Structural analysis of chitosan hydrogels containing polymeric nanocapsules

    Structural analysis of chitosan hydrogels containing polymeric nanocapsules Renata V. Contri, Rosane M.D. Soares, Adriana R. Pohlmann, Silvia S. Guterres PII: DOI: Reference:

S0928-4931(14)00261-6 doi: 10.1016/j.msec.2014.05.001 MSC 4625

To appear in:

Materials Science & Engineering C

Received date: Revised date: Accepted date:

5 February 2014 31 March 2014 3 May 2014

Please cite this article as: Renata V. Contri, Rosane M.D. Soares, Adriana R. Pohlmann, Silvia S. Guterres, Structural analysis of chitosan hydrogels containing polymeric nanocapsules, Materials Science & Engineering C (2014), doi: 10.1016/j.msec.2014.05.001

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ACCEPTED MANUSCRIPT STRUCTURAL ANALYSIS OF CHITOSAN HYDROGELS CONTAINING

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POLYMERIC NANOCAPSULES

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Guterresa

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Renata V. Contria*, Rosane M. D. Soaresb, Adriana R. Pohlmanna,b, Silvia S.

Programa de Pós-Graduação em Ciências Farmacêuticas, Universidade

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b

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Federal do Rio Grande do Sul, Porto Alegre, Brazil;

Instituto de Química, Universidade Federal do Rio Grande do Sul, Porto Alegre,

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*Corresponding author:

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Brazil.

Dr. Renata Vidor Contri

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Faculdade de Farmácia, Universidade Federal do Rio Grande do Sul, Av. Ipiranga, 2752/405 CEP 90610-000, Porto Alegre, RS, Brazil. Phone: 55 51 33085215 Fax: 55 51 33085247 E-mail: [email protected]

E-mail addresses for other authors: [email protected] [email protected] [email protected]

ACCEPTED MANUSCRIPT Abstract The incorporation of different concentrations of polymeric nanocapsule

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suspensions into chitosan hydrogels is proposed, in order to study the structure

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of a formulation with the properties of great tissue adhesion and controlled

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release of the nanoencapsulated drugs, represented here by capsaicinoids. The gels presented acceptable acid pH values and the nanoparticles were visually

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observed in the system. A transition from the micrometer to the nanometer scales suggested that the nanocapsules are initially agglomerated in the hydrogel. A

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sedimentation tendency of the nanocapsules in the system was observed and only physical interaction between the chitosan chains and polymeric

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nanocapsules were verified. The hydrogels, despite the presence of

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nanocapsules, presented shear-thinning properties and an elastic behavior under low and high frequencies, showing a very structured gel network. The observed

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variation in the elasticity of the hydrogels may arise from a decrease in the number of interactions and degree of entanglement between the chitosan chains,

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caused by the presence of nanoparticles. Keywords: chitosan hydrogel, nanocapsules, physical characterization, structure.

ACCEPTED MANUSCRIPT 1. Introduction Chitosan, a cationic biopolymer, has been widely applied in the

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development of differentiated pharmaceutical forms, such as polymeric films,

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hydrogels and beads, which are useful in distinct research areas [1,2,3]. Its

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advantageous properties include biodegradability, bioadherence, wound healing promotion and a bacteriostatic effect [4,5], besides a possible effect on the tight

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junctions between epithelial cells [6], leading to enhancement in the permeation of active compounds into biological tissues. Chitosan hydrogels can be formed

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by simple acidification of the aqueous medium and further entanglement of the chitosan chains due to secondary interactions [2]. The gel characteristics are

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strongly dependent on the chitosan molecular weight and degree of

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deacetylation [7]. Moreover, the addition of crosslinkers brings new properties to the hydrogels, such as thermal and pH responses (physical crosslinking) or a

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very structured hydrogel network (chemical crosslinking) [2,3]. The incorporation of polymeric nanocapsules into chitosan hydrogels is a

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promising system recently proposed by our research group [8]. This innovative system combines the previously-mentioned chitosan properties with the advantages associated with nanocapsules, such as the control of the drug release [9,10], as observed in the controlled release of nanoencapsulated capsaicinoids [8]. The capsaicinoids found in chili peppers present topical analgesic action, due to depletion of P substances, and recently they have been used to treat chronic pain [11]. Since capsaicin and dihydrocapsaicin, which are the major capsaicinoids, are irritant substances and many applications are required to achieve the desired effect [11], their nanoencapsulation could

ACCEPTED MANUSCRIPT increase the patient’s compliance, by means of a decrease in the release rate [8] and in the irritation effect [12].

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In the first work describing the association of polymeric nanocapsules

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and chitosan gel [8], the incorporation of cationic polymeric nanocapsules

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containing capsaicinoids into the referred gel led to a slight increase in the hydrogel viscosity, measured immediately after preparation, suggesting that the

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presence of the nanoparticles has an effect on the chitosan gel structure. In addition, the chitosan hydrogel containing nanocapsules did not show a further

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increase in viscosity during storage, as noted for the chitosan hydrogel containing pure water. Although interesting properties of the novel formulation

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has been already described such as controlled release [8], great skin adhesion

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[13] and decrease on the skin irritation [12], the elucidation of the hydrogel structure in the presence of nanocapsules has not been studied so far, and it is

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of great importance in order to explain such changes in the gel behavior. The incorporation of liposomes [14] and lipid nanoparticles [15] into

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chitosan hydrogels, as well as a rheological study on these types of formulations, has been previously described. The presence of liposomes slightly increased the gelation rate and gel strength of chitosan thermosensitive hydrogels [14]. Also, Souto and co-workers [15] produced a 1% chitosan hydrogel containing solid lipid and nanostructured lipid nanoparticles. Their results showed that pure hydrogels presented a weaker and more sensitive structure when compared to nanoparticle-containing hydrogels. Based on the exposed, the aim of this study was to, by adding different nanocapsule amounts to the hydrogels, investigate the structure of chitosan

ACCEPTED MANUSCRIPT hydrogels containing capsaicinoid-loaded polymeric nanocapsules, as well as to determine the influence of the addition of nanoparticles on chitosan hydrogel

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networks.

2. Materials and Methods

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2.1 Materials

Chitosan (medium molecular weight, deacetylation degree of 75%) and

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Eudragit RS 100® were obtained from Degussa (Darmstadt, Germany) and Sigma-Aldrich (São Paulo, Brazil), respectively. Polysorbate 80, capric/caprylic

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triglycerides and lactic acid 85% were purchased from Labsynth (São Paulo,

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Brazil), Brasquim (Porto Alegre, Brazil) and Via Farma (São Paulo, Brazil).

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Acetonitrile was purchased from Tedia (Fairfield, USA). The capsaicinoids used as model drugs for nanoencapsulation were obtained from Deg (São Paulo, Brazil) (58% of capsaicin and 33% of dihydrocapsaicin). The acetone used was

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purchased from Vetec (Rio de Janeiro, Brazil) and was of analytical grade. 2.2 Production of capsaicinoids-loaded nanocapsules The capsaicinoid-loaded nanocapsules were prepared through the interfacial deposition of the preformed polymer [16], as previously described [8]. Briefly, an organic phase containing 100 mg of Eudragit RS 100 ®, 5 mg of capsaicinoids, 27 mL of acetone and 330 µL of capric/caprylic triglyceride was injected, applying a controlled rate, into an aqueous phase containing 76 mg of polysorbate 80. After mixing for 10 min, the solvents were eliminated using a reduced-pressure rotary evaporator (Büchi) for adjustment of the final volume

ACCEPTED MANUSCRIPT (10 mL). Eudragit RS 100®, the selected polymer, has already been used for the obtainment of nanocapsules for the cutaneous [8], vaginal [17] and ocular

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[18] administration routes.

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2.3 Incorporation of nanocapsules into the chitosan hydrogel

In order to obtain chitosan hydrogels containing nanocapsules, different percentages of nanocapsule suspension and/or water (Table 1) were added to

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the chitosan, which was present at a final concentration of 3%. Prior to use, the chitosan was sieved (sieve number 355) to guarantee a homogeneous system.

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The dispersion was acidified through the addition of lactic acid to give a final concentration of 1%. Manual mixing led to the obtainment of hydrogels. The

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formulations presented theoretical final concentrations of capsaicinoids between

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0.15 (hydrogel B) and 0.5 mg.g-1 gel (hydrogel D).

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2.4 Characterization of the chitosan hydrogels containing nanocapsules 2.4.1 pH value

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The pH values of the hydrogels were determined by potentiometry (potentiometer B474, Micronal) after dilution of the semi-solid formulations in Milli Q® water (1:10 w/v). Data are represented as the mean of three independent experiments. 2.4.2 Particle size distribution The hydrogels were characterized in terms of particle size distribution by laser diffraction (Mastersizer®, Malvern), after their dispersion in distilled water, using 1.38 and 1.52 as the refraction indexes of Eudragit RS 100® (Gels B, C and D) and chitosan (Gels A, B, C and D), respectively. Both refraction indexes

ACCEPTED MANUSCRIPT were used for the data analysis in order to observe the behavior of the nanoparticles and also of the chitosan hydrogel microdomains.

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2.4.3 Zeta potential

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The zeta potential values of the final formulations were determined after dispersion of the hydrogels in a filtered 10 mM NaCl solution (20 mg of gel in 10 mL, vortex mixing). The data are represented as the mean of three different

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measurements.

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2.4.4 Physical stability

The physical stability of the hydrogels was determined by the multiple

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light scattering technique (Turbiscan LAB®, Formulation, France). Each sample

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(around 15 g) was placed into a cylindrical glass cell (25 mm diameter, 55 mm

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height) at 25 oC. The glass cells were completely scanned at time intervals of 5 min for 1 h immediately after preparation of the samples, and also after 5, 10 and 30 days under storage in the same glass cells at 25oC. The transmitted and

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backscattered light as a function of the cell height was obtained during this time interval.

2.4.5 Oscillatory rheology In order to determine the rheological behavior of the chitosan hydrogels, preliminary strain-sweep experiments were carried out to guarantee the linear viscoelastic regime. Oscillatory frequency-sweep measurements were then performed at angular frequencies ranging from 10-2 to 103 Hz at a constant temperature (23.0 ± 0.2 °C) to record the elastic storage modulus (G’), and the viscous loss modulus (G”). All experiments were conducted in a rheometer

ACCEPTED MANUSCRIPT (model MCR 101, Anton-Paar Physica, Germany), with a cone-plate geometry (25 mm in diameter). Steady shear flow curves were also obtained for all of the

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2.4.6 Fourier transform infrared (FT-IR) spectroscopy

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chitosan hydrogel formulations under the same temperature conditions.

The hydrogels under investigation were cast into petri dishes and dried to produce films suitable for FT-IR spectroscopy analysis. The spectra for the films

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were acquired in a Perkin-Elmer PC-16 spectrophotometer, and recorded with 4

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cm-1 of resolution (16 scans) in the range of 4000 to 400 cm. 2.4.7 Differential scanning calorimetry (DSC)

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The thermal properties of the hydrogels were evaluated using a

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differential scanning calorimeter (DSC series Q1000 - TA Instruments),

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operating at a heating rate of 10 oC min-1 from -10 to 300 °C under N2 atmosphere. The weight of each sample was around 9 mg. After the first heating cycle, the samples were cooled to 0 °C and scanned again with heating

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up to 300 °C at the same heating/cooling rate. DSC curves of each hydrogel were obtained from the second heating run at a rate of 10 oC min-1 (after the first run with heating up to 120 ºC and cooling to 25ºC at the same rate of 10 oC min-1). In the present study, the Tg value for Eudragit RS 100® was calculated considering the onset temperature of the change in the heat capacity during the thermal event. 2.4.8 Microscopic evaluation The hydrogel macrostructures were analyzed by optical microscopy (Olympus AX70 photomicroscope), immediately after spreading thin hydrogel

ACCEPTED MANUSCRIPT layers on the glass blades. The presence of the nanocapsules and their morphology after incorporation into the hydrogels were analyzed, after complete

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drying of the water on a smooth glass surface, by scanning electron microscopy

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- SEM (JSM 5800, Exll, Japan, operating at 10kV), at the Electron Microscopy

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Center of the Federal University of Rio Grande do Sul State, Brazil.

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3. Results and Discussion

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3.1 pH value

The pH values observed were 4.52, 4.29, 4.40, and 4.37, for hydrogels

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A, B, C and D, respectively. These relatively low values are due to the lactic

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acid used for the chitosan solubilization. After the addition of lactic acid, the

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amine groups (NH2) of the saccharide become NH3+ and the chitosan becomes soluble in the aqueous medium. As previously described [8], the addition of nanocapsules (pH = 5.65) did

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not affect the pH values of hydrogels. The pH values of the formulations were considered suitable for cutaneous use, since the value measured for the stratum corneum is between 4.0 and 4.5 [19]. 3.2 Particle size distribution The results obtained in the laser diffraction analysis, considering the refraction index of chitosan are shown in Fig. 1. Right after sample addition to the equipment [Fig. 1(a)], all gels presented a micrometric peak, probably due to the presence of microdomains in the chitosan hydrogel network. After 5 min of sample addition to the equipment container [Fig. 1(b)], which maintained the

ACCEPTED MANUSCRIPT sample under constant mixing with distilled water at room temperature, the formulations maintained the micrometric size distribution, however, Hydrogel D

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presented a different profile, showing two micrometric populations. This profile

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suggests a phase separation with two distinct domains. Such domains might

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contain different nanoparticle concentrations, leading to different size distributions.

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The results obtained in the laser diffraction analysis, considering the refraction index of Eudragit RS100 are shown in Fig.2. The results acquired

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immediately after sample addition to the equipment [Fig. 2(a)] showed a micrometric size distribution for the gels, while the nanocapsules in suspension

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showed a nanometric peak. After 5 minutes under mixing in the sample

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apparatus [Fig. 2(b)], a significant change from micrometer to nanometer scale was detected over time for the hydrogel particles. This size transition verifies

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that nanocapsules were present as agglomerates in the hydrogel network and that deagglomeration promptly occurs in aqueous medium. This behavior shows

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that particles were stable after their incorporation into the hydrogel network. In order to gain a better understanding of this phenomenon, the percentage of nanoparticles or microparticles was determined at pre-determined times between 0 and 300 seconds (20 sec, 1 min, 2 min and 5 min), in a mixture with distilled water, and they are shown in Table 2. Interestingly, the presence of particles in the nanometric range was firstly observed for the hydrogel containing the greatest amount of nanocapsules (Gel D). On the other hand, for the hydrogel containing the lowest nanocapsule concentration (Gel B) it took longer for nanoparticles to be observed. This may

ACCEPTED MANUSCRIPT be attributed to the easy dispersion of the agglomerates even when the nanocapsules are present in higher concentration within the hydrogels.

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Nevertheless, a higher concentration of nanocapsules in the hydrogel matrix did

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not lead to non-dispersible nanocapsule agglomerates. This is probably due to

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a stronger repulsion effect when more nanocapsules were present in the matrix. Nanocapsules tend to be repulsed since they present cationic charge, due to the use of Eudragit RS 100® as the polymeric wall. Moreover, the agglomeration

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of nanocapsules, as observed from the laser diffraction results, can disturb the

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chitosan chain entanglement, which is mainly observed for hydrogel D (Figs. 1(a) and (b)). In this regard, the disturbance of the chitosan hydrogel network might have facilitated the dissolution process of the gel and, as consequence,

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3.3 Zeta potential

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the obtainment of nanocapsules with their primary particle size.

The mean zeta potential values measured were 31.4 mV, 32.2 mV and 34.7 mV, for hydrogels B, C and D, respectively. However, the zeta potential of

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the nanocapsules in aqueous suspension was lower (+ 9.62 mV) when compared to the particles dispersed in the hydrogel matrix. This is explained by the cationic character of the chitosan structure. There was a slight increase in the zeta potential (from 31 to 35 mV) as more nanocapsules were added to the hydrogel. This finding is in agreement with the increase in the charge repulsion and a faster transition from nanoparticle agglomerates to the primary particle size. 3.4 Physical stability of hydrogels

ACCEPTED MANUSCRIPT The hydrogels containing nanocapsules did not present a transmittance signal (T), so backscattering profiles were analyzed. In order to visualize the

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related to the first measurement were determined (Fig. 3).

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backscattering variations as a function of time, backscattering signal variations

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The relative profiles showed a decrease in the backscattering signal on the top of the cell, which was proportional to the nanocapsule concentration in

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the hydrogel (hydrogel D > hydrogel C > hydrogel B). Therefore, a tendency toward the sedimentation of the nanocapsules in the semi-solid formulation was

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demonstrated. In this regard, the nanocapsule agglomerates are freely dispersed in the hydrogel network, without chemical interactions occurring

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between the chitosan chains, which would decrease the sedimentation process.

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This result was expected since the chitosan and the nanocapsules have

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positive charges, leading to a tendency toward repulsion. The presence of a higher amount of nanocapsules enhanced the sedimentation process. This observation corroborates the hypothesis that the

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nanocapsules may disturb the chitosan chain entanglements increasing the deposition at the bottom of the cell. The addition of higher amounts of nanocapsules (gel D) might even lead to phase separation into two distinct chitosan microdomains containing different nanocapsule concentrations, as described in section 8.3.2. 3.5 Oscilatory rheology Figure 3 shows the steady shear flow curves for all chitosan hydrogel formulations. All chitosan hydrogels showed pseudoplastic (shear-thinning) behavior. Chitosan chains can form a hydrogel containing ionizable groups due

ACCEPTED MANUSCRIPT to intermolecular Coulomb repulsion, hydrogen bonds, polar forces, and mobile ions, with the development of a high swelling pressure, as noted by several

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authors [20,21,22].

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In our systems, the movement of the entangled chains in solution was

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probably restricted by the close proximity of the neighboring chains. In other words, when the shear rate is increased the subsequent viscosity decrease can

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be explained by the disentanglement of these polymer chains due to the reptation process.

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As the nanoparticle content in the system increased (from hydrogels B to D), reorganization of the intermolecular and/or intramolecular interactions

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appear to have occurred. This behavior is observed for hydrogels A to C (Fig.

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4), where the nanoparticles may have interrupted, to some extent, the chitosan-

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chitosan interactions. Also, the more pronounced shear-thinning behavior (and higher low-shear viscosity) of hydrogel D might be due to an increase in the hydrodynamic friction and a reorganization of the hydrophobic interactions, both

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induced by the agglomeration of Eudragit RS 100® nanoparticles. However, for the intermediate formulations (hydrogel B and C), the nanoparticle concentration was not high enough to induce this disturbance of the chitosan gel. In this case, the flow curves correspond mainly to the free movement of the polymer chains through the system, undisturbed by their neighbors. To elucidate these phenomena, frequency sweep experiments were carried out to investigate the viscoelastic properties of the systems and to establish the three-dimensional structure of these physically crosslinked gel

ACCEPTED MANUSCRIPT networks (Fig. 5). In general, both moduli increased continuously as a function of frequency (Figs. 5 (a) to (d)), indicating the formation of a well-developed

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tridimensional network. The variation in the elasticity of the hydrogels may arise

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from a decrease in the number of interactions and entanglements between the

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chitosan molecules, caused by the presence of the nanoparticles. The differences in the variation in the G’ and G” values led to a crossover

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point in terms of frequency, signifying the transition from liquid-like to a solid-like behavior. Also, at this point, where G’ = G” (known as the gel point), the system

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is considered to have the longest relaxation time. In other words, at this point the energy release needed to return the system to its equilibrium state after

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being strained or deformed is at its lowest value.

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This gelation profile demonstrated that chitosan can behave like a

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hydrogel itself, even in the absence of Eudragit RS 100® nanocapsules. Furthermore, after the crossover point, G’ becomes higher than G” indicating a

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stable and structured hydrogel (i.e. solid-like behavior). Another approach to investigating the stiffness of gels is by considering the loss tangent (tan ), which can be calculated as the G”/G’ ratio at a certain frequency (Fig. 5 (e)). As the oscillation frequency is increased, tan  is reduced continuously for all hydrogels, indicating that the systems become more rigid. After the crossover point (i.e. tan  = 1), however, this stiffening is more noticeable for the hydrogels containing the nanoparticles. Once more, we can state that the Eudragit RS 100® nanocapsules produced more rigid gel structures, which is directly proportional to the content of this component in the chitosan matrix.

ACCEPTED MANUSCRIPT Another interesting parameter related to the gel structure is the relaxation time. This parameter can be calculated as the inverse of the crossover

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frequency [23,24] highlighted in the crossover region. The crossover

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frequencies (Hz) and the calculated relaxation times (sec) observed for each

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chitosan hydrogel were 63.7 Hz and 15.7 sec (Gel A), 66.6 Hz and 15.0 sec (Gel B), 56.1 Hz and 17.8 sec (Gel C) and 51.5 Hz and 19.4 sec (Gel D).

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According to the literature [21,25], polymeric chain segments between crosslinking points are longer in the less densely crosslinked networks and,

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thus, will give rise to lower molecular motion frequencies when compared to those arising from more densely crosslinked networks. However, at higher

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frequencies long chains fail to rearrange themselves in the time scale of the

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imposed motion and, therefore, they stiffen, assuming a more “solid-like” behavior. This stiffening is usually characterized by a sharp increase in G’,

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because shorter polymer chains resulting from densely crosslinked networks exhibit shorter relaxation times. Based on this, we can assume that the neat

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chitosan hydrogel presented a more densely crosslinked network and shorter relaxation time when compared to the other hydrogels. As the content of Eudragit RS 100® nanoparticles increased (from gel B to D), the relaxation time of all hydrogels studied increased and this behavior suggests longer chain segments between the crosslinking points. However, this understanding may be seen to be inconsistent with the shear viscosity and loss tangent behaviors observed. To elucidate this apparent contrast, firstly, two important factors should be considered: chitosan chain entanglement and the presence of nanoparticles in the systems. In addition, the proportion of water available to promote the

ACCEPTED MANUSCRIPT formation of physical junctions (which stabilize the hydrogel network) can play a significant role in terms of the gel structure.

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According to the literature, chitosan must satisfy some specific conditions

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to form a stable polymer network: (i) interchain interactions must be strong

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enough to form semi-permanent junction points in the molecular network; and (ii) the network must be capable of exchanging water molecules with the

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hydrated polymer network [7, 26]. However, these interactions are purely physical and gel formation can be reversed. These aspects corroborate the fact

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that chitosan and its hydrogels showed some differences in the shear viscosity and gel stiffness behavior, probably due to some structural differences and

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variations in the water availability, which directly affect the hydrogel network.

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Moreover, these two diverse behaviors were detected under different conditions, i.e., under shear flow or in oscillatory experiments. The first

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condition was not suitable for measuring the gel stiffness because under these experimental conditions the gel networks were disrupted and the shear

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viscosities were governed only by the flow of the polymeric chains. On the other hand, the G’ and G” values reflect the mechanical moduli of a small amplitude oscillatory movement, which, in our case, can be related only to the gel stiffness and not to hydrodynamic contributions of the nanoparticles in the chitosan matrix. Another significant aspect is that, although physically-bonded hydrogels have the advantage of forming a gel without the use of chemical crosslinking moieties, they can be easily disrupted under shear forces, as demonstrated herein. Chitosan hydrogels with better mechanical properties can be produced using chemicals that produce irreversible polymeric crosslinks. The properties

ACCEPTED MANUSCRIPT of chemically crosslinked hydrogels are mainly dependent on the crosslinking density and the ratio of crosslinker molecules to moles of polymer repeating

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units [26, 27].

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3.6 FT-IR spectroscopy

Regarding the physical interaction between the chitosan and the nanoparticles, the FT-IR spectroscopy analysis (Fig. 6) showed the main

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absorption peaks related to chitosan, Eudragit RS 100® and hydrogels A, B, C and D. As can be observed, as the water content in the hydrogel decreases,

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there was an absorption shift to a lower wavenumber, probably due to the decrease in the hydrogen bonds in the hydrogel. The absorption band at 1158.4

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cm−1 (anti-symmetric stretching of the C–O–C bridge) is assigned to the ether

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bond of chitosan. The peaks at 1016 and 1040.0 cm −1 (skeletal vibrations

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involving C–O stretching) are shown for the asymmetric stretching of the C-O-C bridge. The basic structural features of chitosan are shown at 3241 cm−1 (–OH and –NH2 stretching), 2869 cm−1 and 2800 cm−1 (–CH stretching), 1664 cm−1 (–

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NH2 stretching), 1076cm−1 (C–O–C stretching) and 796 cm−1 (pyranoside ring stretching vibration). The spectrum for Eudragit RS 100®, which is a copolymer of acrylate and methacrylate, is also shown in Fig. 5. The peaks at 1319 cm -1 and 1020 cm-1 are mainly related to C-O double bond stretching mode. The peak at 1600 cm -1 is related to C=O stretching. Interestingly, for the chitosan hydrogel-loaded nanoparticles the peak at 1600 cm-1 shifts, indicating that the hydrogen bonding increased in the presence of chitosan. According to the FT-IR spectra and main absorption peaks, it can be deduced that there was only physical interaction

ACCEPTED MANUSCRIPT between chitosan and Eudragit RS 100® nanoparticles, showing that no covalent chemical bonds or undesirable reactions took place during the

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hydrogel formulation. As mentioned previously, this lack of chemical interaction

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was expected and was one of the reasons for choosing Eudragit RS 100® as

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the nanoparticle polymer. 3.7 DSC

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The thermal analysis showed two main endothermic peaks for hydrogels A, B, C and D (Fig. 7). For comparison purposes, neat chitosan (without being

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prepared as a hydrogel) and Eudragit RS 100® were also analyzed (Fig. 6). Chitosan showed only one endothermic peak, which is associated with a

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crystalline phase transition (Tm = 136.8 ºC). On the other hand, Eudragit RS

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100® presented a glass transition (Tg) at 55 ºC, in agreement with results

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published in the literature [28,29] and also an endothermic peak at Tm = 190.8

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The first endothermic peak in all of the DSC curves for the hydrogels (A, B, C and D) appeared at lower temperatures when compared to neat chitosan; however, the second peak for all hydrogels appeared at higher temperatures. As can be observed, for all hydrogels the water-plasticized chitosan is assumed to appear either in the absence or presence of nanoparticles. Therefore, the Tm values for the hydrogels B, C and D shifted to higher temperatures as the nanocapsule content in the network hydrogels increased, suggesting that the nature and concentration of the water affects the mobility of the polysaccharide chains. Thus, an enhancement of the molecular movement due to dissociation

ACCEPTED MANUSCRIPT of the hydrogen bonds and the beginning of chain scission are expected, due to the variation in the nanocapsule content in the hydrogel.

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Since chitosan presents both amine and hydroxyl groups in the glycoside

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ring, the associations between water and the hydroxyl and amine groups could

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differ in terms of their intensity, and the release of water molecules might preferentially occur from the amine group. Furthermore, the water content can

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also vary according to the degree of deacetylation (DD) and is higher for a lower DD, according to results reported in the literature [7,28].

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3.8 Microscopic evaluation

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The hydrogel macrostructures were visually observed by optical

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microscopy (40 times magnification, data not shown). The presence of clusters could be observed as the concentration of nanocapsules in the hydrogel matrix

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was increased (hydrogels C and D). This is probably related to the nanoparticle agglomerates in the hydrogel formulation. In addition, it was determined by the

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size distribution study that the nanoparticle agglomerates are around 1000 micrometers.

The SEM photomicrographs of the films obtained from the gels are shown in Figure 8. It is possible to observe spherical nanocapsules in the films obtained from hydrogels B, C and D. The nanoparticles can be observed as agglomerates at the hydrogel sample surface. In hydrogel D, it is also possible to see nanocapsules separated from each other. Thus, no damage is caused to the nanocapsules due to the incorporation of the hydrogel or the hydrogel casting to obtain films. The film surfaces became rougher with the addition of

ACCEPTED MANUSCRIPT the nanocapsules. Also, pores can be seen on the surface of the films obtained from gels containing nanocapsules (hydrogels B, C and D).

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3.9 Proposal of hydrogels structures

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Based on all of the results obtained in this study, the hydrogel microdomain structures are proposed in Figure 9. Figure 9(a) shows hydrogel A with a strongly crosslinked structure, without the presence of nanocapsules.

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Figure 9(b) shows hydrogel B with the presence of dense nanoparticle agglomerates, slightly disrupting the chitosan chain entanglement. Figure 9(c)

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shows hydrogel C with the presence of less dense nanoparticle agglomerates, higher in number, disrupting the chain entanglement to a greater extent. Figure

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9(d) shows hydrogel D which, due to the excessive amount of nanocapsules

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microdomains.

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and a reorganization of the chains, presents phase separation in the

4. Conclusion

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The nanocapsules are in the reversible agglomerated form in the innovative formulation proposed, recovering the primary size in aqueous medium. A greater concentration of nanoparticles led to partial disruption of the chitosan chain entanglement up to a concentration where there was a reorganization of the system and a phase separation in the hydrogel microdomains occured. So, it is possible to, by regulating the amount of nanocapsules in the gel, obtain different structures. Although there was a decrease in the number of interactions and entanglements between the chitosan molecules, caused by the presence of nanoparticles, the formation of a well-developed tridimensional network was observed for all cases.

ACCEPTED MANUSCRIPT Acknowledgments: RVC thanks CNPq/Brazil for her fellowship. The authors are grateful to CNPq/MCTI/Brazil, PRONEX and PRONEM FAPERGS/CNPq,

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FINEP, CAPES (Rede Nanotecnologia Farmacêutica) for the financial support.

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References

[1] M.N.V. Ravi Kumar, A review of chitin and chitosan applications. React. Functional Polymers 46 (2000) 1-27.

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[2] J. Berger, M. Reist, J.M. Mayer, O. Felt, N.A. Peppas, R. Gurny, Structure

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and interactions in covalently and ionically crosslinked chitosan hydrogels for biomedical applications, Eur. J. Pharm. Biopharm. 57 (2004) 19-34.

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[3] J. Berger, M. Reist, J.M. Mayer, O. Felt, R. Gurny, Structure and interactions

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in chitosan hydrogels formed by complexation or aggregation for biomedical

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applications, Eur. J. Pharm. Biopharm. 57 (2004) 35-52. [4] Rinaudo, M. Chitin and chitosan: properties and applications. Prog. Polym.

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Sci. 31 (2006) 603-632.

[5] R.A.A. Muzzarelli, Chitins and chitosans for the repair of wounded skin, nerve, cartilage and bone, Carbohydr. Polymers. 76 (2009) 167-182. [6] G. Ranaudi, I. Marigliano, I. Vespignani, G. Perozzi, Y. Sambuy, The effect of chitosan and other polycations on tight junction permeability in the human intestinal Caco-2 cell line, J. Nutritional Biochem. 13 (2002) 157-167. [7] M. Dasha, F. Chiellini, R.M. Ottenbriteb, E. Chiellin, Chitosan—A versatile semi-synthetic polymer in biomedical applications, Prog. Polym. Sci. 36 (2011) 981-1014.

ACCEPTED MANUSCRIPT [8] R.V. Contri, T. Katzer, A.R. Pohlmann, S.S. Guterres, Chitosan hydrogel containing capsaicinoids-loaded nanocapsules: An innovative formulation for

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topical delivery, Soft Mater. 8 (2010) 370-385.

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[9] S.S. Guterres, M.P. Alves, A.R. Pohlmann, Polymeric nanoparticles,

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nanospheres and nanocapsules for cutaneous applications, Drug Target Insights, 2 (2007) 147-157.

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[10] C.E. Mora-Huertas, H. Fessi, A. Elaissari, Polymer-based nanocapsules for

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drug delivery, Int. J. Pharm. 385 (2010) 113–142. [11] M. Hayman, P.C.A. Kam, Capsaicin: A review of its pharmacology and

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clinical applications, Curr. Anaesth. Crit. Care, 19 (2008) 338-343.

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[12] R.V.Contri, L.A. Frank, M. Kaiser, A.R. Pohlmann, S.S. Guterres, The use of nanoencapsulation to decrease human skin irritation caused by

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capsaicionoids, Int. J. Nanomed. 9 (2014) 951 – 962. [13] R.V. Contri, T. Katzer, A.F. Ourique, A.L.M. da Silva, R.C.R. Beck, A.R.

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Pohlmann, S.S. Guterres, Combined effect of polymer nanocapsules and chitosan gel on the increase of capsaicinoids adhesion to the skin surface, J. Biomed. Nanotechnol. 10 (2014) 820-830. [14] E. Ruel-Gariépy, G. Leclairb, P. Hildgenb, A. Guptac, J-C Leroux, Thermosensitive chitosan-based hydrogel containing liposomes for the delivery of hydrophilic molecules. J. Control. Release. 82 (2002) 373-383. [15] E.B. Souto, S.A. Wissing, C.M. Barbosa, R.H. Müller, Evaluation of the physical stability of SLN and NLC before and after incorporation in hydrogel formulations, Eur. J. Pharm. Biopharm. 58 (2004) 83-90.

ACCEPTED MANUSCRIPT [16] H. Fessi, F. Puisieux, J-Ph. Devissaguet, N. Ammoury, S. Benita, Nanocapsule formation by interfacial polymer deposition following solvent

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displacement, Int. J. Pharm. 55 (1989) R1-R4.

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[17] S.S. Santos, A. Lorenzoni, L.M. Ferreira, J. Mattiazzi, A.I.H. Adams, L.B.

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Denardi, S.H. Alves, S.R. Schaffazick, L. Cruz, Clotrimazole-loaded Eudragit® RS100 nanocapsules: Preparation, characterization and in vitro evaluation of

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antifungal activity against Candida species. Mater. Sci. Eng. C. 33 (2013) 1389– 1394.

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[18] T. Katzer, P. Chaves, A. Bernardi, A.R. Pohlmann, S.S. Guterres, R.C.R. Beck, Prednisolone-loaded nanocapsules as ocular drug delivery system:

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Accepted article.

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development, in vitro drug release and eye toxicity, J. Microencapsulation.

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[19] M. Denda, J. Hosoi, Y. Asida, Visual imaging of íon distribution in human epidermis, Biochem. Biophys. Res. Commun. 272 (2000) 134-137.

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[20] S. Abashzadeh, M.H. Hatimiri, F. Atyabi, M. Amini, R. Dinarvand, Novel physical hydrogels compesed of opened ring poly(vinyl pyrrolidone) and chitosan derivatives: Preparation and characterization, J. Appl. Polym. Sci. 121 (2011) 2761-2771. [21] Y. Fan, Y. Liu, J. Xi, R. Guo, Vesicle formation with amphiphilic chitosan derivatives and a conventional cationic surfactant in mixed systems, J. Coll. Interface Sci. 360 (2011) 148-153.

ACCEPTED MANUSCRIPT [22] R. Wang, H. Hu, X. He, W. Liu, H. Li, Q. Guo, L. Yuan, Synthesis and characterization of chitosan/urea formaldehyde shell microcapsules containing

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dicyclopentadiene. J. Appl. Polym Sci. 121 (2011) 2202-2212.

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[23] K.I. Draget, B.T. Stokke, Y. Yuguchi, H. Urakawa, K. Kajiwara, Small-Angle

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X-ray Scattering and Rheological Characterization of Alginate Gels. 3. Alginic Acid Gels, Biomacromolecules. 4 (2003) 1661-1668.

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[24] B.H. Hu, J. Su, P.H. Messersmith, Hydrogels Cross-Linked by Native

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Chemical Ligation, Biomacromolecules, 10 (2009) 2194–2200. [25] S. Argin-Soysal, P. Kofinas, L.O. Martin, Effect of complexation conditions

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on xanthan–chitosan polyelectrolyte complex gels, Food Hydrocoll. 23 (2009)

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202-209.

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[26] G.Q. Yinga, W.Y. Xionga, H. Wanga, Y. Suna, H.Z. Liub, Preparation, water solubility and antioxidant activity of branched-chain chitosan derivatives. Carbohydr. Polymers. 83 (2011) 1787-1796.

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[27] H. Park, K. Park, W.S.W. Shalaby, Biodegradable hydrogels for drug delivery, Technomic Publishing Company, Pennsylvania, 1993. [28] J. Fujimori, Y. Yoshihashi, E. Yonemochi, K. Terada, Application of Eudragit RS to thermo-sensitive drug delivery systems, J. Control. Release 102 (2005) 49-57. [29] N. Pearnchob, R. Bodmeier, Dry polymer powder coating and comparison with conventional liquid-based coatings for Eudragitw RS, ethylcellulose and shellac., Eur. J. Pharm. Biopharm. 56 (2003) 363-369.

ACCEPTED MANUSCRIPT Figure Captions Figure 1. Size distribution of chitosan hydrogels, considering the refraction

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index of chitosan, (a) right after placement in sample container and (b) after 5

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minutes of mixing in the sample container.

Figure 2. Size distribution of chitosan hydrogels, considering the refraction index of Eudragit RS 100® (a) right after placement in sample container and (b)

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after 5 minutes of mixing in the sample container.

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Figure 3. Light backscattering analyses of chitosan hydrogels: (a) Hydrogel B (b) Hydrogel C and (c) Hydrogel D.

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Figure 4. Shear viscosity () as a function of shear rate () for the chitosan

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hydrogels.

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Figure 5. Storage (G’) and loss (G”) moduli as a function of frequency for the chitosan hydrogels [(a) Hydrogel A, (b) Hydrogel B (c) Hydrogel C and (d)

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Hydrogel D] and (e) Loss tangent (tan) as a function of frequency for all chitosan hydrogels. Figure 6. FT-IR spectra of chitosan hydrogels (A, B, C and D), Eudragit RS100 ® and chitosan. Figure 7. DSC thermograms for (a) chitosan hydrogels (A, B, C and D) and (b) Eudragit RS 100® and chitosan (2nd heating cycle). Figure 8. Scanning electron photomicrographs of chitosan films obtained from (a) Hydrogel A, (b) Hydrogel B, (c) Hydrogel C and (d) Hydrogel D (Increase in 10,000 times the real size).

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Hydrogel B, (c) Hydrogel C and (d) Hydrogel D.

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ACCEPTED MANUSCRIPT Table 1. Composition of chitosan hydrogels (the percentages are related to the liquid phase used to formulate each gel).

Nanocapsule

Nanocapsule

liquid

suspension (% of liquid

concentration (x1013

100%

0%

B

70%

30%

C

50%

50%

D

0%

100%

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nanoparticles. g )

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phase)

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phase)

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Hydrogels

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Water (% of

Capsaicinoids concentration (mg. g-1)

---

---

0.81

0.15

1.35

0.25

2.70

0.5

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100

0

100

20

100

0

91

40

64

36

29

60

33

67

25

80

26

74

120

15

85

240

14

86

300

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100

0

9

26

74

71

5

95

75

4

96

6

94

4

96

6

94

3

97

5

95

3

97

5

95

2

98

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Nanometer Micrometer Nanometer Micrometer Nanometer

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Gel C

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Gel B

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Graphical abstract

ACCEPTED MANUSCRIPT Highlights The polymeric nanocapsules are initially agglomerated in the chitosan

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hydrogels.

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The nanocapsules caused a decrease in the degree of chitosan chains entanglement.

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High amounts of nanocapsules led to a reorganization of the microdomains. Sedimentation of nanocapsules and physical interaction with chitosan were

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verified.

The gels presented shear-thinning and elastic properties, besides structured

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network.