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007afdec-bed4-405d-873d-c355ba9add0e.38 | *3.2. BIOPEP Database Analysis*
To evaluate the possible inhibition of antioxidants towards oil oxidation, bioactive peptides are collected from the BIOPEP database and various publications (they are listed in Table S1). Further analysis showed that the peptides whose molecular weight is from 200 to 800 plus a GRAVY value of −2 to 1 as the major proportion of these biopeptides had a possibility of stopping oil oxidation (as presented in Figure S1a,b). Interestingly, it is noted that almost three-quarters of the potential antioxidant peptides containing Tyr, Trp, Cys, or Met residues and mainly located at the C-terminus or N-terminus, especially Tyr residue (as presented in Figure S1c). Thus, it could be presumed that the peptides have a molecular weight of 200 to 800, GRAVY value of −2 to 1, and Tyr, Trp, Cys, or Met residues at N- or C- terminus should be selected as effective candidates for antioxidants.
| doab | 2025-04-07T03:56:59.190626 | 17-11-2022 17:23 | {
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007afdec-bed4-405d-873d-c355ba9add0e.39 | *3.3. Selection of Protease*
To find a targeted peptide that consists of active amino acid residues at the N- or C- terminus, the database ExPASy ENZYME integrating available information about proteolytic sites and enzymes was used to select appropriate protease. The use of ExPASy ENZYME allows us easily to determine the cleavage site between all pairs of amino acids in the N- or C-terminal [34]. According to the preferential cleavage sites searched from ExPASy ENZYME (https://enzyme.expasy.org/enzyme-search-ec.html, accessed on 13 August 2020) and from a review [35], the alkaline proteinase (EC number: 3.4.21.62), chymotrypsin (EC number: 3.4.21.1) and pepsin A (EC number: 3.4.23.1) are likely to hydrolyze proteins to the peptides with Tyr, Trp, or Met as C-terminus or N-terminus (Seen in Table 1). Among the three proteases, alkaline hydrolysates exhibited the highest inhibition effect on linoleic acid oxidation [36]. To further reveal the preference of the proteases for producing Tyr, Trp, and Met residues, alkaline and neutral proteinases were chosen to "really" hydrolyze hazelnut protein. It was seen from Table 1 that alkaline proteinase hydrolysates exhibited a high inhibition rate of 95.11 ± 0.17% after 1 mL of the hydrolysates were added to linoleic acid emulsions incubated at 40 ◦C for 48 h. However, the inhibition rate of linoleic acid by neutral hydrolysates was 81.44 ± 1.94%. In addition, protein hydrolysates hydrolyzed by alkaline + neutral proteinase showed an in-between inhibition rate (83.35 ± 1.02%). Similar to the result reported by Ngamsuk [37], that alkaline proteinase was found to give high activity hydrolysate compared to neutrase and mix.
**Table 1.** Preferential cleavage sites of several proteinases and inhibition rate of hazelnut protein hydrolysates against oxidation of linoleic acid.
Note: 1 means not analyzed.
Furthermore, the purity of hazelnut hydrolysates processed by alkaline proteinase was 73.66 ± 2.50%. It was clear from our data that the proteinase alkaline could be appropriately selected for processing the antioxidant peptides from hazelnut protein. The alkaline protease hydrolysates were freeze-dried for further study.
| doab | 2025-04-07T03:56:59.190702 | 17-11-2022 17:23 | {
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007afdec-bed4-405d-873d-c355ba9add0e.40 | *3.4. Screening of Antioxidant Peptides*
Peptides with molecular weights less than 3 kD, which produced from hazelnut protein were sequenced for the screening of potential antioxidants. Furthermore, the peptides which have Tyr, Trp, and Met residues, whose molecule weight is less than 800 and with GRAVY value of −2 to 1 are desirable for us, as sorted out in Table 2. It was seen that seven peptides from hazelnut protein, designed from No.1 to No.7, were screened as potential antioxidants due to up to the desired requirements. Next, they were artificially prepared as follows regarding the determined amino acid sequences. The five peptides FSEY, QIESW, SEGFEW, IDLGTTY, and GEGFFEM were artificially made based on the active amino acids
of their C-terminal. The peptide AHSVVYAIR (designed as No.6) was synthesized in terms of Tyr-containing residue in its middle position. The peptide NLNQCQRYM (named as No.7) was synthesized because of the existence of Tyr, Cys, and Met residues. Besides this, two reported peptides HLHSAT and ADGF from hazelnut protein were artificially synthesized according to their ability to scavenge ABTS and DPPH radical [22,23]. Thus, the four peptides NLNQCQRYM, AHSVVYAIR, HLHSAT, and ADGF were artificially prepared as the control, and they were designed to clarify the feasibility of amino acid residues in screening antioxidant peptides.
**Table 2.** Sequences of synthetic peptides containing Tyr, Trp, Cys, and Met residues.
Notes: A = Alanine, R = Arginine, N = Asparagine, D = Aspartic Acid, C = Cysteine, E = Glutamic Acid, Q = Glutamine, G = Glycine, H = Histidine, I = Isoleucine, L = Leucine, K = Lysine, M = Methionine, F = Phenylalanine, P = Proline, S = Serine, T = Threonine, W = Tryptophan, Y = Tyrosine,V=Valine. HLHSAT and ADGF are antioxidant peptides obtained from hazelnut in other studies [22,23].
As predicated by us, six synthesized peptides showed a significant impact on retarding the oxidation of linoleic acid (Figure 1d). Peptides FSEY and NLNQCQRYM showed the best antioxidant activity, followed by QIESW, SEGFEW, IDLGTTY, and GEGFFEM. However, the inhibition of these synthesized peptides against linoleic acid oxidation functioned in a dose-dependent manner. Peptide NLNQCQRYM with a concentration higher than 900 μg/mL performed excellent activity in stopping linoleic acid from oxidation, while inhibition rates of the peptides QIESW and SEGFEW which contain Trp residue were less than 80% when their concentrations exceeded 200 μg/mL. It was noted that peptide AHSVVYAIR showed a poor capacity, and its IR% was only 50 even when its concentration was elevated to 5000 μg/mL (Figure 1 not shown). Our work indicated that peptides HLHSAT and ADGF, which have been reported to have ABTS and DPPH radical scavenging capacity [22,23] did not show any inhibition against the oxidation of linoleic acid. Moreover, the peptides that contain Cys residue showed perfect O2 •− scavenging activity compared to others, as these dipeptides did (see Figure 1c). Clearly, data from the artificially synthesized peptides support our assumption that the occurrence of active amino acid residues plays a crucial role in promoting the antioxidant capacity of a peptide. Featuring the properties of the amino acid residues should be a simple and feasible tool for the quick selection of desirable antioxidant peptides from hazelnut protein. Besides this, despite the excellent impact on retarding linoleic acid oxidation of FSEY and NLNQCQRYM, FSEY was selected for further study as the peptide falls within a molecular weight of 200 to 800 and GRAVY value of −2 to 1, as well as containing Tyr residue at the C-terminal.
| doab | 2025-04-07T03:56:59.190829 | 17-11-2022 17:23 | {
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007afdec-bed4-405d-873d-c355ba9add0e.42 | 3.5.1. Frontier Molecular Orbital Energy of Peptides
To convince the antioxidant performance of four active amino acids, 22 synthesized dipeptides were prepared, and their quantum chemical parameters were obtained by DFT calculations [38]. As shown in Table 2, the frontier molecular orbital energy of each dipeptide, expressed by EHOMO (Energy of highest occupied molecular orbital) and ELUMO (Energy of lowest unoccupied molecular orbital), represents the active level of 22 synthetics [26]. Theoretically, a higher EHOMO means more unstable electrons, which are more likely to scavenge free radicals as hydrogen donors. It was seen from Table 3 that the free radical-scavenging ability of these dipeptides, according to their calculated EHOMO, ranked from high to low in the order WH > WP > WY > WC > WD > MW > MH > YH > MY > YP > MP > CH > MD > MC > YD > IR > YC > KP > KD > AH > CP > CD. Moreover, a lower energy gap (E-gap) represents a higher chemical reactivity [39]. It means that the antioxidative activity of 22 dipeptides, based on their energy gap, ranked from strong to poor in the order WH > WC > WY > WP > WD > MW > MC > MH > MD > MY > YH > YC > YD > YP > CH > KD > CD > IR > AH > MP > KP > CP. To evaluate the real antioxidation of the 22 peptides, their inhibition towards linoleic acid oxidation was examined (see Figure 1b and Table 3). As a result, the best-synthesized dipeptide which retards the oxidation of linoleic acid was WY, followed by peptides MY, MW, YH, MH, MC, WC, YC, MD, WD, CP, YD, CH, CD, YP, MP, WH, KD, WP, AH, IR, and KP. It was noted that the dipeptides including KP, KD, IR, and AH with low EHOMO and high E-gap values had a low antioxidant capacity, whereas the dipeptides like WC, WY, MW, MC, MY, MH, and YH which have high EHOMO and low E-gap values showed a higher inhibition towards the oxidation of linoleic acid (see Table 3). Although several dipeptides had a poor activity in inhibiting the oxidation of linoleic acid, most of the synthesized dipeptides showed good antioxidant activity, especially these dipeptides containing W, M, or Y residue. It means that there is a corresponding relationship between the antioxidant activity of a dipeptide and its frontier molecular orbital energy.
**Table 3.** Frontier molecular orbital energy(eV) and GE (mM/mM) values of the dipeptides against the oxidation of linoleic acid.
To clarify the possible active site of each peptide, we constructed the dimensional structures of 22 synthesized dipeptides as well as FSEY. It is well known that HOMO always acts as the active site of any organic compound [40]. As shown in Figures 2 and 3a, the HOMOs of all dipeptides as well as FSEY are located at their active amino acid residues, that is, C, W, M, or Y. It indicates that these amino acids will firstly lose their electrons once when interacting with free radicals [41]. Clearly, data from the analysis of the HOMO site and linoleic acid oxidation inhibition confirmed the significance of active amino acid
residues in determining the activity of antioxidant peptides. The results from Hougland also et al. supported our findings [42].
**Figure 2.** HOMO distribution of the dipeptides. Note: red ball represents the oxygen atom; blue ball represents the nitrogen atom; dark gray ball represents the carbon atom; yellow ball represents the sulfur atom; light gray ball represents the hydrogen atom.
**Figure 3.** Quantum chemical parameters of FSEY. (**a**) shows HOMO distribution of peptide FSEY; (**b**) stands for Fukui functions; (**c**) means the predicting sites of FSEY more prone to a nucleophilic, electrophilic or radical attack. Note: red ball represents the oxygen atom; blue ball represents the nitrogen atom; dark gray ball represents the carbon atom; light gray ball represents the hydrogen atom.
| doab | 2025-04-07T03:56:59.191022 | 17-11-2022 17:23 | {
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007afdec-bed4-405d-873d-c355ba9add0e.43 | 3.5.2. Fukui Function
To predict which atom would be most susceptible to a nucleophilic or electrophilic attack, peptide FSEY was selected to investigate the tendency by Fukui functions. Fukui function is a local reactivity parameter, which is widely used for molecular reactivity analysis, indicating the tendency of a molecule to lose or gain an electron thus predicting which atom in the molecule would be more prone to a nucleophilic or electrophilic attack. When a molecule prefers to accept an electron, the Fukui function is f+, it is the index of nucleophilic attack. While when a molecule has a tendency to lose an electron, the Fukui function is f− and is also termed as the index of electrophilic attack [22]. In our study, the individual atomic charges were calculated by natural population analysis (NPA) with B3LYP/6-311G (d, p) basis set. For all atomic sites of peptide FSEY, their Fukui functions (f−, f+, f0) were presented in Figure 3b,c, respectively.
Blue, red, and green colors in Figure 3b represent nucleophilic, electrophilic, and radical attacks, respectively. It is found that nucleophilic, electrophilic, and radical attacks of peptide FSEY are located at N1, C13, C15, C51, C56, C57, C59, C61, C63, C65, and O66 atoms, especially at the atoms of Tyr residue. These results further highlighted a fact that Tyr residue should act as a biological activity site, as shown in Figure 3c. Moreover, the molecular reactivity site of peptide FSEY, as indicated by Fukui functions, totally corresponded to that of its HOMO.
## *3.6. Effects of Tyrosine Residue's Location on the Antioxidant Activity of Peptides*
To clarify how the position of tyrosine residue governs the antioxidant activity of a peptide against free radicals, three peptides FSEY, FYSE, and YFSE were selected to examine their inhibitions towards the oxidation of linoleic acid (see Figure 4a). The three peptides have the same composition of amino acids but have different locations for tyrosine residue. After incubated with linoleic acid at 40 ◦C for 48 h, the peptide FSEY showed the strongest ability in stopping the oxidation of the fat acid compared to others. The peptide YFSE had the lowest antioxidant activity. It is concluded that if tyrosine residue is located at the C-terminal, the Tyr-containing peptide should have a stronger activity against linoleic acid radical, as shown in Figure 4a.
**Figure 4.** Antioxidant activities of peptides. (**a**) stands for activities of FSEY, FYSE, and YFSE on the oxidation of linoleic acid; (**b**) represents the effects of the antioxidants on oil oxidation, GSH and TBHQ as positive controls.
| doab | 2025-04-07T03:56:59.191212 | 17-11-2022 17:23 | {
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007afdec-bed4-405d-873d-c355ba9add0e.44 | *3.7. Application of Hazelnut Peptide FSEY in Inhibiting Lipid Oxidation*
To validate the antioxidant ability of selected peptides in a real-emulsion, 0.02% hazelnut-original peptide FSEY was added to a hazelnut oil-in-water emulsion system for the evaluation of antioxidant activity against oil rancidity. It was seen that the hazelnutderived peptide FSEY inhibited the rancidity of oil very well by analyzing hydroperoxides on days 1, 3, 6, 10, and 14 (see Figure 4b). Furthermore, the antioxidant activity of peptide FSEY was compared with that of TBHQ, which is a commercial additive for protecting the oil from rancidity, as well as GSH, which is an antioxidant peptide (see Figure 4b). After incubation at 37 ◦C for 14 days, hydroperoxides of the emulsion system was 36.69 μmoL/g oil without antioxidant, whereas that was 16.94 μmoL/g oil, 22.35 μmoL/g oil, and 4.44 μmoL/g oil in the presence of FSEY, GSH, and TBHQ, respectively. It was clear that peptide FSEY showed a higher ability than GSH in controlling lipid oxidation, but lower activity than TBHQ. These results indicated that hazelnut-original peptide FSEY could be used as an antioxidant in the emulsion system for delaying the rancidity of oil. In addition, due to a weak ability in O2 •− scavenging and Fe2+ chelation, we speculate that the peptide FSEY act as a radical scavenger by contributing phenolic hydrogen atom to peroxyl radical.
## **4. Discussion**
Generally, antioxidant peptides against oil oxidation should act as one or more roles, that is, containing hydrophobic amino acids which expose more active sites to terminate lipid chain reaction; having free radical scavenging agents (such as O2 •− and peroxyl radical) or as metal ions chelating agents; and possessing strong lipase-inhibitory activities [2]. Obviously, the efficiency of antioxidation peptides in an emulsion system depends greatly on their ability to present more active sites, scavenge superoxide radicals or peroxyl radicals, and chelate metal ions. In addition, GSH was selected as a positive control for the outstanding antioxidant properties. Our study was pictured in Figure 5. As is shown, we used chemical experiments and physical properties of biopeptides as well as DFT calculations to verify which properties of the amino acid residues could be used to screen antioxidant peptides.
**Figure 5.** Screening antioxidant peptides depending on properties of amino acid residues.
| doab | 2025-04-07T03:56:59.191373 | 17-11-2022 17:23 | {
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007afdec-bed4-405d-873d-c355ba9add0e.45 | *4.1. Chemical Test: We Found Four Key Amino Acid Residues*
The radicals scavenging capacity of a peptide depends greatly upon its amino acid residues, especially upon Tyr, Trp, Met, and Cys. To illustrate this point, various tests firstly were carried out to confirm that Tyr, Trp, and Met and dipeptides containing these residues have excellent antioxidant capacities against the oxidation of linolic acid. It was observed that the absence of these residues caused the dipeptides to lose their activities (see Figure 1b). These amino acids, which are crucial in scavenging free radicals, have been reported by several research works. Amino acids Tyr, Trp, Cys, and Met as well as peptides containing these amino acids showed activities against ABTS radicals and oxygen radicals (ORAC, oxygen radical absorbance capacity) [43], active against ROS or RNS [3], and effectively against AAPH-induced peroxyl radicals [41].
| doab | 2025-04-07T03:56:59.191524 | 17-11-2022 17:23 | {
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007afdec-bed4-405d-873d-c355ba9add0e.46 | *4.2. DFT Calculation: Tyr, Trp, Met, and Cys Are Active Sites*
Theoretically, quantum chemical computations can gain prediction of behaviors of organic compounds, such as their structural features and chemical reactivity, and therefore, help to analyze the relationship between the biological potencies and the type of compounds [38]. The distributions of HOMO correspond to the active sites of the peptides able to scavenge free radicals [26]. By our DFT calculations, HOMOs of the tested peptides are located at their active amino acids, that is, Cys, Trp, Met, or Tyr (see Figure 2). The HOMOs of some peptides including EAAY, PMRGGGYHY, PMRGGYHY, PMRGYHY, PMRYHY, and YHY have been reported to be concentrated on the phenolic hydroxyl structure in Tyr [41]. The peptides PVETVR, QEPLLR, RDPEER, and LDDDGRL have the HOMOs of guanidyl in Arg, and the active sites of peptides KELEEK, DAAGRLQE, and GFAGDDAPRA are located at Lys-Glu, Gly, and Asp [24,39]. Clearly, data from HOMOs addressed that the residues Cys, Trp, Met, or Tyr are key components responsible for the antioxidant activity of the tested peptides in our study.
Generally, a high EHOMO or a low E-gap value means flexible chemical reactivity and could be used to predict the antioxidant activity of each peptide [24]. As predicted in our study, seven synthetic dipeptides, having a higher EHOMO and a lower E-gap value, showed a good ability to inhibit the oxidation of linoleic acid. It is found the seven antioxidant dipeptides possess the active residues Tyr, Trp, or Met. The presence of Tyr, Trp, and Met significantly enhanced the antioxidant activity of these dipeptides compared to other tested peptides (see Table 3). In a similar study, Wang et al. used EHOMO and E-gap to predicate the antioxidant activity of five peptides with only one exception [39]. Experiments conducted by Wu et al. also indicated that EHOMO and E-gap were feasible to describe the antioxidant behaviors of a set of man-made peptides, which were designed from the parent peptide "PMRGGGGYHY" [41]. Consistent with other studies reported [44], the presence of active residues Tyr, Trp, or Met as well as high EHOMO and low E-gap should be the characteristics of a peptide responsible for inhibiting the oxidation of linoleic acid. Amino acids, Tyr and Trp, act as active sites were also confirmed by Molecular docking. Wang et al. found that Trp1 and Tyr4 in peptide WLSYPMNPATGH could form hydrogen bonds with DPPH, which means responsible of Trp and Tyr in scavenging DPPH free radical. These two emerging approaches are helpful in analyzing antioxidative products, meanwhile, DFT calculation is a useful tool in screening antioxidant peptides [45].
| doab | 2025-04-07T03:56:59.191592 | 17-11-2022 17:23 | {
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007afdec-bed4-405d-873d-c355ba9add0e.47 | *4.3. BIOPEP Database Analysis: Rules of Molecular Weight, GRAVY Value and Active Amino Acid Residue's Location*
Few reports have focused on how a molecular weight, GRAVY value, and position of amino acid residues affect the capacity of a peptide against oil oxidation yet. In our study, a peptide in the BIOEPE database falling within a molecular weight of 200 to 800, GRAVY value of −2 to 1, and active amino acid residues at N- or C- terminus produced a strong inhibition towards lipid oxidation, as shown in Table S1 and Figure S1.
Regarding molecular weight or numbers of amino acid residues, numerous studies have indicated that peptides containing amino acid residues between 2 and 11 [46] or weighing less than 1000 Da [47] will exhibit good antioxidant ability. Peptides with more molecular weights (>2000 Da) easily decrease in their antioxidant activity due to the hiding of the active site [47]. It is well-known that the interfacial phase, the contact region between the oil phase and the aqueous phase, is the critical region in the system with regard to the development of lipid peroxidation [48]. Thus, in a given emulsion system, GRAVY value could not be ignored for antioxidant estimation, because a higher GRAVY means higher hydrophobicity. Various researches have proposed that peptides with higher hydrophobicity can protect linoleic acid from oxidation by donating protons to hydrophobic peroxy-radicals [49,50]. In our study, it is seen that peptides from BIOPEP database which able to stop lipid oxidation have GRAVY values ranging from −2 to 1 (see Table S1 and Figure S1b). Thus, a GRAVY value of −2 to 1 was proposed to be an ideal criterion for looking for antioxidant peptides from protein hydrolysate. A recent study has overviewed the roles of amino acid composition and sequence in conferring the antioxidant activities of peptides. The phenolic hydroxyl of Tyr, the indolyl of Trp, the thiol group of Cys, and the thioether of Met are regarded to act as hydrogen donors for free radicals [2,39,51]. Our study has found that the most of antioxidative peptides which are searched from the database BIOPEPE have the residues Tyr, Trp, Cys, or Met. Similarly, the antioxidant activities of peptides LGFEY and LGFYY were attributed to the presence of Tyr residues [52]. Concerning the inhibition of lipid oxidation, most peptides searched from the BIOPEPE database are observed to have active amino acid residues located at C- terminus. In our case, a linoleic acid oxidation system was designed to confirm the strong antioxidant activity of the peptide FSEY having Tyr residue located at the C-terminal (see Figure 4a). HOMO analysis from the synthesized peptides FYSE and YFSE also presents evidence that the reaction sites are all located at Tyr (see Figure S2). Similarly, studies done by Guo et al. [53] and Torkova et al. [54] indicated that peptides having Tyr residue located at the C-terminus strongly scavenged hydroxyl-radical, hydrogen-peroxide, and peroxyl radicals. While
these peptides exhibited a better inhibition against ABTS cation radical when Tyr residue is located at the N-terminus [54]. Clearly, the types and positions of amino acid residues should be considered in searching for an antioxidant peptide from protein.
In conclusion, a wanted antioxidant peptide could be quickly screened by determining the types and location of amino acid residues as well as molecular weight and GRAVY, especially Tyr, Trp, Cys, and Met which act as H donors. Among the four amino acids, Tyr-containing peptides show a prominent antioxidant activity, especially when it is located at the C-terminus.
| doab | 2025-04-07T03:56:59.191855 | 17-11-2022 17:23 | {
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007afdec-bed4-405d-873d-c355ba9add0e.48 | **5. Conclusions**
In our study, amino acids, Met, Tyr, Try, and peptides containing these active amino acids show antioxidant activity against linoleic acid radicals. These amino acid residues, as Tyr residue in peptide FSEY does, act as an active site for scavenging lipid free radicals. More meaningfully, the active amino acid residues located at C-terminal are more active than other positions. Compared to traditional technology for manufacturing bioactive peptides, our work presents a practical route able to successfully screen desirable highactivity antioxidant peptides from hazelnut protein hydrolysates by featuring the properties of amino acid residues. Our technical route consists of two steps. Firstly, peptides from hazelnut protein hydrolyzed by alkaline protease are sequenced; secondly, the peptides falling within a molecular weight of 200 to 800 and GRAVY value of −2 to 1 as well as containing Tyr, Met, Trp residues at C- terminus are selected for antioxidant candidates inhibiting the oxidation of the oil. Using this route successfully releases a peptide from hazelnut protein which inhibits oil oxidation very well. To our knowledge, it is the first attempt to prepare antioxidant peptides based on the properties of amino acid residues. Perhaps, the new findings out of our work will be beneficial for screening bioactive peptides from various protein resources with different purposes.
**Supplementary Materials:** The following are available online at https://www.mdpi.com/article/ 10.3390/antiox11010127/s1, Supplementary Materials to this article contains Table S1. Biopeptides collected from BIOPEP database and other publications, Figure S1. Analysis of the biopeptides harvested from BIOPEP database and other publications, Figure S2. The highest occupied molecular orbital (HOMO) of FYSE and YFSE.
**Author Contributions:** C.S., Writing—original draft, Methodology, Supervision. M.L., Investigation. H.Z., Funding acquisition. Z.L., Writing—review & editing. L.L. and B.Z., Supervision, Writing—review & editing. All authors have read and agreed to the published version of the manuscript.
**Funding:** This work was supported by the Fundamental Research Funds for the Central Universities (No. 2015ZCQ-SW-05).
**Institutional Review Board Statement:** Not applicable.
**Informed Consent Statement:** Not applicable.
**Data Availability Statement:** The data is contained within the article or supplementary material.
**Acknowledgments:** Authors acknowledge Fen Wang (College of Chemistry and Chemical Engineering, Taishan University, Taian, Shandong, China) for her great help in DFT calculations.
**Conflicts of Interest:** The authors declare no conflict 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. The author Lisong Liang is part of the National Innovation Alliance of Hazelnut Industry, the industry 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 | 2025-04-07T03:56:59.192086 | 17-11-2022 17:23 | {
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007afdec-bed4-405d-873d-c355ba9add0e.50 | *Article* **Improving Aqueous Solubility of Natural Antioxidant Mangiferin through Glycosylation by Maltogenic Amylase from** *Parageobacillus galactosidasius* **DSM 18751**
**Jiumn-Yih Wu 1,†, Hsiou-Yu Ding 2,†, Tzi-Yuan Wang 3,†, Yu-Li Tsai 4, Huei-Ju Ting <sup>4</sup> and Te-Sheng Chang 4,\***
**Citation:** Wu, J.-Y.; Ding, H.-Y.; Wang, T.-Y.; Tsai, Y.-L.; Ting, H.-J.; Chang, T.-S. Improving Aqueous Solubility of Natural Antioxidant Mangiferin through Glycosylation by Maltogenic Amylase from *Parageobacillus galactosidasius* DSM 18751. *Antioxidants* **2021**, *10*, 1817. https://doi.org/10.3390/ antiox10111817
Academic Editors: Li Liang and Hao Cheng
Received: 23 October 2021 Accepted: 15 November 2021 Published: 16 November 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/).
**Abstract:** Mangiferin is a natural antioxidant *C*-glucosidic xanthone originally isolated from the *Mangifera indica* (mango) plant. Mangiferin exhibits a wide range of pharmaceutical activities. However, mangiferin's poor solubility limits its applications. To resolve this limitation of mangiferin, enzymatic glycosylation of mangiferin to produce more soluble mangiferin glucosides was evaluated. Herein, the recombinant maltogenic amylase (MA; E.C. 3.2.1.133) from a thermophile *Parageobacillus galactosidasius* DSM 18751T (*Pg*MA) was cloned into *Escherichia coli* BL21 (DE3) via the expression plasmid pET-Duet-1. The recombinant *Pg*MA was purified via Ni2+ affinity chromatography. To evaluate its transglycosylation activity, 17 molecules, including mangiferin (as sugar acceptors), belonging to triterpenoids, saponins, flavonoids, and polyphenol glycosides, were assayed with β-CD (as the sugar donor). The results showed that puerarin and mangiferin are suitable sugar acceptors in the transglycosylation reaction. The glycosylation products from mangiferin by *Pg*MA were isolated using preparative high-performance liquid chromatography. Their chemical structures were glucosyl-*α*-(1→6)-mangiferin and maltosyl-*α*-(1→6)-mangiferin, determined by mass and nucleic magnetic resonance spectral analysis. The newly identified maltosyl-*α*-(1→6)-mangiferin showed 5500-fold higher aqueous solubility than that of mangiferin, and both mangiferin glucosides exhibited similar 1,1-diphenyl-2-picrylhydrazyl free radical scavenging activities compared to mangiferin. *Pg*MA is the first MA with glycosylation activity toward mangiferin, meaning mangiferin glucosides have potential future applications.
**Keywords:** mangiferin; maltogenic amylase; glycosylation; glucoside; *Parageobacillus galactosidasius*
| doab | 2025-04-07T03:56:59.192295 | 17-11-2022 17:23 | {
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007afdec-bed4-405d-873d-c355ba9add0e.51 | **1. Introduction**
Mangiferin is a natural *C*-glucosidic xanthone originally isolated from the *Mangifera indica* (mango) plant. Mangiferin has been reported to possess diverse health-promoting activities, such as antioxidant [1,2], anticancer [3,4], anti-inflammatory [5], and anti-osteoarthritis pain activities [6], allowing it to prevent memory impairment [7], neurodegeneration [8], and organ fibrosis [9]. Furthermore, it offers protection from the deleterious effects of heavy metals [10]. However, the pharmacological use of mangiferin is restricted owing to its poor solubility and low bioavailability [11,12]. As the glycosylates of small molecules have been proven to have better aqueous solubility and bioavailability than the original molecules [13,14], the glycosylation of mangiferin should be further improved for better usage.
Glycosylation of molecules can be achieved using chemical or enzymatic methods; however, enzymatic glycosylation using glycosyltransferases (GTs) and glycoside hydrolases (GHs) offers more advantages than chemical methods [15]. Moreover, GHs use cheaper sugars, such as starch, maltodextrin, maltose, and sucrose, as donors during glycosylation [16], whereas GTs use expensive uridine diphosphate-glucose (UDP-G). Therefore, GHs are preferred for the bioindustrial production of glycosylated molecules. According to the carbohydrate-activating enzyme (CAZy) database, a classification of GH in families based on amino acid sequence similarities and 117 GH families has been discovered to date [17].
Maltogenic amylase (MA; E.C. 3.2.1.133) belongs to the GH13 gene family and hydrolyzes starch to produce maltose [17]. Some specific features of MA were further identified. First, MAs were found to prefer cyclodextrin (CD) to starch as a substrate, whereas typical amylases do not catalyze CD. The sugar preference is due to 130 unique residues at the N-terminal of the MA protein, which would help the enzyme form a dimer and greatly increase its catalytic activities toward CD [18,19]. Second, MAs are intracellular proteins, whereas typical amylases are extracellular proteins. Third, MAs exhibit the bifunctions of hydrolysis and transglycosylation activities [18–28], whereas typical amylases are rarely reported to have transglycosylation activities [18].
Based on the dual functions of hydrolysis and transglycosylation and dual recognition sites on both the *α*-1,4 and *α*-1,6 glycosidic bonds, MAs have also been used in the fine chemical industry to produce novel and branched oligosaccharides from liquefied starch [19–28]. MAs could be further used for the glycosylation of bioactive molecules to develop new drugs in the clinical chemistry field.
In addition to sugars, MAs can glycosylate small and/or bioactive molecules. MAs exhibit transglycosylation reactions in the presence of various acceptor molecules, such as glucose, maltose, and acarbose, by forming *α*-1,3, *α*-1,4, and *α*-1,6 glycosidic linkages [18–28]. MAs have been proven to glycosylate some small molecules, such as hydroquinone [29], caffeic acid [30], ascorbic acid [31], puerarin [32–34], genistin [35], neohesperidin [36], and naringin [37]. For example, the glycosylation of puerarin by two MAs, *Tf*MA from archaeon *Thermofilum pendens* [32] and *Bs*MA from *Bacillus stearothermophilus* [34], has been studied.
In the present study, a maltogenic amylase gene from *Parageobacillus galactosidasius* (*Pg*MA) was cloned into *Escherichia coli* BL21 (DE3) via the expression plasmid pET-Duet-1, and the expressed *Pg*MA was purified. The purified *Pg*MA was characterized and found to glycosylate mangiferin. The novel mangiferin glucosides were isolated for the characterization of both the chemical structures and compounds.
| doab | 2025-04-07T03:56:59.192481 | 17-11-2022 17:23 | {
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007afdec-bed4-405d-873d-c355ba9add0e.53 | *2.1. Reagents and Chemicals*
A Ni2+ affinity column (10 i.d. × 50 mm, Ni Sepharose 6 Fast Flow) used for the purification of the recombinant MA was purchased from GE Healthcare (Chicago, IL, USA). Isopropyl β-D-1-thiogalactopyranoside (IPTG), 1,1-diphenyl-2-picrylhydrazine (DPPH), dimethyl sulfoxide (DMSO), and maltodextrin (dextrose equivalent 4.0–7.0) were bought from Sigma (St. Louis, MO, USA). *α*-CD, *β*-CD, *γ*-CD, soluble starch, and pullulan were purchased from Tokyo Chemical Industry Co., Ltd. (Tokyo, Japan). Restriction enzymes and DNA-modified enzymes were obtained from New England Biolabs (Ipswich, MA, USA). All kits for molecular cloning, including the Geno Plus Genomic DNA Extraction Midiprep System, Mini Plus Plasmid DNA Extraction System, Gel Advanced Gel Extraction Miniprep System, and Midi Plus Ultrapure Plasmid Extraction System, were purchased from Viogene (Taipei, Taiwan). Other reagents and solvents used are commercially available.
| doab | 2025-04-07T03:56:59.192718 | 17-11-2022 17:23 | {
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007afdec-bed4-405d-873d-c355ba9add0e.54 | *2.2. Strains and Plasmids*
*P. galactosidasius* DSM 18751T (BCRC 80657) was obtained from the Bioresources Collection and Research Center (BCRC; Food Industry Research and Development Institute, Hsinchu, Taiwan). *E. coli* BL21 (DE3) and the expression plasmid pET-Duet-1 were obtained from Novagen Inc. (Madison, WI, USA).
| doab | 2025-04-07T03:56:59.192798 | 17-11-2022 17:23 | {
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007afdec-bed4-405d-873d-c355ba9add0e.55 | *2.3. Aligned Amino Acid Sequences*
In total, 588 amino acids of *Pg*MA (OXB94089) and close-related BsMT (AAC46346) were aligned using Clustal W in MEGA X [38].
| doab | 2025-04-07T03:56:59.192841 | 17-11-2022 17:23 | {
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007afdec-bed4-405d-873d-c355ba9add0e.56 | *2.4. Construction of Expression Plasmids*
*P. galactosidasius* DSM 18751T (BCRC 80657) was cultivated in accordance with the BCRC protocol. The genomic DNA of the bacterium was isolated using the Geno Plus Genomic DNA Extraction Midiprep System (Viogene) according to the manufacturer's protocol. The target gene (*Pg*MA) was amplified from the genomic DNA with polymerase chain reaction (PCR). The primer set used in the PCR was as follows: forward 5 -gggggatccgttgaaagaagccatttatcatcg-3 and reverse 5 -gggctcgagtcaattttctacttgatagaggag-3 , which contain BamHI and XhoI restriction sites (underlined mark) for cloning. The amplified DNA fragment (1.8 kb length) was cloned into the expression plasmid pET-Duet-1, named pETDuet-*Pg*MA, which was then transformed into *E. coli* BL21 (DE3) for the recombinant *Pg*MA.
#### *2.5. Production and Purification of Recombinant PgMA in E. coli*
The recombinant *E. coli* harboring the recombinant expression plasmid pETDuet-*Pg*MA was cultivated in Luria–Bertani (LB) medium containing 1% (*w*/*v*) tryptone and sodium chloride and 0.5% (*w*/*v*) yeast extract to the optical density at 560 nm (OD560) of 0.6 and then induced with 0.2 mM of IPTG. After further cultivation at 18 ◦C for 20 h, the cells were centrifuged at 4500× *g* and 4 ◦C for 20 min. The cell pellet was washed and spun down twice with 50 mM of phosphate buffer (PB) at pH 6.8 and then broken with sonication via a Branson S-450D Sonifier (Branson Ultrasonic Corp., Danbury, CT, USA). The sonication program was run for five cycles of 5 s on and 30 s off at 4 ◦C. The mixture was then centrifuged at 15,000× *g* and 4 ◦C for 20 min to remove the cell debris. A supernatant containing the recombinant *Pg*MA fused with a His-tag in its N-terminal was applied in an Ni2+ affinity column. The His-tag-fused *Pg*MA was washed with PB with 25 mM imidazole and eluted with PB containing 250 mM imidazole. The eluate was then concentrated and desalted through Macrosep 10 K centrifugal filters (Pall, Ann Arbor, MI, USA). The concentration of the purified *Pg*MA was determined using the Bradford method [39] and analyzed with sodium dodecyl sulfate (SDS) polyacrylamide gel electrophoresis (PAGE). The purified *Pg*MA was stored in a final concentration of 50% glycerol at −80 ◦C before use.
| doab | 2025-04-07T03:56:59.192872 | 17-11-2022 17:23 | {
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007afdec-bed4-405d-873d-c355ba9add0e.57 | *2.6. Assay of Hydrolysis Activity*
The standard reaction was performed with 1% (*w*/*v*) β-CD, 5.6 μg/mL *Pg*MA, and 50 mM of PB at pH 6 and 60 ◦C for 30 min. After the reaction was stopped by boiling, the amount of reducing sugars produced from each reaction was estimated using the dinitrosalicylic acid method [40]. One unit of MA activity was defined as the amount of the enzyme that released 1 μmol of reducing sugar as maltose per min under the assay condition described earlier. For optimal conditions, the reaction was further performed at different temperatures and pH values, including pH 5 (acetate buffer), pH 6–7 (PB), and pH 8–10 (glycine buffer). Accordingly, the substrate specificity was measured with 1% (*w*/*v*) of the studied sugars, including α-CD, β-CD, γ-CD, soluble starch, and pullulan, performed at pH 7 and 65 ◦C. To realize the effects of metal ions and DMSO on the hydrolysis activity of *Pg*MA, 10 mM of tested metal ion or 5–20% (*v*/*v*) of DMSO was added into the reaction mixture.
| doab | 2025-04-07T03:56:59.193025 | 17-11-2022 17:23 | {
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007afdec-bed4-405d-873d-c355ba9add0e.58 | *2.7. Assay of Transglycosylation Activity*
To determine the transglycosylation activity of the *Pg*MA, β-CD was used as a sugar donor, and the molecules, which belonged to triterpenoids, saponins, flavonoids, or polyphenol glycosides, were tested as sugar acceptors. The reaction mixture containing 5% (*w*/*v*) β-CD, 5.6 μg/mL *Pg*MA, and 1 mg/mL tested molecules, dissolved in DMSO
with 50 mM of PB (pH 7), was incubated at 65 ◦C for 24 h. The reaction mixture was then mixed with an equal volume of methanol and analyzed with high-performance liquid chromatography (HPLC).
| doab | 2025-04-07T03:56:59.193112 | 17-11-2022 17:23 | {
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007afdec-bed4-405d-873d-c355ba9add0e.59 | *2.8. HPLC Analysis*
HPLC was performed with the Agilent 1100 series HPLC system (Santa Clara, CA, USA) equipped with a gradient pump (Waters 600, Waters, Milford, MA, USA). The stationary phase was a C18 column (5 μm, 4.6 i.d. × 250 mm; Sharpsil H-C18, Sharpsil, Bei-jing, China), and the mobile phase was 1% acetic acid in water (A) and methanol (B). The elution condition was a linear gradient from 0 min with 40% B to 20 min with 70% B, an isocratic elution from 20 to 25 min with 70% B, a linear gradient from 25 min with 70% B to 28 min with 40% B, and an isocratic elution from 28 to 35 min with 40% B. All eluants were eluted at a flow rate of 1 mL/min. The sample volume was 10 μL. The detection condition was set at 254 nm.
| doab | 2025-04-07T03:56:59.193163 | 17-11-2022 17:23 | {
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007afdec-bed4-405d-873d-c355ba9add0e.60 | *2.9. Purification of Mangiferin Glycosides*
The purification process was a previously described method [41]. A 100 mL reaction mixture containing 50% (*w*/*v*) maltodextrin, 1 mg/mL mangiferin, 5.6 μg/mL *Pg*MA, and 50 mM PB (pH 7) was incubated at 65 ◦C for 24 h. After the large-scale reaction, an equal volume of methanol was added to stop the transglycosylation. The mixture was then filtrated through a 0.2 μm nylon membrane, and the filtrate was injected in a preparative YoungLin HPLC system (YL9100, YL Instrument, Gyeonggi-do, South Korea) equipped with a preparative C18 reversed-phase column (10 μm, 20.0 i.d. × 250 mm, ODS 3; Inertsil, GL Sciences, Eindhoven, The Netherlands) for the purification of biotransformation products. The operational conditions for the preparative HPLC analysis were the same as those in the HPLC analysis. The elution corresponding to the peak of the metabolite in the HPLC analysis was collected, condensed under a vacuum, and then crystallized by freeze drying. Finally, 20.1 mg of compound (**1**) and 9.3 mg of compound (**2**) were obtained, and the structures of the compounds were confirmed with nucleic magnetic resonance (NMR) and mass spectral analyses. The mass analysis was performed using the Finnigan LCQ Duo mass spectrometer (ThermoQuest Corp., San Jose, CA, USA) with electrospray ionization (ESI). 1H- and 13C-NMR, distortionless enhancement by polarization transfer (DEPT), heteronuclear single quantum coherence (HSQC), heteronuclear multiple bond connectivity (HMBC), correlation spectroscopy (COSY), and nuclear Overhauser effect spectroscopy (NOESY) spectra were recorded on a Bruker AV-700 NMR spectrometer at ambient temperature. Standard pulse sequences and parameters were used for the NMR experiments, and all chemical shifts were reported in parts per million (ppm, *δ*).
The composition of compound (**1**) was as follows: light yellow powder; mp 233–235 ◦C; ESI/MS *m/z*: 583.4 [M-H]−, 565.3, 331.0, 300.9, 259.3; 1H-NMR (DMSO-*d6*, 700 MHz): H*δ* 3.05 (1H, t, *J* = 5.6 Hz, H-4), 3.15 (1H, d, *J* = 6.3 Hz, H-2), 3.20 (1H, t, *J* = 9.1 Hz, H-3 ), 3.29 (1H, t, *J* = 9.1 Hz, H-4 ), 3.33 (1H, m, H-5 ), 3.35 (1H, m, H-5), 3.38 (1H, m, H-3), 3.46 (1H, m, H-6a), 3.52 (1H, d, *J* = 9.1 Hz, H-6b), 3.62 (1H, d, *J* = 9.8 Hz, H-6 a), 3.70 (1H, dd, *J* = 11.2, 4.2 Hz, H-6 b), 4.02 (1H, br, H-2 ), 4.58 (1H, d, *J* = 9.1 Hz, H-1 ), 4.73 (1H, *J* = 4.2 Hz, H-1), 6.36 (1H, s, H-4), 6.86 (1H, s, H-5), and 7.37 (1H, s, H-8). 13C-NMR (DMSO-*d6*, 175 MHz): C*δ* 60.6 (C-6), 66.9 (C-6 ), 70.0 (C-4), 70.2 (C-2 , 4 ), 72.1 (C-2), 72.5 (C-3), 73.2 (C-1 ), 73.3 (C-5), 78.9 (C-3 ), 79.7 (C-5 ), 93.3 (C-4), 98.7 (C-1), 101.3 (C-9a), 102.6 (C-5), 107.5 (C-2), 108.1 (C-8), 111.8 (C-8a), 143.7 (C-7), 150.8 (C-10a), 154.0 (C-6), 156.2 (C-4a), 161.7 (C-1), 163.8 (C-3), and 179.1 (C-9).
The composition of compound (**2**) was as follows: light yellow powder; mp 227–229 ◦C; ESI/MS *m/z*: 745.3 [M-H]−, 727.3, 403.3, 385.0, 331.0, 313.2, 301.2; 1H-NMR (DMSO-*d6*, 700 MHz): H*δ* 3.03 (1H, t, *J* = 9.1 Hz, H-4), 3.19 (1H, dd, *J* = 9.1, 3.5 Hz, H-2), 3.21 (1H, m, H-3 ), 3.23 (1H, m, H-2), 3.30 (1H, m, H-4 ), 3.34 (1H, m, H-4), 3.36 (1H, m, H-5 ), 3.38 (1H, m, H-3), 3.41 (1H, br, H-2 ), 3.42 (1H, m, H-5), 3.44 (2H, m, H-6), 3.46 (1H, m, H-5), 3.55 (2H, m, H-6), 3.60 (1H, m, H-3), 3.64 (1H, m, H-6 a), 3.71 (1H, m, H-6 b), 4.58 (1H, d,
*J* = 9.8 Hz, H-1 ), 4.75 (1H, d, *J* = 3.5 Hz, H-1), 4.95 (1H, d, *J* = 3.5 Hz, H-1), 6.36 (1H, s, H-4), 6.86 (1H, s, H-5), and 7.37 (1H, s, H-8). 13C-NMR (DMSO-*d6*, 175 MHz): C*δ* 60.0 (C-6), 60.7 (C-6), 67.2 (C-6 ), 69.8 (C-4), 70.2 (C-2 , 4 ), 70.8 (C-5), 71.6 (C-2), 72.6 (C-2), 73.1 (C-3), 73.2 (C-1 ), 73.3 (C-3), 73.4 (C-5), 78.9 (C-3 ), 79.7 (C-5 ), 79.8 (C-4), 93.3 (C-4), 98.6 (C-1), 100.9 (C-1), 101.3 (C-9a), 102.6 (C-5), 107.4 (C-2), 108.1 (C-8), 111.7 (C-8a), 143.7 (C-7), 150.8 (C-10a), 154.0 (C-6), 156.2 (C-4a), 161.8 (C-1), 163.8 (C-3), and 179.1 (C-9).
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007afdec-bed4-405d-873d-c355ba9add0e.61 | *2.10. Determination of Aqueous Solubility*
The aqueous solubility of mangiferin and its glucoside derivative were examined as follows: each compound was vortexed in double-deionized H2O for 1 h at 25◦ C. The mixture was centrifuged at 10,000× *g* for 30 min at 25 ◦C. The supernatant was filtrated with 0.2 μM of nylon membrane and analyzed with HPLC. Based on their peak areas, the concentrations of the tested compounds were determined by using calibration curves prepared with HPLC analyses of authentic samples.
| doab | 2025-04-07T03:56:59.193530 | 17-11-2022 17:23 | {
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007afdec-bed4-405d-873d-c355ba9add0e.62 | *2.11. Determination of Antiradical Activity Using a DPPH Assay*
The assay was performed as previously described [42] with minor modifications. The tested sample (dissolved in DMSO) was added to the DPPH solution (1 mM in methanol) to a final volume of 0.1 mL. After 30 min of reaction, the absorbance of the reaction mixture was measured at 517 nm with a microplate reader (Sunrise, Tecan, Männedorf, Switzerland). Ascorbic acid (dissolved in DMSO) was used as a positive antioxidant standard. The DPPH free radical scavenging activity was calculated as follows: DPPH free radical scavenging activity = (OD517 of the control reaction − OD517 of the reaction)/(OD517 of the control reaction).
| doab | 2025-04-07T03:56:59.193588 | 17-11-2022 17:23 | {
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007afdec-bed4-405d-873d-c355ba9add0e.64 | *3.1. Selection of Candidate MAs via Online Genome Sequences*
Maltogenic amylase from *Bacillus stearothermophilus* (*Bs*MA, GenBank accession number: AAC46346) is a well-studied MA for the glycosylation of bioactive molecules [31,33,34,36,37]. Thus, the amino acid sequence of *Bs*MA was selected to search for new MAs from the NCBI GenBank. The highest but distinct MA sequences from bacterial whole genomes were further identified as our study's candidates. The available bacterial strains with known genome sequences in BCRC (Hsinchu, Taiwan) were also included for comparison. Accordingly, an α-glycosidase gene (GenBank accession number: OXB94089.1) from the genome data of *P. galactosidasius* DSM 18751T (GenBank accession number: PRJNA383662) showed the highest homology (79.1%) with *Bs*MA (Figure 1) and the top five candidates with the best-hit of *Bs*MA from NCBI GenBank in Table S1. Therefore, the α-glycosidase gene from the DSM 18,751 strain was identified as a suitable candidate in the present study. Figure 1 shows the α-glycosidase gene from *P. galactosidasius* DSM 18751, which was classified as a maltogenic amylase gene, and the gene product was named *Pg*MA in this study.
**Figure 1.** *Pg*MA (OXB94089) protein sequence. The GH13 family gene sequence identity between *Pg*MA and BsMT (AAC46346) is 79.08% (465/588). The Pfam domains are as follows: (1) alphaamylase\_N (alpha amylase, N-terminal IG-like domain; pfam02903): 1–121 amino acids; (2) alphaamylase (alpha amylase catalytic domain found in cyclomaltodextrinases and related proteins; cd11338): 173–469 amino acids; and (3) malt\_amylase\_C (maltogenic amylase, C-terminal domain; pfam16657), 507–583 amino acids. "." Denotes an identical amino acid; "-" denotes insertions and deletions.
| doab | 2025-04-07T03:56:59.193647 | 17-11-2022 17:23 | {
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007afdec-bed4-405d-873d-c355ba9add0e.65 | *3.2. Production of Recombinant PgMA in E. coli*
To produce the recombinant *Pg*MA, an expression plasmid pETDuet-*Pg*MA was constructed (Figure 2a), and the recombinant *Pg*MA was produced in recombinant *E. coli* BL21 (DE3) and purified as a major band shown by SDS-PAGE with a Ni2+ affinity chromatography (Figure 2b). The purified *Pg*MA showed an estimated 68 kD molecular weight in the SDS-PAGE. The production yield was 18.73 mg/L, and the specific activity of the purified enzyme was determined to be 91.46 U/mg by using β-CD as a substrate at pH 7 and 60 ◦C.
**Figure 2.** Expression of *Pg*MA from *Parageobacillus galactosidasius* DSM 18751T in *Escherichia coli* (DE3). (**a**) Recombinant repression plasmid pETDuet-*Pg*MA. (**b**) Sodium dodecyl sulfate-polyacrylamide gel electrophoresis (SDS-PAGE) analysis of expressed and purified proteins from recombinant *E. coli* harboring pETDuet-*Pg*MA. Lane 1: molecular marker; lane 2: total protein before induction; lane 3: total protein after 20 h of induction; and lane 4: purified protein.
| doab | 2025-04-07T03:56:59.193758 | 17-11-2022 17:23 | {
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007afdec-bed4-405d-873d-c355ba9add0e.66 | *3.3. Determination of Hydrolysis Activity by Recombinant PgMA*
To determine the optimal pH and temperature, β-CD was used as a substrate, and the reaction was performed at different pH levels and temperatures. The results showed that the optimal pH and temperature for the *Pg*MA catalytic reaction were pH 7 (Figure 3a) at 65 ◦C (Figure 3b). Moreover, the addition of Mg2+, K+, or ethylenediaminetetraacetic acid did not significantly affect the activity of *Pg*MA; by contrast, DMSO decreased the activity of *Pg*MA (Figure 3c).
**Figure 3.** Effects of pH (**a**), temperature (**b**), and metal ion or dimethyl sulfoxide (DMSO) (**c**) on the hydrolytic activity of *Pg*MA. The reaction was performed with 1% (*w*/*v*) β-cyclodextrin (CD), 5.6 μg/mL of *Pg*MA, and 50 mM of a different buffer at the tested temperature in the absence or presence of the tested metal ion or DMSO for 30 min. After the reaction was stopped by boiling, the hydrolytic activity of *Pg*MA was determined by measuring the reducing sugars produced from the reaction as described in Section 2.
One of the specific features of MA is that the enzyme prefers CD as its substrate over other polysaccharides, such as starch or pullulan. To characterize the *Pg*MA hydrolysis activity, different polysaccharides were used as a substrate for the *Pg*MA catalytic reaction. The results showed that *Pg*MA exhibited almost equally high specific activities toward α-CD, γ-CD, and β-CD. The specific activities of *Pg*MA toward CD were 65- and 650-fold higher than those toward pullulan and starch, respectively (Figure 4). This CD preference was consistent with other known MAs [20–27]. The 130 residues at the N-terminal of the MA are key for the enzymes to form dimers and largely increase hydrolysis activities toward CD [18,19]. In addition, some recombinant MAs have been purified using Nterminal His-tag fusion [18–27]. The N-terminal His-tag fusion did not seem to affect its dimerization; herein, the recombinant *Pg*MA also remained as the CD preference.
**Figure 4.** Substrate specificity of the hydrolytic activity of *Pg*MA. The reaction was performed with 1% (*w*/*v*) of the tested sugar substrate, 5.6 μg/mL *Pg*MA, and 50 mM of PB at pH 7 and 65 ◦C for 30 min. The hydrolytic activity of *Pg*MA was determined as described in the legend of Figure 3.
| doab | 2025-04-07T03:56:59.193827 | 17-11-2022 17:23 | {
"license": "Creative Commons - Attribution - https://creativecommons.org/licenses/by/4.0/",
"book_id": "007afdec-bed4-405d-873d-c355ba9add0e",
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"title": "Characterization and Encapsulation of Natural Antioxidants: Interaction, Protection and Delivery",
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007afdec-bed4-405d-873d-c355ba9add0e.67 | *3.4. Determination of Transglycosylation Activity by Recombinant PgMA*
Transglycosylation activity is an important property of MAs for biotechnology applications. To clarify the transglycosylation activity of recombinant *Pg*MA, 17 different molecules, including mangiferin (Table S2), belonging to triterpenoids, saponins, flavonoids, flavonoid glycosides, or xanthone glycoside, were used as sugar acceptors with 1% (*w*/*v*) β-CD (as the sugar donor) for activity. The reaction mixture was then analyzed with HPLC. The results showed that only puerarin (Figure 5a) and mangiferin (Figure 5b) could be glycosylated by *Pg*MA.
**Figure 5.** High-performance liquid chromatography (HPLC) analysis of the biotransformation products of puerarin (**a**) and mangiferin (**b**) by *Pg*MA. The reaction was performed with 1% (*w*/*v*) β-CD, 5.6 μg/mL of *Pg*MA, and 1 mg/mL of puerarin or mangiferin at 50 mM of PB (pH 7) and 65 ◦C for 24 h. After the reaction, the reaction mixture was analyzed with HPLC. The conditions for HPLC are described in Section 2.
Except mangiferin and puerarin, the other four tested triterpenoids, two triterpenoids saponins, nine flavonoid aglycones, and glycosides could not act as the sugar acceptors in the transglycosylation of *Pg*MA. *Pg*MA could transglycosylate puerarin, which has the isoflavone-8-*C*-glucosdie structure. However, *Pg*MA could not transglycosylate isoflavone-7-*O*-glucoside (8-hydroxydaidzein-7-*α*-*O*-glucoside) or flavone-8-*C*-glucoside (vitexin). The results imply that *Pg*MA has a narrow and/or specific substrate range. Nevertheless, the
main finding is that *Pg*MA can glycosylate mangiferin, which will expand the biotechnological applications of MAs in the future. MAs have been proven to glycosylate some small molecules, such as hydroquinone [29], caffeic acid [30], ascorbic acid [31], puerarin [32–34], genistin [35], neohesperidin [36], and naringin [37]. Our results also showed that *Pg*MA glycosylated puerarin to three major products, P1, P2, and P3 (Figure 5a). These three major products were not identified in advance because the glycosylation of puerarin has been studied based only on known MAs [32,34]. Li et al. (2004) reported that *Bs*MA glycosylated puerarin to three products (T1, T2, and T3), two of which were identified as maltosyl-α-(1→6)-puerarin (T1) and glucosyl-α-(1→6)-puerarin (T2), while T3 was not identified [34]. Li et al. (2011) further reported that a maltogenic amylase (*Tf*MA) from the archaeon *T. pendens* glycosylated puerarin to a series of products containing glucosyl puerarin and maltosyl puerarin, although they did not identify the exact chemical structures of the products [32]. From the results of the two studies, the P1–P3 products might contain glucosyl and maltosyl puerarin.
The results revealed that *Pg*MA glycosylates mangiferin to produce low amounts of the M1 compound with a yield of 2.3% (Figure 5b). This is the first study to report that MA could glycosylate mangiferin, of which mangiferin glycoside may have better aqueous solubility for different applications. Therefore, we mainly focused on the unknown mangiferin glycoside by *Pg*MA in the following assays.
| doab | 2025-04-07T03:56:59.193956 | 17-11-2022 17:23 | {
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007afdec-bed4-405d-873d-c355ba9add0e.68 | *3.5. Optimization of Biotransformation of Mangiferin by PgMA*
As the yield of the M1 compound from the biotransformation by *Pg*MA is too low to be easily purified, the glycosylation condition must be optimized. The glycosylation reaction was optimized with different sugar donors, concentrations of sugar donors, and reaction times. First, although the yield of the M1 using α-CD showed the best output (Figure 6a), maltodextrin was selected as the sugar donor for experiments due to its highly aqueous solubility at a much lower price.
Second, the GH enzymes contained both hydrolysis and transglycosylation activities. The transglycosylation activity of GH has been reported to increase under low water concentrations [43]. Thus, a solution with a higher sugar concentration and lower water concentration would increase its transglycosylation activity. The transglycosylation activity of *Pg*MA was indeed increased in 50% maltodextrin, the highest soluble concentration. When the maltodextrin concentration was increased from 1% to 50% (*w*/*v*), three compounds (M1, M2, and M3 in Figure 6b) were formed with higher yields (Figure 6c). The maximal yields of M1 and M2 reached 10% and 21%, respectively. The M3 compound was not completely separated with mangiferin, which is similar to the situation of the T3 compound with puerarin by *Bs*MA [34]. Therefore, only M1 and M2 were further studied.
**Figure 6.** *Cont.*
**Figure 6.** Effects of the sugar donor on the glycosylation of mangiferin by *Pg*MA. (**a**) Different sugar donors were used in the glycosylation of mangiferin. (**b**) HPLC analysis of the glycosylation mixture of mangiferin by *Pg*MA with 50% (*w*/*v*) maltodextrin. (**c**) Different maltodextrin concentrations were used in the glycosylation of mangiferin. The reaction was performed with 5.6 μg/mL *Pg*MA, 1 mg/mL mangiferin, and the tested sugar donors at 50 mM of PB (pH 7) and 65 ◦C for 24 h. After the reaction, the reaction mixture was analyzed with HPLC. The conditions for HPLC are described in Section 2. The yield of the product was calculated by dividing the area of the product by that of mangiferin without an enzyme reaction (control reaction).
> Third, the yields of M1 and M2 in 50% maltodextrin by *Pg*MA were further determined under different time courses (Figure 7). The results showed that the yields plateau of M1 and M2 reached 13.2% at 168 h and 33.8% at 72 h, respectively.
**Figure 7.** Time course of the glycosylation of mangiferin by *Pg*MA. The reaction was conducted with 50% (*w*/*v*) maltodextrin, 5.6 μg/mL *Pg*MA, and 1 mg/mL mangiferin at 50 mM of PB (pH 7) and 65 ◦C. At the interval time, the reaction mixture was analyzed with HPLC. The conditions for HPLC and the calculation of the yield were the same as those described in the legend to Figure 6.
| doab | 2025-04-07T03:56:59.194115 | 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.69 | *3.6. Isolation and Identification of Mangiferin Glycosides by PgMA*
The glycosylation of mangiferin by *Pg*MA was scaled up to 100 mL. The products M1 and M2 were purified using preparative HPLC. From the 100 mL reaction, 20.1 mg of compound (**1**) (M1) and 9.3 mg of compound (**2**) (M2) were isolated. The molecular weights of the purified products were then determined with mass spectrometry. The mass spectrometry of compound (**1**) revealed an [M–H]− ion peak at *m/z*: 583.4 in the electrospray ionization mass spectrum (ESI-MS) corresponding to the molecular formula C25H28O16 (Figure S1). The mass data imply that M1 contains one glucosyl moiety attached to the mangiferin structure. In the mass data of M2, an [M–H]− ion peak at *m/z*: 745.3 in the ESI-MS corresponded to the molecular formula C31H38O21 (Figure S2), which implies that compound (**2**) contains two glucosyl moieties attached to the mangiferin structure. To identify the structures in advance, the structures of both compounds were determined using NMR spectroscopy. 1H and 13C NMR, including the DEPT, HSQC, HMBC, COSY, and NOESY spectra, were obtained.
The characteristic 1H and 13C NMR sugar signals in compound (**1**) were assigned to *C*-glucosyl and *O*-glucosyl moieties by one-dimensional (1-D) and 2-D NMR experiments. The 1H spectrum of compound (**1**) in DMSO-*d6* showed three singlets at 6.36, 6.86, and 7.37 ppm and a complex 10-spin system between 3.0 and 5.0 ppm. Analysis of this secondorder system revealed coupling constants typical of two glucose moieties. The compound (**1**) glucosidic linkage of the *C*-glucosyl moiety on the xanthone C-2 was revealed by the presence of HMBC correlations between C-2/H-1 (107.5/4.58 ppm), and the anomeric proton H-1 at 4.58 (d, *J* = 9.1 Hz) indicated a *C*-*β*- configuration of mangiferin that was confirmed by the data reported in the literature [44]. The mangiferin *O*-glucosyl moiety was a doublet signal at H-1 (4.73 ppm, d, *J* = 4.2 Hz) with the corresponding carbon atom at C-1 (98.7 ppm) assigned to the anomeric proton and indicating an *O*-*α*-configuration by HSQC, which is in the *O*-*α*-configuration. The H-1 (*δ* = 4.73 ppm) of mangiferin and the HMBC cross signaled H-1/C-6 (4.73/66.9 ppm) and H-6 a, 6 b/C-1 (3.46, 3.52/98.7 ppm). The significant downfield shift of the 13C signal of C-6 indicated the connection of the second glucosyl moiety, which confirmed the *α*-(1→6) linkage of the second glucosyl moiety. The NMR signals were identified as shown in Table S3. The compound (1) was thus confirmed as glucosyl-*α*-(1→6)-mangiferin (Figures S3–S9).
The 1H spectrum of compound (**2**) in the same compound (**1**) solvent also showed three singlets at 6.36, 6.86, and 7.37 ppm and a complex 11-spin system between 3.0 and 5.0 ppm. Analysis of this second-order system revealed coupling constants typical of three glycose moieties, which included the chemical shifts listed in Table S3. The glucosyl moiety chemical shifts of C-2 at 107.4 ppm and H-1 at 4.58 ppm (d, *J* = 9.8 Hz) according to the corresponding HMBC indicated a C-C bond between the sugar and the aglycone of mangiferin (*C*-glucosyl-xanthone) and were confirmed by the data reported in the literature [44]. The *O*-maltosyl moiety connected to mangiferin was confirmed by HMBC from the anomeric carbon C-6 (66.9 ppm), and the corresponding anomeric proton H-1 at 4.75 (d, *J* = 3.5 Hz) indicated an *O*-*α*-configuration. The maltose doublet signal at *δ*<sup>H</sup> H-1 (d, *J* = 3.5 Hz) and H-1 4.95 (d, *J* = 3.5 Hz) with the corresponding carbon atom at C-1 (98.6 ppm) and C-1 (100.9 ppm) was assigned to the anomeric proton and indicated two *O*-*α*-configurations by HSQC. The HMBC cross peaks of C-1/H-6 (98.6/3.64, 3.71 ppm) and C-1/H-4 (100.9/3.34 ppm) confirmed the *α*-(1→4) between the two-glucosyl moiety. Our experimental 1H and 13C chemical shifts listed in Table S2 confirmed compound (**2**) as maltosyl-*α*-(1→6)-mangiferin (Figures S10–S16). Figure 8 summarizes the biotransformation process of mangiferin by *Pg*MA.
| doab | 2025-04-07T03:56:59.194266 | 17-11-2022 17:23 | {
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007afdec-bed4-405d-873d-c355ba9add0e.70 | *3.7. Characterizations of Mangiferin Glucosides*
The low aqueous solubility of mangiferin restricts its usage as a pharmaceutical agent. However, the glycosylation of mangiferin may mitigate such a restriction. Only a few studies have reported different glycosylation agents for mangiferin. Wu et al. (2013) used β-fructofuranosidase (E.C. 3.2.1.26; GH 32 family) to glycosylate mangiferin into fructosylβ-(2→6)-mangiferin and found that its DPPH radical scavenging activity was similar to that of mangiferin [45]. Nguyen et al. (2020) used dextransucrase (E.C. 2.4.1.5; GH 70 family) from *Leuconostoc mesenteroides* to glycosylate mangiferin into glucosyl-α-(1→6)-mangiferin (**1**) [46]. They found that the aqueous solubility of glucosyl-*α*-(1→6)-mangiferin (**1**) was 2300-fold higher than that of mangiferin. In this study, the amount of purified glucosyl-*α*- (1→6)-mangiferin (**1**) was too low to repeat the solubility experiment. The solubility of the newly identified maltosyl-*α*-(1→6)-mangiferin (**2**) was determined. The results showed that the solubility of maltosyl-*α*-(1→6)-mangiferin (**2**) was 5500-fold higher than that of mangiferin (Table 1). Thus, maltosyl-*α*-(1→6)-mangiferin (**2**) possesses higher solubility than glucosyl-*α*-(1→6)-mangiferin (**1**). It has been reported that the more sugar moieties in the glycosylated compounds, the higher the aqueous solubility of the glycosylated compounds [29–37].
**Figure 8.** Biotransformation process of mangiferin by *Pg*MA.
**Table 1.** Aqueous solubility of mangiferin and maltosyl-*α*-(1→6)-mangiferin (**2**).
<sup>1</sup> The mean (*n* = 2) is shown, and the standard deviations are represented by error bars. <sup>2</sup> The folds of the aqueous solubilities of the mangiferin glucoside derivatives are expressed as relative to that of mangiferin normalized to 1.
Mangiferin exhibits a wide pharmacological profile, and its antioxidant property is well known from previous studies. Furthermore, it has been associated with the redox aromatic system of the xanthone nucleus [3–10,12]. Thus, the antioxidative activities of the two mangiferin glucosides were determined using DPPH free radical scavenging assay. The assay showed that the antioxidant activity levels of mangiferin and its two glucosides were all higher than those of ascorbic acid (Figure 9). In other words, the antioxidant activities of the two mangiferin glucosides are comparable with those of mangiferin. The *ortho*-dihydroxyl groups on the benzene ring of the mangiferin structure have been reported to play a key role in exerting its antioxidant activity [1,2]. Both mangiferin glucosides remained the key functional groups after glycosylation; therefore, most of the antioxidant activity remained in the mangiferin derivatives. These glycoside derivatives (glucoside and fructoside) might possess different pharmacological properties. A futher study will focus on the bioactivities and bioavailability of these mangiferin derivatives
**Figure 9.** The 1,1-diphenyl-2-picrylhydrazyl (DPPH) free radical scavenging activity of mangiferin, mangiferin glucosides, and ascorbic acid. The DPPH scavenging activity was determined as described in Section 2. The IC50 values represent the concentrations required for 50% DPPH free radical scavenging activity. The mean (*n* = 3) is shown, and the standard deviations are represented by error bars. M1 and M2 are glucosyl-*α*-(1→6)-mangiferin and maltosyl-*α*-(1→6)-mangiferin, respectively.
| doab | 2025-04-07T03:56:59.194489 | 17-11-2022 17:23 | {
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007afdec-bed4-405d-873d-c355ba9add0e.71 | **4. Conclusions**
The recombinant *Pg*MA from *P. galactosidasius* DSM 18751<sup>T</sup> was confirmed to exhibit the bifunctions of hydrolysis and transglycosylation activities. The recombinant *Pg*MA can glycosylate mangiferin and produce glucosyl-*α*-(1→6)-mangiferin and maltosyl-*α*- (1→6)-mangiferin with a high maltodextrin concentration. The novel maltosyl-*α*-(1→6) mangiferin showed much higher aqueous solubility than that of mangiferin. The two mangiferin glucosides exhibited similar DPPH antioxidative activity compared to mangiferin. To our knowledge, *Pg*MA is the first MA identified with glycosylation activity toward mangiferin. With higher water solubility and compatible antioxidant activity, the two mangiferin glucoside derivatives have better pharmaceutical applicability.
**Supplementary Materials:** The following materials are available online at https://www.mdpi.com/ article/10.3390/antiox10111817/s1. Table S1: Top five candidates with best-hit of *Bs*MA from NCBI GenBank. Table S2. The list of the tested molecules in the transglycosylation reaction of *Pg*MA. Table S3: 1H and 13C NMR assignments in DMSO-*d6* at 700 and 175 MHz for compounds (**1**) and (**2**). Figure S1: Mass–mass analysis of glucosyl-*α*-(1→6)-mangiferin (**1**) in the negative mode. Figure S2: Mass–mass analysis of maltosyl-*α*-(1→6)-mangiferin (**2**) in the negative mode. Figure S3: Onedimensional NMR spectrum (1H-NMR, 700 MHz, DMSO-*d6*) of the glucosyl-*α*-(1→6)-mangiferin (**1**). Figure S4: One-dimensional NMR spectrum (13C-NMR, 175 MHz, DMSO- *d6*) of the glucosyl-*α*- (1→6)-mangiferin (**1**). Figure S5: One-dimensional NMR spectrum (DEPT-135, 175 MHz, DMSO-*d6*) of the glucosyl-*α*-(1→6)-mangiferin (**1**). Figure S6: Two-dimensional NMR spectrum (1H-13C HSQC, 700 MHz, DMSO-*d6*) of the glucosyl-*α*-(1→6)-mangiferin (**1**). Figure S7: Two-dimensional NMR spectrum (1H-13C HMBC, 700 MHz, DMSO-*d6*) of the glucosyl-*α*-(1→6)-mangiferin (**1**). Figure S8: Two-dimensional NMR spectrum (1H-1H COSY, 700 MHz, DMSO-*d6*) of the glucosyl-*α*-(1→6) mangiferin (**1**). Figure S9: Two-dimensional NMR spectrum (1H-1H NOESY, 700 MHz, DMSO-*d6*) of the glucosyl-*α*-(1→6)-mangiferin (**1**). Figure S10: One-dimensional NMR spectrum (1H-NMR, 700 MHz, DMSO-*d6*) of the maltosyl-*α*-(1→6)-mangiferin (**2**). Figure S11: One-dimensional NMR spectrum (13C-NMR, 175 MHz, DMSO-*d6*) of the maltosyl-*α*-(1→6)-mangiferin (**2**). Figure S12: Onedimensional NMR spectrum (DEPT-135, 175 MHz, DMSO-*d6*) of the maltosyl-*α*-(1→6)-mangiferin (**2**). Figure S13: Two-dimensional NMR spectrum (1H-13C HSQC, 700 MHz, DMSO-*d6*) of the maltosyl*<sup>α</sup>*-(1→6)-mangiferin (**2**). Figure S14: Two-dimensional NMR spectrum (1H-13C HMBC, 700 MHz, DMSO-*d6*) of the maltosyl-*α*-(1→6)-mangiferin (**2**). Figure S15: Two-dimensional NMR spectrum (1H- 1H COSY, 700 MHz, DMSO-*d6*) of the maltosyl-*α*-(1→6)-mangiferin (**2**). Figure S16: Two-dimensional NMR spectrum (1H-1H NOESY, 700 MHz, DMSO-*d6*) of the maltosyl-*α*-(1→6)-mangiferin (**2**).
**Author Contributions:** Conceptualization: T.-S.C.; data curation and methodology: Y.-L.T., T.-S.C. and H.-Y.D.; project administration: T.-S.C. and J.-Y.W.; writing—original draft, review, and editing: T.-S.C., T.-Y.W., J.-Y.W., H.-J.T. and H.-Y.D. All authors have read and agreed to the published version of the manuscript.
**Funding:** This research was funded by the Ministry of Science and Technology of Taiwan under grant number MOST 110-2221-E-024-002 to T.-S.C. and grant number MOST 110-2221-E-507-002 to J.-Y.W.
**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 or supplementary material.
**Conflicts of Interest:** The authors declare no conflict of interest.
| doab | 2025-04-07T03:56:59.194689 | 17-11-2022 17:23 | {
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007afdec-bed4-405d-873d-c355ba9add0e.73 | *Article* **The Fine-Tuned Release of Antioxidant from Superparamagnetic Nanocarriers under the Combination of Stationary and Alternating Magnetic Fields**
**Lucija Mandi´c, Anja Sadžak , Ina Erceg, Goran Baranovi´c and Suzana Šegota \***
Ruder Boškovi´ ¯ c Institute, 10000 Zagreb, Croatia; [email protected] (L.M.); [email protected] (A.S.); [email protected] (I.E.); [email protected] (G.B.)
**\*** Correspondence: [email protected]
**Abstract:** Superparamagnetic magnetite nanoparticles (MNPs) with excellent biocompatibility and negligible toxicity were prepared by solvothermal method and stabilized by widely used and biocompatible polymer poly(ethylene glycol) PEG-4000 Da. The unique properties of the synthesized MNPs enable them to host the unstable and water-insoluble quercetin as well as deliver and localize quercetin directly to the desired site. The chemical and physical properties were validated by X-ray powder diffraction (XRPD), field emission scanning electron microscopy (FE–SEM), atomic force microscopy (AFM), superconducting quantum interference device (SQUID) magnetometer, FTIR spectroscopy and dynamic light scattering (DLS). The kinetics of in vitro quercetin release from MNPs followed by UV/VIS spectroscopy was controlled by employing combined stationary and alternating magnetic fields. The obtained results have shown an increased response of quercetin from superparamagnetic MNPs under a lower stationary magnetic field and s higher frequency of alternating magnetic field. The achieved findings suggested that we designed promising targeted quercetin delivery with fine-tuning drug release from magnetic MNPs.
**Keywords:** release kinetics; superparamagnetism; stationary magnetic field; alternating magnetic fields; magnetic nanoparticles; quercetin
| doab | 2025-04-07T03:56:59.194861 | 17-11-2022 17:23 | {
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007afdec-bed4-405d-873d-c355ba9add0e.74 | **1. Introduction**
Quercetin (C15H10O7, 3,3 ,4 ,5,7-pentahydroxyflavone) is a major member of the flavonols, a subclass of flavonoids, natural polyphenols [1]. It is an important component of the human's daily diet and widely distributed in vegetables and fruits such as onions, tomatoes, berries, grapes, nuts, as well as in many flowers and leaves [2,3]. In addition, quercetin exhibits a wide range of biological and pharmacological activities, including antioxidant, anti-inflammatory, antibacterial, anti-anaemic, and anticarcinogenic activities [1,3–5]. Extensive studies have reported that quercetin can inhibit the proliferation of several types of cancers such as lung, prostate, breast cancer, and pancreatic tumour cells [1]. In addition, the main feature of quercetin is its antioxidant potential of OH groups in the structure that can bind to reactive oxygen species (ROS) and maintain cell viability. Quercetin has been shown to decrease the activity of antioxidant and apoptotic proteins and increase the levels of antiapoptotic proteins [6]. However, its therapeutic and clinical properties are limited due to its hydrophobic nature and low stability in the physiological medium. The problem with instability, low solubility and poor bioavailability can be successfully overcome by their loading in drug delivery systems including nanoparticles (NPs) [3,5,7,8]. The rapid growth of nanotechnology is the key to a revolutionary platform for chemical, physical, biological and mechanical properties of various materials [9,10]. There is tremendous interest in nanomaterials or NPs in the biomedical field [11]. For example, a variety of NPs is envisioned to be used in medical applications such as cancer detection, magnetic resonance imaging, cardiovascular and neurological treatment diseases,
**Citation:** Mandi´c, L.; Sadžak, A.; Erceg, I.; Baranovi´c, G.; Šegota, S. The Fine-Tuned Release of Antioxidant from Superparamagnetic Nanocarriers under the Combination of Stationary and Alternating Magnetic Fields. *Antioxidants* **2021**, *10*, 1212. https://doi.org/10.3390/ antiox10081212
Academic Editors: Li Liang and Hao Cheng
Received: 15 June 2021 Accepted: 26 July 2021 Published: 28 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/).
targeted drug delivery, hyperthermia, bioseparation, and gene transfer [3,9,10]. In recent years, magnetic nanoparticles (MNPs) have been highlighted among the many types of NPs [12–14]. Essentially, researchers were attracted by their excellent unique physical, chemical and magnetic properties [15]. In 1957, Gilchrist and coworkers showed the first use of magnetic particles for inductive heating of lymph nodes in dogs [16,17].
In 1983, Widder and coworkers reported the first use of MNPs containing doxorubicin to treat Yoshida rat sarcoma with an external magnetic field. These results represented a compelling advance in chemotherapy treatment, with complete cancer remission demonstrated in 77% of animals in the magnetically localized doxorubicin-magnetite microparticles [17,18]. MNPs have received much attention as target drugs that can replace traditional chemotherapy without the side effects [19]. Several inorganic magnetic nanoparticles (MNPs) have the potential to be used for drug delivery, but MNPs are the only magnetic materials approved by the Food and Drug Administration (FDA) for human use [3,19–21]. Numerous physical, biological and chemical preparation methods have been accepted for the MNP's synthesis [22]. Magnetite NPs are most commonly used in biological applications due to their unique physicochemical properties such as particle size, size distribution, shape and high surface area [23,24]. They exhibit interesting properties such as superparamagnetism, high field irreversibility and high saturation field [24–26]. Due to these properties, the superparamagnetic NPs can become magnetized when the external magnetic field is used and they do not remain magnetized when the field is turned off. The localization of drugs with MNPs in combination with an external magnetic field and their retention until the completion of therapy [26] represent a promising strategy of drug delivery with the controlled release [27]. The effectiveness of magnetic delivery systems includes the field strength, gradient, magnetic and physicochemical properties of the NPs [28]. Moreover, the main challenge of bare MNPs is to avoid their agglomeration due to their van der Waals and magnetic dipole–dipole attraction forces [13,29]. Considering their hydrophobic surface and rapid clearance from the blood through the reticuloendothelial system (RES), they are not suitable for drug delivery systems [22]. Therefore, to overcome this inconvenience, it was necessary to coat the magnetite NPs to reduce the aggregation tendency, protect their surface from oxidation, and make the particles biocompatible and stable [30].
Various polymers have been used for drug delivery, with polyethylene glycol (PEG) being the gold standard and the most commonly used polymer [31]. PEG is approved by the Food and Drug Administration (FDA) for internal use in humans and its products have been on the market for 20 years [23,31]. Since 1994, Gref and co-workers have reported on the PEG coating and demonstrated that the naked particles were removed from the liver only 5 min after injection [31]. PEG coating is extensively used in the preparation of NPs for biomedical applications due to its many advantages, such as stability in physiological media, prolonged half-life in the body, biocompatibility, and water solubility. Moreover, PEG coating prevents or reduces aggregation and confers better physical stability to drugs through steric and hydric repulsion [19,31].
In this study, we synthesized MNPs, which are known to have excellent biocompatibility and negligible toxicity [26], allowing their application in therapy. Prepared by the solvothermal method and stabilized by the widely used PEG, MNPs possess unique properties, such as colloidal stability, dispersibility, high porosity, high loading capacity, and specific morphological, thermal, and magnetic properties, especially superparamagnetism, which enable them to host the unstable or water-insoluble drugs and to direct and localize the drugs to the specific site in the tissue. The synthesized MNPs were fully characterized in terms of structural, morphological and magnetic properties. In addition, the kinetics of quercetin from the MNPs were controlled in vitro by simply varying stationary and alternating magnetic fields, resulting in fine-tuned manipulation of the released quercetin as a model drug. It should be noted that in this study cytotoxicity has been considered, but it was not a priority at this stage of our research. Regarding the measurement and result in the cytotoxicity literature of MNPs, we quite rightly expected at least the same cytotoxicity
as the results obtained by Barreto et al. [3], Hua et al. [32] and Luo et al. [33]. It was shown that the system developed provides prolonged quercetin release, which is an important characteristic of targeted drug delivery systems. The enhanced quercetin release at the lower stationary magnetic field and higher frequency alternating magnetic field, together with the synergism of chemical and physical, i.e., superparamagnetic properties of MNPs, demonstrate the great potential of MNPs as a promising targeted drug delivery system with high potential for their, both therapeutic and diagnostic activity
| doab | 2025-04-07T03:56:59.195104 | 17-11-2022 17:23 | {
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007afdec-bed4-405d-873d-c355ba9add0e.76 | *2.1. Chemicals*
Iron (III) chloride hexahydrate (97%) was purchased from Alfa Easar (Ottawa, ON, Canada). Ammonium acetate and polyethylene glycol (PEG, Mw = 4000 Da) were obtained from Sigma Aldrich (St. Louis, MO, USA). Ethylene glycol and ethanol (96%) were purchased from Lach-ner (Neratovice, Czech Republic). Compressed nitrogen was purchased from Messer (Bad Soden am Taunos, Germany). Silicon oil was received from Acros organics (Waltham, MA, USA). Phosphate buffered saline (PBS) buffer (PBS tablets, pH 7.4, *<sup>I</sup>*<sup>c</sup> = 150 × <sup>10</sup>−<sup>3</sup> mol dm−3) were purchased from Sigma Aldrich (St. Louis, MO, USA). Deionized water Millipore mili Q-H2O was used to prepare the PBS medium. Quercetin (≥99%) was supplied by Lach-ner (Neratovice, Czech Republic). A molecular weight cut-off dialysis bag (MWCO, 8Kd) was purchased from Thermo Fisher Scientific (Waltham, MA, USA).
| doab | 2025-04-07T03:56:59.195607 | 17-11-2022 17:23 | {
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007afdec-bed4-405d-873d-c355ba9add0e.77 | *2.2. Synthesis of Bare and PEG Coated Magnetite MNPs*
The modified solvothermal reaction was used for the preparation of mesoporous magnetic nanoparticles [33,34]. Briefly, 1.35 g FeCl3 × 6H2O, 3.85 g of CH3COONH4, 1.0 g PEG (M = 4 kD) and 70 mL of ethylene glycol were added to a 250 mL two-necked flask equipped with a magnetic rod. The mixture was stirred vigorously for 1 h at 160 ◦C with a Heidolph MR Hei-Standard mixer (Schwabach, Germany). The chemical reaction was under the protection of an inert nitrogen atmosphere to form a homogeneous brownish solution (see Figure 1). After an hour, the system was stopped and cooled to room temperature. The mixture was transferred into a Teflon-coated stainless-steel autoclave (BHL 800 Berghof, Eningen, Germany) which is connected to a temperature controller. The autoclave was heated to 200 ◦C and maintained for 19 h and, afterwards, it was cooled to 50 ◦C. To remove the solvent, MNPs were centrifugated (Universal 320 Hettich Zentrifugen, Tuttlingen, Germany) at 8000 rpm for 10 min. After separation, MNPs were washed several times with ethanol and dispersed with a shaker (IKA Shaker Vortex 1, Staufen, Germany) between each wash. Finally, MNPs were left to dry in a desiccator for further characterization. For comparison purpose and characterization of NPs, bare magnetite MNPs were resynthesized each time using the same procedure.
**Figure 1.** Schematic illustration of mesoporous MNPs preparation using solvothermal method.
| doab | 2025-04-07T03:56:59.195682 | 17-11-2022 17:23 | {
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007afdec-bed4-405d-873d-c355ba9add0e.78 | *2.3. Characterization of Synthesized Magnetite MNPs*
The structural characteristics were determined by X-ray powder diffraction using Philips MPD 1880 diffractometer (Brooklyn, NY, USA) with monochromatic CuKα radiation (*λ* = 0.154 nm) at room temperature. The structural features of prepared samples were recorded at 2*θ* angles in the range of 10◦–70◦ with a step of 0.02◦ and fixed time of 10 s per step. X-ray diffraction in polycrystalline is used to confirm crystalline size and structure of bare and coated magnetite MNPs obtained by solvothermal method. Field emission scanning electron microscope, JEOL JSM-7000F (Tokyo, Japan), was used to determine the morphology, particle size distribution and surface texture of bare and PEG-coated magnetite MNPs. FE-SEM was linked to the EDS/INCA 350 (energy dispersive X-ray analyzer) manufactured by Oxford Instruments Ltd., London, UK. The morphology of the MNPs has been further investigated using atomic force microscopy (AFM). The samples for AFM imaging were prepared by deposition of a magnetite MNPs suspension on the mica substrate. The MNPs are rinsed three times with 50 μL of MiliQ water to remove all residual impurities. AFM images were obtained by scanning the magnetite MNPs on the mica surface in the air using MultiMode Scanning Probe Microscope with Nanoscope IIIa controller (Bruker, Billerica, MA, USA) with SJV-JV-130 V ("J" scanner with vertical engagement); Vertical engagement (JV) 125 μm scanner (Bruker Instruments, Inc., Bruker, Billerica, MA, USA). Tapping mode was performed for imaging using silicon tip (R-TESPA, Bruker, Nom. Freq. 300 kHz, Nom. spring constant of 40 N/m) at 25.0 ◦C, allowing thermal equilibration before each imaging. AFM images were collected using random spot surface sampling (at least two areas per sample) for each analysed sample. All the images were processed by first-order flattening only and analysed using the NanoScope Analysis software (Version 5.31r1). Morphology analysis was investigated by Transmission Electron Microscope (TEM), Zeiss EM10 (Oberkochen, Germany), operated at 100 kV. In this propose, MNPs coated with PEG were dispersed in Milli-Q H2O and bare MNPs were dispersed in ethanol, ultrasonicated and placed on carbon coated copper grids. After air drying, samples were photographed by transmission electron microscopy. Samples were analysed with ImageJ and 50 particles were counted for each image.
The MNPs mesoporosity determination has been performed using nitrogen adsorption–desorption measurements on an ASAP2020 (Micromeritics, Norcross, GA, USA) accelerated surface area analyzer at 77 K. Before measuring, the samples were degassed at reduced pressure andat 120 ◦C for at least 6 h. All measurements have been made in duplicate. The specific surface area, the pore volume and the pore size are determined using Brunauer–Emmett–Teller (BET) analysis. To confirm the superparamagnetic property of the synthesized MNPs, magnetization measurements were performed. Magnetization of the powder samples of MNPs was measured using a commercial Quantum Design MPMS-5 SQUID magnetometer (San Diego, CA, USA). The powder samples were placed into a small ampoule whose diamagnetic contribution was properly subtracted. In addition, the field dependence of the magnetization (M(H)), including magnetic hysteresis loops, was measured at 300 K under fields up to 10 kOe.
| doab | 2025-04-07T03:56:59.195802 | 17-11-2022 17:23 | {
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007afdec-bed4-405d-873d-c355ba9add0e.79 | *2.4. The Determination of Loaded Quercetin into Magnetic Mesoporous MNPs*
The quercetin loading efficiency into mesoporous MNPs has been confirmed using Brunauer–Emmett–Teller (BET) analysis. The difference in the specific surface area, the pore volume and the pore size before and after immersing the MNPs into the quercetin solution confirmed the loading of quercetin into MNPs. FTIR spectra were obtained by Alpha-T FTIR Spectrometer (Bruker, Billerica, MA, USA). All spectra were recorded between 4000 and 350 cm−<sup>1</sup> at a nominal resolution of 4 cm−<sup>1</sup> at 25 ◦C, and the total number of recordings was 16. Dried samples were mixed with KBr powder and then they were pressed to produce pellets. TG analysis data were carried out on a TG/DTA simultaneous analyser DTG-60H with a 10 ◦C/min heating rate under a nitrogen atmosphere. The measurements were recorded in a range of room temperature up to 1200 ◦C.
UV/VIS spectrophotometer was used to study the quercetin loading efficiency of the synthesized MNPs. The loading efficiency was calculated by measuring the absorbance of the supernatant with the WTW photoLab® 7600 UV-VIS Spectrophotometer (Xylem, Rye Brook, NY, USA) at 374 nm. Measurements were performed in a rectangular quartz cuvette with a 10 mm optical path length and covers at 25 ◦C.
A total of 500 mg of quercetin was dissolved in 100 mL of ethanol and suspended in an ultrasonic bath (Bandelin Sonorex Super RK 100 H, Berlin, Germany) for 1 h at room temperature. The 25 mL aliquot of quercetin solution and 100 mg of coated MNPs were transferred into a 50 mL Falcon conical centrifuge tube. The mixture was mechanically stirred at a thermocontrol shaker (Barnstead Lab-line 4450 e-class) for 24 h at 200 rpm and 25 ◦C. Afterwards, the quercetin-loaded MNPs were separated from unloaded quercetin by centrifugation (Universal 320 Hettich Zentrifugen, Tuttlingen, Germany) at 8000 rpm for 15 min. Compared with quercetin concentration supernatant before adding the synthesized MNPs, the concentration loss was determined using a calibration curve in pure EtOH. The coefficient of determination was 0.9978, and the determined molar absorption coefficient of quercetin at temperature 298 K and 374 nm is 19,131 mol−<sup>1</sup> cm−<sup>1</sup> dm3. The loading efficiency (*LE*) was calculated by measuring the absorbance of the supernatant with the Photolab 7600 UV-VIS spectrophotometer (Xylem, New York, NY, USA) at *λ* = 374 nm. The absorbance (*λ* = 374 nm) was collected and converted to concentration by using the equation from the calibration curve. Therefore, the drug loading efficiency was calculated as the following equation:
$$LE = \frac{m\_{\text{embeddeded}}}{m\_{\text{NP}}} \times 100\% \tag{1}$$
where *m*embedded represent the mass of quercetin incorporated in nanoparticles, and *m*NP is the total mass of MNPs which is used for loading.
#### Size Distribution of Magnetic MNPs Using Dynamic and Electrophoretic Light Scattering
Hydrodynamic diameter (*d*H) and zeta potential (*ζ*) of suspended MNPs were measured by photon correlation spectrophotometer, Zetasizer Nano ZS (Malvern, UK) with green laser (*λ* = 532 nm) using the M3-PALS technique. All measurements were conducted at 25 ◦C in PBS (pH = 7.4) buffer. The hydrodynamic diameter was determined from the peak maximum of the volume size function. The zeta potential (*ζ*) was calculated from the electrophoretic mobility using a Smoluchowski approximation (*f*(κa) = 1.5). The *d*<sup>H</sup> values were obtained as an average value of 10 measurements, while the zeta potential values were reported as an average of 3 independent measurements. The results were collected by the Zetasizer software 6.32 (Malvern Instruments, Malvern, UK).
| doab | 2025-04-07T03:56:59.196300 | 17-11-2022 17:23 | {
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007afdec-bed4-405d-873d-c355ba9add0e.80 | *2.5. Release Study of Quercetin under Stationary and Alternating Magnetic Fields*
The apparatus setup scheme shown in Figure 2 enables the fine-tuning of the quercetinrelease kinetic profile under applied combined stationary and alternating magnetic fields. The magnetic field enforced in the experiment is a combination of stationary and alternating magnetic fields. The position of the permanent magnet fixed and stable towards the fixed magnetic coil placed within the reactor ensures the permanent and alternating magnetic fields perpendicular to each other [35]. Without going into more detail, we showed in previous work [35] that, by applying external magnetic fields, MNPs behave as Brownian particles with quasi-periodic movement that enables loaded drug molecules to became enhanced released from the MNPs. A magnetic field is an effective stimulus with deep penetration capacity. The high-frequency alternating magnetic field (HF-AMF), from 50 to 400 kHz, and low-frequency alternating magnetic field (LF-AMF), from 0.1 to 5 kHz, have been employed to trigger the release of drugs from nanocarriers [36]. In our previous work, we employed the LF-AMF to induce the flavonoid release from the magnetic aggregates [35]. To get more insights into the effect of the combination of applied permanent and AMF to drug release, we designed sophisticated HF-AMF instrument to induce increased drug release at HF-AMF having in mind Brezovitch criterion [37]. He proposed a safety limit
where the product of magnetic field and amplitude frequency (H0×f) should not exceed 4.85 × 108 A m−<sup>1</sup> <sup>s</sup>−<sup>1</sup> to avoid any harmful effect on the organism. In our study, the product amounts 4 mA m−<sup>1</sup> s−1, 1 mA m−<sup>1</sup> s−<sup>1</sup> and 0.3 mA m−<sup>1</sup> for 100 kHz, 50 kHz and 10 kHz, respectively, indicating that we chose good frequencies and applied magnetic fields for any possible safe application to patients. In consideration of the design of our experiment, where there were relatively high frequencies of alternating magnetic fields (from 10 to 100 kHz), it was expected that the release of quercetin would be significantly enhanced by the influence of higher frequency.
**Figure 2.** Experimental setup of the release kinetics and schematic illustration of a reactor.
The drug release was calculated by measuring the absorbance of quercetin released in the supernatant at *λ* = 330 nm by the UV-Vis spectrophotometer. The releasing kinetics of quercetin from magnetite MNPs was investigated at three temperatures (25 ◦C, 30 ◦C and 37 ◦C) in the mixture PBS/EtOH (vol. 50/50) in which the solubility of quercetin totals to 5.66 mg/mL, 5.83 mg/mL and 6.02 mg/mL at 25 ◦C, 30 ◦C and 37 ◦C, respectively [38]. For the use of NPs as a drug delivery system, a physiological temperature of 37 ◦ C is of crucial importance. However, before the application of MNPs, they must be adequately stored. There are studies on the storage of MNPs at low temperatures, and the influence of low temperatures on the magnetic properties of MNPs [39]. Although the temperature effects on the magnetic properties of magnetite are very weak, MNPs contain a coating of organic material, in our case PEG 4000. Since PEG 4000 has crystallization temperatures at around 32 ◦C [40], which means that by changing the temperature of the MNPs suspension from the storage temperature to the ambient temperature, and further to the physiological temperature of 37 ◦C, the surface properties of MNPs and the relaxation kinetics could be changed. For this reason, measurements were made not just at 37 ◦C, but also at 25 and 30 ◦C to see how much the temperature change affects the drug-release kinetics. In addition, similar studies of the mechanism of release kinetics from NPs at different temperatures (10, 22 and 37 ◦C) have been conducted, for example, in study by Gronczewska [41], in that drug release and matrix degradation of polymer microspheres with different glass transition temperatures were investigated at various temperatures in order to clarify the effect of temperature on mechanisms of drug release. Then, 100 mg of quercetin-loaded MNPs were transferred in a dialysis bag (MWCO 8 kD, Thermo Fisher Scientific (Waltham, MA, USA)), after which 1 mL of PBS/EtOH (Vol. % = 50:50) medium was added and closed with dialysis bag clip holders. The dialysis bag was placed in a glass cylindrical reactor with a thermostatic jacket and flange size LF 100 containing 150 mL of PBS/EtOH mixture. The overhead stirrer (Ministar 20 control, IKA-Werke GmbH&Co (Staufen, Germany)) is going through the centre neck flat flange lid. The stirrer was set to 200 rpm. At selected time intervals, 1 mL of supernatant was replaced with fresh PBS/EtOH mixture through one angled side neck flat flange lid. The reactor is connected to a refrigerated–heating circulator (Corio CD-201F Julabo GmbH (Seelbach, Germany)) to control the appropriate temperature. The controlled release of quercetin was tested using appropriate external alternating (10 kHz, 50 kHz and 100 kHz) and stationary magnetic fields (*B* = 7.9 mT
and 11.0 mT) at controlled electric current (*I* = 100 mA). An external alternating magnetic field was achieved with a function generator (Wavetek 164 30 MhZ (San Diego, CA, USA) connected to the coil (N = 270, *l* = 4 cm). Experiments were performed using the magnetic field system set up from permanent disk-shaped magnets (rare earth) and solenoid with permalloy core connected to signal generator alternating 100 mA current. A reactor with the sample was placed among the two magnetic fields. Defining the *O*xy plane as the surface of the liquid, the permanent field was along the *O*<sup>z</sup> axis and the alternating field along the *O*<sup>x</sup> axis. Weak fields were applied in all experiments: the strength of the static permanent magnetic fields at the appropriate distance between the membrane dialysis bag and the permanent magnet was *B* = 7.9 mT and 11.0 mT. They were placed on a stand inside the reactor and used as sources of the permanent magnetic field. The function generator is connected to the oscilloscope (DS1000Z, Rigol Technologies (Beijing, China), which allowed the observation of the sinus waveform signal. The release kinetics of quercetin from magnetite MNPs was quantified by UV/VIS spectrophotometry (Photolab 7600 UV/VIS spectrophotometer Xylem (New York, NY, USA)). The linearity of the calibration was found to be valid from 1 × <sup>10</sup>−<sup>6</sup> mol dm−<sup>3</sup> to 1 × <sup>10</sup> <sup>−</sup><sup>4</sup> mol dm−<sup>3</sup> with correlation coefficients for quercetin all approaching 1.00. All release kinetics experiments have been performed in duplicate.
#### **3. Results and Discussion**
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007afdec-bed4-405d-873d-c355ba9add0e.81 | *3.1. Characterization of Synthesized Magnetite MNPs*
The X-ray powder diffraction patterns of the synthesized bare and PEG-coated MNPs are presented in Figure 3A,B, respectively. Characteristic peaks exhibited in the XRPD pattern are well-matched with the magnetite diffraction peaks and confirm the cubic inverse spinel structure of MNPs. The sharp diffraction of three characteristic peaks (220), (311) and (400) also indicate the spinel structure of magnetite [42]. The formation of pure cubic magnetite is confirmed by the value of the calculated lattice parameter "*a*" which has been determined to be 8.389 Å [43]. Using Scherrer's equation, the average crystallite size of bare and PEG-coated MNPs were calculated to be 25 nm and 19 nm, respectively. In addition, the decreased average crystallite size, another piece of evidence suggesting that PEG decreases the crystallinity of MNPs at a lower intensity and with broader diffraction peaks than PEG-coated MNPs.
The size and morphology of bare and PEG-coated mesoporous MNPs were observed by field-emission scanning electron microscope (FE-SEM), atomic force microscopy (AFM) and transmission electron microscopy (TEM), as shown in Figure 3A,B,G–J,K–L). Both bare and PEG-coated MNPs maintain a uniform spherical shape, some of them agglomerated due to magneto–dipole interactions between MNPs. While bare MNPs showed cluster structure with a very rough surface, the process of PEG coating revealed the flattened surface of mesoporous MNPs (Figure 3I,J). These findings have been confirmed by AFM imaging where the roughness of the MNPs surface has been increased by PEG coating, for almost 100%, from 5.58 ± 1.06 nm to 10.9 ± 2.1 nm, confirming the effective coverage of the bare MNPs by PEG [43]. The less agglomerated texture of the PEG-coated MNPs can be related to the effect of the PEG layer during the synthesis of MNPs. In addition, a size histogram of bare mesoporous MNPs obtained using SEM micrographs shows a broader size distribution than PEG-coated MNPs (*d*ave = 103.4 ± 0.7 nm and 101.0 ± 0.9 nm for bare and PEG-coated MNPs, respectively), indicating that polymer decreased the magnetic interaction among the particles and prevent their agglomeration. The cluster structure of mesoporous MNPs has been confirmed using AFM. The average size of bare MNPs and PEG-coated MNPs corresponds to results obtained by SEM, it was even 10% larger due to the convolution effect. However, in both 2D height images, it is shown that the cluster structure MNPs consists of smaller substructures, in the range of 15 to 35 nm, which roughly correspond to the dimension of the crystallite size obtained by X-ray powder diffraction. Hence, AFM revealed that only several nanometers increase in roughness is observed when PEG 4 kD is used, suggesting that the PEG layer is largely twisted the
magnetite surface, rather than stretch out linearly [44]. Furthermore, this PEG layer should decrease the magnetic interactions among the MNPs and prevents their agglomeration. The mean diameter of bare MNPs obtained by TEM is larger (143 ± 30 nm) than obtained by SEM (103.4 ± 0.7 nm), indicating a higher degree of polydispersity. However, the mean diameter of PEG-coated MNPs (96 ± 10 nm) corresponds the value obtained by SEM (101.0 ± 0.9 nm). Increasing the size of the PEG-coated MNPs can be attributed to the successful coating of PEG. As shown in Figure 3K,L, both MNPs maintain a typical spherical shape.
Nitrogen adsorption–desorption isotherms of bare MNPs confirmed their mesoporosity (see Figure 4). The surface area, pore size and total pore volume were calculated to be 20.5 m2g−1, 17.7 nm and 0. 09 cm3 g−1, respectively, strongly supporting the fact that the bare MNPs have mesoporous structure. The PEG coating of bare MNPs led to a slight decrease in surface area (19.3 m2 g−1) but an increase in pore size and total pore volume (24.6 nm<sup>1</sup> and 0.11 cm<sup>3</sup> g<sup>−</sup>1, respectively).
FTIR spectrum (Figure 5A) of bare magnetite MNPs shows bands at 585 and 395 cm−<sup>1</sup> corresponding to the symmetric stretching vibration mode of the Fe-O bond. The absorption maxima at 3452 cm−<sup>1</sup> and 1644 cm−<sup>1</sup> suggest the presence of O-H linkages. In the pure PEG, the bands at 1341 and 1100 cm−<sup>1</sup> belong to the C-O-C ether bond asymmetric and symmetric stretching vibrations. The band at 2890 cm−<sup>1</sup> is attributed to -CH2- stretching vibration in PEG. In addition, absorption bands at 1283 and 1465 cm−<sup>1</sup> attributed to the vibration of–CH2 and at 964 cm−<sup>1</sup> corresponds to the CH out-of-plane vibration [18]. The hydroxyl groups were also confirmed at 3444 and 1631 cm<sup>−</sup>1. The presence of characteristic FTIR bands of PEG in PEG-coated MNPs spectrum confirmed the successful coating of PEG on the surface of magnetite MNPs. The PEG-coated MNPs spectrum shows a strong C-O-C ether stretch at 1110 and 1381 cm−<sup>1</sup> [44]. In addition, the O-H linkages at the 1642 and 3445 cm−<sup>1</sup> bands exhibited enhanced intensity which also indicates that PEG modified the surface of MNPs. The transmittance bands at 589 and 399 cm−<sup>1</sup> confirm the symmetric stretching mode of the Fe-O bond. The results indicate that PEG is successfully functionalizing the surface of MNPs.
In order to confirm the superparamagnetic properties of synthesized bare and PEGcoated MNPs, measurements of the magnetization curve have been performed. The S-shaped hysteresis loops are a typical feature of superparamagnetic MNPs and obtained result is very similar to our previous work (77 emu g−1) on Fe3O4 MNPs [35]. The magnetization curve clearly shows that magnetization depends on the applied magnetic field, but not on the sign of the applied field. Magnetic characterization at 300 K indicates that the bare MNPs have saturation magnetization at the maximum field of 5 kOe value of 76.71 emu g−1, which is lower than obtained value for the bulk Fe3O4 (*M*<sup>s</sup> = 92 emu g−1) [42] or *M*s = 96.43 emu g−<sup>1</sup> [19].
The observed decrease in the saturation magnetization could be explained either by the use of PEG for the surface modification process [43] that causes the softening of the magnetization or by the difference in particle size [45]. In addition, the saturation magnetization of PEG-coated MNPs at the maximum field of 5 kOe is 74.75 emu g−1, somewhat lower than bare MNPs (Figure 5B). However, the magnetization measurement of synthesized MNPs confirmed their superparamagnetic properties, thus confirming their further application in release studies under applied magnetic fields.
| doab | 2025-04-07T03:56:59.196945 | 17-11-2022 17:23 | {
"license": "Creative Commons - Attribution - https://creativecommons.org/licenses/by/4.0/",
"book_id": "007afdec-bed4-405d-873d-c355ba9add0e",
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"author": "",
"title": "Characterization and Encapsulation of Natural Antioxidants: Interaction, Protection and Delivery",
"publisher": "MDPI - Multidisciplinary Digital Publishing Institute",
"isbn": "9783036554563",
"section_idx": 81
} |
007afdec-bed4-405d-873d-c355ba9add0e.82 | *3.2. Loading of Flavonoid into Magnetite MNPs*
The FTIR spectrum (Figure 6A,B) of quercetin detected OH group stretching at 3403 cm−<sup>1</sup> and 3328 cm−1. The band at 1664 cm−<sup>1</sup> corresponds to the C = O aryl ketonic stretching and the bands at 1617 cm−1, 1558 cm−<sup>1</sup> and 1520 cm−<sup>1</sup> correspond to C=C aromatic ring stretching. OH bending of the phenol functional group at 1375 cm−<sup>1</sup> and 1314 cm−<sup>1</sup> belongs to the in-plane bending of C–H in aromatic hydrocarbon. Bands at 933 cm<sup>−</sup>1, 820 cm−1, 639 cm−<sup>1</sup> and 602 cm−<sup>1</sup> correspond to aromatic C–H out-of-plane bendings. The C–O stretching in the aryl ether ring and the C–O stretching in phenol
corresponds to 1244 cm−<sup>1</sup> and 1210 cm−<sup>1</sup> transmittance maxima. The band at 1167 cm−<sup>1</sup> is attributed to the C–CO–C stretch and bending mode in ketone, respectively. The FTIR spectra of quercetin-loaded MNPs show the broadening of the OH band at 3446 cm−1, which confirms the entrapment of quercetin in MNPs [46]. The interval from 1560 to 816 cm−<sup>1</sup> matches very well with pure quercetin peaks and indicates successful loading [46].
**Figure 3.** XRD pattern of (**A**) mesoporous bare MNPs Fe3O4; (**B**) PEG-coated MNPs; SEM images of (**C**) bare Fe3O4 MNPs showing the rough rounded cluster with extensive open porosity ranging from 30 to 200 nm in diameter; (**D**) SEM image of the surface of PEG-coated Fe3O4 MNPs ranging in di-ameter from 50–130 nm; Histogram of bare MNPs Fe3O4 (**E**) and PEG-coated Fe3O4 MNPs (**F**); (2D height topographic AFM image of single (**E**) bare magnetite Fe3O4 MNPs (**G**) and (**F**) PEG-coated Fe3O4 MNPs (**H**) showing cluster structure details; 2D-amplitude AFM image of single (**G**) bare magnetite (**I**) and (**H**) PEG-coated MNPs (**J**) showing subcluster structures in size range from 10 to 30 nm; TEM images of (**K**) bare Fe3O4 MNPSs; (**L**) TEM image of the surface of PEG-coated Fe3O4 MNPs.
The loading of quercetin has been also confirmed using nitrogen adsorption–desorption isotherms of quercetin-loaded PEG MNPs. In comparison to PEG covered MNPs, the surface area has been decreased by almost 19% to 15.7 m<sup>2</sup> g−1, pore size decreased to 14.2 nm for 42.4%, while total pore volume amounting 0.06 cm<sup>3</sup> g<sup>−</sup>1, decreased for 49% to 0.06 cm3 g<sup>−</sup>1, strongly supporting the fact that the quercetin has been effectively loaded into MNPs.
The thermogravimetric study was performed to confirm the quercetin loading in MNPs. Figure 6C shows comparative weight loss for quercetin and quercetin-loaded MNPs. A thermogravimetric study of quercetin reveals that the compound undergoes a three-stage thermal decomposition. The first stage of mass loss begins at 30 ◦C and continues up to 133 ◦C. A mass loss of 3.52% is attributed to dehydration or loss of water molecules on the surface of quercetin. In the temperature range 133 ◦C to 385 ◦C, the compound experiences a weight loss of 28.5% due to the melting of quercetin. The final thermal decomposition is observed in the temperature range of 385 ◦C to 1110 ◦C, and the weight loss of quercetin is 67% [47]. In the case of quercetin-loaded MNPs, the weight loss in the temperature range from 30 ◦C to 1200 ◦C is about 12.2%, which is attributed to the decomposition of organic compounds from MNPs. This data results in the great thermal stability of quercetin when it has been encapsulated in MNPs. In the case of PEG-coated MNPs, there is an increase in the weight gain resulting from the burning of the PEG and oxygenation of Fe3O4 starting at 270 ◦C under the continuous flow of oxygen at high temperatures and finishing at 450 ◦C [48]. However, the weight loss of PEG from the PEG-coated MNPs amounts to 4%. It is assumed that the thermal decomposition of PEG occurs at both C-O and C-C bonds of the backbone chain [49]. The influence of the quercetin loading into magnetite MNPs on the size and morphology of the MNPs has been further investigated. 2D height AFM image (Figure 6D) and 2D-amplitude AFM image (Figure 6E) revealed that distinct subcluster structure containing MNPs has been retained irrespective of quercetin loading, with size grains from 20 to 50 nm in diameter. The roughness value after quercetin loading decreased from (10.9 ± 2.1) nm to (4.86 ± 1.1) nm additionally confirming the successful loading of quercetin to PEG loaded MNPs.
**Figure 4.** N2 absorption-desorption isotherm for (**A**) bare MNPs, (**B**) PEG-coated MNPs and (**C**) quercetin-loaded PEG\_MNPs.
**Figure 5.** (**A**) FTIR spectra of MNPs, PEG and PEG-coated MNPs (**B**) Magnetic behavior of MNPs. The hysteresis loop shows a slight decrease in the magnetization behavior after a thin PEG layer coating. The magnetic hysteresis loops of mesoporous MNPs at 300 K. Inset within Figure 5 (**B**) shows the magnetic coercivity *H*c = 53.3 Oe.
**Figure 6.** FTIR spectra of (**A**) quercetin and quercetin-loaded MNPs; (**B**) zoomed spectrum of quercetin-loaded MNPs in the region of quercetin reach bands; (**C**) TGA curves of PEG-coated MNPs, quercetin and quercetin-loaded MNPs. Morphology of quercetin-loaded MNPs on the 2D height AFM image (**D**) and 2D-amplitude AFM image (**E**).
In addition, UV/VIS spectroscopy is used to study quercetin loading efficiency. Compared with quercetin concentration before adding the synthesized MNPs, the concentration loss was determined using a calibration curve in pure ethanol (EtOH). The coefficient of determination was 0.9978, and the determined molar absorption coefficient of quercetin at 298 K and 374 nm is 19,131 mol−<sup>1</sup> dm3 cm<sup>−</sup>1. The loading efficiency (LE) was determined to be (20.2 ± 1.3%) calculated from 17 independent experiments. Our results suggest significant improvement of the loading efficiency of the quercetin compared with a loading efficiency of solid lipid NPs (13.20 ± 0.18%) [1]. However, the limited LE is due to the reduced specific loading site of quercetin induced by PEG coatings [50]. Despite this, the PEG –MNPs provide the capability to load antioxidant quercetin with low aqueous solubility which reflects the potential of the synthesized MNPs as drug delivery carriers.
| doab | 2025-04-07T03:56:59.197447 | 17-11-2022 17:23 | {
"license": "Creative Commons - Attribution - https://creativecommons.org/licenses/by/4.0/",
"book_id": "007afdec-bed4-405d-873d-c355ba9add0e",
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"title": "Characterization and Encapsulation of Natural Antioxidants: Interaction, Protection and Delivery",
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"isbn": "9783036554563",
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007afdec-bed4-405d-873d-c355ba9add0e.83 | *3.3. Homogeneity and Stability of Synthesized Mesoporous MNPs*
One of the essential features for successful drug delivery is stability and homogeneous dispersion of NPs in buffer. Zeta potential was used to determine the stability of the colloidal suspension of bare MNPs, PEG-coated MNPs and quercetin-loaded MNPs. It has been shown that PEG-stabilized NPs exhibit longer bloodstream circulation time and higher resistance to protein binding [44]. Due to its unique properties and its biocompatibility, PEG is selected as the stabilizing agent in this study. The surface charge of NPs has an important effect on the blood circulation time, the pharmacokinetics of NPs and the zeta potential above ±30 mV indicated to be relevant for stability of NPs in aqueous suspensions [12,51,52]. The zeta potential of MNPs (Table 1) was determined in phosphatebuffered solution (PBS). Bare MNPs exhibited a zeta potential of (−30.6 ± 0.7) mV, while PEG coating increased the absolute zeta potential value to (−35.1 ± 1.5) mV. Indeed, the higher negative zeta potential value indicated that PEG-coated MNPs possessed higher stability after PEG coating. The zeta potential of quercetin-loaded MNPs (−31.3 ± 0.8) mV decreased slightly in comparison to PEG-coated MNPs indicating quercetin adsorption on the PEG layer of MNPs. However, the observed change in zeta potential value did not decrease the stability of MNPs. Moreover, the successful loading of quercetin into PEG-coated MNPs has been confirmed also by electrophoretic measurements.
Furthermore, the hydrodynamic diameter is lower which could be attributed that PEG coating while quercetin loading provides better colloidal stability and reduces aggregation. The volume size distributions of all samples, i.e., bare MNPs, PEG-coated MNPs and quercetin-loaded MNPs were unimodal. In Table 1, it can be seen that the highest polydispersity index was observed to be 0.54 ± 0.1 for the bare synthesized MNPs suspension. This result is consistent with the results obtained from the size distribution of bare MNPs using SEM where the size distribution of bare MNPs was broader than the size distribution of the PEG-coated MNPs. The observed discrepancy between SEM and DLS data, particularly in the polydispersity index (PDI), can be explained by the fact that the SEM micrographs were taken in a dried state, whereas the DLS experiment was carried out in suspension. The PDI results reported in Table 1 also support the conclusion that PEG coatings, even presented in suspension, decreased the process of aggregation of bare MNPs and break down the cluster. The average hydrodynamic diameter of both, bare and PEG-coated MNPs also supports the above findings. In Figure 7, it is shown the volume size distributions of bare MNPs, PEG-coated MNPs and quercetin-loaded PEG\_MNPs. The PDI of PEG-coated MNPs is lower than PDI of loaded MNPs (PDI = 0.47 ± 0.1 and 0.52 ± 0.07, respectively) indicating narrower size distribution of the PEG-coated MNPs than quercetin-loaded MNPs (see Figure 7). However, the cluster sizes of quercetin-loaded MNPs were within the range of (681 ± 73) nm, which is the smallest range of all analysed MNPs. Thus, the PEG coating on the surface of Fe3O4 decreased the size compared to the bare MNPs and indicates that the PEG coating prevents or reduces aggregation to some extent.
**Table 1.** Zeta potential and hydrodynamic diameter of bare MNPs, PEG-coated MNPs and quercetinloaded MNPs.
| doab | 2025-04-07T03:56:59.197793 | 17-11-2022 17:23 | {
"license": "Creative Commons - Attribution - https://creativecommons.org/licenses/by/4.0/",
"book_id": "007afdec-bed4-405d-873d-c355ba9add0e",
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"title": "Characterization and Encapsulation of Natural Antioxidants: Interaction, Protection and Delivery",
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"isbn": "9783036554563",
"section_idx": 83
} |
007afdec-bed4-405d-873d-c355ba9add0e.84 | *3.4. Release Study*
Before we start to evaluate the quercetin-release kinetics from MNPs into PBS/EtOH (Vol. % 50:50), we want to emphasize that the results were obtained by applying external stationary and alternating magnetic fields. The underlying idea was to enable fine-tuned and controlled quercetin-release kinetics, which is certainly very important in a widespread, high-demand application of high-demand drug delivery via nanocarriers.
**Figure 7.** Volume size distributions of bare MNPs, PEG-coated MNPs and quercetin-loaded PEG\_MNPs.
The in vitro quercetin release profile from synthesized and carefully designed MNPs was studied in duration up to 8 h with a dialysis membrane and in the presence of external magnetic fields. The results are shown as the cumulative released mass of quercetin in Figure 8. Within 8 h, only up to 5% of quercetin has been released depending on the experimental conditions.
A similar sustained-release profile of quercetin from alginate NPs at pH 7.4 has been reported earlier [53], where only 10% of quercetin release was observed after 12 h, while after 9 days, only 50% of the quercetin got released. In another study, the quercetin release from the functionalized magnetite NPs was conducted in acidic and basic pH and cumulative release reached 3.7% after 6 h [14]. A similar prolonged release has also been found for quercetin release from polylactide NPs, which showed almost 60% quercetin released after 4 days [54]. The observed slow quercetin-release kinetics offers prolonged exposure to the drug and improves its efficiency compared with free drugs [55]. This is considered as an advantage because the burst release of drugs leads to a significant premature quantity of the drug that can result in toxicity [21]. For example, the quercetin at a concentration between 0 and 200 × <sup>10</sup>−<sup>6</sup> mol dm−<sup>3</sup> could decrease antioxidant activity while quercetin at a concentration of (0.2–1) × <sup>10</sup>−<sup>6</sup> mol dm−<sup>3</sup> possesses pro-oxidant activity [56]. In addition, quercetin release was affected by oxidative degradation process in PBS solution after continuously stirring for 6 h; this is yet another reason to employ nanocarriers for quercetin delivery [14,57,58]. Therefore, we prepared MNPs onto which quercetin easily adsorbs and has a prolonged stability and duration. The net result is quercetin release for a prolonged time. A first-order release profile of quercetin from Fe3O4-quercetin-copolymer NPs was revealed by Barreto et al. [3]. On the other hand, the release rate of quercetin from superparamagnetic magnetite NPs coated with chitosan, PEG and dextran was found to be of zero-order kinetics (linear with time) [41].
The assumptions in release kinetics experiments were as follows:
(i) The total amount of quercetin remained practically constant during the whole release experiment; (ii) both the quercetin solution within the membrane interior and in the membrane exterior were homogeneous due to the constant stirring and the fact that quercetin solution was never saturated; (iii) the thickness of the membrane provides the equal rate constants from the membrane interior to the membrane exterior, and vice versa; (iv) the volume within the membrane interior *V*<sup>i</sup> and the volume exterior to the membrane *V*<sup>o</sup> were constant during all performed release experiments. This is because *V*<sup>i</sup> = 1 mL and *V*<sup>o</sup> = 150 mL, *V*<sup>i</sup> < *V*o.
As a preliminary approach in working out the data, a simple model of single exponential decay was meant to be used in which the fraction of the released drug is Φ = 1 − *exp*(−*kt*), where *Φ* is a fraction of drug present in the outer volume *V*<sup>0</sup> (other fractions are in the inner volume *V*i. *Φ*i, and in the membrane, *Φ*m) [59]. The coefficient *k* is related to the apparent kinetics and, as such, cannot provide information about the actual release rate from the nanocarriers into the interior volume *V*i. However, the problem with this simple formula is the equilibrium value Φ<sup>o</sup> = 1 which is achieved when the elapsed
time is sufficiently large (*t* → +∞). In our release kinetics experiments, it is invariably between 0.04 and 0.10. Thus, the dialysis bag with its content has to be considered as a source of drug molecules. It is not possible to know the total mass of the drug that is amenable to be released, but it can be estimated from the value of *m*0, which occurs in another simple model:
$$m(t) = m\_0 \left(1 - e^{-kt}\right), \frac{m}{m\_0} = 1 - e^{-kt} \tag{2}$$
where *m*(*t*) is the released mass, not a fraction of, at time *t*, *m*<sup>0</sup> is the total released quercetin mass from the dialysis bag after infinite time, and *k* is the rate coefficient of actual release kinetics of the dialysis bag membrane. The problem is that *m*<sup>0</sup> is not experimentally well defined, i.e., it is only approximately constant across the series of experiments. Fitting the release experimental data obtained at various magnetic fields and at three different temperatures using this simple model (Table 2 and Figures 8 and 9) resulted as expected in a fairly narrow interval of *m*<sup>0</sup> values with average *m*<sup>0</sup> = (1.48 ± 0.34) mg.
Each experiment was done in duplicate. This was fully justified to avoid averaging the measurement results and use the mean values of the two fitting procedures because the product, *kt*, was always rather small (Table 2). This was the case because, if *<sup>m</sup>*<sup>1</sup> <sup>=</sup> *<sup>m</sup>*0,1- <sup>1</sup> − *<sup>e</sup>*−*k*1*<sup>t</sup>* , *<sup>m</sup>*<sup>2</sup> <sup>=</sup> *<sup>m</sup>*0,2- <sup>1</sup> − *<sup>e</sup>*−*k*2*<sup>t</sup>* and *m* = *m*<sup>0</sup> - <sup>1</sup> <sup>−</sup> *<sup>e</sup>*−*kt* where *m* = 1/2(*m*<sup>1</sup> + *m*2) and *m*<sup>0</sup> = 1/2(*m*0.1 + *m*0.2), the following is obtained:
$$
\varepsilon^{-kt} = \frac{m\_{0.1}}{m\_{0.1} + m\_{0.2}} \varepsilon^{-k\_1 t} + \frac{m\_{0.2}}{m\_{0.1} + m\_{0.2}} \varepsilon^{-k\_2 t} \tag{3}
$$
Since the product *kt* is always very small, it turns out that a simple formula,
$$k = \frac{m\_{0.1}}{m\_{0.1} + m\_{0.2}}k\_1 + \frac{m\_{0.2}}{m\_{0.1} + m\_{0.2}}k\_2\tag{4}$$
can be used. Furthermore, it is very often *<sup>m</sup>*0,1<sup>≈</sup> *<sup>m</sup>*0,2 which gives *<sup>k</sup>* <sup>≈</sup> <sup>1</sup> <sup>2</sup> (*k*1+ *k*2).
It is important to emphasize that the intent of this study was not only to estimate the time required for the complete quercetin release from nanocarriers, but also to investigate and demonstrate how the stationary and alternating field affect the rate of quercetin release. In our previous work [35], we have shown how the simultaneous application of stationary and alternating field can accelerate the release of drug from aggregates of MNPs. Being relatively unstable and dysfunctional, aggregates vigorously moved under the influence of the alternating field which resulted in drug release enhancement. MNPs that were additionally functionalized and stabilized by the PEG layer were used for this purpose in the present study. Figure 8 shows the release kinetics of quercetin at the temperature of 30 ◦C in the absence of the magnetic field and under an alternating field of 10 kHz, 50 kHz and 100 kHz at a constant stationary magnetic field *B* = 11 mT.
Since we used a dialysis membrane bag, the first step was to perform calibration experiments with free quercetin following the same protocol as in experiments with MNPs to get information about membrane permeation kinetics during the first several hours (*<sup>k</sup>* = 6.617 × <sup>10</sup>−<sup>3</sup> min<sup>−</sup>1; *<sup>k</sup>* = 9.637 × <sup>10</sup>−<sup>3</sup> min−<sup>1</sup> and *<sup>k</sup>* = 14.592 × <sup>10</sup>−<sup>3</sup> min−<sup>1</sup> at 25 ◦C, 30 ◦<sup>C</sup> and 37 ◦C, respectively). We selected the MWCO cellulose membrane (8 kD) membrane based on the porosity of the dialysis membrane as well as to avoid possible adverse interactions between quercetin and the membrane materials. The cellulose membrane has recently been used in the measurement of both, drug diffusion and drug release rates from varied formulations, such as creams and hydrogels [7].
The rate constant of the membrane when there was non-saturated quercetin solution in the membrane bag indicated the barrier effects of dialysis membrane and showed faster membrane permeation kinetics at higher temperatures (*k* = 0.0066 min−1, 0.0096 min−<sup>1</sup> and 0.0146 min−<sup>1</sup> at 25 ◦C, 30 ◦C and 37 ◦C, respectively), as expected. The rate constants obtained in our experiments are larger than those obtained in release kinetics of doxorubicin by Yu et al., where *<sup>k</sup>* = 0.019 ± 0.003 *<sup>h</sup>*−<sup>1</sup> = 0.0003 min−<sup>1</sup> [60].
With no magnetic field and at 30 ◦C, the rate constant is *<sup>k</sup>* = 0.0019 ± 0.0001 min−1. At 10 kHz, 50 kHz and 100 kHz and under *B* =11.0 mT the rate constant values were *<sup>k</sup>* = 0.0032 ± 0.0009 min−1, *<sup>k</sup>* = 0.0019 ± 0.0001 min−<sup>1</sup> and *<sup>k</sup>* = 0.0034 ± 0.0013 min−1, respectively. Thus, the release of the quercetin is the faster at the highest field frequency. It is evident that the alternating magnetic field can accelerate the quercetin release at a given stationary magnetic field.
**Table 2.** The experimental release kinetics under the permanent magnetic fields of 7.9 mT and 11.0 mT and three frequencies (10 kHz, 50 kHz and 100 kHz) at temperatures 25 ◦C, 30 ◦C and 37 ◦C.
**Figure 8.** The representative cumulative release profiles for the quercetin from MNPs through dialysis membrane under the stationary magnetic field *B* = 11 mT and alternating field with frequencies 10 kHz, 50 kHz and 100 kHz at 30 ◦C.
**Figure 9.** The representative cumulative release profiles for the quercetin from MNPs through dialysis membrane at 25 ◦C using two stationary magnetic fields and three different frequencies of alternating magnetic field (**A**) 100 kHz, (**B**) 50 kHz and (**C**) 10 kHz.
Our next task was to see the effect a stationary magnetic field on release kinetics. Figure 9 shows the dependence of quercetin-release kinetics at 25 ◦C under the alternating field frequency of 10 kHz and stationary magnetic field of *B* = 7.9 mT and 11 mT (Figure 9C). Under the stationary magnetic field of 7.9 mT, the membrane bag has released quercetin with a rate constant *<sup>k</sup>* = 0.0043 ± 0.0003 min−1. When a stronger stationary field of 11 mT is applied, the release kinetics becomes slower, *<sup>k</sup>* = 0.0024 ± 0.0012 min−1. The opposite effect on the release kinetics was observed at the highest 100 kHz frequency (Figure 9A), where an increase in the rate constant with increasing stationary field by almost 95% was observed (at *<sup>B</sup>* = 7.9 mT and *<sup>B</sup>* = 11 mT, *<sup>k</sup>* = 0.0015 ± 0.0002 min−1; *<sup>k</sup>* = 0.0029 ± 0.0004 min<sup>−</sup>1, respectively).
Since the opposite effects of the influence of a stationary magnetic field on the rate constant are obtained at different frequencies of the alternating field, our next analysis focuses on the measurements of the rate constants at the same stationary field and frequency, but at different temperatures. Comparing the constants of the apparent release rate of quercetin at a stationary field of 7.9 mT with an increase in temperature from 25 to 37 ◦C (*<sup>k</sup>* = 0.0022 ± 0.0004 min−1, *<sup>k</sup>* = 0.0022 ± 0.0008 min−<sup>1</sup> and *<sup>k</sup>* = 0.0025 ± 0.0002 min−<sup>1</sup> for 25 ◦C, 30 ◦C and 37 ◦C, respectively), it can be seen that the rate constant slightly increases with increasing temperature only above 30 ◦C. The same effect was observed under the stronger stationary field (*B* = 11 mT, see Table 2). At higher temperatures kinetic energy of quercetin molecules is larger or, putting differently, their diffusivity is larger and more quercetine is released and detected within the same time interval. If we compare the rate constants at the same temperature (e.g., 30 ◦C), the rate constant obtained at a stronger stationary field (*<sup>B</sup>* = 11 mT) and 50 KHz, is smaller (*<sup>k</sup>* = 0.0019 ± 0.0001 min−1) than at a weaker field of 7.9 mT (*<sup>k</sup>* = 0.0022 ± 0.0008 min−1). It is obvious that when overcoming stationary field, the thermal energy of the MNPs is large enough to increase the movement of the MNPs and increase quercetin release. The influence of the temperature under the constant stationary field and the frequency of the alternating field clearly shows that the amount of quercetin released increases with increasing temperature at almost all applied frequencies of the alternating fields except at *B* = 11 mT and frequencies *f* = 100 and 50 kHz and at *B* = 7.9 mT and *f* = 50 kHz, which is also reflected in the magnitudes of the quercetin release rate constants. For example, at *<sup>B</sup>* = 11 mT and *<sup>f</sup>* = 100 kHz, *<sup>k</sup>* = 0.0029 ± 0.0004 min<sup>−</sup>1; *<sup>k</sup>* = 0.0034 ± 0.0013 min−<sup>1</sup> and *<sup>k</sup>* = 0.0038 ± 0.0013 min−<sup>1</sup> for 25 ◦C, 30 ◦C and 37 ◦C, respectively.
Thus, it was shown here that by choosing the temperature, the quercetin release rates can cover a wide range of values. We have shown that the synthesized MNPs are suitable nanocarriers for quercetin, especially because the required drug dose can be delivered in a prolonged time. Since the average half-life of quercetin absorbed in the human body is 3.5 h [61], this study represents a significant improvement for flavonoid delivery, which, when loaded into MNPs, remains stable in a prolonged period of time.
## **4. Conclusions**
In summary, superparamagnetic magnetite nanoparticles (MNPs) were prepared by solvothermal method and stabilized by biocompatible poly (ethylene glycol) PEG-4000 Da. The X-ray powder diffraction patterns of the synthesized bare and PEG-coated MNPs confirmed the cubic inverse spinel structure of MNPs. The size and morphology of bare and PEG-coated MNPs have been obtained by SEM, TEM and AFM analysis. By AFM is showed that bare MNPs have cluster structure with a very rough surface and when bare MNPs is coated with PEG, the roughness of the MNPs surface has been increased by PEG coating, for almost 100%, from 5.58 ± 1.06 nm to 10.9 ± 2.1 nm, confirming the effective coverage of the bare MNPs by PEG. A size histogram of bare mesoporous MNPs obtained using SEM micrographs shows a broader size distribution than PEG-coated MNPs (*d*ave = 103.4 ± 0.7 nm and 101.0 ± 0.9 nm for bare and PEG-coated MNPs, respectively), indicating that PEG decreased the magnetic interaction among the particles and prevent their agglomeration. Nitrogen adsorption–desorption of bare MNPs and PEG-coated MNPs confirmed their mesoporosity. The PEG molecules were successfully coated on the surface of MNPs, as revealed by FTIR spectroscopy. The PEG-coated MNPs spectrum showed a strong C-O-C ether stretch at 1110 and 1381 cm−1. The results of the saturation magnetization confirmed the superparamagnetic properties of synthesized bare and PEG-coated MNPs. The stability of MNPs improved after PEG modification, indicated by the increase in zeta potential from (−30.6 ±0.7) mV to (−35.1 ± 1.5) mV.The loading of quercetin into MNPs was confirmed by FTIR spectroscopy and thermogravimetric analysis. The UV/VIS spectra of the supernatant revealed a loading efficiency of (20.2 ± 1.3%). The quercetin release studies in vitro followed by UV/VIS spectroscopy have shown the prolonged quercetin-release kinetics from MNPs that could be controlled
by using combined stationary and alternating magnetic fields. The prolonged quercetin release, as an important characteristic for targeted drug delivery, the study of the kinetic parameters of the quercetin release process and the increased response of quercetin release under both the lower stationary magnetic field (7.9 mT) and the higher frequency of alternating magnetic field (100 kHz) suggest that the fine tuning of the release as desired along with synergism of physicochemical and superparamagnetic properties enables the great potential of MNPs as a promising targeted flavonoid delivery system.
**Author Contributions:** S.Š. designed research; L.M., I.E. and A.S. performed research; L.M., G.B. and S.Š. analysed data and contributed to the discussion; L.M., S.Š. and G.B. wrote the paper; all authors approved the final version of the paper. All authors have read and agreed to the published version of the manuscript.
**Funding:** This work was supported by the Croatian Science Foundation under the project IP-2016-06- 8415. The funders had no role in the design and conduct of this study, collection and interpretation of the data, or preparation and approval of the manuscript.
**Institutional Review Board Statement:** Not applicable.
**Informed Consent Statement:** Not applicable.
**Data Availability Statement:** Data available on request due to restrictions e.g., privacy or ethical. The data presented in this study are available on request from the corresponding author. The data are not publicly available due to extreme quality of data.
**Acknowledgments:** The authors thank T. Mrla for experimental setup and instrument development for controlling the electric current (*I* = 100 mA).
**Conflicts of Interest:** The authors declare no conflict of interest.
| doab | 2025-04-07T03:56:59.198007 | 17-11-2022 17:23 | {
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007afdec-bed4-405d-873d-c355ba9add0e.86 | *Article* **Construction of** *Polygonatum sibiricum* **Polysaccharide Functionalized Selenium Nanoparticles for the Enhancement of Stability and Antioxidant Activity**
**Wanwen Chen 1,2, Hao Cheng 1,\* and Wenshui Xia 1,2**
**Abstract:** Although selenium nanoparticles (SeNPs) have attracted great attention due to their potential antioxidant activity and low toxicity, the application of SeNPs is still restricted by their poor stability. A combination of polysaccharides and SeNPs is an effective strategy to overcome the limitations. In this study, *Polygonatum sibiricum* polysaccharide (PSP) was used as a stabilizer to fabricate SeNPs under a simple redox system. Dynamic light scattering, transmission electron microscopy, energy dispersive X-ray, ultraviolet-visible spectroscopy, Fourier transform infrared, and X-ray photoelectron spectrometer were applied to characterize the synthesized PSP-SeNPs. The stability and the antioxidant activity of PSP-SeNPs were also investigated. The results revealed that the zero-valent and well-dispersed spherical PSP-SeNPs with an average size of 105 nm and a negative ζ-potential of −34.9 mV were successfully synthesized using 0.1 mg/mL PSP as a stabilizer. The prepared PSP-SeNPs were stable for 30 days at 4 ◦C. The decoration of the nanoparticle surface with PSP significantly improved the free radical scavenging ability of SeNPs. Compared to the H2O2-induced oxidative stress model group, the viability of PC-12 cells pretreated with 20 μg/mL PSP-SeNPs increased from 56% to 98%. Moreover, PSP-SeNPs exhibited a higher protective effect on the H2O2-induced oxidative damage on PC-12 cells and lower cytotoxicity than sodium selenite and SeNPs. In summary, these results suggest the great potential of PSP-SeNPs as a novel antioxidant agent in the food or nutraceuticals area.
**Keywords:** selenium nanoparticles; *Polygonatum sibiricum* polysaccharide; stability; antioxidant
| doab | 2025-04-07T03:56:59.199138 | 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.87 | **1. Introduction**
Selenium is an essential micronutrient for humans and animals [1]. It is an integral component of more than 30 kinds of selenoproteins and selenium-containing enzymes, such as selenoprotein P (SelP), selenoprotein S (SelS), selenoprotein M (SelM), subfamilies of thioredoxin reductases (TrxR), glutathione peroxidases (GPx), and iodothyronine deiodinases (ID), that play a key role in regulating redox balance and preventing cellular damage from radicals [2,3]. However, at least one billion people in the world are at risk of selenium deficiency at present because the intake of selenium is insufficient to meet the daily requirement [4]. Epidemiological studies established that selenium deficiency is associated with many diseases, including premature aging, a decline in sperm motility, myocardial failure, neurological diseases, endemic osteoarthropathy (Keshan disease), and ischemic heart disease [5]. Although high-dose sodium selenite, methyl selenium, and selenocysteine exhibit excellent bioactivities, they can also result in serious toxicity problems, leading to many diseases [6]. Thus, it is of great importance to seek novel selenium species as food supplements or additives.
**Citation:** Chen, W.; Cheng, H.; Xia, W. Construction of *Polygonatum sibiricum* Polysaccharide Functionalized Selenium Nanoparticles for the Enhancement of Stability and Antioxidant Activity. *Antioxidants* **2022**, *11*, 240. https:// doi.org/10.3390/antiox11020240
Academic Editor: Stanley Omaye
Received: 20 December 2021 Accepted: 26 January 2022 Published: 26 January 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/).
Selenium nanoparticles (SeNPs) have gained much attention owing to their unique physical, chemical, and antioxidant activities [7]. Moreover, SeNPs have higher bioavailability and lower toxicity in comparison to other chemical forms of selenium, making them the promising alternative selenium source in food dietary [8]. However, SeNPs alone with valence state zero are highly unstable in an aqueous solution and easily transform to aggregate, resulting in lower bioactivity and further limiting their practical application [9]. Many efforts have been made to develop a simple, efficient, and green strategy for the dispersion and stabilization of SeNPs using bioactive templates [10]. Natural polysaccharides not only have complex structures, large specific surface areas, and ionizable functional groups but also possess excellent biocompatibility and biodegradability [11]. These features could decrease the surface energy of SeNPs, further preventing aggregation through electrostatic interaction or hydrogen bonds. Thus, polysaccharides applied as carriers to fabricate SeNPs with desired characteristics, such as stability and functionality, using the green chemical method is drawing much attention recently. For example, numerous studies reported that chitosan (CS) could be used as templates to prepare uniform SeNPs and the ligated SeNPs remain stable for over 1 month [12]. However, the superior properties of CS are limited due to its water insolubility and our previous research also found that CS-SeNPs aggregated under alkaline conditions (pH ≥ 9) [13]. Several polysaccharides derived from fungi [14], fruit [15], and medicinal plants [16] have been demonstrated to enhance the antioxidant activity of SeNPs. Recently, medicinal plant polysaccharides have attracted increasing attention due to their significant bioactivities with no side effects [17]. Therefore, it can be expected that the combination of medicinal plant polysaccharides with SeNPs will reduce the inherent limitations and enhance the benefits of selenium and polysaccharides.
*Polygonatum sibiricum* is a traditional Chinese herbal medicine, belonging to the *Liliaceae* family, which has been introduced in the 2015 edition of pharmacopeia [18]. China has abundant resources of *Polygonatum sibiricum*, especially in the south of the Yangtze River [19]. The constituents of *P. sibiricum* include polysaccharides, saponins, flavonoids, alkaloids, lignin, vitamins, and a variety of trace elements, of which polysaccharides are the major pharmacologically active ingredients [20]. In the last three years, *Polygonatum sibiricum* polysaccharides (PSP) are demonstrated to exhibit a wide range of pharmacological activity [21], such as osteogenic activity [22], anti-diabetes [23], immunological activity [24], and especially antioxidant activity, which makes them suitable for application in functional foods and therapeutic agents. PSP demonstrated strong antioxidant properties, which could attenuate D-gal-induced heart aging [25] and protect the mice livers against ethanolinduced oxidative damage via inhibiting oxidative stress [26]. However, no study has been reported using PSP as a decorator to functionalize SeNPs.
In this study, considering the antioxidant activity of PSP as well as the drawbacks of SeNPs, a combined strategy was conducted to fabricate SeNPs using PSP as a stabilizer in the redox system of sodium selenite (Na2SeO3) and ascorbic acid (Vc) through a simple chemistry approach. The synthesized PSP functionalized SeNPs (PSP-SeNPs) were characterized by dynamic light scattering (DLS), transmission electron microscopy (TEM), energy dispersive X-ray (EDX), ultraviolet-visible spectroscopy (UV-vis), Fourier transform infrared (FTIR), and X-ray photoelectron spectrometer (XPS). The physicochemical stabilities of synthesized nanoparticles under varying conditions, including ionic strength, pH, and temperature, were analyzed. In addition, the antioxidant activity of PSP and PSP-SeNPs was quantified by ABTS and DDPH free radical scavenging assays. Moreover, the protective effect on the H2O2-induced cell death was also investigated by MTT assay.
| doab | 2025-04-07T03:56:59.199291 | 17-11-2022 17:23 | {
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007afdec-bed4-405d-873d-c355ba9add0e.89 | *2.1. Reagents*
Commercial *Polygonatum sibiricum* polysaccharide (PSP) with a purity of 95% and a molecular weight of 14 kDa was obtained from Qiannuo Biotechnology Co. Ltd. (Xi'an, China), sodium selenite (Na2SeO3), hydrogen peroxide (H2O2), ascorbic acid (Vc), potassium persulfate (K2S2O8), 1, 1-diphenyl-2-picrylhydrazyl (DPPH), 2, 2-azinobis (3-ethylbenzothiazoline-6sulfonic acid) and diammonium salt (ABTS) were purchased from Sinopharm Chemical Reagent Co., Ltd. (Shanghai, China). All chemicals used were of analytical grade, and the water used in all experiments was obtained from the Milli-Q system.
| doab | 2025-04-07T03:56:59.199660 | 17-11-2022 17:23 | {
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007afdec-bed4-405d-873d-c355ba9add0e.90 | *2.2. Preparation of SeNPs and PSP Stabilized SeNPs*
PSP-SeNPs were prepared according to the procedure described by Ye et al. with minor modification [8]. PSP stock solution (5 mg/mL) was freshly prepared. Where 1 mL of sodium selenite solution (50 mM) was mixed with various volumes of PSP solution under stirring for 5 min. Then 1 mL of ascorbic acid solution (200 mM) was added dropwise into the mixture, and it was reconstituted to a final volume of 10 mL with Milli-Q water. The reaction was carried out at room temperature for 30 min. Finally, the solution was dialyzed using regenerated cellulose tubes (Mw cutoff 3500 Da) against ultrapure water for 48 h at 4 ◦C. The final concentrations of PSP were 0.01, 0.05, 0.075, 0.1, 0.125, 0.15, 0.25 mg/mL. SeNPs were synthesized in the absence of PSP through the same procedure as above. The resultant products were lyophilized to obtain the freeze-dried nanocomposites. The concentration of selenium was determined by the Optima 8300 inductively coupled plasma optical emission spectrometer (ICP-OES, PerkinElmer, Billerica, MA, USA).
| doab | 2025-04-07T03:56:59.199718 | 17-11-2022 17:23 | {
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007afdec-bed4-405d-873d-c355ba9add0e.91 | *2.3. Characterization*
The mean diameter, size distribution, and ζ-potential of nanocomposites were determined using a Zetasizer Nano ZS analyzer (Malvern Instruments Corporation, Worcestershire, UK). The morphology was observed using transmission electron microscopy (TEM) (JEOL, JEM-2100, Tokyo, Japan). Samples for TEM observation were prepared by placing one drop of SeNPs and PSP-SeNPs aqueous solution on a carbon-coated copper grid and dried at room temperature. The micrographs were acquired at the accelerating voltage of 200 kV. The elemental composition and distribution were determined by the energy dispersive X-ray (EDX) analysis performed on a high-resolution transmission electron microscopy (HRTEM) (JEOL, JEM-2100, Tokyo, Japan). The ultraviolet-visible (UV-vis) spectrophotometer (UV-1800, Shimadzu Corporation, Tokyo, Japan) was used to measure the UV-vis absorption spectra of SeNPs and PSP-SeNPs solutions in the wavelength range of 190–800 nm with an interval of 1.0 nm. The Fourier transform infrared (FTIR) spectra were recorded on a Nicolet iS 10 instrument (Thermo Fisher Scientific, Waltham, MA, USA). Each sample was grounded with KBr, pressed into uniform pellets, and scanned in the wavenumber range of 4000–400 cm−<sup>1</sup> with a resolution of 4.0 cm−<sup>1</sup> using pure KBr as the background. The X-ray photoelectron spectrometer (XPS) was used to analyze the valence states of the elements. The XPS patterns were operated on a Thermo Scientific ESCALab 250Xi+ (Thermo Fisher Scientific, Waltham, MA, USA) using 150 W monochromated Al Kα radiation.
| doab | 2025-04-07T03:56:59.199812 | 17-11-2022 17:23 | {
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007afdec-bed4-405d-873d-c355ba9add0e.92 | *2.4. Stability of PSP-SeNPs*
The stability of PSP-SeNPs under various conditions was investigated according to the methods described previously [27]. To determine the effect of ionic concentration on stability, 10 mL of PSP-SeNPs were mixed with different concentrations of NaCl solution (10, 50, and 100 mM). The effect of pH on the stability of NPs was analyzed by adjusting the pH of PSP-SeNPs to 2, 3, 4, 5, 6, 7, 8, 9, and 10 using 0.1 M HCl or NaOH. Where 10 mL of PSP-SeNPs were incubated in a water bath at different temperatures (25, 50, 70, and 90 ◦C) to investigate the effect of temperature on the stability of PSP-SeNPs. After being stabilized for 1 h, their mean diameter and ζ-potential were determined using a Zetasizer Nano ZS analyzer. In addition, PSP-SeNPs solutions were stored at 4 ◦C for 30 days to investigate the short-term storage stability by determining the mean diameter and ζ-potential.
#### *2.5. Antioxidant Assays*
| doab | 2025-04-07T03:56:59.199923 | 17-11-2022 17:23 | {
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007afdec-bed4-405d-873d-c355ba9add0e.93 | 2.5.1. DPPH Radical Scavenging Ability
The DPPH radical scavenging activity was determined referring to the methods reported previously with minor modifications [14]. Various concentrations of PSP, SeNPs, PSP-SeNPs, and Vc at 0.01, 0.05, 0.1, 0.25, 0.5, 0.75, 1.0 mg/mL were prepared. Further, 2 mL of the sample solutions were mixed with an equal volume of freshly prepared DPPH solution (50 mg/L) in ethanol. The mixture was shaken vigorously and incubated in darkness at 33 ◦C for 30 min. The absorbance was measured at 517 nm using a UVvis spectrophotometer. Vc was used as a positive control. The scavenging activity was calculated as follows:
$$\text{DPPH radical savingability} \left(\% \right) = \left(1 - \frac{\text{A}\_{\text{a}} - \text{A}\_{\text{b}}}{\text{A}\_{\text{c}}} \right) \times 100 \tag{1}$$
where Aa is the absorbance of the sample mixed with DPPH solution, Ab is the absorbance of the sample in the absence of the DPPH solution, Ac is the absorbance of the DPPH solution without the sample as a blank control.
| doab | 2025-04-07T03:56:59.200000 | 17-11-2022 17:23 | {
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007afdec-bed4-405d-873d-c355ba9add0e.94 | 2.5.2. ABTS Radical Cation Decolonization Assay
The assay of ABTS radical cation scavenging ability was performed as described previously with some modification [28]. ABTS and potassium persulfate (K2S2O8) were dissolved in distilled water. A stock solution of ABTS•<sup>+</sup> was prepared by mixing 7.4 mM ABTS solution with 2.6 mM K2S2O8 solution. The mixture was incubated for 12 h in the dark to reach equilibrium. The ABTS•<sup>+</sup> stock solution was diluted with sodium phosphate buffer (pH 7.4) to obtain an optical density of 0.70 ± 0.02 at 734 nm. Then 1 mL of different concentrations of PSP, SeNPs, PSP-SeNPs, and Vc (0.01, 0.05, 0.1, 0.25, 0.5, 0.75, 1.0 mg/mL) was added to 4 mL of diluted ABTS•<sup>+</sup> solution. The mixture was vigorously blended and incubated at room temperature for 6 min in darkness. The absorbance was measured at 734 nm using a UV-vis spectrophotometer. The ability to scavenge ABTS•<sup>+</sup> was calculated by Equation (2).
$$\text{ABTS}^{\bullet+} \text{radical scanning ability} \left( \% \right) = \left( 1 - \frac{\text{A}\_{\text{d}} - \text{A}\_{\text{e}}}{\text{A}\_{\text{f}}} \right) \times 100 \tag{2}$$
where Ad is the absorbance of the sample mixed with the ABTS•<sup>+</sup> solution, Ae is the absorbance of the sample in the absence of the ABTS•<sup>+</sup> solution, Af is the absorbance of the ABTS•<sup>+</sup> solution without the sample.
| doab | 2025-04-07T03:56:59.200068 | 17-11-2022 17:23 | {
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007afdec-bed4-405d-873d-c355ba9add0e.95 | *2.6. Cells Culture and MTT Assays*
PC-12 cells were cultured in Dulbecco's modified Eagle's medium (DMEM) supplemented with 10% fetal bovine serum (FBS) and 1% antibiotic mixture (100 U/mL penicillin and 100 μg/mL streptomycin). The cytotoxic effects of different selenium concentrations of PSP-SeNPs, SeNPs, and Na2SeO3 on cells were tested using MTT assays [15]. Cells were seeded in a 96-well plate at a density of 1 × <sup>10</sup><sup>4</sup> cells/well and incubated at 37 ◦C in a CO2 incubator (5% CO2 and 95% relative humidity) for 24 h. Then the medium was removed and cells were treated with different concentrations of samples prepared in DMEM with 10% FBS for an additional 24 h. After incubation, 20 μL of MTT (5 mg/mL) was added to each well and incubated at 37 ◦C for 3 h. Then the supernatant was removed and 150 μL of DMSO was added. The absorbance was measured by a microplate reader at 570 nm. The cell viability was calculated by Equation (3).
$$\text{Cell viability } (\%) = \text{OD}\_{\text{sample}} / \text{OD}\_{\text{control}} \times 100 \tag{3}$$
where ODsample is the absorbance of the treated cells and ODcontrol is the absorbance of the control cells.
To determine the protective effect of PSP-SeNPs, SeNPs, and Na2SeO3 on H2O2 induced cell cytotoxicity, cells were pre-incubated with different selenium concentrations of samples prepared in DMEM with 10% FBS for 24 h. After incubation, the medium was removed and cells were washed with PBS. Then cells were treated with a medium
containing 500 μM H2O2 for 12 h. The medium was removed and the cell viability was determined by MTT assay as described above.
| doab | 2025-04-07T03:56:59.200150 | 17-11-2022 17:23 | {
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007afdec-bed4-405d-873d-c355ba9add0e.96 | *2.7. Statistical Analysis*
All the experiments were performed at least in triplicate. The results were expressed as mean ± standard deviation (SD). Statistical analysis was carried out using paired t-tests for comparing means of two samples by the SPSS 20.0 statistical software (IBM, Armonk, NY, USA). Statistical differences between samples were performed with a level of significance of *p* < 0.05.
| doab | 2025-04-07T03:56:59.200253 | 17-11-2022 17:23 | {
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007afdec-bed4-405d-873d-c355ba9add0e.97 | **3. Results**
## *3.1. The Synthesis of SeNPs and PSP-SeNPs*
In the present study, SeNPs and PSP-SeNPs were prepared using a simple redox system of ascorbic acid and sodium selenite in the absence and presence of PSP as the stabilizer and capping agent. The visual color of the reaction solution is an indicator to preliminary infer the formation of selenium nanoparticles [29]. As shown in Figure 1, the red color of the solution indicated the SeO3 <sup>2</sup><sup>−</sup> was successfully reduced to either monoclinic or amorphous SeNPs [16]. In addition, the SeNPs in the presence of PSP showed a uniform red color and were stable in the aqueous solution. However, SeNPs without the decoration of PSP aggregated into precipitation after 1 day of storage, whereas no significant changes were observed in the solution of PSP-SeNPs. This might be attributed to the high surface energy, leading to the aggregation of SeNPs [9]. Hence, PSP plays a key role in the formation and stabilization of SeNPs.
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007afdec-bed4-405d-873d-c355ba9add0e.98 | *3.2. The Size and ζ-Potential Analysis of SeNPs and PSP-SeNPs*
The concentration of the polysaccharides is an important factor that influences the size of SeNPs, further affecting their functionality in food or medical application [30]. Thus, the effect of PSP concentrations on the hydrated particle size and the corresponding polydispersity index (PDI), as well as the ζ-potential of nanoparticles in the aqueous solution was investigated first. The particle size of barely SeNPs was up to 157 nm (Figure 2A). The addition of PSP at different concentrations could decrease the average size of SeNPs. The average diameter of PSP-SeNPs significantly decreased from 151 to 132 nm as the concentration of PSP increased from 0.01 to 0.075 mg/mL. PSP-SeNPs showed the smallest average size of 114 nm at the PSP concentration of 0.1 mg/mL, whereas further increases in PSP concentration from 0.125 to 0.25 mg/mL resulted in an increase in the size from 123 to 152 nm. It might be due to PSP at a low concentration was not enough to control the formation of SeNPs and prevent them from aggregation [31]. On the other hand, too high
PSP concentration represented more PSP chains coated on the surface of SeNPs, resulting in a larger hydration particle size [32]. As shown in Figure 2B, SeNPs in the absence of PSP exhibited a negative ζ-potential at −20.3 mV. The ζ-potential values of PSP-SeNPs were determined to be approximately −24.7, −26.6, −29.6, −30.4, −32.8, −34.9 mV at the PSP concentration of 0.01, 0.05, 0.075, 0.1, 0.125, 0.25 mg/mL. The absolute ζ-potential values of PSP-SeNPs increased with the PSP concentration increasing, further demonstrating that negatively charged PSP was exposed on the surface of SeNPs. Moreover, the higher magnitude of ζ-potential represents greater stability of nanoparticles [13], suggesting that the SeNPs decorated with PSP possess higher stability than barely SeNPs. PSP-SeNPs prepared by 0.1 mg/mL PSP were used in the following experiments.
**Figure 2.** Size distribution (**A**) and ζ-potential (**B**) of SeNPs and PSP-SeNPs prepared with different concentrations of PSP (0.01–0.25 mg/mL). Values marked with \*: *p* < 0.05, \*\*: *p* < 0.01, and \*\*\*: *p* < 0.001 indicated significant differences when compared to SeNPs.
## *3.3. Morphological and Structural Characterizations of SeNPs and PSP-SeNPs*
The morphology and size of SeNPs and PSP-SeNPs were further characterized by TEM. Figure 3A,B exhibited the TEM images of SeNPs in the absence of PSP. The results showed that adjacent SeNPs agglomerated together and presented a dendritic structure. The large-sized cluster and aggregates can also be easily visualized. However, the SeNPs in the presence of 0.1 mg/mL PSP (Figure 3C,D) exhibited a homogeneous and monodisperse spherical structure with an average size of about 105 nm, confirming the important role of PSP in regulating and stabilizing SeNPs. It should be pointed out that the hydrodynamic radius of the nanoparticles provided in the DLS analysis was larger than the size observed in the TEM image, which was sensitive to the electron-rich nanoparticles. The HRTEM image (Figure 3E) of an individual PSP-SeNPs showed a distinct lattice fringe with an interplanar spacing of 0.43 nm, revealing the excellent crystallinity of PSP-SeNPs. The elemental composition and distribution of the PSP-SeNPs were further determined by EDX. As shown in Figure 3F, the strong C, O, and Se element peaks were observed in EDX spectra. The PSP-SeNPs had a 63.10% weight percentage of C atom, together with 10.95% O atom and 25.94% Se atom. Furthermore, no other peaks for other elements were detected, indicating that PSP was successfully coated on the surface of SeNPs and confirming the purity of PSP-SeNPs [33].
**Figure 3.** TEM images of SeNPs (**A**,**B**) and PSP-SeNPs in the presence of 0.1 mg/mL PSP (**C**,**D**). HRTEM of an individual PSP-SeNPs (**E**) and typical EDX from HRTEM (**F**).
| doab | 2025-04-07T03:56:59.200387 | 17-11-2022 17:23 | {
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007afdec-bed4-405d-873d-c355ba9add0e.99 | *3.4. The Stability of SeNPs and PSP-SeNPs*
Stability is an important factor influencing the functionality and applications of nanomaterials. In this study, the effect of pH, temperature, and ionic strength on the stability of PSP-SeNPs was investigated. As shown in Figure 4A, the average size of PSP-SeNPs significantly decreased from 1262 to 186 nm when pH was increased from 2 to 3. It could be observed that no obvious changes occurred in the average size at pH range from 4 to 10. Similar results were also described previously on the stability of Polyporus umbellatus polysaccharide (PUP) coated SeNPs [34]. This might be ascribed to the protonation of PSP at pH 2 that weakened the electrostatic interactions between SeNPs and PSP, leading to the aggregation of nanoparticles. Moreover, the ζ-potential of PSP-SeNPs kept increasing with pH increased and reached the highest value of −32.6 mV at pH 7. A further increase in pH did not significantly affect the ζ-potential of PSP. It has been reported that the ζ-potential of nanoparticles was highly associated with the pKa value of the polysaccharides. The pH value higher than the pKa of polysaccharides resulted in more deprotonated characteristic groups, contributing to the increase in ζ-potential [27]. The average size of PSP-SeNPs increased from 113 to 191 nm, accompanied by the temperature increase from 25 ◦C to 90 ◦C with a constant ζ-potential at around −31 mV (Figure 4B). The result indicated that heating could increase the chances and strength of collisions, resulting in a larger size [29]. As shown in Figure 4C, the particle size of PSP-SeNPs exhibited a slight increase in 10 and 50 mM NaCl with decreased ζ-potential, and steeply increased to 882 nm in a high concentration of NaCl at 100 mM. High ion strength could remarkably reduce the surface
charge of nanoparticles due to the electrostatic interaction between positive charged Na+ and negatively charged PSP-SeNPs, resulting in the decrease of the electrostatic repulsion among nanoparticles [35]. It was observed that PSP-SeNPs were stable at about 113 nm for at least 20 days of storage (Figure 4D). The stability of PSP-SeNPs was higher than that of SeNPs decorated with a hyperbranched polysaccharide from Lignosus rhinocerotis 14. It should be pointed out that SeNPs in the absence of PSP precipitated after 1-day storage (Figure 1). Moreover, the particle size of PSP-SeNPs only increased from 113 to 123 nm after 30 days of storage and the ζ-potential of PSP-SeNPs presented at around −30 mV during the storage time, suggesting that PSP-SeNPs had better stability.
**Figure 4.** Effect of pH (**A**), temperature (**B**), ion strength (**C**), and storage time (**D**) on the average size and ζ-potential of PSP-SeNPs. Values marked with \*: *p* < 0.05, \*\*: *p* < 0.01, and \*\*\*: *p* < 0.001 indicated significant differences when compared to the conditions of pH: 7, temperature: 25 ◦C, NaCl: 0 mM, or storage time: 0 day.
| doab | 2025-04-07T03:56:59.200632 | 17-11-2022 17:23 | {
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007afdec-bed4-405d-873d-c355ba9add0e.100 | *3.5. Characterization and Possible Stabilizing Mechanism of PSP-SeNPs*
The UV-vis spectra of PSP and PSP-SeNPs in the range of 190 to 800 nm were presented in Figure 5A. It was shown that no characteristic absorption peaks were observed on the UV-vis spectra of PSP at the concentration of 0.01 mg/mL. The PSP-SeNPs exhibited wide absorption bands with a maximum absorption peak at about 288 nm. The characteristic absorption peak corresponded to a localized surface plasmon response (LSPR), further demonstrating the formation of nanoparticles [36].
FTIR spectra were performed to clarify the interaction between PSP and SeNPs. In the spectrum of PSP (Figure 5B), the broad absorption band at nearly 3390 cm−<sup>1</sup> was assigned to the O-H stretching vibration. The peak presented at 2927 cm−<sup>1</sup> was attributed to the C-H stretching vibration. The signals that occurred in the region of 1200–1000 cm−<sup>1</sup> were associated with the C-O stretching vibration, indicating the existence of a pyranose ring [37]. The FTIR spectrum of PSP-SeNPs was similar to that of the pure PSP, indicating the presence of PSP on the surface of SeNPs. In addition, the O-H stretching vibration occurred red-shift from 3390 cm−<sup>1</sup> to 3376 cm−1, suggesting the formation of hydrogen bonds between SeNPs and the PSP chains [38]. Based on the above results, we proposed that the interaction mechanism was similar to the combination of arabinogalactans/and SeNPs as described previously [36]. Briefly, the SeO3 <sup>2</sup><sup>−</sup> reacted with the -OH group in the PSP molecule to form special chain-shaped intermediates first, then reduced to the element Se by ascorbic acid. The Se atom further aggregated into the nucleus to form SeNPs as
the reaction processed and the -OH groups of PSP were bound to the surface of SeNPs to prevent the aggregation of nanoparticles.
**Figure 5.** UV-vis spectra (**A**), FTIR spectra (**B**), XPS spectra (**C**), and XPS spectra of Se 3d (**D**) of PSP and PSP-SeNPs.
The XPS spectra were further used to analyze the valence state of selenium. The peaks of Se 3d and 3p orbitals at the binding energy of 55.6 and 179.3 eV (Figure 5C) indicated the zero-valent state of Se within the PSP-SeNPs [10]. As shown in Figure 5D, the peaks of Se 3d5/2 and Se 3d3/2 were up-shifted from 55.1 and 55.9 (SeNPs) to 55.4 and 56.2 (PSP-SeNPs), respectively. The results indicate that the Se 3d orbit participated in the formation of PSP-SeNPs [39], confirming that PSP was successfully conjugated to the SeNPs. Meanwhile, no peak was found at 59.5 eV, which represented the typical Se 3d signal of Se (IV), suggesting that Se (IV) was completely reduced to elemental selenium [40].
| doab | 2025-04-07T03:56:59.200915 | 17-11-2022 17:23 | {
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007afdec-bed4-405d-873d-c355ba9add0e.101 | *3.6. Antioxidant Assays*
The DPPH and ABTS radical scavenging activity were measured in our study to evaluate the antioxidant activity of PSP, SeNPs, and PSP-SeNPs. As shown in Figure 6A, PSP exhibited a low DPPH radical scavenging ability at the tested concentrations. Both SeNPs and PSP-SeNPs had a concentration-dependent DPPH radical scavenging effect at 0.01–1.0 mg/mL. PSP-SeNPs showed a higher scavenging ability than SeNPs. The scavenging effect of PSP-SeNPs reached 59% at the concentration of 1.0 mg/mL, whereas SeNPs could only scavenge 43% DPPH radical at the same concentration. This might be attributed to the enhanced hydrogen-donating ability of PSP-SeNPs to form a stable DPPH-H molecule [41]. Compared to the DPPH radical, all the tested samples performed more efficiently in scavenging ABTS radical (Figure 6B). Similar to the DPPH scavenging assay, the ABTS radical scavenging capacity of PSP-SeNPs was significantly stronger than that of PSP and SeNPs. At 1.0 mg/mL, the scavenging effects of PSP, SeNPs, and PSP-SeNPs were 20%, 62% and 89%, respectively. It has been reported that the DPPH scavenging ability of gum arabic-selenium nanocomposites was lower than 60% at 1.0 mg/mL [42]. The ABTS radical scavenging activity of SeNPs functionalized with a polysaccharide from *Rosa roxburghii* fruit only reached about 50% at 1.0 mg/mL 15. The free radical scavenging ability of PSP-SeNPs synthesized in our study was higher than the above nanoparticles. Moreover, the results showed that the surface decoration of SeNPs with PSP
could remarkably improve the antioxidant activity of SeNPs and PSP. PSP-SeNPs with a smaller size could provide more radical reactive sites due to their larger specific surface area, resulting in higher antioxidant activity [29,43]. However, barely SeNPs were easily aggregated with a decreased active surface to react with the free radicals, further reducing their biological activities [43].
**Figure 6.** Antioxidant activities of PSP, SeNPs, and PSP-SeNPs in vitro. (**A**) DPPH radical scavenging activity. (**B**) ABTS radical scavenging activity. Ascorbic acid (Vc) is used as a positive control. Values marked with \*: *p* < 0.05, \*\*: *p* < 0.01, and \*\*\*: *p* < 0.001 indicated significant differences when compared to SeNPs at the same concentration.
| doab | 2025-04-07T03:56:59.201101 | 17-11-2022 17:23 | {
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007afdec-bed4-405d-873d-c355ba9add0e.102 | *3.7. Effects of PSP-SeNPs on H2O2-Induced PC-12 Cells Toxicity*
Although the free radical scavenging assays proved the excellent antioxidant activity of PSP-SeNPs, the antioxidant assays based on chemical reactions may not necessarily reflect the behavior of antioxidants in biological systems [16]. Thus, the effect of different selenium species on oxidative stress-induced damage to PC-12 cells was further investigated by MTT assay. As depicted in Figure 7A, the cell viability was higher than 90% when incubated with SeNPs and PSP-SeNPs at the concentration of 1–20 μg/mL. However, the cell viability dramatically decreased to 67% after treatment with 20 μg/mL Na2SeO3, suggesting that both SeNPs and PSP-SeNPs showed lower cytotoxicity than Na2SeO3.
**Figure 7.** Effects of sodium selenite (Na2SeO3), SeNPs, and PSP-SeNPs on the viability of PC-12 cells (**A**). Values marked with \*: *p* < 0.05 and \*\*\*: *p* < 0.001 indicated significant differences when compared to the control group. The protective effect against H2O2 (0.5 μM)-induced PC-12 cells toxicity by MTT assay (**B**). Values marked with \*: *p* < 0.05 and \*\*: *p* < 0.01 indicated significant differences when compared to the H2O2 treated group.
The overproduction of reactive oxygen species (ROS) is considered to be the main cause of oxidative damage [44]. Herein, exogenous H2O2 was used as an inducer of cell damage in our model. PC-12 cells incubated with 500 μM H2O2 showed a remarkable decrease of cell viability reaching 56% (Figure 7B). However, the viability of PC-12 cells decreased to 55%, 50%, and 43% when pretreated with Na2SeO3 at concentrations of 1, 10, and 20 μg/mL, respectively. Interestingly, compared with the H2O2-induced oxidative stress model group, cells pretreated with SeNPs or PSP-SeNPs alleviated the H2O2-induced toxicity on PC-12 cells in a concentration-dependent manner, as reflected by the increase in cell viability. The viability of PC-12 cells pretreated with 20 μg/mL SeNPs or PSP-SeNPs significantly increased to 79% and 98%, respectively. In addition, the protective effect of PSP-SeNPs on H2O2-induced oxidative damage on PC-12 cells was better than that of SeNPs. The results confirmed that PSP-SeNPs had excellent antioxidant activity in cells, which may be associated with the free radical scavenging ability.
| doab | 2025-04-07T03:56:59.201231 | 17-11-2022 17:23 | {
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007afdec-bed4-405d-873d-c355ba9add0e.103 | **4. Conclusions**
Our present study provided a facile approach for the synthesis of size-controlled SeNPs by using PSP as a stabilizer in the redox system of sodium selenite and ascorbic acid. The synthesized PSP-SeNPs presented a monodisperse spherical structure with zero-valent Se. The interaction between the hydroxyl groups of PSP chains and the surface of SeNPs contributed to the stable structure of PSP-SeNPs. Furthermore, PSP-SeNPs exhibited stronger free radical scavenging ability and a higher protective effect against H2O2-induced PC-12 cell death than SeNPs. Our findings not only provide the foundations for the utilization of PSP in the development of stable SeNPs but also emphasize the potential application of PSP-SeNPs as an antioxidant in food additives, dietary supplements, and nutraceuticals.
**Author Contributions:** Conceptualization, W.C. and H.C.; methodology, W.C.; software, W.C.; validation, W.C.; formal analysis, W.C. and H.C.; investigation, W.C.; resources, W.X.; data curation, H.C.; writing—original draft preparation, W.C.; writing—review and editing, H.C.; visualization, W.X.; supervision, W.X.; project administration, W.C. and H.C.; funding acquisition, W.C. and H.C. All authors have read and agreed to the published version of the manuscript.
**Funding:** This research was funded by the China Postdoctoral Science Foundation (Grant No. 2021M691288) and the National Natural Science Foundation of China (Grant No. 32101939).
**Institutional Review Board Statement:** Not applicable.
**Informed Consent Statement:** Not applicable.
**Data Availability Statement:** The data presented in this study are available in this manuscript.
**Conflicts of Interest:** The authors declare no conflict of interest.
#### **References**
| doab | 2025-04-07T03:56:59.201363 | 17-11-2022 17:23 | {
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007afdec-bed4-405d-873d-c355ba9add0e.104 | *Article* **Improving Physicochemical Stability of Quercetin-Loaded Hollow Zein Particles with Chitosan/Pectin Complex Coating**
**Muhammad Aslam Khan 1,2, Chufan Zhou 1,2, Pu Zheng 3, Meng Zhao <sup>4</sup> and Li Liang 1,2,\***
**Abstract:** Hollow nanoparticles are preferred over solid ones for their high loading capabilities, sustained release and low density. Hollow zein particles are susceptible to aggregation with a slight variation in the ionic strength, pH and temperature of the medium. This study was aimed to fabricate quercetin-loaded hollow zein particles with chitosan and pectin coating to improve their physicochemical stability. Quercetin as a model flavonoid had a loading efficiency and capacity of about 86–94% and 2.22–5.89%, respectively. Infrared and X-ray diffraction investigations revealed the interaction of quercetin with zein and the change in its physical state from crystalline to amorphous upon incorporation in the composite particles. The chitosan/pectin coating improved the stability of quercetin-loaded hollow zein particles against heat treatment, sodium chloride and in a wide range of pH. The complex coating protected quercetin that was encapsulated in hollow zein particles from free radicals in the aqueous medium and enhanced its DPPH radical scavenging ability. The entrapment of quercetin in the particles improved its storage and photochemical stability. The storage stability of entrapped quercetin was enhanced both at 25 and 45 ◦C in hollow zein particles coated with chitosan and pectin. Therefore, composite hollow zein particles fabricated with a combination of polysaccharides can expand their role in the encapsulation, protection and delivery of bioactive components.
**Keywords:** hollow zein particle; chitosan; pectin; quercetin; coating
| doab | 2025-04-07T03:56:59.201504 | 17-11-2022 17:23 | {
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007afdec-bed4-405d-873d-c355ba9add0e.105 | **1. Introduction**
Hollow zein particles have been fabricated by wrapping sodium carbonate (Na2CO3) nanoprecipitate as sacrificial templet with zein in ethanol–water binary mixture followed by antisolvent precipitation [1]. Hollow particles in the loading and controlled release of bioactive components were preferred over their solid counterpart for more surface area and low density [2], but proteinaceous nature limits their utility as an efficient delivery system due to destabilization around pI (5–6.5), presence of counterion and high temperatureinduced denaturation [3,4]. Numerous strategies have been adopted to overcome instability issues of zein particles, for instance, coating with proteins [5,6], polysaccharides [7] and lipids [8]. Pectin is an anionic biodegradable polymer found in the plant cell wall and mainly made up of methyl esterified 1-4 linked α-D-galacturonic acid and 1-2-linked α-Lrhamnopyranose. Composite hollow zein particles with casein and pectin were developed by heating at 80 ◦C and 6.2 pH for 1 h to attain outstanding stability under simulated gastrointestinal conditions. Still, these composite particles were limited only to encapsulate and deliver heat-sensitive bioactives [9].
Chitosan-coated solid zein particles were fabricated through hydrophobic, hydrogen and van der Waals interactions at pH 4, improving the entrapment, photo/thermal
**Citation:** Khan, M.A.; Zhou, C.; Zheng, P.; Zhao, M.; Liang, L. Improving Physicochemical Stability of Quercetin-Loaded Hollow Zein Particles with Chitosan/Pectin Complex Coating. *Antioxidants* **2021**, *10*, 1476. https://doi.org/10.3390/ antiox10091476
Academic Editor: Elisabetta Esposito
Received: 26 August 2021 Accepted: 13 September 2021 Published: 16 September 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/).
protection and controlled release of bioactive components [10–12]. Chitosan, a N-acetyl-Dglucosamine and D-glucosamine β1-4 linked cationic polymer, is obtained by deacetylation of chitin and considerably used for the stabilization of delivery systems [13]. However, the deprotonation of the amine groups of chitosan above pka (~pH 6.5) reduces charge density, and chitosan competes with counterions with the increase in ionic strength, heading towards destabilization [12,14]. It has been reported that pectin imparted good pH and heating stability to zein core–shell nanoparticles, but the endurance for increasing ionic strength was extremely weak [15,16]. Chitosan and pectin could form polyelectrolyte complexes via electrostatic interaction [17], which was used to improve the physiochemical stability of nanoliposome [18]. Therefore, a combination of chitosan and pectin may synergistically and resourcefully bear variation in pH, temperature and counterions.
Quercetin is a flavonoid with antioxidant, anticarcinogenic, antiviral and anti-inflamm atory properties. Its low solubility in water and chemical instability have been addressed through biopolymer-based nano/micro-delivery systems for the application in functional foods [19,20]. In the current work, composite hollow zein particles were fabricated with chitosan-pectin complex coating for the encapsulation and protection of quercetin. The particles were characterized for size, ζ-potential and loading efficiency of quercetin. The lyophilized samples of quercetin-loaded composite particles were analyzed with infrared and X-ray diffraction techniques. Moreover, the particle dispersions were subject to varying pH, ionic strength and temperature conditions to assess physical stability. Finally, the antioxidant activity, photochemical and storage stability of quercetin encapsulated in composite hollow zein particles were examined to judge the protective effects of the particles. This study focused on the fabrication of composite hollow zein particles with improved physical stability and better protective effect on flavonoids through tailoring a complex polysaccharide interfacial layer.
| doab | 2025-04-07T03:56:59.201641 | 17-11-2022 17:23 | {
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007afdec-bed4-405d-873d-c355ba9add0e.106 | **2. Materials and Methods**
#### *2.1. Materials*
Zein from corn (~98%) was purchased from J&K Chemical Ltd. (Shanghai, China). Sodium carbonate (Na2CO3, ~99.8%) was purchased from Sinopharm Chemical Reagent Co., Ltd. (Shanghai, China). Pectin from citrus peel was purchased from Shanghai Sangong Bioengineering Co., Ltd. (Shanghai, China). Chitosan (low molecular weight, 50–190 KDa) and quercetin (≥95%, HPLC) were purchased from Sigma-Aldrich Co. (Shanghai, China). Ultra-pure water obtained using a Milli-Q direct water purification system equipped with Quantum TEX column (Molsheim, France) was used throughout all the experiments.
| doab | 2025-04-07T03:56:59.201914 | 17-11-2022 17:23 | {
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007afdec-bed4-405d-873d-c355ba9add0e.107 | *2.2. Preparation of Blank and Quercetin-Loaded Hollow Zein, Hollow Zein-Chitosan, and Hollow Zein-Chitosan/Pectin Particles*
Hollow zein (HZ) particles were fabricated with sodium carbonate sacrificial templet by mixing 1.75 mL of absolute ethanol and 2.5 mL of 50 mg/mL zein in 70% (*v/v*) ethanol– water binary mixture with 0.75 mL of 0.5, 1 and 2 (*w/v*%) Na2CO3 aqueous solution under magnetic stirring at 1000 rpm for 1 min followed by adding to a 20 mL ultra-pure water [1,21]. The HZ particles were mixed with 0.5, 1 and 2 mg/mL chitosan in 1% (*v/v*) acetic acid at a 1:1 volume ratio under stirring at 1000 rpm for 30 min. Ethanol in the particle dispersion was removed under vacuum with a rotary evaporator RE-52C (Shanghai Tianheng Instrument Co. Ltd., Shanghai, China) at 35 ◦C for 30 min. Unstable particles were separated by centrifugation the particle dispersion at 2000× *g* for 15 min; the supernatant was then centrifuged at 15,000× *g* for 30 min to remove unabsorbed chitosan, and the precipitate was redispersed in an equal volume of distilled water to obtained chitosan-coated hollow zein (HZ-chi) particles. The aqueous solutions of pectin at 0.01, 0.025, 0.05 and 0.1 mg/mL were added to the dispersion of HZ-chi particles under stirring for 30 min at pH 4–4.5 to obtain pectin- and chitosan-coated hollow zein (HZ-chi/pec) particles. The quercetin-loaded hollow particles were prepared by adding 2, 3, 4 and 5 mg of
quercetin in 50 mg/mL zein stock solution in ethanol–water binary mixture corresponding to 100, 150, 200 and 250 μg/mL of quercetin in the final particle dispersion.
| doab | 2025-04-07T03:56:59.201970 | 17-11-2022 17:23 | {
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007afdec-bed4-405d-873d-c355ba9add0e.109 | 2.3.1. Particle Size and ζ-Potential
Samples were diluted by 200 folds with distilled water and measured at 25 ◦C and analyzed on a NanoBrook Omini particle size analyzer (Brookhaven Instrument, New York, NY, USA) at a scattering angle of 90◦. NNLS function was used to acquire the size distribution, while phase analysis light scattering (PALS) was employed to estimate the ζ-potential. Samples were prepared in triplicates for each measurement.
| doab | 2025-04-07T03:56:59.202089 | 17-11-2022 17:23 | {
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007afdec-bed4-405d-873d-c355ba9add0e.110 | 2.3.2. Loading Efficiency and Loading Capacity of Quercetin
Quercetin-loaded hollow zein and hollow zein-chitosan particles were centrifuged at 2000× *g* for 15 min to remove unencapsulated quercetin, and the supernatant and samples were diluted 50-fold in ethanol for the measurement of quercetin absorption at a λmax of 373 nm using a UV1800 UV-Vis spectrophotometer (Shimadzu Corporation, Tokyo, Japan) with a standard curve constructed from 1 to 20 μg/mL of quercetin dissolved in ethanol (a correlation coefficient of 0.999, Figure S4), adopting the method of Wang et al. [22]. Samples were prepared in triplicates for each measurement. Loading efficiency and capacity of quercetin in the particles were determined by the following equations.
$$\text{Loading efficiency} \left( \% \right) = \frac{\text{Queretion in supernantant}}{\text{Total added queue}} \times 100 \tag{1}$$
$$\text{Loading capacity} \left( \% \right) = \frac{\text{Quercetin entrapped in particles } \left( \mu \text{g} \right)}{\text{Zein and chipsan } \left( \mu \text{g} \right)} \times 100 \tag{2}$$
| doab | 2025-04-07T03:56:59.202142 | 17-11-2022 17:23 | {
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007afdec-bed4-405d-873d-c355ba9add0e.111 | 2.3.3. Microstructural Analysis
Lyophilized particles were mounted to the surface of double-sided carbon tape and coated with a thin layer of gold. The morphology of particles was observed with a Hitachi SU8010 FE-SEM (Hitachi, Co., Tokyo, Japan) operated at an accelerating voltage of 8 kV.
| doab | 2025-04-07T03:56:59.202219 | 17-11-2022 17:23 | {
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007afdec-bed4-405d-873d-c355ba9add0e.112 | 2.3.4. Infrared Spectroscopy (IR)
Samples were freeze-dried and pressed into a transparent pellet with KBr, Infrared spectra of particles and raw materials were collected in the range of 400–4000 cm−<sup>1</sup> on Nicolet™ iS™ 10 FT-IR Spectrometer (Thermo Fisher Scientific, Waltham, MA, USA) at a 4 cm−<sup>1</sup> resolution and 16 scans.
| doab | 2025-04-07T03:56:59.202259 | 17-11-2022 17:23 | {
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007afdec-bed4-405d-873d-c355ba9add0e.113 | 2.3.5. X-ray Diffraction (XRD)
A Bruker D2 PHASER (Brucker, Odelzhausen, Germany) X-ray diffractometer, operated at 30 kV, 10 mA was used to obtain X-ray diffractograms of quercetin, zein, chitosan, pectin and quercetin-loaded hollow zein particles coated with chitosan and pectin. The data were collected over an angular range from 5◦ to 40◦ 2θ in continuous mode using step size and time of 0.02◦ and 5 s, respectively.
| doab | 2025-04-07T03:56:59.202302 | 17-11-2022 17:23 | {
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007afdec-bed4-405d-873d-c355ba9add0e.114 | *2.4. Stability Assessment of Particles under Stressed Condition*
| doab | 2025-04-07T03:56:59.202345 | 17-11-2022 17:23 | {
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007afdec-bed4-405d-873d-c355ba9add0e.115 | 2.4.1. Sodium Chloride Stability
Freshly prepared quercetin-loaded HZ, HZ-chi and HZ-chi/pec particles were exposed to 50, 100, 200, 300, 400, 500 mM sodium chloride under continuously mixing for 30 min followed by a 5 min rest [8]. Particle size and ζ-potential were measured, as mentioned above in Section 2.3.1.
| doab | 2025-04-07T03:56:59.202370 | 17-11-2022 17:23 | {
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007afdec-bed4-405d-873d-c355ba9add0e.116 | 2.4.2. pH Stability
The pH values of quercetin-loaded HZ, HZ-chi and HZ-chi/pec particles were adjusted from 2 to 9 with 0.1 mM hydrochloric acid and sodium hydroxide and then continuously stirred for 30 min [8]. Particle size and ζ-potential were analyzed by diluting them in pH-adjusted Milli-Q water [23].
## 2.4.3. Temperature Durability
Quercetin-loaded HZ, HZ-chi and HZ-chi/pec particles were incubated in a water bath at 30, 40, 50, 60, 70, 80 and 90 ◦C for 30 min and evaluated for particle size and ζ-potential [8].
| doab | 2025-04-07T03:56:59.202408 | 17-11-2022 17:23 | {
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007afdec-bed4-405d-873d-c355ba9add0e.117 | *2.5. Antioxidant Activity*
ABTS and DPPH assays of quercetin-loaded HZ and HZ-chi/pec particles were estimated according to the method of Dong et al. and Pan et al. [24,25]. In brief, equal volumes of potassium persulfate (2.6 mM) and ABTS (7.4 mM) were mixed and allowed to react and generate ABTS+ for 12 h in the dark. After the diluted ABTS+ solution with an absorbance of 0.7 at 734 nm was mixed with samples at a volume ratio of 2:1 in the dark for 6 min, the absorbance was measured at 734 nm on Synergy H1 Microplate Reader (BioTek Instruments, Inc., Winooski, VT, USA). Likewise, after 0.1 mM DPPH in ethanol was added to samples in equal volumes and allowed to react for 30 min in the dark, the absorbance was recorded at 517 nm. The ABTS<sup>+</sup> and DPPH scavenging capacity was calculated with the help of the following equation,
$$\text{Free radical scavering capacity (\%)} = (\text{A}\_{\text{Control}} - \text{A}\_{\text{Sample}}) / \text{A}\_{\text{Control}} \times 100 \tag{3}$$
where AControl and ASample are the absorbances of the free radical solution without and with samples, respectively.
| doab | 2025-04-07T03:56:59.202554 | 17-11-2022 17:23 | {
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007afdec-bed4-405d-873d-c355ba9add0e.118 | *2.6. Functional Characteristics of Particles*
#### 2.6.1. Photochemical Stability of Quercetin
Photochemical stability of pristine and encapsulated quercetin was evaluated by the procedure presented by Sun et al. [26]. Quercetin dispersed in water and encapsulated in HZ and HZ-chi/pec particles at 5 μg/mL were irradiated up to 120 min with a 365 nm ultraviolet lamp (VWR International Inc., West Chester, PA, USA). Samples were collected at 0, 15, 30, 60, 90 and 120 min, and the content of quercetin was analyzed with the help of UV-Vis spectrophotometer mentioned above in Section 2.3.2.
| doab | 2025-04-07T03:56:59.202916 | 17-11-2022 17:23 | {
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007afdec-bed4-405d-873d-c355ba9add0e.119 | 2.6.2. Storage Stability of Quercetin and Particles
Samples were stored at 25 and 45 ◦C for 28 days inside an LRH-250F incubator (Yiheng Scientific Instrument Co., Ltd. Shanghai, China). The stability of quercetinloaded particles was analyzed in terms of particle size and ζ-potential during storage. The retention of quercetin was expressed as percent retention and calculated by using the following equation:
$$\text{Quercective iteration (\%)} = \text{Q}\_{\text{l}}/\text{Q}\_{\text{i}} \times 100\tag{4}$$
where Qi and Qt are the content of quercetin at the beginning and specific time intervals during storage.
| doab | 2025-04-07T03:56:59.202973 | 17-11-2022 17:23 | {
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007afdec-bed4-405d-873d-c355ba9add0e.120 | *2.7. Statistical Analysis*
All experiments were done in triplicates. The results were expressed in mean plus standard deviation and analyzed for a significant difference (*p* < 0.05) with IBM SPSS statistics 20.0 software package (IBM, Armonk, NY, USA).
| doab | 2025-04-07T03:56:59.203033 | 17-11-2022 17:23 | {
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007afdec-bed4-405d-873d-c355ba9add0e.121 | **3. Results and Discussion**
*3.1. Characterization of Hollow Zein Particles*
3.1.1. Effect of Na2CO3 and Chitosan on Hollow Zein Particles
The size of hollow zein particles is greatly influenced by the concentration of Na2CO3 used for the preparation of sacrificial templet [1,27]. The smallest particles of 76 nm were fabricated with 1% Na2CO3 (Figure 1A). At 0.5 and 2% Na2CO3, the size of HZ particles was 145 and 210 nm, respectively. The reason for the bigger particles is due to the formation of a thicker zein shell around all the available sodium carbonate nuclei formed in ethanolic conditions at a Na2CO3 concentration of 0.5% but the formation of bigger Na2CO3 nanocrystal in size due to excessive aggregation and crystal growth at 2% [27]. The pH values of the HZ particle dispersions prepared with 0.5, 1 and 2% Na2CO3 were 9.04, 10.31 and 10.77, respectively, which is above the pI of zein [28]. The ζ-potential of HZ particles was −8, −26 and −24 mV when prepared with 0.5, 1 and 2% Na2CO3 (Figure 1B), receptively. There was a slight increase in the particle size of HZ particles upon the addition of chitosan (Figure 1A), except that precipitation was observed at 0.05% chitosan and 2% Na2CO3. The increase in the particle size was the most obvious when the chitosan concentration was 0.1% at 1% and 2% Na2CO3. In the presence of chitosan, the pH of particle dispersions shifted to 4.0–4.5. Chitosan interacts with proteins below their pI through hydrophobic, electrostatic, van der Waals and hydrogen bonding [12,29]. The ζ-potential of HZ-chi particles ranged between +30 and +58 mV (Figure 1B). The amine groups in chitosan contribute to the positive surface charge of composite zein particles [30,31]. These results suggest the formation of a chitosan shell.
**Figure 1.** Particle size distribution (**A**) and ζ−potential (**B**) of hollow zein particles fabricated with 0.5% (black), 1% (red) and 2% (blue) Na2CO3 and coated with chitosan at various concentrations (Mark at the right). \* indicates unstable particles.
| doab | 2025-04-07T03:56:59.203070 | 17-11-2022 17:23 | {
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007afdec-bed4-405d-873d-c355ba9add0e.122 | 3.1.2. Fabrication of Pectin-Coated Composite Particles
HZ-chi particles prepared with 1% Na2CO3 were further coated with pectin to form a second layer. The addition of pectin did not influence particle PDI (Table 1). When the concentration of chitosan was 0.5 mg/mL, the size of HZ-chi particles increased from 87 to 170 and 180 nm upon adding 0.01 and 0.025 mg/mL of pectin, respectively. When the concentration of chitosan was 2 mg/mL, the size of HZ-chi particles increased from 84 to 194 and 192 nm upon adding 0.01 and 0.025 mg/mL of pectin, respectively. Meanwhile, a reduction in the ζ-potential positive values was observed, showing insufficient pectin to fully cover the surface of particles. With further increasing the pectin concentration to 0.05 and 0.1 mg/mL in the presence of 0.5 mg/mL chitosan and to at 0.05 mg/mL pectin in the presence of 2 mg/mL, the particle dispersion became unstable with the formation of precipitates, due to charge neutralization [7]. The neutralization was observed at 0.01 and 0.025 mg/mL pectin in the presence of 1 mg/mL chitosan (Table 1). However, in the presence of 1 mg/mL chitosan, an increment in size to 248 and 219 nm was observed
at 0.05 and 0.1 mg/mL pectin, respectively, and ζ-potentials changed to negative values, suggesting the formation of pectin surface layer. An absolute value of ζ-potential above 20 mV is so high enough to ensure the physical stability of biopolymer-based particles [32]. Therefore, the hollow zein particles prepared with 1 mg/mL chitosan and 0.1 mg/mL pectin were used for further study on their physicochemical and functional attributes.
**Table 1.** Effect of pectin on size, PDI and ζ-potential of hollow zein particles coated with chitosan.
Values with different letters (upper case A and B for the concentration of chitosan; lower case a, b and c for the concentration of pectin) are significantly different in rows (*p* < 0.05); – represents the aggregation followed by precipitation of nanoparticles.
| doab | 2025-04-07T03:56:59.203202 | 17-11-2022 17:23 | {
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007afdec-bed4-405d-873d-c355ba9add0e.123 | 3.1.3. Encapsulation of Quercetin
Quercetin was used as a model bioactive flavonoid to be encapsulated in hollow zein particles. The loading efficiencies of quercetin were 72.71–79.86% in HZ particles, while the loading efficiencies increased to 90.58–93.86% at the flavonoid concentrations of 100–200 μg/mL and 85.84% at 250 μg/mL in HZ-chi particles (Table 2). The higher loading efficiency of quercetin in the presence of chitosan is attributed to its interaction with chitosan through electrostatic and hydrogen bonding, in addition to hydrophobic interaction with zein [33,34]. The loading capacity of quercetin in the absence and presence of chitosan remained similar and increased from about 2% to 6%. This is different from a decreasing trend that was commonly observed in composite protein particles with increasing the content of the wall materials [35,36].
**Table 2.** Loading efficiency and loading capacity of quercetin in hollow zein and zein-chitosan particles.
Values with different letters (upper case A, B and C for the concentration of quercetin in column; lower case a and b in the row for the type of particles) are significantly different (*p* < 0.05).
<sup>250</sup> 72.71 <sup>±</sup> 3.82 Aa 85.84 <sup>±</sup> 4.16 Bb 6.29 <sup>±</sup> 0.98 Aa 5.89 <sup>±</sup> 1.08 Aa
SEM images showed that quercetin-loaded HZ, HZ-chi and HZ-chi/pec were spherical with a smooth surface (Figure 2). The loading of quercetin increased the size of HZ particles without and with chitosan and/or pectin coating (Figure S1A). A similar trend was previously reported for the encapsulation of resveratrol and curcumin in composite hollow zein particles [11,37]. The loading of quercetin did not affect the ζ-Potential of HZ particles but increased ζ-potential absolute values of HZ-chi and HZ-chi/pec particles (Figure S1C), possibly due to rearrangement and exposure of more charged groups upon flavonoid inclusion [33].
**Figure 2.** SEM images of quercetin-loaded hollow zein (qHZ, (**A**)), zein-chitosan (qHZ-chi, (**B**)) and zein-chitosan/pectin (qHZ-chi/pec, (**C**)) particles. The concentrations of chitosan and pectin were 1 and 0.1 mg/mL, respectively.
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007afdec-bed4-405d-873d-c355ba9add0e.124 | *3.2. Physical Characterization* 3.2.1. XRD
Quercetin is a crystalline solid in its pristine form [38], as evident from sharp X-ray diffraction patterns at 8, 10, 12, 13.01, 15, 17 and 27 of diffraction angle (2θ) (Figure 3). The distinct peaks of quercetin with lower intensities were apparent in its mixture of zein, chitosan and pectin. In the diffractograms of qHZ, qHZ-chi and qHZ-chi/pec particles, the crystalline peaks of quercetin disappeared, advocating the transformation from crystalline to an amorous state upon encapsulation in particles [10,39]. Pure zein and chitosan showed mild crystallinity with moderate peaks at 9.56 and 20.24 2θ because of respective α-helixes and crystal lattice structure [40,41]. When zein was structured into HZ, HZ-chi, HZ-chi/pec, qHZ, qHZ-chi and qHZ-chi/pec particles, the diffraction pattern raised from α-helixes in zein and crystal lattice of chitosan became flattened due to interaction between the polymers during the formation of the composite particles [42,43].
**Figure 3.** XRD diffraction patterns of quercetin, mixture (quercetin, zein, chitosan and pectin), zein, chitosan, pectin, HZ, HZ-chi, HZ-chi/pec, qHZ, qHZ-chi, qHZ-chi/pec.
| doab | 2025-04-07T03:56:59.203450 | 17-11-2022 17:23 | {
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007afdec-bed4-405d-873d-c355ba9add0e.125 | 3.2.2. IR Spectroscopy
OH stretching of zein and HZ particles was at 3318 cm−<sup>1</sup> but changed to 3316 and 3405 cm−<sup>1</sup> in HZ-chi and HZ-chi/pec particles, indicating hydrogen bonding among zein, chitosan and pectin (Figure 4B,C). Likewise, amide I and II absorption band of zein changed from 1658 and 1541 cm−<sup>1</sup> to 1656 and 1536 cm−<sup>1</sup> in HZ particles, to 1658 and 1542 cm−<sup>1</sup> in HZ-chi particles, and to 1654 and 1635 cm−<sup>1</sup> in HZ-chi/pec particles. The variations of stretching and bending vibrations of C=O and N-H indicate that hydrophobic and electrostatic interactions occurred during the fabrication of HZ, chitosan and chitosan/pectin coated HZ particles [29]. The distinct absorption peaks of quercetin IR spectrum at 3386, 1655, 1599, 1373, 1257 and 1160 cm−<sup>1</sup> (Figure 4A) signify O-H, C=O, C=C, C-OH, C-O-C
and C-OH (B ring), respectively [34,44]. The absorption bands were clearly seen with lower intensities in the physical mixture of quercetin, zein, chitosan and pectin (Figure 4A). However, the characteristic absorption bands of quercetin were invisible in HZ, HZ-chi and HZ-chi/pec particles, indicating the entrapment of quercetin in the particles and limited starching and bending of various bonds [42]. The encapsulation of quercetin resulted in the variation of OH stretching of HZ, HZ-chi and HZ-chi/pec particles (Figure 4A,B), suggesting the flavonoid interaction with wall materials [45]. Upon encapsulation of quercetin in hollow zein particle, the amide I peak of zein was unchanged, whereas the amide II showed a redshift to 1537, 1543 and 1539 cm−<sup>1</sup> in HZ, HZ-chi and HZ-chi/pec particles (Figure 4A). These shifts in the amide II result from hydrophobic, electrostatic and hydrogen bonding of quercetin with wall materials [46,47].
**Figure 4.** IR spectra of quercetin, mixture (quercetin, zein, chitosan and pectin), quercetin-loaded HZ−chi and HZ−chi/pec particles (**A**), HZ, HZ−chi and HZ−chi/pec blank particles (**B**), and raw materials zein, chitosan and pectin (**C**).
| doab | 2025-04-07T03:56:59.203533 | 17-11-2022 17:23 | {
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007afdec-bed4-405d-873d-c355ba9add0e.126 | *3.3. Stability of Hollow Particles*
## 3.3.1. Salt Endurance
Colloidal stability against salt was assessed by exposing particles to various concentrations of NaCl. Quercetin-loaded hollow zein particles without and with chitosan were highly unstable and precipitated out even at 50 mM NaCl (Figure S2A,B), attributed to the screening effect and charge neutralization by counterions [7,43]. Quercetin-loaded HZchi/pec showed a remarkable endurance to precipitation up to 500 mM NaCl (Figure S2C). The size of quercetin-loaded HZ-chi/pec particles kept a single peak and increased gradually as the concentration of NaCl increased up to 200 mM (Figure 5A). Meanwhile, their ζ-potential changed to −29 mV (Figure 5B). It is possible that the NaCl ions screen the repulsion among the polysaccharide chain, leading to a greater adsorption of pectin on the particle surface [48,49]. These results are different from the sedimentation of pectin-coated zein particle at 70 mM NaCl reported by Huang et al. [16]. It can be thus speculated that the complex coating of chitosan and pectin provided better stability against salt than did by pectin alone. The size of quercetin-loaded HZ-chi/pec particles became bigger and had two peaks (Figure 5A), and their ζ-potential absolute values significantly decreased upon further increasing the concentration of NaCl. These results suggest the salt at high concentrations reduced the electrostatic repulsion between particles, resulting in their aggregation [46].
**Figure 5.** Size distribution (**A**) and ζ−potential (**B**) of quercetin−loaded hollow zein particles coated with chitosan and pectin as a function of NaCl concentration.
| doab | 2025-04-07T03:56:59.203654 | 17-11-2022 17:23 | {
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007afdec-bed4-405d-873d-c355ba9add0e.127 | 3.3.2. pH Stability
Protein-based particles are liable to aggregation around pI [6], limiting their application. Quercetin-loaded hollow zein particles were sable with similar size distribution at pH 2, 3 and 4 (Figure 6A) but precipitated at pH 5 around pI, due to charge neutralization and lack of repulsive forces [50,51]. It is difficult to re-disperse the precipitated zein particles above the pI of zein. Quercetin-loaded hollow zein-chitosan particles showed two size distributions above and below pH 4 (Figure 6A). The change in the particle size below pH 4 is attributed to swelling and dissolution of the polymers with higher charge density [18], while the formation of bigger aggregates is at pH 5, close to the pI of zein [7]. The particles with chitosan were unstable at pH 6–9 due to the deprotonation of chitosan amine groups at pka and above [52,53]. It is noted that quercetin-loaded HZ-chi/pec particles showed good stability across the investigated pH range of 2–5 (Figure 6A). The ζ-potential of quercetin-loaded HZ-chi/pec was the highest at pH 5 and decreased as pH decreased (Figure 6B), due to deionization of the carboxylic groups of pectin below its pka (3.5) [16]. At pH 6–9, the size of quercetin-loaded HZ-chi/pec particles was around 220 nm and less than those at lower pH. Their ζ-potential also decreased slightly as the pH increased from pH 5. Karim et al. also reported a decline in the ζ-potential and size of pectin/chitosan-coated nanoliposome at pH 8 compared with that of pH 5 [18]. These changes might be due to the detachment of loosely adsorbed pectin molecules from the surface, since chitosan possesses less charged groups as the pH increases [54]. These findings suggest that the double coating with chitosan and pectin provides excellent protection against aggregation, especially around the pI of zein and pka of chitosan.
**Figure 6.** Size distribution (**A**) and ζ−potential (**B**) of quercetin−loaded hollow zein particles without (blue) and with chitosan (red) and chitosan/pectin (black) coating at various pH values.
| doab | 2025-04-07T03:56:59.203760 | 17-11-2022 17:23 | {
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007afdec-bed4-405d-873d-c355ba9add0e.128 | 3.3.3. Temperature Stability
Bioactive-component-loaded carrier systems for food application might be exposed to different temperature treatments during the production cycles. Figure 7 shows the effect of temperature on the size distribution and ζ-potential of quercetin-loaded HZ, HZ-chi and HZ-chi/pec particles. Quercetin-loaded HZ particles had ζ-potential values between +41–+48 mV (Figure 7B). Size of quercetin-loaded HZ particles increased as temperature increased and showed two peaks above 40 ◦C (Figure 7A), possibly attributed to rearrangement of zein molecules and exposure of nonpolar groups followed by a collapse of hollow structure [55]. This is different from solid zein particles, those were colloidally stable when heated at pH 4 and 80 ◦C for 120 min [43]. Likewise, Figure 7A shows that size of quercetin-loaded HZ-chi particles increased with increasing temperature, but two peaks were observed above 70 ◦C, suggesting that the chitosan coating improves the particle stability against heat treatment. Their ζ-potential was +32 mV at 30 ◦C and increased to a range of +45 and +48 mV at 40–80 ◦C and to +56 mV at 90 ◦C. The changes might be due to that the realignment of zein and chitosan at higher temperatures possibly leads to inter/intramolecular interactions [5,56]. The quercetin-loaded HZ-chi/pec particles were stable when the temperature was increased to 40 ◦C (Figure 7A). Then, their size gradually increased upon further increase in temperature and showed two peaks at 90 ◦C. Their ζ-potential was kept between −25 and −32 mV at all the investigated temperatures. These results indicate that the pectin coating further improves the colloidal stability of quercetin-loaded HZ-chi particles. It is presumed that steric stabilization of pectin coating inhibits the collision of zein particles with more exposed reactive functional groups at the higher temperature, thus preventing increment in size and particle aggregation [57].
**Figure 7.** Effect of temperature on size distribution qHZ (blue), qHZ−chi (red) and qHZ−chi/pec (black) (**A**) and ζ−potential (**B**) of quercetin−loaded composite hollow zein particles.
| doab | 2025-04-07T03:56:59.203888 | 17-11-2022 17:23 | {
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007afdec-bed4-405d-873d-c355ba9add0e.129 | 3.3.4. Storage Stability
After storage at 25 and 45 ◦C for 7 days, quercetin-loaded HZ and HZ-chi particles precipitated, while no precipitation was observed for quercetin-loaded HZ-chi/pec particles (Figure S3). Figure 8 shows size distribution and ζ-potential of quercetin-loaded HZ-chi/pec particles during storage. Their size distribution remained a single peak during storage at 45 ◦C and increased to around 325 nm after 6 days. At 25 ◦C, the size distribution kept a single peak around 220 nm after 6 days and then showed two peaks. These results indicate quercetin-loaded HZ-chi/pec particles are more stable at 45 ◦C than 25 ◦C, possibly due to the high temperature facilitates adsorption of pectin to the particles more effectively and improves its complex formation capacity [58,59]. At 25 ◦C, a smaller size, around 90 nm was observed after storage for 13 days (Figure 8), probably due to the detachment, depolymerization and hydrolysis of galacturonic acid glycan chains of pectin at pH<5[60]. At 25 ◦C, a larger size around 900–1500 nm was observed after 20 days (Figure 8). The ζ-potential of HZ-chi/pec particles became more negative over time, both at 25 and 45 ◦C. These changes indicate the realignment of pectin during storage and exposure of more charged COO- groups at the interface, leading to an upsurge in ζ-potential [18,54]. Pectin
coating substantially enhances the colloidal stability of quercetin-loaded HZ-chi/pec compared with that of qHZ and qHZ-chi (Figure S3A,B) by preventing aggregation of particles through electrostatic repulsions and steric stabilization as well [61].
**Figure 8.** Size distribution (**A**) and ζ−potential (**B**) of quercetin−loaded hollow zein particles coated with chitosan and pectin at 25 ◦C (red) and 45 ◦C (black) during storage.
| doab | 2025-04-07T03:56:59.204012 | 17-11-2022 17:23 | {
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007afdec-bed4-405d-873d-c355ba9add0e.130 | *3.4. Antioxidant Activity*
The antioxidant activity of encapsulated quercetin was investigated by ABTS and DPPH radical scavenging assays (Figure 9). The ABTS+ scavenging capacity of quercetinloaded HZ was lower than the quercetin dispersed in water and ethanol. The reason for the lower scavenging capacity of encapsulated quercetin is being embedded in the hydrophobic pockets of protein and inaccessible to ABTS+ [30]. Furthermore, the hydroxyl groups in the B ring of quercetin are the main contributor of H+ and are involved in hydrogen bonding in the composite particles, thus unavailable to scavenge the free radicals. Earlier, quercetin encapsulated in SPI and solid zein particles showed a similar reduction in the ABTS<sup>+</sup> savaging capacity. On the other hand, the interfacial chitosan-pectin coating in quercetin-loaded HZ-chi/pec particles may increase the accessibility of hydrophilic ABTS+ to quercetin, demonstrating higher scavenging of ABTS<sup>+</sup> compared with that of qHZ (Figure 9). DPPH scavenging capacity of quercetin was improved upon encapsulation in HZ and HZ-chi/pec particles by compared with that of dispersed in ethanol and water (Figure 9). Earlier, it has been reported that both zein and DPPH being soluble in ethanol–water binary medium facilitate scavenging of DPPH by hydrophobic antioxidants encapsulated in the zein particles [62,63]. The ABTS and DPPH radical scavenging assay revealed that by encapsulating quercetin in composite hollow zein particles, it could be better protected from free reactive radicals in the surrounding medium along with an improved or sustained antioxidant activity.
**Figure 9.** ABTS+ and DPPH scavenging capacity of quercetin, blank and quercetin-loaded HZ and HZ-chi/pec particles.
| doab | 2025-04-07T03:56:59.204104 | 17-11-2022 17:23 | {
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007afdec-bed4-405d-873d-c355ba9add0e.131 | *3.5. Stability of Quercetin*
### 3.5.1. Photochemical Stability
Quercetin is prone to degradation upon exposure to ultraviolet light due to oxidative decarboxylation of the C ring [22]. The retention of unencapsulated quercetin sharply decreased to 25% after 120 min of irradiation (Figure 10A). The quercetin loaded in HZ and HZ-chi/pec particles was more stable, with retention of about 80% after 120 min of irradiation. These results suggesting the hollow particle provide the excellent stability of quercetin against ultraviolet light. The protection results from the physical barrier and light-scattering effect of the particles [61]. The similar retention of quercetin in HZ and HZchi/pec particles (Figure 10A) indicates that the protection was fundamentally attributed to zein. A comparable protective effect has previously been reported by encapsulating quercetin in WPI/lotus root amylopectin, pea protein-isolated and zein/soluble soybean polysaccharide composite nanoparticles due to hydrogen and hydrophobic interaction between quercetin and proteins [39,43,64].
**Figure 10.** Retention of quercetin free and encapsulated in hollow zein particles coated without (qHZ) and with (qHZ-chi/pec) 1 mg/mL chitosan and 0.1 mg/mL pectin under irradiation (**A**) and during storage at 25 ◦C and 45 ◦C (**B**).
#### 3.5.2. Storage Stability
Liu and coworkers reported a drastic decrease in the content of quercetin to around 38% in 3 days of storage at room temperature [39]. The degradation of quercetin was faster at 45 ◦C than 25 ◦C, with the retention of quercetin being 42% and 16% after 2 days (Figure 10B), respectively. Its complete loss was observed at 25 ◦C and 45 ◦C after 4 days. The degradation of quercetin is attributed to its auto-oxidation in the aqueous medium, which is much pronounced at elevated temperature [19,65]. The retention of quercetin was significantly greater in HZ-chi/pec particles during storage, being 84% and 67% at 25 and
45 ◦C after 27 days (Figure 10B), respectively. The ABTS<sup>+</sup> scavenging capacity of quercetinloaded HZ-chi/pec particles was lower than quercetin alone (Figure 9), indicating that the entrapment of quercetin reduces its accessibility to ABTS+ [30]. It is thus speculated that the entrapment in HZ-chi/pec particles (Table 2) provides a physical barrier between quercetin and environment-sensitive factors, resulting in the improved stability of quercetin (Figure 10B). Moreover, the hydrophobic interaction and hydrogen bond of quercetin with excipient biopolymer (Figure 4) may inhibit the flavonoid autoxidation and prolong its storage life.
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007afdec-bed4-405d-873d-c355ba9add0e.132 | **4. Conclusions**
The hollow zein particles coated with chitosan and pectin were prepared with 1% Na2CO3 as a sacrificial template. The hollow particles coated with 1 mg/mL chitosan and 0.1 mg/mL pectin had a size of 219 nm and ζ-potential of −28 mV. Chitosan coating improved the loading efficiency of quercetin in hollow zein particles. The coating of chitosan/pectin improved the stability of quercetin-loaded hollow zein particles against heat treatment, pH variation and salt. The entrapment in the hollow particles improved the photostability and storage stability of quercetin. The storage stability was better at 25 ◦C for entrapped quercetin but at 45 ◦C for hollow zein particles coated with chitosan and pectin. These findings will extend the application of composite hollow zein particles for the incorporation of bioactive components in functional products.
**Supplementary Materials:** The following are available online at https://www.mdpi.com/article/10 .3390/antiox10091476/s1, Figure S1: Size distribution (A) and ζ-potential (B) of hollow zein (HZ) particles coated with chitosan (HZ-chi) and with chitosan and pectin (HZ-chi/pec) without and with quercetin. Figure S2: Visual appearance of quercetin-loaded hollow zein particles (A) coated with chitosan (B) and with chitosan and pectin (C) at 50–500 mM NaCl. Figure S3: Visual appearance of qHZ, qHZ-chi, qHZ-chi/pec at 25 ◦C (A), 45 ◦C (B). Figure S4: Calibration curve of quercetin.
**Author Contributions:** M.A.K. and C.Z.; Conceptualization, Investigation, Writing—original draft preparation, Writing—review and editing, P.Z. and M.Z.; Writing—review and editing, L.L.; Writing review and editing, Reviewing, Supervision. All authors have read and agreed to the published version of the manuscript.
**Funding:** This work received support from the National Natural Science Foundation of China (NSFC Project 31571781).
**Institutional Review Board Statement:** Not applicable.
**Informed Consent Statement:** Not applicable.
**Data Availability Statement:** The data presented in this study are available in this manuscript.
**Conflicts of Interest:** The authors declare no conflict of interest.
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007afdec-bed4-405d-873d-c355ba9add0e.134 | *Article* **Whey Protein Isolate-Xylose Maillard-Based Conjugates with Tailored Microencapsulation Capacity of Flavonoids from Yellow Onions Skins**
**S, tefania Adelina Milea, Iuliana Aprodu, Elena Enachi, Vasilica Barbu, Gabriela Râpeanu, Gabriela Elena Bahrim and Nicoleta Stănciuc \***
> Faculty of Food Science and Engineering, Dunarea de Jos University of Galati, 111 Domnească Street, 800201 Galat,i, Romania; [email protected] (S, .A.M.); [email protected] (I.A.); [email protected] (E.E.); [email protected] (V.B.); [email protected] (G.R.); [email protected] (G.E.B.)
**\*** Correspondence: [email protected]
**Abstract:** The objective of this study is to encapsulate flavonoids from yellow onion skins in whey protein isolates (WPI) and xylose (X), by Maillard-based conjugates, as an approach to improve the ability to entrap flavonoids and to develop powders with enhanced antioxidant activity. WPI (0.6%, *w*/*v*) was conjugated to X (0.3%, *w/v*) through the Maillard reaction at 90 ◦C for 120 min, in the presence of a flavonoid-enriched extract. Two variants of powders were obtained by freeze-drying. The glycation of WPI allowed a better encapsulation efficiency, up to 90.53 ± 0.29%, corresponding to a grafting degree of 30.38 ± 1.55%. The molecular modelling approach was used to assess the impact of X interactions with α-lactalbumin and β-lactoglobulin on the ability of these proteins to bind the main flavonoids from the yellow onion skins. The results showed that X might compete with quercetin glucosides to bind with α-lactalbumin. No interference was found in the case of βlactoglobulin. The microstructural appearance of the powders revealed finer spherosomes in powder with WPI–X conjugates via the Maillard reaction. The powders were added to nachos, followed by a phytochemical characterization, in order to test their potential added value. An increase in antioxidant activity was observed, with no significant changes during storage.
**Keywords:** glycation; flavonoids; microencapsulation; onion skins; antioxidant activities
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007afdec-bed4-405d-873d-c355ba9add0e.135 | **1. Introduction**
The recent trends in food intakes have experienced a transition to a more healthy oriented nutrition, which shifted eating habits to natural foods, paving the way for the extraction and identification of new biologically active compounds, with beneficial or even therapeutic functions, with a considerable emphasis put on well-being and the prevention of disease. Therefore, obtaining and incorporating bioactive-enriched plant extracts in food may significantly contribute to lowering the risk of specific illnesses [1]. Certain advantages may result from the use of bioactive-enriched plant extracts when compared with individual or synthetic compounds, particularly in terms of the synergistic actions of different molecules [2].
Onion (*Allium cepa* L.) is cultivated around the world, being the second most grown horticultural crop after tomatoes. It has been estimated that more than 550,000 tonnes of onion skin bio-waste was generated by the use of the 89 million tonnes onion harvest [3]. The onion waste represents an environmental problem since it is not suitable for animal feeding and so is usually sent to landfill. Onion skins and the outer layers contain significant quantities of fiber and phenolic compounds, such as flavonoids, glucosides, phenolic acids, and organosulfur compounds [3]. In particular, the onion solid waste is rich in quercetin, quercetin glucosides, quercetin polymers, ferulic acid, gallic acid, and
**Citation:** Milea, S, .A.; Aprodu, I.; Enachi, E.; Barbu, V.; Râpeanu, G.; Bahrim, G.E.; St ˘anciuc, N. Whey Protein Isolate-Xylose Maillard-Based Conjugates with Tailored Microencapsulation Capacity of Flavonoids from Yellow Onions Skins. *Antioxidants* **2021**, *10*, 1708. https:// doi.org/10.3390/antiox10111708
Academic Editors: Li Liang and Hao Cheng
Received: 1 October 2021 Accepted: 26 October 2021 Published: 27 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/).
kaempferol, with significant beneficial effects [4] associated with biological activities such as: antidiabetic, antioxidant, anti-inflammatory, anticancer, antimicrobial, and enzyme inhibitory effects [5]. Thus, it can be appreciated that onion has nutritional complexity and holds suitable potential for functional food development, as a source of antioxidant, antimicrobial, anticancer, and antibrowning compounds [6].
Nowadays, the food industry is focusing on implementing methods for the valorization of onion solid waste, as a natural resource with a high amount of value-added ingredients, into eco-friendly functional foods [7]. However, adding polyphenols in a free form in foods may lead to chemical instability due to the unsaturated bonds contained in their molecular structures. The stability is affected by the presence of oxidants, heat, light, and enzymes during storage [8]. Suitable techniques to protect phenolic compounds from chemical damage before their industrial application carry out microencapsulation using different methods, such as freeze-drying [9], which may overcome the drawbacks of their instability, improve their bioavailability as well as shelf life [10] and widen the industrial applications in the food, pharmaceutical and cosmetics industries [11]. In our recent study, different delivery systems were developed for extracts enriched in onion skin flavonoids using a unique combination of whey protein isolates, whey proteins hydrolysates, pectin, and maltodextrin as coating materials [12]. The coating materials should have thermal or mechanical stability to protect the core materials from external factors. Since proteins have amphiphilic properties, they can correlate with the interaction of various chemical groups. However, when using proteins as coatings, some limitations should be considered, given by several external factors, including pH variation, ionic strength and in vitro proteolysis by pepsin, which lead to the degradation of protective walls, causing the release and degradation of bioactives during digestion [13]. These authors tested various structural designs, such as Maillard-based conjugation, to modify the structure and properties of whey proteins and to produce more stable delivery systems with excellent properties. The functional and physico-chemical properties gained with glycation reaction refer to significantly improved emulsifying properties, thermal stability, antioxidant properties, antibacterial activity, and water solubility [14], simultaneously with enhancing the thermal stability of proteins over a wide range of pH and thermal aggregation values [13]. Numerous studies are focused on whey protein as an encapsulating material, but, to the best of our knowledge, there are no studies using whey protein isolates (WPI) in conjugate form with xylose (X) as an encapsulation material for flavonoids. Therefore, the aim of this study was to test the possibility of using WPI–X Maillard-based conjugates as coating materials for flavonoids extracted from yellow onions skins. Flavonoids were isolated by means of solid–liquid extraction in combination with ultrasound-assisted extraction using ethanol as solvent. The WPI–X conjugates were generated via heating in an alkali environment, whereas the flavonoid microcapsules were generated using freeze-drying. Two powders were obtained, using WPI-X conjugates with and without heating, and the resulting freeze-dried powders were characterized in terms of their encapsulation efficiency, phytochemical content and antioxidant activity. Structural and morphological particularities of the samples were analyzed using confocal laser electron microscopy. In order to test the added value, the powders were added to the recipe of a food product (nachos), followed by phytochemical characterization. The results obtained in this study could bring certain benefits in terms of exploiting the bioactive potential of phytochemicals and glycation reaction for developing formulas with improved functional properties.
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007afdec-bed4-405d-873d-c355ba9add0e.137 | *2.1. Chemicals*
Whey protein isolate (WPI) (protein content of 95%) was purchased from Fonterra (New Zealand). Xylose (X) (about 99% purity) and the reagents used to determine the total phenolic compounds, total flavonoid content, 2,2-diphenyl-1-picrylhydrazyl (DPPH), and 6-hydroxy-2,5,7,8-tetramethylchroman 2-carboxylic acid (Trolox) were purchased from Sigma-Aldrich Corp. (St. Louis, MO, USA). All other reagents were of analytical grade.
| doab | 2025-04-07T03:56:59.205187 | 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.138 | *2.2. Ethanolic Ultrasound-Assisted Extraction of Flavonoids from Onion Skins*
Yellow onions were purchased from a local market (Galati, Romania) in June 2020. The outer layers of onions were collected, cleaned with distilled water and dried. Before extraction, in order to obtain a homogeneous batch of particle size, the onion skins were ground to sizes smaller than 0.5 mm × 0.5 mm and used for further extraction. The flavonoidic extract was obtained by mixing 50 g of the onion skins with 450 mL of 70% ethanol solution and glacial acetic acid (ratio 9:1, *v*/*v*). The extraction was performed using a sonication bath at 40 ◦C for 30 min. In order to obtain flavonoids-enriched extracts, the extraction was repeated three times, while the supernatants were centrifuged at 5000× *g* for 10 min at 4 ◦C, collected and concentrated under reduced pressure at 40 ◦C. The obtained extract was characterized in terms of selected phytochemicals and used for microencapsulation experiments.
| doab | 2025-04-07T03:56:59.205236 | 17-11-2022 17:23 | {
"license": "Creative Commons - Attribution - https://creativecommons.org/licenses/by/4.0/",
"book_id": "007afdec-bed4-405d-873d-c355ba9add0e",
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"author": "",
"title": "Characterization and Encapsulation of Natural Antioxidants: Interaction, Protection and Delivery",
"publisher": "MDPI - Multidisciplinary Digital Publishing Institute",
"isbn": "9783036554563",
"section_idx": 138
} |
007afdec-bed4-405d-873d-c355ba9add0e.139 | *2.3. Preparation of WPI–X–Flavonoid Conjugates*
WPI–X conjugates were prepared according to the method described by Jia et al. [13] with minor modifications. An amount of 60 mg/mL of WPI was first dissolved in 100 mL ultrapure water under gentle stirring, followed by the addition of 30 mg/mL of X (mass ratio of 2:1). The mixture was allowed to hydrate for 14 h at room temperature (25 ◦C). After complete hydration, about 750 mg of concentrated flavonoid-enriched extract was added and the mixture was allowed to dissolve by ultrasonication for 1 h at 35 ◦C. The resulting solution was divided into two: variant 1 (coded V1) and variant 2 (coded V2). The pH of V2 solution was adjusted at 9.0 and placed in a sealed screw-top glass tube. In order to promote the Maillard conjugation, Variant 2 was heated to 90 ◦C in water bath for 3 h. After heating, the temperature of Variant 2 was lowered to 25 ◦C in an ice bath. Both variants were freeze-dried (CHRIST Alpha 1–4 LD plus, Osterode am Harz Germany) at −42 ◦C under a pressure of 10 Pa for 48 h and stored at −4 ◦C.
| doab | 2025-04-07T03:56:59.205321 | 17-11-2022 17:23 | {
"license": "Creative Commons - Attribution - https://creativecommons.org/licenses/by/4.0/",
"book_id": "007afdec-bed4-405d-873d-c355ba9add0e",
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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": 139
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007afdec-bed4-405d-873d-c355ba9add0e.140 | *2.4. Characterization of the Extract and of Microencapsulated Powders*
Both the extract and the powders were characterized for flavonoids, total phenolic compound contents and antioxidant activity using the aluminum chloride method, Folin– Ciocâlteu and DPPH method, respectively, as described by Milea et al. [12]. The total flavonoid contents are expressed in mg quercetin equivalents (QE)/g dry weight (DW), whereas the total polyphenol contents are expressed in mg gallic acid equivalents (GAE)/g DW. In each case, the concentrations of bioactives and antioxidant activity were expressed through selected standard calibration curves.
The encapsulation efficiency of the powders was calculated as described by Saénz et al. [15]. In brief, the microencapsulation efficiency was determined by assessing the surface flavonoid contents (SFC) and total flavonoid contents (TFC) of the powders, expressed as mg QE/g DW. In order to quantify the SFC, 50 mg of powders was mixed with 5 mL of ethanol and methanol (ratio 1:1, *v*/*v*). These dispersions were stirred at room temperature for 1 min and then centrifuged at 4000× *g* and 4 ◦C for 10 min. For TFC, 50 mg of powder was accurately weighed and dispersed in 5 mL of a mixture of ethanol, acetic acid, and water (50:8:42, *v*/*v*/*v*). The resulting dispersion was vortexed (1 min), followed by ultrasonication for 30 min at 40 ± 1.0 ◦C, to break the microcapsules. The supernatant was centrifuged at 14,000× *g* for 10 min and then filtered. The content of flavonoids in the resulting supernatants was measured by the aluminum chloride method, as explained by Milea et al. [12]. The microencapsulation efficiency (ME, %) was calculated using Equation (1):
$$\text{ME} \left( \% \right) = \frac{\text{TFC} - \text{SFC}}{\text{TFC}} \times 100 \tag{1}$$
The antioxidant activity was assessed using the DPPH method. Briefly, 0.1 mL of supernatant resulted from TFC determination was added to 3.9 mL of DPPH stock solution. The DPPH stock solution was prepared mixing 3 g of DPPH with 100 mL of methanol. Simultaneously, a control sample was prepared by adding 0.1 mL methanol to 3.9 mL
DPPH. The absorbance for both samples was read at 515 nm after 1.5 h. The results were calculated using a calibration curve and are expressed in mMol Trolox/g DW.
| doab | 2025-04-07T03:56:59.205422 | 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",
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"isbn": "9783036554563",
"section_idx": 140
} |
007afdec-bed4-405d-873d-c355ba9add0e.141 | *2.5. Browning Index and Grafting Degree Measurement of the Powders*
The browning intensity was measured using a spectrophotometric method with minor modifications [16]. A volume of 5 mL of a mixture consisting of equal volume of acetic acid (2% *v*/*v*) and formaldehyde (1% *v*/*v*) was added to 0.1 mL of 1 mg/mL of samples and centrifuged for 10 min at 4500× *g*. The resulting supernatant (5 mL) was mixed with 5 mL of ethanol and the mixture was centrifuged again. Absorbances of the supernatant at 420 and 600 nm were measured. The difference between the two absorbance values was used to evaluate the browning index.
The grafting degree was determined using the o-phthalaldehyde (OPA) method, according to Jia et al. [13]. A volume of 4 mL of OPA was added to 0.2 mL of the diluted samples (0.1 mg/mL) in test tubes. Upon homogenization, all tubes were placed in a water bath at 35 ◦C for 1 min. The absorbance was measured at 340 nm. A blank sample was also made using the same volume of ultrapure water. Grafting degree was determined using Equation (2):
$$\text{Grafting Degree} \left( \% \right) = 100 \times \frac{\text{A}\_0 - \text{A}\_\kappa}{\text{A}\_0} \tag{2}$$
where A0 and As are the absorbance of blank sample and absorbance of tested samples, respectively.
| doab | 2025-04-07T03:56:59.205568 | 17-11-2022 17:23 | {
"license": "Creative Commons - Attribution - https://creativecommons.org/licenses/by/4.0/",
"book_id": "007afdec-bed4-405d-873d-c355ba9add0e",
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"title": "Characterization and Encapsulation of Natural Antioxidants: Interaction, Protection and Delivery",
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"isbn": "9783036554563",
"section_idx": 141
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007afdec-bed4-405d-873d-c355ba9add0e.142 | *2.6. In Silico Investigations*
In agreement with the experimental approach, the molecular modeling tools were used to simulate X binding by the two major whey proteins. The three-dimensional molecular models of the α-lactalbumin (α-LA, PDB ID: 1F6S) [17] and β-lactoglobulin monomers (β-LG, PDB ID: 4DQ3) [18] from the RCSB Protein Data Bank were optimized and relaxed at 25 ◦C using GROMACS 4.6 software [19], in agreement with the protocol previously described by Aprodu et al. [20]. The equilibrated protein models were used as receptors for the X binding. The PatchDock algorithm [21], which is very efficient for performing protein–small ligand docking, was used to identify the most probable binding site of X molecules to the WP. Matching the receptor and ligand molecules was carried out through rigid body docking, which is based on the shape complementarity principles. The algorithm employed involves the following major stages: the surface of the receptor was first segmented to identify the so-called hot spot residues on the concave, convex or flat geometric patches, which were selected for a further surface patch matching step with the ligand. The resulting complexes were filtered to disqualify the solutions, involving steric clashes, and finally ranked based on the geometric shape complementarity score. The best three WPI–X fits were selected based on the binding energy values among the potential docking models generated by the PatchDock algorithm [21]. An in-depth analysis of the binding pockets was carried out using the PDBePISA [22] tools and DoGSiteScorer web server [23] to identify the extent to which protein glycation affects flavonoids binding.
#### *2.7. Confocal Laser Microscope Spectroscopy*
A confocal laser scanning microscopy analysis of the samples was employed to assess the structural appearance of the microencapsulated powders. The CLSM images were captured with a Zeiss confocal laser scanning system (LSM710) equipped with several types of lasers such as a diode laser (405 nm), Ar laser (458, 488, 514 nm), DPSS laser (diode pumped solid state—561 nm) and HeNe laser (633 nm). The powders were observed with a 20× apochromatic objective, at zoom values of 1 and 0.6, respectively. The obtained 3D images were rendered and processed by ZEN 2012 SP1 software (Black Edition).
| doab | 2025-04-07T03:56:59.205655 | 17-11-2022 17:23 | {
"license": "Creative Commons - Attribution - https://creativecommons.org/licenses/by/4.0/",
"book_id": "007afdec-bed4-405d-873d-c355ba9add0e",
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"title": "Characterization and Encapsulation of Natural Antioxidants: Interaction, Protection and Delivery",
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007afdec-bed4-405d-873d-c355ba9add0e.143 | *2.8. Formulation of a Value-Added Food Product*
To support the multifunctional properties, the powders were added as an ingredient in a nacho recipe at a ratio of 3%. The recipe involved mixing of corn (250 g) and wheat flour (100 g) in a ratio of 2.5:1, onion (100 mg), pepper (10 mg), oil (20 mL), salt (10 mg), powders (3%) and water (150 mL). The corresponding samples were coded as N1 and N2. The control sample was nachos without the addition of microencapsulated powder (C). Samples were homogenized and allowed to stand for 1 h at room temperature to equilibrate. After homogenization, the nachos were formed and cooked for 6 min in an oven (3 min on each side) at 200 ◦C. The storage stability of bioactives was tested at 0 days and after 28 days at 25 ◦C.
#### *2.9. Statistical Analyses*
All analyses were performed in triplicate and data are reported as mean ± standard deviation (SD). After running the normality and homoscedasticity tests, experimental data were subjected to one-way analysis of variance (ANOVA) in order to identify significant differences. The Tukey method with a 95% confidence interval was employed for post hoc analysis; *p* < 0.05 was considered to be statistically significant. The statistical analysis was carried out using Minitab 18 software.
| doab | 2025-04-07T03:56:59.205818 | 17-11-2022 17:23 | {
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"title": "Characterization and Encapsulation of Natural Antioxidants: Interaction, Protection and Delivery",
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"section_idx": 143
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007afdec-bed4-405d-873d-c355ba9add0e.145 | *3.1. Phytochemical Characterization of the Yellow Onion Skin Extract*
The solid–liquid ethanolic ultrasound-assisted method applied in this study allowed us to obtain a bioactive-enriched extract, containing flavonoids of 228.7 ± 3.0 mg QE/g DW, with a total polyphenolic content of 96.1 ± 2.7 mg GAE/g DW, and yielding an antioxidant activity of 495.9 ± 2.4 mM TEAC/g DW. In our previous study, different extraction techniques were tested in order to select the most suitable method to obtain flavonoid-enriched extracts from yellow onion skins [24]. The results showed a satisfactory content in phytochemicals, when comparing the ultrasound-assisted technique versus the conventional solid–liquid extraction. However, the selection of the ultrasound-assisted extraction in this study was based on the reduction time and protection of thermolabile compounds. Therefore, Constantin et al. [24] reported similar values for flavonoid contents in ultrasoundassisted extracts of 230.6 ± 8.4 mg QE/g DW. Additionally, Milea et al. [12] extracted the biologically active compounds from yellow onion skins using a similar method and reported flavonoids of 97.3 ± 3.0 mg QE/g DW, polyphenols of 55.3 ± 2.5 mg GAE/g DW and an antioxidant activity of 345.0 ± 2.7 mM TE/g DW. Singh et al. [25] used an ultrasoundassisted method to extract the bioactive compounds from onions. The extraction with 70% ethanol showed similar values for flavonoid extraction of 212.3 ± 14.6 mg QE/g and a higher amount of phenolic compounds (418.0 ± 34.4 mg GAE/g). On the other hand, Pobłocka-Olech et al. [26] extracted flavonoids from different varieties of yellow onion skins using only methanol and obtained a lower level compared with the current results, between 2.4 and 12.2 mg QE/g. Benito-Román et al. [27] performed a comparative study of polyphenols from onion wastes between conventional and ultrasound-assisted extraction. They reported smaller values for flavonoid contents, ranging from 7.7 ± 0.1 to 23.8 mg QE/g dry onion skins (DOS). The different values could be explained by the distinct selected parameters, different origin of raw materials or by the method of expressing final results (DW/DOS). Likewise, the experimental conditions allowed us to extract a significant amount of polyphenols (73.3 ± 1.8 mg GAE/g DOS), which were further increased to 102.1 ± 5.1 mg GAE/g DOS.
The difference between results is due to the extraction method. As is known, ultrasoundassisted method simplifies and accelerates the extraction because the high-intensity ultrasounds increase pressure and temperature, causing a disruption of the cell wall of the matrix, with the subsequent release of polyphenols. Moreover, this technique offers the advantages of lower extraction times and temperatures compared to conventional extraction techniques [28].
| doab | 2025-04-07T03:56:59.205921 | 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.146 | *3.2. Correlation between Browning Intensity, Grafting Degree and Microencapsulation Efficiency*
An effective method to improve the functional properties of proteins, including the ability to include and protect low molecular bioactives, is based on the interaction with polysaccharides and smaller carbohydrates, via Maillard conjugation [29]. The Maillard reaction is a complex reaction occurring between amines and carbonyls [30], which involves, first, to consumption of the free amino group by the carbonylation reaction, mainly coming from the free amino group on the side chain such as lysine and arginine, or the free amino group on the N-terminus of the peptide chain of the protein molecule. Therefore, the carbonyl condensation between the reducing sugar and nucleophilic amino group could be analyzed by the loss of amino acids in the reaction [31]. Ideally, as reported by Jiménez-Castanõ et al. [32], to produce a glycol conjugate destined for incorporation into food, to avoid the formation of the highly colored, insoluble, nitrogen-containing polymeric compounds, referred to as melanoidins, the Maillard reaction needs to be performed under carefully controlled conditions to prevent the later stage changes. As reported by Jia et al. [13], protein glycation by the Maillard reaction might be favored in alkali conditions, while glycation is inhibited by the partial denaturation of the protein in acid conditions. These authors suggested a higher grafting degree and lower browning intensity at pH 9.0 after heating for 3 h. Therefore, these parameters were selected in our study to promote Maillard-based conjugates between WPI and X.
The brown-colored pigment formation in foods is caused by the Maillard reactions or caramelization [16]; therefore, the browning index is generally accepted as an indicator of the Maillard reaction. The brown pigment formation in the microencapsulated powders was evaluated by absorbance measurements at 420 nm and 600 nm, respectively. As expected, the browning intensity was higher (0.12 ± 0.01) for the heat-treated variant (Variant 2) than for the untreated variant (0.09 ± 0.01) (Variant 1). A proportional increase was observed between the browning intensity and antioxidant activity of the powders, indicating the strong antioxidant potency of the glycated variant due to the heating process. The powders showed significant differences in antioxidant activity (*p* < 0.05), with values of 179.7 ± 4.5 mMol TE/g DW for V1 and 184.4 ± 0.7 mMol TE/g DW for V2. Suminar et al. [33] explained that this was probably caused by reducing sugar reacting more easily with amino acids in heating conditions and producing antioxidant activity. The formation of the conjugate was also confirmed by grafting degree, which is able to reflect the level of glycation [13]. In the present study, a correlation between the grafting degree and encapsulation efficiency can be observed in both variants. Thereby, a grafting degree of 22.6 ± 2.5% and an encapsulation efficiency of 86.7 ± 1.4% were observed for V1. A significantly higher (*p* < 0.05) values were estimated for Variant 2, with a grafting degree of 30.4 ± 1.6% and an encapsulation efficiency of 90.5 ± 0.3%. Therefore, it can be appreciated that the Maillard-based conjugates showed a higher ability to entrap the flavonoids from yellow onion skin extract. These results indicate that the glycated form of the powder has a positive effect on the encapsulation efficiency.
In the conditions applied in our study, the Maillard-based conjugates between WPI and X caused structural changes in proteins that improved the ability to entrap flavonoids, in good agreement with reports of Liu et al. [34], Xu et al. [35] and Liu et al. [36]. The glycation degree can be correlated with the decrease in available -NH2 groups. For example, Shang et al. [37] suggested a dramatic loss in Lys and Arg, whereas a significant decrease in Tyr and Cys was also found, due to the formation of the dehydroalpropyl side chain. These authors also reported a transition toward a higher molecular weight distribution of WPI heated in the presence of X, at 90 ◦C and 95 ◦C and pH 9.0, whereas the contents of protein polymers larger than 40 kDa increased with the reaction time, thus indicating a protein crosslinking phenomenon. The heat-induced glycation reaction between WPI and X molecules might induce the formation of hydrogen bonds, thus weakening the interaction between molecules, and result in a reduction in the β-sheet and β-turns but an increase in the random coil [37].
In another study, lycopene was encapsulated in whey protein isolate and xylooligosaccharides conjugates and presented values ranging from 10.0 ± 0.4% to 27.0 ± 0.5%, depending on the other parameters of the reaction (temperature, time, pH). Muhoza et al. [38] evaluated the possibility of glycating the casein by the Maillard reaction with dextran for delivering coenzyme Q10. These authors reported that when the reaction time was less than 8 h, the grafting degree of the mixture about 20%. Ghatak and Iyyaswami [39] encapsulated quercetin from dry onion peels and obtained an encapsulation yield between 40.4% and 96.4%. They found that the highest encapsulation yield was achieved under the following process conditions: casein concentration of pH 7.09 for the crude extract containing the quercetin concentration of 16.27 M. Akdeniz et al. [40] reported the encapsulation efficiency values of phenolic from onion skins as being between 55.6 and 89.2% for different coating material combinations, with a maximum value found for maltodextrin:casein in a ratio of 6:4.
| doab | 2025-04-07T03:56:59.206103 | 17-11-2022 17:23 | {
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007afdec-bed4-405d-873d-c355ba9add0e.147 | *3.3. Molecular Modeling*
Details on how protein glycation with X impacts further flavonoids binding were collected by means of an in silico approach. The potential binding sites for X were predicted for both α-LA and β-LG molecules by performing molecular docking simulations. The best three fits involving α-LA and β-LG as receptors, decided based on the binding energy values and the interface area, were further analyzed in detail (Table 1). In the case of α-LA, mainly two binding sites located on the protein surface appeared to accommodate the ligands used in the study. The highest affinity of the receptor for its ligand was estimated based on the lowest binding energy. Thus, α-LA exhibited the highest affinity toward X molecules when bound to the cavity involving residues of the α-LA core (Phe53, Gln54, Tyr103, Trp104) and the amino-terminal section of the Leu105-Leu110 helix. In addition to the hydrophobic contacts, three hydrogen bonds of 3.68 Å, 2.37 Å, and 2.83 Å involving Thr33, Leu105 and Ala106, respectively, contributed to the attraction between α-LA and the X molecule hosted within this cavity. Two different relative binding positions of the X molecules with respect to the α-LA receptor, sharing common amino acid residues in contact with the ligand, were predicted with high scores. None of the two X binding modes appeared to affect the attachment of the main flavonoids prevailing in the onion skin extract, namely quercetin-4 -*O*-monoglucoside (QMG) and quercetin-3,4 -*O*-diglucoside (QDG) [41], to the α-LA. In agreement with Horincar et al. [41], the α-LA molecule accommodates, with high specificity, the same binding site of both QDG and QMG ligands (Figure 1). The amino acids establishing direct contacts with QMG are Leu3, Glu11, Leu12, Lys13, Asp14, Thr38, Leu52, Leu85, Thr86, Asp88, Ile89, Met90 and Lys93, whereas the residues responsible for the QDG binding are Glu1, Leu3, Arg10, Glu11, Leu12, Lys13, Thr38, Leu52, Asp83, Leu85, Thr86, Asp88 and Ile<sup>89</sup> [41].
In addition, this wide pocket with a volume of 435.8 Å3 appears to be able to accommodate an X molecule, which overlaps the QMG without interfering with QDG binding. It should be noted that α-LA shows a better affinity towards QMG and QDG (binding energy of −24.21 kcal/mol and −32.01 kcal/mol, respectively) with respect to X (binding energy of −7.48 kcal/mol).
On the other hand, the β-LG molecule is able to accommodate the X molecule in three different pockets with volumes ranging from 137.86 to 217.15 Å (Table 1), without interfering with QMG or QDG binding (Figure 1).
**Table 1.** Molecular details on the interactions between the main whey proteins (α-lactalbumin (α-LA) and β-lactoglobulin monomer (β-LG)) equilibrated at 25 ◦C, xylose (X) and the major flavonoids from onion skins (quercetin-4 -*O*-monoglucoside (QMG) and quercetin-3,4 -*O*-diglucoside (QDG) [41].
β-LG binds the X molecules more tightly compared to α-LA; the X binding energy by the β-LG monomer varies between −16.41 and −13.00 kcal/mol. In addition, the binding of two X molecules to the β-LG pockets involves hydrogen bonds established with Glu<sup>158</sup> in the case of complex 1 and Met24, Asp137 and Leu149 in the case of complex 2, as presented in Table 1. The in silico results successfully complement the experimental findings, adding valuable details on how whey protein glycation with X further impacts flavonoid binding. These atomic level observations indicate that, upon glycation, the β-LG molecule might play a major role in flavonoid biding.
**Figure 1.** Superposition of the models showing the most probable complexes formed between (**a**) α-lactalbumin and (**b**) β-lactoglobulin (represented in blue in surf style) and xylose (represented in orange in licorice style—models 1 and 2), quercetin-4 -*O*-monoglucoside (represented in red in licorice style) and quercetin-3,4 -*O*-diglucoside (represented in green in licorice style). Images were prepared using VMD software [42].
| doab | 2025-04-07T03:56:59.206562 | 17-11-2022 17:23 | {
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007afdec-bed4-405d-873d-c355ba9add0e.148 | *3.4. Phytochemical Profile of the Powders*
The two powders showed a total flavonoid content of 97.7 ± 3.7 mg QE/g DW in V1 and 120.0 ± 1.6 mg QE/g DW in V2. Total polyphenolic content showed no significant (*p* < 0.05) differences of 45.1 ± 0.6 mg GAE/g DW and 46.9 ± 1.5 mg GAE/g DW in V1 and V2, yielding a corresponding antioxidant activity of 179.7 ± 4.5 mMol TE/g DW and 184.4 ± 0.7 mMol TE/g DW, respectively. Therefore, the glycation of WPI allowed a better encapsulation of flavonoids, yielding a powder with a higher antioxidant activity. To the best of our knowledge, no other studies are available that exploit the potential of WPI conjugates as biopolymeric wall materials used in the microencapsulation of flavonoids.
In a previous study, Milea et al. [43] encapsulated flavonoids from yellow onion skins using maltodextrin, pectin and whey protein hydrolysates as coating materials in different ratios. The concentration of flavonoids, polyphenols and the antioxidant activity of the freeze-dried variants showed comparable levels, as flavonoids varied from 98.1 ± 0.5 to 103.7 ± 0.6 mg QE/g DW, whereas significant higher polyphenol contents (varying
from 53.5 ± 1.7 to 69.3 ± 1.0 mg GAE/g DW) and antioxidant activities (varying from 280.6 ± 3.1 to 337.6 ± 0.9 mM TE/g DW) were reported for different variants. Horincar et al. [41] used different combinations of biopolymeric coatings based on whey protein isolate and chitosan, maltodextrin and pectin as adjuvants for encapsulation. These authors obtained two variants of freeze-dried powder with different profiles. Therefore, lower values for total flavonoid content of 5.8 ± 0.2 mg QE/g DW and antioxidant activity of 175.9 ± 1.5 mM TE/g DW were suggested in coatings with WPI-chitosan. When using a more complex biopolymeric wall material, including WPI-maltodextrin-pectin, these authors obtained a powder with significant higher flavonoid content and antioxidant activity of 104.9 ± 5.0 mg QE/g DW and 269.2 ± 3.6 mM TE/g DW, respectively.
| doab | 2025-04-07T03:56:59.206835 | 17-11-2022 17:23 | {
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"section_idx": 148
} |
007afdec-bed4-405d-873d-c355ba9add0e.149 | *3.5. Structure and Morphology of the Powder*
The confocal laser scanning microscopy technique allows the simultaneous identification of several compounds, at the surface of the particles using specific samples that usually discharge light at different wavelengths. This type of analysis also permits the visualization of the internal morphology of any type of particle by different fluorophores labels, hence displaying the compositional evolution of the targeted molecules, which represents the main aim of any study that regards the protection of valuable molecules such as antioxidant compounds, mainly polyphenols, through an encapsulation process. Therefore, it can be applied as a nondestructive visualization technique for microparticles.
By using a confocal laser scanning Carl Zeiss 710 microscope with the ZEN 2012 SP1 software (Black Edition), the images of V1 and V2 powders (Figure 2) were captured, both in the native state without any other additional dye added (Figure 2a,c) and stained with Congo Red (Figure 2b,d, respectively). In the native state, the powders showed an autofluorescence (in the range of 520–580 nm) due to the rich content of polyphenols among which quercetin predominates [39]. The biologically active compounds were captured in the WPI–X matrix (with a displayed autofluorescence showed in blue). Several irregular scaly formations could be observed with larger dimensions for the V2 (199.6–253.3 μm), compared to V1 (94.5–104.8 μm), probably due to the WPI–X conjugates. The microstructure was rather similar to that reported by Horincar et al. [41], who used different polymers as the encapsulating matrices such as chitosan, maltodextrin and pectin.
Through the fluorescent staining with Congo Red, a fluorophore usually used to highlight the fluorescence of proteins, several spherosomes were revealed, highlighting the encapsulated flavonoids in the WPI–X matrix. Nonetheless, the displayed formations were larger (up to 51.5 μm) and less numerous in the V1 sample where a significant amount of nonencapsulated flavonoids was visualized (Figure 2b).
The interaction of the WPI with the small carbohydrates via the Maillard reaction favored a better incorporation of the biologically active compounds from the onion extract into finer spherosomes (approximately 20 μm) or in the form of scales with digitiform extensions. The fluorophore bound to the conjugated proteins and generated a fine, orange wall with a fluorescent emission between 600 and 620 nm around the bioactives.
.**-**2
.2
| doab | 2025-04-07T03:56:59.206952 | 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": 149
} |
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