), and above 310 ◦ C (loss of chitosan cross-linked with TPP). By comparison with the TGA thermogram of chitosan particles, AP-loaded chitosan particles showed four-step weight loss (Fig. 3Ac and d). This new range of weight loss appeared at temperatures ranging from 250 to 310 ◦ C, corresponding to the Td of AP. The percent weight loss at this temperature range was thus used to compute the amount of loaded AP (see Supplementary material). The loading capacity (LC) and encapsulation efficiency (EE) were then calculated using Equations (1) and (2), respectively, and tabulated in Table 1. The LC of AP was in a range of 8–20% when the initial AP varied from 25 to 150% (w/w) of chitosan. In addition, the LC increased with an increase of the initial AP content. This result was in agreement with the findings regarding the loading of ammonium glycyrrhizinate or BSA into chitosan-TPP nanoparticles, which have been reported by Wu et al. [11] and Xu and Du Table 1 Loading capacity and encapsulation efficiency of AP in AP-loaded chitosan nanoparticles. Sample
Fig. 5. Z-average diameter of chitosan particles and AP-loaded chitosan particles with different CTS to AP weight ratios. Data are the mean ± standard deviation (n = 10).
CTS:AP
TPP (%)
1:0.25 1:0.50 1:1.00 1:1.50
0.5 0.5 0.5 0.5
a b
LCa (%)
EEb (%)
8.46 8.45 13.87 19.78
76.67 68.78 43.27 38.91
LC = (weight of loaded AP/weight of sample) × 100. EE = (weight of loaded AP/weight of AP in feed) × 100.
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Fig. 7. TEM micrographs at 80 kV of (a) chitosan particles (100,000×) and (b) AP-loaded chitosan particles with CTS to AP weight ratio of 1:1.00 (100,000×).
[10], respectively. The EE was about 39–77%. This value decreased with an increase of the initial AP content, a result which could be due to the saturation of AP loading into chitosan nanoparticles. 3.4. In vitro release of AP from chitosan nanoparticles The release profiles of AP from AP-loaded chitosan nanoparticles were investigated in vitro at ambient condition for 5 h. The media used were ethanol and tris buffer (pH∼8.0) because AP dissolved well in both solvents. The solubility of AP was restricted in phosphate-buffered saline (PBS) (pH∼7.4) and citric acid/trisodium citrate buffers (pH∼3.0); as a result, the amount of AP could not be determined by a spectrophotometer. The amount of AP released at different times was measured by a spectrophotometer at a wavelength of ∼247 nm. The release profiles of AP in ethanol (Fig. 8) and tris buffer (Fig. 9) were similar. There were two stages of AP release, depending on the release rate (the slope of the release profile). For the initial stage (i.e., the first hour for ethanol and the first 20 min for tris buffer), the release rate
Fig. 8. In vitro release profiles of AP in ethanol at ambient temperature from APloaded chitosan particles prepared by using different TPP concentrations: (a)–(d) 0.5%, (e) 2% and (f) 4%; and different CTS to AP weight ratios: (a) 1:0.25, (b) 1:0.50, (c) 1:1.00 and (d)–(f) 1:1.50. Data are the mean ± standard deviation (n = 3).
was rapid, especially in the case of tris buffer (Fig. 9). The mechanism of AP release at this stage could be explained by the diffusion of AP localized at the particle surface, which might be involved with the concentration gradient. The diffusion of AP was enhanced at high pH due to the deprotonation of chitosan; as a result, the electrostatic interaction between ammonium ions on chitosan chains and phosphoric groups of TPP molecules weakened or disappeared, and AP was eventually released very quickly (Scheme 1). In the second stage, the release rate was relatively slow, i.e., almost zero (Figs. 8 and 9). This might be due to the inability of ethanol and tris buffer to break or destroy the nanoparticles, resulting in no additional release of AP at this stage. The release of AP in ethanol reached a plateau within 1 h, while reaching a plateau in tris buffer required a shorter time (20 min). This might be a result of the weakening of electrostatic interaction between the cationic material and anionic TPP, which caused both faster release and shorter release periods, as mentioned above. However, the amount of AP released in ethanol was higher than that in tris buffer, which might be due to the superior solubility of AP in ethanol.
Fig. 9. In vitro release profiles of AP in tris buffer at ambient temperature from AP-loaded chitosan nanoparticles prepared by using CTS to AP weight ratio of 1:1.50, and different TPP concentrations: (a) 0.5%, (b) 2% and (c) 4%. Data are the mean ± standard deviation (n = 3).
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Scheme 1. Schematic drawing representing the loss of cross-linked structure via electrostatic interaction between ammonium ions on chitosan chains and phosphoric groups of TPP molecules (a) due to the deprotonation of chitosan in tris buffer (pH ∼ 8) (b).
The amount of AP released in ethanol was influenced by the amount of AP entrapped. The high LC provided a fast release rate and a concomitantly high amount of released AP (Fig. 8a-d). This might be explained by a wider AP concentration gap between the polymeric particles and the release medium, which caused a higher diffusion rate [10]. The concentration of TPP also affected the amount of released AP. AP was more easily released when TPP concentration was low (Figs. 8d–f and 9). This could be a result of low density structure, which is in agreement with previous studies [9,14,19]. 4. Conclusion Ascorbyl palmitate (AP)-loaded chitosan nanoparticles were prepared by droplet formation via an oil-in-water emulsion, followed by droplet solidification via ionic gelation using tripolyphosphate (TPP) as a cross-linking agent. The obtained APloaded chitosan nanoparticles were spherical, with an average size of 60–100 nm as observed by SEM, and 30–60 nm by TEM. The loading capacity (LC) and encapsulation efficiency (EE) of AP in the nanoparticles was about 8–20% and 39–77%, respectively, when the weight ratio of AP to chitosan was in the range of 0.25–1.50. By increasing the weight ratio of AP to chitosan, LC increased, while EE decreased. The release of AP in ethanol or tris buffer (pH∼8.0) via the diffusion mechanism was completed within 1 h (ethanol) and 20 min (tris buffer). The amount of released AP increased with increasing LC and with decreasing TPP concentration. Acknowledgements This work has been supported by the Thailand Research Fund (IRPUS/I250B02002); the Faculty of Agro-Industry, Kasetsart Uni-
versity, Thailand (AI-01-50); and the International Laboratories Corp., Ltd., Thailand. Appendix A. Supplementary data Supplementary data associated with this article can be found, in the online version, at doi:10.1016/j.colsurfb.2009.11.007. References [1] E. DeRitter, Science 113 (1951) 628–631. [2] R. Austria, A. Semenzato, A. Bettero, J. Pharm. Biomed. Anal. 15 (1997) 795– 801. [3] J. Kristl, B. Volk, M.G. Perlin, M.S. Entjurc, P. Jurkovic, Eur. J. Pharm. Sci. 19 (2003) 181–189. [4] V. Teeranachaideekul, R.H. Muller, V.B. Junyaprasert, Int. J. Pharm. 340 (2007) 198–206. [5] S. Lee, J. Lee, Y.W. Choi, Biol. Pharm. Bull. 30 (2007) 393–396. [6] V. Teeranachaideekul, E. Souto, R. Müller, V.B. Junyaprasert, J. Microencapsul. 25 (2008) 111–120. [7] A. Tangsumranjit, Y. Pellequer, H. Lboutounne, Y.C. Guillaume, A. Lamprecht, J. Millet, J. Drug Del. Sci. Technol. 16 (2006) 161–163. [8] S. Lee, J. Lee, W.C. Young, Bull. Korean Chem. Soc. 28 (2007) 99–102. [9] J.A. Ko, H.J. Park, S.J. Hwang, J.B. Park, J.S. Lee, Int. J. Pharm. 249 (2002) 165– 174. [10] Y. Xu, Y. Du, Int. J. Pharm. 250 (2003) 215–226. [11] Y. Wu, W. Yang, C. Wang, J. Hu, S. Fu, Int. J. Pharm. 295 (2005) 235–245. [12] K.G. Desai, H.J. Park, J. Microencapsul. 22 (2005) 179–192. [13] L.Y. Wang, Y.H. Gu, Q.Z. Zhou, G.H. Ma, Y.H. Wan, Z.G. Su, Colloids Surf. B 50 (2006) 126–135. [14] K.G. Desai, H.J. Park, J. Microencapsul. 23 (2006) 91–103. [15] Q. Gan, T. Wang, Colloids Surf. B 59 (2007) 24–34. [16] W. Ajun, S. Yan, G. Li, L. Huili, Carbohydr. Polym. 75 (2009) 566–574. [17] K.G. Desai, C. Lui, H.J. Park, J. Microencapsul. 23 (2006) 79–90. [18] D.R. Bhumkarl, V.B. Pokharkar1, AAPS Pharm. Sci. Tech. 7 (2) (2006) E1–E6. [19] D.V. Ratnam, D.D. Ankola, V. Bhardwaj, D.K. Sahana, M.N.V. Ravi Kumar, J. Controlled Release 113 (2006) 189–207.