Chitosan/dextran multilayer microcapsules for polyphenol co-delivery

Materials Science and Engineering C 46 (2015) 374–380

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Chitosan/dextran multilayer microcapsules for polyphenol co-delivery Marco Paini a,b,⁎, Bahar Aliakbarian a,b, Alessandro A. Casazza a,b, Patrizia Perego a,b, Carmelina Ruggiero c, Laura Pastorino c a b c

Department of Civil, Chemical and Environmental Engineering, University of Genoa, via Opera Pia 15, 16145 Genoa, Italy Research Center for Biologically Inspired Engineering in Vascular Medicine and Longevity (BELONG), Via Montallegro 1, 16145 Genoa, Italy Department of Informatics, Bioengineering, Robotics and Systems Engineering, University of Genoa, Via Opera Pia 13, 16145 Genoa, Italy

a r t i c l e

i n f o

Article history: Received 16 September 2014 Accepted 21 October 2014 Available online 23 October 2014 Keywords: Nanostructured polymeric capsules Layer-by-layer Controlled release Polyphenols Acidic environment

a b s t r a c t Polysaccharide-based nanostructured polymeric microcapsules were fabricated by the electrostatic layerby-layer self-assembly technique and used to encapsulate mixtures of four different polyphenols in order to achieve their controlled release. The real-time fabrication of the dextran/chitosan multilayer was monitored by quartz crystal microbalance with dissipation monitoring, and the morphology of the nanostructured polymeric capsules was characterized by scanning electron microscopy. The polyphenol encapsulation was obtained by reversible permeability variation of the capsule shell in ethanol:water mixtures. The loading efficiency in different water:ethanol mixtures and the release rate in acidic conditions were characterized by UV spectroscopy and HPLC. The higher loading efficiency was obtained with an ethanol:water 35:65 phenolic solution, equal to 42.0 ± 0.6%, with a total release of 11.5 ± 0.7 mg of total polyphenols per 11.3 μL of microcapsules after 240 min of incubation in acidic environment. The results suggest that polysaccharide-based capsules can be successfully used to encapsulate and release low water-soluble molecules, such as polyphenols. © 2014 Elsevier B.V. All rights reserved.

1. Introduction Polyphenols are secondary metabolites present in plants. They are a large family of substances, ranging from simple molecules to complex structures [1]. These compounds show a wide spectrum of biological properties such as antioxidant, anti-inflammatory, antibacterial and antiviral activities [2]. Antioxidant properties make polyphenols potential therapeutic agents against serious diseases, like cancer, diabetes and cardiovascular disorders [3–8], acting against reactive oxygen species generated by exogenous chemicals or endogenous metabolism [9] and preventing cell damages caused by oxidative stress [10]. Several limitations have been associated with low bioavailability of polyphenols, including limited stability in environmental conditions, such as temperature, light, moisture, pH, oxygen concentration [11], low water solubility and rapid catabolism in the upper gastrointestinal tract and liver [12] and fast excretion through urinary system [13]. In vitro studies have shown that biological effects of polyphenols are extremely dose dependent and are evidenced at much higher concentrations than those present in natural sources [14]. Abbreviations: CAE, caffeic acid equivalents; CHI, chitosan; DEX, dextran sulfate; LE, loading efficiency; TPC, total phenolic concentration; NPC, nanostructured polymeric capsules;QCM-D, quartz crystal microbalance with dissipation monitoring;SEM,scanning electron microscopy. ⁎ Corresponding author at: Department of Civil, Chemical and Environmental Engineering, University of Genoa, via Opera Pia 15, 16145 Genoa, Italy. E-mail address: [email protected] (M. Paini).

http://dx.doi.org/10.1016/j.msec.2014.10.047 0928-4931/© 2014 Elsevier B.V. All rights reserved.

The development of effective encapsulation strategies of such molecules is therefore most desirable in order to fully exploit their therapeutic potential. Encapsulation in micro/nanoscale delivery systems can improve polyphenols half-life in vivo, preserving their biological activities, and can enhance their bioavailability. Moreover, encapsulation in a nanoengineered carrier can be used in order to achieve targeted delivery of the molecules into the diseased tissue with a controlled release profile [15]. Several encapsulation strategies for polyphenols have been proposed so far, mainly based on the use of liposomes, micelles, emulsions and spray drying techniques [16]. Engineering the structure, and thus the function, of the delivery system at the nanoscale resolution plays a pivotal role in the design of new successful treatment regimes. Moreover, cost is an important factor for the industrialization of such nanoformulation. In this framework, nanostructured polymeric capsules (NPC), obtained by the electrostatic layer-by-layer selfassembly technique, have been shown to possess great potentialities [17]. NPC are fabricated by the alternate adsorption of oppositely charged polyelectrolytes onto the surface of micro/nanoscale templates, usually carbonate particles [18]. Once the polyelectrolyte multilayer which constitutes the capsules shell has been deposited, the template is removed by dissolution in acidic medium or by chelating agents [19, 20]. By this technique it is possible to fabricate hollow polyelectrolyte capsules whose shell thickness ranges from few nanometers to tens of nanometers and with a predetermined composition, structure and thus functionality. An interesting property of such systems is the possibility to vary shell permeability as a consequence of a specific stimulus

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[21–25], usually pH, in order to load and, if required, to release cargo molecules. The assembly process is versatile and not very expensive, requiring only simple laboratory equipment. Therefore, NPC can be regarded as a very promising carrier for industrial scale-up. The main requirement to be satisfied for drug delivery systems is the use of fully biocompatible and biodegradable carriers. In this respect, the use of biopolymers having well characterized physicochemical properties is highly desirable. Natural polysaccharides have received great attention in the last few years, due to their unique features for applications in the field of drug delivery systems [26,27]. Polysaccharides are highly stable, safe, non-toxic and biodegradable, and can be easily chemically modified, resulting in a wide range of derivatives exhibiting different characteristics. Moreover polysaccharides, and specifically chitosan, have natural bioadhesive properties towards biological tissues that could prolong the residence time and therefore increase the absorbance of loaded drugs [28]. Chitosan, a copolymer of glucosamine and N-acetyl glucosamine, is a polycationic, biocompatible and biodegradable natural biopolymer mainly derived from the outer shells of crustaceans such as crabs and shrimps. Chitosan has different functional groups that can be modified with a wide range of ligands. Because of its properties, chitosan has great potential in biomedical applications, including drug delivery and tissue engineering [29]. Due to its cationic nature, chitosan is a good candidate for the layer-by-layer technique. One of the polysaccharides which displays complexing properties with chitosan is the polyanion dextran sulfate, which is obtained by the esterification of dextran using sulfuric acid. Several dextran sulfate/drug conjugates have been proposed as drug delivery systems [30]. In the present work, we describe the fabrication of biocompatible and biodegradable NPC, composed by cationic chitosan deposited in alternation with anionic dextran sulfate, and their use for the encapsulation of an ensemble of polyphenolic molecules for a synergistic effect. The deposition process and the structural properties of the chitosan/dextran sulfate multilayer were characterized by quartz crystal microbalance with dissipation monitoring (QCM-D). Then the assembly procedure was used for the deposition of the multilayers onto the surface of CaCO3 microparticles, followed by their removal under treatment with the chelating agent EDTA. The hollow NPC were then used for the encapsulation of polyphenols by means of the reversible permeability increase of their shell in ethanol: water mixtures containing polyphenols [31]. Loaded and unloaded NPC were structurally characterized by scanning electron microscopy. The polyphenol release in simulated gastric environment was characterized by means of UV–vis spectroscopy and HPLC. Finally, the influence of the thickness of the NPCs shell on the release rate was evaluated.

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2.2. Quartz crystal microbalance with dissipation monitoring The build-up of DEX/CHI multilayer was monitored by QCM-D (QCM-Z500, KSV Instruments, Helsinki, Finland). This technique has been extensively described [32,33], and allows to evaluate simultaneously the normalized resonant frequency (Δf) and energy dissipation shifts (ΔD) [32]. A quartz crystal with gold plated polished electrodes is excited at its fundamental frequency (5 MHz) and at the 3rd, 5th, 7th, 9th and 11th overtones (12, 25, 35, 45 and 55 MHz). As the mass is deposited onto the crystal surface, the oscillation frequency decreases. If the deposited mass is rigidly attached to the crystal, the frequency decrease is proportional to the mass and can be calculated using the Sauerbrey equation [34]. However, for viscoelastic materials the deposited mass introduces a dissipative energy damping. Using a Voigt-based model [35], the QCM-D response of a viscoelastic material can be modeled and the properties of added layers such as mass, density and thickness can be obtained. In this model, the adsorbed film is represented by a single Voigt element consisting of a parallel combination of a spring and dashpot to represent the elastic (storage) and inelastic (damping) behavior of a material, respectively. Before adsorption, the quartz crystals were cleaned with H2SO4 at 150 °C for 20 min followed by washing in pure water. A PTFE liquid chamber with a volume of 2 mL was used in the experiments. Polysaccharide solutions were alternatively introduced into the measurement chamber and left in contact with the quartz crystal. After each adsorption step, pure water was poured into the chamber and left in contact with the crystal for 1 min in order to remove the unabsorbed polysaccharides. The data analysis was performed using the QCM Impedance Analysis software (KSV Instruments, version 3.11). 2.3. NPC preparation NPC were assembled onto calcium carbonate sacrificial microparticles (6 μm in diameter), obtained by mixing calcium chloride and sodium carbonate solutions according to the reaction [36,37]: CaCl2 þ Na2 CO3 ¼ CaCO3 þ 2NaCl:

2. Materials and methods

108 CaCO3 microparticles were covered by successively deposited layers of anionic DEX and cationic CHI. Polysaccharides were left to adsorb onto the microparticle surface for 20 min, after each deposition step the dispersion of covered particles was centrifuged (2500 rpm for 5 min) and the precipitated covered particles were separated from the solution. These particles were washed three times in pure water, with successive centrifugation and separation steps. Four or eight bilayers were deposited onto the surface of the microparticle. Microparticles were then dissolved by their dispersion in EDTA solution at a concentration of 0.5 M at pH 7 followed by three washing steps in pure water. Knowing the diameter and the number of the prepared NPC, the internal volume of the NPC batch was calculated, resulting equal to 11.30 ± 0.90 μL.

2.1. Chemicals

2.4. NPC loading

Ethanol, methanol, acetic acid, acetonitrile, Folin–Ciocalteu reagent, medium MW chitosan (CHI), dextran sulfate sodium salt (DEX) from Leuconostoc spp. (MW 9000–20,000), NaCl, CaCO3, Na2CO3, ethylenediaminetetraacetic acid (EDTA) and standards of tyrosol, caffeic acid, vanillic acid and p-coumaric acid were purchased from Sigma-Aldrich (St. Louis, MO, USA). The polysaccharides were used as received. CHI and DEX solutions were prepared with a concentration of 0.5 mg/mL in NaCl 0.5 M. Chitosan was dissolved by addition of acetic acid to a final concentration of 0.3% (v/v) under continuous stirring overnight. Standards of tyrosol, caffeic acid, vanillic acid and p-coumaric acid were mixed and dissolved in ethanolic (20%, 35% and 50% v/v) solutions at a concentration of 2.5 mg/mL for each molecule.

The phenolic concentrations in the loading solutions were determined using the Folin–Ciocalteu assay [38]: 0.2 mL of diluted solution and 0.5 mL of Folin–Ciocalteu reagent were added to 4.8 mL of deionized water and, after mixing, 1 mL of a 20% Na2CO3 solution was added. Deionized water was added in order to reach a final volume of 10 mL. Solutions were mixed and incubated at room temperature in dark conditions for 1 h. Sample aliquots were used for the determination of total phenolic concentration (TPC) using a UV–vis spectrophotometer (Perkin Elmer, Wellesley, USA) at a wavelength of 725 nm. Concentrations were expressed as mg of caffeic acid equivalents (CAE) per mL. Caffeic acid was chosen as a standard due to its wide use as reference when working with phenolic compounds [39–41]. Absorbance

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Fig. 1. (a) Plot of surface coverage (ΔM/A, expressed in ng/cm2) as a function of layers (x axis) during the alternated adsorption of CHI and DEX. (b) Dissipation changes measured by QCMD during the build-up of a (CHI/DEX)4 multilayer. The data for the 3rd, 5th and 7th overtones are presented.

values were interpolated by a linear calibration curve (Eq. (1)) obtained with the same method by standard methanolic solutions of caffeic acid (10–1000 μg/mL), with an R2 of 0.996. ABS725

nm

¼ 0:002  TPC−0:004

ð1Þ

All loading solutions (35 mL) were prepared and immediately used for incubation with 11.30 ± 0.90 μL of NPC. Namely, NPC were exposed to loading solutions for 24 h with continuous stirring at 300 rpm in dark conditions at room temperature. During this period, the loading of phenolic compound was favored by both the presence of ethanol in solution and by the acidic environment generated by the presence of polyphenols. Afterwards, NPC were centrifuged, the supernatant was collected and pure water was added in order to close the NPC shells. The collected supernatant was used to calculate the loading efficiency (LE) according to Eq. (2).

LE ð%Þ ¼

  TPC−Csup =TPC  100

ð2Þ

where TPC and Csup are total phenolic concentrations of the initial solution and in the supernatant after the loading phase, respectively evaluated with the Folin–Ciocalteu assay. 2.5. Scanning electron microscopy (SEM)

Fig. 2. Scanning electron microscopy image of (DEX/CHI)4 NPC loaded in ethanol:water 35:65 (v/v) solution.

SEM measurements were carried out by a Zeiss Supra microscope. Samples were prepared dropping the solutions onto silicon supports with subsequent drying. To this purpose, silicon supports, having an area of 0.5 × 0.5 mm, were cleaned with H2SO4 at 150 °C for 20 min followed by extensive washing in pure water.

M. Paini et al. / Materials Science and Engineering C 46 (2015) 374–380 Table 1 TPC (expressed as mgCAE/mL) and amount of polyphenols (in mgCAE) released from nanocapsules (11.3 μL) prepared in solutions with different ethanol:water ratios after 120 min of incubation in water at pH 1.8.

H2O:EtOH 100:0 v/v H2O:EtOH 80:20 v/v H2O:EtOH 65:35 v/v H2O:EtOH 50:50 v/v

5.2 6.6 7.9 10.4

± ± ± ±

Table 2 Composition of loading solutions and release media (pH 1.8) after 240 min of incubation of nanocapsules expressed as single phenolic compounds (%) analyzed by HPLC.

Polyphenols released (mgCAE)

TPC (mgCAE/mL) 0.9a 0.2a,b 0.5b 0.2c

8.2 8.6 11.5 7.8

± ± ± ±

0.1a 0.1b 0.7b 1.4c

Different letters within the column indicate a significant difference between data at p b 0.05. CAE: caffeic acid equivalents; TPC: total phenolic concentration.

377

Tyrosol Caffeic acid Vanillic acid p-coumaric acid

Loading solution

Release after 240 min

H2O

H2O

32.50 12.58 32.83 19.39

EtOH:H2O 35:65 ± ± ± ±

0.83a 0.08b 0.10c 0.12d

29.44 18.75 28.94 22.87

± ± ± ±

2.66a 0.51b 0.57c 1.84c

17.59 32.64 27.29 22.48

EtOH:H2O 35:65 ± ± ± ±

1.99a 1.02b 1.88a,b 2.67a,b

46.63 20.50 20.87 12.01

± ± ± ±

1.88a 0.36b 1.50b 0.37c

Different letters within the column indicate a significant difference between data at p b 0.05. Values are means ± s.d. of three replicate analyses.

2.6. Polyphenol release from NPC in a gastric simulated environment The loaded NPC were centrifuged at 2500 rpm for 5 min and then resuspended in 5 mL of deionized water at pH 1.8. The pH was adjusted using HCl 37% w/v. At different times (0, 15, 30, 45, 60, 120, 180, 240 min), 1 mL of the solution was analyzed with a modified Folin– Ciocalteu assay to quantify the amount of total phenolic compounds. Briefly, 0.1 mL of Folin–Ciocalteu reagent and 0.2 mL of a 20% Na2CO3 solution were added to 1 mL of sample. Solutions were mixed and left at room temperature in the dark for 1 h. The total phenolic concentration (TPC) was calculated using the same spectrophotometer above. TPC was expressed as mgCAE per mL of acidified water. The method response was described with a linear equation (Eq. (3)), using standard methanolic solutions of caffeic acid (50–2000 μg/mL), with an R2 of 0.923. ABS725

nm

¼ 0:2071  TPC−0:0003

ð3Þ

100% B in 5 min, isocratic at 100% B for 5 min, followed by returning to the initial conditions (10 min) and column equilibration (12 min). The flow rate, the temperature of the column and the injection volume were 1 mL/min, 30 °C and 20 μL, respectively. The samples were detected at 280 nm. Before injection, samples were filtered through a 0.22 μm membrane filter. The concentration of each phenolic compound was calculated based on each standard solution. 2.8. Statistical analysis All experiments were carried out in triplicate. Analysis of variance (ANOVA) and Tukey's post hoc test (p b 0.05), using “Statistica” software version 8.0 (StatSoft, Tulsa, USA), were used. The statistically significant differences were shown in tables and figures using different letters.

2.7. HPLC analysis

3. Results and discussion

The initial loading solutions and samples after 240 min of incubation in acidic environment were analyzed via HPLC (Hewlett Packard, 1100 Series, Palo Alto, CA, USA), equipped with a C18 reverse-phase column (Model 201TP54, Vydac, Hesperia, CA, USA) coupled with a UV–vis detector, as described by [42]. The mobile phase was water/acetic acid (99:1%, v/v) as solvent A and methanol/acetonitrile (50:50%, v/v) as solvent B, while the solvent gradient changed according to the following conditions: from 5% to 30% B in 25 min, from 30% to 40% B in 10 min, from 40% to 48% B in 5 min, from 48% to 70% B in 5 min, from 70% to

3.1. Quartz crystal microbalance with dissipation monitoring Fig. 1 shows the build-up of a (DEX/CHI)4 multilayer. The increase in surface coverage (Fig. 1a) with each adsorption step indicates that mass is deposited, while ΔD (Fig. 1b) is almost constant showing a strong interaction among layers. It could also be observed that the CHI harmonics are separated, while for DEX they are superimpose, indicating that CHI has a more viscoelastic behavior than DEX. The total thickness of (DEX/CHI) bilayer was calculated to be about 9 nm.

14.00 12.00

mgCAE

10.00 8.00 6.00 4.00

2.00 0.00 0

30

60

90

120

150

180

210

240

t (min) Fig. 3. Release profiles of nanocapsules loaded with phenolic compounds solubilized in water (solid line) and ethanol:water 35:65 (v/v) (dashed line). Results are expressed as mg of CAE (caffeic acid equivalents).

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2.50

mgCAE

2.00

1.50

1.00

0.50

0.00

0

30

60

90

120

150

180

210

240

t (min) Fig. 4. Release profiles of 8-bilayer nanocapsules loaded with phenolic compounds solubilized in an ethanol:water 35:65 (v/v) solution. Results are expressed as mg of CAE (caffeic acid equivalents).

3.2. NPC preparation and loading The protocol used for the deposition of the multilayer onto the quartz crystal surface was used for the deposition onto CaCO3 microparticles. Four bilayers were deposited, and then the microparticles were removed to obtain hollow NPC which were loaded with polyphenols in ethanol:water mixtures as previously described. Fig. 2 shows the SEM image of one loaded (DEX/CHI)4 NPC. The real concentration (TPC) of loading solutions with different ethanol:water ratios was calculated with the Folin–Ciocalteu assay, and the amount of polyphenols loaded in the NPC was evaluated after 120 min of incubation at pH 1.8, as shown in Table 1. It may be noted that TPC increased with increasing fraction of ethanol in the solvent mixture, until complete dissolution with ethanol: water ratio 50:50, with 10.4 ± 0.2 mgCAE/mL (approximately equal to initial theoretical TPC). Increasing the solubility of polyphenols led to an increased amount of loaded polyphenols, from 8.2 ± 0.1 mgCAE in absence of ethanol to 11.5 ± 0.7 mgCAE for an ethanol:water ratio of 35:65. However, a further increase in the ethanol fraction up to 50% v/v resulted in a decrease in the amount of released phenolic compounds. The reversible reorganization of polyions in water and ethanol: water mixture resulting in a permeability change was already reported for synthetic polyelectrolytes [37,43]. However, the mechanism of pore creation is not fully understood. As it relates to polysaccharide networks, the presence of ethanol in the solution allows polysaccharide molecules to come closer together, interact and form “clumps” [36]. We reckon that an increase in the ethanol fraction up to 50% v/v resulted in large defects in the NPC shell, which could be not fully reversed when redispersed in pure water. For the subsequent analyses, a solvent with ethanol:water ratio 35:65 (v/v) was selected which resulted to maximize solubilization of phenolic compounds and minimize structural instability of nanocapsules. The LE of the nanocapsules has been evaluated comparing the phenolic concentrations of the supernatant after the first centrifugation step and the corresponding TPC before the loading phase. For the sample loaded in water environment, LE is 29.6 ± 2.0%, while in the presence of ethanol at 35% v/v LE significantly increases (p b 0.05) to 42.0 ± 0.6%. This increase can be explained considering both the higher solubility of phenolics in aqueous-ethanol loading solution and the shell structure modification induced by ethanol.

The behavior of the NPC in acidic environment, simulating gastric fluids, was evaluated at subsequent times of incubation. Fig. 3 shows the release trends of phenolic compounds from nanocapsules loaded in absence and in presence of ethanol (35% v/v) in the loading solution. Polyphenols released rapidly in the first 30 min for the nanocapsules loaded in absence of ethanol, and in the first 60 min for the capsules loaded in presence of ethanol. An initial release at t0 was detected, probably due to the polyphenols adherent to the external layer of the NPCs. The total amount of polyphenols released is higher when ethanol is present in the loading solution (11.6 ± 0.1 mgCAE after 240 min of incubation), as a direct consequence of the increased polyphenol dissolution in the solvent mixture. In absence of ethanol in the loading solution, the maximum amount of polyphenols encapsulated in the nanocapsules was statistically significantly lower (p b 0.05) and equal to 9.2 ± 0.1 mgCAE after 240 min of incubation. At the end of the incubation, acidic solutions and initial loading solutions (pure water and ethanol:water 35:65 v/v) were analyzed by HPLC, in order to evaluate the content of the phenolic components in the loading solutions and in the release media (Table 2). These results confirm the limited solubilities of some phenolic species when pure water is used as solvent: caffeic acid and p-coumaric acid represent only 12.58 ± 0.08% and 19.39 ± 0.12% of the phenolic compounds in solution, respectively, both lower than the theoretical 25%. The addition of ethanol led to an increase of the percentage of these compounds to 18.75 ± 0.51% and 22.87 ± 1.84%, respectively. The concentrations of all the phenolic species are about 25%. After 240 min of release, different behaviors have been noted using different solvents: with pure water, the percentage of tyrosol released from NPC is sensibly lower (p b 0.05) than that of other compounds, equal to 17.59 ± 1.99% of the released polyphenols, while in presence of ethanol this compound has the highest percentage. This may be explained considering that both polyphenols and NPC have different steric and electric properties in the two solvents, which can lead to different selectivities with respect to tyrosol and to acidic species (caffeic, vanillic and p-coumaric acids) during the loading phase. Finally NPC having the shell architecture (DEX/CHI)8 were fabricated and loaded in order to check the possibility of increasing the release time by increasing the shell thickness. Fig. 4 shows the release profile of the 8-bilayer capsules in acidic environment (pH 1.8). The release profile for (DEX/CHI)8 NPC was found to be similar to the one for (DEX/CHI)4 NPC, with a complete release in the first hour

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of incubation. However, the amount of phenolics released is definitely lower, equal to 2.0 ± 0.1 mg after 240 min of incubation. This can be explained considering that phenolics have to cross a higher number of layer in order to reach the hollow core of the capsule, which lead to a lower LE, equal to 21.2 ± 2.7%. Considering these results, increasing the number of bilayers is not advisable, neither to delay the release nor to load a higher amount of phenolic compounds in nanocapsules. 4. Conclusions Polyphenol mixtures, composed of four different molecules, were encapsulated in polysaccharide based microcapsules. The cationic polysaccharide chitosan and the anionic polysaccharide dextran sulfate were used for the nanostructured capsule formation in a layerby-layer fashion. QCM-D was employed to characterize the stepby-step deposition of the polysaccharide multilayer. The influence of different binary mixtures of ethanol and water on the loading efficiency of the microcapsules was evaluated in order to find the optimal loading conditions. Release rates in acidic environment were determined by UV spectroscopy and HPLC and the influence of the shell thickness was also taken into account. Our results suggest that polysaccharide capsule can be used to encapsulate mixtures of low water-soluble molecules, such as polyphenols, in order to obtain a synergistic effect. References [1] A. Munin, F. Edwards-Lévy, Pharmaceutics 3 (2011) 793–829. [2] B. Aliakbarian, A.A. Casazza, P. Perego, Food Chem. 128 (2011) 704–710. [3] B. Aliakbarian, D. Palmieri, A.A. Casazza, D. Palombo, P. Perego, Nat. Prod. Res. 26 (2012) 2280–2290. [4] J.A. Baur, D.A. Sinclair, Nat. Rev. Drug Discov. 5 (2006) 493–506. [5] D. Delmas, A. Lancon, D. Colin, B. Jannin, N. Latruffe, Curr. Drug Targets 7 (2006) 423–442. [6] M.C. Desco, A. Asensi, R. Marquez, J. Martinez-Valls, M. Vento, F.V. Pallardo, J. Sastre, J. Vina, Diabetes 51 (2002) 1118–1124. [7] M.S. Jang, E.N. Cai, G.O. Udeani, K.V. Slowing, C.F. Thomas, C.W.W. Beecher, H.H.S. Fong, N.R. Farnsworth, A.D. Kinghorn, R.G. Mehta, R.C. Moon, J.M. Pezzuto, Science 275 (1997) 218–220. [8] A. Kumari, S.K. Yadav, Y.B. Pakade, V. Kumar, B. Singh, A. Chaudhary, S.C. Yadav, Colloids Surf. B 82 (2011) 224–232. [9] S.R. Georgetti, R. Casagrande, C.R.F. Souza, W.P. Oliveira, M.J.V. Fonseca, LWT Food Sci. Technol. 41 (2008) 1521–1527. [10] A. Moure, J.M. Cruz, D. Franco, J.M. Dominguez, J. Sineiro, H. Dominguez, M.J. Nunez, J.C. Parajo, Food Chem. 72 (2001) 145–171. [11] Z.X. Fang, B. Bhandari, Food Chem. 129 (2011) 1139–1147. [12] S. Das, K.Y. Ng, J. Pharm. Sci. US 99 (2010) 840–860. [13] M.R. Serrano-Cruz, A. Villanueva-Carvajal, E.J.M. Rosales, J.F.R. Davila, A. DominguezLopez, LWT Food Sci. Technol. 50 (2013) 554–561. [14] T.G. Shutava, S.S. Balkundi, P. Vangala, J.J. Steffan, R.L. Bigelow, J.A. Cardelli, D.P. O'Neal, Y.M. Lvov, ACS Nano 3 (2009) 1877–1885. [15] S. Mura, J. Nicolas, P. Couvreur, Nat. Mater. 12 (2013) 991–1003. [16] Z.X. Fang, B. Bhandari, Trends Food Sci. Technol. 21 (2010) 510–523. [17] L. Pastorino, S. Erokhina, V. Erokhin, Curr. Org. Chem. 17 (2013) 58–64. [18] G.B. Sukhorukov, E. Donath, S. Davis, H. Lichtenfeld, F. Caruso, V.I. Popov, H. Mohwald, Polym. Adv. Technol. 9 (1998) 759–767. [19] A.A. Antipov, G.B. Sukhorukov, E. Donath, H. Mohwald, J. Phys. Chem. B 105 (2001) 2281–2284. [20] G.B. Sukhorukov, H. Mohwald, Trends Biotechnol. 25 (2007) 93–98. [21] S. Erokhina, L. Benassi, P. Bianchini, A. Diaspro, V. Erokhin, M.P. Fontana, J. Am. Chem. Soc. 131 (2009) 9800–9804. [22] S. Erokhina, O. Konovalov, P. Bianchini, A. Diaspro, C. Ruggiero, V. Erokhin, L. Pastorino, Colloids Surf. A 417 (2013) 83–88. [23] L. Pastorino, S. Erokhina, F. Caneva-Soumetz, C. Ruggiero, J. Nanosci. Nanotechnol. 9 (2009) 6753–6759. [24] L. Pastorino, S. Erokhina, F.C. Soumetz, P. Bianchini, O. Konovalov, A. Diaspro, C. Ruggiero, V. Erokhin, J. Colloid Interface Sci. 357 (2011) 56–62. [25] N. Habibi, L. Pastorino, F.C. Soumetz, F. Sbrana, R. Raiteri, C. Ruggiero, Colloids Surf. B 88 (2011) 366–372. [26] Z.H. Liu, Y.P. Jiao, Y.F. Wang, C.R. Zhou, Z.Y. Zhang, Adv. Drug Deliv. Rev. 60 (2008) 1650–1662. [27] G. Saravanakumar, D.G. Jo, J.H. Park, Curr. Med. Chem. 19 (2012) 3212–3229. [28] H. Liu, B. Chen, Z.W. Mao, C.Y. Gao, J. Appl. Polym. Sci. 106 (2007) 4248–4256.

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Marco Paini has obtained his graduation in Biotechnology in 2009 and his M.Sc. degree in Industrial Biotechnology in 2011 from the University of Modena and Reggio Emilia, Italy. He is currently a Ph.D. student in Material, Process and Chemical Engineering in the Department of Civil, Chemical and Environmental Engineering of the University of Genoa, Italy. His main research interests include antioxidant extraction from food industry residues, encapsulation techniques and food engineering.

Bahar Aliakbarian, Ph.D. in Material, Process and Chemical Engineering, is a post-doc fellow in the Food Technology and Biotechnology Laboratory in the Department of Civil, Chemical and Environmental Engineering of the University of Genoa, Italy. Her studies focused on extraction and evaluation of the biological properties of antioxidant compounds, nutraceutical and functional food development, encapsulation techniques and tissue engineering. She maintains scientific relationships with important international universities, and during her research she authored more than 30 international scientific papers and participated at several international conferences.

Alessandro Alberto Casazza, Ph.D. in Material, Process and Chemical Engineering, is a post-doc fellow in the Food Technology and Biotechnology Laboratory in the Department of Civil, Chemical and Environmental Engineering of the University of Genoa, Italy. His studies focused on the extraction of antioxidant compounds from food industry residues through conventional and non-conventional technologies. He also worked on the lipid extraction from microalgae and on biomass pyrolysis for energy production. His main research activity is documented by 26 scientific papers and 7 abstracts published in international journals, 2 contributions in volume and participations in several international conferences.

Patrizia Perego is a Professor of Chemical Plants and Deputy Dean of the Polytechnic School in the University of Genoa (Italy), Department of Civil, Chemical and Environmental Engineering. She authored over 100 international scientific papers and several contributions in volumes, and she maintains scientific relationships with important national and international universities and companies. Her research interests include nutraceutical and functional food development, valorization of food industry residues, technologies for food preservation, industrial process optimization and production and characterization of probiotics.

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M. Paini et al. / Materials Science and Engineering C 46 (2015) 374–380 Carmelina Ruggiero received the M.Sc. degree in Electrical Engineering from University of Genoa, Genoa, Italy, in 1971 and the Ph.D. and DIC degrees from Imperial College, London, U.K., in 1977. She is currently Professor of Bioengineering at University of Genoa and Head of the Bioengineering and Medical Informatics Laboratory at the Department of Informatics, Bioengineering, Robotics and Systems engineering (DIBRIS), University of Genoa. Her main research interests include nanobioscience, computer modeling, biomedical signals, and image analysis.

Laura Pastorino received the M.Sc. degree in Chemical Engineering (1999) and the Ph.D. in Biophysics (2003) from the University of Genoa, Genoa, Italy. She is currently an assistant professor of Bioengineering at the Department of Informatics, Bioengineering, Robotics and Systems Engineering, University of Genoa, Genoa, Italy. Her research interests include nanobiotechnology for drug delivery and tissue engineering.