Effect of chitosan coating on the properties of nanoliposomes loaded with flaxseed-peptide fractions: Stability during spray-drying

Journal Pre-proofs Effect of chitosan coating on the properties of nanoliposomes loaded with flaxseed-peptide fractions: stability during spray-drying Khashayar Sarabandi, Seid Mahdi Jafari PII: DOI: Reference:

S0308-8146(19)32091-6 https://doi.org/10.1016/j.foodchem.2019.125951 FOCH 125951

To appear in:

Food Chemistry

Received Date: Revised Date: Accepted Date:

12 January 2019 6 November 2019 23 November 2019

Please cite this article as: Sarabandi, K., Mahdi Jafari, S., Effect of chitosan coating on the properties of nanoliposomes loaded with flaxseed-peptide fractions: stability during spray-drying, Food Chemistry (2019), doi: https://doi.org/10.1016/j.foodchem.2019.125951

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Effect of chitosan coating on the properties of nanoliposomes loaded with flaxseed-peptide fractions: stability during spray-drying

Running title: Spray-drying stabilization of chitosan-coated nanoliposome

Khashayar Sarabandi, Seid Mahdi Jafari*

Faculty of Food Science & Technology, Gorgan University of Agricultural Sciences and Natural Resources, Gorgan, Iran *Corresponding

author, TelFax: +98 17 324 26 432. Email address: [email protected] (SM Jafari). *Corresponding

author Graphical abstract.

Highlights

Loading of bioactive peptide fractions into nanoliposomes was performed. Nanoliposomes coated with 0.4% chitosan had high physical stability and EE. FTIR/SEM showed entrapment of liposomes in reservoir-type and irregular structures. Chitosan coating leads to the retention of the spray-dried nanoliposomes.

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Abstract In this study, production of peptide fractions with different molecular weights (10, 30, and 100 KDa) from flaxseed hydrolysates was investigated. The peptide-fraction with lowest MW (10 KDa) showed the highest antioxidant activity (AA); therefore, selected and loaded into nanoliposomes. Then, the effect of chitosan coating (0.1-0.8% w/v) on the physicochemical features, stability, and release rate of loaded nanoliposomes was assessed. Nanoliposomes coated with 0.4% chitosan exhibited a higher physical stability and encapsulation efficiency (EE) compared with other samples. Then, stabilization of uncoated/coated nanoliposomes by spray-drying within the maltodextrin matrix was investigated. FTIR results indicated incorporation of peptides into the polar and non-polar regions of vesicles and formation of the hydrogen bonds with 1

phosphatidylcholine. Chitosan coating significantly enhanced the physical stability, EE, and AA retention of nanoliposomes after powder reconstitution process by increasing the bilayer membrane rigidity against the dehydration tension and decreasing the leakage of loaded peptides. Keywords: Chitosan coating; nanoliposomes; stabilization; spray-drying; chemical structure

1. Introduction The importance and healthful benefits of the bioactive compounds (such as vitamins, phytosterols, polyphenols, fatty acids, peptides and hydrolysates, enzymes, probiotics, and anthocyanins) have led to the increasing tendency towards the use of these compounds in the production of various types of functional beverages and food products. In contrast, there are some disadvantages for these compounds, including low solubility, bioavailability, and biostability, physicochemical incompatibility and instability in the environmental conditions, food products and the digestive system, and undesirable flavor and odor, which are among the challenges that complicate the shelflife and direct use of these compounds in food systems (Sarabandi, Mahoonak, Hamishekar, Ghorbani, & Jafari, 2018; Shishir, Xie, Sun, Zheng, & Chen, 2018; Udenigwe, 2014). Therefore, numerous studies have been conducted in recent years on the design and production of various delivery systems with the aim of enhancing the bioavailability, maintaining the biological activity, and stabilizing these bioactive compounds. The most important delivery systems are the lipid nanocarriers such as nanoliposomes (Sarabandi, Mahoonak, Hamishehkar, Ghorbani, & Jafari, 2019; Akhavan, Assadpour, Katouzian, & Jafari, 2018). The properties of nanoliposomes include the application of food grade compounds in their formulation as well as the capability of encapsulation of various hydrophilic, hydrophobic, and amphiphilic bioactive compounds such as the essence oils, essential oils, vitamins, polyphenols, fatty acids, phytosterols, proteins, peptides, enzymes, and anthocyanins (da Rosa Zavareze et al., 2014; Katouzian & Jafari, 2016; Mohan, McClements, & Udenigwe, 2016; Ramezanzade, Hosseini, & Nikkhah, 2017). However, the thermodynamic instability and processes such as agglomeration, 2

coalescence, flocculation, precipitation, high release rate, and microbial contamination are among the major challenges of the colloidal systems (Faridi Esfanjani, Assadpour, & Jafari, 2018; Yousefi, Ehsani, & Jafari, 2019). These disadvantages reduce the shelf life and result in the leakage, and loss of loaded compounds (Huang, Su, Zheng, & Tan, 2017; McClements, 2018). One of the methods to increase the physicochemical and microbial stability of bioactive-loaded carriers is to remove their water content and convert them into the powdered form. Freeze-drying (Alemán, Marín, Taladrid, Montero, & Gómez-Guillén, 2019; Stark, Pabst, & Prassl, 2010) and spray-drying (Jafari, Assadpoor, He, & Bhandari, 2008; Tewa-Tagne, Briançon, & Fessi, 2007) are the most common methods of powder production and increasing the stability of delivery systems. However, given the high cost and the long time of freeze-drying process, the use of spray-drying method as an economical, flexible, and high-speed process can be regarded to be a suitable and efficient alternative (Akbarbaglu et al., 2019; Assadpour & Jafari, 2019; Sarabandi, Jafari, Mahoonak, & Mohammadi, 2019). Despite the application of fillers such as maltodextrin (as a protective matrix to trap the nanoparticles), processes such as thermal stress and dehydration during the drying process can damage the vesicular membrane and cause a substantial loss of the loaded compounds after the liposomal reconstitution (Wang et al., 2015). Therefore, developing approaches aimed at increasing the membrane rigidity against heat during the spray-drying process is of great importance. Some studies have examined the effect of coating nanoliposomes containing blackberry extract (Gültekin-Özgüven, Karadağ, Duman, Özkal, & Özçelik, 2016) and cacao hull extract (Altin, Gültekin-Özgüven, & Ozcelik, 2018) with chitosan as well as coating with calcium alginate (Wang et al., 2015) on the stability of nanocarriers during the spray-drying process. In this study, the bioactive peptides obtained from enzymatic hydrolysis of flaxseed protein were selected as the core material. The importance of these peptides is in their various pharmaceutical, antioxidant, and health properties (Sarmadi & Ismail, 2010). Given these properties, flaxseed proteins are considered as additives in the formulation of various dietary supplements and production of the functional products (Akbarbaglu et al., 2019). In addition, the peptide fractions 3

with different molecular weights have different antioxidant power and biological activity (Hosseini, Ramezanzade, & Nikkhah, 2017; Mohan et al., 2016). Therefore, in this study, selection of the optimum fraction for loading in the nanoliposomes and its effect on the stability of system were considered. The behavior and effect of loading the peptide fractions within the nanocarriers vary given the differences in the functional characteristics and the complex structure of the antioxidant peptides with other bioactive compounds such as polyphenols. To the best of our knowledge, the effect of chitosan coating on enhancing the stability of the nanoliposomes containing flaxseed peptide-fractions has not been evaluated before and after the spray-drying solidification process. Considering the above issues, the objectives in this study included: 1) Coating of nanoliposomes containing flaxseed peptide fractions with different concentrations of chitosan and selecting the optimum treatment; 2) Evaluation of the physicochemical characteristics, stability, and release rate of the nanocarriers; and 3) Investigation of the effect of chitosan coating on the physical, functional, and stabilization features, encapsulation efficiency (EE) retention, antioxidant activity (AA), chemical structure (FTIR), and morphology (SEM) of the nanoliposomes containing the flaxseed peptide fractions before and after the spraydrying process. 2. Materials and methods The defatted-flaxseed meal (cold pressed) was purchased from local market. The chemicals including Alcalase 2.4 L. (Novo Nordisk, Bagsvaerd, Denmark), ABTS (2,2′-azino-bis (3ethylbenzothiazoline-6-sulfonic acid) diammonium salt), DPPH (1,1-Diphenyl-2-picrylhydrazyl), chitosan (medium molecular weight) and comasi brilliant blue (G250) were purchased from Sigma– Aldrich Co. (St. Louis, MO, USA). Lecithin (lipoid, Germany), Alpha-deoxyribose was obtained from Fluka (Stockholm, Sweden). Potassium persulphate and Tween-80 were obtained from Merck (Darmstadt, Germany). Maltodextrin (DE18) was purchased from Pooran powder (Isfahan, Iran). Other chemicals used were of analytical grade. 2.1. Flaxseed protein extraction and preparation of protein hydrolysates 4

The preparation, extraction, and production of the protein concentrate from flaxseed meal were carried out in accordance with the method described by Akbarbaglu et al. (2019). Briefly, the mucilage removal process (with 0.5M NaHCO3 solution at 50 °C for 1 h), separation of the remaining oil from the meal flour (with hexane at a ratio of 1:4 for 1 h), extraction (in the solution with pH = 10), protein precipitation (after adjusting the mixture pH to 4.2 with 0.5 M HCl), neutralization (with 0.5 M NaOH up to pH 7), and finally freeze-drying (Christ, Germany) were performed. The dissolution process of flaxseed protein concentrate (at a concentration of 5% w/v) was conducted in 0.01 M phosphate buffer saline for 30 min. Then, the enzymatic hydrolysis of proteins was performed with alcalase (pH = 8, 50 ˚C) at 2% enzyme to substrate ratio for 4 h. The activity of the enzymes was stopped after transferring the reaction medium to a water bath (95 ˚C for 15 min). The resulting dispersion was centrifuged (7,000 × g for 10 min) and the supernatant containing the hydrolysate was separated (Akbarbaglu et al., 2019). 2.2. Separation of bioactive-peptide fractions The fractionation process of hydrolysates was performed using ultrafiltration membrane (Amicon ultra centrifugal filter, Millipore, UK) with molecular weight cut-off of 10, 30 and 100 KDa. The supernatant containing hydrolysates were passed through filters with 100 KDa (PF-100 with MW=30-100 KDa), 30 KDa (PF-30 with MW=10-30 KDa) and 10 KDa (PF-10 with MW<30 KDa), respectively. The resulting peptide fractions were freeze-dried and stored at -18 ˚C until experiments. 2.3. Antioxidant characterization 2.3.1. DPPH free radical scavenging A sample of 1.5 mL peptide-containing solution (10 mg/mL) was mixed with 1.5 mL of 0.2 mM DPPH solution, and then the resulting solution was kept in the dark for 30 min. In the next step, the mixture was centrifuged (4,000 × g, 10 min) and the supernatant adsorption was read at 517 nm

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(Sarabandi et al., 2018). The DPPH free radical scavenging was calculated using the following equation:

Inhibition  %  1   sample Abs /blank Abs    100 (1) 2.3.2. ABTS free radical scavenging The ABTS solution was prepared from the ABTS and potassium persulfate mixture with a final concentration of 7 mM and 2.45 mM, respectively. After 16 h of storage in the dark, the mixture was diluted with 0.2 M PBS (pH 7.4) (absorbance of 0.70 at 734 nm). Then, 30 µL of the peptide solution (10 mg/mL) was added to 3 mL of the ABTS solution, vortexed for 30 s, and kept in the dark for 6 min. The absorbance was then read at 734 nm. In order to determine the Trolox equivalent antioxidative capacity (TEAC), a standard curve of the reaction of different concentrations of Trolox (50-1000 μM) with ABTS was plotted (Akbarbaglu et al., 2019). 2.4. Preparation and chitosan coating of loaded nanoliposomes The production of nanoliposomes was carried out according to Sarabandi et al. (2019). 0.09 g of phosphatidylcholine, 0.01 g of cholesterol, and 1 drop (0.02 g) of Tween-80 were dissolved in 10 mL of absolute ethanol for 20 min. The resulting solution was transferred to a round-bottom flask and the solvent evaporation process and the formation of a thin layer were performed at 60 ˚C and the rotational speed of 70 rpm using a rotary evaporator (Laborota 4002, Heidolph, Germany). Complete removal of the solvent residues was performed after 16 h of storage in the desiccator. For hydration of the thin layer and for the formation of the liposomes, 10 mL of the peptide-fraction solution (5 mg/mL) was added to the film. The resulting mixture was vortexed for 2 min and stirred in the rotary (60 ˚С). This process was repeated in 3 cycles. The particle size reduction was performed in 10 cycles (1 min on and 1 min off) using an ultrasound probe (UP200H, Hielsher, Germany) at the frequency of 20 kHz. Coating of the nanoliposomes was performed based on the method of Li, Paulson, and Gill (2015) with some modifications. After the initial trial and error, chitosan was dissolved in 1% acetic acid (v/v) at various concentrations (0.0, 0.1, 0.2, 0.4, 0.6, and 0.8% w/w) for 16 h. Then, the same 6

volume of each chitosan solution was added gently (drop-wise) to the nanoliposome solution during stirring (300 rpm). The coated samples were kept at 4 ˚C until further tests. 2.5. Determination of mean particle size, PDI, and zeta potential The mean particle size, PDI, and zeta potential of the nanoliposomes were measured after dilution of each sample with distilled water (100-fold) using the dynamic light scattering (DLS) system (NanoSizer 3000, Malvern Instruments, Malvern, UK) at 25 °C and 90° angle. 2.6. Evaluation of encapsulation efficiency (EE) Two mL from each of the newly prepared samples was transferred to an Amicon ultra centrifugal filter (molecular weight cutoff = 100 kDa, Millipore, UK) and centrifuged at 3,000 × g for 10 min. Then, the amount of the peptides passing through the filter (free peptides) was determined by the Bradford (1976) method. Eventually, EE was obtained by calculating the concentration of the protein loaded into the nanoliposomes to the total protein (in percentage) (Sarabandi et al., 2019). 2.7. Effect of storage conditions on physical stability and EE To determine the effect of each storage condition, 2 mL of each sample was transferred to the microtubes and stored in the refrigerator (4 ˚C) and ambient temperature (25 ˚C) for 30 days. Then, changes in the mean particle size and EE of the nanoliposomes (resulting from the peptide release) were measured on the basis of the previously described methods (Li et al., 2015). 2.8. Spray-drying of nanoliposomes To prepare the feed, 2 g of maltodextrin was dissolved in 20 mL of distilled water for 1 h at 300 × g. Then, 10 mL of the liposomal solution (un-coated and coated sample at optimum concentration) was gradually added to 10 mL of the maltodextrin solution during stirring at 200 × g. Prior to the spray-drying, the mixture was kept at room temperature for 20 min. The feed-to-powder conversion process was performed with an in pilot-scale spray dryer (Büchi B-191; Büchi LaboratoriumsTecnik, Switzerland). The process parameters included the inlet-air temperature (130 ± 1 ˚C), outlet air temperature (73 ± 2 ˚C), feed rate (5 mL/min), drying air speed (0.54 m3 h-1), nozzle diameter

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(0.5 mm), and air pressure (5.4 bar). The spray-dried powders were collected and packed in airtight bags and stored in refrigerator until experiments (Sarabandi et al., 2018). 2.9. Physicochemical and morphological properties of spray-dried nanoliposomes Physical and functional properties of powders such as moisture content, water activity, bulk density, flowability (angle of repose), solubility, hygroscopicity and particle size were calculated using the methods described by Sarabandi et al. (2017). 2.9.1. FTIR spectroscopy To evaluate the chemical structure, each sample consisting of the pure hydrolysates, empty and coated freeze-dried nanoliposomes, chitosan, maltodextrin, and spray-dried nanoliposomes with a ratio of 1 to 100 was mixed with potassium bromide (KBr) and pressed to take a disk shape. Finally, the FTIR spectroscopy of the samples was performed at a frequency of 4000 to 400 cm-1 using a FTIR spectrophotometer (Shimadzu 8400, Japan). 2.9.2. Morphological features The structural properties of the pure (freeze-dried) peptides, primary nanoliposomes, and the spraydried liposomes were evaluated. For the liquid samples, each nanoliposome sample was diluted (50fold) with distilled water. Then, 1 to 2 drops of each sample were poured onto the laboratory slide and covered with lamina. In the next step, the samples were dried at 37 °C four 1 h. The samples were then coated with a thin gold layer to examine the morphological characteristics. Subsequently, the morphological characteristics were evaluated under 25 kV accelerating voltage using the scanning electron microscopy (SEM) (HITACHI PS-230, Japan). 2.10. Evaluation of properties of reconstituted nanoliposomes 2.10.1. Physical properties Ten mg of each dried sample was dissolved in distilled water at 300 × g for 30 min. Then the mean particle size, PDI, and zeta potential of the reconstituted samples were calculated based on the previously methods described. 2.10.2. Retention of EE and antioxidant activity 8

The effect of coating process on the retention of EE and antioxidant activity (AA) of the nanoliposomes after spray-drying and reconstitution were evaluated. Then, the retention percentage of each of these indices after reconstruction of the spray-dried nanoliposomes was calculated on the basis of the previously described methods and using the following equation: Retention of EE or AA (%) = (EE or AA after reconstitution/EE or AA in feed solution)×100

(2)

2.11. Statistical analysis The data statistical analysis was carried out using the SPSS software (version 19.0, SPSS Inc., Chicago, IL) and one-way analysis of variance (ANOVA). All tests were performed in 3 replications. Moreover, the Duncan’s test was employed at 5% significance level to compare the mean values. 3. Results and discussion 3.1. Antioxidant properties of peptide fractions The amino acid composition is one of the most important indices for the biological activity of bioactive peptides (Akbarbaglu et al., 2019). This study evaluated the effect of content and composition of total amino acids (especially hydrophobic) on the antioxidant properties of peptide fractions (Fig. 1a). Our results showed that the amount of hydrophobic antioxidant amino acids (alanine, valine, isoleucine, leucine, tyrosine, phenylalanine, tryptophan, and methionine) and other antioxidants (histidine and lysine) in peptide fractions with low molecular weight (PF-10 kDa) are more than other fractions. Similar results were observed in the evaluation of the composition of free and total amino acids in peptide fractions derived from cod protein hydrolysate (Farvin et al., 2016). The results of the scavenging of DPPH and ABTS free radicals in peptide fractions (Fig. 1b) indicated that the molecular weight of fractions affected antioxidant activity. Among different samples, PF-10 had a higher DPPH radical scavenging (about 49%) than PF-100 (about 42%). This difference can be attributed to the greater amount of free and total hydrophobic amino acids and antioxidants in this sample (Akbarbaglu et al., 2019).

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These amino acids act as proton-donating compounds and induce DPPH radical scavenging and stable diamagnetic molecule formation (You, Zhao, Cui, Zhao, & Yang, 2009). Also, the higher amount of hydrophobic amino acids in PF-10 further activates this fraction in reaction with the DPPH lipophilic radical (Sarabandi et al., 2018). The activity of peptide fractions in the ABTS radical scavenging (as an index of lipophilic and hydrophilic antioxidant compound activity) was also evaluated. The highest percentage of this radical scavenging (about 82%) was observed in PF10. This finding indicates higher electron and hydrogen donation of this fraction compared with other samples. In a similar study, the antioxidant activity of peptides derived from canola meal protein hydrolysate was measured. Among different samples, the highest ABTS radical scavenging activity was observed in peptide fractions less than 1 kDa (Alashi et al., 2014). Considering these cases, PF-10 was selected for loading in nanoliposomes and further studies. 3.2. Physical properties of nanoliposomes The effect of different concentrations of chitosan coating on the mean particle size, polydispersity index (PDI) and zeta potential of nanoliposomes loaded with PF-10 were evaluated, as shown in Table 1A. It was revealed that chitosan-coated nanoliposomes had a higher particle size and PDI. However, the magnitude of these changes varied depending on the concentration of chitosan. For example, coating with 0.1% chitosan increased the mean size and PDI of the nanoparticles from 86 nm to 554 nm and from 0.326 to 0.643, respectively. These changes imply system instability due to the adherence of particles to each other, the bridging flocculation, and the production of heterogeneous liposomal systems with wide particle size distribution (da Rosa Zavareze et al., 2014; Li et al., 2015; Sarabandi, Mahoonak, et al., 2019). Among the other samples, nanoliposomes coated with 0.4% chitosan had the lowest particle size (132 nm) and PDI (0.371). These findings indicated a more homogeneous system than other samples. A similar study of coated nanoliposomes loaded with black mulberry extract observed severe instability and a significant increase in particle size at lower chitosan concentrations (less than 0.2%) (Gültekin-Özgüven et al., 2016). Moreover, Li et al. (2015) coated nanoliposomes containing salmon protein hydrolysate with chitosan (0– 10

0.6%). They reported that the lowest uniformity (PDI = 0.87) was observed in nanoparticles coated with 0.3% chitosan. This study also reported the instability of the system, particle aggregation, and the production of aggregates with the size above 1000 nm at low chitosan concentrations (less than 0.1%). The zeta potential of empty nanoliposomes (-16 mV) was affected by the phosphate groups. However, loading of peptide fraction with the zeta potential of -31 mV resulted in the surface charge shift of the nanoparticles from -16 mV to -18 mV (Table 1A). The negative charge of the peptide solution can be attributed to the high concentration of glutamic and aspartic amino acids in its composition (Corrêa et al., 2019). Loading of peptides in monolayer membranes and surface areas, as well as the reaction of phospholipids with peptide fractions, are the reasons for the zeta potential change of nanoliposomes (da Rosa Zavareze et al., 2014). Similar studies showed that the encapsulation of whey protein (Mohan et al., 2016) and casein (Sarabandi et al., 2019) hydrolysate changed the zeta potential of nanoliposomes. On the other hand, coating the nanoliposomes with chitosan resulted in the charge shift of the particles from negative to positive values, due to locating amino groups of chitosan with positive charge on the surface of the charged nanovesicles as a result of the ionic reaction between them (Ramezanzade et al., 2017). Also, no change in zeta potential was observed after increasing the chitosan concentration from 0.6 to 0.8% due to the saturation of vesicles surface with the coating compounds. A similar study reported the zeta potential of nanoliposomes loaded with trout protein hydrolysate (Ramezanzade et al., 2017) and salmon peptides (Li et al., 2015) after coating with chitosan. 3.3. Encapsulation efficiency (EE) EE is one of the effective indicators of the performance of nanocarriers in enhancing the stability and preservation of nutritional and pharmaceutical compounds. The value of this indicator was affected by the concentration of coating with chitosan (Table 1A). As the concentration of chitosan increased from 0.1 to 0.2%, EE increased from about 76% to 88% which can be attributed to coating of pores in the surface of the membrane and inhibiting the release of loaded compounds (Li 11

et al., 2015). However, coating the nanoparticles with concentrations above 0.4% chitosan resulted in lower EE. These changes can be related to the effect of different coating concentrations on the zeta potential and stability of nanoliposomes (Li et al., 2015). These findings were in agreement with the results obtained of Ramezanzade et al. (2017) and Li et al. (2015), respectively, who evaluated the effect of chitosan concentrations on the EE of nanoliposomes loaded with rainbow trout and salmon protein hydrolysates; respectively. 3.4. Effect of storage conditions on physical stability and EE One of the most important goals of designing and producing colloidal systems is to maintain their physicochemical stability and encapsulation efficiency during storage (Sarabandi et al., 2019). In this study, the physical stability of nanoliposomes coated with different concentrations of chitosan was investigated after 30 days storage at room temperature (25˚C) and refrigerator (4˚C), as depicted in Fig. 2a. The results showed that the maximum and minimum changes in particle size were observed in nanoliposomes maintained at ambient temperature and refrigerator; respectively. For example, the mean particle size for uncoated nanoliposomes increased from 86 nm to 347 nm and 148 nm after storage at 25˚C and 4˚C, respectively, due to maintaining zeta potential and physical stability, increasing membrane stability, as well as lower accumulation of nanoparticles resulting from their lower molecular mobility at the refrigerator temperature (Gibis, Zeeb, & Weiss, 2014; Tan, Feng, Zhang, Xia, & Xia, 2016). However, coating the nanoparticles especially with 0.4% chitosan resulted in the highest physical stability and the smallest change in the particle size (from about 131 nm to 289 nm and 150 nm) after storage under the above conditions. The higher physical stability of nanoliposomes after coating (especially at optimum concentration) can be attributed to the role of chitosan layer by creating electrostatic repulsion between the particles and preventing aggregation and integration processes (Fig. 2b) (Li et al., 2015). Also, lower physical stability (increased turbidity as a result of increasing particle size) of nanoliposomes without coating (Fig. 2c) compared to the coated samples can be observed after 4 months of storage (Fig. 2d) (data not shown). 12

The effect of storage temperature and different concentrations of chitosan on the release of loaded peptides and encapsulation efficiency was also investigated (Fig. 2e-f). The sustained release of loaded compounds can be attributed to the diffusion-controlled mechanism (Hosseini et al., 2017). Among the samples stored at 4 ˚C for 30 days, the highest (80%) and the lowest (53%) EE retention were related to coated and non-coated nanoliposomes; respectively. However, keeping the samples at ambient temperature reduced the EE value. After 30 days of storage, approximately 73% (in 0.6% chitosan-coated sample) and 44% (uncoated sample) of EE were maintained; respectively. The reduced release rate of the loaded peptides at low temperature can be attributed to the decrease in membrane flexibility and permeability, as well as the lower molecular mobility of phospholipids at low temperature (Zhao et al., 2011). The role of chitosan coatings on the retention of EE can also be attributed to the coating of the membrane surface pores, reducing the leakage of loaded compounds, and increasing the stability of nanoparticles (Li et al., 2015). In another study, the increased stability and the reduced release of loaded trout protein hydrolysates were reported after coating of nanoliposomes (Ramezanzade et al., 2017). Nanoliposomes coated with 0.4% chitosan were selected as the optimum treatment (optimum coating concentration). Finally, this sample along with uncoated nanoliposomes were used to evaluate the effect of coating process on the physicochemical properties and stability of nanoliposomes during spray drying and after reconstitution 3.5. Physical properties of spray-dried nanoliposomes Moisture content, water activity (less than 0.6), and angle of repose in the powders indicated good microbiological stability and flowability, respectively (Sarabandi et al., 2017), as shown in Table 1B. In addition, the solubility of powders was more than 95%, and no difference was observed among the samples. On the other hand, hygroscopicity was measured as an index of determining physical stability, preservation of functional properties and flowability of powders. This index was higher in chitosan-coated sample (about 16%) than non-coated sample (14%), which could be related to the effect of chitosan on increased moisture uptake during storage. However, the 13

hygroscopicity of spray-dried samples was much lower than that of the pure peptide fraction (about 42%). This finding indicates the increased stability of spray-dried liposomes during storage (Akbarbaglu et al., 2019). Also, the average particle size of powders was obtained between 8 and 9 µm. Regarding the threshold for particle size detection in foods (such as ice cream, dessert, sauce, and chocolate) which is between 10 and 50 µm (Imai, Saito, Hatakeyama, Hatae, & Shimada, 1999), the production of coarse particles in the formulation of fortified food products results in a sandy and undesirable feeling in the mouth (Sarabandi et al., 2018). The average particle size of the spray-dried liposomes enables direct use of these powders in the production of functional formulations. 3.6. FTIR results The effect of loading peptide fractions on the chemical structure of nanoliposomes and their possible interactions between their functional groups were investigated in this study. In addition, the chemical structure of the nanoliposomes entrapped in the maltodextrin matrix was also evaluated after the spray-drying process (Fig. 3). The major areas in the chemical structure of the peptides are mainly related to O-H stretch (3737 cm-1 and 2933 cm-1), N-H stretch (3431 cm-1), C=O stretch and amide-Ι region (1647 cm-1), C-N stretch and N-H deformation (1537 cm-1), C=O stretch (1071 cm-1) and N-H bending (670 cm-1). The most important spectra in empty nanoliposomes are O-H stretch (3438cm-1), stretch vibrations of CH2 (2927cm-1), stretch vibrations of C=O (1736 cm-1), symmetrical-PO2- stretch vibrations (1246 cm-1 and 1124 cm-1) and asymmetrical stretch vibrations of the choline group (N+CH3) belonging to the polar region of phosphatidylcholine (966 cm-1) (Sarabandi et al., 2019). The encapsulation of peptide fractions into nanoliposomes resulted in changes in these frequencies. The most important of these changes can be the increase of intensities of the peaks in the frequencies of 3438 cm-1 and 2927 cm-1 to 3434 cm-1 and 2929 cm-1. The reason for these changes can be attributed to the formation of hydrogen bonds between the O-H and N-H groups, the placement of peptides in the monolayer membrane, and the formation of the ionic complex with 14

phosphatidylcholine (da Rosa Zavareze et al., 2014). Moreover, increasing the intensity and shifting frequencies of 1736 cm-1 to 1740 cm-1 (hydrogen bonding between carbonyl phosphatidylcholine group and peptide), 1246 cm-1 to 1247 cm-1 and 1124 cm-1 to 1095 cm-1 (hydrogen bond formation between PO2 and peptides and their location in the polar region of vesicle) was observed. On the other hand, the peak intensity at the frequency of 966 cm-1 increased as a result of the peptides in the inner and polar regions of phosphatidylcholine (Sarabandi et al., 2019). However, some of these peaks were replaced with chitosan after coating the nanoliposomes. Some of these changes include shifting the frequency of 3434 cm-1 to 3439 cm-1 (hydrogen bonding between amino or hydroxyl groups of chitosan and carboxyl or amino groups of peptide), and reducing the intensity and shifting frequency of 1436 cm-1 to 1462 cm -1 (reduction of acyl chain mobility in the monolayer membrane resulting in decreased membrane fluidity) (Hasan et al., 2016; Ramezanzade et al., 2017). Spray drying of nanoliposomes using maltodextrin carrier also caused changes in the frequencies of carrier matrix. The peaks related to maltodextrin were mainly at 3388 cm-1 (O-H stretch), 2923 cm-1 (C-H stretch), 1652 cm-1 (H2O absorbed in amorphous region), 1455 cm-1 (CH2 bending), 1155 cm-1 (C-O stretch, C-O-H bending), 1017 cm-1 (C-O stretch), 850 cm-1 (CH2 and C1-H deformation), 764 cm-1 (C-C stretch) and 701 cm-1 (structural status of pyranose ring) (Akbarbaglu et al., 2019; Sarabandi et al., 2019). The results indicated a maltodextrin spectroscopy pattern. However, displacements at frequencies 3388 cm-1 to 3413 cm-1, 2923 cm-1 to 2927 cm-1, 1652 cm-1 to 1647 cm-1 and 1455 cm-1 to 1419 cm-1, 1017 cm-1 to 1025 cm-1were observed after spray drying of nanoliposomes in maltodextrin, due to the formation of some hydrogen bonds, hydrophobic and ionic interactions between the coated nanoliposomes and the carrier structure. The results of this section also indicated the incorporation of nanoliposomes in the maltodextrin matrix. In another study, a complex was formed between carboxymethyl starches and lecithin, and the resulting mixture was spray-dried. The results indicated the ionic reactions between CMS and the choline lecithin group (Friciu, Le, Ispas-Szabo, & Mateescu, 2013). Also, no new peaks were formed after

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the spray drying process. Similar findings were observed in the study carried out by Akbarbaglu et al. (2019) after spray drying of linseed hydrolysates. 3.7. Morphological features The morphological structure of the carriers reflects the effect of production process, type and composition of the formulation on the physical, functional, performance and stability properties of the produced particles (Sarabandi et al., 2017). The structural properties of pure freeze-dried peptide fraction (Fig. 4a), coated nanoliposomes (Fig. 4b), and spray-dried nanoliposomes (Fig. 4c and d) were investigated in this study. SEM images indicated the fractured sheet/laminar structures on freeze-dried peptides. These images indicate the impact of the freeze-drying process on the production of structures with high pores (Sarabandi et al., 2018). On the other hand, the aggregates of relatively spherical and adherent particles with smooth surfaces were observed in the produced nanoliposomes. Spherical particles have a higher controlled release ability and protection of encapsulated compounds due to less surface contact with the environment. The cause of particle adhesion can be attributed to the influence of the nanoparticles drying process during the preparation and taking SEM images resulted in their relative deformation (Rabelo, Oliveira, da Silva, Prata, & Hubinger, 2018). The SEM image showed the production of nano-sized particles which confirms the DLS results. The images of spray-dried nanoliposome also showed the production of irregularly shaped, wrinkled particles with different sizes. The high rate of moisture evaporation during the spray drying process results in rapid crust formation (Gültekin-Özgüven et al., 2016). Moreover, the structure and surface properties of spray-dried particles are affected by feed composition, carrier type, and process air temperature (Akbarbaglu et al., 2019). 3.8. Properties of reconstituted nanoliposomes 3.8.1. Physical properties It is important to maintain the physicochemical stability and encapsulation efficiency of nanoliposomes after spray drying and reconstitution process. The effect of chitosan coating on the particle size, PDI, zeta potential and EE of spray-dried nanoliposomes was investigated after 16

reconstitution (Table 1C). The mean particle size in uncoated nanoliposomes increased more than 5fold after reconstitution. In addition, the PDI for these nanoparticles changed from 0.326 to 0.701. These changes indicate damage to the liposomal membrane due to the stresses of drying, reduced hydration and adhesion of the nanoparticles after reconstitution as well as reduced homogeneity of the colloidal system (Wang et al., 2015). However, coating the nanoliposomes with chitosan significantly increased the stability and reduced changes in their size and PDI after reconstitution; which could reveal the increased liposomal membrane stability and reduced deformation of chitosan-coated nanoparticles during the drying and reconstitution steps. After the particle reconstitution, zeta potential of the uncoated nanoliposomes changed significantly, due to the release of loaded peptides from the vesicular structure. Also, EE of uncoated and coated nanoliposomes decreased by about 20% and 9%, respectively, after reconstitution. The reason for preserving physical and EE during storage and after nanoliposomal reconstitution could be due to the role of chitosan coating (as a protective compound) in reducing liposomal membrane mobility, creating electrostatic repulsion, maintaining monolayer integrity against unstable mechanisms such as aggregation, flocculation, coalescence, deposition, as well as reduced leakage of loaded compounds during storage and after reconstitution (Li et al., 2015; Tan et al., 2016). The effect of coating nanoliposomes with calcium alginate on their physical stability and EE retention after freeze-drying and spray drying was investigated in a similar study. The results showed that EE for coated and uncoated nanoliposomes decreased from about 81% to 67% and from 87% to 36%, respectively. The value of this index reached 58% (coated sample) and 47% (uncoated sample), respectively after spray drying (Tewa-Tagne et al., 2007). 3.8.2. Retention of antioxidant activity (AA) It is important to maintain the biological and antioxidant activity of bioactive compounds during the processes. The effect of chitosan coating on the retention of DPPH and ABTS free radical scavenging activity in nanoliposomes before and after spray drying has been shown in Fig. 5. There was no difference between the retention of these two indices in coated and uncoated nanoliposomes. 17

However, the spray drying process and thermal stress resulted in a decrease in the antioxidant activity of loaded nanoliposomes. About 10% and 6% of the DPPH and ABTS free radical scavenging activity were lost after spray drying. However, the decrease of these indices in the coated samples was about 4% and 1%, respectively; which indicates the effect of chitosan coating on the retention of primary EE and protection of peptide fractions against thermal stresses during spray drying. The retention of antioxidant activity in this study was higher than that reported for nano-composites containing cacao-hull extract. About 39% of total phenol, 30% of flavonoids and 47% of total extract antioxidant activity were retained after spray drying of nanoliposomes (Altin et al., 2018). Spray drying of nanoliposomes loaded with black mulberry extract was performed in a similar study, in which about 69% and 56% of total phenolic and anthocyanin compounds of the extract were retained in chitosan-coated particles (Gültekin-Özgüven et al., 2016). Increased antioxidant stability of quercetin (Hao et al., 2017) and carotenoids (Tan et al., 2016) were reported in nanoliposomes coated with chitosan in another study. 4. Conclusion In this study, loading of peptide fractions from flaxseed protein hydrolysate in nanoliposomes was performed. Then, the effect of different concentrations of chitosan-coating on the physicochemical, stability and release properties of the nanoliposomes was evaluated. The results showed that stability and maintenance of EE in nanoliposomes were affected by the storage temperature and the concentration of chitosan. Chemical structure evaluation (FTIR) indicated the interactions between peptides and liposomes and their placement into the internal parts of vesicles and monolayer membrane. Then, spray drying stabilization of loaded nanoliposomes by converting them into powder was performed. Investigation of the physicochemical and hygroscopicity properties of the spray-dried nanoliposomes indicated the production of powders with appropriate physicochemical stability, functionality, and flowability. The morphological characteristics of the particles were affected by the type of process. SEM images of nanoliposomes also confirmed the DLS results. The 18

effect of chitosan coating on the physicochemical and antioxidant properties of nanoliposomes during spray drying was also investigated. The results indicated increased lipid membrane stability to thermal stress and dehydration, thereby maintaining physical properties, zeta potential, EE and antioxidant activity of coated nanoliposomes after reconstitution. Acknowledgment This study was financially supported by the Iran National Science Foundation (INSF), Grant No. 98001878. References Akbarbaglu, Z., Jafari, S. M., Sarabandi, K., Mohammadi, M., Heshmati, M. K., & Pezeshki, A. (2019). Influence of spray drying encapsulation on the retention of antioxidant properties and microstructure of flaxseed protein hydrolysates. Colloids and Surfaces B: Biointerfaces. 178, 421-429. Akhavan, S., Assadpour, E., Katouzian, I., & Jafari, S. M. (2018). Lipid nano scale cargos for the protection and delivery of food bioactive ingredients and nutraceuticals. Trends in Food Science & Technology, 74, 132-146. Alashi, A. M., Blanchard, C. L., Mailer, R. J., Agboola, S. O., Mawson, A. J., He, R., … Aluko, R. E. (2014). Antioxidant properties of Australian canola meal protein hydrolysates. Food Chemistry, 146, 500–506. Alemán, A., Marín, D., Taladrid, D., Montero, P., & Gómez-Guillén, M. C. (2019). Encapsulation of antioxidant sea fennel (Crithmum maritimum) aqueous and ethanolic extracts in freezedried soy phosphatidylcholine liposomes. Food Research International, 119, 665–674. Altin, G., Gültekin-Özgüven, M., & Ozcelik, B. (2018). Chitosan coated liposome dispersions loaded with cacao hull waste extract: Effect of spray drying on physico-chemical stability and in vitro bioaccessibility. Journal of Food Engineering, 223, 91–98. Assadpour, E., & Jafari, S. M. (2019). Advances in spray-drying encapsulation of food bioactive ingredients: from microcapsules to nanocapsules. Annual Review of Food Science and 19

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flavonoid quercetin with chitosan-coated nano-liposomes. LWT-Food Science and Technology, 85, 37–44. Hasan, M., Messaoud, G. Ben, Michaux, F., Tamayol, A., Kahn, C. J. F., Belhaj, N., … ArabTehrany, E. (2016). Chitosan-coated liposomes encapsulating curcumin: study of lipid– polysaccharide interactions and nanovesicle behavior. RSC Advances, 6(51), 45290–45304. Hosseini, S. F., Ramezanzade, L., & Nikkhah, M. (2017). Nano-liposomal entrapment of bioactive peptidic fraction from fish gelatin hydrolysate. International Journal of Biological Macromolecules, 105, 1455–1463. Huang, M., Su, E., Zheng, F., & Tan, C. (2017). Encapsulation of flavonoids in liposomal delivery systems: the case of quercetin, kaempferol and luteolin. Food & Function, 8(9), 3198–3208. Imai, E., Saito, K., Hatakeyama, M., Hatae, K., & Shimada, A. (1999). Effect of physical properties of food particles on the degree of graininess perceived in the mouth. Journal of Texture Studies, 30(1), 59–88. Jafari, S. M., Assadpoor, E., He, Y., & Bhandari, B. (2008). Encapsulation efficiency of food flavours and oils during spray drying. Drying Technology, 26(7), 816–835. Katouzian, I., & Jafari, S. M. (2016). Nano-encapsulation as a promising approach for targeted delivery and controlled release of vitamins. Trends in Food Science and Technology. 53, 3448. Li, Z., Paulson, A. T., & Gill, T. A. (2015). Encapsulation of bioactive salmon protein hydrolysates with chitosan-coated liposomes. Journal of Functional Foods, 19, 733–743. McClements, D. J. (2018). Encapsulation, protection, and delivery of bioactive proteins and peptides using nanoparticle and microparticle systems: A review. Advances in Colloid and Interface Science, 253, 1–22. Mohan, A., McClements, D. J., & Udenigwe, C. C. (2016). Encapsulation of bioactive whey peptides in soy lecithin-derived nanoliposomes: influence of peptide molecular weight. Food Chemistry, 213, 143–148. 21

Rabelo, R. S., Oliveira, I. F., da Silva, V. M., Prata, A. S., & Hubinger, M. D. (2018). Chitosan coated nanostructured lipid carriers (NLCs) for loading Vitamin D: A physical stability study. International Journal of Biological Macromolecules, 119, 902–912. Ramezanzade, L., Hosseini, S. F., & Nikkhah, M. (2017). Biopolymer-coated nanoliposomes as carriers of rainbow trout skin-derived antioxidant peptides. Food Chemistry, 234, 220–229. Sarabandi, K., Peighambardoust, S. H., Mahoonak, A. S., & Samaei, S. P. (2017). Effect of carrier types and compositions on the production yield, microstructure and physical characteristics of spray dried sour cherry juice concentrate. Journal of Food Measurement and Characterization. https://doi.org/10.1007/s11694-017-9540-3 Sarabandi, K., Jafari, S. M., Mahoonak, A. S., & Mohammadi, A. (2019). Application of gum Arabic and maltodextrin for encapsulation of eggplant peel extract as a natural antioxidant and color source. International Journal of Biological Macromolecules, 140, 59–68. Sarabandi, K., Mahoonak, A. S., Hamishehkar, H., Ghorbani, M., & Jafari, S. M. (2019). Protection of casein hydrolysates within nanoliposomes: Antioxidant and stability characterization. Journal of Food Engineering, 251, 19-28. Sarabandi, K., Mahoonak, A. S., Hamishekar, H., Ghorbani, M., & Jafari, S. M. (2018). Microencapsulation of casein hydrolysates: physicochemical, antioxidant and microstructure properties. Journal of Food Engineering, 237, 86-95. Sarmadi, B. H., & Ismail, A. (2010). Antioxidative peptides from food proteins: a review. Peptides, 31(10), 1949–1956. Shishir, M. R. I., Xie, L., Sun, C., Zheng, X., & Chen, W. (2018). Advances in micro and nanoencapsulation of bioactive compounds using biopolymer and lipid-based transporters. Trends in Food Science & Technology, 78, 34–60. Stark, B., Pabst, G., & Prassl, R. (2010). Long-term stability of sterically stabilized liposomes by freezing and freeze-drying: Effects of cryoprotectants on structure. European Journal of Pharmaceutical Sciences, 41(3–4), 546–555. 22

Tan, C., Feng, B., Zhang, X., Xia, W., & Xia, S. (2016). Biopolymer-coated liposomes by electrostatic adsorption of chitosan (chitosomes) as novel delivery systems for carotenoids. Food Hydrocolloids, 52, 774–784. Tewa-Tagne, P., Briançon, S., & Fessi, H. (2007). Preparation of redispersible dry nanocapsules by means of spray-drying: development and characterisation. European Journal of Pharmaceutical Sciences, 30(2), 124–135. Udenigwe, C. C. (2014). Bioinformatics approaches, prospects and challenges of food bioactive peptide research. Trends in Food Science & Technology, 36(2), 137–143. Wang, L., Hu, X., Shen, B., Xie, Y., Shen, C., Lu, Y., … Wu, W. (2015). Enhanced stability of liposomes against solidification stress during freeze-drying and spray-drying by coating with calcium alginate. Journal of Drug Delivery Science and Technology, 30, 163–170. You, L., Zhao, M., Cui, C., Zhao, H., & Yang, B. (2009). Effect of degree of hydrolysis on the antioxidant activity of loach (Misgurnus anguillicaudatus) protein hydrolysates. Innovative Food Science & Emerging Technologies, 10(2), 235–240. Yousefi, M., Ehsani, A., & Jafari, S. M. (2019). Lipid-based nano delivery of antimicrobials to control food-borne bacteria. Advances in Colloid and Interface Science, 270, 263-277. Zhao, L., Xiong, H., Peng, H., Wang, Q., Han, D., Bai, C., … Deng, B. (2011). PEG-coated lyophilized proliposomes: preparation, characterizations and in vitro release evaluation of vitamin E. European Food Research and Technology, 232(4), 647–654. Fig 1. (a) HPLC chromatogram and amino acid composition; and (b) antioxidant activities of different peptide fractions Fig 2. The changes in (a) mean particles size; (b) schematic of stabilized nanovesicles via chitosan coating; changes in the turbidity of (c) uncoated and (d) coated nanoliposomes after 3 months storage in ambient temperature; the effect of storage temperature (e) 25˚С and (f) 4 ˚С on the encapsulation efficiency of nanoliposomes

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Fig 3. FTIR spectra of pure peptides, empty, loaded nanoliposomes, pure chitosan, coated nanoliposomes, pure maltodextrin and spray-dried nanoliposomes Fig 4. Morphological properties of (a) freeze-dried pure peptide; (b) nanoliposomes; (c and d) spray-dried nanoliposomes Fig 5. Antioxidant activity of uncoated (UL) and coated (CL) nanoliposomes before and after reconstitution (R) process

Table 1. (A) Physical properties and encapsulation efficiency of chitosan-coated nanoliposomes containing flaxseed peptide fractions Chitosan

Average size

Polydispersity

Zeta potential

Encapsulation

(% w/v)

(nm)

index (PDI)

(mV)

efficiency (%)

Control

81.49±2.52e

0.284±0.01g

-16.36±0.93e

-

0

86.19±1.17e

0.326±0.01f

-18.61±1.37f

84.01±1.91b

0.1

554.73±18.01a

0.643±0.01a

6.73±1.01d

76.63±1.47c

0.2

182.83±2.32b

0.485±0.01b

24.93±0.89c

88.51±1.31a

0.4

132.56±1.62d

0.371±0.01e

29.67±0.90b

90.73±1.60a

0.6

152.21±4.43c

0.423±0.01d

31.83±1.51a

75.53±2.15c

0.8

173.66±2.04b

0.452±0.01c

32.52±1.06a

67.96±2.40d

Data are presented as mean ± standard deviation (n=3) and different letters in the same column indicate significant differences at the 5% level in Duncan’s test.

(B) Physical properties of spray-dried chitosan-coated nanoliposomes containing flaxseed peptide fractions Chitosan

Moisture

Water

Bulk density

Angle of

Solubility

Hygroscopicity

Mean particle

(% w/v)

content (%)

activity

(g/mL)

repose (˚)

(%)

(%)

size (μm)

0

3.21±0.22a

0.301±0.01a

0.283±0.01a

35.5±1.7a

95.6±1.2a

14.5±1.4b

8.9±0.6a

0.4

3.26±0.08a

0.311±0.01a

0.297±0.01a

34.7±1.1a

97.2±1.3a

16.5±0.8a

8.4±0.2a

Data are presented as mean ± standard deviation (n=3) and different letters in the same column indicate significant differences at the 5% level in Duncan’s test.

(C) Changes in Mean particle size, PDI, Zeta potential and encapsulation efficiency values before and after reconstitution of spray-dried nanoliposomes Mean particle size (nm) Chitosan (% w/v)

Primary

Reconstituted

Polydispersity index Primary

Reconstituted

0

86.2±1.2b

428.3±6.9a

0.326±0.01b

0.701±0.01a

0.4

132.7±1.5b

233.4±5.7a

0.371±0.01b

0.392±0.01a

Zeta potential (mV) Chitosan (% w/v)

Primary

Reconstituted

Encapsulation efficiency (%) Primary

Reconstituted

0

-18.6±1.3a

-25.7±1.4b

84.0±1.9a

65.1±2.7b

0.4

29.7±0.9a

26.1±0.6b

90.7±1.6a

81.8±2.1b

24

Data are presented as mean ± standard deviation (n=3) and different letters in the same row indicate significant differences at the 5% level in Duncan’s test.

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