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007afdec-bed4-405d-873d-c355ba9add0e.150 | *3.6. Characterization of New Formulated Food Product*
To test selected functionality, the powders were added to a recipe of nachos in a ratio of 3%. Therefore, three variants of nachos were obtained according to the two variants, coded as N1 (3% addition of variant V1) and N2 (3% addition of variant V2) and a blank without powder (C). The obtained food products were analyzed in terms of bioactive stability for 28 days over storage at 25 ◦C. As expected, the differences in bioactives and antioxidant activity between samples correlated with the added quantity of powder (Table 2).
**Table 2.** Phytochemical profile of added-value nachos and stability during 28 days of storage.
Means on the same row that do not share letter (a, b) are significantly different, based on Tukey method and 95% confidence.
During the storage test, no significant decrease (*p* > 0.05) was found in the flavonoid contents of N1, in contrast with N2, where a significant decrease of 9% was observed (*p* < 0.05). From Table 2, a significant increase in antioxidant activity values for both variants, during storage, can be observed, probably due to the release of some other compounds, apart from flavonoids, such as phenolics from microcapsules. Therefore, an increase in antioxidant activity was found in both variants, at approximatively 26%. Milea et al. [12] reported a decrease of 43% for flavonoids, 35% for polyphenols and 8% for antioxidant activity, in the case of a new formulated soft cheese with the addition of 1% microencapsulated powder, and 47% for flavonoids and 31% and 9% for polyphenols and antioxidant activity in the case of a soft cheese with 2% microencapsulated powder.
| doab | 2025-04-07T03:56:59.207134 | 17-11-2022 17:23 | {
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007afdec-bed4-405d-873d-c355ba9add0e.151 | **4. Conclusions**
In this study, flavonoid-loaded microcapsules using whey protein isolates glycated with xylose via the Maillard reaction were successfully obtained. The liquid–solid, ultrasoundassisted extraction method was applied to obtain a flavonoid-enriched extract. The extract showed a significant content of flavonoids and a satisfactory antioxidant activity. Whey protein isolates and xylose were used, in nonglycated and glycated forms, as possible candidates for the microencapsulation of flavonoid-enriched onion skin extracts by freezedrying. Both powders were characterized, showing significant amounts of polyphenols, flavonoids and a remarkable antioxidant capacity. A positive correlation was found between the browning index and antioxidant activity, and consecutively between the grafting degree and microencapsulation efficiency. The confocal laser scanning microscopy confirmed the higher ability of the whey protein isolate–xylose conjugates to entrap flavonoids. The molecular docking studies allowed the identification of the potential zones from αlactalbumin and β-lactoglobulin surfaces involved in the interaction with xylose molecules. Xylose appeared to attach with high affinity to the α-lactalbumin protein pocket involved in flavonoid binding. In the case of β-lactoglobulin, the tested ligands docked to different sites; smaller cavities located on the protein surface are preferred by xylose. The powder was added to nachos, and a slightly decrease in phytochemicals was found during storage. However, the antioxidant activity of the added-value products increased, probably due to the release of some other bioactives from microcapsules. Based on the reported results, the protein–monosaccharide Maillard-type conjugates are a good alternative for food ingredient carriers and promising attractive methods of delivery.
**Author Contributions:** Conceptualization, N.S.; methodology, N.S.; software, S, .A.M.; validation, I.A. and N.S.; formal analysis, S, .A.M., E.E. and V.B.; resources, G.E.B. and G.R.; writing—original draft preparation, S, .A.M., I.A. and N.S.; writing—review and editing, I.A. and N.S.; supervision, N.S.; project administration, N.S.; funding acquisition, G.E.B. and G.R. All authors have read and agreed to the published version of the manuscript.
**Funding:** This work was supported by a grant of the Romanian Ministry of Research and Innovation, CCCDI-UEFISCDI, project number PN-III-P1-1.2-PCCDI-2017-0569-PRO-SPER (10PCCI), within the PNCDI programme.
**Institutional Review Board Statement:** Not applicable.
**Informed Consent Statement:** Not applicable.
**Data Availability Statement:** Data are contained within the article.
**Acknowledgments:** The Integrated Centre for Research, Expertise and Technological Transfer in Food Industry is acknowledged for providing technical support.
**Conflicts of Interest:** The authors declare no conflict of interest.
| doab | 2025-04-07T03:56:59.207238 | 17-11-2022 17:23 | {
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007afdec-bed4-405d-873d-c355ba9add0e.153 | *Article* **Resveratrol Stabilization and Loss by Sodium Caseinate, Whey and Soy Protein Isolates: Loading, Antioxidant Activity, Oxidability**
**Xin Yin 1,2, Hao Cheng 1,2, Wusigale 3,4, Huanhuan Dong 1,2, Weining Huang 1,2 and Li Liang 1,2,\***
**Abstract:** The interaction of protein carrier and polyphenol is variable due to their environmental sensitivity. In this study, the interaction between resveratrol and whey protein isolate (WPI), sodium caseinate (SC) and soy protein isolate (SPI) during storage were systematically investigated from the aspects of polyphenol loading, antioxidant activity and oxidability. It was revealed that resveratrol loaded more in the SPI core and existed both in the core of SC micelles and on the particle surface, while WPI and resveratrol mainly formed in complexes. The loading capacity of the three proteins ranked in order SC > SPI > WPI. ABTS assay showed that the antioxidant activity of the protein carriers in the initial state was SC > SPI > WPI. The results of sulfhydryl, carbonyl and amino acid analysis showed that protein oxidability was SPI > SC > WPI. WPI, with the least oxidation, improved the storage stability of resveratrol, and the impact of SC on resveratrol stability changed from a protective to a pro-degradation effect. Co-oxidation occurred between SPI and resveratrol during storage, which refers to covalent interactions. The data gathered here suggested that the transition between the antioxidant and pro-oxidative properties of the carrier is the primary factor to investigate its protective effect on the delivered polyphenol.
**Keywords:** protein; resveratrol; loading; antioxidant activity; oxidability; stability
| doab | 2025-04-07T03:56:59.207433 | 17-11-2022 17:23 | {
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"title": "Characterization and Encapsulation of Natural Antioxidants: Interaction, Protection and Delivery",
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007afdec-bed4-405d-873d-c355ba9add0e.154 | **1. Introduction**
Protein-based assemblies including molecular complexes, nano-/micro-particles, and their stabilized emulsions and emulsion gels have been expected to protect antioxidants [1,2]. Even though the stabilization mechanism of polyphenols in proteins is not fully clear, there is a hypothesis that proteins express a protective effect by shielding the environmental accessibility of polyphenols and scavenging the free radical [3]. However, amphiphilic and hydrophilic polyphenols cannot be completely encapsulated in a carrier, and a portion of polyphenols are still in the free form. Meanwhile, it is worth noting that antioxidants can be converted into pro-oxidants under certain conditions and proteins may generate reactive oxidative species [4]. The imbalance between pro-oxidation and anti-oxidation in the physiological system eventually leads to the oxidation of biomolecules, the so-called oxidative stress [5]. The interaction between proteins and polyphenols might also be complicated and changeable, since they are both environment sensitive.
It has been reported that bioactive components including vitamins, polyunsaturated fatty acids and polyphenols may also affect the delivery carriers. The photo-decomposition of folic acid caused the indirect oxidation of the whey protein isolate (WPI), which enhanced the protein antioxidant activity, leading to increased protection for the folic acid [6].
**Citation:** Yin, X.; Cheng, H.; Wusigale; Dong, H.; Huang, W.; Liang, L. Resveratrol Stabilization and Loss by Sodium Caseinate, Whey and Soy Protein Isolates: Loading, Antioxidant Activity, Oxidability. *Antioxidants* **2022**, *11*, 647. https://doi.org/10.3390/ antiox11040647
Academic Editor: Monica Rosa Loizzo
Received: 1 March 2022 Accepted: 25 March 2022 Published: 28 March 2022
**Publisher's Note:** MDPI stays neutral with regard to jurisdictional claims in published maps and institutional affiliations.
**Copyright:** © 2022 by the authors. Licensee MDPI, Basel, Switzerland. This article is an open access article distributed under the terms and conditions of the Creative Commons Attribution (CC BY) license (https:// creativecommons.org/licenses/by/ 4.0/).
β-Carotene, one of the major carotenoids, produced free radicals that can accelerate the oxidation of WPI in oil-in-water emulsions [7], while phenolic anthocyanins provided protection against the oxidation of Trp [8]. The oxidation of fish protein rich in polyunsaturated fatty acids was promoted by lipid oxidation products, especially secondary oxidation products, and the oxidation of protein and lipids occurred in parallel, showing a good correlation [9]. Although protein–polyphenol interaction has been investigated for the modification of protein structure and colloidal stability [10,11], its impact on the stability of polyphenols has rarely been reported until now. It is thus necessary to clarify the mechanism of proteins on the stability of polyphenols in more depth.
Resveratrol (*trans*-3,5,4 -trihydroxy-stilbene) is known as a polyphenolic compound with antioxidant activity. However, resveratrol is prone to oxidation, which limits its application in commercial products. Various proteins (e.g., zein, gliadin and ovalbumin) have been reported to stabilize resveratrol [12,13], but bovine serum albumin (BSA) accelerates the degradation of resveratrol [14]. Proteins in the molecular level and in the form of micelles might provide a different microenvironment and unique carrying properties for targeted antioxidants [15]. β-casein in the molecular level improved the storage stability of both *cis-* and *trans*-resveratrol better than β-casein micelles, although β-casein micelles could inhibit the transformation of resveratrol from *trans*-isomer to *cis*-isomer to a certain extent [16]. The stabilization effect on resveratrol is dependent on the type and concentration of protein carriers [17], but lacking systematic comparison study. Therefore, there is a growing demand for clarifying the theoretical basis to select suitable proteins as carrier materials for resveratrol.
Sodium caseinate (SC) and WPI are major milk proteins, and SC has a disordered structure and is more hydrophobic properties than WPI, while WPI contains two major globular proteins β-lactoglobulin and α-lactalbumin [18]. Soy protein isolate (SPI) is mainly composed of 7S and 11S globulins. WPI, SC and SPI are generally recognized as safe (GRAS), and their assemblies are commonly used to protect antioxidants against oxidation and degradation [19,20]. In the present study, WPI, SC, and SPI at various concentrations were used to investigate their effect on the storage stability of resveratrol and the polyphenol impact on the composition of proteins. The data gathered here should help guide the shelf life of the protein–polyphenol system used in commercial products.
| doab | 2025-04-07T03:56:59.207572 | 17-11-2022 17:23 | {
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007afdec-bed4-405d-873d-c355ba9add0e.155 | **2. Materials and Methods**
#### *2.1. Materials*
WPI (≥92%) was obtained from Davisco International Inc (Le Sueur, MN, USA). SPI (≥90%) was from Shandong Xiya Chemical Industry Co., Ltd. (Linshu, Shandong, China). SC, resveratrol (trans-isomer, >99%) and polydatin (HPLC grade, >95%) were purchased from Sigma-Aldrich Co. (St. Louis, MO, USA). 2,2 -azino-bis-3- ethylbenzthiazoline-6-sulphonic acid (ABTS) was purchased from Aladdin Bio-Chem Technology Co., Ltd. (Shanghai, China). Other agents were of analytical grade and purchased from SinoPharm CNCM Ltd. (Shanghai, China).
#### *2.2. Sample Preparation*
WPI, SPI or SC powder was dissolved in ultrapure water. The solutions were adjusted to pH 12 with 2 M NaOH and hydrated fully with magnetic stirring for 1 h, and then neutralized the pH to 7 with 2 M HCl under agitation for another 1 h. Stock solutions of proteins were 0.02%, 0.2% and 2.0% (*w*/*v*). Stock solution of resveratrol was prepared at a concentration of 2 mM by dissolving in 70% (*v*/*v*) ethanol. The resveratrol solution was added into protein solutions and diluted with water at pH 7 under stirring for 30 min. The final concentrations were 0.01%, 0.10% and 1.00% for proteins and 25, 50 and 100 μM for resveratrol. The 0.02% (*w*/*v*) sodium azide was added to solutions as an antimicrobial agent.
#### *2.3. Fluorescence Spectroscopy*
Fluorescence of pyrene as a probe was measured on a Cary Eclipse fluorescence spectrophotometer (Agilent Co., Ltd., New York, NY, USA) equipped with 10 mm quartz cuvettes. The spectral resolution was 2.5 nm for both excitation and emission. Pyrene in acetone was added into samples with its final concentration of 1 μM under stirring for at least 12 h before measurement. Fluorescence emission spectra were scanned from 350 to 600 nm with the excitation wavelength of 335 nm and the ratio of the intensity of the first and third bands (I1/I3) was calculated [21].
Fluorescence emission spectra of resveratrol in the absence or presence of proteins were recorded from 330 to 600 nm at an excitation wavelength of 320 nm. Slit widths with a nominal band-pass of 5 nm were used for both excitation and emission. Background of proteins was subtracted from raw spectra.
### *2.4. Particle Size and ζ-Potential*
Size distribution by the intensity and ζ-potential were determined by a NanoBrooker Omni Particle size Analyzer (Brookhaven Instruments Ltd., New York, NY, USA) with a He/Ne laser (λ = 633 nm) at a scattering angle of 173◦. They were obtained using an NNLS model and Smoluchowski model through phase analysis light-scattering (PALS) measurement, respectively.
| doab | 2025-04-07T03:56:59.207905 | 17-11-2022 17:23 | {
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007afdec-bed4-405d-873d-c355ba9add0e.156 | *2.5. Color Evaluation*
The color parameters of protein-resveratrol solutions before and after storage at 45 ◦C for 30 days were measured using a ColorQuest XE colorimeter (ColorQuest XE, Hunter Lab, Reston, VA, USA) and calculated using the Hunter Lab color scale (L\*a\*b\*). L\* represents the lightness (black = 0 to white = 100), a\* varies from red (positive) to green (negative), and b\* varies from yellow (positive) to blue (negative). The total color difference (ΔE) was calculated from the tristimulus color coordinates using the following equation:
$$
\Delta \mathbf{E} = \left[ (\mathbf{L}^\* - \mathbf{L}\_i^\*)^2 + (\mathbf{a}^\* - \mathbf{a}\_i^\*)^2 + (\mathbf{b}^\* - \mathbf{b}\_i^\*)^2 \right]^{1/2} \tag{1}
$$
where, Li\*, ai\*, bi\* are the initial values of the CIE L\*a\*b\* color coordinates of freshlyprepared samples, and L\*, a\*, b\* are the color coordinates of samples after 30 days. Additionally, the difference in chroma (ΔC\*) value, which represents the color intensity of samples, was analyzed by the following equation [22]:
$$
\Delta \mathbf{C}^\* = \left[ \left( \mathbf{a}^\* - \mathbf{a}\_i^\* \right)^2 + \left( \mathbf{b}^\* - \mathbf{b}\_i^\* \right)^2 \right]^{1/2} \tag{2}
$$
| doab | 2025-04-07T03:56:59.208205 | 17-11-2022 17:23 | {
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007afdec-bed4-405d-873d-c355ba9add0e.157 | *2.6. Resveratrol Quantification*
An exactly 0.5 mL sample was mixed with 0.5 mL polydatin (internal standard, 50 μM) in methanol and then added into 4 mL methanol under vortexing for 60 s. After the mixture was centrifuged at 15,000× *g* for 60 min, the supernatant was measured on the Alliance HPLC system equipped with a 2695 separation module and 2998 PDA detector (Waters, Milford, MA, USA). The mobile phase was a mixture of methanol and distilled water (50:50, *v*/*v*), the flow rate was 1 mL min−1, and the column temperature was 35 ◦C. Both *trans*-resveratrol and polydatin were analyzed at 306 nm [23].
| doab | 2025-04-07T03:56:59.208296 | 17-11-2022 17:23 | {
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007afdec-bed4-405d-873d-c355ba9add0e.158 | *2.7. Loading Efficiency*
Loading efficiency of resveratrol was determined by isoelectric precipitation method [24]. The samples with WPI, SPI and SC were adjusted to pH 4.8–4.6 using 0.1 M NaOH or HCl. Loading efficiency of resveratrol was calculated according to following formulation:
$$\text{Loading efficiency} \left(\% \right) = \left(1 - \frac{\text{C}\_{\text{s}}}{\text{C}\_{0}} \right) \times 100 \tag{3}$$
where, C0 and Cs were resveratrol in samples and in the supernatant, centrifuged at 5000× *g* for 20 min, respectively.
#### *2.8. Antioxidant Activity*
ABTS assay was analyzed according to previous methods [25]. In brief, 7.4 mM ABTS and 2.6 mM K2S2O8 were mixed in the dark for 12 h to produce ABTS· <sup>+</sup> solution, which was diluted and mixed with samples or buffer at a volume ratio of 19:1 and kept in the dark for 6 min. The absorbance was measured at 729 nm using a UV-1800 UV–Vis spectrophotometer (Shimadzu Co., Tokyo, Japan). The radical-scavenging activity was calculated as follows:
$$\text{Scavenging capacity} \left( \% \right) = \frac{\text{A}\_{\text{\textdegree}} - \text{A}\_{\text{\textdegree}}}{\text{A}\_{\text{\textdegree}}} \times 100 \tag{4}$$
where Ac and As are the absorbance of radical plus buffer and sample, respectively.
#### *2.9. Sulfhydryl Analysis*
Samples were mixed at a volume ratio of 1:2 with 0.1 M phosphate buffer at pH 8.0 without and with 8 M urea for free and total sulfhydryl determination, respectively. Then absorbance at 412 nm was measured, after 10 mM DTNB was added under vigorous stirring and incubated in the dark for 1 h. Both reagent and sample blanks were subtracted. Content of free and total sulfhydryl was calculated by using a molar extinction coefficient of 13,600 M<sup>−</sup>1cm−<sup>1</sup> and expressed as nmol per mg protein [26].
| doab | 2025-04-07T03:56:59.208369 | 17-11-2022 17:23 | {
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007afdec-bed4-405d-873d-c355ba9add0e.159 | *2.10. Carbonyl Analysis*
Protein solutions in the presence and absence of resveratrol during storage at 45 ◦C were mixed at a volume ratio of 1:2 with 10 mM DNPH in 2 M HCl. After 10% (*w*/*v*) trichloroacetic acid was added and centrifuged at 10,000× *g* for 5 min, precipitate was washed with 50% ethyl acetate and then dissolved in 6 M guanidine HCl in 20 mM phosphate buffer at pH 2.3. Absorbance at 370 nm was measured and carbonyl content was calculated using an extinction coefficient of 22,000 M−1cm−<sup>1</sup> and expressed as nmol per mg protein [27].
## *2.11. Amino Acid Analysis*
Amino acids except tryptophan were analyzed through acid hydrolysis of proteins by mixing 4 mL of samples with the same volume of 12 M HCl under blown nitrogen for 3 min, followed by hydrolysis at 120 ◦C for 22 h. Then a certain amount of NaOH was added to neutralize, and water was added to give a total volume of 25 mL. Tryptophan was determined by alkaline hydrolysis of proteins with 10 M NaOH and neutralized with a certain amount of HCl. The supernatant was centrifuged after filtering with filter paper. Amino acids were analyzed on the Agilent 1100 HPLC system equipped with an Agilent Hypersil ODS column (Angelon Co., Ltd., New York, NY, USA). Proline was detected at 262 nm, and the other amino acids were detected at 338 nm [6].
| doab | 2025-04-07T03:56:59.208483 | 17-11-2022 17:23 | {
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007afdec-bed4-405d-873d-c355ba9add0e.160 | *2.12. Statistical Analysis*
All experiments were repeated three times. Data are presented as mean ± standard deviation. An analysis of variance (ANOVA) of the data was carried out and identified using the Duncan procedure. All statistical analyses were performed using the software package SPSS 20.0 (SPSS Inc., Chicago, IL, USA). A *p* value < 0.05 was considered significant.
#### **3. Results**
| doab | 2025-04-07T03:56:59.208573 | 17-11-2022 17:23 | {
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007afdec-bed4-405d-873d-c355ba9add0e.161 | *3.1. Particle Characterization*
Pyrene is often used to investigate the association of macromolecules and the critical micelle concentration (CMC). Its intensity ratio I1/I3 decreased as the hydrophobicity of surrounding microenvironment increased [28]. The I1/I3 ratio of pyrene in water was 1.75 (±0.01). When the concentration of SC was 0.01%, the I1/I3 ratio was 1.67 (Figure 1), and its size distribution had three peaks around 1.5, 25 and 215 nm by intensity (Figure 2A). According to the submicelle model, each casein forms small submicelle units through hydrophobic interactions, and these subunits use calcium phosphate as the cement and further aggregates together to form SC micelles [29]. The relatively low concentration of SC solution is not sufficient to drive the formation of micelles, 0.01% SC mainly dissolved in the molecular level [30]. As the protein concentrations increased above 0.5%, the I1/I3 of SC gradually decreased to about 1.10 (Figure 1). The size distribution of SC at 0.1% showed a major peak around 230 nm and a minor peak around 25 nm, while only a peak at around 380 nm was observed at 1% (Figure 2A). These results indicate that SC aggregates to form micelles at 1% concentration [31]. As for WPI and SPI, the I1/I3 ratios of 0.01% protein were respectively 1.35 and 1.40 (Figure 1). Meanwhile, WPI had two peaks around 220 and 520 nm (Figure 2B), and SPI had two peaks around 110 and 380 nm (Figure 2C). The relatively low I1/I3 and large particle size suggest that WPI and SPI had already aggregated at 0.01%. The size peaks of WPI were not dependent on its concentrations (Figure 2B), while SPI became bigger with increasing concentrations, with two major peaks around 180 and 660 nm at 1% (Figure 2C). This is consistent with the results of the I1/I3 of pyrene. From Figure 1, the I1/I3 ratios of WPI decreased slightly from 1.35 at 0.01% to 1.18 at 0.1%, and then remained unchanged as the protein concentration further increased. The I1/I3 ratios decreased as the concentrations of proteins increased, reaching around 0.80 for 2% SPI (Figure 1). By comparative analysis, these results indicate that SPI particles have the most hydrophobic core, which is consistent with the highest content of hydrophobic amino acids (Tables 1 and 2 vs. Table 3). It has been reported that SPI had lower solubility compared with WPI and SC, while hydrophilic groups and/or water molecules were entrapped in the core of WPI particles [32,33].
**Figure 1.** I1/I3 of whey protein isolate (WPI), sodium caseinate (SC) and soy protein isolate (SPI) solutions at various concentrations.
Since WPI, SC or SPI have an isoelectric point (pI) around pH 4.5~5, the ζ−potential values of their particles are negative at pH 7.0 (Figure 3). ζ-Potential absolute values of the protein particles ranked in order WPI > SC > SPI at the same concentration (Figure 3). This is consistent with their molar ratio of acidic (Asp and Glu) and basic (His, Lys and Arg) amino acids being around 2.38 for WPI, 2.03 for SC, and 1.98 for SPI, calculated from the data in Tables 1–3. Together with the most hydrophobic core of SPI particles in Figure 1, these results indicate that more negatively-charged groups were masked in SPI particles than WPI and SC particles. ζ-Potential absolute values of all complex particles decreased as the protein concentration increased (Figure 3), suggesting that negatively-charged groups were entrapped in the aggregated particle core.
**Figure 2.** Size distribution of SC (**A**), WPI (**B**) and SPI (**C**) particles in the absence and presence of 25, 50 and 100 μM resveratrol. The protein concentrations were 0.01%, 0.1% and 1%.
**Table 1.** Amino acid composition of whey protein isolate (WPI) at 1% in the absence and presence of 100 μM resveratrol (RES) before and after storage for 30 days.
Note: Different lower-case letters in the same row represent significantly different mean values (*p* < 0.05).
**Table 2.** Amino acid composition of sodium caseinate (SC) in the absence and presence of resveratrol (RES) before and after storage for 30 days.
Note: Different lower-case letters in the same row represent significantly different mean values (*p* < 0.05).
**Table 3.** Amino acid composition of soy protein isolate (SPI) in the absence and presence of resveratrol (RES) before and after storage for 30 days.
Note: Different lower-case letters in the same row represent significantly different mean values (*p* < 0.05).
**Figure 3.** ζ−Potential of WPI (black), SC (red) and SPI (blue) particles in the absence and presence of 25, 50 and 100 μM resveratrol. The protein concentrations were 0.01%, 0.1% and 1%.
ζ−Potential absolute values of the protein particles decreased as the concentration of resveratrol increased, which was most pronounced at the protein concentration of 0.01% (Figure 3). The particles of SC, WPI and SPI became more homogeneous upon loading of resveratrol (Figure 2). These results are consistent with the formation of uniform particles of WPI with naringenin, a polyhydroxy flavonoid [34]. Meanwhile, the size distribution of all protein-resveratrol particles increased as the polyphenol concentration increased (Figure 2), which is consistent with the effect of hesperetin or hesperidin concentration on their individual particles with β-conglycinin, one of the major fractions of soy proteins, possibly due to polyphenols acting as bridging agents for protein molecules [35]. SC-resveratrol, WPI-resveratrol and SPI-resveratrol particles had a size distribution around 200–300 nm, 150–250 nm and 100–200 nm, respectively. At 25 μM resveratrol, the size distribution of
SC-resveratrol particles was close to the largest size distribution of SC particles (Figure 2A), while WPI-resveratrol and SPI-resveratrol particles had a size distribution close to the smallest ones of pure protein (Figure 2B,C). These results suggest that the addition of resveratrol favors the aggregation of SC but inhibits the formation of large WPI and SPI aggregates.
#### *3.2. Resveratrol Loading*
| doab | 2025-04-07T03:56:59.208623 | 17-11-2022 17:23 | {
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007afdec-bed4-405d-873d-c355ba9add0e.162 | 3.2.1. Microenvironment of Resveratrol
Pure resveratrol showed a crystalline structure, with sharp peaks at 6.58, 13.26, 16.39, 19.27, 22.40, 23.66, 25.28, 28.36 on a 2θ scale (Figure S2). Its characteristic peaks with less intensity were still observed in its physical mixtures with the proteins but disappeared in its protein particles (Figure S2), indicating that the polyphenol was amorphous when loaded in the protein particles [36]. From Figure 4, resveratrol in the absence of protein emits a relatively weak fluorescence, owing to its proton transfer tautomer fluorescence band [37]. The λmax of resveratrol around 400 nm shifted to 392, 388 and 383 nm in the presence of 0.1% WPI, SC and SPI, respectively. At the same time, the fluorescence intensity at λmax was 1.75, 3.06 and 4.09 times that of resveratrol alone. Similar changes were previously observed in the presence of β-lactoglobulin (β-LG) and bovine serum albumin (BSA), with respective fluorescence intensity at λmax of 393 and 379 nm being 1.21 and 4.92 times that of resveratrol alone [38]. These results indicate that the microenvironment of resveratrol was more hydrophobic in protein particles, and the order of hydrophobicity was SPI > SC > WPI. The hydrophobicity of the resveratrol microenvironment (Figure 4) is consistent with the aggregation degree of pure protein at 1% but not that at 0.1% (SPI > WPI > SC, Figure 1). This is possibly attributed to the different impact of resveratrol loading on the aggregation of the three proteins. As discussed above, the added resveratrol as bridging agent favors the aggregation of SC (Figure 2).
**Figure 4.** Fluorescence emission spectra of resveratrol in the absence (control) and presence of WPI, SC and SPI. Concentrations of resveratrol and proteins were 25 μM and 0.1%, respectively.
| doab | 2025-04-07T03:56:59.208938 | 17-11-2022 17:23 | {
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007afdec-bed4-405d-873d-c355ba9add0e.163 | 3.2.2. Loading Efficiency of Resveratrol
When the concentration of proteins was 0.01%, loading efficiencies of resveratrol were between 2% and 8% (Figure 5). The polyphenol loading efficiencies at 25 and 50 μM were greater in WPI and SPI particles than in SC particles. This may be due to WPI and SPI existing in the aggregate form at 0.01%, while SC exists in the molecular state (Figure 1). The loading efficiencies of resveratrol at 100 μM were 4% in all the protein particles (Figure 5), therefore resveratrol mainly exists in the free state in the presence of 0.01% proteins. As the concentration of WPI increased from 0.1% to 1%, the loading efficiencies of resveratrol increased by around 10%, of which the highest was 28%. The highest loading efficiencies of resveratrol in the presence of 0.1% SC and SPI were 31% and 27%, which further increased
at 1% SC and SPI to around 80% and 76%, respectively. In the case of 0.1% and 1% proteins, the loading efficiencies of resveratrol ranked in order SC > SPI > WPI. As mentioned above, resveratrol is conducive to the micellization of SC (Figures 2 and 4). The complex of resveratrol and protein masked the charged group, and the absolute value of the ζ-potential of the system decreased in the order of WPI > SC > SPI (Figure 3). It is speculated that the loading of resveratrol in SC particles not only depends on the transfer of the hydrophobic environment, but also refers to the bridging of resveratrol to submicelles. These results supported the hypothesis that the resveratrol was mainly located in the hydrophobic core of SPI, while both entrapped in the hydrophobic core and partially bound to the surface of the SC micelles. For WPI, more resveratrol complexed with the protein. Meanwhile, the loading efficiencies of the remaining resveratrol in protein particles were similar before and after storage at 45 ◦C for 30 days (Figure 5 and Figure S3).
**Figure 5.** Loading efficiency of resveratrol in its complex particles with WPI (black), SC (red) and SPI (blue) at 0.01%, 0.1% and 1%.
#### 3.2.3. Antioxidant Activity
It has been reported that casein contains more powerful antioxidant peptides than whey protein [39]. The presence of small peptides and C-terminal aromatic tyrosine residues contribute to the radical scavenging ability of SPI [40]. From Figure 6, the ABTS· + scavenging capacity of proteins ranked in order SC > SPI > WPI under the same concentration. Resveratrol contains three phenolic hydroxyl groups and possesses antioxidant activity [41]. When the concentrations of resveratrol were 25, 50 and 100 μM, its ABTS· + scavenging capacities were 11%, 20% and 44% (Figure 6), respectively. The scavenging capacities of WPI-resveratrol particles were similar to the sum of the individual capacity at the polyphenol concentrations of 25 and 50 μM (Figure 6A,B), suggesting an additive effect. At 100 μM, the scavenging capacities of WPI-resveratrol particles were less than the sum of the individual capacities (Figure 6C), suggesting partial screening of total antioxidant activity. As for SPI, resveratrol at 25 and 50 μM showed an additive effect with 0.01% and 0.1% protein but a masking effect with 1% protein (Figure 6A,B), and the masking effect was also observed at 100 μM resveratrol with all the investigated concentrations (Figure 6C). A masking effect was also observed in the case of SC, except for 25 and 50 μM resveratrol and 0.01% protein, which showed an additive effect (Figure 6). The masking effect is due to the protein–polyphenol interaction and the encapsulation of polyphenol in particles masking the phenolic hydroxyl groups [42,43]. It is worth noting that at 1% SPI and SC systems, the masked antioxidant activity was almost equal to that of resveratrol alone. This further confirms that resveratrol is mainly embedded in the hydrophobic core of SPI aggregations and SC micelles.
| doab | 2025-04-07T03:56:59.209068 | 17-11-2022 17:23 | {
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007afdec-bed4-405d-873d-c355ba9add0e.164 | *3.3. Protein Oxidation*
#### 3.3.1. Sulfhydryl Groups
It is well known that the ability of susceptible proteins for scavenging and generating reactive oxygen radical changes with the environment, which is related to the oxidation of the protein [44]. Protein oxidation is commonly accompanied by a decrease in the number of sulfhydryl (SH) groups [45]. The surface and total sulfhydryl contents of WPI were 12 and 17 nmol/mg, respectively, and for SPI, they were 3 nmol/mg and 5 nmol/mg, respectively. The lower sulfhydryl content of SPI in the initial state compared to WPI reflects that the initial oxidation state of SPI is greater than WPI, which may be related to the protein extraction process. As reported in the process of preparing SPI from defatted soy flour, lipoxygenase (LOX) was inevitably present in the system. The weakly alkaline extraction conditions caused LOX in the soybean flour to catalyze the oxidation of residual lipids [27]. The free and total sulfhydryl content of WPI and SPI decreased after storage (Figure 7), indicating that the accessible cysteine residues located at both surface and buried in the protein were attacked by free radicals [46]. The decrease in free and total sulfhydryl contents of SPI was greater in the presence than in the absence of resveratrol, while their contents of WPI were not affected by resveratrol (Figure 7). The interference of environmental factors on WPI and SPI sulfhydryl groups has been studied. After ultrasonic treatment, the disulfide bonds of SPI were destroyed, which significantly increased the free sulfhydryl content [47]. However, sonication did not change the thiol content of the whey protein concentrate. As reported, the oxidative susceptibility of free SH groups may depend on the constituent of mixture proteins. The intramolecular positions of the free thiol groups in β-lactoglobulin and α-lactalbumin may make WPI less sensitive [48].
| doab | 2025-04-07T03:56:59.209558 | 17-11-2022 17:23 | {
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007afdec-bed4-405d-873d-c355ba9add0e.165 | 3.3.2. Carbonyl Groups
Carbonyl groups (aldehydes and ketones) are produced on the side chains of the protein when they are oxidized [49]. The carbonyl contents of WPI, SC and SPI were 1.32, 1.63 and 2.26 nmol/mg (Figure 8), respectively. Due to the preparation process of SC, it contains about 6% of small ions in addition to the pure casein, mainly calcium, phosphate, magnesium and citric acid [15], leading to the worst oxidation stability. The extraction of SPI from soy flour may accelerate its carbonylation [50], since soy protein is extremely vulnerable to the attack of peroxyl radicals, and its degree of oxidation is related to the residual lipid content and LOX activity during the preparation process [27,51]. The carbonyl content of WPI increased during storage, reaching 2.7 nmol/mg after 30 days, which was invariable with the addition of resveratrol (Figure 8). The carbonyl content of SPI and SC increased as the concentration of resveratrol increased after 10 days. When the resveratrol concentrations were 0, 25, 50 and 100 μM, the carbonyl content of SPI increased from 3.95 to 5.11 nmol/mg, while the carbonyl content of SC increased from 2.77 to 3.29 nmol/mg after 30 days. The increase in carbonyl content may be related to the formation of peroxides in the system, which is generated by oxygen molecules attacking free radicals. The formation of peroxides on the α-carbon or other carbons of protein amino acid residues will result in an increase in the carbonyl content [27]. From Figure 8, it indicated that the peroxide content in the three protein solutions was in the order of SPI > SC > WPI, and the addition of resveratrol to SC and SPI solutions produced more peroxides. Together with the sulfhydryl contents in Figure 7, these results indicated that the SPI was more labile to oxidation than SC in the presence of resveratrol.
**Figure 6.** ABTS· <sup>+</sup> scavenging capacity of resveratrol, WPI (black), SC (red), SPI (blue) and WPIresveratrol, SC-resveratrol and SPI-resveratrol complex nanoparticles. The concentrations of proteins were 0.01%, 0.1% and 1%, while the concentrations of resveratrol were 25 (**A**), 50 (**B**) and 100 (**C**) μM.
**Figure 7.** Surface (**A**) and total (**B**) sulfhydryl content of WPI-resveratrol and SPI-resveratrol complex particles before (no pattern) and after (sparse pattern) storage at 45 ◦C for 30 days. The concentration of proteins was 1%.
#### 3.3.3. Amino Acid Composition
The oxidative attack of proteins modifies the side-chain groups of amino acid residues [52]. Tables 1–3 show the amino acid composition of WPI, SC and SPI in the absence and presence of resveratrol before and after storage for 30 days. The addition of resveratrol had no significant effect on the amino acid composition of the proteins before storage. The content of Cys ranked in order WPI > SPI > SC, and the surface and total sulfhydryl contents of SC were too low to be detected by the method of sulfhydryl analysis with DTNB (Figure 7). In the case of WPI alone, the content of Trp, Tyr, Thr, Lys, Met and Phe reduced after storage (Table 1), consistent with the indirect oxidation of WPI caused by the photodecomposition of folic acid [6]. Resveratrol had no effect on the change in the amino acid contents of WPI (Table 1). As for SC alone, the content of Trp, Tyr, Thr, Lys, Met, Asp and Arg reduced after storage and was more pronounced in the presence of resveratrol (Table 2). In addition, the content of Glu, Ser, Gly also reduced in the presence of resveratrol. The losses of Trp were about 11% for WPI and 79% and 87% for SC in the absence and presence of resveratrol, respectively. These results are consistent with a previous study that the tryptophan oxidation product, kynurenine, was higher in casein than β-LG upon photo-oxidation induced by
riboflavin [4]. In the case of SPI alone, Asp, Ser, His, Gly, Thr, Tyr, Cys, Val, Met, Lys, Trp reduced after storage and was more pronounced in the presence of resveratrol (Table 3). In addition, the content of Glu also reduced in the presence of resveratrol. The reduction in the kinds and contents of total amino acids ranked in the order of SPI > SC > WPI (Tables 1–3).
**Figure 8.** Carbonyl content of proteins in WPI-resveratrol (**A**), SC-resveratrol (**B**) and SPI-resveratrol (**C**) complex nanoparticles with various resveratrol concentrations during storage at 45 ◦C for 30 days. The concentration of proteins was 1%.
| doab | 2025-04-07T03:56:59.209709 | 17-11-2022 17:23 | {
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007afdec-bed4-405d-873d-c355ba9add0e.166 | *3.4. Storage Stability of Resveratrol*
By visual observation, all protein-resveratrol solutions were transparent and colorless except that SPI-resveratrol solutions were turbid at 1% protein (Table S1 and Figure S1). No significant change was observed for WPI-resveratrol solutions at 45 ◦C after 30 days. However, SPI-resveratrol and SC-resveratrol solutions changed from colorless to light yellow after storage. It has been reported that the wine with resveratrol changed from colorless to light yellow, due to its sensitivity to atmospheric oxidation [53]. After storage at 45 ◦C for 30 days, the total color difference (ΔE) and chroma change (ΔC\*) of resveratrol alone increased, respectively, from 1.24 to 3.06 and from 1.17 to 2.94, as its concentration increased from 25 to 100 μM (Table 4). The ΔE and ΔC\* of WPI, SC, SPI, and WPI-resveratrol solutions were less than those of resveratrol alone. However, the ΔE and ΔC\* of SC-resveratrol and SPI-resveratrol solutions increased as the polyphenol concentration increased and were greater than the sum of correspondingly individual values at each concentration. A previous study also reported that WPI as emulsifier showed a better effect on inhibiting color changes of lutein-loaded emulsions relative to SC [54].
**Table 4.** Total color difference (ΔE) and chroma change (ΔC\*) of WPI-resveratrol, SC-resveratrol and SPI-resveratrol complex solutions before and after storage at 45 ◦C for 30 days. The concentration of proteins was 1%.
Note: Different lower-case letters in the same column represent significantly different mean values, different upper-case letters in the same row represent significant different mean values (*p* < 0.05).
Resveratrol alone degraded during storage at 45 ◦C and its content remained 68–74% after 30 days (Figure 9). The retention of resveratrol was improved by WPI, and the protective effect decreased slightly as the protein concentration increased. After 30 days of storage, the retention of resveratrol at 25 μM was around 88, 84, and 74% at 0.01, 0.1, and 1% WPI (Figure 9A), respectively, and the polyphenol retention was proportional to its initial concentration (Figure 9). In contrast, the loss of resveratrol was accelerated by SPI, the effect of which was more pronounced when the protein concentrations were 0.1% and 1% than 0.01% (Figure 9). SC also accelerated the degradation of resveratrol, the effect of which was less than that of SPI and decreased as the polyphenol concentration increased. The retention of resveratrol was consistent with the color change of its corresponding samples (Table 4).
**Figure 9.** Retention of resveratrol alone (green) and in its WPI (black), SC (red) and SPI (blue) complex particles at various protein concentrations during storage at 45 ◦C. The concentrations of resveratrol were 25 (**A**), 50 (**B**) and 100 (**C**) μM.
| doab | 2025-04-07T03:56:59.210048 | 17-11-2022 17:23 | {
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007afdec-bed4-405d-873d-c355ba9add0e.167 | **4. Discussion**
Resveratrol self-aggregates at a concentration higher than 40 μM, due to the hydrophobic stacking of aromatic phenol rings [55]. The aggregation of resveratrol reduces its contact
with the external environment and affects its antioxidant activity with the highest value observed at a concentration of 30 μM [56]. Therefore, the retention of resveratrol increased from 68% to 74% as its concentration increased from 25 to 100 μM (Figure 9). α-Tocopherol (Log P ~ 8.84, https://go.drugbank.com/drugs/DB00163/ accessed on 8 July 2021), a hydrophobic vitamin E, was reported both bound in the molecular level and encapsulated as the aggregate in WPI particles, while naringenin (LogP ~ 2.84, https://go.drugbank. com/drugs/DB03467/ accessed on 8 July 2021), a polyhydroxy flavonoid, was bound in the molecular level [34]. Resveratrol (LogP ~ 3.4, https://go.drugbank.com/drugs/DB02709/ accessed on 8 July 2021) is more hydrophobic than naringenin but more hydrophilic than α-tocopherol. When calculated, based on the loading efficiency of resveratrol in Figure 5, the encapsulated amount of resveratrol in protein particles increased as the polyphenol concentration increased (Figure S3). It is thus possible that the aggregated resveratrol in protein particles increased as its concentration increased, which was supported by the transfer from the additive to the masking effect of total antioxidant activity (Figure 6). Therefore, the polyphenol retention increased with its concentration in protein particles (Figure 9).
For WPI, the solvent-accessible (bounded in the molecular level and in free state) resveratrol can scavenge and control the free radicals in the system within a certain range. Its oxidation was the least and not affected by resveratrol during storage for 30 days (Figures 7 and 8 and Table 1). At the same time, the stability of resveratrol was improved by WPI, with a retention of above 74% after 30 days (Figure 9). It is thus speculated that there is no reciprocal oxidation between WPI and resveratrol during storage. As the concentration of WPI increased, the loading efficiency of resveratrol increased (Figure 5), but the polyphenol stability decreased (Figure 9). These results suggest that the loaded microenvironment is not conducive to the polyphenol stability, compared to the free part in the WPI solution. The protective effect of WPI on resveratrol stability might not be attributed to the complex property of the protein.
For SPI, the encapsulated resveratrol located in the hydrophobic core could not exert its antioxidant capacity. Thus its oxidation was the most at the beginning and accelerated by resveratrol during storage after 10 days (Figures 7 and 8 and Table 3). At the same time, the stability of resveratrol decreased upon loading in SPI particles (Figure 9). These results suggest the occurrence of reciprocal oxidation between SPI and resveratrol. The co-oxidation has been reported for whey protein and Antarctic krill oil in oil-in-water emulsion [57]. The initial state of the SPI system contained more peroxides than SC and WPI (Figures 7 and 8), free radicals and hydroperoxides generated during protein oxidation may accelerate the degradation of resveratrol [58] (Figure 9). It has also been reported that ascorbic acid acted as a co-oxidant by generating superoxide anions in the presence of air and extracting hydrogen from the carrier [59]. Resveratrol is oxidized to generate H2O2 [60]. When the retention of resveratrol was between 59 and 73% after 10 days (Figure 9), the polyphenol may act as a co-oxidant to accelerate the oxidation of SPI (Figure 8).
However, most of the resveratrol in the SC system was encapsulated in the hydrophobic core of the protein, but also partially bounded with submicelles in the molecular level, which can play their antioxidant effect to a certain extent. The oxidation of SC was more pronounced than that of WPI but less than that of SPI at the beginning and during storage in the absence and presence of resveratrol (Figures 7 and 8 and Table 2). At the same time, the impact of SC on resveratrol stability basically changed from a protective to a harmful effect during storage (Figure 9). The antioxidant activity of SC was greater than that of WPI and SPI (Figure 6), and the loading efficiencies of resveratrol in SC particles were greater than those in SPI and WPI particles at protein concentrations of 0.1% and 1% (Figure 5). Therefore, the stability of resveratrol was initially improved by SC (Figure 9). A stable protein carrier can maintain the stability of polyphenols through scavenging free radicals and isolating the interference of external unfavorable factors [61]. Then, with the increasing oxidation of SC, the ability to scavenge free radicals was not enough to resist the auto-oxidation of SC. The system was out of balance and the protein changed from antioxidant to pro-oxidant to cause the co-oxidation with resveratrol (Figures 8 and 9).
According to the molecular mechanism of the protein–polyphenol interaction, the di-phenol part of polyphenol is easily oxidized by molecular oxygen and side-chain amino groups under certain conditions to form quinine, which can form a dimer in a side reaction and interact with the amino group of polypeptide or the irreversible reaction of the sulfhydryl side chain leads to the formation of protein cross-links. The closer the distance between the formed oxidation product and the α-carbon or other carbons of protein amino acid residues, the more easily the reaction occurs (Figure 4). Meanwhile, quinine can undergo condensation reactions to form high molecular weight, highly reactive brown tannins [17], which is verified in Table 4 and Figure S1. The formation of a covalent EGCGprotein complex involved the reaction of dimer quinone with protein nucleophilic side chains, such as lysine and cysteine residues, which is consistent with the results of amino acid composition in SC/SPI-resveratrol complex particles after storage (Tables 1–3). It has been assumed that the structure of SC and SPI gradually became flexible during storage and the exposed active groups benefited from the covalent interactions of protein-resveratrol complexations [62].
| doab | 2025-04-07T03:56:59.210325 | 17-11-2022 17:23 | {
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007afdec-bed4-405d-873d-c355ba9add0e.168 | **5. Conclusions**
WPI improved the storage stability of resveratrol, but SPI accelerated the loss of resveratrol, while the impact of SC on resveratrol stability basically changed from a protective to a harmful effect. The stability of polyphenols increased as the polyphenol concentration increased but decreased as the protein concentration increased. The loading efficiency of resveratrol in protein particles and the initial antioxidant activity of proteins were not the dominant factors to affect the storage stability of resveratrol. The effect of proteins on the stability of resveratrol was mainly dependent on their oxidation sensitivity. The co-oxidation of resveratrol with SPI and SC occurred during storage. The oxidation degree of WPI was the least and not affected by resveratrol. The results obtained suggest that WPI might be a better material to design an effective carrier for the long-term protection of resveratrol than SPI and SC. To our knowledge, it is the first time that the important role of protein oxidability on the stability of polyphenols during storage has been reported and provides useful guidelines for the long-term protection of polyphenols by protein-based carriers.
**Supplementary Materials:** The following supporting information can be downloaded at: https: //www.mdpi.com/article/10.3390/antiox11040647/s1, Figure S1: Appearance of WPI-resveratrol, SC-resveratrol and SPI-resveratrol complex nanoparticles before (A–C, respectively) and after (a–c, respectively) storage at 45 ◦C for 30 days. The concentration of proteins from left to right was 0.01%, 0.1% and 1%. The concentrations of resveratrol from left to right was 25, 50 and 100 μM; Figure S2: XRD patterns of resveratrol (black), proteins (blue), their physical mixtures (red) and resveratrolloaded protein particles (green). The concentration of protein was 1%; Figure S3: Loading efficiency of resveratrol in its complex particles with WPI (black), SC (red) and SPI (blue) at 0.01%, 0.1% and 1% after storage at 45 ◦C; Table S1: Turbidity of WPI-resveratrol, SC-resveratrol and SPI-resveratrol complex nanoparticles at various concentrations of proteins and resveratrol.
**Author Contributions:** X.Y.: conceptualization, investigation, writing—original draft, formal analysis. H.C.: resources, methodology. W.: writing—review and editing. H.D.: methodology, writing review and editing. W.H.: conceptualization. L.L.: conceptualization, resources, writing—review and editing, supervision. All authors have read and agreed to the published version of the manuscript.
**Funding:** This research was funded by the Postgraduate Research & Practice Innovation Program of Jiangsu Province (KYCX20-1863).
**Institutional Review Board Statement:** Not applicable.
**Informed Consent Statement:** Not applicable.
**Data Availability Statement:** The data are contained within the article and supplementary materials.
**Conflicts of Interest:** The authors declare no conflict of interest.
| doab | 2025-04-07T03:56:59.210752 | 17-11-2022 17:23 | {
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007afdec-bed4-405d-873d-c355ba9add0e.171 | **Encapsulation of Phenolic Compounds from a Grape Cane Pilot-Plant Extract in Hydroxypropyl Beta-Cyclodextrin and Maltodextrin by Spray Drying**
**Danilo Escobar-Avello 1,2, Javier Avendaño-Godoy 3, Jorge Santos 4,5, Julián Lozano-Castellón 1,6, Claudia Mardones 7, Dietrich von Baer 7, Javiana Luengo 3, Rosa M. Lamuela-Raventós 1,6, Anna Vallverdú-Queralt 1,6,\* and Carolina Gómez-Gaete 2,3,\***
**Abstract:** Grape canes, the main byproducts of the viticulture industry, contain high-value bioactive phenolic compounds, whose application is limited by their instability and poorly solubility in water. Encapsulation in cyclodextrins allows these drawbacks to be overcome. In this work, a grape cane pilot-plant extract (GCPPE) was encapsulated in hydroxypropyl beta-cyclodextrin (HP-*β*-CD) by a spray-drying technique and the formation of an inclusion complex was confirmed by microscopy and infrared spectroscopy. The phenolic profile of the complex was analyzed by LC-ESI-LTQ-Orbitrap-MS and the encapsulation efficiency of the phenolic compounds was determined. A total of 42 compounds were identified, including stilbenes, flavonoids, and phenolic acids, and a complex of (*epi*)catechin with *β*-CD was detected, confirming the interaction between polyphenols and cyclodextrin. The encapsulation efficiency for the total extract was 80.5 ± 1.1%, with restrytisol showing the highest value (97.0 ± 0.6%) and (*E*)-resveratrol (32.7 ± 2.8%) the lowest value. The antioxidant capacity of the inclusion complex, determined by ORAC-FL, was 5300 ± 472 μmol TE/g DW, which was similar to the value obtained for the unencapsulated extract. This formulation might be used to improve the stability, solubility, and bioavailability of phenolic compounds of the GCPPE for water-soluble food and pharmaceutical applications.
**Keywords:** microencapsulation; cyclodextrin; vine shoots; food waste; *Vitis vinifera* L.; polyphenols; stilbenoids; mass spectrometry; Fourier Transform Infrared Spectroscopy; Scanning Electron Microscopy
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007afdec-bed4-405d-873d-c355ba9add0e.172 | **1. Introduction**
The generation of food and agricultural waste is a growing problem, with negative impacts on the economy, environment, and human health. Therefore, the integral valorization of these wastes by conversion into bioenergy or recovery of chemical compounds for biobased products is a technological challenge for achieving a circular economy. Alternative ways of disposing of food waste include the valorization of byproducts as a source of phenolic compounds used to fortify high-consumption foods or to formulate new functional foods [1].
**Citation:** Escobar-Avello, D.; Avendaño-Godoy, J.; Santos, J.; Lozano-Castellón, J.; Mardones, C.; von Baer, D.; Luengo, J.; Lamuela-Raventós, R.M.; Vallverdú-Queralt, A.; Gómez-Gaete, C. Encapsulation of Phenolic Compounds from a Grape Cane Pilot-Plant Extract in Hydroxypropyl Beta-Cyclodextrin and Maltodextrin by Spray Drying. *Antioxidants* **2021**, *10*, 1130. https://doi.org/10.3390/ antiox10071130
Academic Editors: Li Liang, Hao Cheng and Daniel Franco Ruiz
Received: 10 June 2021 Accepted: 13 July 2021 Published: 15 July 2021
**Publisher's Note:** MDPI stays neutral with regard to jurisdictional claims in published maps and institutional affiliations.
**Copyright:** © 2021 by the authors. Licensee MDPI, Basel, Switzerland. This article is an open access article distributed under the terms and conditions of the Creative Commons Attribution (CC BY) license (https:// creativecommons.org/licenses/by/ 4.0/).
Grape canes of *V. vinifera* L. produced during pruning are the main byproduct of viticulture, and millions of tons are generated worldwide every year [2,3]. With the aim of developing a circular economy by an integrated biorefinery strategy, grape canes are being investigated as a high-value resource due to their attractive chemical composition and potential industrial applications. These include SO2 substitution in wine to improve wine quality [4], usage as a cosmetic ingredient [5] and as a filler in food packaging [4,6], and for the recovery of hemicellulosic oligosaccharides, lignin, and cellulosic substrates [7]. Above all, however, grape canes have been studied for their nutraceutical applications, as they contain high-added-value phenolic compounds with wide-ranging biological properties.
Grape canes contain a complex mixture of phenolic compounds, including phenolic acids (hydroxybenzoic acid and hydroxycinnamic acids), flavonoids (mainly proanthocyanidins, flavonols, flavanonol and flavanones), and stilbenes (monomers, dimers, and oligomers). The most abundant phenolic compounds are proanthocyanidins and stilbene oligomers [8,9]. The phenolic composition of *V. vinifera* plants is genetically determined and strongly influenced by environmental conditions such as water stress [10]. Other determining factors, which particularly affect the stilbenes content, are the cultivar [11], the geographic region of cultivation [12], and the time, temperature, and humidity of storage after pruning [13]. Moreover, the yields of phenolic compounds such as resveratrol and *ε*-viniferin are highly dependent on variables of the extraction methods such as grape cane particle size, the type of solvent, temperature, duration, and the effects of light [14]. Furthermore, the phenolic profile of grape canes can also be altered by the extraction and scale-up process, where oxidation, degradation, or polymerization of proanthocyanidins have been observed, as well as the formation of phenolic aldehydes [2]. Phenolic compounds are known to be unstable and sensitive to high temperature, light, pH, and oxidative and degradative enzymes, which affects the phenolic profile of extracts. It is therefore important to find a strategy to protect phenolic compounds, and preserve their biological activities and properties. Enhancing their bioaccessibility and bioavailability and promoting their transport for absorption by the human body are also of great interest. Accordingly, new approaches, such as encapsulation with cyclodextrins, have been developed to overcome these drawbacks. [15].
Cyclodextrins are highly biocompatible and have been approved by the Food and Drug Administration (FDA) as safe for humans. Several research studies have described the complexation of phenolic compounds with cyclodextrins [15], which provides protection from environmental conditions and improves bioactive shelf-life. Cyclodextrins, which have a truncated cone-shaped structure, possess a hydrophobic interior and a hydrophilic outer surface. Complexation in cyclodextrins improves the water solubility of phenolic compounds, which is otherwise relatively poor. The stability of the complex is maintained via hydrophobic forces, van der Waals interactions and hydrogen bonding [15]. Therefore, encapsulation of the bioactive molecule by cyclodextrin alters the physicochemical properties of both agents. Nevertheless, the effect of cyclodextrins on the profile of phenolic compounds from grape canes after encapsulation needs further study.
In this context, we encapsulated a previously characterized grape cane pilot-plant extract [2] in HP-*β*-CD by a spray-drying technique, using maltodextrin (MD) as a coating material. The physicochemical properties and parameters of the encapsulation process were determined, and the complex formation was verified by scanning electron microscopy (SEM) and Fourier Transform Infrared Spectroscopy with diamond attenuated total reflectance (FTIR-ATR). In addition, the phenolic profile of the inclusion complex was investigated using LC-ESI-LTQ-Orbitrap-MS, and encapsulation efficiency and antioxidant capacity were determined. The microencapsulated extract is envisaged as a functional ingredient of food, cosmetics, biomaterials, and other biobased products.
| doab | 2025-04-07T03:56:59.211160 | 17-11-2022 17:23 | {
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007afdec-bed4-405d-873d-c355ba9add0e.174 | *2.1. Chemicals and Reagents*
Gallic, 4-hydroxybenzoic, and ellagic acids, catechin, epicatechin, (*E*)-resveratrol, (*E*)-*ε*-viniferin, (*E*)-piceatannol, eriodictyol, taxifolin, quercetin, quercetin-3-*O*-glucoside, and quercetin-3-*O*-glucuronide were purchased from Sigma-Aldrich (St. Louis, MO, USA). Gallic acid and kaempferol-3-*O*-glucoside were acquired from Extrasynthèse (Genay, Auvergne-Rhône-Alpes, France). Isohopeaphenol and hopeaphenol were kindly given by the research group of Prof. Dr. Peter Winterhalter (Institute of Food Chemistry, Technical University Braunschweig, Lower Saxony, Germany). Light exposure was avoided when manipulating the standards.
HPLC-grade acetonitrile, formic acid, ethanol, and water were purchased from Merck (Darmstadt, Hesse, Germany). Ultrapure water was generated by a Milli-Q water purification system Millipore (Bedford, Massachusetts, USA). Potable ethanol (96%) from molasses employed for pilot-plant scale extraction was purchased from Oxiquim S.A. (Coronel, Concepción, Chile).
Hydroxypropyl beta-cyclodextrin (HP-*β*-CD, Kleptose®, HP oral grade) was purchased from Roquette Frères (Lestrem, Lillers, France). Maltodextrin (MD) (dextrose equivalent 16.5–19.5) was obtained from Merck KGaA, (Darmstadt, Hesse, Germany).
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007afdec-bed4-405d-873d-c355ba9add0e.175 | *2.2. Pilot-Plant Scale Extraction*
Grape cane extraction was performed on a pilot-plant scale (750 L) at 80 ◦C for 100 min, following our previously reported extraction process [2].
After winter pruning, we collected a total of 500 kg of wet sample grape canes (*V. vinifera* L. cv. Pinot noir) from plants in an organic vineyard at Viña De Neira, located in Ránquil, Itata Valley, Biobio Region, Chile. All samples were cut into 30–50 cm long pieces and stored for three months at room temperature (19 ◦C ± 5) and 30–70% relative humidity, according to previous reports by Riquelme et al. [16] and Patent [13]. We used 67 kg of dry grape canes from a total sample of 500 kg of wet samples for the pilot-plant extraction, before which grape canes were crushed in a Retsch grinder (model SM) at 300–2000 rpm until particle size was less than 1 cm. After extraction, the ethanol used as a solvent for the extraction was removed and recovered by distillation (absolute pressure 0.05 bar). The liquid extract was collected in a dark container and protected from light.
## *2.3. Preparation of Microcapsules by Spray-Drying*
The GCPPE was in a mixed ethanol/water solution (30:70 *v*/*v*). HP-*β*-CD was used to prepare the microcapsules in a proportion of 2.2% *w*/*v* with the extract. MD in a proportion of 10% *w*/*v* was used as the coating material. HP-*β*-CD was slowly added to a beaker containing 200 mL of GCPPE to avoid its agglomeration. The mixture was continuously stirred at room temperature and protected from the light for 24 h, during which the encapsulation took place. Then, MD was added slowly, and the mixture was stirred for a few minutes [17]. The microencapsulated and unencapsulated GCPPE were dried using a Büchi Mini Spray Dryer B-290 (Büchi, Flawil, Switzerland). Inlet temperature was maintained at 130 ◦C, while the outlet air temperature was 71 ◦C. Air inlet and airflow were 35–40 m3/h and 473 L/h, respectively. The spray dryer was equipped with a nozzle tip diameter of 0.7 mm and a peristaltic pump operated at 6–7 mL/min. The dried powder was collected and stored in an amber airtight container at 4 ◦C until analysis.
| doab | 2025-04-07T03:56:59.211707 | 17-11-2022 17:23 | {
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007afdec-bed4-405d-873d-c355ba9add0e.176 | *2.4. Determination of the Physical Properties of the Microencapsulated Powders* 2.4.1. Moisture Content and Total Solids
The moisture content of the samples was calculated from the weight loss after heating the sample to 105 ◦C for 6 h [18].
2.4.2. Process Yield (PY%)
The yield of the powder process was calculated considering the number of solids introduced into the spray drying system and the powder obtained at the end of the technological process [19]. The results were determined according to Equation (1):
$$\text{PY } (\%) = \frac{\text{Power after spray dry}}{\text{Solds introduced in the feeding}} \times 100\tag{1}$$
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007afdec-bed4-405d-873d-c355ba9add0e.177 | 2.4.3. Bulk Density
To determine the bulk density, about 10 g of the spray-dried sample was weighed, placed in a 25 mL graduated test tube, and the occupied volume was recorded [18].
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007afdec-bed4-405d-873d-c355ba9add0e.178 | 2.4.4. Angle of Repose
The angle of repose was determined using a fixed funnel by the following Equation (2) as in Dadi et al. [18]:
$$\text{Angle of response } (^{\circ}) = \tan^{-1} \text{ (H/R)}\tag{2}$$
where H is the height of the pile and R is its radius at the base.
#### 2.4.5. Size Distribution
The particle size distribution was determined in a particle size analyzer by laser diffraction with a Mastersizer 3000 (Malvern Instruments, Worcestershire, UK). Samples were dispersed in MilliQ water (900 mL) under constant stirring (2300 RPM) using a Hydro EV dispersion unit to achieve a homogeneous suspension. The particle size distribution in the powder (*span*) was calculated using Equation (3):
$$Spam = \begin{pmatrix} \mathbf{d}90 \ -\mathbf{d}\_{10} \end{pmatrix} / \mathbf{d}\_{50} \tag{3}$$
where d90, d10, and d50 are the equivalent volume diameters at 90%, 10%, and 50% cumulative volume, respectively [20].
| doab | 2025-04-07T03:56:59.211987 | 17-11-2022 17:23 | {
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007afdec-bed4-405d-873d-c355ba9add0e.179 | *2.5. Scanning Electron Microscopy*
The microcapsules and GCPPE were analyzed by scanning electron microscopy (SEM) using the JSM 6380 LV system (JEOL Techniques Ltd., Tokyo, Japan). The microscope was operated at 20 kV accelerating voltage. The samples were coated with a gold layer of about 150 Å in thickness, using an Edwards S 150 sputter coater (Agar Scientific, Standsted, UK) [21].
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007afdec-bed4-405d-873d-c355ba9add0e.180 | *2.6. Fourier Transform Infrared Analysis*
The FTIR absorption spectra of the individual samples of GCPPE, HP-*β*-CD, the inclusion complex (IC) (GCPPE+HP-*β*-CD+MD), and the physical mixture were analyzed separately. The physical mixture (PM) (GCPPE/HP-*β*-CD/MD) was prepared by accurately weighing HP-*β*-CD (100 mg), GCPPE (100 mg), and MD (20 mg), which were ground in a mortar until the mixture was homogeneous. The resulting physical mixture was immediately analyzed by FTIR-ATR.
The FTIR spectra were recorded using a Bruker Alpha T FTIR (Bruker, Germany) spectrophotometer equipped with a diamond attenuated total reflectance (ATR) unit. The 32 scans were acquired over a spectral range of 4000–500 cm−<sup>1</sup> with a resolution of 4 cm−<sup>1</sup> [22,23]. All spectra were acquired and processed using the OPUS 7.0 software.
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007afdec-bed4-405d-873d-c355ba9add0e.181 | *2.7. LC-ESI-LTQ-Orbitrap-MS Analyses*
Liquid chromatography (LC) analysis was conducted using an Accela chromatograph (Thermo Scientific, Hemel Hempstead, UK) equipped with a quaternary pump, photodiode array detector, and thermostated autosampler. Chromatographic separation was performed in an Atlantis T3 column 2.1 × 100 mm, 3μm (Waters, Milford, MA, USA). Gradient elution
of analytes was performed utilizing H2O/0.1% HCOOH (solvent A) and CH3CN (solvent B) at a continuous flow rate of 0.350 mL/min, and an injection volume of 5 μL. The following gradient was applied: 0 min, 2% B; 0–2 min, 8% B; 2–12 min, 20% B; 12–13 min, 30% B; 13–14 min, 100% B; 14–17 min, 100% B; 17–18 min, 2% B and the column was equilibrated to the initial conditions for 5 min [24].
The LC equipment was coupled to an LTQ-Orbitrap Velos mass spectrometer (Thermo Scientific, Hemel Hempstead, UK) employed for accurate mass measurements and equipped with an electrospray ionization (ESI) source operating in negative mode. The working parameters were as follows: source voltage, 4 kV; sheath gas, 20 a.u. (arbitrary units); auxiliary gas, 10 a.u.; sweep gas, 2 a.u.; and capillary temperature, 275 ◦C. Default values were used for most of the other acquisition parameters (FT Automatic gain control target <sup>5</sup>·10<sup>5</sup> for MS mode and 5·10<sup>4</sup> for MSn mode). Samples were analyzed in FTMS mode with a resolving power of 30,000 (FWHM at *m*/*z* 400) and data-dependent MS/MS events were acquired with a resolving power of 15,000. The most intense ions detected in the FTMS spectrum were chosen for the data-dependent scan. The parent ions were fragmented by high-energy C-trap dissociation by normalized collision energy of 35 V and an activation time of 10 ms. The mass range in FTMS mode was from *m*/*z* 100 to 1500. Instrument control and data recovery were conducted using Xcalibur 3.0 software (Thermo Fisher Scientific). The tentative identification of analytes was performed by comparing MS/MS spectra with fragments found in databases and the literature when no standard compound was available [25].
Individual compounds were semi-quantified using pure standards or the most similar compounds. Some analytes, such as glycosylated forms, dimers, or oligomers, were semi-quantified using the aglycone form of the monomer [2].
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007afdec-bed4-405d-873d-c355ba9add0e.182 | *2.8. Encapsulation Efficiency*
Encapsulation efficiency was calculated by considering the nonencapsulated compounds present in the GCPPE before the encapsulation process and the encapsulated compounds after the spray drying process. The identification and quantification of phenolic compounds from GCPPE were specified in a previous article [2] (see detail Supplementary Materials Table S1). Extraction of polyphenols from the microcapsules was performed according to the procedure of Robert et al. [26] with minor modifications. Before the analysis, samples were dissolved in highly pure deionized water containing 0.1% *v*/*v* formic acid (1 mg/mL). The GCPPE was centrifuged at 4000 rpm for 5 min at 4 ◦C. The supernatant was recovered, and the extraction procedure was repeated twice. The supernatants were combined and evaporated under nitrogen flow, and the residue was reconstituted in 0.1% aqueous formic acid (5 mL). The samples were filtered through 0.20 μm PTFE membrane filters (Waters Corporation, Milford, CT, USA) into an amber vial. Subsequently, both samples were analyzed using LC -ESI-LTQ-Orbitrap- MS to determine the individual degree of encapsulation according to the method described above (Section 2.7). We estimated the encapsulation efficiency (EE), according to Radünz et al. [27], based on Equation (4):
$$\text{EE} \left( \% \right) = \frac{\text{Phenodic compound of GCPPE} - \text{Phenodic compound of the capsule}}{\text{Phenodic compound of GCPPE}} \times 100\tag{4}$$
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007afdec-bed4-405d-873d-c355ba9add0e.183 | *2.9. Antioxidant Capacity Assay*
The assay of oxygen radical absorbance capacity using fluorescein (ORAC-FL) was carried out according to the method reported by Ou et al. [28]. The calibration curves were prepared with Trolox, and results reported as μmol Trolox equivalents (TE) by grams of dried weight (DW) (TE/g DW). All assays were performed in triplicate and protected from light.
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007afdec-bed4-405d-873d-c355ba9add0e.184 | *2.10. Statistical Analysis*
All samples were run at least three times, and results are expressed as means ± standard deviations. A *p*-value < 0.05 was considered statistically significant using Student's
*t*-test with 95% confidence. Statistical analyses were determined using GraphPad Prism 8.0.1 (GraphPad Software, San Diego, CA, USA).
#### **3. Results and Discussion**
#### *3.1. Physical Characterization of the Microencapsulated Powder*
Table 1 shows the physicochemical properties and process parameters for the GCPPE and inclusion complex (IC) (GCPPE+HP-*β*-CD+MD). The process yield obtained by spraydrying for IC (GCPPE+HP-*β*-CD+MD) was 83.8 ± 2.6%, which was two-fold higher than for GCPPE alone (38.4 ± 1.2%). Similarly, the total solids increased 2.6-fold for the microencapsulated formulation. The high yield of powdered microparticles can be attributed to the rapid formation of the drying crust, which prevents the powder from adhering to the drying chamber [29]. Our result constitutes an improvement on the yield (64.5 ± 1.5%) reported by Davidov-Pardo et al. [30], who microencapsulated a grape seed extract using MD. The high values obtained in our work are promising for the development of industrial-scale applications.
**Table 1.** Physical characteristics and process parameters for the grape cane phenolic extract (GCPPE) and inclusion complex (GCPPE+HP-β-CD+MD).
Results are expressed as means ± standard deviations, and values with different superscripts letters in a column indicate significant differences at *p* < 0.05.
> The particle size distribution and median particle diameter were smaller in the IC (GCPPE+HP-*β*-CD+MD) than in the GCPPE alone; the particle sizes were 10.9 μm and 17.5 μm, respectively. According to the literature, the diameter of spray-dried particles depends on the properties of the material, the drying conditions, the atomization method used, and the concentration and viscosity of the encapsulated material [31]. The *span* values of the IC (GCPPE+HP-*β*-CD+MD) and GCPPE were very similar, 6.14 and 6.15, respectively, and were higher than those reported for an aqueous grape skin extract microencapsulated with Arabic gum, polydextrose, and partially hydrolyzed guar gum, which ranged from 1.91 to 5.99 [32]. A lower *span* value is a desirable result, as it indicates a more homogeneous particle size distribution [32].
> The bulk density was 0.10 ± 0.01 g/mL and 0.19 ± 0.01 g/mL for the GCPPE and IC (GCPPE+HP-*β*-CD+MD), respectively, being lower than the values reported for encapsulated rosemary essential oil (0.25–0.34 g/mL) [31] or soy milk (0.21–0.22 g/mL) [20]. The slightly higher bulk density of the IC (GCPPE+HP-*β*-CD+MD) vs. the GCPPE indicates an improved powder flow, as a more densely packed powder reflects weaker forces between the particles [20]. Density is an important factor for the packaging, transportation, and marketing of a microencapsulated product. A dry product with high density can be stored in a smaller container compared to a less dense product [31]. The flowability of the samples was also determined by the angle of repose, which was 34.8◦ ± 0.5 for the GCPPE and 36.9◦ ± 1.3 for the IC (GCPPE+HP-*β*-CD+MD), showing no statistical difference between them. A similar value was obtained in a study on the microencapsulation of bioactive products from a *Moringa stenopetala* leaf extract using MD, where the angle of repose was 37.26◦ ± 1.01 [18].
## *3.2. Surface Morphology: SEM Analysis*
SEM can be used to determine the surface morphology of materials and is recognized as an auxiliary method for monitoring the formation of inclusion complexes. The structure and size of the GCPPE and IC (GCPPE+HP-*β*-CD+MD) in the solid state obtained from the spray-drying process were analyzed through microscopy. Microcapsules should preferably have a slightly spherical form and a uniform and smooth cover with minimum fractures and signs of collapse [33]. The SEM micrographs showed that the GCPPE was composed of a mixture of non-spherical particles with irregular surfaces and other larger spherical microparticles (Figure 1A–C). In contrast, the IC (GCPPE+HP-*β*-CD+MD) was spherical, and without visible pores on a smooth surface; microparticles of a variable size but with similar morphology were observed together (Figure 1D–F). These significant morphological changes are probably due to a loss of crystallinity of the guest molecule after its inclusion in the cyclodextrin [34]. The SEM results provided evidence for the formation of the IC (GCPPE+HP-*β*-CD+MD), which was subsequently supported by mass spectrometry and FTIR analysis.
**Figure 1.** Scanning electron microscopy micrographs of the GCPPE (**A**–**C**) and IC (GCPPE+HP-*β*-CD+MD) (**D**–**F**) at different magnifications.
| doab | 2025-04-07T03:56:59.212599 | 17-11-2022 17:23 | {
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007afdec-bed4-405d-873d-c355ba9add0e.185 | *3.3. FTIR Analysis of Spray-Dried Powders*
FTIR-ATR is a useful method to detect the formation of inclusion complexes, which are revealed by changes in the FTIR spectra, such as the reduction, disappearance, or shift of absorption bands, due to weak intermolecular interactions [35]. The FTIR spectra of the GCPPE, HP-*β*-CD, IC (GCPPE+HP-*β*-CD+MD), and physical mixture (PM) (GCPPE/HP-*β*-CD/MD) are shown in Figure 2.
**Figure 2.** FTIR-ATR spectra of the GCPPE (**red**), HP-*β-*CD (**purple**), IC (GCPPE+HP-*β*-CD+MD) (**blue**), and PM (GCPPE/HP-*β*-CD/MD) (**green**).
The FTIR spectrum of the GCPPE (Figure 2, red) showed specific bands associated with phenolic compounds (in bold). The peaks at 1602 cm−<sup>1</sup> and 1448 cm−<sup>1</sup> were owing to the C=C stretching vibration of the phenolic aromatic ring and the C–H bending vibrations of the CH2 groups. The peak at 1239 cm−<sup>1</sup> was assigned to C=O stretching of gallic or ellagic acid components due to the presence of hydrolyzable tannins [36]. The band at 1513 cm−<sup>1</sup> was due to the C–C benzene skeletal vibrations of stilbenoids. The strong band at 1032 cm−<sup>1</sup> is attributed to C–O stretching in phenolic compounds [22]. Moreover, the band at 831 cm−<sup>1</sup> was due to the C-H out-of-plane bending vibrations of aromatic compounds. On the other hand, the signals at 2918 cm−<sup>1</sup> and 2850 cm−<sup>1</sup> were related to the CH2 asymmetric and symmetric stretch vibration in aliphatic hydrocarbons [37]. Additionally, a peak at 1103 cm−<sup>1</sup> was observed owing to the C–H in-plane bending vibration [36].
The FTIR spectrum of HP-*β*-CD (Figure 2, purple) showed bands at 3341 cm−<sup>1</sup> due to O–H stretching vibrations and 2927 cm−<sup>1</sup> due to C–H stretching vibrations. The peaks at 1645 cm−<sup>1</sup> correspond to the bending of H–O–H, at 1151 cm−<sup>1</sup> to C–O vibration, and at 1020 cm−<sup>1</sup> to the C–O–C symmetric stretching vibration. The peak at 849 cm−<sup>1</sup> was due to an α-type glycosidic bond [38]. The band at 1080 cm−<sup>1</sup> was ascribed to C–C stretching vibrations, and the peak at 1410 cm−<sup>1</sup> to C–C–H and O–C–H bending [39]. The presence of the hydroxypropyl group was recognized by a peak at 2970 cm−<sup>1</sup> corresponding to the antisymmetric vibration of methyl groups. Additionally, a peak was observed at 1367 cm<sup>−</sup>1, which was ascribed to the bending vibration of the methyl group [40].
The spectrum of the IC (GCPPE+HP-*β*-CD+MD) (Figure 2, blue) shows that some of the characteristic peaks of the GCPPE and HP-*β*-CD have shifted, decreased, or disappeared. The bands at 1645 cm−<sup>1</sup> and 1020 cm−<sup>1</sup> of HP-*β*-CD have shifted to 1634 cm−<sup>1</sup> and 1015 cm−1, respectively, in the IC. Two peaks of the GCPPE at 1239 cm−<sup>1</sup> and 1513 cm−1, ascribed to the presence of hydrolysable tannins and stilbenes, respectively, have shifted to 1246 cm−<sup>1</sup> and 1514 cm<sup>−</sup>1, showing a sharp reduction in intensity due to the complexation, whereas the peak at 2850 cm−<sup>1</sup> disappeared. Furthermore, the bands of the GCPPE at 1602 cm−<sup>1</sup> and 1448 cm<sup>−</sup>1, related to the phenolic aromatic ring, completely disappeared in the IC (GCPPE+HP-*β*-CD+MD). These findings may indicate that the phenolic rings became embedded in the HP-*β*-CD cavities.
The FTIR spectrum of the PM (Figure 2, green) showed a simple overlap between the individual components of GCPPE, HP-*β*-CD, and MD. Insignificant variations in intensity were detected, indicating a mixture of the three components without interactions between them.
| doab | 2025-04-07T03:56:59.212889 | 17-11-2022 17:23 | {
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007afdec-bed4-405d-873d-c355ba9add0e.186 | *3.4. Phenolic Profile of the IC (GCPPE+HP-β-CD+MD) by LC-ESI-LTQ-Orbitrap-MS*
We performed a targeted analysis of phenolic compounds in the IC (GCPPE+HP-*β*-CD+MD) using LC-ESI-LTQ-Orbitrap-MS. Table 2 shows 42 identified compounds with their accurate mass, theoretical mass, retention times (min), molecular formula, error (ppm) between the found mass and the accurate mass of each compound, and the MS/MS fragment ions with their respective intensities used for identification. The identification was further supported by comparisons with mass spectra databases and the literature. In addition, 13 phenolic compounds were identified by comparing the retention times and their masses with pure standards. We have reported the fragmentation patterns of most of these compounds in previous studies using an analytical [8] and pilot-scale extraction [2].
*Antioxidants* **2021**, *10*, 1130
**Table 2.** Identification of phenolic compounds in the IC (GCPPE+HP-*β*-CD+MD), an adduct of *β*-CD and a complex between *β*-CD with (*epi*)catechin using LC-ESI-LTQ-Orbitrap-MS in negativemode.
153
*Antioxidants* **2021**, *10*, 1130
**Table2.***Cont.*
Of the 42 compounds identified in the present work, 22 had been previously detected in both the analytical and pilot-plant extractions [2,8]: gallic acid (*m*/*z* 169.0138, −2.41 ppm), monogalloyl-glucose (*m*/*z* 331.0662, −2.65 ppm), protocatechuic acid-*O*-hexoside (1) (*m*/*z* 315.0718, −1.07 ppm), protocatechuic acid (*m*/*z* 153.0190, −2.16 ppm), protocatechuic acid-*O*-hexoside (2) (*m*/*z* 315.0717, −1.30 ppm), syringic acid hexoside (*m*/*z* 359.0933, 2.21 ppm), catechin (*m*/*z* 289.0722, 1.43 ppm), epicatechin (*m*/*z* 289.0715, −1.00 ppm), restrytisol (A or B) (*m*/*z* 471.1441, −1.86 ppm), taxifolin (*m*/*z* 303.0508, −0.78 ppm), astilbin (*m*/*z* 449.1084, −1.12 ppm), stilbenoid heterodimer (caraphenol B/C) (*m*/*z* 469.1292, −0.06 ppm), eriodictyol-*O*-glucoside (*m*/*z* 449.1089, −0.04 ppm), stilbenoid dimer (*m*/*z* 469.1292, −0.07 ppm), pallidol (*m*/*z* 453.1349, 2.24 ppm), (*E*)-resveratrol (*m*/*z* 227.0715, 0.77 ppm), stilbenoid dimer (resveratrol dimer) (*m*/*z* 453.1351, 1.57 ppm), resvertarol-*O*hexoside (*m*/*z* 615.1868, −0.62 ppm), eriodictyol (*m*/*z* 287.0558, −0.92 ppm), hopeaphenol (*m*/*z* 905.2607, −2.24 ppm), isohopeaphenol (*m*/*z* 905.2588, 2.15 ppm) and (*E*)-ε-viniferin (*m*/*z* 453.1348, 1.03 ppm). These findings indicate that these compounds are stable through each step of the production process, including microencapsulation.
On the other hand, seven compounds identified in the microencapsulated extract had previously been detected only in the analytical extraction [8]: procyanidin dimer (1) (*m*/*z* 577.1348, −0.69 ppm), procyanidin dimer (2) (*m*/*z* 577.1346, −0.35 ppm), epicatechin gallate (*m*/*z* 441.0821, −1.30 ppm), (*E*)-piceatannol (*m*/*z* 243.0665, 0.79 ppm), viniferin diglycoside (*m*/*z* 777.2397, −0.36 ppm), (*E*)-ω-viniferin (*m*/*z* 453.1346, 0.62 ppm), and stilbenoid tetramer (vitisin A/B/C/D) (*m*/*z* 905.2599, −3.05 ppm). Thus, although these compounds were not detected in the pilot-scale extraction, they were recovered after the microencapsulation process. Similar to our results, the incorporation of *β*-CD enabled an effective and selective recovery of flavan-3-ols [41] and stilbenes [42], resulting in a cleaner analytical extract phenolic profile. Additionally, three compounds previously detected in the pilot-scale extraction [2]: protocatechuic aldehyde (*m*/*z* 137.0241, −2.02 ppm), kaempferol-3-*O*-glucoside (*m*/*z* 447.0931, −0.49 ppm), and ethyl protocatechuate (*m*/*z* 181.0504, −1.02 ppm), were also recovered and identified in the microencapsulated extract.
Finally, ten compounds were identified only in the IC (GCPPE+HP-*β*-CD+MD), and not in the analytical or pilot extracts: five flavonoids, three stilbenes, an adduct of *β*-CD, and a complex of *β*-CD with (*epi*)catechin.
*Flavonoids.* The taxifolin isomer (*m*/*z* 303.0503, −2.32 ppm) showed ions at *m/z* 285.0390, owing to the initial loss of a water molecule and ions at *m*/*z* 177.0184 and 125.0237 due to cleavage of the C ring, respectively. Quercetin (*m*/*z* 301.0354, 0.06 ppm) was identified and confirmed by comparison with a pure standard. Myricetin (*m*/*z* 317.0301, −0.63 ppm), tentatively identified by its fragmentation pattern, produced ions at *m*/*z* 178.9981 (1,2A−) and 151.0032 (1,3A−) due to retro-Diels–Alder fragmentation [43], and at *m*/*z* 192.0058 due to the loss of the B ring. Although they were not identified in our previous studies, these compounds have been recovered, detected, and quantified by other authors using microwave-assisted, subcritical water, and conventional extraction techniques [44].
Dihydrokaempferol-*O*-rhamnoside (engeletin) (*m*/*z* 433.1139, −0.38 ppm), a compound previously reported in grape stems [45], was tentatively identified and showed fragment ions at *m*/*z* 269.0446, 287.0550, and 259.0603. An undefined tetrahydroxyisoflavanone (*m*/*z* 287.0561, −0.18 ppm) gave product ions at *m*/*z* 259.0602, 243.0652, and 201.0547, and was provisionally identified as 2,6,7,4 -tetrahydroxyisoflavanone, based on the exact mass and fragmentation pattern. However, as dihydrokaempferol and eriodictyol chalcone have similar structures, the identity of this compound could not be accurately defined using our spectrometric approach.
*Stilbenes*. A stilbenoid dimer (maackin, Figure 3A) (*m*/*z* 485.1242, 0.01 ppm) showed product ions at *m*/*z* 467.1125, 375.0865, and 363.0863, which were generated by the loss of a water molecule (18 Da), resorcinol (110 Da), and 2-hydroxy-4-methylenecyclohexa-2,5-dienone (122 Da), respectively [46]. This compound has a structure consisting of two
piceatannol units, and the most likely assignment is maackin A, which was identified previously in *V. vinifera* stalks [47].
**Figure 3.** Representative stilbenes tentatively identified in the microencapsulated extract. (**A**) Maackin (C28H22O8); (**B**) Viniferol E (C56H44O13); (**C**) Viniphenol A (C84H64O18).
A stilbenoid tetramer (Figure 3B) (*m*/*z* 923.2680, 0.68 ppm) was tentatively identified as viniferol E and yielded product ions at *m*/*z* 905.2576, 881.2573, 801.2318, 783.2209 and 707.1898. The product ions at *m*/*z* 905.2576 and 881.2573 were due to a loss of H2O (18 Da) and C2H2O (42 Da), respectively, and at *m*/*z* 801.2318 probably to the loss of the group C7H6O2 (122 Da). The ion at *m*/*z* 801.2318 was further fragmented to ions at *m*/*z* 783.2209 and *m*/*z* 707.1898 by the loss of a water molecule (18 Da) and a phenol group (94 Da), respectively. Viniferol E was previously detected and quantified from grapevine canes by subcritical water extraction [48].
A stilbenoid hexamer, viniphenol A (Figure 3C) (*m*/*<sup>z</sup>* 679.1969 [M − 2H]2<sup>−</sup>, −1.12 ppm), was detected as a doubly charged ion with product ions at *m*/*z* 905.2584, 585.1543, 491.1126, 453.1333 and 359.0914. The product ion at *m*/*z* 905.2584 shows the presence of a stilbenoid tetramer molecule, probably formed by the loss of a stilbenoid dimer (454 Da) from the deprotonated hexamer. The high-intensity product ion at *m*/*z* 585.1543 could be attributed to the loss of a phenol group (94 Da) from the stilbenoid trimer (*m*/*z* 679). The product ion at *m*/*z* 585.1543 underwent fragmentation to ions at *m*/*z* 491.1126 and *m*/*z* 359.0914, which could be attributed to the loss of a phenol group (94 Da) and a stilbenoid dimer (226 Da), respectively. Finally, a low intensity stilbenoid dimer fragment was observed at *m*/*z* 453.1333. Viniphenol A was previously isolated from *V. vinifera* stalks by centrifugal partition chromatography, while its structure was proposed based on the analysis of spectroscopic data and molecular modeling under NMR conditions [47].
According to the supplier, the HP-*β*-CD used for encapsulation has a maximum *β*-CD impurity of 1.5%. The presence of [*β*-CD+ HCOO]— (*m*/*<sup>z</sup>* 1179.3679, −0.04 ppm) and a complex of (*epi*)catechin with *<sup>β</sup>*-CD [*β*-CD +(*epi*)catechin]— (*m*/*<sup>z</sup>* 1423.5708, −0.71 ppm) were detected and identified by comparison with the mass spectra reported by Zy˙ zelewicz et al. [ ˙ 49]. The detection of this complex by mass spectrometry confirmed the interaction between polyphenols and cyclodextrins, in agreement with the SEM and FTIR analysis.
| doab | 2025-04-07T03:56:59.213090 | 17-11-2022 17:23 | {
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007afdec-bed4-405d-873d-c355ba9add0e.187 | *3.5. Encapsulation Efficiency*
GCPPE is a complex mixture of various compounds that have different physical and chemical properties and abilities to form interactions and bind within the HP-*β*-CD cavity. As shown earlier, the phenolic compounds in the microencapsulated extract are phenolic acids and derivatives, flavonoids, and stilbenes. The individual EE (%) for twenty of these
compounds was calculated and presented in Table 3. The other compounds were identified but not quantified due to their low amounts (see detail Supplementary Materials Table S1).
**Table 3.** Encapsulation efficiency (%).
Results are given as means ± standard deviations. For each phenolic class, we calculated the weighted average of the concentration of the respective metabolites. The total weighted average was calculated by weighting all quantified compounds. All encapsulation efficiencies were calculated according to Equation (4) (Section 2.8).
The average EE for all the analyzed compounds was 80.5 ± 1.1%. Several authors have studied the encapsulation of phenolic compounds from wine byproducts using different encapsulation materials. Davidov-Pardo et al. [30] reported a similar polyphenol EE of 82% for a commercial grape seed extract microencapsulated by spray drying using MD as the wall material. Moschona and Liakopoulou-Kyriakides [50] found a low EE of 55% to 79% for grape marc and lees phenolic extracts from white and red wine encapsulated with alginate and chitosan. Lavelli and Sri Harsha [51] also reported a low EE of 68% in a study where alginate hydrogel was used as an agent to encapsulate phenolic compounds from grape skin. Another study using grape skin extracts, encapsulated in water-in-oilin-water (W/O/W) double emulsions, observed an improved EE of 87.74 ± 3.12% for anthocyanins [52]. The EE depends on a variety of factors, such as the technique used, the solubility and size of the guest molecule relative to the cavity of the host, the concentrations of the host and guest molecules, the binding constant within the guest and host, etc. [53]. As we did not find a similar study in the literature that employed the same raw material, encapsulant, and analytical methods to determine the concentration of guest molecules, we also analyzed the individual encapsulation of twenty compounds present in the IC (GCPPE+HP-*β*-CD+MD).
The average EE for the phenolic acids and derivatives was 81.5 ± 0.7%, the lowest value being obtained for protocatechuic acid-*O*-hexoside 1 (54.3 ± 2.4%) and the highest for hydroxybenzaldehyde (95.8 ± 0.6%). The EE for protocatechuic acid was 66.8 ± 9.8%, higher than the value reported by Taofiq et al. [54], who determined an EE of 50.3% for protocatechuic acid encapsulated by the atomization/coagulation technique, using sodium alginate in combination with calcium chloride (CaCl2) to promote alginate gelation. The EE for hydroxybenzaldehyde obtained here is higher than the value (46.50%) reported for *p*-hydroxybenzaldehyde in a *β*-CD inclusion complex [55].
The EE for gallic acid, 83.4 ± 0.6%, was slightly higher than the value reported by da Rosa et al. [56], who determined an EE of 80.0 ± 1.4% for microencapsulated gallic acid using *β*-CD and the lyophilization method. However, Olga et al. [53] obtained a higher EE of 89.22% in a complex with HP-*β*-CD, a result that was reduced to 77.34% when the gallic acid was co-encapsulated with *trans*-ferulic acid. The authors suggested a possible antagonistic relationship between the two phenols in the HP-*β*-CD cavity.
The EE of 87.6 ± 0.5% observed for ellagic acid pentoside was considerably higher than the 55.2% reported for ellagic acid (aglycone) encapsulated with polyvinyl alcohol [57]. The EE for caftaric acid—a hydroxycinnamic acid—was very high (87.1 ± 2.2%) in comparison with the values reported for other hydroxycinnamic acids, such as the essential oil-encapsulated chitosan-*ρ*-coumaric acid (42 ± 1%) [58] and chlorogenic acid (77.5%) in *β*-CD nanosponges [59], respectively. Finally, to our knowledge, this is the first time that the EE for ethyl protocatechuate (69.7 ± 5.5%) and protocatechuic aldehyde (75.6 ± 4.2%) has been reported.
The average EE for flavonoids was 85.2 ± 2.4%. The values for astilbin isomer (2) and astilbin (1) were particularly high, 92.0 ± 1.1% and 82.7 ± 2.3%, respectively. Zheng and Zhang [60] reported a lower EE of 80.1% for astilbin encapsulated with zein−caseinate nanoparticles by the antisolvent method.
Quercetin-3-*O*-glucuronide (88.0 ± 3.1%) and quercetin-*O*-glucoside (81.8 ± 7.0) also had high EE values. Tchabo et al. [61] obtained a lower EE (63.90–66.45%) for quercetin-3-*O*-glucoside in a spray-dried mulberry leaf extract prepared with MD, yet the EE was higher (91.71–93.95%) when sodium carboxymethyl cellulose was used. To the best of our knowledge, no EE values have been previously reported for quercetin-3-*O*-glucuronide.
The EE for eriodictyol was 74.2 ± 4.7%, similar to the almost 70% reported for naringenin (another flavanone) in polymer PLGA nanoparticles prepared by an emulsiondiffusion-evaporation method [62].
The stilbene group had the lowest mean EE (78.6 ± 1.9%), mainly because of the low value obtained for (*E*)-resveratrol (32.7 ± 2.8%). Nevertheless, previous studies have shown that the cyclodextrin encapsulation of resveratrol increases its solubility, stability, and bioactivity (antioxidant and anticarcinogenic properties) [15]. Furthermore, the HP-*β*-CD complex exhibits a strong H-bonding interaction with molecules such as oxyresveratrol [63]. In our study, we found a high EE (97.0 ± 0.6%) for restrytisol (A or B), an oxidized resveratrol dimer, which may be related to the van der Waals force interaction and hydrogen bonding between the guest compound and the HP-*β*-CD.
A good EE was obtained for stilbene dimers such as pallidol (74.6 ± 3.8%), (*E*)-ε-viniferin (76.8 ± 3.5%), and an undefined resveratrol dimer (94.1 ± 2.1%). Previously, ε-viniferin, a resveratrol dimer, was encapsulated in phospholipid-based multi-lamellar liposomes called spherulites or onions. This formulation gave a lower EE (58 ± 3%) than in our study, but increased the water solubility of this stilbene more than five-fold and provided protection against its UV-induced isomerization [64]. Finally, the low EE (65.8 ± 8.6%) obtained for the stilbenoid tetramer is probably due to its large size and polar surface area.
| doab | 2025-04-07T03:56:59.213517 | 17-11-2022 17:23 | {
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007afdec-bed4-405d-873d-c355ba9add0e.188 | *3.6. Antioxidant Capacity*
Several authors suggest that the antioxidant capacity of phenolic compounds is improved by encapsulation in cyclodextrins [15]. The ORAC value has been applied to standardize the antioxidant activity of herbal extracts and foods, and is widely used as an accurate indicator of antioxidant activity in vivo [65].
The antioxidant capacity of the IC (GCPPE+HP-*β*-CD+MD) by ORAC-FL was 5300 ± 472 μmol TE/g DW, similar to the 4612 ± 155 μmol TE/g DW reported in our previous GCPPE study [2]. The GCPPE microencapsulated with HP-*β*-CD retains its antioxidant capacity and its formulation may improve stability, solubility, and bioavailability for applications in the food, cosmetic and pharmaceutical industries.
## **4. Conclusions**
Phenolic compounds from a grape cane pilot-plant extract were successfully encapsulated in an inclusion complex (GCPPE+HP-*β*-CD+MD). The microencapsulated extract was rich in stilbenes, especially oligomers, flavonoids, and phenolic acids. A complex of (*epi*)catechin and *β*-CD was detected by mass spectrometry, which confirmed the interaction between polyphenols and cyclodextrin. The formation of the inclusion complex was also supported by FTIR-ATR and SEM analyses. HP-*β*-CD provided a high EE for phenolic compounds, with a mean of 80.5 ± 1.1%, the highest values being obtained for restrytisol (97.0 ± 0.6%), stilbenoid heterodimer (1) (96.8 ± 0.4%) and hydroxybenzaldehyde (95.8 ± 0.6%), and the lowest for (*E*)-resveratrol (32.7 ± 2.8%). The antioxidant capacity of the inclusion complex was similar to the unencapsulated extract. Considering the protection afforded the phenolic compounds by the inclusion complex, it is expected that the formulation may improve their stability, solubility, and bioavailability in water-soluble applications for the food and pharmaceutical industries.
| doab | 2025-04-07T03:56:59.213893 | 17-11-2022 17:23 | {
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007afdec-bed4-405d-873d-c355ba9add0e.189 | **5. Patents**
The preparation of grape canes before extraction in the pilot plant followed the procedure described in the Patent [13]. The formulation of the microparticles of phenolic compounds from the extract of grape canes of *V. vinifera* and the application of HP-*β*-CD to form the IC and the use of MD as a coating agent are described in the Patent [17].
**Supplementary Materials:** The following are available online at https://www.mdpi.com/article/10 .3390/antiox10071130/s1, Table S1: Identification and quantification of phenolic compounds from GCPPE using LC-ESI-LTQ-Orbitrap-MS in negative mode.
**Author Contributions:** Conceptualization, D.E.-A., C.G.-G., C.M., J.L., D.v.B. and A.V.-Q.; methodology, D.E.-A., J.A.-G., J.S., J.L.-C., C.G.-G., J. L and A.V.-Q.; validation, A.V.-Q. and C.G.-G.; formal analysis, D.E.-A., J.A.-G., J.L.-C. and J.S.; investigation, D.E.-A., J.A.-G. and J.S.; resources, A.V.-Q., C.G.-G., R.M.L.- R., C.M., J.L. and D.v.B.; data curation, D.E.-A., J.A.-G. and J.S.; writing—original draft preparation, D.E.-A.; writing—review and editing, D.E.-A., A.V.-Q., C.G.-G., C.M., J.A.-G., D.v.B., R.M.L.-R. and J.S.; visualization, D.E.-A.; supervision, A.V.-Q., C.G.-G., R.M.L.-R. and C.M.; project administration, D.E.-A., A.V.-Q., C.G.-G., C.M. and R.M.L.-R.; funding acquisition, D.E.-A., C.M., A.V.-Q. and R.M.L.-R. All authors have read and agreed to the published version of the manuscript.
**Funding:** This research was supported by the Agencia Nacional de Investigación y Desarrollo (ANID)/PCI-REDES170051; ANID PIA/APOYO CCTE AFB170007; CORFO 14 IDL2- 30156, from Chile; CICYT [AGL2016-75329-R], CIBEROBN from the Instituto de Salud Carlos III, ISCIII from the Ministerio de Ciencia, Innovación y Universidades, (AEI/FEDER, UE) and Generalitat de Catalunya (GC) [2017SGR 196]. Danilo Escobar-Avello is grateful to ANID/Scholarship Program/DOCTORADO BECAS CHILE/2017—72180476. Javier Avendaño-Godoy is grateful to ANID/Scholarship Program/DOCTORADO NACIONAL/2020-21202096. Anna Vallverdú-Queralt thanks to the Ministry of Science, Innovation and Universities for the Ramon y Cajal contract (RYC-2016-19355).
**Institutional Review Board Statement:** Not applicable.
**Informed Consent Statement:** Not applicable.
**Data Availability Statement:** Data is contained within the article and Supplementary Materials.
**Acknowledgments:** The authors would like to thank the CCiT-UB for the mass spectrometry equipment and to Yamil Neira from Viña de Neira, who provided the grape cane samples.
**Conflicts of Interest:** Rosa M. Lamuela-Raventós reports receiving lecture fees from Cerveceros de España and Wine in moderation. She also received lecture fees and travel support from Adventia. The other authors declare no conflicts of interest. The funders had no role in the design of the study; in the collection, analyses, or interpretation of data; in the writing of the manuscript, or in the decision to publish the results.
| doab | 2025-04-07T03:56:59.214118 | 17-11-2022 17:23 | {
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007afdec-bed4-405d-873d-c355ba9add0e.191 | *Article* **Thermo-Responsive Gel Containing Hydroxytyrosol-Chitosan Nanoparticles (Hyt@tgel) Counteracts the Increase of Osteoarthritis Biomarkers in Human Chondrocytes**
**Anna Valentino 1,†, Raffaele Conte 2,†, Ilenia De Luca 1, Francesca Di Cristo 3, Gianfranco Peluso 1,4, Michela Bosetti 5,\* and Anna Calarco 1,\***
**Abstract:** Although osteoarthritis (OA) is a chronic inflammatory degenerative disease affecting millions of people worldwide, the current therapies are limited to palliative care and do not eliminate the necessity of surgical intervention in the most severe cases. Several dietary and nutraceutical factors, such as hydroxytyrosol (Hyt), have demonstrated beneficial effects in the prevention or treatment of OA both in vitro and in animal models. However, the therapeutic application of Hyt is limited due to its poor bioavailability following oral administration. In the present study, a localized drug delivery platform containing a combination of Hyt-loading chitosan nanoparticles (Hyt-NPs) and in situ forming hydrogel have been developed to obtain the benefits of both hydrogels and nanoparticles. This thermosensitive formulation, based on Pluronic F-127 (F-127), hyaluronic acid (HA) and Hyt-NPs (called Hyt@tgel) presents the unique ability to be injected in a minimally invasive way into a target region as a freely flowing solution at room temperature forming a gel at body temperature. The Hyt@tgel system showed reduced oxidative and inflammatory effects in the chondrocyte cellular model as well as a reduction in senescent cells after induction with H2O2. In addition, Hyt@tgel influenced chondrocytes gene expression under pathological state maintaining their metabolic activity and limiting the expression of critical OA-related genes in human chondrocytes treated with stressors promoting OA-like features. Hence, it can be concluded that the formulated hydrogel injection could be proposed for the efficient and sustained Hyt delivery for OA treatment. The next step would be the extraction of "added-value" bioactive polyphenols from by-products of the olive industry, in order to develop a green delivery system able not only to enhance the human wellbeing but also to promote a sustainable environment.
**Keywords:** hydroxytyrosol-chitosan nanoparticles; injectable hydrogel; anti-inflammatory; anti-oxidative; osteoarthritis
| doab | 2025-04-07T03:56:59.214336 | 17-11-2022 17:23 | {
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007afdec-bed4-405d-873d-c355ba9add0e.192 | **1. Introduction**
Osteoarthritis (OA) is a chronic inflammatory degenerative disease affecting millions of people worldwide. OA leads to cartilage deterioration, inflammation of the synovial membrane, and subchondral bone sclerosis due to abnormal bone remodeling caused by an overproduction of enzymes degrading the extracellular matrix [1–3]. This disease considerably reduces the quality of life for patients and is associated with pain, transient morning stiffness, and crepitus felt in a joint on moving it [4–6]. A large body of evidence
**Citation:** Valentino, A.; Conte, R.; De Luca, I.; Di Cristo, F.; Peluso, G.; Bosetti, M.; Calarco, A. Thermo-Responsive Gel Containing Hydroxytyrosol-Chitosan Nanoparticles (Hyt@tgel) Counteracts the Increase of Osteoarthritis Biomarkers in Human Chondrocytes. *Antioxidants* **2022**, *11*, 1210. https://doi.org/10.3390/ antiox11061210
Academic Editors: Li Liang and Hao Cheng
Received: 20 May 2022 Accepted: 19 June 2022 Published: 20 June 2022
**Publisher's Note:** MDPI stays neutral with regard to jurisdictional claims in published maps and institutional affiliations.
**Copyright:** © 2022 by the authors. Licensee MDPI, Basel, Switzerland. This article is an open access article distributed under the terms and conditions of the Creative Commons Attribution (CC BY) license (https:// creativecommons.org/licenses/by/ 4.0/).
supports the involvement of inflammation and reactive oxygen species (ROS) production by chondrocytes in OA cartilage [7–9].
To date, numerous pharmacological and non-pharmacological therapies have been developed for the management of OA. However, the current therapies are limited to palliative care and do not exclude the necessity of surgical intervention [10,11]. Under alternative or adjuvant therapeutic schemes, regular dietary intake of natural functional foods containing polyphenols and other phytochemicals such as fruits, vegetables, whole grains, legumes, and olive oil [12–17] has been associated to a protective role in chronic disease prevention such as OA because of their anti-inflammatory and antioxidant properties [18].
Among all, Hydroxytyrosol (Hyt), a polyphenol found mainly in olive oil and raw olives, exerts strong antioxidant activity (as a potent radical scavenger and metals chelator) and acts as an anti-inflammatory as well as antithrombotic, antitumor, antimicrobial, and neuroprotective agent [19–22].
Fucelli et al. demonstrated the ability of Hyt to reduce inflammatory markers, such as Cyclooxygenase-2 (COX2) and Tumor Necrosis Factor alfa (TNF-α) and reduces oxidative stress on a mouse model of systemic inflammation [23]. Pre-treatment of Balb/c mice with Hyt (40 and 80 mg/Kg b.w.) prevented all lipopolysaccharide-induced effects and decreased oxidative stress. In another study, Cetrullo et al. demonstrated that Hyt inhibits the inflammatory response in vascular endothelial cells, macrophages, and monocytes [24]. Furthermore, Hyt reduces oxidative stress and damage, exerts pro-survival and antiapoptotic actions, and favorably influences the expression of critical OA-related genes in human chondrocytes treated with stressors promoting OA-like features [25].
However, the amount of Hyt obtained through its natural sources' consumption is considerably lower than the recommended daily intake able to exert its claimed healthpromoting properties [26]. Furthermore, the local therapeutic concentration of Hyt following oral administration is limited due to its poor bioavailability and enzyme degradation. Encapsulation of Hyt could be a functional alternative strategy to preserve its the biological activity and to ensure controlled release of the latter increasing the residence time inside the joint. Chitosan biopolymer has been extensively used as a matrix for the encapsulation of a wide range of natural products due to its beneficial properties including biodegradability, biocompatibility, and low cost [27]. Moreover, the ionic gelation method allows to obtain drug-loaded chitosan nanoparticles with a controlled size and satisfactory encapsulation capacity, protecting polyphenols from enzymatic oxidation or degradation. However, polymeric particles present some significant limitations such as initial burst release, escape from the joint's cavity, and in vivo rejection.
To overcome the above-mentioned drawbacks, in the present work, a localized drug delivery platform containing a combination of Hyt-loading chitosan nanoparticles (Hyt-NPs) and in situ forming hydrogel have been developed to derive the benefits of both hydrogels and nanoparticles. This thermo-sensitive formulation, based on Pluronic F-127 (F-127), hyaluronic acid (HA), and Hyt-NPs (called Hyt@tgel) presents the unique ability to be injected in a minimally invasive way into a target region as a freely flowing solution. When the temperature rises near the body temperature of 37 ◦C, hydrogel has in situ sol-to-gel transition accommodating the shape to the geometry of the treated area.
HA, a naturally polysaccharide, represents one of the largest components of the extracellular matrix of articular cartilage and plays an important endogenous role in the protection of articular cartilage decreasing the gene expression of inflammatory cytokines. Moreover, HA degrade ECM enzymes with stimulating in vitro chondrocytes proliferation, and chondrogenesis by directing mesenchymal stromal cells (MSCs) differentiation and increasing type 2 collagen production [28–30]. Although HA represents a conventional treatment in knee OA management, several lines of clinical evidence have questioned the effectiveness of such therapies due to HA prompt in vivo degradation mediated by hyaluronidases and oxidative stress [31,32]. To prolong HA residence time and confer optimized product functionality, Pluronic F-127 (F-127) consisting of hydrophilic poly (ethylene oxide) (PEO) and hydrophobic poly (propylene oxide) (PPO) (PEO-PPO-PEO) was added. Moreover, reported by Young-seok Jung and colleagues [33], the addition of high-molecular-weight HA (Mw: ~1000 kDa) increases the mechanical strength of thermosresponsive hydrogel hindering the interactions between water and poloxamer molecules due to HA-assisted inter-micellar packing. Starting from the results obtained in the work of Young-seok Jung, nanocomposite hydrogel formulations (Hyt@tgels) were optimized to ensure gelation around 37 ◦C, as well as allowing Hyt release exerting antioxidant and antiinflammatory activity on an in vitro induced inflammatory environment mimicking OA.
Based on the above, the developed platform may serve as both a Hyt delivery system and as a tissue engineering scaffold to stimulate the regeneration of a lesioned tissue and to prevent chondrocytes senescence providing an alternative and potentially more effective loco-regional approach to manage OA.
| doab | 2025-04-07T03:56:59.214520 | 17-11-2022 17:23 | {
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007afdec-bed4-405d-873d-c355ba9add0e.193 | **2. Materials and Methods**
#### *2.1. Materials*
Hyt (>98% purity), chitosan medium molecular weight (50,000–190,000 Da, 75–85% deacetylated, viscosity < 200 mPa.s, 1% in acetic acid), lactic acid (DL-Lactic acid, powder), sodium tripolyphosphate (TPP, technical grade), Pluronic F-127, and Fluorescein isothiocyanate (FITC), 3,3 ,5,5 -Tetramethylbenzidine (TMB), Thiobarbitoric acid (TBA), Dichloro-dihydro-fluorescein diacetate (DCFH-DA) and Ultrapure HA in the form of sodium hyaluronate medium molecular weight were purchased from Sigma Aldrich (Milan, Italy) and used as received. All other reagents used in the experiment were of analytical grade and when not indicated were purchased from Sigma Aldrich (Milan, Italy).
#### *2.2. Preparation and Physico-Chemical Characterization of Hyt-Loading Nanoparticles (Hyt-NPs)*
A series of three Hyt-loading nanoparticles (Hyt-NPs) with varying chitosan concentration (0.1%, 0.5%, and 1% *w*/*w*) were obtained, with slight modifications, according to the well-known ionotropic gelation method [34]. Briefly, chitosan solution in 1% (*v*/*v*) lactic acid was prepared and stirred overnight at room temperature. TPP (5 mg/mL) and Hyt (10 mg) were dissolved in double distilled water to achieve different CS:TPP mass ratios. All solutions were filtered using 0.45 μm pore size membrane filters. TPP/Hyt solution was added dropwise into the chitosan solution under magnetic stirring (750 rpm) until a translucent Hyt-NPs suspension was formed. The suspension was stirred for 1 h at 1000 rpm at 25 ◦C to allow complete interaction. Then, the solution containing nanoparticles was ultrasonicated for 5 min at 40 kHz. Finally, Hyt-NPs were collected by cooling centrifugation (Frontiers 5718R, OHAUS, Milan, Italy) at 15,000 rpm for 45 min at 4 ◦C and washed with deionized water. FITC-loaded NPs were obtained by adding hydrophilic fluorescent probe into the TPP aqueous phase instead of Hyt, and the NPs prepared as described previously. Blank NPs were produced as negative control.
Particle Size (hydrodynamic diameter), polydispersity index (PDI), and zeta potential measurements were carried out on freshly prepared samples as reported in Conte et al. [35]. All samples were diluted in deionized water and measured at 25 ◦C using a Malvern Zetasizer (Malvern Instruments Ltd., Malvern, UK). The reported data are an average value of three measurements of the same sample. The particle size was confirmed by NanoSight NS300 Nanoparticles Tracking Analysis (NTA, Malvern Instruments, Amesbury, United Kingdom, UK). The fresh nanoparticle dispersions were centrifuged at 14,000 rpm for 30 min (5718R, OHAUS, Nänikon, Switzerland). The amount of drug entrapped in NPs was determined in triplicate indirectly by analyzing the amount of free Hyt in supernatant. The free Hyt in supernatant was quantified as described in Section 2.4.4 paragraph.
The encapsulation efficiencies of a series of Hyt-loaded nanoparticles were determined based on the following equation:
Encapsulation Efficiency (EE %) <sup>=</sup> Total amount of Hyt loaded <sup>−</sup> Free Hyt in supernatant Total amount of Hyt loaded <sup>×</sup> <sup>100</sup>
| doab | 2025-04-07T03:56:59.214949 | 17-11-2022 17:23 | {
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007afdec-bed4-405d-873d-c355ba9add0e.194 | *2.3. Hyt-Loaded Hydrogel (Hyt@tgel) Preparation*
Injectable hydrogels were prepared according to the cold method as reported by [33]. Briefly, HA (100 mg) and Pluronic F-127 concentrations of 12–25% (*w*/*v*, 5 mL) were mixed in double distilled water at a temperature below 4 ◦C to form hydrogels. The polymer solution was left for at least 24 h to ensure the complete dissolution. Meanwhile, lyophilized Hyt-NPs (1, 5, and 10 mg) were added to the polymer dispersion and stirred for 1 h at 4 ◦C.
#### *2.4. Hyt@tgel Characterization*
| doab | 2025-04-07T03:56:59.215167 | 17-11-2022 17:23 | {
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007afdec-bed4-405d-873d-c355ba9add0e.195 | 2.4.1. Gelation Time and Syringeability
The sol-gel phase transition (gelation time) of Hyt@tgels was determined by modified test tube inversion method [36]. An aliquot (1.0 mL) of each sample was prepared in a glass tube and then placed in a low temperature digital water bath. The solution was heated at the rate of 0.5 ◦C/min and after each minute the glass vial rotated 90◦ to check the gelling of the sample. Tsol-gel was determined as the temperature at which the gel did not exhibit gravitational flow during a period of 2 min when the tube was reversed. Averages and standard deviations of each sample were determined in triplicate.
Syringeability was assayed after injection of Hyt@tgel through a syringe with a 30-gauge needle. The solutions which were easily passed from the syringe were termed as pass and the solutions which were difficult to pass were termed as fail.
| doab | 2025-04-07T03:56:59.215222 | 17-11-2022 17:23 | {
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007afdec-bed4-405d-873d-c355ba9add0e.196 | 2.4.2. Mechanical Strength Test
The mechanical strength of Hyt@tgels was measured in relation to its viscosity with a Brookfield viscometer (RVDV-II + P, Brookfield, WI, USA) set at 200 rpm router speed with increasing temperatures (20–65 ◦C, Equilibrium time: 1 ◦C/2 min (>35 ◦C) or 5 ◦C/10 min (<35 ◦C)).
| doab | 2025-04-07T03:56:59.215307 | 17-11-2022 17:23 | {
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007afdec-bed4-405d-873d-c355ba9add0e.197 | 2.4.3. Short-Term Stability Studies
The physico-chemical stability of Hyt@tgels was determined upon 14-day storage at different temperatures (4.0 ± 0.5 ◦C and 25.0 ± 0.5 ◦C). At predetermined times, aliquots were centrifuged (12,000× *g*, 4 h, 20 ◦C) to separate nanoparticles from the hydrogel. All samples were analyzed for particle size, PDI, and % drug entrapment efficiency, and the results were compared with the initial values. The Hyt stability during storage was confirmed by HPLC analysis as described in Section 2.4.4 paragraph.
| doab | 2025-04-07T03:56:59.215344 | 17-11-2022 17:23 | {
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007afdec-bed4-405d-873d-c355ba9add0e.198 | 2.4.4. In Vitro Hyt Release
The cumulative Hyt release from the hydrogel formulations was determined using the dialysis bag method in phosphate buffer saline (PBS, pH 7.4). The Hyt@tgels formulations (1 mL) were sealed in pre-swollen cellulose membrane dialysis bags (3.5–5.0 kDa cut-off, Spectrum) and immersed into 5 mL of PBS buffer (pH 7.4) in a water bath at 37 ◦C shaken at 100 rpm for 5 days. At set time intervals, 5 mL of the release media was collected for Hyt analysis and replaced with the same volume of fresh PBS to maintain the sink conditions. Hyt released in the PBS media from the hydrogels was measured with liquid chromatography–tandem mass spectrometry (LC-MS/MS) as reported by [37]. The LC-MS/MS system consisted of a Shimadzu NexeraXR UHPLC (Shimadzu Italy, Milan, Italy) coupled to an LCMS 8060 turbo spray ionization triple-quadrupole mass spectrometer (LCMS 8060, Shimadzu Italy, Milan, Italy). The whole system was controlled by Lab Solution software. Separation of analytes was achieved using a 2.6 μm Kinetex polar C18 column (Phenomenex, Torrance, CA, USA). The mobile phase included Buffer A (0.1% formic acid in water) and Buffer B (0.1% formic acid in acetonitrile) in isocratic flow. The total run time was 7 min for each injection. The mass spectrometer was operated in the turbo-spray mode with negative ion detection. The detection and quantification of Hyt was accomplished by multiple reaction monitoring (MRM) with the transitions *m*/*z* 153.05 → 123.0 (quantifier); 153.05 → 93.0 (qualifier). The instrumental parameters tuned to maximize the MRM signals were nebulizing gas flow 3 L/min, heating gas flow 10 L/min, interface temperature 370 ◦C, DL temperature 250 ◦C, heat block temperature 450 ◦C, and drying gas flow 10 L/min.
| doab | 2025-04-07T03:56:59.215394 | 17-11-2022 17:23 | {
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007afdec-bed4-405d-873d-c355ba9add0e.200 | 2.5.1. Cell Culture and Treatment
Human chondrocyte cells line C20A4 was obtained from American Type Culture Collection (ATCC, Manassas, VA, USA). It was maintained at 37 ◦C in a humidified atmosphere containing 5% CO2 in Dulbecco's modified Eagle's Medium/Nutrient Mixture F-12 (DMEM/F12) supplemented with 10% fetal bovine serum (FBS), 1% L-glutamine, 50 U/mL penicillin, 50 mg/mL streptomycin, 50 μg/mL ascorbic acid, and 50 μM α-tocopherol (Euroclone, Milan, Italy). Cells were tested for contamination, including *Mycoplasma*, and used within 2–4 months. All experiments were performed with an 80% confluent monolayer. The protective effects of Hyt were studied with the acute toxicity model by pre-treating cells with Hyt@tgel for 24 and 96 h followed by 24 h H2O2 (230 μM) [38] treatment in the absence of hydroxytyrosol. A shorter exposure (4 h) was used to investigate effects on mRNA expression.
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007afdec-bed4-405d-873d-c355ba9add0e.201 | 2.5.2. Intracellular Oxidative Stress
DCFH-DA assay was used to measure the production of intracellular reactive oxygen species (ROS) in C20A4 cells according to the manufacturer's protocol. Following treatment, cells were labeled with DCFH-DA (25 μM) for 1 h in the dark. The fluorescence was measured every 5 min for 1 h, with an excitation wavelength of 485 nm and an emission wavelength of 535 nm using a microplate reader (Cytation 3).
The malondialdehyde (MDA) concentration, as a lipid peroxidation index, was determined using the thiobarbituric acid reactive substances (TBARS) assay, according to the manufacturer's protocol. The basal concentration of MDA was established adding about 600 μL of TBARS solution to 50 μg of total protein dissolved in 300 μL of Milli-Q water. The mix was incubated for 40 min at 100 ◦C prior to centrifugation at 14,000 rpm for 2 min. The supernatant was analyzed with a microplate reader at a wavelength of 532 nm [39].
Total SOD-like activity was assessed with the SOD Assay Kit-WST according to the manufacturer's protocol. The activity was expressed as units per mg of protein, where one unit of enzyme inhibits reduction of cytochrome C by 50% in a coupled system formed by xanthine and xanthine oxidase.
## 2.5.3. Enzyme-Linked Immunosorbent Assay (ELISA)
Secreted IL-6, IL-8, and TNF-α protein levels were measured in supernatants of chondrocytes treated as reported in paragraph 2.5.1. Briefly, 100 μL of samples and standards were added into the wells already pre-coated with antibody specific for IL-6, IL-8, or TNF-α, and incubated for 2 h at 37 ◦C. Unbound substances were removed and 100 μL of biotinconjugated antibody specific for IL-6, IL-8, or TNF-α was added to the well. After washing, 100 μL of avidin conjugated Horseradish Peroxidase (HRP) was added to the wells and incubated for 1 h at 37 ◦C, followed by addition of 90 μL of TMB substrate solution, and then incubation for 15–30 min at 37 ◦C. Stop solution was added to each well, the plate was gently tapped for thorough mixing, and the color intensity measured at 450 nm using a Cytation 3 Microplate Reader (ASHI, Milan, Italy).
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007afdec-bed4-405d-873d-c355ba9add0e.202 | 2.5.4. Quantitative Senescence-Associated Beta-Galactosidase Assay
4-methylumbelliferyl-β-D-galactopyranoside (4-MUG) was used as substrate of βgalactosidase for the quantitative SA-β-gal assay [40]. 4-MUG does not fluoresce until cleaved by the enzyme to generate the fluorophore 4-methylumbelliferone. The assay was carried out on lysates obtained from cells that were grown as reported above. The production of the fluorophore was monitored at an emission/excitation wavelength of 365/460 nm.
#### 2.5.5. RNA Isolation, Reverse Transcription, and Quantitative Real-Time PCR (qRT-PCR)
Total RNA was extracted from cell cultures using TriFast (EuroClone, Milan, Italy), according to the manufacturer's protocol, and mRNA levels quantified by RT-PCR amplification as reported by Calarco el al. [41]. For retro-transcription, total RNA (0.5 μg) was treated as described in EuroClone standard protocol and amplified by qPCR. Specific primers for SRY-Box Transcription Factor 9 (*SOX9*), Collagen Type II Alpha 1 Chain (*COL2A1*), Aggrecan (*ACAN*), Cartilage Oligomeric Matrix Protein *(COMP*), Interleukin-6 (*IL-6*), Interleukin-8 (*IL-8*), tumor necrosis factor (*TNF*)-*α*, Matrix Metallopeptidase 3 and 13 (*MMP-3* and *13*), and β-Actin (*ACTB*) were used and listed in Table 1. qRT-PCR was run on a 7900 HT fast real-time PCR System (Applied Biosystem, Milan, Italy). The reactions were performed according to the manufacturer's instructions using SYBR Green PCR Master mix (Euroclone, Italy). Data were normalized using the housekeeping gene (ACTB). All reactions were run in triplicate and the results expressed as mean <sup>±</sup> SD. The 2−ΔΔCt method was used to determine the relative quantification.
**Table 1.** Primers used for qRT-PCR.
| doab | 2025-04-07T03:56:59.216103 | 17-11-2022 17:23 | {
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007afdec-bed4-405d-873d-c355ba9add0e.203 | *2.6. Statistical Analysis*
Statistical comparisons between the different experimental groups and controls were made using GraphPad Prism 6 software (GraphPad Software Inc., San Diego, CA, USA). Each experiment was performed at least three times and all quantitative data are expressed as mean ± standard deviation (SD).
| doab | 2025-04-07T03:56:59.216338 | 17-11-2022 17:23 | {
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007afdec-bed4-405d-873d-c355ba9add0e.204 | **3. Results and Discussion**
## *3.1. Preparation and Physicochemical Characterization of Hyt-Loaded Nanoparticles (Hyt NPs)*
OA, the most common musculoskeletal disease in the elderly population, involves the inflammatory immune response at both local (joint site) and systemic levels leading to severe articular joint pain and reduced joint mobility. To date, local anti-inflammatory treatment is usually insufficient because of their short intra-articular half-lives, while systemic administration is associated with more adverse events [42–44]. Chitosan-based nanoparticles have been extensively used as ideal drug carriers for wide range of biomedical applications due to their good compatibility and degradability [45,46].
In this work, Hyt-loaded nanoparticles (Hyt NPs) were successfully produced by the ionic gelation method, using tripolyphosphate (TPP) as the crosslink. Although the ionic gelation process represents a simple and robust route to obtain chitosan NPs in aqueous medium and under mild conditions, the optimal process parameters were determined to achieve NPs with high drug loading and narrow polydispersity index (PDI). Indeed, the ratio of chitosan/TPP, the chitosan concentration, and the concentration of the encapsulated drug could interfere with the NP size and size distribution during NP formation.
Results of polymer ratio on particle size and size distribution, polydispersity index (PDI), zeta potential (ZP), and encapsulation efficiency (EE%) of nine batches of Hyt NPs studied are summarized in Table 2. Particle size of the prepared formulations was in a nanometric range varying between 510.14 ± 13.21 nm (CS:TPP 1:1) and 137.56 ± 3.13 nm (CS:TPP 10:1) demonstrating that the size of the nanoparticles depends greatly on the ratio of CS to TPP. This behavior is achieved by the interaction of the phosphate charged groups of TPP with the –NH3 <sup>+</sup> groups within the CS structure. Indeed, as the amount of TPP increases the particle size decreases because of the increment in the cross-linking of CS macromolecules mediated by TPP, leading to a minimum particle size at 10:1 CS:TPP ratio.
**Table 2.** Effect of chitosan concentration and chitosan/TPP ratio on the size (hydrodynamic diameter), polydispersity index (PDI), zeta potential (ZP), and encapsulation efficiency (EE) of Hyt-loading nanoparticles (Hyt NPs). The Hyt concentration was kept constant at 10 mg.
Note: In the same column, value with the same subscript letter (a–c) were not significantly different (*p* > 0.05). Data were mean of three replications ± standard deviation (SD).
Moreover, all formulations present a narrow size distribution and high positive surface charge indicating their better stability to aggregation due to the repulsive forces exerted by the positive surface charge. As reported in Table 2, the EE of Hyt NPs enhanced with an increase in CS:TPP ratio ranging between 18.31 ± 1.23% of 1:1 and 74.18 ± 3.16% of 10:1. This could likely be attributed to the number of crosslinking units associated with different TPP concentrations [47]. Moreover, the reduction in nanoparticle size obtained with 10:1 CS:TPP ratio resulted in increment of space for drug encapsulation.
According to the above results, the chitosan and TPP ratio of 10:1 was chosen for further study as the obtained Hyt NPs showed the highest EE with acceptable particle size and distribution.
Figure 1 shows representative images of: size (1A), zeta potential (1B) distribution, screenshot of nanoparticles tracking analysis video (NTA, 1C), and measurements (1D) of Hyt-NPs synthetized in the optimal condition.
#### *3.2. In Vitro Hydrogel Formulation (Hyt@tgel) and Hyt Release*
To obtain a sustained and localized drug delivery of Hyt at body temperature, different amounts of Hyt NPs were dispersed into injectable hydrogels composed of 20 wt% of Pluronic F127 and 1 wt% Hyaluronic acid (Hyt@tgel). According to Young-seok et al. [33] the Hyt@tgel formulation was optimized to reduce the Pluronic F-127 concentration needed to obtain gelation at body temperature. Moreover, the hydrophobic interaction between acetyl groups on HA and methyl groups on Pluronic could enhance the mechanical strength of the resulting hydrogel at temperatures above the critical gelation temperature (CGT). As shown in Figure 2A, the addition of different concentrations of Hyt-NPs (1, 5, and 10 mg) did not significantly affect the Hyt@tgel gelation temperature, suggesting that hydrogel structure organization was maintained after nanoparticles dispersion. These results are in agreement with previous studies at the same Pluronic concentrations [48,49]. The gelation time of the Hyt@tgel at 35 ◦C was slightly increased by Hyt NPs incorporation (Figure 2B). In particular, the presence of high nanoparticle concentrations increases the gelation time by 0.5 min (10 mg, Hyt@tgel10) and 0.2 min (5 mg, Hyt@tgel5) with respect to Hyt@tgel alone (10.6 min). Long in vivo gelation time, in fact, can cause nanoparticle loss by diffusion into the surrounding tissue. On the contrary, gelation that occurs too quickly could lead to
clogging of the injection needle resulting in incomplete administration. Based on suitable gelation time and temperature, further analyses were conducted only on the Hyt@tgel10 sample. As shown in Figure 2C, Hyt@tgel10 demonstrated easy injectability through hypodermic needles at room temperature, while when the temperature increases at 35 ◦C, the extrusion needs the application of an extra force due to the increase in the viscosity. When the gel concentration reached the critical gelation concentration, the Hyt@tgel10 passed from an aqueous solution to a gel as the temperature was increased from 4 to 35 ◦C as demonstrated by the inversion test tube (Figure 2D). Hyt@tgel10 exhibited a viscous flowable form at low temperature becoming a semi-solid gel after incubation at temperature higher than 30 ◦C. This behavior was confirmed by the measure of viscosity as a function of temperatures (Figure 2E).
There is a substantial body of evidence that encapsulation enhances the bioactivity of compounds improving their stability in aqueous medium and increasing upon the delivery at the target site. Chen et al. demonstrated the ability of chitosan microspheres dispersed in a thermally responsive chitosan hydrogel to load anti-inflammatory drugs. After injection into the knee joints of OA rabbits, drugs were released for more than 7 days in a controlled manner [50]. According to Wang et al., curcumin-loaded HA/chitosan nanoparticles exhibited a good sustained-release property leading to inflammation and cartilage apoptosis inhibition acting on the NF-κB pathway [51].
**Figure 2.** Characterizations of Hyt@tgel. Gelation temperature (**A**) and gelation time at 37 ◦C (**B**) of different hydrogel compositions (Hyt@tgel, Hyt@tgel1, Hyt@tgel5, and Hyt@tgel10). Representative photographs of the Hyt@tgel10 syringeability (**C**) and inverted test tube (**D**) obtained at 4 and 35 ◦C. Phenol red was added to facilitate hydrogel monitoring. (**E**) Solution viscosity measurement of the Hyt@tgel10 as a function of temperature. (**F**) Cumulative Hyt release from Hyt NPs and Hyt@tgel10 in phosphate buffer saline (PBS) after 24 h (and eight days). Six different experiments were conducted, and the results expressed as the mean of the values obtained (mean ± SD).
Chitosan nanoparticles have been recognized as a useful drug delivery tool in OA for their ability to prolong the drug retention time. To evaluate the sustained release properties of Hyt-NPs, an in vitro drug release study of Hyt from Hyt-NPs and from HYt@tgel10 was carried out using dialysis membrane against phosphate buffer saline (PBS). As shown in Figure 2F, the in vitro release of Hyt by chitosan NPs exhibited a fast drug release rate with 41% of Hyt released within the first hour, with the majority of the release occurring during the initial 2 days (75%). On the contrary, Hyt release rate from Hyt@tgel10 significantly slowed down (*p* < 0.05) with only 10% of Hyt released after 1 h, followed by a prolonged Hyt release up to 1 week. The slow Hyt release from the hydrogel could be attributed to the densely packed inter-micellar structure due to the presence of HA. Moreover, the highly packed super-molecular structure could reduce the diffusion coefficients inside of the hydrogel leading to a prolonged drug release.
Physical stability of nanoparticles in Hyt@tgel was investigated at 4 and 25 ◦C over 14 days by measuring size and PDI. As reported in Table 3, Hyt NPs were stable when stored at both low and room temperatures without a significant increase in particle size and PDI. Moreover, the drug encapsulation efficiency, assessed in parallel, demonstrated no decrease in the Hyt retention rate over the 14-day period confirming the protective effect of chitosan nanoparticles on biomolecules.
**Table 3.** Particle size and entrapment efficiency studies of Hyt@tgel10 before and after 14-day storage.
Note: Data were expressed as mean ± standard deviation, *n* = 3.
Taken together, the physicochemical behavior of Hyt@tgel10 is consistent with a potential use as a device to be injected through a syringe, because the sol-to-gel transition temperature is between room temperature and physiological temperature.
| doab | 2025-04-07T03:56:59.216381 | 17-11-2022 17:23 | {
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007afdec-bed4-405d-873d-c355ba9add0e.205 | *3.3. Oxidative Damage Protection of Hyt@tgel*
Several studies have concluded that OA progression is significantly related to an imbalance between the production of reactive oxygen species (ROS) and their clearance by an antioxidant defense system [52,53]. During OA pathogenesis, chondrocytes become both source and target of elevated amounts of reactive chemical species, particularly oxygen and nitrogen species triggering a vicious circle that leads to further damage of cartilage cells and matrix [54]. A wide body of evidence suggested that Hyt has antioxidant activity by inhibition and/or scavenging of reactive oxygen species (ROS) [54,55]. Moreover, a gene expression profiling study has suggested that Hyt affect the expression of genes involved in oxidative stress, inflammation, cell proliferation, or differentiation, suggesting that the beneficial effects of this molecule may be multifactorial and context-dependent [56]. The efficiency of Hyt@tgel10 to reduce the intracellular ROS generation was assessed in C20A4 cells in the presence of hydrogen peroxide (H2O2) (Figure 3A,B). The stimulation of chondrocytes with H2O2 mimics the in vivo condition observed in OA cartilage inducing the production of cellular and mitochondrial ROS and producing proinflammatory and procatabolic responses [57,58]. A significant increase (*p* < 0.001) in chondrocyte intracellular oxidants by approximately 2.8 times was obtained with respect to untreated cells (Control) after H2O2 24 h treatment (Figure 3A). A short-time pre-incubation (24 h) with Hyt@tgel10 considerably reduced (*p* < 0.01) H2O2-induced ROS production of about 1.4-fold with respect to the H2O2 group. Moreover, the protective effect of released Hyt was greatly enhanced ((*p* < 0.001) by a longer pre-treatment (96 h) resulting in slight fluorescence increase with respect to control cells. MDA, a lipid peroxidation end product, is abundant in synoviocytes from patients with OA. Under oxidative stress, polyunsaturated fatty acids of cellular membrane lipids represent the prime targets of ROS attack. The lipid peroxidation leads to the formation of chemically reactive lipid aldehydes, such as MDA, capable of causing severe damage to nucleic acids and proteins, altering their functions and leading to the loss of both structural and metabolic function of cells [59]. As reported in Figure 3B, treatment of cells with H2O2 increased intracellular lipid peroxidation to 2-fold relative to control (*p* < 0.001). Conversely, the presence of Hyt@tgel10 for 24 h markedly diminished (*p* < 0.01) the MDA level (1.1-fold) compared with H2O2 treated cells, with a marked decrease (*p* < 0.001) after 96 h leading the MDA formation to levels almost similar to control.
**Figure 3.** Antioxidant capacity of Hyt@tgel in H2O2-treated chondrocytes. C20A4 cells were incubated in the presence of hydrogel for 24 and 96 h and then treated with hydrogen peroxide for 24 h. (**A**) ROS release was determined by oxidized H2DCFDA (DCF). (**B**) Malondialdehyde quantity was used as a marker of lipid peroxidation. (**C**) Superoxide dismutase (SOD2) activity measured by assay kit. (**D**) SOD2 mRNA transcription level. Results are expressed as the mean of three independent experiments ± S.D (*n* = 3). \*\* *p* < 0.01, \*\*\* *p* < 0.001 versus untreated cells (control). ### *p* < 0.001 versus H2O2 group.
To prevent an accumulation of ROS-mediated damage, chondrocytes produce a number of antioxidant enzymes including the superoxide dismutases (SOD), catalase, and glutathione peroxidase [60]. The three SOD family members SOD1, SOD2, and SOD3 transform O2 − into hydrogen peroxide (H2O2), limiting the formation of highly aggressive compounds such as ONOO− and OH−. All SOD are expressed at lower levels in OA cartilage compared to normal control cartilage, at both the messenger RNA (mRNA) and protein level. In particular, Ruiz-Romero et al. demonstrated through a proteomics approach, a significant decrease in the major mitochondrial antioxidant protein manganese-superoxide dismutase (SOD2) in the superficial layer of OA cartilage. This SOD2 reduction makes cartilage more susceptible to ROS damage suggesting a central role of mitochondrial redox imbalance in OA pathogenesis [61]. To verify if the antioxidant actions of Hyt have been related not only to its free radical scavenging activity, but also to the ability to enhance the endogenous defense system by inducing antioxidant/detoxifying enzymes activity, SOD2 activity was assayed. As shown in Figure 3C, treatment with H2O2 leads to decrease in antioxidant enzyme activity of about 54% with respect to untreated cells. When chondrocytes were pre-treated with Hyt@tgel10 for 24 and 96 h, SOD2 activity was 18% and 42%, respectively, higher than that in H2O2-treated cells, demonstrating a good ability to protect mitochondria from oxidative damage. Moreover, Hyt@tgel10 pretreatment restored the SOD2 transcript to above their control levels, by significantly increasing its expression by 2.4-fold for 24 h and 5.5-fold after 96 h over the H2O2-depressed level (Figure 3D).
Taken together, the results reported herein confirm a key role of Hyt@tgel10 pretreatment to effectively suppress the production of intracellular ROS and lipid peroxidation and also elevated the activity of antioxidant enzymes such as SOD, limiting oxidative stress-induced damage in the OA in vitro model.
## *3.4. Hyt@tgel Suppresses Inflammatory Response in Chondrocytes*
Increases in the levels of the cytokines in joints plays a central role in the pathogenesis of OA by modulating oxidative stress, cartilage ECM turnover, and chondrocytes apoptosis [62,63]. The current drugs for treating OA are developed primarily to relieve pain and control symptoms, failing to cure the disease [63]. Epidemiologic studies demonstrated the lower incidence of inflammatory chronic disease, such as OA in people of the Mediterranean basin. One of the possible reasons is that Mediterranean people have a high intake of olive and olive oil rich in polyphenolic compounds with antioxidant and anti-inflammatory properties [14,64,65]. During the pathophysiological processes of OA, cytokines, hormone-like proteins, are responsible for the loss of metabolic homeostasis of tissues forming joints by promoting catabolic and destructive processes. Olive-oil-rich extracts inhibit the production of proinflammatory cytokines, including IL-1β, TNF-α, IL-6, and prostaglandin E2 in arthritic joints [66,67]. Richard and colleagues demonstrated a pivotal role of Hyt extracted from olive vegetation water in diminished secretion of cytokines (IL-1 α, IL-1 β, IL-6, IL-12, TNF-α), and chemokines (CXCL10/IP-10, CCL2/MCP-1) in murine macrophages (RAW264.7 cells) stimulated with lipopolysaccharide (LPS) [68]. Another study showed a decrease in the severity of the disease and an overall anti-IL-1β effect after treatment with olive and grape seed extract in animal models of post-traumatic OA [69]. In the present study, secreted IL-6, IL-8, and TNF-α were detected in the supernatant of chondrocytes cell line C20A4 by enzyme-linked immunosorbent assays (ELISA). As expected, incubation of cells for 24 h with Hyt@tgel10 significantly reduces the amount of released cytokines with respect to the control in a time-dependent manner (Figure 4A–C). Consistently, the protective effects of Hyt were confirmed also by RT-qPCR analysis. As reported in Figure 4D–F, the mRNA levels of all tested cytokines (relative to the housekeeping gene) were significantly upregulated (*p* < 0.01) in H2O2-treated cells, compared with the control group. As expected, the H2O2-driven release of IL-6, IL-8, and TNF-α was decreased by Hyt@tgel10 pre-treatment with a 50% reduction in interleukin expression levels with respect to H2O2-treated cells.
**Figure 4.** Hyt@tgel10 inhibits H2O2-induced inflammatory response in chondrocytes. The effect of Hyt on the production of IL-6 (**A**,**D**), IL-8 (**B**,**E**), and TNF-α (**C**,**F**) was measured by ELISA assay (**A**–**C**) and qRT-PCR (**D–F**). C20A4 cells were pre-treated with Hyt@tgel10 for 24 h, then stimulated with H2O2 for 24 h (ELISA assay) or 4 h (qRT-PCR). Results are expressed as the mean of three independent experiments ± S.D (*n* = 3). ### *p* < 0.001 H2O2 vs. CTL, \* *p* < 0.05, \*\* *p* < 0.01, and \*\*\* *p* < 0.005 Hyt@tgel10 vs. H2O2.
| doab | 2025-04-07T03:56:59.216904 | 17-11-2022 17:23 | {
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007afdec-bed4-405d-873d-c355ba9add0e.206 | *3.5. Hyt@tgel Protects against H2O2-Mediated Chondrocyte ECM Degradation*
Once damaged, the cartilage is enabled to repair itself due to its special physiological structure. In the early stages of OA, the production of inflammatory mediators including cytokines and prostaglandins by the cartilage and synovial cells lead to activation of matrix metalloproteinases (MMPs) [70]. Among them, matrix metalloproteinases MMP-3 and MMP-13 can further promote cartilage inflammation, chondrocyte apoptosis, and ROS production, via a positive feedback loop [71,72]. Emerging evidence has shown that MMP13 is considered a significant biomarker to assess OA therapeutic effects and OA progression [73,74]. In this context, bioactive molecules able to suppress these inflammatory mediators or block the involved signaling pathway may help to reduce the OA pathological process [75]. To further corroborate the Hyt@tgel anti-inflammatory action, the expression of catabolic genes such as those coding for MMP-3 and MMP-13 were evaluated (Figure 5).
**Figure 5.** Hyt released by Hyt@tgel10 prevents the expression of OA-related genes in chondrocytes treated with H2O2. (**A**) C20A4 chondrocytes were pre-treated with Hyt@tgel10 for 24 h, then stimulated with H2O2 for 4 h (±SD, *n* = 3, \*\* *p* < 0.01 vs. H2O2 group). (**B**) Beta-galactosidase senescence assay. The graph shows the mean percentage value of senescent cells in every experimental condition (±SD, *n* = 3, \*\*\* *p* < 0.005).
Compared with the untreated group, the mRNA expression of MMP-3 and MMP-13 increased significantly (*p* < 0.01) in H2O2-treated cells, while Hyt@tgel10 pre-treatment was able to reduce about 40% of the increases provoked by H2O2. These results, in line with Facchini et al. [25] corroborated the capacity of Hyt to antagonize the activation of pro-inflammatory pathways like NF-κB even in chondrocytes.
Activation of catabolic enzymes degrades proteoglycan and collagen in the articular cartilage. Moreover, inflammatory states lead to de-differentiation of chondrocytes accompanied by decreased expression of chondrocyte-specific proteins [76]. As reported in Figure 5A, the expression of SOX9 [77] (an early marker of the formation of a cartilage-like tissue matrix), COL2A1, ACAN [78], and COMP (markers of the final stage of chondrogenic differentiation) was significantly rescued by incubation with Hyt@tgel10. These data indicate that Hyt released by Hyt@tgel influenced the ECM balance and gene expression in the chondrocytes under pathological state maintaining their metabolic activity and proliferation in their differentiated phenotype.
Although various cell types are involved in OA pathology, chondrocytes play a major role in OA induction by cellular senescence [79]. It has been shown that chondrocytes have telomere shortening with age. For this reason, chondrocyte senescence, caused by chronic stress in the cells or caused by post-traumatic inflammation, is believed to be closely related to OA [80]. Therefore, the regulation of hypertrophic or senescent chondrocytes using natural phytochemicals known to have a powerful anti-inflammatory and antioxidant activity could be a potential therapeutic target to slow or stop the progression of OA [76]. Data demonstrated that senescence was noticeably reduced in cells treated with Hyt@tgel as detected in an in-situ beta-galactosidase assay (Figure 5B). In particular, Hyt@tgel10 treatment reduced more than two times the percentage of senescent cells compared to untreated chondrocytes.
Two different mechanisms of senescence are suggested in chondrocytes: replicative senescence and stress-induced premature senescence [81,82]. Upregulation of inflammatory cytokines expression induces senescence directly, while downregulation of chondrocyte phenotypic maintenance genes such as SOX9, BMP-2, IGF-1, and ACAN induces senescence indirectly. Thus, the association between aging and/or trauma, reduces the number of healthy and functioning chondrocytes, promoting cartilage degeneration and eventually leads to osteoarthritic pathophysiology [10]. Therefore, the reduction of this cell population lends further credit to hydroxytyrosol ability to preserve chondrocytes from senescence after Hyt@tgel10 treatment.
| doab | 2025-04-07T03:56:59.217387 | 17-11-2022 17:23 | {
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007afdec-bed4-405d-873d-c355ba9add0e.207 | **4. Conclusions**
OA is mainly caused by trauma induced by an external force or cartilage damage ac-cumulated during aging. This study provided new insights into the therapeutic effects of intra-articular injection of Hyt-loaded chitosan nanoparticles embedded into thermosensitive hydrogels (Hyt@tgel10). The hydrogel exhibited a sol-gel transition behavior and a gelation time consistent with its therapeutic application. Moreover, Hyt released from hydrogel was able to protect chondrocytes from ROS damage and to revert the activation of inflammatory factors, limiting, in an in vitro model, the vicious cycle typical of OA progression. Hence, it can be concluded that the formulated hydrogel injection could be proposed for the efficient and sustained Hyt delivery for OA treatment. The next step would be the extraction of "added-value" bioactive polyphenols from by-products of the olive industry, in order to develop a green delivery system able not only to enhance human wellbeing but also to promote a sustainable environment.
**Author Contributions:** Conceptualization, A.V., R.C. and A.C.; investigation, A.V., R.C., F.D.C. and I.D.L.; writing—original draft preparation, A.V. and R.C.; writing—review and editing, M.B. and A.C.; supervision, M.B. and A.C.; funding acquisition, M.B., G.P. and A.C. All authors have read and agreed to the published version of the manuscript.
**Funding:** This work was financially supported by the PON 03 PE\_00110\_1/ptd1\_000410 Titolo: Sviluppo di nanotecnologie Orientate alla Rigenerazione e Ricostruzione tissutale, Implantologia e Sensoristica in Odontoiatria/oculistica (SORRISO); POR Campania FESR 2014\_2020 "Tecnologie abilitanti per la sintesi eco-sostenibile di nuovi materiali per la restaurativa dentale"—ABILTECH; EU funding within the Horizon 2020 Program, under the MSCA-RISE 2016 Project "VAHVISTUS" (Grant 734759); POR 2014–2020 FESR MISE Prog. n.F/200004/01-02/X45: Micro-Poli, Titolo: Micronanodispositivi veicolanti polifenoli isolati da scarti della filiera olivicola come nuovi integratori alimentari; Fondazione Cariplo, "grant no. 2018-1001", Economia Circolare-Ricerca per un Futuro Sostenibile" program, "High added-value bioactive polyphenols recovered from waste of olive oil production" research project.
**Institutional Review Board Statement:** Not applicable.
**Informed Consent Statement:** Not applicable.
**Data Availability Statement:** The data presented in this study are available in the article.
**Acknowledgments:** The authors gratefully acknowledge Orsolina Petillo for her assistance with cell culture (IRET-CNR).
**Conflicts of Interest:** The authors declare no conflict of interest.
| doab | 2025-04-07T03:56:59.217645 | 17-11-2022 17:23 | {
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007afdec-bed4-405d-873d-c355ba9add0e.209 | *Article* **Preparation, Characterization, and Antioxidant Activity of Nanoemulsions Incorporating Lemon Essential Oil**
**Ting Liu 1,2, Zhipeng Gao 3, Weiming Zhong 3, Fuhua Fu 1,2, Gaoyang Li 1,2, Jiajing Guo 1,2,\* and Yang Shan 1,2,\***
**Abstract:** Lemon essential oil (LEO) is a kind of citrus essential oil with antioxidant, anti-inflammatory, and antimicrobial activities, but low water solubility and biological instability hinder its industrial application. In this study, LEO was nanoemulsified to solve these problems. The preparation procedure of lemon essential oil nanoemulsions (LEO-NEs) was optimized, and the physicochemical characterization and antioxidant activities were explored. Single-factor experiments (SFEs) and response surface methodology (RSM) were conducted for the effects on the mean droplet size of LEO-NEs. Five factors of SFE which may influence the droplet size were identified: HLB value, concentration of essential oil, concentration of surfactant, ultrasonic power, and ultrasonic time. On the basis of the SFE, the RSM approach was used to optimize the preparation procedure to obtain LEO-NEs with the smallest droplet size. LEO-NEs exhibited good antioxidant activity when the HLB value was 13, content of surfactant was 0.157 g/mL, ultrasonic time was 23.50 min, and ultrasonic power was 761.65 W. In conclusion, these results can provide a good theoretical basis for the industrial application of lemon essential oil.
**Keywords:** lemon essential oil; nanoemulsions; ultrasonication; response surface methodology; antioxidant activities
| doab | 2025-04-07T03:56:59.217849 | 17-11-2022 17:23 | {
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"title": "Characterization and Encapsulation of Natural Antioxidants: Interaction, Protection and Delivery",
"publisher": "MDPI - Multidisciplinary Digital Publishing Institute",
"isbn": "9783036554563",
"section_idx": 209
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007afdec-bed4-405d-873d-c355ba9add0e.210 | **1. Introduction**
Essential oils (EOs), as volatile products of secondary plant metabolism, are well known for their antioxidant [1,2], anti-inflammatory [3], and antimicrobial [4] activities. Citrus EOs have high yield and demand in EO, which are major by-products of citrus processing. Lemon essential oil (LEO) is a kind of citrus EOs, which is commonly used for flavoring and fragrance. The FDA has also deemed LEO safe for use as a preservative or flavoring agent [5]. Furthermore, some researchers reported that LEO had antioxidant activity using DPPH, ABTs, and β-Carotene bleaching assays [6,7]. The antioxidant activity of the LEO is related to the preservation of food and the prevention of diseases. Thus, it has the prospect to replace synthetic preservatives [8].
However, the greatest impediment to the widespread use of LEO is its insolubility in water, and other disadvantages include volatility, low stability, and sensitivity to the environment. LEO could be encapsulated in emulsions to reduce its hydrophobicity, but conventional emulsions are thermodynamically unstable and the components tend to separate from each other [9]. These problems can be solved by nanoemulsions (NEs) prepared using emerging nanotechnology [10]. A NE is a type of drug delivery system with a simple preparation process and stable formulation quality. It has a certain kinetic
**Citation:** Liu, T.; Gao, Z.; Zhong, W.; Fu, F.; Li, G.; Guo, J.; Shan, Y. Preparation, Characterization, and Antioxidant Activity of Nanoemulsions Incorporating Lemon Essential Oil. *Antioxidants* **2022**, *11*, 650. https:// doi.org/10.3390/antiox11040650
Academic Editors: Li Liang and Hao Cheng
Received: 11 February 2022 Accepted: 25 March 2022 Published: 28 March 2022
**Publisher's Note:** MDPI stays neutral with regard to jurisdictional claims in published maps and institutional affiliations.
**Copyright:** © 2022 by the authors. Licensee MDPI, Basel, Switzerland. This article is an open access article distributed under the terms and conditions of the Creative Commons Attribution (CC BY) license (https:// creativecommons.org/licenses/by/ 4.0/).
and thermodynamic stability [10], which can effectively improve the stability of the drug after emulsification on the one hand, and reduce the irritation of drug delivery on the other [11]. The droplet size of the NE is relatively small (20–200 nm) [12,13]. Meanwhile, the particle size of NEs determines its surface and interface properties. NEs with small particle size have a low particle weight and high surface-to-volume ratio, and the Brownian motion of small particle NEs can overcome gravity, which can reduce the occurrence of coalescence, aggregation, and flocculation [14,15]. However, the small droplets in oil-inwater nanoemulsions are mainly composed of oil and dispersed in water with surfactant, whereby their minimum particle size is limited by the oil [16]. Currently, the methods for preparing NEs are classified into high-energy emulsification and low-energy emulsification methods according to the physicochemical mechanism of droplet rupture. Ultrasound is a widely used high-energy process to prepare NEs. It consumes less surfactant with smaller particles compared to the low-energy method [17]. Meanwhile, it provides better control of the system and has a lower production cost than other high-energy methods (microfluidization, high-pressure homogenization) [18].
Thus, the main purpose of this study was to employ the ultrasonic method for preparing LEO-NEs with small particle size, good stability, and high antioxidant activity. SFEs and RSM were employed to prepare optimized LEO-NEs and investigate the individual effects of the independent variables on the droplet size. The findings can provide a basis for formulating and rationalizing the application of LEO-NEs and lay the foundation for their scale-up production in the cosmetics and the food industries.
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007afdec-bed4-405d-873d-c355ba9add0e.212 | *2.1. Materials and Chemicals*
Lemon was obtained from Sichuan Province. Tween-80 and Span-80 were purchased from Sinopharm Chemical Reagent Co., Ltd. (Shanghai, China). the total antioxidant capacity assay kits with DPPH and ABTS were purchased from Suzhou Comin Biotechnology Co., Ltd. (Suzhou, China). Ultrapure water (MILLI Q) was used in the experiments.
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007afdec-bed4-405d-873d-c355ba9add0e.214 | 2.2.1. Extraction and GC-MS Analysis of Lemon Essential Oil (LEO)
The method of extracting essential oil referred to Guo et al. [4]. LEO was extracted from a mixture of lemon peel and water by steam distillation. Sodium chloride was added in the extraction process, and anhydrous sodium sulfate was added to dry the essential oil after extraction. The determination of LEO components was determined according to procedures reported earlier [19]. LEO was analyzed by GC–MS using an Agilent 7890A GC with a Gerstel MPS autosampler and an Agilent 5975C MSD detector. The carrier gas was helium with a flow rate of 1 mL/min. The temperature was programmed as follows: the initial temperature of 40 ◦C was maintained for 1 min; the temperature was increased to 220 ◦C at a rate of 3 ◦C/min for 25 min; the final temperature of 250 ◦C was reached at a rate of 5 ◦C/min for 10 min. MS conditions were 70 eV EI and an ion source temperature of 230 ◦C. The mass-to-charge (*m*/*z*) range was set to 35–350 atomic units. The National Institute of Standards and Technology (NIST 08) was used to compare the data of the LEO components.
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007afdec-bed4-405d-873d-c355ba9add0e.215 | 2.2.2. Preparation of Lemon Oil-Based Nanoemulsions (LEO-NEs)
The LEO-NEs were formed from LEO, a mixture of two surfactants (Tween-80 and Span-80), and deionized water. A procedure for the oil–water mixtures was followed to obtain 20 mL; the pre-emulsion was centrifuged for 5 min at 10,000 rpm using a high-speed homogenizer (F6/10, Jingxin, Shanghai, China). The homogenate was processed further by an ultrasonicator (JY92-11D, Jingxin, Shanghai, China). During the ultrasonication process, samples were put in ice water for a low-temperature environment.
| doab | 2025-04-07T03:56:59.218518 | 17-11-2022 17:23 | {
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007afdec-bed4-405d-873d-c355ba9add0e.216 | *2.3. Optimization and Statistical Design of LEO-NEs*
#### 2.3.1. Single-Factor Experiments (SFE)
Single-factor experiments were designed to investigate the effects of hydrophilic–lipophilic balance (HLB) value, content of Span-80 and Tween-80 (STmix), concentration of essential oil, ultrasonic time, and ultrasonic power on the mean droplet size, which can also provide a reasonable data range for the design of the response surface methodology. Specific parameters are presented in Table 1. The HLB value represented the combined affinity of hydrophilic and oleophilic groups in emulsifier molecules for oil or water [20]. Different HLB values of surfactants can contribute to the formation of two types of emulsions: water-in-oil (W/O) emulsion and oil-in-water (O/W) emulsion. To prepare the O/W LEO-NEs with hydrophilicity, an oil-in-water emulsifier with a high HLB value (8–15) was chosen. According to Nirmal et al. [21], different combinations of STmix were used to create surfactant HLB values ranging from 8–15, as shown in Table 2.
**Table 1.** Variables of single-factor experiments (SFE) and Response surface methodology (RSM).
**Table 2.** Different combinations of Span 80 and Tween 80 used to create surfactant HLB value.
#### 2.3.2. Response Surface Methodology (RSM) Design
The levels of the independent variables to be used in the Box–Behnken designs were determined by the results of the SFE. The RSM explored the effects of the selection factor over 29 runs. In this work, the BBD with four variables (factor A was the HLB value, factor B was the STmix content, factor C was the ultrasonic time, and factor D was the ultrasonic power) at three levels (−1, 0, 1) was carried out to evaluate the effect on the dependent variable. The mean droplet size (Y) was the response value. The optimum formulation was chosen by the analysis of the RSM. The specific parameters are shown in Table 1.
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007afdec-bed4-405d-873d-c355ba9add0e.218 | 2.4.1. Mean Droplet Size and Polydispersity Index (PDI) of LEO-NEs
The mean droplet size, particle size distribution, and PDI were measured using an NS-90 nano-granularity analyzer (Malvern Instruments Ltd., Malvern, UK). The average diameter of the particles indicated the average particle size. The intensity of particles of different diameters indicated the particle size distribution. To avoid bubbles and multiple light scattering, the LEO-NE was diluted 50-fold with ultrapure water. Three sets of measurements were performed in each sample to determine the mean droplet size and PDI of LEO-NEs in 1 mL of the diluted samples.
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007afdec-bed4-405d-873d-c355ba9add0e.219 | 2.4.2. Transmission Electron Microscopy (TEM) Images of LEO-NEs
The particle morphology of the LEO-NEs with a 20-fold dilution was observed by TEM (Hitachi HT-7700, Tokyo, Japan). Dilution was undertaken to prevent interparticle aggregation.
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007afdec-bed4-405d-873d-c355ba9add0e.220 | 2.4.3. DPPH Radical-Scavenging Activity
The DPPH scavenging assay using 0.5 g/mL of LEO and LEO-NEs (stored for 7 days) was measured following the kit instructions. The EOs were diluted with extraction buffer in the kit. Firstly, 380 μL of Reagent 1 was added to 20 μL of sample and then shaken vigorously for 20 min. The change in absorbance was measured at 515 nm by the microplate reader (Thermo Scientific, Waltham, MA, USA). The percentage inhibition free radical scavenging rate of DPPH was calculated as follows:
DPPH scavenging activity (Inhibition%) = [(Acontrol − Asample)/Acontrol] × 100 (1)
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007afdec-bed4-405d-873d-c355ba9add0e.221 | 2.4.4. ABTs Radical-Scavenging Activity
The ABTs scavenging assay of 0.5 g/mL of LEO and LEO-NEs (stored for 7 days) was performed following the kit instructions. The change in absorbance was measured at 734 nm by the microplate reader. The percentage inhibition of ABTs was calculated with the following formula:
ABTs scavenging activity (Inhibition%) = [(Acontrol − Asample + Ablank)/Acontrol] × 100 (2)
## *2.5. Data Analysis*
The results of the single-factor experiments were analyzed by Graphpad Prism version 8 software. The statistical analysis of the results of the response surface test was performed by Design-Expert version 13 software. All of the components, as well as their probable interactions, were examined using statistical parameters for analyses of variance (ANOVAs), such as degrees of freedom, F-ratios, and *p*-values. The model with a good fit to the data was selected (*p* < 0.05).
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007afdec-bed4-405d-873d-c355ba9add0e.222 | **3. Results**
#### *3.1. Chemical Composition of the Lemon Essential Oil*
A lemon-like odor liquid oil isolated by steam distillation from lemon peels was transparent and colorless. The components of the LEO identified are given in Table 3. Analysis of the volatile constituents of the LEO compounds by GC–MS identified 15 compounds that accounted for more than 0.5%, with a total of 96.36%. The major components detected in LEO were limonene (48.54%), α-pinene (30.90%), β-citral (3.65%), and β-myrcene (3.01%). As seen, the main constituents of the EO in this study were composed of monoterpene hydrocarbons (83.53%), including limonene, α-pinene, β-myrcene, and terpinolene.
**Table 3.** Chemical composition (%) of the essential oil isolated from lemon peels.
### *3.2. Single-Factor Experiments*
The effects of parameters on the mean droplet size of LEO-NEs were investigated using singlefactor experiments, including the HLB value (8, 9, 10, 11, 12, 13, 14 and 15), concentration of essential oil (0.05, 0.06, 0.07, 0.08, 0.09 and 1 g/mL), concentration of surfactant (0.0125, 0.025, 0.05, 0.1 and 0.2 g/mL), ultrasonic power (100, 300, 500, 700 and 900 W) and ultrasonic time (0, 10, 20, 30 and 40 min). The ranges for parameter values of RSM were set to the right and left of the optimum values.
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007afdec-bed4-405d-873d-c355ba9add0e.223 | 3.2.1. Effect of HLB Value on the Mean Droplet Size of LEO-NEs
The HLB value of the surfactant can assist in identifying the best-suited stabilizer. When the HLB value of the STmix couple matches the HLB value required for the EO to form nanoemulsions, NEs with a small droplet size can be produced [22]. It was a crucial step to select an appropriate HLB value to obtain LEO-NEs with the smallest particle size. In the present work, the impact of the HLB value on the mean droplet size of LEO-NEs was studied first. As indicated in Figure 1a, when the HLB value changed from 8–12, the mean droplet size progressively declined, while the mean droplet size exhibited an upward trend when the HLB value was above 12. Furthermore, the particle size of LEO-NEs grew considerably when the HLB value increased from 14 to 15. Therefore, the optimum HLB value for the smallest droplet of LEO-NEs was 12.
**Figure 1.** Effects of HLB value (**a**), essential oil concentration (**b**), Surfactant concentration (**c**), ultrasound time (**d**) and ultrasonic power (**e**) on the mean droplet size of NEO-NEs.
## 3.2.2. Effect of Essential Oil Concentration on the Mean Droplet Size of LEO-NEs
To explore the effect of essential oil concentration on the mean droplet size of LEO-NEs, the formulation was performed with different LEO concentrations (ranging from 0.05 to 0.1 g/mL). As shown in Figure 1b, a significant increase in the mean droplet size was observed when the LEO content was changed from 0.05 g/mL to 0.1 g/mL. The nanoemulsion with a low concentration of LEO was more suitable for production applications. According to our results, the essential oil concentration of 0.05 g/mL in LEO-NEs was finally chosen for the subsequent experiments.
#### 3.2.3. Effect of Surfactant Concentration on the Mean Droplet Size of LEO-NEs
STmix with different concentrations was used in the NEs system. As shown in Figure 1c, a sharp decrease in the mean droplet size from 133.71 to 75.66 nm was observed when STmix concentration increased from 0.0125 to 0.1 g/mL. On the other hand, it remained almost constant when increasing the surfactant concentration from 0.1 to 0.2 g/mL. Therefore, the 0.1 g/mL surfactant concentration was selected for subsequent experiments.
#### 3.2.4. Effect of Ultrasonic Time on the Mean Droplet Size of LEO-NEs
Various ultrasonic time was used to prepare the NEs, with the aim of investigating the effects on the mean droplet size of LEO-NEs. As shown in Figure 1d, when the ultrasonic time was 0, which means that the emulsion was not treated by ultrasound, the mean droplet size fluctuated over a wide range, and the repeatability of the experiment was poor. Meanwhile, a layering phenomenon was observed after staying still at room temperature overnight. The smallest particle size was observed at the ultrasonic time of 20 min. The increase in ultrasonic time can promote the integration of water and oil. Longer ultrasonic times, on the other hand, may result in higher degradation or disintegration of bioactive chemicals in LEO, as well as energy waste [23]. Therefore, ultrasonic time of 20 min was selected for the subsequent studies considering both saving energy and achieving the best results.
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007afdec-bed4-405d-873d-c355ba9add0e.224 | 3.2.5. Effect of Ultrasonic Power on the Mean Droplet Size of LEO-NEs
To study the effects of ultrasonic power on the mean droplet size of LEO-NEs, the preparation process was carried out with different ultrasonic powers ranging from 100 to 900 W. As shown in Figure 1e, the value of droplet size decreased with the increase in ultrasonic power from 100 to 700 W. However, the particle size increased instead when the ultrasonic power was increased from 700 to 900 W. Excessive ultrasonic power may induce a rise in the number of bubbles in solvents during cavitation, lowering the efficiency of the ultrasound energy delivered into the medium [24]. As a result, the ultrasonic power of 700 W was chosen for further experiments.
#### *3.3. Response Surface Optimization of LEO-NEs*
The preparation process was further optimized using BBD experiments to obtain the best experimental parameters. The BBD of the response surface was used to optimize the formulation and preparation of LEO-NEs. The following regression equation model was obtained by regression analysis:
As shown in Table 4, the generation of a model with no significant lack of fit implied it suitability. The *R2* > 90% indicated that the model could accurately reflect the change in the response value when the fitness was high. The coefficient of variance (CV), which is the ratio of the estimated standard error to the mean of the observed responses, is related to the model reproducibility. Our model had a CV of 9.97% (<10%), which is usually considered to be sufficiently reproducible. The ratio of adequate precision reflects the ratio of response to the deviation, and its value was 11.840 (>4), indicating an adequate signal.
An analysis of variance of the regression coefficient revealed that C was extremely significantly different (*p* < 0.01), while A significantly differed in its linear effect (*p* < 0.05), remaining factors indicated a non-significant difference. The main effect relationship of each factor could be ranked as ultrasonic time > HLB value > ultrasonic power > surfactant content. Among interaction effects, B2 had extremely significant differences (*p* < 0.01), while BC and C<sup>2</sup> significantly differed (*p* < 0.05). The remaining effects were not significant (*p* > 0.1). The experimental values of mean droplet size of nanoemulsions are presented in Table 5.
**Table 4.** ANOVA of RSM outcome <sup>α</sup>.
<sup>α</sup> *R2* = 0.97; adj. *R<sup>2</sup>* = 0.85; C.V. (%) = 9.97; adequate precision = 11.84.
**Table 5.** Experimental values of mean droplet size of nanoemulsions obtained from BBD experimental design.
Finally, the response surface was plotted using Design-expert 13. The effect of the two-factor interaction on the size of the mean droplet is intuitively shown in Figure 2. The response surface slope was steeper in Figure 2d, indicating that the interplay of surfactant content and ultrasound time had a bigger impact on LEO-NE particle size. The gradient of the response surface was moderate, as shown in Figure 2c, showing that the interaction of HLB value and ultrasonic power had less of an effect on the droplet size.
#### *3.4. Physicochemical Properties and Stability of LEO-NEs*
The above regression model was used to generate the optimum process parameters and validation results. When the HLB value (A) was 13, surfactant content (B) was 0.157 g/mL, ultrasonic time (C) was 23.50 min, and ultrasonic power (D) was 761.65 W, the predicted minimum mean droplet size was 66.82 nm. The actual mean particle diameter was 64.60 nm, and the PDI was 0.255. The
particle size distribution of NEO-NEs is shown in Figure 3a. Then, the morphological changes and the changes in particle size of NEs during the storage period were observed, and the difference in antioxidant activity between emulsion and essential oil was compared.
**Figure 2.** Response surface plot showing the significant (*p* < 0.05) interaction effect for mean droplet size as a function of (**a**) HLB value of STmix and STmix content, (**b**) HLB value of STmix and ultrasonic time, (**c**) HLB value of STmix and ultrasonic power, (**d**) STmix content and ultrasonic time, (**e**) STmix content and ultrasonic power, and (**f**) ultrasonic time and ultrasonic power.
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007afdec-bed4-405d-873d-c355ba9add0e.225 | 3.4.1. Morphological Observation of LEO-NEs
TEM was used to describe the morphology of the LEO-NEs, as shown in Figure 3a, the droplets were well distributed and spherical. However, the diameters of the particles in the NEs were not the same, ranging from 50–100 nm, in line with those reported by the particle size meter.
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007afdec-bed4-405d-873d-c355ba9add0e.226 | 3.4.2. Changes in Particle Size of LEO-NEs during the Storage Period
Figure 3b demonstrates the changes in the mean droplet of LEO-NEs for 1 week at a storage temperature of 25 ◦C. The particle size of the LEO-NEs changed little during a week, ranging from 62.96 nm to 64.60 nm.
**Figure 3.** Physicochemical Properties and Stability of LEO-NEs: (**a**) particle size distribution and transmission electron microscopy (TEM) of droplets of LEO-NEs; (**b**) effect of storage time on particle size of LEO-NEs; (**c**) antioxidant activity comparison between LEO-NEs and LEO.
#### 3.4.3. Antioxidant Activity of LEO-NEs
The DPPH and ABTs assays were used to measure the free-radical-scavenging potential to compare the difference between LEO-NEs and EO at the same concentration (0.05 g/mL). It has been shown that EOs exhibit antioxidant activities due to large amounts of polyphenol compounds. Figure 3c shows that the antioxidant activity of the LEO-NEs was higher than that of LEO with the same concentration, whereby the inhibition of DPPH free radicals by LEO-NEs (57.61%) was much better than that by LEO (8.74%), but the inhibition of ABTs free radicals by LEO-NEs (31.74%) was similar to that by LEO (30.61%).
#### **4. Discussion**
Coinciding with the reports by other Hirai et al. [25], Aguilar et al. [26], Perdones et al. [27] and Campolo et al. [28], limonene was the most abundant component in LEO while its content may vary. LEO was rich in constituents with monoterpene structure (limonene, α-pinene, etc.) which have been proven to possess antioxidant activity [29]. For example, limonene was shown to prevent neuronal suffering [30], oxidative stress on lymphocytes, and mitochondrial dysfunction [29] through its antioxidant activity. In addition, LEO components present in other studies were not detected in this experiment such as β-phellandrene [25], camphene, and sabinene [26]. Differences in LEO composition may be due to differences in geographic location, environmental factors, plant age, developmental stage, harvest time, extraction site, and extraction method [31].
In this study, LEO-NEs were prepared by the ultrasonic method using STmix as an emulsifier. The influence of each factor was studied by SFEs. Firstly, when the HLB values of STmix ranged from 8–14, the mean droplet size of NEs was less than 200 nm, while NEs could not be formed when HLB was 15. The large range of suitable HLB values means that many kinds of surfactants can be used to prepare LEO-NEs. Tween-80 was not suitable for this experiment; however, it was also used to make lemon LEO-NEs in other experiments. Mossa et al. [13] reported that the droplet size of LEO-NEs was 131.9 nm, while the particle size of the LEO-NEs was 181.5 nm in the study of Yazgan [32]. Although these studies were able to form NEs with Tween-80, the particle sizes were larger than 100
nm. Furthermore, the droplet diameter of LEO-NEs was 91 nm [33] and 135 nm [34] when produced with Tween-80 using the high-pressure homogenizer method. These results indicate that different essential oil components and different emulsification methods may lead to different particle sizes when constructing NEs.
When the concentration of essential oil was 0.05–0.1 g/mL, the particle size increases with the increase in concentration of essential oil, indicating its greater impact on particle size. However, previous studies showed that the particle size would not increase when the essential oil exceeds a certain amount if the concentration of surfactant micelles remains not changed, because of a saturation with lemon oil, whereby any further lemon oil droplets added to the nanoemulsions would not dissolve [35]. This phenomenon did not occur in our experiments because the concentration range of LEO was not large enough. An increase in surfactant concentration can also lead to a decrease in particle size. The surfactants can affect inter-particle interactions in emulsions, whereby a the higher surfactant concentration results in weaker inter-particle interactions and smaller droplets formed [36]. The effect of surfactant concentration on mean particle size may be related to the surfactant dose required to cover the surface of the formed droplets, whereby self-emulsification would be more dependent on surfactant concentration [37]. In the process of ultrasonic preparation of NEs, the particle size did not decrease when the surfactant concentration increased to a certain amount. In addition, the dependence of the mean particle size on surfactant concentration did not depend strongly on storage time and temperature [12].
Ultrasonic cavitation is a feasible and energy-efficient method for preparing NEs, which offers improvements in terms of stability and decreases the Ostwald ripening rate. During the ultrasonication processes, soundwave energy causes cavities and sinusoidal pressure variations in the liquid–liquid interphase, resulting in a shockwave action on the particle surface and a reduction in particle size [38]. The particle size of nanoemulsions prepared with the ultrasonic method is generally determined by the sonication time and sonication power, but is insensitive to ultrasonication amplitude [39]. Understanding the dynamic routes is critical for reducing processing time and avoiding energy oversupply. When the ultrasonic time reached a certain value, the particle size reached the minimum. Increasing the ultrasonic time would not lead to a significant change in particle size. The increase in ultrasonic power led to a decrease and then increase in particle size, coinciding with the report of Kentish et al. [40]. In addition, Floris et al. [41] reported that high ultrasonic power may destroy bioactive substances.
The small size of the particles in NEs would result in less agglomeration or precipitation and higher stability of the system [42,43]. RSM was used to optimize NEs to obtain the smallest droplet size. The interaction between surfactant concentration and ultrasonic time had the greatest effect on particle size. However, the particle size did not decrease indefinitely, as it was limited by the ingredients of the essential oil. The optimal preparation conditions obtained by RSM were similar to those obtained by SFE, and the conditions predicted by RSM were relatively more precise.
Due to the mass transfer of oil molecules, droplets in NEs change from smaller droplets to larger droplets through an intermediate water phase, which is called Ostwald ripening. Ostwald ripening leads to droplet growth and phase separation [44]. From the TEM image and the particle size change during storage, particle diameter does not exceed 200 nm; hence, the ripening phenomenon was not serious in LEO-NEs. However, the TEM images revealed that the diameters of the particles in the nanoemulsion were not the same. The TEM image was similar to that presented by Kaur et al. [45] and Zhong et al. [46]. In previous studies, the structure of NE was presented a spherical substance consisting of several small spherical packets [46]. The particle size of LEO-NEs had the tendency to decrease in 1 week, possibly due to the EOs in the NEs undergoing a small amount of evaporation, thereby reducing the content of essential oil. In the study of Zhong et al. [46], there was a tendency for the particle size to increase with storage time, which may have been due to Ostwald ripening.
The prepared NE was not only stable but also had sustained-release activities. The study of antioxidant activities is essential as reflected in the reduction in reactive oxygen species (ROS) in the food and cosmetics industries. We could find that the encapsulation of essential oils in NEs helped to enhance their antioxidant activities when comparing the antioxidant activity of essential oils and NEs. DPPH scavenging activity refers to the ability to reduce the stable DPPH free radical to its reduced form DPPH-H [47]. ABTs scavenging activity refers to the ability to decolorize the radical cation (ABTS•+) [48]. Due to the different principles of determination, the two results are not necessarily related. The different methods employed to indicate antioxidant activity can comprehensively profile the antioxidant activities of LEO-NEs. According to a previous study [6], the DPPH radical scavenging activity and ABTs radical-scavenging activity of pure LEO were 32.85% and 41.57% respectively. These results are close to the antioxidant capacity of LEO-NEs in our study, but the composition and
determination method of the essential oil had an impact on the results. In addition, the antioxidant activity of LEO and LEO-NEs may be due to components in LEO with antioxidant activity. However, The antioxidant activity of LEO should not only consider the primary constituents [49]. The main antioxidant components in lemon essential oil need to be further studied.
| doab | 2025-04-07T03:56:59.219517 | 17-11-2022 17:23 | {
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007afdec-bed4-405d-873d-c355ba9add0e.227 | **5. Conclusions**
This study explored the optimum preparation procedure of LEO-NEs using SFEs and RSM. The optimal parameters were as follows: HLB value of 13, surfactant content of 0.157 g/mL, ultrasonic time of 23.50 min, and ultrasonic power of 761.65 W. The optimized mean droplet size was 64.60 nm. In addition, the TEM images and storage results demonstrated the good dispersion and stability of LEO-NEs. The antioxidant activity experiments showed that LEO-NEs had better antioxidant capacity than essential oils. Some of the characteristics of LEO-NEs investigated in this study and future endeavors may lay the foundation for the practical application of antioxidant activity and other biological activities of LEO-NEs.
**Author Contributions:** Conceptualization, J.G.; methodology, T.L.; software, T.L. andW.Z.; writing—review and editing, T.L., Y.S. and J.G.; visualization, T.L., W.Z. and Z.G.; supervision, G.L., F.F. and Y.S.; project administration and funding acquisition, Y.S. All authors contributed to the article and approved the submitted version. All authors have read and agreed to the published version of the manuscript.
**Funding:** This work was financially supported by the Agricultural Science and Technology Innovation Project of Hunan Province, China, (2021CX05, the Changsha Municipal Natural Science Foundation (kq2202332, kq2014070), the Agricultural Science and Technology Innovation Fund of Hunan (2020CX47), the Key Laboratory of Agro-Products Processing, Ministry of Agriculture and Rural Affairs of China (S2021KFKT-22), the National Natural Science Foundation of China (32073020), and the Hunan innovative province construction project (2019NK2041).
**Institutional Review Board Statement:** Not applicable.
**Informed Consent Statement:** Not applicable.
**Data Availability Statement:** Data is contained within the article.
**Conflicts of Interest:** The authors declare no conflict of interest.
| doab | 2025-04-07T03:56:59.220153 | 17-11-2022 17:23 | {
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007afdec-bed4-405d-873d-c355ba9add0e.229 | *Article* **Engineering of Liposome Structure to Enhance Physicochemical Properties of** *Spirulina plantensis* **Protein Hydrolysate: Stability during Spray-Drying**
**Maryam Mohammadi 1,2,\*, Hamed Hamishehkar 3, Marjan Ghorbani 4, Rahim Shahvalizadeh 1,2, Mirian Pateiro 5,\* and José M. Lorenzo 5,6**
- [email protected]
**Abstract:** Encapsulating hydrolysates in liposomes can be an effective way to improve their stability and bioactivity. In this study, *Spirulina* hydrolysate was successfully encapsulated into nanoliposomes composed of different stabilizers (cholesterol or γ-oryzanol), and the synthesized liposomes were finally coated with chitosan biopolymer. The synthesized formulations were fully characterized and their antioxidant activity evaluated using different methods. Then, stabilization of coated nanoliposomes (chitosomes) by spray-drying within the maltodextrin matrix was investigated. A small mean diameter and homogeneous size distribution with high encapsulation efficiency were found in all the formulations, while liposomes stabilized with γ-oryzanol and coated with chitosan showed the highest physical stability over time and preserved approximately 90% of their initial antioxidant capacity. Spray-dried powder could preserve all characteristics of peptide-loaded chitosomes. Thus, spray-dried hydrolysate-containing chitosomes could be considered as a functional food ingredient for the human diet.
**Keywords:** *Spirulina platensis*; bioactive peptides; encapsulation; liposomes; chitosome
| doab | 2025-04-07T03:56:59.220308 | 17-11-2022 17:23 | {
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007afdec-bed4-405d-873d-c355ba9add0e.230 | **1. Introduction**
The microalga *Spirulina* has gained more attention in areas such as the pharmaceutical, food, poultry, and aquaculture industries for its nutritional and health benefits [1]. Certain therapeutic effects of *Spirulina* (reduced hyperlipidemia, obesity, and blood cholesterol; antioxidant and anticancer activity; immune system improvement; and increased beneficial intestinal bacteria) have been proven by pre-clinical and clinical studies, which are related to their bioactive constitution, e.g., phycocyanins, carotenoids, phenolic compounds, and polyunsaturated fatty acids. The green-blue microalgae are a rich source of proteins (60–70% of dry matter) and, due to the absence of cellulose in the cell wall, are very digestible. Thus, they have gained more attention in recent years as a food supplement, especially for athletes and vegetarians [2].
However, the undesirable taste, low digestibility, and high allergenicity of algae-based protein isolates for monogastric animals and humans have limited their application in the food industry. Protein hydrolysates and peptides can be excellent alternatives to overcome the problems associated with the direct consumption of protein isolates, as they have
**Citation:** Mohammadi, M.; Hamishehkar, H.; Ghorbani, M.; Shahvalizadeh, R.; Pateiro, M.; Lorenzo, J.M. Engineering of Liposome Structure to Enhance Physicochemical Properties of *Spirulina plantensis* Protein Hydrolysate: Stability during Spray-Drying. *Antioxidants* **2021**, *10*, 1953. https://doi.org/10.3390/ antiox10121953
Academic Editors: Li Liang and Hao Cheng
Received: 18 October 2021 Accepted: 2 December 2021 Published: 6 December 2021
**Publisher's Note:** MDPI stays neutral with regard to jurisdictional claims in published maps and institutional affiliations.
**Copyright:** © 2021 by the authors. Licensee MDPI, Basel, Switzerland. This article is an open access article distributed under the terms and conditions of the Creative Commons Attribution (CC BY) license (https:// creativecommons.org/licenses/by/ 4.0/).
nutritional and health-promoting features and act as natural antioxidant agents in food preservation [3,4]. Although algae-derived peptides have various advantages, their instability during storage and under harsh conditions (e.g., in the gastrointestinal tract), low absorption efficiency, bitterness, reaction with the food matrix, and possible inactivation inhibit the application of hydrolysates in foods and beverages. Incorporating these bioactive compounds in lipid-based nanocarriers such as liposomes can be an appropriate solution to cover all of these problems and increase their efficacy under different conditions [5].
Liposomes have an enclosed vesicular structure and are able to accommodate both water-soluble and hydrophobic compounds in their internal aqueous core and bilayer space, respectively [6]. Moreover, the typical constituents of liposomes are completely natural, and their nontoxicity has led to the broad application of these vesicular systems in the encapsulation of various bioactive compounds [7,8]. However, the major drawbacks of this versatile carrier are the fluidity of the intravesicular space and the flexible bilayer structure, which can lead to the physical instability of vesicles (aggregation/flocculation and fusion/coalescence), resulting in changes in size and loss of liposome-incorporated bioactive materials over time. A possible solution to this problem is to engineer the liposomal structure [9]. Mostly, cholesterol has been applied as a stabilizing factor in vesicular systems because it can increase the packing of phospholipid molecules, reduce the fluidity of intravesicular space, and consequently create a more rigid and stable structure over time and under severe shear stress [10]. Moreover, the above-mentioned problems can be improved by depositing an oppositely charged biopolymer such as chitosan around the liposome surface through electrostatic interaction [11,12]. Several studies on improving hydrolysate stability during storage and processing using chitosan coating have been conducted [13,14].
To make a formulation that is more stable over time and more appropriate for industrial application, it can be transformed into powder form by spray-drying or freeze-drying. Freeze-drying is a more expensive technology and requires more time and energy compared with spray-drying. Thus, spray-drying technology is an economical strategy to make powdered liposomal dispersions, and it has a wide range of use in the food industry compared to other drying techniques. However, there have been no studies regarding the simultaneous use of different stabilizers and coating materials to increase the stability and bioactivity of liposome-containing *Spirulina* hydrolysates. Therefore, the aims of this research were as follows: (1) to explore the effect of cholesterol and γ-oryzanol as stabilizing agents and chitosan as a coating material on the mean diameter and encapsulation efficiency of *Spirulina* hydrolysate-loaded liposome, (2) to examine the antioxidant capacity of synthesized liposomes using the different methods, and (3) to estimate the stability of synthesized liposomes during storage. Finally, the optimum formulation for easy usage was converted into powder form and its physicochemical and structural properties and antioxidant activity were evaluated.
| doab | 2025-04-07T03:56:59.220420 | 17-11-2022 17:23 | {
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007afdec-bed4-405d-873d-c355ba9add0e.232 | *2.1. Materials and Reagents*
Pepsin from powdered porcine gastric mucosa (activity ≥ 250 units/mg solid), 1,1 diphenyl-2-picrylhydrazyl (DPPH), 2,2 -azino-bis(3-ethylbenzothiazoline-6-sulfonic acid) diammonium salt (ABTS), 2,4,6(tripyridyl)-1,3,5-triazine (TPTZ), cholesterol, and Coomassie brilliant blue (G250) were purchased from Sigma-Aldrich (St. Louis, MO, USA); potassium persulfate, iron (III) chloride hexahydrate, trichloroacetic acid (TCA), ferrous chloride, γoryzanol, iron sulfate, and maltodextrin were obtained from Merck (Darmstadt, Germany). *Spirulina platensis* powder was purchased from Noor Daro Gonbad (Gonbad Kavous, Iran).
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007afdec-bed4-405d-873d-c355ba9add0e.233 | *2.2. Protein Hydrolysis*
For protein hydrolysis, lyophilized *Spirulina platensis* protein was dispersed in distilled water (DW) to achieve a protein concentration of 3% (*w*/*v*). Protein hydrolysates were produced by pepsin protease for 240 min. The hydrolysis conditions were set as follows: pH 2, temperature 37 ◦C, enzyme-to-substrate (E/S) ratio of 6% (*w/w*). The hydrolysis reaction was performed in a shaker incubator (Unimax 1010; Heidolph, Schwabach, Germany), then the enzymes were thermally inactivated (90 ◦C, 10 min), and the solution was cooled down to room temperature. The hydrolysate solution was then centrifugated at 4550× *g* for 10 min and the supernatant was collected for further analysis [4].
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007afdec-bed4-405d-873d-c355ba9add0e.234 | *2.3. Degree of Hydrolysis (DH)*
The extent of enzymatic hydrolysis can be defined by the degree of hydrolysis (DH), which is a key factor determining the chain length of peptides, and thereby their functional properties. A higher DH corresponds to mean shorter peptide length and vice versa. This index is significantly influenced by hydrolysis time and the type of enzyme used. To determine DH, 1 mL of hydrolysate was added to 1 mL of TCA (0.44 M), followed by centrifugation at 7800× *g* for 10 min at 4 ◦C. The collected supernatants were analyzed for soluble protein by the Bradford assay. DH was estimated using the following equation [15]:
$$\text{DH} \left( \% \right) = \frac{\text{TCA} - \text{Soluble Protein of Hydrolysis}}{\text{Total Protein of Sample (non-hydrylyzed)}} \times 100\tag{1}$$
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007afdec-bed4-405d-873d-c355ba9add0e.235 | *2.4. Amino Acid Profile*
An RP-HPLC apparatus (Young Lin Acme 9000, YL Instruments, Anyang, Korea) equipped with a reverse-phase column (150 mm × 4.6 mm × 5 μm; RP-C18 ODS-A, Barcelona, Spain), a fluorescence detector (LC305; Lab Alliance, State College, PA, USA), and a mobile phase of acetate buffer (50 mM at pH 3.4, with a flow rate of 1.3 mL/min) were used to determine the amino acids in *Spirulina* protein hydrolysates. For this purpose, the hydrolysate was intensively treated with HCl (6 M) at 110 ◦C for 24 h. The digested sample was derivatized with orthophthaldehyde and injected to the HPLC column. The amount of amino acids in hydrolysates was expressed as mg/100 g protein. The biological value (*BV*) and amino acid score (*AAS*) of hydrolysates, as nutritional parameters, were determined using the following equations [16]:
$$AAS = \frac{\% \text{ Essertial amino acids in sample}}{\% \text{ Essenential amino acids recommended by FAO}} \tag{2}$$
$$BV\left(\%\right) = 10^{2.15} \times Ly^{0.41} \times \left(Phc + Tyr\right)^{0.6} \times \left(Met + Cys\right)^{0.77} \times Thr^{0.24} \times Tr p^{0.21} \tag{3}$$
where each amino acid symbol is expressed as % amino acid in sample/% amino acid FAO pattern.
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007afdec-bed4-405d-873d-c355ba9add0e.236 | *2.5. Preparation of Spirulina Hydrolysate (HS)-Loaded Liposomes*
HS-encapsulated nanoliposomes were prepared using the thin layer hydration method as described by Mohammadi et al. [7] with slight modifications. For this procedure, 1.2 % (*w*/*v*) Phospholipon 90 G (soybean lecithin of ~90% phosphatidylcholine; Lipoid GmbH, Ludwigshafen, Germany) and 2 stabilizing agents (cholesterol or 0.15% (*w*/*v*) γ-oryzanol) were dissolved in 15 mL of 96% ethanol and stirred on a hotplate at 50 ◦C for complete solubilization. Subsequently, the solvent was evaporated using a rotary evaporator (Heidolph, Germany) at 50 ◦C until a thin film was formed in the roundbottomed flasks. The resulting lipid films were hydrated with 15 mL of DW containing HS at 0.3% (*w*/*v*) with continuous agitation on a rotary evaporator at 55 ◦C, followed by sonication using a sonication probe (130 W, 20 kHz; Vibra-Cell Sonics & Materials, Newtown, CT, USA) at 80% sonication strength for 10 min. During sonication, the sample was placed into an ice bath to avoid overheating of dispersion. To prepare the empty liposomes, the same method was applied, except HS was excluded in the hydration step and the thin layer was hydrated only with DW.
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007afdec-bed4-405d-873d-c355ba9add0e.237 | *2.6. Preparation of Chitosan-Coated Nanoliposomes*
For coating with chitosan, prepared nanoliposomes were added to chitosan solution (0.4%, *w*/*v*) dissolved in acetic acid (1% *v/v*) in a drop-wise manner with a volume ratio of 1:1 and stirred for 2 h.
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007afdec-bed4-405d-873d-c355ba9add0e.239 | 2.7.1. Measurement of Particle Size and Zeta Potential (ζ)
Liposome dispersions were diluted 1:10 with DW before analysis by a zetasizer (Zetasizer Nano ZS, Malvern Instruments Ltd., Malvern, UK).
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007afdec-bed4-405d-873d-c355ba9add0e.240 | 2.7.2. Encapsulation Efficiency
Encapsulation efficiency (*EE*) was determined by separating encapsulated hydrolysates from free ones using an Amicon filter (Amicon Ultra-15, with molecular weight cutoff of 30 kDa; Millipore Corp., Cork, Ireland), followed by centrifugation at 3000 rpm for 10 min. Free and total hydrolysates were determined by calculating the protein amount using the Bradford method as described previously.
*EE* was determined according to following equation:
$$EE = \frac{\text{Total protein content} - \text{Amount of free hydrolysisate}}{\text{Total protein content}} \times 100\tag{4}$$
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007afdec-bed4-405d-873d-c355ba9add0e.241 | 2.7.3. Scanning Electron Microscopy (SEM)
To investigate the morphological features of the vesicles, γ-oryzanol liposome (with and without chitosan coating) was dispersed onto the laboratory lamel and dried at 37 ◦C, then transferred to adhesive-coated aluminum pin stubs. The stubs were coated with a thin layer of gold and examined using a scanning electron microscope (MIRA3, TESCAN, Brno, Czech Republic) [17].
#### 2.7.4. Transmission Electron Microscopy (TEM) Measurements
For TEM measurement, 5 μL of each sample was placed onto a copper grid coated with carbon film for 3 min before being blotted off using filter paper. After that, 10 μL of contrast dye containing 2% uranyl acetate was placed onto the grid, left for 2 min, and blotted off with filter paper. Finally, the grids were loaded onto a specimen holder and then into a transmission electron microscope (100 Kv; LEO 906, Zeiss, Oberkochen, Germany).
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007afdec-bed4-405d-873d-c355ba9add0e.242 | 2.7.5. Fourier-Transform Infrared Spectroscopy (FT-IR)
To determine the functional groups of liposomes, lyophilized samples (ALPHA 1–4 LD freeze dryer, Martin Christ, Osterode am Harz, Germany) were formed into KBr pellets with a mass ratio of 1:100 [17]. The samples were analyzed using FTIR (4300, Shimadzu, Kyoto, Japan) from 4000 to 400 cm−<sup>1</sup> with a minimum of 256 scans/spectrum and a constant scan speed of 4◦/s.
| doab | 2025-04-07T03:56:59.221387 | 17-11-2022 17:23 | {
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007afdec-bed4-405d-873d-c355ba9add0e.243 | 2.7.6. Determination of Total Phenolic Content (TPC)
TPC was performed according to the Folin–Ciocalteu method described by de Araujo et al. [18]. Briefly, 300 μL of sample was mixed with 125 μL of Folin–Ciocalteu reagent and 1825 μL of DW. After the mixture was vortexed for 5 min at ambient temperature, 250 μL of sodium carbonate solution (20%, *w*/*v*) was added and it was vortexed for another 5 min. Then, the mixture was placed in a water bath at 40 ◦C for 30 min. The samples were then centrifuged at 10,000 rpm for 10 min, and the absorbance of the upper phase was measured at 765 nm by a spectrophotometer (Ultrospec 2000; Scinteck, Cambridge, UK). The results were expressed as mg gallic acid per g sample using the following formula [18]:
$$\mathbf{C} = c \frac{V}{\mathbf{m}} \tag{5}$$
where *C* is the total phenolic content (mg GAE/g dry extract), *c* is the concentration of gallic acid obtained from the calibration curve (mg/mL), *V* is the volume of extract (mL), and m is the mass of the extract (g).
| doab | 2025-04-07T03:56:59.221434 | 17-11-2022 17:23 | {
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007afdec-bed4-405d-873d-c355ba9add0e.244 | 2.7.7. Antioxidant Activity of Protein Hydrolysates
| doab | 2025-04-07T03:56:59.221515 | 17-11-2022 17:23 | {
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007afdec-bed4-405d-873d-c355ba9add0e.245 | 2.7.7.1. DPPH Radical Scavenging Activity
To measure DPPH radical scavenging activity, 1 mL of each concentration of hydrolysates was added to 1 mL of DPPH solution (0.1 mM), followed by incubation for 30 min in the dark. The absorbance was read at 517 nm and the DPPH radical scavenging activity was calculated by the following equation [19]:
$$\text{DPPH} \,\text{radial\,\,scavenging\,\,activity\,\,(\%)} = \frac{A\_{\text{control}} - A\_{\text{sample}}}{A\_{\text{control}}} \times 100\tag{6}$$
where *Acontrol* and *Asample* are the absorbance of the control and sample, respectively.
| doab | 2025-04-07T03:56:59.221539 | 17-11-2022 17:23 | {
"license": "Creative Commons - Attribution - https://creativecommons.org/licenses/by/4.0/",
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007afdec-bed4-405d-873d-c355ba9add0e.246 | 2.7.7.2. ABTS Radical Scavenging Activity
The mixture of ABTS (7 mM) and potassium persulfate (2.45 mM) with a volume ratio of 1:1 generates a green-blue reagent (ABTS+) after 12–16 h incubation in the dark, and has maximum absorption at 734 nm. When this cationic radical is exposed to the hydrogen donating compound, the green-blue is decolorized and the color intensity is measured at 734 nm. To measure ABTS radical scavenging activity, 40 μL of the prepared hydrolysate concentration was mixed with 4 mL of diluted ABTS solution, vortexed vigorously for 30 s, and incubated in the dark for 6 min. The absorbance was measured at 734 nm. The ABTS radical scavenging activity was calculated by the following equation [20]:
$$\text{ABTS radical saving activity (\%)} = \frac{A\_{\text{control}} - A\_{\text{sample}}}{A\_{\text{control}}} \times 100\tag{7}$$
#### 2.7.7.3. Ferric Reducing/Antioxidant Power (FRAP) Assay
The capability of hydrolysate to reduce Fe+3 ions present in the complex to a Fe+2 form with 2,4,6-tri (2-pyridyl)-s-triazine (TPTZ) was determined by the ferric ion reducing capacity (FRAP) assay as described previously. The FRAP reagent was freshly prepared by mixing TPTZ (10 mM) dissolved in 40 mM HCL, iron (III) chloride hexahydrate (20 mM) dissolved in water, and acetate buffer (0.3 mM) at pH 3.6 at a ratio of 1:1:10 (*v/v/v*) and warming it to 37 ◦C. Then, 900 μL of the working solution was mixed with 100 μL of different concentrations of hydrolysate and the corresponding nano-formulated system, followed by incubation of the mixture at 37 ◦C for 30 min, and the resulting blue color absorbance at 595 nm was recorded. Different concentrations of FeSO4, in the range 0–1 mM, were used as the calibration curve [21].
| doab | 2025-04-07T03:56:59.221586 | 17-11-2022 17:23 | {
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007afdec-bed4-405d-873d-c355ba9add0e.247 | *2.8. Storage Stability of HS-Loaded Liposome and Chitosome*
In order to perform this test, HS-loaded liposomes (stabilized with γ-oryzanol) and the corresponding chitosomes were stored at 4 ◦C for 1 month. The mean diameter, PDI, ζ-potential, and precipitation were monitored during storage by DLS as described in Section 2.7. The residual antioxidant activity of selected formulations was controlled by ABTS assay.
| doab | 2025-04-07T03:56:59.221798 | 17-11-2022 17:23 | {
"license": "Creative Commons - Attribution - https://creativecommons.org/licenses/by/4.0/",
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007afdec-bed4-405d-873d-c355ba9add0e.248 | *2.9. Spray-Drying of Chitosomes*
Before spray drying, the γ-oryzanol-stabilized chitosome dispersion was mixed with maltodextrin solution (40% *w*/*v*) at a mass ratio of 40:60, and stirred overnight at room temperature. Then, the resulting dispersions were spray-dried using a Mini Spray Dryer (Büchi Labortechnik, Flawil, Switzerland). The inlet and outlet air temperature were 130 and 75 ◦C, respectively. Dried powders were stored in airtight containers and placed in a desiccator at room temperature.
| doab | 2025-04-07T03:56:59.221847 | 17-11-2022 17:23 | {
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"title": "Characterization and Encapsulation of Natural Antioxidants: Interaction, Protection and Delivery",
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007afdec-bed4-405d-873d-c355ba9add0e.249 | *2.10. Characterization of Spray-Dried Powder*
Production yield was measured by calculating the mass ratio of the produced powder to the total solid content in the feed. Other physical properties of the spray-dried powder, such as moisture content, bulk density, and solubility, were computed using the methods described by Sarabandi et al. [20].
The particle morphology of spray-dried powder was evaluated by scanning electron microscopy (SEM; MIRA3, TESCAN, Brno, Czech Republic).
To determine the size, polydispersity, and ζ of reconstituted nanoliposomes, the powder was dissolved in an appropriate concentration and its particle size and ζ potential were determined by a zetasizer.
#### *2.11. Statistical Analysis*
Statistical analysis was performed using SPSS software (version 24.0, IBM, Chicago, IL, USA). Normal distribution and variance homogeneity had been previously tested (Shapiro– Wilk). Data of 3 repetitions were subjected to analysis of variance (ANOVA), followed by Tukey's test at a 5% significance level.
## **3. Results and Discussion**
#### *3.1. Characterization of Hydrolyzed Spirulina Protein (HS)*
Extracted *Spirulina* isolate was hydrolyzed by pepsin enzyme. The hydrolysis degree was found to be 16.5% over 4 h hydrolysis time. The solubility of hydrolysate under harsh acidic conditions was improved after enzymatic hydrolysis, but the highest solubility was obtained under alkaline pH conditions.
Figure 1 shows the amino acid composition of pepsin-hydrolyzed peptides. According to the obtained profile, the hydrolysate was rich in acidic amino acids (aspartic and glutamic acid), arginine, valine, lysine, alanine, glycine, threonine, and leucine. All the essential amino acids (except sulfur-containing amino acids) were present in the hydrolysate at concentrations higher than the FAO recommended levels for adults. Moreover, the hydrolysate showed good nutritional value as determined by amino acid score (72%) and biological value (78%), and good antioxidant activity (IC50 1 mg/mL).
**Figure 1.** Amino acid composition of pepsin-hydrolyzed peptides: (1) aspartic acid; (2) glutamic acid; (3) asparagine; (4) histidine; (5) serine; (6) glutamine; (7) arginine; (8) glycine; (9) threonine; (10) alanine; (11) tyrosine; (12) methionine; (13) valine; (14) phenylalanine; (15) isoleucine; (16) leucine; (17) lysine; (18) tryptophan.
Compared to native protein (with a DPPH IC50 value of 3 mg/mL), the hydrolysates had a significantly lower IC50 value of 1.0 mg/mL, indicating their effectiveness in scavenging DPPH radicals. The ABTS IC50 of hydrolysate was estimated to be 2 mg/mL. Compared with native protein (with an IC50 value of 4.5 mg/mL), the hydrolysates had lower IC50 values, indicating their effectiveness against ABTS radicals.
Encapsulating these bioactive compounds in lipid-based nanocarriers such as liposomes can be an appropriate solution to cover all of the mentioned problems and increase their efficacy under different conditions.
| doab | 2025-04-07T03:56:59.221893 | 17-11-2022 17:23 | {
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007afdec-bed4-405d-873d-c355ba9add0e.250 | *3.2. Characterization of Uncoated and Chitosan-Coated Liposomal Dispersions*
In this study, HS was encapsulated into the liposomal carrier and its experimental characteristics were investigated. Two stabilizers (cholesterol and γ-oryzanol) were applied in the preparation of primary liposome dispersion, then the resulting nanoliposomes were coated with cationic chitosan polymer at a final concentration of 0.2% *w*/*v* (this concentration resulted in the smallest particle size and highest surface charge on the coated liposome dispersions), and their effects on the physicochemical properties of the resulting formulations were examined. The mean particle diameter and ζ of primary and chitosan-coated nanoliposomes (chitosomes) are shown in Table 1. Liposomes stabilized by cholesterol and γ-oryzanol had a small particle size and homogeneous size distribution, and their surface charge was between −11 and −14 mV. Following the addition of the chitosan polymer, the particle size and PDI of nanoliposomes increased and ζ changed from negative to positive values (approximately 29 mV), confirming that cationic chitosan successfully covered the primary liposomes. These findings are in accordance with those of Altin et al. [22], who reported that surface coating of primary liposomes containing phenolic extract from cocoa hull waste with cationic chitosan by electrostatic deposition increased the particle size of liposomes and the secondary liposomes had a positive charge.
SEM and TEM images of primary liposomes (γ-oryzanol-liposomes) and the corresponding chitosomes are shown in Figure 2. The cholesterol-liposome and γ-oryzanolliposome had similar shape and morphology. The results obtained from the zetasizer apparatus were somewhat confirmed by SEM and TEM. The SEM images show spherical particles with a small particle size < 100 nm and narrow distribution. In TEM images, the spherical structure and monodispersed distribution of primary liposomes are very clear [23].
**Figure 2.** (**a**,**b**) SEM and (**c**,**d**) TEM images of primary liposomes stabilized with γ-oryzanol (**a**,**c**) and corresponding γ-oryzanol-chitosomes (**b**,**d**).
**Table 1.** Characteristics of hydrolysate-loaded γ-oryzanol-liposome and chitosome.
| doab | 2025-04-07T03:56:59.222321 | 17-11-2022 17:23 | {
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007afdec-bed4-405d-873d-c355ba9add0e.251 | *3.3. Encapsulation Efficiency (EE)*
The *EE* of the primary HS-loaded liposomes stabilized by cholesterol and γ-oryzanol is given in Table 1. Overall, both liposomes showed high *EE* > 85%, indicating that both stabilizing agents had good potential for encapsulation of HS. Incorporating sterol compounds in the liposome structure significantly increased the rigidity of the liposome membrane; thus, the system could encapsulate a larger amount of hydrophilic bioactive material. In another study, high *EE* was reported for fish hydrolyzed collagen-loaded liposomes stabilized by cholesterol and glycerol [24]. In another study, orange seed protein hydrolysates were produced using alcalase and pepsin enzymes, which were incorporated into uncoated liposomes and chitosome systems. The hydrolysates produced with alcalase showed a higher *EE* than those produced with pepsin. The authors suggested that this difference may be related to the higher DH of alcalase hydrolysate (approximately 24%), the lower molecular weight of resulting peptides compared to pepsin hydrolysate, and the easy incorporation of alcalase hydrolysate into the aqueous core of liposomes [14]. The authors also claimed that incorporating peptides into the chitosomes led to increased *EE* of vesicles compared to plain liposomes. This may be related to occupying pores in the surface of the liposome surface preventing the leakage of incorporated bioactive materials. These findings were consistent with those reported by [14,20].
| doab | 2025-04-07T03:56:59.222457 | 17-11-2022 17:23 | {
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007afdec-bed4-405d-873d-c355ba9add0e.252 | *3.4. Determination of Total Phenolic Content and Antioxidant Activity of Liposomes and Chitosomes*
Liposomal nanocarriers can be applied for encapsulation of both liposoluble and hydrophilic antioxidant and phenolic compounds to improve their bioavailability. The total phenolic and antioxidant capacity of uncoated liposomes and chitosome dispersions are shown in Table 2. There was no significant difference (*p* < 0.05) between HS and HS-loaded cholesterol-liposomes by TPC, DPPH, ABTS, or FRAP assay, indicating that the phenolic compounds and, subsequently, the antioxidant properties of HS were properly preserved in the nanoliposomal carrier. Preservation of the antioxidant activity of anthocyanin-rich black carrot extract after 21 days of storage by encapsulating in liposomes has been reported [25].
**Table 2.** Antioxidant activity and total phenol content of hydrolysate-loaded γ-oryzanol-liposomes and chitosomes.
Means in same column with different superscripts (a, b) are statistically different (*p* < 0.05).
After the cholesterol-liposome surface was coated with chitosan, the antioxidant activity remained unchanged as compared to the uncoated liposome. The antioxidant activity of chitosan has been reported by others [22]. The authors suggested that the phenolic bioactive material could be partially located on the surface of the liposome, and consequently chitosan–phenolic compound conjugates might be formed, and these couples synergistically improve the antioxidant activity [22].
Conversely, in another study, after coating the surface of sour cherry extract-loaded liposomes with cationic chitosan, the TPC content decreased from 38.19 to 31.23 mg/L. The authors suggested that the available chitosan on the liposome surface might block the availability of phenolic compounds on the surface of uncoated liposomes [26].
γ-Oryzanol, a plant sterol with a structure similar to cholesterol, has a wide capacity for scavenging free radicals, consequently preventing lipid oxidation. The HS-loaded γ-oryzanol-liposomes showed higher TPC and antioxidant capacity. This was attributed to the cooperative scavenging capacity of γ-oryzanol with HS in a liposome system [27]. This cooperative antioxidative effect was reported by Li et al. [28]. Sage extract (SE) and zein hydrolysate in combination showed higher antioxidant activity than the simple sum of their individual effects [28].
| doab | 2025-04-07T03:56:59.222552 | 17-11-2022 17:23 | {
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007afdec-bed4-405d-873d-c355ba9add0e.253 | *3.5. Fourier-Transform Infrared Spectroscopy (FTIR)*
The structural changes in synthesized liposomes with and without HS and the successful chitosan coating were confirmed by FTIR spectroscopy. From the IR spectrum of the HS (Figure 3a), a broadband at 3300 cm−<sup>1</sup> was attributed to O–H and N–H stretching and two bands at 2926 and 2853 cm−<sup>1</sup> were related to CH2 stretching vibrations of aliphatic chains. The amide region bands (1658 cm−<sup>1</sup> corresponding to protein amide I, 1550 cm−<sup>1</sup> corresponding to protein amide II, and 1247 cm−<sup>1</sup> related to protein amide III) were clearly visible in the IR spectrum of the HS [29].
**Figure 3.** FTIR spectra of (**a**) *Spirulina* hydrolysate (HS); (**b**) blank nanoliposomes stabilized with γ-oryzanol; (**c**) HS-loaded γ-oryzanol nanoliposomes; and (**d**) HS-loaded γ-oryzanol-chitosomes.
Blank nanoliposomes (Figure 3b) were observed at the following wavenumbers: 3438 cm−<sup>1</sup> related to hydroxyl stretch vibration, 2925 cm−<sup>1</sup> attributed to stretch vibrations of a methylene group, 1735 cm−<sup>1</sup> mainly related to the stretching vibration of the polar head ester groups of phospholipids, 1654 cm−<sup>1</sup> related to C=C stretching vibrations, 1246 and 1109 cm−<sup>1</sup> corresponding to symmetric and antisymmetric stretch vibrations of a phosphate group, and 958 cm−<sup>1</sup> related to asymmetrical stretch vibrations of N+/CH3.
Comparing the IR spectra of HS-loaded γ-oryzanol-liposomes (Figure 3c) and corresponding blank liposomes (Figure 3b), a great similarity between their spectra can be observed. The incorporation of hydrolysate into the liposomal carrier resulted in a shift in some frequencies. The most important of these changes were slight shifts at 1735 and 1654 cm−<sup>1</sup> to 1737 and 1658 cm−<sup>1</sup> for HS-loaded liposomes, which may correspond to the possible interaction of HS with carbonyl ester groups at the interfacial part of the liposomal bilayers [30].
For the HS-loaded γ-oryzanol-chitosomes (Figure 3d), a shift from 3379 to 3415 cm−<sup>1</sup> was detected, which may correspond to hydrogen bonding between hydroxyl or amino groups of chitosan and carboxylic acid or amino groups of hydrolysates. Moreover, after chitosan coating, significant changes in the absorption bands of acyl chains (3000–2800 cm–1) were detected. These peaks were converted into two narrow and intense peaks at 2862 and 2924 cm−<sup>1</sup> in the case of chitosan-coated liposomes. Further evidence for electrostatic conjugation of chitosan on liposome surface was a considerable shift to higher frequencies in the carbonyl group (from 1737 to 1739 cm–1). This indicates that the carbonyl groups are involved with cationic groups of chitosan, resulting in the destruction of some hydrogen bonds [31].
| doab | 2025-04-07T03:56:59.222703 | 17-11-2022 17:23 | {
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007afdec-bed4-405d-873d-c355ba9add0e.254 | *3.6. Storage Stability of HS-Loaded Liposomes and Chitosomes*
Regarding our previous research on the effect of temperature on the physical stability of vitamin D3-loaded liposomes and the instability and aggregation of the formulation at ambient temperature due to higher fluidity of the lipid bilayer and higher loss of encapsulated bioactive material, refrigerator temperature (4 ◦C) was selected to examine the physical stability of selected formulations [7]. Liposomes as vesicular droplets have permeable and flexible bilayers and a high tendency to fuse and aggregate, resulting in the release of encapsulated bioactive materials during storage. In this study, stabilizing agents (cholesterol and γ-oryzanol) and coating material (chitosan) were tested to enhance the physical stability of liposomes. HS-loaded γ-oryzanol-liposomes and the corresponding chitosomes had no marked difference (*p* < 0.05) in magnitude, PDI, and ζ after 30 days of storage at 4 ◦C (Table 3). Moreover, both samples preserved more than 90% of their initial antioxidant activity in ABTS radical scavenging activity (*p* < 0.05). When the storage time was expanded, slight lipid oxidation may have occurred in unsaturated fatty acids of the phospholipid bilayers, leading to decreased antioxidative activity of the tested formulations. On the other hand, no significant precipitate was observed, especially in the HS-loaded chitosome system, during storage. When storage time was expanded to 2 months, the chitosome system showed higher physical stability compared to the uncoated liposomes. In contrast, the stability of HS-loaded cholesterol-liposomes in terms of mean diameter, PDI, and ζ was much lower than that of HS-loaded γ-oryzanol-liposomes at 4 ◦C. When storage time was expanded, the mean diameter and PDI of these formulations significantly increased (*p* < 0.05), and observable precipitates were separated into round-bottomed Falcon tubes.
**Table 3.** Physical stability of hydrolysate-loaded γ-oryzanol-liposomes and chitosomes during one month at 4 ◦C.
Means in same column with different superscripts (a, b, c, d) are statistically different (*p* < 0.05).
On the other hand, the residual antioxidant activity of HS-loaded cholesterol-liposomes was 50% of its initial antioxidant activity. This observed instability of HS-loaded cholesterolliposomes could have resulted from the poor coating of liposomal space cores with cholesterol stabilizing agents, and possible replacement of empty spaces with hydrophilic domain residues (glycine, arginine, and lysine) of hydrolysates. Nevertheless, some hydrophobic domains of *Spirulina* hydrolysates could interact with acyl chains of the lipid bilayer of liposome through hydrophobic binding, leading to more flexibility and fluidity of liposomes [32]. HS-loaded γ-oryzanol-chitosomes showed better results in terms of particle size, PDI, aggregation, and antioxidant activity compared to HS-loaded cholesterolliposomes during storage. In summary, HS-loaded γ-oryzanol-liposomes and HS-loaded γ-oryzanol-chitosomes showed promising storage stability with no precipitate or change in mean diameter and a slight decrease in antioxidant activity at 4 ◦C. Thus, γ-oryzanol as a stabilizing agent and chitosan polymer as a coating material were able to reduce membrane fluidity and flexibility, contributing to stability.
In another study, different stabilizers (cholesterol and glycerol) were used to encapsulate peptides obtained from defatted Asian sea bass skin. Regarding the results, both formulations showed small particle size and high encapsulation efficiency. However, after the lyophilization process, hydrolysate-loaded cholesterol-liposomes showed higher stability and higher antioxidant activity in the gastrointestinal tract than lyophilized glycineliposomes during storage at 25 ◦C for 28 days [24]. It was reported that curcumin-loaded chitosomes showed higher physical stability than curcumin-loaded uncoated liposomes due to the lower flexibility of chitosan-coated liposomes and, as a result, lower membrane fusion between droplets [33].
#### *3.7. Production Yield and Physicochemical Properties of Spray-Dried Chitosomes*
The powder moisture content was found to be 4.65 ± 0.61%, which is lower than the specified minimum moisture content of many powders used in food applications to inhibit microbiological spoilage and lipid oxidation and extend the product's shelf life.
The bulk density of the powder was 0.31 ± 0.01 g cm−3. A higher bulk density has several advantages, including more convenient storage conditions due to lower space requirements for storage and greater protection against oxidation during storage due to the presence of less air in powder. In addition, the solubility and production yield of powder were found to be 96% and 57%, respectively.
## *3.8. Microstructure and Particle Size Distribution*
The morphology of dried particles is mostly affected by the evaporation rate and viscoelastic properties of shell material. SEM images (Figure 4) show mostly spherical particles with little evidence of roughness or fracturing in the microcapsules. Moreover, most of the spray-dried powders showed a well-defined spherical shape with a particle size of around 1–3 μm, which is smaller than the threshold diameter used in food fortification (10–50 μm) [34]. The coarse powder results in a sandy and unfavorable mouthfeel in the fortified food products. These results are in accordance with those reported by Sarabandi et al. [13].
**Figure 4.** (**a**) SEM image of spray-dried hydrolysate-loaded γ-oryzanol-chitosomes; (**b**) particle size result of reconstituted γ-oryzanol-chitosome powder.
| doab | 2025-04-07T03:56:59.222857 | 17-11-2022 17:23 | {
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} |
007afdec-bed4-405d-873d-c355ba9add0e.256 | 3.9.1. Physical Properties
It is essential to conserve the physicochemical properties of nanoliposomes after the spray-drying process. The effect of spray-drying on particle size, PDI, and ζ of reconstituted chitosomes was investigated (Table 4). As expected, the chitosan coating increased the physical stability of nanocarriers after spray-drying, and slight changes were observed in the size and ζ of reconstituted chitosomes, but the PDI for these systems changed from 0.33 to 0.51. In a previously reported study, the effect of spray-drying on the physical properties of reconstituted liposomal powders was investigated. Regarding those results, the uncoated liposome systems were immediately unstable, as the maltodextrin was added to the dispersion as a wall matrix, i.e., these systems were not spray-dried. However, the coated systems did not significantly affect the spray-drying process and the diameter of the liposomal powder was smaller than that of liposomal dispersion before thermal processing. The authors suggested that when coated liposomal systems are added to a solution containing salts or sugars, which have the potential to induce an osmotic driving force and reduce water activity, migration of water molecules happens from the core to the liposomal surface, reducing the concentration gradient between the internal and external aqueous phases of the liposomes, thus reducing the size of the liposomal powder [35]. Moreover, the addition of maltodextrin as a hydrophilic nonionic polysaccharide had no effect on the ζ of coated liposomes.
**Table 4.** Physicochemical properties of reconstituted hydrolysate-loaded γ-oryzanol-chitosomes.
Means in same column with different superscripts (a, b) are statistically different (*p* < 0.05).
#### 3.9.2. Retention of Antioxidant Activity (AA)
The effect of spray-drying and thermal stress on the retention of ABTS radical scavenging activity in the chitosome system is shown in Table 4. There was no significant difference between these two indices in chitosome system retention before and after the spray-drying process (*p* < 0.05), which shows the positive effect of chitosan coating on the preservation of the biological activity of peptide fractions against thermal stresses during spray-drying. Our results are consistent with those reported by Sarabandi et al. [13]. In another study, about 39% of total phenols, 30% of flavonoids, and 47% of radical scavenging activity of extract were maintained after the spray-drying of nanoliposomes [22]. The spray-drying of nanoliposomes loaded with black mulberry extract resulted in the preservation of approximately 69% of total phenolic and 56% of anthocyanin compounds in chitosan-coated nanoliposomes [36].
| doab | 2025-04-07T03:56:59.223159 | 17-11-2022 17:23 | {
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"url": "https://mdpi.com/books/pdfview/book/6198",
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"title": "Characterization and Encapsulation of Natural Antioxidants: Interaction, Protection and Delivery",
"publisher": "MDPI - Multidisciplinary Digital Publishing Institute",
"isbn": "9783036554563",
"section_idx": 256
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007afdec-bed4-405d-873d-c355ba9add0e.257 | **4. Conclusions**
*Spirulina plantensis* hydrolysate was successfully encapsulated into nanoliposomes using the thin-layer hydration method of sonication. This study shows that stabilizing the liposome structure with γ-oryzanol and covering the liposomes with a polycationic chitosan polymer provided long-term physical stability, and the system preserved its antioxidant capacity over time. Thus, this study suggests that chitosomes stabilized with γ-oryzanol could be used as a promising delivery system to protect against the loss of hydrolysate under processing or storage conditions.
**Author Contributions:** Methodology, M.M.; formal analysis, M.M. and R.S.; data curation and preparation, M.M.; Project administration, H.H.; writing—original draft preparation, M.M. and R.S.; Review and editing, H.H., M.G., M.P., and J.M.L. All authors have read and agreed to the published version of the manuscript.
**Funding:** This research received no external funding.
**Institutional Review Board Statement:** Not applicable.
**Informed Consent Statement:** Not applicable.
**Data Availability Statement:** Data is contained within the article.
**Acknowledgments:** The authors would like to acknowledge the Drug Applied Research Center, Tabriz University of Medical Sciences, for financial support.
**Conflicts of Interest:** The authors declare no conflict of interest.
| doab | 2025-04-07T03:56:59.223329 | 17-11-2022 17:23 | {
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"book_id": "007afdec-bed4-405d-873d-c355ba9add0e",
"url": "https://mdpi.com/books/pdfview/book/6198",
"author": "",
"title": "Characterization and Encapsulation of Natural Antioxidants: Interaction, Protection and Delivery",
"publisher": "MDPI - Multidisciplinary Digital Publishing Institute",
"isbn": "9783036554563",
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007afdec-bed4-405d-873d-c355ba9add0e.259 | *Article* **Yogurt Fortification by the Addition of Microencapsulated Stripped Weakfish (***Cynoscion guatucupa***) Protein Hydrolysate**
**Karina Oliveira Lima 1, Meritaine da Rocha 2, Ailén Alemán 3, María Elvira López-Caballero 3,\*, Clara A. Tovar 4, María Carmen Gómez-Guillén 3, Pilar Montero 3,\* and Carlos Prentice 1,†**
**Abstract:** The aim of the present work was to fortify yogurt by adding a stripped weakfish (*Cynoscion guatucupa*) protein hydrolysate obtained with the enzyme Protamex and microencapsulated by spray drying, using maltodextrin (MD) as wall material. The effects on the physicochemical properties, syneresis, texture, viscoelasticity, antioxidant and ACE inhibitory activities of yogurt after 1 and 7 days of storage were evaluated. In addition, microbiological and sensory analyses were performed. Four yogurt formulations were prepared: control yogurt (without additives, YC), yogurt with MD (2.1%, YMD), with the free hydrolysate (1.4%, YH) and the microencapsulated hydrolysate (3.5%, YHEn). Yogurts to which free and microencapsulated hydrolysates were added presented similar characteristics, such as a slight reduction in pH and increased acidity, with a greater tendency to present a yellow color compared with the control yogurt. Moreover, they showed less syneresis, the lowest value being that of YHEn, which also showed a slight increase in cohesiveness and greater rheological stability after one week of storage. All yogurts showed high counts of the microorganisms used as starters. The hydrolysate presence in both forms resulted in yogurts with antioxidant activity and potent ACE-inhibitory activity, which were maintained after 7 days of storage. The incorporation of the hydrolysate in the microencapsulated form presented greater advantages than the direct incorporation, since encapsulation masked the fishy flavor of the hydrolysate, resulting in stable and sensorily acceptable yogurts with antioxidant and ACE inhibitory activities.
**Keywords:** fish protein hydrolysate; microencapsulation; yogurt; physicochemical properties; antioxidant activity; antihypertensive activity
| doab | 2025-04-07T03:56:59.223445 | 17-11-2022 17:23 | {
"license": "Creative Commons - Attribution - https://creativecommons.org/licenses/by/4.0/",
"book_id": "007afdec-bed4-405d-873d-c355ba9add0e",
"url": "https://mdpi.com/books/pdfview/book/6198",
"author": "",
"title": "Characterization and Encapsulation of Natural Antioxidants: Interaction, Protection and Delivery",
"publisher": "MDPI - Multidisciplinary Digital Publishing Institute",
"isbn": "9783036554563",
"section_idx": 259
} |
007afdec-bed4-405d-873d-c355ba9add0e.260 | **1. Introduction**
Milk and dairy products, such as yogurt, are widely appreciated and consumed worldwide because of their sensory and nutritional characteristics [1,2]. Compared with milk, yogurt is more nutritious and an excellent source of protein, calcium, phosphorus, riboflavin, thiamine, vitamin B12, niacin, magnesium, and zinc [3]. However, despite their beneficial health effects, these products are generally not considered to be an important source of bioactive compounds [2].
There are some studies related to the addition to yogurt of bioactive compounds, such as *Spirulina platensis* [4,5], rice bran [6], strawberry pulp [1], mushroom extracts (*Agaricus bisporus*) [7], fish collagen [8], monk fruit extract (*Siraitia grosvenorii*) [9] and *Ficus glomerata* Roxb fruit extract [10], in which the addition of these compounds increased the antioxidant and/or angiotensin-converting enzyme (ACE) inhibitory capacity. This may be
**Citation:** Lima, K.O.; da Rocha, M.; Alemán, A.; López-Caballero, M.E.; Tovar, C.A.; Gómez-Guillén, M.C.; Montero, P.; Prentice, C. Yogurt Fortification by the Addition of Microencapsulated Stripped Weakfish (*Cynoscion guatucupa*) Protein Hydrolysate. *Antioxidants* **2021**, *10*, 1567. https://doi.org/10.3390/ antiox10101567
Academic Editors: Li Liang and Hao Cheng
Received: 3 August 2021 Accepted: 28 September 2021 Published: 1 October 2021
**Publisher's Note:** MDPI stays neutral with regard to jurisdictional claims in published maps and institutional affiliations.
**Copyright:** © 2021 by the authors. Licensee MDPI, Basel, Switzerland. This article is an open access article distributed under the terms and conditions of the Creative Commons Attribution (CC BY) license (https:// creativecommons.org/licenses/by/ 4.0/).
an ideal strategy for facilitating the consumption of bioactive compounds and increasing the functionality of food, as people of all ages widely accept and consume yogurt.
Correspondingly, proteins derived from fish by-products represent a very interesting source of bioactive peptides due to their low cost and the established requirement of reducing agro-industrial waste [11]. Fish protein hydrolysates are sources of peptides with diverse bioactivities, including antioxidant and antihypertensive actions [12]. Some peptides present antihypertensive activity as they are able to inhibit the angiotensin-converting enzyme (ACE), which has a fundamental role in the regulation of blood pressure [12–14]. Moreover, antioxidants can also delay oxidative stress, involved in the occurrence of various diseases, including hypertension and aging [15].
To improve bioactivity, protein hydrolysates and bioactive peptides can be incorporated into different foods, such as dairy products, for their functional properties. However, scarce information about the incorporation of fish hydrolysates into yogurt is available. In one study, bovine and fish skin hydrolysates induced greater syneresis and less firmness and viscoelasticity in skimmed bovine milk yogurt than caseinate hydrolysate, which could be attributed to their hindering effects on yogurt acidification [16]. Moreover, the incorporation of fish protein hydrolysates can be difficult due to their high hygroscopicity, bitter taste, chemical instability, interaction with the food matrix, incompatibility and limited bioavailability [17–19]. Microencapsulation is a process in which the compounds of interest are coated or incorporated into a protective matrix, and is considered effective for overcoming the limitations mentioned above [20].
Among the numerous encapsulation techniques, spray drying is widely used in the food industry [21,22]. This technique consists of atomizing a formulation containing the protective matrix and bioactive compounds in small drops, followed by subsequent rapid drying through a stream of hot air to produce dry microparticles [14,23]. In previous studies, Lima et al. [24] used spray drying to encapsulate a stripped weakfish hydrolysate in maltodextrin. The microencapsulated hydrolysate was characterized, and the stability and biological properties were evaluated in vitro and in vivo in a model of *Caenorhabditis elegans*; the results showed improvements in growth and reproduction rate as well as a protective effect on nematodes exposed to oxidative stress upon consumption of the encapsulates.
Based on the above-mentioned information, the objective of this study was to develop a functional yogurt by incorporating a microencapsulated protein hydrolysate from stripped weakfish (*Cynoscion guatucupa*) in the formulation. For this purpose, sensory and microbiological analyses were performed to confirm that a quality product was obtained. Subsequently, the effects on the physicochemical and rheological properties, texture, and bioactivity of the yogurts after 1 and 7 days of storage were evaluated.
| doab | 2025-04-07T03:56:59.223589 | 17-11-2022 17:23 | {
"license": "Creative Commons - Attribution - https://creativecommons.org/licenses/by/4.0/",
"book_id": "007afdec-bed4-405d-873d-c355ba9add0e",
"url": "https://mdpi.com/books/pdfview/book/6198",
"author": "",
"title": "Characterization and Encapsulation of Natural Antioxidants: Interaction, Protection and Delivery",
"publisher": "MDPI - Multidisciplinary Digital Publishing Institute",
"isbn": "9783036554563",
"section_idx": 260
} |
007afdec-bed4-405d-873d-c355ba9add0e.262 | *2.1. Materials*
The by-products of stripped weakfish (*Cynoscion guatucupa*) were obtained from a fishing company in the city of Rio Grande (RS, Brazil). The carcasses and trimmings were processed in a meat–bone separator (High Tech, HT250C, Chapecó, Brazil), discarding the skin and bones. The resulting muscle was packed in plastic bags and stored at −18 ◦C until use. Protamex enzyme purchased from Sigma-Aldrich (St. Louis, MO, USA) was used in the hydrolysis process. Maltodextrin with a dextrose equivalent (DE) of 5 was purchased from Manuel Riesgo S.A. (Madrid, Spain). All chemical reagents used in this study were of analytical grade.
| doab | 2025-04-07T03:56:59.223996 | 17-11-2022 17:23 | {
"license": "Creative Commons - Attribution - https://creativecommons.org/licenses/by/4.0/",
"book_id": "007afdec-bed4-405d-873d-c355ba9add0e",
"url": "https://mdpi.com/books/pdfview/book/6198",
"author": "",
"title": "Characterization and Encapsulation of Natural Antioxidants: Interaction, Protection and Delivery",
"publisher": "MDPI - Multidisciplinary Digital Publishing Institute",
"isbn": "9783036554563",
"section_idx": 262
} |
007afdec-bed4-405d-873d-c355ba9add0e.263 | *2.2. Protein Hydrolysate*
The fish muscle protein hydrolysate, with a degree of hydrolysis of 5%, was produced using the enzyme Protamex at 50 ◦C and pH 7 as previously described in Lima et al. [24]. The lyophilized hydrolysate was stored at −18 ◦C until use. Hydrolysate characteristics (amino acid profile, Fourier transform infrared spectroscopy, morphological and biological activity) were previously described in Lima et al. [24,25].
| doab | 2025-04-07T03:56:59.224067 | 17-11-2022 17:23 | {
"license": "Creative Commons - Attribution - https://creativecommons.org/licenses/by/4.0/",
"book_id": "007afdec-bed4-405d-873d-c355ba9add0e",
"url": "https://mdpi.com/books/pdfview/book/6198",
"author": "",
"title": "Characterization and Encapsulation of Natural Antioxidants: Interaction, Protection and Delivery",
"publisher": "MDPI - Multidisciplinary Digital Publishing Institute",
"isbn": "9783036554563",
"section_idx": 263
} |
007afdec-bed4-405d-873d-c355ba9add0e.264 | *2.3. Microencapsulation*
The hydrolysate was microencapsulated with maltodextrin (MD) by spray drying as described by Sarabandi et al. [21], with some modifications. The MD and the hydrolysate (60:40 *w*/*w*) were dissolved in distilled water constituting 10% of the total solids and stirred for 3 h at room temperature. Subsequently, the solutions were atomized in a Mini Spray Dryer B-290 (Büchi, Switzerland) at 0.3 L/h with an inlet temperature of 130 ◦C, aspiration rate 100% and an outlet temperature of 70 ± 2 ◦C. Thus, a microencapsulated hydrolysate (microcapsules composed of maltodextrin and hydrolysate) was obtained. The characteristics and stability of the encapsulation have been previously described in Lima et al. [24].
| doab | 2025-04-07T03:56:59.224114 | 17-11-2022 17:23 | {
"license": "Creative Commons - Attribution - https://creativecommons.org/licenses/by/4.0/",
"book_id": "007afdec-bed4-405d-873d-c355ba9add0e",
"url": "https://mdpi.com/books/pdfview/book/6198",
"author": "",
"title": "Characterization and Encapsulation of Natural Antioxidants: Interaction, Protection and Delivery",
"publisher": "MDPI - Multidisciplinary Digital Publishing Institute",
"isbn": "9783036554563",
"section_idx": 264
} |
007afdec-bed4-405d-873d-c355ba9add0e.265 | *2.4. Yogurt Preparation*
Yogurts were made with UHT (Ultra High Temperature) whole cow milk (fat 3.6%; protein 3.1% and carbohydrates 4.6%) and natural yogurt was purchased from a local market (Madrid, Spain). Yogurts were prepared in a Thermomix (Vorwerk & Co., Wuppertal, Germany). Firstly, the milk was heated to 40–45 ◦C while stirring for 3 min, followed by the addition of natural yogurt and stirring for 5 min. Later, the tested ingredients were added, and the mixture was subsequently stirred for another 5 min to constitute the different samples. The temperature was maintained during the mixing steps. For each 100 mL of milk, 12.5 g of natural yogurt and different concentrations of the tested ingredients were added, in order to maintain the same concentration of wall material and hydrolysate present in the microencapsulated hydrolysate. Four lots of yogurts were then obtained: control yogurt (without additional ingredients; lot YC); yogurt with the addition of 2.1 g maltodextrin (wall material; lot YMD); yogurt with the addition of 1.4 g of free protein hydrolysate (lot YH) and yogurt with the addition of 3.5 g of microencapsulated hydrolysate (lot YHEn, composed of 2.1g MD and 1.4 g of hydrolysate). Later, the formulations were transferred to disposable plastic cups of 100 mL and incubated at 43 ◦C until reaching pH 4.6 [26]. The yogurts were stored under refrigeration (6 ± 1 ◦C) for 7 days.
| doab | 2025-04-07T03:56:59.224181 | 17-11-2022 17:23 | {
"license": "Creative Commons - Attribution - https://creativecommons.org/licenses/by/4.0/",
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"title": "Characterization and Encapsulation of Natural Antioxidants: Interaction, Protection and Delivery",
"publisher": "MDPI - Multidisciplinary Digital Publishing Institute",
"isbn": "9783036554563",
"section_idx": 265
} |
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