http://informahealthcare.com/mnc ISSN: 0265-2048 (print), 1464-5246 (electronic) J Microencapsul, 2014; 31(4): 363–372 ! 2014 Informa UK Ltd. DOI: 10.3109/02652048.2013.858792
ORIGINAL ARTICLE
Development of a novel drug delivery system: chitosan nanoparticles entrapped in alginate microparticles Ghislain Garrait1,2, Eric Beyssac1, and Muriel Subirade2
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1
Universite´ d’Auvergne, UFR Pharmacie, Equipe d’Accueil Conception, Inge´nierie et De´veloppement de l’Aliment et du Me´dicament (EA CIDAM), Clermont-Ferrand F-63001, France and 2Faculte´ des Sciences de l’Agriculture et de l’Alimentation, INAF/STELA, Universite´ Laval, Sainte-Foy, Que´bec, Canada Abstract
Keywords
A novel carrier using chitosan nanoparticles entrapped into alginate microparticles is proposed for protecting molecules of interest from degradation in the digestive tract. The effects of polymer concentration, sonication, stirring, pH, and processing conditions on the physical characteristics of the carrier were studied. FITC and RBITC were used to localise the polymers within particles using CLSM. Diffusion of amaranth red (AR) from nanoparticles was quantified during dissolution under gastric and intestinal conditions. Under optimal preparation conditions, the size distribution of nanoparticles loaded with AR was uniform (690 nm) with an encapsulation efficacy of 21.9%. Alginate microparticles (285 mm) containing a homogenous distribution of nanoparticles and polymers were obtained. At gastric pH, the carrier released less than 5% of the loaded AR and, at intestinal pH, the release was rapid and complete. The drug carriers developed shows a promising use as a vehicle suitable to protect molecules of interest after oral administration.
Alginate microparticle, chitosan nanoparticle, gastro-resistant, novel drug delivery system
Introduction The oral route is the most convenient, safe and widely accepted for administering drugs. However, acid and pepsin in the stomach and pancreatic enzymes in the small intestine can degrade pharmaceutical payloads or cause substantial losses thereof (Bernkop-Schnurch et al., 2004). In addition, many bioactive dietary compounds or drugs are absorbed poorly through the intestinal mucosa because of their physicochemical properties (i.e. high molecular mass, hydrophilic character). Carrier systems for protecting therapeutic or nutritional molecules against the harsh environment of the gastrointestinal tract (GIT) and providing efficient and site-specific delivery are therefore also receiving much attention. Among these carriers, nanoparticles present certain advantages as interaction with the biological environment, enhancement of absorption, and improvement retention time (Cheng et al., 2008). Nanoparticles are solid colloidal carriers ranging from 10 to 1000 nm in diameter and may be composed of synthetic or natural polymers. However, the preparation of synthetic polymer nanoparticles involves heat or organic solvent, which can affect bioactive compound structure or stability. In contrast, natural polymers are often water-soluble, biodegradable, biocompatible, less toxic and amenable to simple preparation techniques under mild conditions. Among the natural polymers, alginate and chitosan have been studied extensively, making them very attractive as drug delivery carriers.
Address for correspondence: Muriel Subirade, Faculte´ des Sciences de l’Agriculture et de l’Alimentation, INAF/STELA, Universite´ Laval, Pavillon Paul-Comtois, Que´bec QC G1K 7P4, Canada. Tel: +1 418 656 2131x4278. Fax: +1 418 656 2131 3353. E-mail: [email protected]
History Received 7 May 2013 Revised 16 September 2013 Accepted 15 October 2013 Published online 1 April 2014
Alginate is an anionic and water-soluble polysaccharide and can be cross-linked under mild conditions in the presence of any divalent cation (Ca2þ, Sr2þ, Cu2þ or Zn2þ) (Wee and Gombotz, 1998). Alginate gelation may be used to form particles that protect payload compounds against the gastric environment. In addition, alginate microparticles possess a bioadhesive property that allows them to stick to the intestinal mucosa and thus increase drug residence time (Chickering et al., 1997). Due to its various properties, alginate has been used widely to develop drug delivery systems especially for oral route use (Wee and Gombotz, 1998). However, significant drug leakage through pores is well known (Liu and Krishnan, 1999). Thus, alginate has been combined successfully with chitosan to form a chitosan–alginate complex with lower porosity limiting the leakage of the encapsulated molecules (Douglas and Tabrizian, 2005). Chitosan is a cationic polysaccharide obtained from the partial deacetylation of chitin. It carries a positive charge and can react with negatively charged surfaces (e.g. mucus) to prolong retention time and improve the bioavailability of the payload (Deacon et al., 2000). These properties all make this polymer suitable for use as a drug delivery carrier. Indeed, chitosan has been used widely to develop particles for controlled release of active agents via several routes of administration (e.g. oral, nasal). The main drawback of administering chitosan carriers via the oral route is the easy dissolution of the polymer matrix exposing acid-sensitive payloads to degradation (George and Abraham, 2006). Alginate interacts via anionic carboxylic groups with cationic amino groups of chitosan to form chitosan–alginate particles. This complex is stable under acidic conditions and thus protects pharmaceutical preparations from degradation (Takka and Gurel, 2010), but becomes soluble at intestinal pH, allowing rapid and extensive release of the loaded compound (Woitiski et al., 2010).
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The objective of this study was to develop a novel drug delivery system using chitosan nanoparticles entrapped into alginate microparticles. The effects of processing conditions on the physical characteristics (e.g. size, shape) of the chitosan nanoparticles and alginate microparticles were investigated. Parameters such as polymer and sodium sulphate concentrations, sonication, stirring and addition of surfactant were examined. Fluorescein 5(6)-isothiocyanate (FITC) and rhodamine B isothiocyanate (RBITC) were used to localise the polymers within the particles. Amaranth red (AR), used as a model, was loaded into the nanoparticles, and release kinetics in simulated gastric or intestinal fluids were investigated.
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Materials
Chitosan nanoparticles Preparation of chitosan nanoparticles The effects of different factors (chitosan concentration, sodium sulphate concentration, sonication and stirring) on the particle size were studied (Table 1). All experiments were performed in triplicate. Chitosan nanoparticles were prepared by the precipitation/ coaversion method described previously (Borges et al., 2005). Briefly, a solution (200 mL) of 2% (v/v) acetic acid and 1% (w/v) Tween 80 was prepared, in which low-molecular-weight chitosan powder was dissolved (Table 1). Sodium sulphate solution (3.5 mL) was then added dropwise (1 mL/min) with or without continuous sonication (Sonicator model XL 2020, 550 W, output control ‘‘1’’, Misonix, Farmingdale, NY) at 4 C for 15 min and with or without stirring (100 rpm) for 60 min at room temperature (RT). Sodium sulphate induced the production of chitosan particles by a decrease in polymer solubility inducing a precipitation. An opalescent suspension was formed spontaneously and the particles therein were recovered by centrifugation (5000 g, 15 min, RT), re-suspended and centrifuged five times in 40 mL of ultrapure water. Particles thus obtained were characterised immediately. Table 1. Preparation conditions to study the effects of chitosan and sodium sulphate concentrations, stirring and sonication on particles size distribution.
Chitosan concentration (w/v) Sodium sulphate concentration (w/v) Sonication (min) Stirring (min)/(rpm) NA, not applicable.
AR (0.01%, w/v) was added to 200 mL of acetic acid/Tween 80 solution with stirring. After 15 min, 0.25% (w/v) chitosan was dissolved therein by stirring for 1 h. An aliquot was taken to determine the total amount of AR at the beginning of the experiment. Sodium sulphate solution (3.5 mL, 10%, w/v) was then added to form chitosan nanoparticles by continuous sonication (15 min, 4 C) with stirring (100 rpm, 60 min, RT). The nanoparticles obtained were washed (to remove unloaded AR) and centrifuged (5000 g, 15 min, RT) five times with 40 mL of ultrapure water. Nanoparticle size was determined immediately and dye encapsulated in nanoparticles was quantified directly to determine AR encapsulation efficacy and content. Alginate microparticles
Low-molecular-weight chitosan with deacetylation up to high as 75% according to specifications and viscosity ranging from 20 to 300 cps (1% solution in 1% acetic acid, 25 C), acetic acid 99%, TweenÕ 80, alginic acid sodium salt (15 cps for 1% w/v aqueous solution, mannuronic/guluronic acid ratio of 0.65), calcium chloride, FITC, RBITC, enzymes and other chemicals used for dissolution experiments were all purchased from the Sigma Chemical Company (St Louis, MO). AR was obtained from Sensient Colors Canada Ltd (Kingston, ON) and sodium sulphate was obtained from Fluka Chemical Corp. (Ronkonkoma, NY). All solutions were prepared in ultrapure water. All other chemicals were of reagent grade.
Formulation
Preparation of amaranth-red-loaded chitosan nanoparticles
1 0.25; 0.5; 1; 2% 10% 15 60/100
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Preparation of alginate microparticles loaded with chitosan nanoparticles Alginate microparticles were obtained using an EncapsulatorÕ (model IER50R, Inotech, Switzerland), of which the vibrating nozzle breaks a laminar liquid jet (i.e. alginate solution containing chitosan nanoparticles) into equal-sized micro-droplets, which were collected in a solution of calcium chloride to stabilise them as microparticles. Several experimental conditions were tested in order to obtain a homogeneous suspension of nanoparticles in alginate. Nanoparticle suspensions (0.2, 0.25, 0.3, 0.35, 0.4, 0.6 and 0.8% w/v) were combined with 4% sodium alginate solution at a nanoparticle:alginate ratio of 7:1 (final alginate concentration of 0.5%), stirred for 30 min and then run through the Encapsulator (700–1200 Hz, amplitude 4, 0.6 kV, 0.6 Bar, nozzle diameter 100 or 150 mm). The droplets formed were collected in 0.1 M calcium chloride solution. A nanoparticle:alginate ratio of 4:1 was studied using the 150-mm nozzle. The microparticles formed were observed by optical microscopy and the particle diameter was determined using an image-processing software. Evaluation of AR release during alginate microparticle preparation To avoid release of AR from chitosan nanoparticles into the alginate during microparticle formation, we investigated the impact of pH and alginic acid concentration. A suspension of AR-loaded nanoparticles (0.2% w/v in ultrapure water) was added dropwise to 4% alginic acid solution (pH adjusted to 4, 5, 6, 7 or 8 with 1 M hydrochloric acid, n ¼ 2 for each pH value) with stirring until a final alginate concentration of 0.8% w/v was reached (nanoparticle:alginate ratio of 4:1). In a second experiment, alginic acid solution (4% or 3%, pH 5) was diluted to 0.5, 0.6 and 0.8% with 0.2% nanoparticle suspension (n ¼ 2 for each test). Vigorous stirring was maintained for 15 min and an aliquot (4 mL) was centrifuged (10 000 g, 10 min, RT) and AR in the supernatant was measured as absorbance at 520 nm. The solutions were run through the 150-mm nozzle and the droplets were collected in 0.1 M calcium chloride solution. The dye content of the recovered particles was assayed. Finally, in another attempt to reduce AR release during microparticle formation, chitosan nanoparticles were re-suspended at 0.2% w/v in 1% v/v Tween 80. This suspension was added dropwise to 4% alginate solution (pH 5) at a nanoparticle:alginate ratio of 7:1 (final alginate concentration 0.5% w/v). The mixture was run through the 150-mm nozzle and droplets were collected in 0.1 M calcium chloride solution.
DOI: 10.3109/02652048.2013.858792
Microparticle morphology, size and AR content were then determined. Freeze-drying
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Freshly prepared alginate microparticles entrapping AR-loaded chitosan nanoparticles were dispersed in ultrapure water containing 1% (v/v) Tween 80. The mixture was stirred for 30 min then filtered on large pore size, frozen in liquid nitrogen and lyophilised (Model Freezone 4.5, Labconco, Kansas City, MI) for 48 h. The dry particles were stored at 20 C until further use in order to avoid the degradation of alginate microparticles entrapping AR-loaded chitosan nanoparticles. To determine their morphology and size, the microparticles were dispersed in 0.9% NaCl solution (50 mL) and stirred for 1 h. Characterisation of particles Particle size analysis Measurement of size and size distribution of freshly prepared nanoparticles was based on the static light scattering using a Mastersizer 2000 (Malvern Instruments, Southborough, MA). Particles were suspended in distilled water containing 1% v/v Tween 80. All size measurements were performed at 25 C and at a 90 scattering angle with 180 s of recording. The mean hydrodynamic diameter was generated by cumulative analysis (no less than triplicate with independent particle batches). Microparticle size was also evaluated using an optical microscope (Olympus BX50WI, Olympus, Melville, NY) fitted with a digital camera (model U-TV1 X, Olympus Optical, Tokyo, Japan). Microparticles were visualised at 40 magnification and the diameter of 50 randomly selected microparticles was determined using an image processing Image-Pro PlusÕ software (Media Cybernetics, Inc., Rockville, MD). Surface particle morphology Nanoparticle morphology was examined by transmission electron microscopy (TEM) using a Jeol 1230 microscope (Tokyo, Japan). A drop of freshly prepared nanoparticles in ultrapure water was air-dried onto a copper grid. After negative staining with uranyl acetate, the sample was desiccated for at least 72 h at RT and then examined at 30 and 50 kV. Nanoparticles were visualised using different accelerating voltages in order to improve resolution and modify magnification. Microparticles were also examined at 40 and 100 using the optical microscope. AR content of particles To evaluate the AR encapsulation efficacy and content of the nanoparticles, 25 mg of nanoparticles were disrupted in 25 mL of simulated gastric fluid (SGF) (USP standard XXX, 2 g of sodium chloride, 7 mL of 37% hydrochloric acid and 1 L of ultrapure water, pH 1.2) with vigorous magnetic stirring at RT for 30 min. The resulting solution was centrifuged at 10 000 rpm for 20 min at RT and absorbance at 520 nm by the supernatant was measured using a UV-visible spectrophotometer (model 8453, HewlettPackard, Palo Alto, CA). The absorbance was compared to a standard curve prepared at the same time. AR encapsulation efficacy (EE) was obtained from the relationship EE ¼ 100A/B, where A is the AR encapsulated in the chitosan nanoparticles and B is the total amount of AR added. AR content into alginate microparticles was determined by disrupting microparticles (6 g) in 50 mL of a mixture of 0.2 M sodium bicarbonate and 0.06 M tri-basic sodium citrate at
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pH 8.0 (Li et al., 2007), centrifuging, measuring the supernatant OD at 520 nm. AR content was obtained from the following relationship: 100 amount of AR encapsulated in chitosan nanoparticlesðgÞ AR content ð%Þ ¼ amount of chitosan nanoparticlesðgÞ Polymer distribution within particles The localisation of both polymers within the particles was determined using confocal laser scanning microscopy (CLSM). The labelling protocol was based on a method previously published (Huang et al., 2002). Chitosan was labelled with FITC while alginate was labelled with RBITC as published previously (Mladenovska et al., 2007). FITC and RBITC were chosen to allow simultaneous detection in different channels. FITC-labelled chitosan nanoparticles were prepared by dissolving labelled polymer (0.25%, w/v) in acetic acid/Tween 80 solution and adding 3.5 mL of sodium sulphate solution (10%, w/v) dropwise with continuous sonication at 4 C for 15 min and with magnetic stirring (100 rpm) for 60 min at RT. Particles were recovered by centrifugation (5000 g, 15 min, RT), washed twice and used to produce alginate microparticles. Labelled microparticles were obtained using a FITC-labelled chitosan nanoparticle suspension (0.2%, w/v) stirred with RBITClabelled alginate solution (final concentration 0.8% w/v) for 30 min and run through the encapsulator fitted with the 150-mm nozzle. The droplets were collected in 0.5 M calcium chloride. The double-labelled microparticles were observed using the Olympus light microscope Fluoview FV 300 Laser Scanning Confocal Imaging System (Paris, France) equipped with an argon ion laser (EM 488 nm) and green (543 nm) and red (633 nm) helium–neon ion laser. The laser was adjusted to excitation/ emission wavelengths of 488/520 nm for FITC and 543/572 nm for RBITC. All confocal fluorescence pictures were taken with a 40 objective (oil immersion). In vitro release studies Dissolution assay in simulated gastric and intestinal fluids Dry microparticles (0.2 g) were dispersed in 50 mL of SGF with or without pepsin (3.2 g, 924 units/mg of protein) or in simulated intestinal fluid (SIF; 6.8 g of monobasic potassium phosphate dissolved in 250 mL of ultrapure water and added to 190 mL of 0.2 N sodium hydroxide and 400 mL of ultrapure water and adjusted to pH 7.4 with 0.2 N sodium hydroxide) with or without pancreatin (10 g, activity equivalent to USP specification) as described in USP standard XXX (2007) using paddle apparatus II. Agitator speed was set at 90 rpm and the temperature was maintained at 37 0.5 C. Samples were withdrawn at 15-min or 1 h intervals and particle shape was observed microscopically (40 and 100). The diameter of 50 randomly selected particles was determined using an image processing Image-Pro PlusÕ software. An equal volume of medium was added to the release mixture after each sampling to maintain a constant volume of 50 mL. Dissolution assay in simulated gastric or intestinal fluids lasted 6 h (n ¼ 6 for each test). Another dissolution experiment (n ¼ 6) has been performed with two steps: 1 h in gastric followed by 6 h in intestinal medium. AR release in simulated gastric and intestinal fluids The release of AR from chitosan/alginate microparticles was also tested, under the same conditions as described above for particle
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dissolution. Samples withdrawn at 15-min or 1 h intervals were centrifuged (5000 g, 10 min, RT) and the AR content of the supernatant was determined by measuring absorbance at 520 nm. Statistical analysis Values are presented as mean the standard error of the mean. Comparisons between groups were based on a one-way ANOVA. A difference between means was considered significant if p 0.05.
Results To obtain nanoparticle size uniformity, chitosan concentration (0.25, 0.50, 1 and 2%), sodium sulphate concentration (7.5, 10, 12.5 and 15%) and the effect of stirring during the ultrasound (sonication) treatment were studied. As sodium sulphate was added, chitosan solutions changed from clear to opalescent or turbid, indicating the formation of particles or aggregates. Using 10% sodium sulphate, sonication and stirring, particle mean hydrodynamic diameter increased from 520 nm to 1980 nm as chitosan concentration increased from 0.25 to 2% (Figure 1A). At 0.25% and 0.50% chitosan, particle mean diameters were, respectively, 520 and 530 nm, with the 410–550 nm size range accounting for, respectively, 14% and 10%. Moreover, the values of distribution width (span) were low (around 1.6). It was noted that particle monodispersity was obtained at these two chitosan concentrations. Contrariwise a polydispersity was observed when 1% and 2% chitosan was used to develop nanoparticles with a high span calculation. At 0.25% chitosan, the size of the particles obtained by sonication with stirring increased with the sodium sulphate concentration (Figure 1B). Indeed, the mean hydrodynamic diameters of particles varied from 520 nm to 840 nm for sodium sulphate at 7.5% and 15%, respectively. In the same way, the span differed from 1.6 to 2.3. At 10% sodium sulphate, stirring resulted
Preparation of chitosan nanoparticles loaded with AR The nanoparticles formed had a mean hydrodynamic diameter of 690 20 nm. The AR encapsulation efficacy was 21.9 0.5% and the content of AR was 0.18 0.1 mg/mg nanoparticles (around 4.5 mg of AR, n ¼ 10). Preparation of alginate microparticles loaded with chitosan nanoparticles To obtain alginate microparticles with chitosan nanoparticles distributed evenly throughout, several experimental parameters including nanoparticle concentration and final alginate concentration were studied. At higher nanoparticle concentrations (0.25, 0.30, 0.35, 0.40, 0.60, 0.80%, w/v, final alginate concentration of 0.5%), immediate precipitation was observed, leading to the formation of agglomerates. At a nanoparticle concentration of 0.2%, mean particle diameters of 228 18 mm (Figure 4A) and 283 26 mm were obtained using, respectively, the 100-mm and 150-mm nozzles (Figure 4B). All particles appeared smooth-surfaced and well dispersed in the solution, as observed by light microscopy (Figure 4). However, their shape was not spherical, due likely to the low viscosity of the nanoparticle/alginate solution. Using a nanoparticle/alginate solution ratio of 4:1 (final alginate concentration of 0.8%), a more viscous solution was obtained. Under this condition, chitosan–alginate particles about 265 mm in diameter and of spherical shape were obtained using the 150-mm nozzle (Figure 4C and D).
(A) 14
Chitosan concentrations 0.25% 0.50% 1.00% 2.00%
12 Volume (%)
Figure 1. Effect of chitosan (A) and sodium sulphate (B) concentrations on particle size distribution. Chitosan (A) powder (0.25; 0.5; 1 or 2%, w/v) was dissolved in 200 mL acetic acid solution (1%, v/v). Then, 3.5 mL of sodium sulphate solution at 10% (w/v) was added dropwise (1 mL/min) with continuous sonication (4 C, 15 min) and with stirring (100 rpm, 60 min, room temperature). Chitosan powder (0.25, w/v) was dissolved into 200 mL acetic acid solution (1%, v/v). Then, 3.5 mL of sodium sulphate solution (B) at 7.5; 10; 12.5 or 15% (w/v) was added dropwise (1 mL/min) with continuous sonication (4 C, 15 min) and with stirring (100 rpm, 60 min, room temperature).
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Preparation of chitosan nanoparticles
in a lower mean hydrodynamic diameter (520 nm vs. 570 nm, Figure 2A). Without sonication, the polydispersity of the particles obtained was high and the mean hydrodynamic diameter was 6000 nm. Sonication produced a uniform size distribution with a mean radius of 520 nm (Figure 2B). Scanning electron micrograph (Figure 3) indicated the formation of smooth-surfaced spherical nanoparticles in the size range of 100 nm.
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DOI: 10.3109/02652048.2013.858792
(A)
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Figure 2. Effect of stirring (A) and sonication (B) on particle size distribution. Chitosan powder (0.25%, w/v) was dissolved in 200 mL acetic acid solution (1%, v/v). Then, 3.5 mL of sodium sulphate solution at 10% (w/v) was added dropwise (1 mL/min) with (100 rpm, 60 min, room temperature) or without stirring (A) and with continuous (4 C, 15 min) or without sonication (B).
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Figure 3. Transmission electron microscopy (TEM) image of chitosan nanoparticles. Nanoparticles were formed by the addition of 3.5 mL of sodium sulphate (10%, w/v) to 200 mL of chitosan (0.25%, w/v). The formation process was performed with stirring (100 rpm, 60 min, room temperature) and sonication (4 C, 15 min). Nanoparticles obtained were examined by TEM (A: 30 kV; B: 50 kV).
Evaluation of AR release during alginate microparticle preparation Preliminary experiments indicated that pH plays a major role in AR loss from chitosan nanoparticles during the alginate microparticle formation process. In fact, up to 45% of the dye diffuses into the alginate portion as soon as the two solutions are mixed at pH 6–8, while flocculation is observed as the pH is adjusted to 4. A pH of 5.0 appeared to reduce AR loss from the nanoparticles to around 25% and minimise flocculation. Furthermore, increasing the alginate concentration from 0.5% to 0.8% increased shedding of AR into the supernatant from 23% to 63%, even though the particle size remained similar (around 285 mm). As observed by light microscopy, mixed chitosan/ alginate microparticles appeared smooth-surfaced, well dispersed in the solution and spherical at 0.8% alginate or less spherical at 0.5% alginate (data not shown). The non-ionic surfactant Tween 80 was effective at reducing premature dye loss to 7.80 0.12%. The amount of AR thus
retained in the mixed chitosan/alginate microparticles was more than 42 0.4 mg per gram. Light microscopy observations of these particles indicated a size range essentially unchanged (284 8 mm with Tween 80 and 283 26 without) and a smooth and spherical shape. Freeze-drying conditions The mean hydrodynamic diameter of amaranth-loaded mixed chitosan/alginate microparticles re-hydrated after lyophilising was significantly smaller (267 22 mm) than that of the fresh condition (284 8 mm). In contrast, particle shape was unchanged, with smooth and spherical surface. Polymer distribution within particles Imaging with CLSM indicated that the FITC-labelled chitosan nanoparticles were evenly distributed throughout the microparticle wall and matrix (Figure 5A). Using RBITC, homogenous
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Figure 4. Light microscopy observations of mixed chitosan/alginate microparticles. Nanoparticles suspension (0.2%, w/v) are combined to a 4% sodium alginate solution at a ratio nanoparticles:alginate of 7:1 (A and B) or 4:1 (C and D). Then, formed mixture is extruded through a nozzle with a diameter of 100 (A) or 150 mm (B–D). The droplets formed are collected in a solution of calcium chloride to 0.1 M and microparticles obtained are observed under 40 or 100 magnification.
Figure 5. Polymers distribution within particles by confocal laser scanning microscopy. After their formation, mixed microparticles were observed by CLSM (under 40 magnification); FITC labelled chitosan (A); RBITC labelled alginate (B); image obtained by superposition (C and D).
DOI: 10.3109/02652048.2013.858792
distribution of alginate throughout the particle wall and matrix was observed (Figure 5B). Observation of three-dimensional reconstructions by imaging several coplanar sections throughout the mixed chitosan/alginate microparticle (Figure 5C and D) further indicated that the distribution of alginate and chitosan was homogenous throughout the particles. In vitro release studies
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Degradation assay in simulated gastric and intestinal fluids Based on the light microscopy observation (40), microparticle size decreased significantly after 15 min in SGF, from 284 8 mm (control) to 192 8 mm with pepsin, and 220 19 mm without (Figure 6). Regardless of the presence or absence of enzyme, the microparticles retained their smooth surface until the end of the test. However, they lost their spherical shape and numerous creases appeared. No breakdown of alginate was observed. In SIF with or without pancreatin, microparticles were degraded rapidly, disappearing within 15 min, while no breakdown of chitosan nanoparticles was observed (Figure 6). Particle behaviour was thus similar in the presence or absence of enzymes, marked by a significant decrease in size and changes in shape in SGF within 1 h and rapid, complete breakdown with release of chitosan nanoparticles in SIF. AR release in simulated gastric and intestinal fluids Figure 7 displays AR release profiles in the presence of digestive enzymes (pepsin or pancreatin). In SGF, initial AR release began within 15 min (2.8 1.1%), and then reached only 5.0 0.5% at 3 h and 6.6 0.8% at the end of the experiment (6 h). The release into SGF fluid is probably due to dye onto particle surface. In SIF, more than 85% of the encapsulated dye was released within the
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first 15 min (Figure 7A), followed by slower release, reaching 88 3% and 92 2%, respectively, after 3 h and 6 h. When chitosan/alginate microparticles were first dispersed in SGF with pepsin for 1 h and then transferred to SIF with pancreatin for another 6 h, the same AR release profile was observed (Figure 7B). It thus appeared that less than 5% of the encapsulated dye was released into the gastric medium (after 1 h), while more than 85% was recovered in the intestinal medium within the first 15 min and reached around 92% at the end of the experiment.
Discussion To develop a food-grade carrier for protecting bioactive compounds or drugs from gastric acid and pepsin, two natural polymers were chosen, namely chitosan and alginate. Chitosan nanoparticles were developed to protect a model molecule (AR) from enzymatic attack in the small intestine. Furthermore, because of their spherical structure and small size (51 mm), these nanoparticles can enter freely into tissues through fine capillaries, diffuse between cells and are readily internalised by cells (Ermak and Giannasca, 1998). However, the glycosidic bonds of chitosan are rapidly hydrolyzed under gastric conditions (George and Abraham, 2006). To prevent breakdown of the nanoparticles, they were entrapped in microparticles of alginate, which is not degraded under gastric conditions. Our intention was to ensure protection of the nanoparticles until they reach the lumen of the small intestine, where they may be released in order to deliver the entrapped active compound. Absorption of intact nanoparticles into the bloodstream could thus be possible. In preliminary tests, it was found that nanoparticles were formed only with relatively low concentrations of polymer and flocculant. When sodium sulphate is added to a solution of chitosan in acetic acid, a decrease in polymer solubility occurs
Figure 6. Morphology of mixed chitosan/alginate microparticles in simulated gastric (SGF) or intestinal (SIF) fluids. Dry microparticles (0.2 g) were dispersed in 50 mL of 0.9% NaCl solution (control), SGF (pH 1.2, with or without pepsin) or SIF (pH 7.4, with or without pancreatin) under stirring (90 rpm) for 6 h. The morphology of microparticles was monitored microscopically under 40 (control and SGF conditions) and 1000 (SIF condition) magnifications. Arrows indicate chitosan nanoparticles.
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Figure 7. In vitro dissolution tests of mixed chitosan/alginate microparticles. Dry microparticles (0.2 g) were dispersed in 50 mL of SGF (pH 1.2, with pepsin-A), SIF (pH 7.4, with pancreatin-A) or SGF followed by SIF (B). Values represent the means (n ¼ 6 for each test) the standard error.
due to increased hydrogen bonding between the polysaccharide chains, resulting in precipitation (Wang et al., 2011). Under these mild conditions requiring no organic solvent, chitosan nanoparticles can be easily loaded with an expensive biologically active compound at minimal loss (Berthold et al., 1996). The mean hydrodynamic diameter of 520 nm obtained under optimal preparation conditions is consistent with the results of Borges et al. (2005) who prepared 684 nm nanoparticles. In our experiment, mastersizer measurement of these particles showed a unimodal distribution, indicating the development of uniform nanoparticles. It was also demonstrated that continuous stirring during nanoparticle formation had no significant impact (Figure 2A) on the mean hydrodynamic diameter (520 nm vs. 570 nm). Stirring was therefore maintained, since it promotes interaction between the molecule of interest and the chitosan. In contrast, sonication is a key factor in nanoparticle development. The mean hydrodynamic diameter (Figure 2B) of the chitosan particles made without sonication was 6000 nm with a bimodal distribution (span value around 2.5), indicating aggregation of the nanoparticle with increasing particle size and polydispersity, and gross precipitation was noticed other than uniform particles. This result could be due to poor homogenisation of polymer and flocculant. The high shear fields produced by sonication thus appear essential for breaking large agglomerations into sub-micron-sized spheres (Bihari et al., 2008). TEM images (Figure 3) confirm the formation of nanoparticles, showing small,
smooth-surfaced spherical particles around 100 nm. As reported by Borges et al. (2005), the discrepancy between the TEM observation and the size range determined by dynamic light scattering (520 nm) is likely due to TEM sample preparation, which involves desiccation for 72 h. The food dye AR possesses three anionic groups (SO2 3 ) and, therefore, have a strong affinity with the cationic groups (NHþ 3 ) of chitosan. AR was loaded into the chitosan nanoparticles by mixing dye solution with chitosan solution prior to adding sodium sulphate. Strong electrostatic interaction between the high anionic charge density of AR and the amino groups of chitosan likely contributed to encapsulation efficiency. Encapsulation efficiency may be affected by the nature of the loaded molecule. For example, Chen and Subirade (2007) have shown that chitosan/ b-lactoglobulin nanoparticles were able to entrap another dye (brilliant blue) with an efficiency of about 60%. Using a precipitation method, recombinant human interleukin-2 was entrapped in chitosan microparticles with an encapsulation efficiency of 98% (Ozbas-Turan et al., 2002). The encapsulation of AR influenced nanoparticle diameter, since dye-loaded nanoparticles were somewhat bigger (690 nm). A reasonable explanation for this could be conformational change of the chitosan molecule, since the dye was incorporated into the chitosan solution before the formation of nanoparticles. It may be presumed that docking of the dye molecules on the polysaccharide chains increased the steric volume of the chitosan and that this
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DOI: 10.3109/02652048.2013.858792
additional volume was retained when sodium sulphate was added to cause precipitation. Having established the appropriate conditions and technique for obtaining chitosan nanoparticles, the next step was to protect the nanocarrier and loaded molecule against gastric conditions. Chitosan can be hydrolyzed under acidic conditions, by breakage of the hemiacetal glycosidic bonds (Wang et al., 2011). Sodium alginate is more resistant, and since it is an anionic polysaccharide with charges opposite those of chitosan, it was chosen as a natural polymer to protect chitosan nanoparticles from gastric hydrolysis. However, our goal was to obtain alginate microparticles with chitosan nanoparticles evenly distributed throughout. It was, therefore, necessary to formulate a suspension of nanoparticles in sodium alginate solution that would remain homogenous during microparticle formation. At chitosan nanoparticle concentrations above 0.2%, the formation of agglomerates was immediate, due likely to excessive interaction of chitosan cationic charges with anionic charges of alginate. At 0.2% chitosan the sought stable suspension of nanoparticles in sodium alginate solution was obtained (at a nanoparticle:alginate ratio of 7:1 or 4:1) and chitosan nanoparticles entrapped in alginate microparticles (around 280 mm) were successfully prepared. However, a major drawback of this technique is the loss of loaded compound from the nanoparticles during the process. Amaranth dye partitioned between the chitosan and alginate phases. Since the pKa of sodium alginate is between 3.4 and 4.4, this polymer was negatively charged under our conditions and may have competed with AR for cationic sites on chitosan and repelled the anionic charges of the dye. Borges et al. (2005) have described this phenomenon in the context of chitosan nanoparticles coated with sodium alginate, observing more than 60% of ovalbumin desorption when nanoparticles were added to sodium alginate solution. At pH 4, in the range of the pKa of alginate and hence equilibrium between negative and positive charges, rapid flocculation was observed. However, at pH 5, the competition between AR and polymer for chitosan cationic charges is reduced since there is a decrease of anionic charges of sodium alginate (dye desorption of 25%). The sodium alginate concentration can also modify the AR encapsulation efficiency. Indeed, the AR desorption in the medium is increased when nanoparticles suspension is mixed with the higher alginate concentration probably due to the presence of many anionic charges. A sodium alginate pH of 5 and a final concentration of 0.5% (nanoparticle:alginate ratio of 7:1) appeared to provide the smallest loss of dye. Under these process conditions, the size of particles remained near 280 mm. In the range of sodium alginate pH and concentrations studied, the viscosity of the mixed solution did not change much (results not shown) and did not appear to affect the microparticle formation process using the EncapsulatorÕ. The reduced loss of AR from the nanoparticles in the presence of the non-ionic surfactant Tween 80 could be due to interference with electrostatic interactions at the boundary between the anionic sodium alginate polymer and the cationic chitosan. The surfactant could also coat the nanoparticles and provide steric interference with dye diffusion. Since the long-term stability of the hydrated chitosan/alginate microparticles is poor at ambient temperature, freeze-drying was investigated as a preservation method. However, this process may cause particle aggregation, leading in some cases to irreversible fusion and increased size distribution (Abdelwahed et al., 2006). After freeze-drying with trehalose as a cryoprotectant (Dulieu and Bazile, 2005), our reconstituted microparticles showed a decrease in the average hydrodynamic diameter, an increased polydispersity index and deformities (data not shown). Using the non-ionic surfactant Tween 80 at a concentration of 1%, the
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chitosan/alginate microparticle physical and chemical characteristics (e.g. morphology and AR content) were preserved. It has been shown previously that low concentrations of this surfactant protect proteins entrapped in freeze-dried particles (Hillgren and Alden, 2002). Analysis by CLSM revealed a homogenous dispersion of nanoparticles throughout the alginate microparticles (Figure 5C and D). The association between RBITC and FITC fluorescence was intimate, as would be expected given that the strong electrostatic attraction should exist between the polymers due to opposite charges at the pH of the formulation. The slow release of AR into SGF (with or without pepsin) and microparticle shrinkage from 284 mm to about 200 mm with loss of porosity suggest that hydrogen bonding between alginate and chitosan hydroxyl groups was increased, which would be expected since the degree of ionisation of alginate is low at this pH, due to protonation of carboxylic groups (Chen et al., 2004). This condition reduced AR diffusion from the microparticles to about 5% over 6 h. Chitosan or chitosan/alginate particles have been considered unsuitable for oral delivery of therapeutic proteins (Chen et al., 2004) due to the rapid dissolution of chitosan at gastric pH. Indeed, protonation of the amino groups leads to chitosan chain repulsion, allowing diffusion of proton and counter ions along with water into the carrier and ultimately loss of particle integrity (Suksamran et al., 2011). By encapsulating chitosan nanoparticles inside alginate microparticles as in our study, polymer dissolution and drug release into the gastric environment are slowed. This micro-carrier system may, therefore, allow a pH-sensitive or polypeptide payload to pass through the stomach intact. At pH 7.4, the carboxyl groups of alginate are completely ionised (un-protonated), while chitosan amino groups are at most 9% protonated (Dambies et al., 2001). This would reduce hydrogen bonding and weaken the interaction between the two polymers, leading to rapid breakdown of the carriers and release of nanoparticles followed by release of the amaranth dye into the medium. However, dye release from chitosan nanoparticles in SIF is believed to occur by a diffusion process rather than erosion or dissociation of the polymeric matrix. At intestinal pH, the chitosan amino groups no longer attract the AR anionic groups, allowing dye release within 15 min. The release of this surrogate molecule from the nanoparticles is thus directly related to the surrounding pH. The low pH of the gastric environment guards against release, while the increased pH in the intestine allows it. However, this release pattern is likely a characteristic of our surrogate (which bears three anionic groups) and could be modified significantly in the case of a payload with a different ionic charge. Chitosan nanoparticles released in the small intestine should be able to interact with the negatively charged glycocalyx on the apical membrane of GIT cells. Since the pH in this environment is around 6.8, the nanoparticles should still bear enough positive charge to do so. Indeed, Behrens et al. (2002) have shown that intestinal cells interact more with chitosan than with polystyrene (a non-bioadhesive nanoparticle model). Moreover, chitosan is able to open the tight junctions between epithelial cells, thus facilitating the transport of carried molecules through the intestinal epithelium (Wong, 2009). This absorption-enhancing property is attributed in part to the capacity of chitosan to interact with negatively charged sites of ZO-1 proteins on cell surfaces, inducing redistribution of F-actin, a cytoskeleton protein regulating paracellular flow (Wong, 2010). Our nanometer-sized carrier could thus penetrate the intestinal mucosa within 30–60 min through intercellular spaces between enterocytes and M cells lining Peyer’s patches, as Carino and Mathiowitz (1999) have demonstrated with nanoparticles of sizes ranging from 10 to
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1000 nm (Carino and Mathiowitz, 1999). A compound that was not released too quickly from the nanoparticles could thus be delivered in the bloodstream to produce a therapeutic effect with greater efficiency.
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Conclusions The oral route of drug delivery is the most convenient and usually the safest and least expensive, as well as the one most often used. However, bioactive compounds or drugs advancing through the digestive tract may be degraded as they encounter gastric acid and/or digestive enzymes. There is, therefore, a need for carriers that protect pharmaceutical or nutraceutical payloads and facilitate their uptake by intestinal cells. In order to develop such a carrier, we chose two natural non-toxic polymers, namely chitosan and alginate. This is the first description of chitosan nanoparticles loaded with a compound entrapped in alginate microparticles. Using CLSM with different fluorescent labels for each of polymer, we were able to observe a homogenous distribution of both polymers throughout the microparticle matrix. The capacity of the carrier to control the release of the compound was then studied in a simulated gastric and intestinal environment with or without digestive enzymes. The less than 5% release of AR in the gastric environment suggests that this novel drug carrier and delivery system could protect pharmaceutical payloads until they reach the intestine, where release may be rapid and nearly complete, at least in the case of a molecule bearing a significant anionic charge. In such cases, the pH determines the strength of the electrostatic attraction binding the molecule to the carrier by varying the amount of positive charge on the chitosan chains. Meanwhile, the chitosan nanoparticles released in the intestinal environment remain intact. Because of their size, they are capable of crossing the intestinal boundary membrane and entering the bloodstream, where they could deliver drugs. These results confirm the possibility of using chitosan nanoparticles entrapped in alginate microparticles as a new drug delivery system, in particular to protect and enhance uptake of bioactive food-related compounds (e.g. polyphenolic compounds, vitamin D, curcumin) or poorly absorbed drugs (e.g. insulin, heparin, calcitonin). The surrogate compound (AR) used in the present study will be replaced in order to study candidate molecules identified for specific therapeutic targets.
Acknowledgements This study benefited from support provided by the De´le´gation Ge´ne´rale pour l’Armement (DGA) and by the Natural Sciences and Engineering Research Council of Canada (NSERC) Research Chair in Proteins, Biosystems and Functional Foods. The authors thank Claire Philippe for skilled technical assistance.
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ORIGINAL ARTICLE
Development of a novel drug delivery system: chitosan nanoparticles entrapped in alginate microparticles Ghislain Garrait1,2, Eric Beyssac1, and Muriel Subirade2
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1
Universite´ d’Auvergne, UFR Pharmacie, Equipe d’Accueil Conception, Inge´nierie et De´veloppement de l’Aliment et du Me´dicament (EA CIDAM), Clermont-Ferrand F-63001, France and 2Faculte´ des Sciences de l’Agriculture et de l’Alimentation, INAF/STELA, Universite´ Laval, Sainte-Foy, Que´bec, Canada Abstract
Keywords
A novel carrier using chitosan nanoparticles entrapped into alginate microparticles is proposed for protecting molecules of interest from degradation in the digestive tract. The effects of polymer concentration, sonication, stirring, pH, and processing conditions on the physical characteristics of the carrier were studied. FITC and RBITC were used to localise the polymers within particles using CLSM. Diffusion of amaranth red (AR) from nanoparticles was quantified during dissolution under gastric and intestinal conditions. Under optimal preparation conditions, the size distribution of nanoparticles loaded with AR was uniform (690 nm) with an encapsulation efficacy of 21.9%. Alginate microparticles (285 mm) containing a homogenous distribution of nanoparticles and polymers were obtained. At gastric pH, the carrier released less than 5% of the loaded AR and, at intestinal pH, the release was rapid and complete. The drug carriers developed shows a promising use as a vehicle suitable to protect molecules of interest after oral administration.
Alginate microparticle, chitosan nanoparticle, gastro-resistant, novel drug delivery system
Introduction The oral route is the most convenient, safe and widely accepted for administering drugs. However, acid and pepsin in the stomach and pancreatic enzymes in the small intestine can degrade pharmaceutical payloads or cause substantial losses thereof (Bernkop-Schnurch et al., 2004). In addition, many bioactive dietary compounds or drugs are absorbed poorly through the intestinal mucosa because of their physicochemical properties (i.e. high molecular mass, hydrophilic character). Carrier systems for protecting therapeutic or nutritional molecules against the harsh environment of the gastrointestinal tract (GIT) and providing efficient and site-specific delivery are therefore also receiving much attention. Among these carriers, nanoparticles present certain advantages as interaction with the biological environment, enhancement of absorption, and improvement retention time (Cheng et al., 2008). Nanoparticles are solid colloidal carriers ranging from 10 to 1000 nm in diameter and may be composed of synthetic or natural polymers. However, the preparation of synthetic polymer nanoparticles involves heat or organic solvent, which can affect bioactive compound structure or stability. In contrast, natural polymers are often water-soluble, biodegradable, biocompatible, less toxic and amenable to simple preparation techniques under mild conditions. Among the natural polymers, alginate and chitosan have been studied extensively, making them very attractive as drug delivery carriers.
Address for correspondence: Muriel Subirade, Faculte´ des Sciences de l’Agriculture et de l’Alimentation, INAF/STELA, Universite´ Laval, Pavillon Paul-Comtois, Que´bec QC G1K 7P4, Canada. Tel: +1 418 656 2131x4278. Fax: +1 418 656 2131 3353. E-mail: [email protected]
History Received 7 May 2013 Revised 16 September 2013 Accepted 15 October 2013 Published online 1 April 2014
Alginate is an anionic and water-soluble polysaccharide and can be cross-linked under mild conditions in the presence of any divalent cation (Ca2þ, Sr2þ, Cu2þ or Zn2þ) (Wee and Gombotz, 1998). Alginate gelation may be used to form particles that protect payload compounds against the gastric environment. In addition, alginate microparticles possess a bioadhesive property that allows them to stick to the intestinal mucosa and thus increase drug residence time (Chickering et al., 1997). Due to its various properties, alginate has been used widely to develop drug delivery systems especially for oral route use (Wee and Gombotz, 1998). However, significant drug leakage through pores is well known (Liu and Krishnan, 1999). Thus, alginate has been combined successfully with chitosan to form a chitosan–alginate complex with lower porosity limiting the leakage of the encapsulated molecules (Douglas and Tabrizian, 2005). Chitosan is a cationic polysaccharide obtained from the partial deacetylation of chitin. It carries a positive charge and can react with negatively charged surfaces (e.g. mucus) to prolong retention time and improve the bioavailability of the payload (Deacon et al., 2000). These properties all make this polymer suitable for use as a drug delivery carrier. Indeed, chitosan has been used widely to develop particles for controlled release of active agents via several routes of administration (e.g. oral, nasal). The main drawback of administering chitosan carriers via the oral route is the easy dissolution of the polymer matrix exposing acid-sensitive payloads to degradation (George and Abraham, 2006). Alginate interacts via anionic carboxylic groups with cationic amino groups of chitosan to form chitosan–alginate particles. This complex is stable under acidic conditions and thus protects pharmaceutical preparations from degradation (Takka and Gurel, 2010), but becomes soluble at intestinal pH, allowing rapid and extensive release of the loaded compound (Woitiski et al., 2010).
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The objective of this study was to develop a novel drug delivery system using chitosan nanoparticles entrapped into alginate microparticles. The effects of processing conditions on the physical characteristics (e.g. size, shape) of the chitosan nanoparticles and alginate microparticles were investigated. Parameters such as polymer and sodium sulphate concentrations, sonication, stirring and addition of surfactant were examined. Fluorescein 5(6)-isothiocyanate (FITC) and rhodamine B isothiocyanate (RBITC) were used to localise the polymers within the particles. Amaranth red (AR), used as a model, was loaded into the nanoparticles, and release kinetics in simulated gastric or intestinal fluids were investigated.
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Materials
Chitosan nanoparticles Preparation of chitosan nanoparticles The effects of different factors (chitosan concentration, sodium sulphate concentration, sonication and stirring) on the particle size were studied (Table 1). All experiments were performed in triplicate. Chitosan nanoparticles were prepared by the precipitation/ coaversion method described previously (Borges et al., 2005). Briefly, a solution (200 mL) of 2% (v/v) acetic acid and 1% (w/v) Tween 80 was prepared, in which low-molecular-weight chitosan powder was dissolved (Table 1). Sodium sulphate solution (3.5 mL) was then added dropwise (1 mL/min) with or without continuous sonication (Sonicator model XL 2020, 550 W, output control ‘‘1’’, Misonix, Farmingdale, NY) at 4 C for 15 min and with or without stirring (100 rpm) for 60 min at room temperature (RT). Sodium sulphate induced the production of chitosan particles by a decrease in polymer solubility inducing a precipitation. An opalescent suspension was formed spontaneously and the particles therein were recovered by centrifugation (5000 g, 15 min, RT), re-suspended and centrifuged five times in 40 mL of ultrapure water. Particles thus obtained were characterised immediately. Table 1. Preparation conditions to study the effects of chitosan and sodium sulphate concentrations, stirring and sonication on particles size distribution.
Chitosan concentration (w/v) Sodium sulphate concentration (w/v) Sonication (min) Stirring (min)/(rpm) NA, not applicable.
AR (0.01%, w/v) was added to 200 mL of acetic acid/Tween 80 solution with stirring. After 15 min, 0.25% (w/v) chitosan was dissolved therein by stirring for 1 h. An aliquot was taken to determine the total amount of AR at the beginning of the experiment. Sodium sulphate solution (3.5 mL, 10%, w/v) was then added to form chitosan nanoparticles by continuous sonication (15 min, 4 C) with stirring (100 rpm, 60 min, RT). The nanoparticles obtained were washed (to remove unloaded AR) and centrifuged (5000 g, 15 min, RT) five times with 40 mL of ultrapure water. Nanoparticle size was determined immediately and dye encapsulated in nanoparticles was quantified directly to determine AR encapsulation efficacy and content. Alginate microparticles
Low-molecular-weight chitosan with deacetylation up to high as 75% according to specifications and viscosity ranging from 20 to 300 cps (1% solution in 1% acetic acid, 25 C), acetic acid 99%, TweenÕ 80, alginic acid sodium salt (15 cps for 1% w/v aqueous solution, mannuronic/guluronic acid ratio of 0.65), calcium chloride, FITC, RBITC, enzymes and other chemicals used for dissolution experiments were all purchased from the Sigma Chemical Company (St Louis, MO). AR was obtained from Sensient Colors Canada Ltd (Kingston, ON) and sodium sulphate was obtained from Fluka Chemical Corp. (Ronkonkoma, NY). All solutions were prepared in ultrapure water. All other chemicals were of reagent grade.
Formulation
Preparation of amaranth-red-loaded chitosan nanoparticles
1 0.25; 0.5; 1; 2% 10% 15 60/100
2
3
4
0.25%
0.25%
0.25%
7.5; 10; 12.5; 15% 15 60/100
10%
10%
NA 60/100
15 NA
Preparation of alginate microparticles loaded with chitosan nanoparticles Alginate microparticles were obtained using an EncapsulatorÕ (model IER50R, Inotech, Switzerland), of which the vibrating nozzle breaks a laminar liquid jet (i.e. alginate solution containing chitosan nanoparticles) into equal-sized micro-droplets, which were collected in a solution of calcium chloride to stabilise them as microparticles. Several experimental conditions were tested in order to obtain a homogeneous suspension of nanoparticles in alginate. Nanoparticle suspensions (0.2, 0.25, 0.3, 0.35, 0.4, 0.6 and 0.8% w/v) were combined with 4% sodium alginate solution at a nanoparticle:alginate ratio of 7:1 (final alginate concentration of 0.5%), stirred for 30 min and then run through the Encapsulator (700–1200 Hz, amplitude 4, 0.6 kV, 0.6 Bar, nozzle diameter 100 or 150 mm). The droplets formed were collected in 0.1 M calcium chloride solution. A nanoparticle:alginate ratio of 4:1 was studied using the 150-mm nozzle. The microparticles formed were observed by optical microscopy and the particle diameter was determined using an image-processing software. Evaluation of AR release during alginate microparticle preparation To avoid release of AR from chitosan nanoparticles into the alginate during microparticle formation, we investigated the impact of pH and alginic acid concentration. A suspension of AR-loaded nanoparticles (0.2% w/v in ultrapure water) was added dropwise to 4% alginic acid solution (pH adjusted to 4, 5, 6, 7 or 8 with 1 M hydrochloric acid, n ¼ 2 for each pH value) with stirring until a final alginate concentration of 0.8% w/v was reached (nanoparticle:alginate ratio of 4:1). In a second experiment, alginic acid solution (4% or 3%, pH 5) was diluted to 0.5, 0.6 and 0.8% with 0.2% nanoparticle suspension (n ¼ 2 for each test). Vigorous stirring was maintained for 15 min and an aliquot (4 mL) was centrifuged (10 000 g, 10 min, RT) and AR in the supernatant was measured as absorbance at 520 nm. The solutions were run through the 150-mm nozzle and the droplets were collected in 0.1 M calcium chloride solution. The dye content of the recovered particles was assayed. Finally, in another attempt to reduce AR release during microparticle formation, chitosan nanoparticles were re-suspended at 0.2% w/v in 1% v/v Tween 80. This suspension was added dropwise to 4% alginate solution (pH 5) at a nanoparticle:alginate ratio of 7:1 (final alginate concentration 0.5% w/v). The mixture was run through the 150-mm nozzle and droplets were collected in 0.1 M calcium chloride solution.
DOI: 10.3109/02652048.2013.858792
Microparticle morphology, size and AR content were then determined. Freeze-drying
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Freshly prepared alginate microparticles entrapping AR-loaded chitosan nanoparticles were dispersed in ultrapure water containing 1% (v/v) Tween 80. The mixture was stirred for 30 min then filtered on large pore size, frozen in liquid nitrogen and lyophilised (Model Freezone 4.5, Labconco, Kansas City, MI) for 48 h. The dry particles were stored at 20 C until further use in order to avoid the degradation of alginate microparticles entrapping AR-loaded chitosan nanoparticles. To determine their morphology and size, the microparticles were dispersed in 0.9% NaCl solution (50 mL) and stirred for 1 h. Characterisation of particles Particle size analysis Measurement of size and size distribution of freshly prepared nanoparticles was based on the static light scattering using a Mastersizer 2000 (Malvern Instruments, Southborough, MA). Particles were suspended in distilled water containing 1% v/v Tween 80. All size measurements were performed at 25 C and at a 90 scattering angle with 180 s of recording. The mean hydrodynamic diameter was generated by cumulative analysis (no less than triplicate with independent particle batches). Microparticle size was also evaluated using an optical microscope (Olympus BX50WI, Olympus, Melville, NY) fitted with a digital camera (model U-TV1 X, Olympus Optical, Tokyo, Japan). Microparticles were visualised at 40 magnification and the diameter of 50 randomly selected microparticles was determined using an image processing Image-Pro PlusÕ software (Media Cybernetics, Inc., Rockville, MD). Surface particle morphology Nanoparticle morphology was examined by transmission electron microscopy (TEM) using a Jeol 1230 microscope (Tokyo, Japan). A drop of freshly prepared nanoparticles in ultrapure water was air-dried onto a copper grid. After negative staining with uranyl acetate, the sample was desiccated for at least 72 h at RT and then examined at 30 and 50 kV. Nanoparticles were visualised using different accelerating voltages in order to improve resolution and modify magnification. Microparticles were also examined at 40 and 100 using the optical microscope. AR content of particles To evaluate the AR encapsulation efficacy and content of the nanoparticles, 25 mg of nanoparticles were disrupted in 25 mL of simulated gastric fluid (SGF) (USP standard XXX, 2 g of sodium chloride, 7 mL of 37% hydrochloric acid and 1 L of ultrapure water, pH 1.2) with vigorous magnetic stirring at RT for 30 min. The resulting solution was centrifuged at 10 000 rpm for 20 min at RT and absorbance at 520 nm by the supernatant was measured using a UV-visible spectrophotometer (model 8453, HewlettPackard, Palo Alto, CA). The absorbance was compared to a standard curve prepared at the same time. AR encapsulation efficacy (EE) was obtained from the relationship EE ¼ 100A/B, where A is the AR encapsulated in the chitosan nanoparticles and B is the total amount of AR added. AR content into alginate microparticles was determined by disrupting microparticles (6 g) in 50 mL of a mixture of 0.2 M sodium bicarbonate and 0.06 M tri-basic sodium citrate at
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pH 8.0 (Li et al., 2007), centrifuging, measuring the supernatant OD at 520 nm. AR content was obtained from the following relationship: 100 amount of AR encapsulated in chitosan nanoparticlesðgÞ AR content ð%Þ ¼ amount of chitosan nanoparticlesðgÞ Polymer distribution within particles The localisation of both polymers within the particles was determined using confocal laser scanning microscopy (CLSM). The labelling protocol was based on a method previously published (Huang et al., 2002). Chitosan was labelled with FITC while alginate was labelled with RBITC as published previously (Mladenovska et al., 2007). FITC and RBITC were chosen to allow simultaneous detection in different channels. FITC-labelled chitosan nanoparticles were prepared by dissolving labelled polymer (0.25%, w/v) in acetic acid/Tween 80 solution and adding 3.5 mL of sodium sulphate solution (10%, w/v) dropwise with continuous sonication at 4 C for 15 min and with magnetic stirring (100 rpm) for 60 min at RT. Particles were recovered by centrifugation (5000 g, 15 min, RT), washed twice and used to produce alginate microparticles. Labelled microparticles were obtained using a FITC-labelled chitosan nanoparticle suspension (0.2%, w/v) stirred with RBITClabelled alginate solution (final concentration 0.8% w/v) for 30 min and run through the encapsulator fitted with the 150-mm nozzle. The droplets were collected in 0.5 M calcium chloride. The double-labelled microparticles were observed using the Olympus light microscope Fluoview FV 300 Laser Scanning Confocal Imaging System (Paris, France) equipped with an argon ion laser (EM 488 nm) and green (543 nm) and red (633 nm) helium–neon ion laser. The laser was adjusted to excitation/ emission wavelengths of 488/520 nm for FITC and 543/572 nm for RBITC. All confocal fluorescence pictures were taken with a 40 objective (oil immersion). In vitro release studies Dissolution assay in simulated gastric and intestinal fluids Dry microparticles (0.2 g) were dispersed in 50 mL of SGF with or without pepsin (3.2 g, 924 units/mg of protein) or in simulated intestinal fluid (SIF; 6.8 g of monobasic potassium phosphate dissolved in 250 mL of ultrapure water and added to 190 mL of 0.2 N sodium hydroxide and 400 mL of ultrapure water and adjusted to pH 7.4 with 0.2 N sodium hydroxide) with or without pancreatin (10 g, activity equivalent to USP specification) as described in USP standard XXX (2007) using paddle apparatus II. Agitator speed was set at 90 rpm and the temperature was maintained at 37 0.5 C. Samples were withdrawn at 15-min or 1 h intervals and particle shape was observed microscopically (40 and 100). The diameter of 50 randomly selected particles was determined using an image processing Image-Pro PlusÕ software. An equal volume of medium was added to the release mixture after each sampling to maintain a constant volume of 50 mL. Dissolution assay in simulated gastric or intestinal fluids lasted 6 h (n ¼ 6 for each test). Another dissolution experiment (n ¼ 6) has been performed with two steps: 1 h in gastric followed by 6 h in intestinal medium. AR release in simulated gastric and intestinal fluids The release of AR from chitosan/alginate microparticles was also tested, under the same conditions as described above for particle
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dissolution. Samples withdrawn at 15-min or 1 h intervals were centrifuged (5000 g, 10 min, RT) and the AR content of the supernatant was determined by measuring absorbance at 520 nm. Statistical analysis Values are presented as mean the standard error of the mean. Comparisons between groups were based on a one-way ANOVA. A difference between means was considered significant if p 0.05.
Results To obtain nanoparticle size uniformity, chitosan concentration (0.25, 0.50, 1 and 2%), sodium sulphate concentration (7.5, 10, 12.5 and 15%) and the effect of stirring during the ultrasound (sonication) treatment were studied. As sodium sulphate was added, chitosan solutions changed from clear to opalescent or turbid, indicating the formation of particles or aggregates. Using 10% sodium sulphate, sonication and stirring, particle mean hydrodynamic diameter increased from 520 nm to 1980 nm as chitosan concentration increased from 0.25 to 2% (Figure 1A). At 0.25% and 0.50% chitosan, particle mean diameters were, respectively, 520 and 530 nm, with the 410–550 nm size range accounting for, respectively, 14% and 10%. Moreover, the values of distribution width (span) were low (around 1.6). It was noted that particle monodispersity was obtained at these two chitosan concentrations. Contrariwise a polydispersity was observed when 1% and 2% chitosan was used to develop nanoparticles with a high span calculation. At 0.25% chitosan, the size of the particles obtained by sonication with stirring increased with the sodium sulphate concentration (Figure 1B). Indeed, the mean hydrodynamic diameters of particles varied from 520 nm to 840 nm for sodium sulphate at 7.5% and 15%, respectively. In the same way, the span differed from 1.6 to 2.3. At 10% sodium sulphate, stirring resulted
Preparation of chitosan nanoparticles loaded with AR The nanoparticles formed had a mean hydrodynamic diameter of 690 20 nm. The AR encapsulation efficacy was 21.9 0.5% and the content of AR was 0.18 0.1 mg/mg nanoparticles (around 4.5 mg of AR, n ¼ 10). Preparation of alginate microparticles loaded with chitosan nanoparticles To obtain alginate microparticles with chitosan nanoparticles distributed evenly throughout, several experimental parameters including nanoparticle concentration and final alginate concentration were studied. At higher nanoparticle concentrations (0.25, 0.30, 0.35, 0.40, 0.60, 0.80%, w/v, final alginate concentration of 0.5%), immediate precipitation was observed, leading to the formation of agglomerates. At a nanoparticle concentration of 0.2%, mean particle diameters of 228 18 mm (Figure 4A) and 283 26 mm were obtained using, respectively, the 100-mm and 150-mm nozzles (Figure 4B). All particles appeared smooth-surfaced and well dispersed in the solution, as observed by light microscopy (Figure 4). However, their shape was not spherical, due likely to the low viscosity of the nanoparticle/alginate solution. Using a nanoparticle/alginate solution ratio of 4:1 (final alginate concentration of 0.8%), a more viscous solution was obtained. Under this condition, chitosan–alginate particles about 265 mm in diameter and of spherical shape were obtained using the 150-mm nozzle (Figure 4C and D).
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Figure 1. Effect of chitosan (A) and sodium sulphate (B) concentrations on particle size distribution. Chitosan (A) powder (0.25; 0.5; 1 or 2%, w/v) was dissolved in 200 mL acetic acid solution (1%, v/v). Then, 3.5 mL of sodium sulphate solution at 10% (w/v) was added dropwise (1 mL/min) with continuous sonication (4 C, 15 min) and with stirring (100 rpm, 60 min, room temperature). Chitosan powder (0.25, w/v) was dissolved into 200 mL acetic acid solution (1%, v/v). Then, 3.5 mL of sodium sulphate solution (B) at 7.5; 10; 12.5 or 15% (w/v) was added dropwise (1 mL/min) with continuous sonication (4 C, 15 min) and with stirring (100 rpm, 60 min, room temperature).
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in a lower mean hydrodynamic diameter (520 nm vs. 570 nm, Figure 2A). Without sonication, the polydispersity of the particles obtained was high and the mean hydrodynamic diameter was 6000 nm. Sonication produced a uniform size distribution with a mean radius of 520 nm (Figure 2B). Scanning electron micrograph (Figure 3) indicated the formation of smooth-surfaced spherical nanoparticles in the size range of 100 nm.
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Figure 2. Effect of stirring (A) and sonication (B) on particle size distribution. Chitosan powder (0.25%, w/v) was dissolved in 200 mL acetic acid solution (1%, v/v). Then, 3.5 mL of sodium sulphate solution at 10% (w/v) was added dropwise (1 mL/min) with (100 rpm, 60 min, room temperature) or without stirring (A) and with continuous (4 C, 15 min) or without sonication (B).
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Figure 3. Transmission electron microscopy (TEM) image of chitosan nanoparticles. Nanoparticles were formed by the addition of 3.5 mL of sodium sulphate (10%, w/v) to 200 mL of chitosan (0.25%, w/v). The formation process was performed with stirring (100 rpm, 60 min, room temperature) and sonication (4 C, 15 min). Nanoparticles obtained were examined by TEM (A: 30 kV; B: 50 kV).
Evaluation of AR release during alginate microparticle preparation Preliminary experiments indicated that pH plays a major role in AR loss from chitosan nanoparticles during the alginate microparticle formation process. In fact, up to 45% of the dye diffuses into the alginate portion as soon as the two solutions are mixed at pH 6–8, while flocculation is observed as the pH is adjusted to 4. A pH of 5.0 appeared to reduce AR loss from the nanoparticles to around 25% and minimise flocculation. Furthermore, increasing the alginate concentration from 0.5% to 0.8% increased shedding of AR into the supernatant from 23% to 63%, even though the particle size remained similar (around 285 mm). As observed by light microscopy, mixed chitosan/ alginate microparticles appeared smooth-surfaced, well dispersed in the solution and spherical at 0.8% alginate or less spherical at 0.5% alginate (data not shown). The non-ionic surfactant Tween 80 was effective at reducing premature dye loss to 7.80 0.12%. The amount of AR thus
retained in the mixed chitosan/alginate microparticles was more than 42 0.4 mg per gram. Light microscopy observations of these particles indicated a size range essentially unchanged (284 8 mm with Tween 80 and 283 26 without) and a smooth and spherical shape. Freeze-drying conditions The mean hydrodynamic diameter of amaranth-loaded mixed chitosan/alginate microparticles re-hydrated after lyophilising was significantly smaller (267 22 mm) than that of the fresh condition (284 8 mm). In contrast, particle shape was unchanged, with smooth and spherical surface. Polymer distribution within particles Imaging with CLSM indicated that the FITC-labelled chitosan nanoparticles were evenly distributed throughout the microparticle wall and matrix (Figure 5A). Using RBITC, homogenous
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Figure 4. Light microscopy observations of mixed chitosan/alginate microparticles. Nanoparticles suspension (0.2%, w/v) are combined to a 4% sodium alginate solution at a ratio nanoparticles:alginate of 7:1 (A and B) or 4:1 (C and D). Then, formed mixture is extruded through a nozzle with a diameter of 100 (A) or 150 mm (B–D). The droplets formed are collected in a solution of calcium chloride to 0.1 M and microparticles obtained are observed under 40 or 100 magnification.
Figure 5. Polymers distribution within particles by confocal laser scanning microscopy. After their formation, mixed microparticles were observed by CLSM (under 40 magnification); FITC labelled chitosan (A); RBITC labelled alginate (B); image obtained by superposition (C and D).
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distribution of alginate throughout the particle wall and matrix was observed (Figure 5B). Observation of three-dimensional reconstructions by imaging several coplanar sections throughout the mixed chitosan/alginate microparticle (Figure 5C and D) further indicated that the distribution of alginate and chitosan was homogenous throughout the particles. In vitro release studies
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Degradation assay in simulated gastric and intestinal fluids Based on the light microscopy observation (40), microparticle size decreased significantly after 15 min in SGF, from 284 8 mm (control) to 192 8 mm with pepsin, and 220 19 mm without (Figure 6). Regardless of the presence or absence of enzyme, the microparticles retained their smooth surface until the end of the test. However, they lost their spherical shape and numerous creases appeared. No breakdown of alginate was observed. In SIF with or without pancreatin, microparticles were degraded rapidly, disappearing within 15 min, while no breakdown of chitosan nanoparticles was observed (Figure 6). Particle behaviour was thus similar in the presence or absence of enzymes, marked by a significant decrease in size and changes in shape in SGF within 1 h and rapid, complete breakdown with release of chitosan nanoparticles in SIF. AR release in simulated gastric and intestinal fluids Figure 7 displays AR release profiles in the presence of digestive enzymes (pepsin or pancreatin). In SGF, initial AR release began within 15 min (2.8 1.1%), and then reached only 5.0 0.5% at 3 h and 6.6 0.8% at the end of the experiment (6 h). The release into SGF fluid is probably due to dye onto particle surface. In SIF, more than 85% of the encapsulated dye was released within the
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first 15 min (Figure 7A), followed by slower release, reaching 88 3% and 92 2%, respectively, after 3 h and 6 h. When chitosan/alginate microparticles were first dispersed in SGF with pepsin for 1 h and then transferred to SIF with pancreatin for another 6 h, the same AR release profile was observed (Figure 7B). It thus appeared that less than 5% of the encapsulated dye was released into the gastric medium (after 1 h), while more than 85% was recovered in the intestinal medium within the first 15 min and reached around 92% at the end of the experiment.
Discussion To develop a food-grade carrier for protecting bioactive compounds or drugs from gastric acid and pepsin, two natural polymers were chosen, namely chitosan and alginate. Chitosan nanoparticles were developed to protect a model molecule (AR) from enzymatic attack in the small intestine. Furthermore, because of their spherical structure and small size (51 mm), these nanoparticles can enter freely into tissues through fine capillaries, diffuse between cells and are readily internalised by cells (Ermak and Giannasca, 1998). However, the glycosidic bonds of chitosan are rapidly hydrolyzed under gastric conditions (George and Abraham, 2006). To prevent breakdown of the nanoparticles, they were entrapped in microparticles of alginate, which is not degraded under gastric conditions. Our intention was to ensure protection of the nanoparticles until they reach the lumen of the small intestine, where they may be released in order to deliver the entrapped active compound. Absorption of intact nanoparticles into the bloodstream could thus be possible. In preliminary tests, it was found that nanoparticles were formed only with relatively low concentrations of polymer and flocculant. When sodium sulphate is added to a solution of chitosan in acetic acid, a decrease in polymer solubility occurs
Figure 6. Morphology of mixed chitosan/alginate microparticles in simulated gastric (SGF) or intestinal (SIF) fluids. Dry microparticles (0.2 g) were dispersed in 50 mL of 0.9% NaCl solution (control), SGF (pH 1.2, with or without pepsin) or SIF (pH 7.4, with or without pancreatin) under stirring (90 rpm) for 6 h. The morphology of microparticles was monitored microscopically under 40 (control and SGF conditions) and 1000 (SIF condition) magnifications. Arrows indicate chitosan nanoparticles.
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Figure 7. In vitro dissolution tests of mixed chitosan/alginate microparticles. Dry microparticles (0.2 g) were dispersed in 50 mL of SGF (pH 1.2, with pepsin-A), SIF (pH 7.4, with pancreatin-A) or SGF followed by SIF (B). Values represent the means (n ¼ 6 for each test) the standard error.
due to increased hydrogen bonding between the polysaccharide chains, resulting in precipitation (Wang et al., 2011). Under these mild conditions requiring no organic solvent, chitosan nanoparticles can be easily loaded with an expensive biologically active compound at minimal loss (Berthold et al., 1996). The mean hydrodynamic diameter of 520 nm obtained under optimal preparation conditions is consistent with the results of Borges et al. (2005) who prepared 684 nm nanoparticles. In our experiment, mastersizer measurement of these particles showed a unimodal distribution, indicating the development of uniform nanoparticles. It was also demonstrated that continuous stirring during nanoparticle formation had no significant impact (Figure 2A) on the mean hydrodynamic diameter (520 nm vs. 570 nm). Stirring was therefore maintained, since it promotes interaction between the molecule of interest and the chitosan. In contrast, sonication is a key factor in nanoparticle development. The mean hydrodynamic diameter (Figure 2B) of the chitosan particles made without sonication was 6000 nm with a bimodal distribution (span value around 2.5), indicating aggregation of the nanoparticle with increasing particle size and polydispersity, and gross precipitation was noticed other than uniform particles. This result could be due to poor homogenisation of polymer and flocculant. The high shear fields produced by sonication thus appear essential for breaking large agglomerations into sub-micron-sized spheres (Bihari et al., 2008). TEM images (Figure 3) confirm the formation of nanoparticles, showing small,
smooth-surfaced spherical particles around 100 nm. As reported by Borges et al. (2005), the discrepancy between the TEM observation and the size range determined by dynamic light scattering (520 nm) is likely due to TEM sample preparation, which involves desiccation for 72 h. The food dye AR possesses three anionic groups (SO2 3 ) and, therefore, have a strong affinity with the cationic groups (NHþ 3 ) of chitosan. AR was loaded into the chitosan nanoparticles by mixing dye solution with chitosan solution prior to adding sodium sulphate. Strong electrostatic interaction between the high anionic charge density of AR and the amino groups of chitosan likely contributed to encapsulation efficiency. Encapsulation efficiency may be affected by the nature of the loaded molecule. For example, Chen and Subirade (2007) have shown that chitosan/ b-lactoglobulin nanoparticles were able to entrap another dye (brilliant blue) with an efficiency of about 60%. Using a precipitation method, recombinant human interleukin-2 was entrapped in chitosan microparticles with an encapsulation efficiency of 98% (Ozbas-Turan et al., 2002). The encapsulation of AR influenced nanoparticle diameter, since dye-loaded nanoparticles were somewhat bigger (690 nm). A reasonable explanation for this could be conformational change of the chitosan molecule, since the dye was incorporated into the chitosan solution before the formation of nanoparticles. It may be presumed that docking of the dye molecules on the polysaccharide chains increased the steric volume of the chitosan and that this
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additional volume was retained when sodium sulphate was added to cause precipitation. Having established the appropriate conditions and technique for obtaining chitosan nanoparticles, the next step was to protect the nanocarrier and loaded molecule against gastric conditions. Chitosan can be hydrolyzed under acidic conditions, by breakage of the hemiacetal glycosidic bonds (Wang et al., 2011). Sodium alginate is more resistant, and since it is an anionic polysaccharide with charges opposite those of chitosan, it was chosen as a natural polymer to protect chitosan nanoparticles from gastric hydrolysis. However, our goal was to obtain alginate microparticles with chitosan nanoparticles evenly distributed throughout. It was, therefore, necessary to formulate a suspension of nanoparticles in sodium alginate solution that would remain homogenous during microparticle formation. At chitosan nanoparticle concentrations above 0.2%, the formation of agglomerates was immediate, due likely to excessive interaction of chitosan cationic charges with anionic charges of alginate. At 0.2% chitosan the sought stable suspension of nanoparticles in sodium alginate solution was obtained (at a nanoparticle:alginate ratio of 7:1 or 4:1) and chitosan nanoparticles entrapped in alginate microparticles (around 280 mm) were successfully prepared. However, a major drawback of this technique is the loss of loaded compound from the nanoparticles during the process. Amaranth dye partitioned between the chitosan and alginate phases. Since the pKa of sodium alginate is between 3.4 and 4.4, this polymer was negatively charged under our conditions and may have competed with AR for cationic sites on chitosan and repelled the anionic charges of the dye. Borges et al. (2005) have described this phenomenon in the context of chitosan nanoparticles coated with sodium alginate, observing more than 60% of ovalbumin desorption when nanoparticles were added to sodium alginate solution. At pH 4, in the range of the pKa of alginate and hence equilibrium between negative and positive charges, rapid flocculation was observed. However, at pH 5, the competition between AR and polymer for chitosan cationic charges is reduced since there is a decrease of anionic charges of sodium alginate (dye desorption of 25%). The sodium alginate concentration can also modify the AR encapsulation efficiency. Indeed, the AR desorption in the medium is increased when nanoparticles suspension is mixed with the higher alginate concentration probably due to the presence of many anionic charges. A sodium alginate pH of 5 and a final concentration of 0.5% (nanoparticle:alginate ratio of 7:1) appeared to provide the smallest loss of dye. Under these process conditions, the size of particles remained near 280 mm. In the range of sodium alginate pH and concentrations studied, the viscosity of the mixed solution did not change much (results not shown) and did not appear to affect the microparticle formation process using the EncapsulatorÕ. The reduced loss of AR from the nanoparticles in the presence of the non-ionic surfactant Tween 80 could be due to interference with electrostatic interactions at the boundary between the anionic sodium alginate polymer and the cationic chitosan. The surfactant could also coat the nanoparticles and provide steric interference with dye diffusion. Since the long-term stability of the hydrated chitosan/alginate microparticles is poor at ambient temperature, freeze-drying was investigated as a preservation method. However, this process may cause particle aggregation, leading in some cases to irreversible fusion and increased size distribution (Abdelwahed et al., 2006). After freeze-drying with trehalose as a cryoprotectant (Dulieu and Bazile, 2005), our reconstituted microparticles showed a decrease in the average hydrodynamic diameter, an increased polydispersity index and deformities (data not shown). Using the non-ionic surfactant Tween 80 at a concentration of 1%, the
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chitosan/alginate microparticle physical and chemical characteristics (e.g. morphology and AR content) were preserved. It has been shown previously that low concentrations of this surfactant protect proteins entrapped in freeze-dried particles (Hillgren and Alden, 2002). Analysis by CLSM revealed a homogenous dispersion of nanoparticles throughout the alginate microparticles (Figure 5C and D). The association between RBITC and FITC fluorescence was intimate, as would be expected given that the strong electrostatic attraction should exist between the polymers due to opposite charges at the pH of the formulation. The slow release of AR into SGF (with or without pepsin) and microparticle shrinkage from 284 mm to about 200 mm with loss of porosity suggest that hydrogen bonding between alginate and chitosan hydroxyl groups was increased, which would be expected since the degree of ionisation of alginate is low at this pH, due to protonation of carboxylic groups (Chen et al., 2004). This condition reduced AR diffusion from the microparticles to about 5% over 6 h. Chitosan or chitosan/alginate particles have been considered unsuitable for oral delivery of therapeutic proteins (Chen et al., 2004) due to the rapid dissolution of chitosan at gastric pH. Indeed, protonation of the amino groups leads to chitosan chain repulsion, allowing diffusion of proton and counter ions along with water into the carrier and ultimately loss of particle integrity (Suksamran et al., 2011). By encapsulating chitosan nanoparticles inside alginate microparticles as in our study, polymer dissolution and drug release into the gastric environment are slowed. This micro-carrier system may, therefore, allow a pH-sensitive or polypeptide payload to pass through the stomach intact. At pH 7.4, the carboxyl groups of alginate are completely ionised (un-protonated), while chitosan amino groups are at most 9% protonated (Dambies et al., 2001). This would reduce hydrogen bonding and weaken the interaction between the two polymers, leading to rapid breakdown of the carriers and release of nanoparticles followed by release of the amaranth dye into the medium. However, dye release from chitosan nanoparticles in SIF is believed to occur by a diffusion process rather than erosion or dissociation of the polymeric matrix. At intestinal pH, the chitosan amino groups no longer attract the AR anionic groups, allowing dye release within 15 min. The release of this surrogate molecule from the nanoparticles is thus directly related to the surrounding pH. The low pH of the gastric environment guards against release, while the increased pH in the intestine allows it. However, this release pattern is likely a characteristic of our surrogate (which bears three anionic groups) and could be modified significantly in the case of a payload with a different ionic charge. Chitosan nanoparticles released in the small intestine should be able to interact with the negatively charged glycocalyx on the apical membrane of GIT cells. Since the pH in this environment is around 6.8, the nanoparticles should still bear enough positive charge to do so. Indeed, Behrens et al. (2002) have shown that intestinal cells interact more with chitosan than with polystyrene (a non-bioadhesive nanoparticle model). Moreover, chitosan is able to open the tight junctions between epithelial cells, thus facilitating the transport of carried molecules through the intestinal epithelium (Wong, 2009). This absorption-enhancing property is attributed in part to the capacity of chitosan to interact with negatively charged sites of ZO-1 proteins on cell surfaces, inducing redistribution of F-actin, a cytoskeleton protein regulating paracellular flow (Wong, 2010). Our nanometer-sized carrier could thus penetrate the intestinal mucosa within 30–60 min through intercellular spaces between enterocytes and M cells lining Peyer’s patches, as Carino and Mathiowitz (1999) have demonstrated with nanoparticles of sizes ranging from 10 to
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1000 nm (Carino and Mathiowitz, 1999). A compound that was not released too quickly from the nanoparticles could thus be delivered in the bloodstream to produce a therapeutic effect with greater efficiency.
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Conclusions The oral route of drug delivery is the most convenient and usually the safest and least expensive, as well as the one most often used. However, bioactive compounds or drugs advancing through the digestive tract may be degraded as they encounter gastric acid and/or digestive enzymes. There is, therefore, a need for carriers that protect pharmaceutical or nutraceutical payloads and facilitate their uptake by intestinal cells. In order to develop such a carrier, we chose two natural non-toxic polymers, namely chitosan and alginate. This is the first description of chitosan nanoparticles loaded with a compound entrapped in alginate microparticles. Using CLSM with different fluorescent labels for each of polymer, we were able to observe a homogenous distribution of both polymers throughout the microparticle matrix. The capacity of the carrier to control the release of the compound was then studied in a simulated gastric and intestinal environment with or without digestive enzymes. The less than 5% release of AR in the gastric environment suggests that this novel drug carrier and delivery system could protect pharmaceutical payloads until they reach the intestine, where release may be rapid and nearly complete, at least in the case of a molecule bearing a significant anionic charge. In such cases, the pH determines the strength of the electrostatic attraction binding the molecule to the carrier by varying the amount of positive charge on the chitosan chains. Meanwhile, the chitosan nanoparticles released in the intestinal environment remain intact. Because of their size, they are capable of crossing the intestinal boundary membrane and entering the bloodstream, where they could deliver drugs. These results confirm the possibility of using chitosan nanoparticles entrapped in alginate microparticles as a new drug delivery system, in particular to protect and enhance uptake of bioactive food-related compounds (e.g. polyphenolic compounds, vitamin D, curcumin) or poorly absorbed drugs (e.g. insulin, heparin, calcitonin). The surrogate compound (AR) used in the present study will be replaced in order to study candidate molecules identified for specific therapeutic targets.
Acknowledgements This study benefited from support provided by the De´le´gation Ge´ne´rale pour l’Armement (DGA) and by the Natural Sciences and Engineering Research Council of Canada (NSERC) Research Chair in Proteins, Biosystems and Functional Foods. The authors thank Claire Philippe for skilled technical assistance.
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