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Physical, thermal, and storage stability of multilayered emulsion loaded with β-carotene

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  • Carotenoids are colored bioactive substances increasingly used due to their antioxidant properties, vitamin A precursor role, and ability to function as a natural food color. Knowledge of carotenoid behavior during high-heat processing and subsequent storage in emulsified food matrix is essential to expand their application natural food colors and neutraceuticals. Firstly, the physical, thermal, and colloidal stability of emulsions constructed from octenyl succinic anhydride-modified starch (OSA starch)-chitosan multilayered interfaces were investigated. Results of charge reversal from −32.4 ± 1.9 mV to +38.0 ± 0.8 mV indicate that multilayered interfaces were formed in emulsions. As measured by Z-average size, the emulsions were stable after the thermal treatment at 121 °C for 60 min, thus demonstrating a novel heat-stable multilayered emulsion. Subsequently, a select multilayered emulsion was loaded with β-carotene, and its storage stability was assessed. The degradation of β-carotene in an oil-in-water emulsion was better described with zeroth order kinetics; β-carotene dissolved in bulk oil was better described using a second-order kinetic equation. The presence of an encapsulating material around the oil droplets loaded with β-carotene enhanced its stability, which makes it instrumental in extending shelf-life and maintaining a consistent appearance. The results can be used to predict the availability of β-carotene during storage.
  • The Lonicera Linn. genus is a constituent member of the Caprifoliaceae family[1]. It is the largest genus in this family and comprises at least 200 species with a notable presence in North Africa, North America, Asia, and Europe[1]. Members of the Lonicera genus possess a wide range of economic benefits from their use as ornamental plants to food and as plants credited with numerous health benefits. Conspicuous among the numerous members of this genus with known medicinal uses are L. japonica, L. macranthoides, L. hypoglauca, L. confusa, and L. fulvotomentosa[2]. Though these species feature prominently in the Chinese Pharmacopoeia, other species such as L. acuminata, L. buchananii, and L. similis are recognized as medicinal resources in certain parts of China[1]. Among the aforementioned species, L. japonica takes precedence over the rest due to its high medicinal and nutritional value[3,4]. For instance, the microRNA MIR2911, an isolate from L. japonica, has been reported to inhibit the replication of viruses[57]. Also, the water extract of L. japonica has been used to produce various beverages and health products[8]. The Lonicera genus therefore possesses huge prospects in the pharmaceutical, food, and cosmetic industries as an invaluable raw material[9].

    The main active constituents of the Lonicera genus include organic acids, flavonoids, iridoids, and triterpene saponins. Chlorogenic acids, iridoids, and flavones are mainly credited with the anti-inflammatory, antiviral, anticancer, and antioxidant effects of the various Lonicera species[1013]. Their hepatoprotective, immune modulatory, anti-tumor and anti-Alzheimer’s effects are for the most part ascribed to the triterpene saponins[1416]. As stated in the Chinese Pharmacopoeia and backed by the findings of diverse research groups, the plants of the Lonicera genus are known to possess high amounts of organic acids (specifically chlorogenic acid) and pentacyclic triterpenoid saponins[2,1719]. The flower and flower bud have traditionally served as the main medicinal parts of the Lonicera genus even though there is ample evidence that the leaves possess the same chemical composition[20]. A perusal of the current scientific literature reveals the fact that little attention has been devoted to exploring the biosynthesis of the chemical constituents of the Lonicera genus with the view to finding alternative means of obtaining higher yields. It is therefore imperative that priority is given to the exploration of the biosynthesis of these bioactive compounds as a possible means of resource protection. There is also the need for further research on ways to fully tap the medicinal benefits of other plant parts in the Lonicera genus.

    Here, we provide a comprehensive review of relevant scientific literature covering the structure, pharmacology, multi-omics analyses, phylogenetic analysis, biosynthesis, and metabolic engineering of the main bioactive constituents of the Lonicera genus. Finally, we proffer suggestions on the prospects of fully exploiting and utilizing plants of the Lonicera genus as useful medicinal plant resources.

    A total of at least 400 secondary metabolites have been reported for the Lonicera genus. These metabolites are categorized into four main groups (Fig. 1a), including not less than 50 organic acids, 80 flavonoids, 80 iridoids, and 80 triterpene saponins[2123]. Organic acids are mainly derivatives of p-hydroxycinnamic acid and quinic acid. Among the organic acids, chlorogenic acids are reported to be the main bioactive compounds in L. japonica[2426]. The organic acids are most abundant in the leaves, while the least amounts are found in the stem of L. japonica. The flowers of the plant are known to contain moderately high amounts of organic acids[27]. The basic core structure of the flavonoids is 2-phenylchromogen. Luteolin and its glycoside which are characteristic flavonoids of the Lonicera genus are most abundant in L. japonica[28]. On the whole, the flavonoid contents in L. japonica are also highest in the leaves, available in moderate amounts in the flowers, and in lowest amounts in the stem[21]. The core structures of the iridoids are iridoid alcohols, the chemical properties of which are similar to hemiacetal. The iridoids often exist in the form of iridoid glycosides in plants. Secoiridoids glycosides are predominant in the Lonicera genus[25]. In L. japonica, the contents of the iridoids are most abundant in the flowers, moderate in leaves, and lowest in the stem[21]. The characteristic saponins of the Lonicera genus are mainly pentacyclic triterpenoids, including the hederin-, oleanane-, lupane-, ursulane- and fernane-types, etc[22]. The hederin-type saponins are reported in the highest amounts in L. macranthoides[17] (Fig. 1b).

    Figure 1.  Core structures of main secondary metabolites and their distribution in five species of Lonicera. (a) 1 and 2, the main core structures of organic acids; 3, the main core structures of flavonoids; 4, the main core structures of iridoids; 5, the main core structures of triterpene saponins. (b) Comparison of dry weight of four kinds of substances in five species of Lonicera[17,28].

    The similarities between chlorogenic acid (CGA) and flavonoids can be traced back to their biosynthesis since p-coumaroyl CoA serves as the common precursor for these compounds[29]. p-coumaroyl CoA is obtained through sequential catalysis of phenylalanine and its biosynthetic intermediates by phenylalanine-ammonia-lyase (PAL), cinnamate 4-hydroxylase (C4H) and 4-coumarate CoA ligase (4CL)[3033].

    CGA is a phenolic acid composed of caffeic acid and quinic acid and is the most important bioactive compound among the organic acids. Its biosynthesis has been relatively well-established; three main biosynthetic routes have been propounded (Fig. 2a). One route relates to the catalysis of caffeoyl-CoA and quinic acid by hydroxycinnamoyl-CoA quinate transferase (HQT)/hydroxycinnamoyl CoA shikimate/quinate hydroxycinnamoyl transferase (HCT) to produce CGA[3437]. The HQT-mediated pathway has been deemed the major route for CGA synthesis in in different plant species[38,39]. The second biosynthetic route stems from the biosynthesis of p-coumaroyl quinate through the catalytic effect of HCT/HQT and subsequent hydroxylation of p-coumaroyl quinate under the catalysis of p-coumarate 3'-hydroxylase (C3’H)[34,36,37]. For the third route, caffeoyl glucoside serves as the intermediate to form CGA, a process that is catalyzed by hydroxycinnamyl D-glucose: quinic acid hydroxycinnamyl transferase (HCGQT)[40,41].

    Figure 2.  Biosynthetic pathways of main bioactive constituents of Lonicera. (a) Biosynthetic pathways of chlorogenic acid. (b) Biosynthetic pathways of luteoloside. (c) Biosynthetic pathways of secologanin. (d) Biosynthetic pathways of hederin-type triterpene saponins. PAL, phenylalanine ammonia-lyase; C4H, cinnamate 4-hydroxylase; 4CL, 4-hydroxycinnamoyl CoA ligase; HCT, hydroxycinnamoyl CoA shikimate/quinate hydroxycinnamoyl transferase; C3’H, p-coumaroyl 3-hydroxylase; HQT, hydroxycinnamoyl-CoA quinate transferase; UGCT, UDP glucose: cinnamate glucosyl transferase; CGH, p-coumaroyl-D-glucose hydroxylase; HCGQT, hydroxycinnamoyl D-glucose: quinate hydroxycinnamoyl transferase; CHS, Chalcone synthase; CHI, Chalcone isomerase; FNS, Flavone synthase; F3H, flavonoid 30-monooxygenase/flavonoid 30-hydroxylase; UF7GT, flavone 7-O-β-glucosyltransferase; GPS, Geranyl pyrophosphatase; GES, geraniol synthase; G8O, geraniol 10-hydroxylase/8-oxidase; 8HO, 8-hydroxygeraniol oxidoreductase; IS, iridoid synthase; IO, iridoid oxidase; 7DLGT, 7-deoxyloganetic acid glucosyltransferase; 7DLH, 7-deoxyloganic acid hydroxylase; LAMT, loganic acid O-methyltransferase; SLS, secologanin synthase; FPS, farnesyl pyrophosphate synthase; SS, squalene synthase; SE, squalene epoxidase; β-AS, β-amyrin synthase; OAS, oleanolic acid synthetase.

    The key enzymes in the biosynthesis of p-coumaroyl CoA, and invariably CGA, thus, PAL, C4H, and 4CL have been established in diverse studies such as enzyme gene overexpression/knockdown[42], enzyme activity analysis[33] and transcriptomics[18]. However, the centrality of HQT in the biosynthesis of CGA remains disputable. While some studies have reported a strong correlation between HQT expression level with CGA content and distribution[18,35,39,43,44], others found no such link[45], bringing into question the role of HQT as a key enzyme in CGA biosynthesis.

    Few studies have been conducted on the regulation of CGA biosynthesis in the Lonicera genus. It was found that overexpression of the transcription factor, LmMYB15 in Nicotiana benthamiana can promote CGA accumulation by directly activating 4CL or indirectly binding to MYB3 and MYB4 promoters[46]. LjbZIP8 can specifically bind to PAL2 and act as a transcriptional repressor to reduce PAL2 expression levels and CGA content[47]. Under NaCl stress, increased PAL expression promoted the accumulation of phenolic substances in leaves without oxidative damage, a condition that was conducive to the accumulation of the bioactive compounds in leaves[48].

    Luteolin and its derivative luteolin 7-O- glucoside (luteoloside) are representative flavonoids of the Lonicera genus. Similar to CGA, luteolin is biosynthesized from p-coumaroyl CoA but via a different route. The transition from p-coumaroyl CoA to luteolin is underpinned by sequential catalysis by chalcone synthetase (CHS), chalcone isomerase (CHI), flavonoid synthetase (FNS), and flavonoid 3'-monooxygenase/flavonoid 3'-hydroxylase (F3'H)[45,49,50] (Fig. 2b). Luteoloside is synthesized from luteolin by UDP glucose-flavonoid 7-O-β-glucosyltransferase (UF7GT)[51]. Similar to CGA biosynthesis, the key enzymes of luteolin synthesis include PAL, C4H, and 4CL in addition to FNS[33,45,52]. The content of luteoloside was found to be highly abundant in senescing leaves relative to other tissues such as stem, flowers, and even young leaves[52]. Through transcriptomic analysis, luteoloside biosynthesis-related differentially expressed unigenes (DEGs) such as PAL2, C4H2, flavone 7-O-β-glucosyltransferase (UFGT), 4CL, C4H, chalcone synthase 2 and flavonoid 3'-monooxygenase (F3'H) genes were found to be upregulated in the senescing leaves. Biosynthesis-related transcription factors such as MYB, bHLH, and WD40 were also differentially expressed during leaf senescence[52], while bHLH, ERF, MYB, bZIP, and NAC were differentially expressed during flower growth[53]. Further analysis of the transcription factors revealed that MYB12, MYB44, MYB75, MYB114, MYC12, bHLH113, and TTG1 are crucial in luteoloside biosynthesis[52,53].

    The biosynthesis of terpenoids mainly involves three stages; formation of intermediates, formation of basic structural skeleton, and modification of basic skeleton[54]. The intermediates of terpenoids are mainly formed through the mevalonate (MVA) and methylerythritol phosphate (MEP) pathways, and eventually converted to the universal isoprenoid precursors, isopentenyl pyrophosphate (IPP) and its isomer dimethylallyl pyrophosphate (DMAPP) through a series of enzyme-catalyzed reactions. Under the catalysis of geranyl pyrophosphatase (GPS), IPP is then converted to geranyl pyrophosphate (GPP). Different terpenoids are subsequently derived from GPP as the intermediate product. For instance, in the formation of secoiridoid, GPP first removes the phosphoric acid group to obtain geraniol, second through a series of reactions such as oxidation and cyclization, the skeleton of iridoid, namely iridodial, can be obtained. Finally, through a series of reactions, the basic carbon skeleton of the secoiridoid, namely secologanin, is obtained[5561] (Fig. 2c). In the formation of triterpene saponins, the key step lies in the formation of the precursor, 2,3-oxidosqualene, a reaction that is catalyzed by squalene epoxidase (SE). There are many pentacyclic triterpenes in the Lonicera genus, the most important type being the hederin-type saponins with hederagenin as aglycones. Hederin-type saponins are produced after the synthesis of oleanolic acid from β-amyrin and catalyzed by β-starch synthetase (β-AS) and Oleanolic acid synthase (OAS)[62,63]. The skeletal modification of the triterpenoid saponins is mainly achieved via the activities of the CYP450 enzymes and UDP-glycosyltransferase (UGT). Hence, the corresponding aglycones are first obtained via oxidation by the CYP450 enzymes (e.g., CYP72A), and further subjected to glycosylation by the appropriate UGT enzyme[6365] (Fig. 2d). Skeletal formations of the iridoids and triterpene saponins in general have been well researched, but the same cannot be said about the enzymes involved in biosynthesis of these groups of compounds in the Lonicera genus. To fully utilize the iridoids and triterpene saponins in the Lonicera genus, it is necessary to further explore their biosyntheses with the view to enhancing and optimizing the process.

    Given the importance of the bioactive compounds in the Lonicera genus, continual isolation of these compounds using the traditional methods are not only tedious and time-consuming, but also unsustainable. With the development and application of microbial metabolic engineering, different strategies have been introduced to produce these bioactive compounds by heterologous synthesis (Table 1).

    Table 1.  Biosynthesis of Lonicera-specialized metabolites using metabolic engineering.
    Engineering bacteriaOperational methodsProductsYieldRefs
    S. cerevisiaeEliminate the tyrosine-induced feedback inhibition, delete genes involved in competing pathways and overexpress rate-limiting enzymesCaffeic acid569.0 mg/L[69]
    S. cerevisiaeEmploye a heterologous tyrosine ammonia lyase and a 4HPA3H complex composed of HpaB and HpaC derived from different speciesCaffeic acid289.4 mg/L[73]
    S. cerevisiaeSupply and recycle of three cofactors: FADH2, S-adenosyl-L-methion, NADPHCaffeic acid
    Ferulic acid
    Caffeic acid: 5.5 g/L;
    Ferulic acid: 3.8 g/L
    [117]
    E. coliKnocking out competing pathwaysCaffeic acid7,922 mg/L[118]
    E. coliArtificial microbial community, a polyculture of three recombinant Escherichia coli strainsChlorogenic acid250 μM[68]
    Cell-free biosynthesisExtract and purify spy-cyclized enzymes (CFBS-mixture)Chlorogenic acid711.26 mg/L[70]
    S. cerevisiaeThree metabolic engineering modules were systematically optimized: shikimate pathway and carbon distribution, branch pathways, CGA pathway genesChlorogenic acidFlask fermentation: 234.8 mg/L;
    Fed-batch fermentation:
    806.8 mg/L
    [119]
    E. coliUsing modular coculture engineering: construction of the defective strain improves the production and utilization of precursor substancesChlorogenic acid131.31 mg/L[122]
    E. coliIntroduce heterologous UDP-glucose biosynthetic genesLuteolin34 mg/L[120]
    Y. lipolyticaOverexpression of the key genes involved in the mevalonate pathway, the gene encoding cytochrome P450 (CYP716A12) to that encoding NADPH-P450 reductaseOleanolic acid129.9 mg/L[85]
    S. cerevisiaeImprove the pairing efficiency between Cytochrome P450 monooxygenase and reductase and the expression level of key genesOleanolic acid606.9 mg/L[121]
    S. cerevisiaeHeterologous expression and optimization of CrAS, CrAO, and AtCPR1, and regulation of ERG1 and NADPH regeneration systemOleanolic acid433.9 mg/L[123]
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    Due to the demand for CGA in the food, pharmaceutical, chemical, and cosmetic industries, the traditional means of obtaining the same requires a relatively longer period for plant maturation to obtain low yields of the desired product. This therefore brings into question the sustainability and efficiency of this approach. The alternative and sustainable approach has been to produce CGA using synthetic biology and metabolic engineering.

    Current research has sought to utilize Escherichia coli (and its mutant strain) and Saccharomyces cerevisiae to synthetically generate CGA and other flavonoids[6673]. For instance, Cha et al. employed two strains of E. coli to produce a relatively good yield of CGA (78 mg/L). Their approach was based on the ability of one strain to generate caffeic acid from glucose and the other strain to use the caffeic acid produced and quinic acid as starting materials to synthesize CGA[66]. Using a bioengineered mutant of E. coli (aroD mutant), Kim et al. increased the yield of CGA to as high as 450 mg/L[67]. Others have sought to increase the yield of CGA by employing a polyculture of three E. coli strains that act as specific modules for the de novo biosynthesis of caffeic acid, quinic acid and CGA. This strategy eliminates the competition posed by the precursor of CGA (i.e., caffeic acid and quinic acid) and generally results in improved production of CGA[68]. Saccharomyces cerevisiae is a chassis widely used for the production of natural substances from plants with an intimal structure that can be used for the expression of cytochrome P450 enzymes that cannot be expressed in E. coli. Researchers have used yeast to increase the production of organic acids[69]. A de novo biosynthetic pathway for the construction of CGA in yeast has been reported new cell-free biosynthetic system based on a mixture of chassis cell extracts and purified Spy cyclized enzymes were adopted by Niu et al. to a produce the highest yield of CGA reported so far up to 711.26 ± 15.63 mg/L[70].

    There are many studies on the metabolic engineering for the synthesis of flavonoids, but few on luteolin and its glycosides. Strains of E. coli have been engineered with specific uridine diphosphate (UDP)-dependent glycosyltransferase (UGT) to synthesize three novel flavonoid glycosides. These glycosides were quercetin 3-O-(N-acetyl) quinovosamine (158.3 mg/L), luteolin 7-O-(N-acetyl) glucosaminuronic acid (172.5 mg/L) and quercetin 3-O-(N-acetyl)-xylosamine (160.8 mg/L)[71]. Since most of the flavonoid glycosides synthesized in E. coli are glucosylated, Kim et al. in their bid to synthesize luteolin-7-O-glucuronide, deleted the araA gene that encodes UDP-4-deoxy-4-formamido-L-arabinose formyltransferase/UDP-glucuronic acid C-4'' decarboxylase in E.coli and were able to obtain a yield of 300 mg/L of the desired product[72].

    Terpenoidal saponins are mostly derived from slow-growing plants and usually possess multiple chiral centers[74]. Traditional isolation and even chemical synthesis of the terpenoidal saponins are both tedious and uneconomical for large-scale production. Therefore, it is necessary to find other ways to synthesize these compounds known to have diverse pharmacological functions.

    Heterologous synthesis has become an important way to improve the target products. With the development of synthetic biology, heterologous synthesis of triterpene saponins involves chassis of both plant and microbial origin. In this regard, Nicotiana benthamiana is a model plant species for the reconstruction of the biosynthetic pathways of different bioactive compounds including monoterpenes, hemiterpenes, and diterpenes[59,7577]. Aside from Nicotiana benthamiana, other plants have also been used as heterologous hosts[78]. Heterologous synthesis using microbial hosts mainly involves Saccharomyces cerevisiae and Escherichia coli[7981], and other microorganisms[82,83]. Comparatively, plants as biosynthetic hosts have the advantages of an established photosynthetic system, abundant supply of relevant enzymes, and presence of cell compartments, etc. They are however not as fast growing as the microorganisms, and it is also difficult to extract and separate the desired synthesized compounds from them as hosts.

    Although heterologous synthesis has many advantages, the premise of successful construction of synthetic pathway in host is to elucidate the unique structure of the compound and the key enzyme reaction mechanism in the biosynthetic pathway. There is little research on metabolic engineering of the hederin-type pentacyclic triterpene saponins in Lonicera, but there are studies on the heterologous synthesis of its aglycone precursor, oleanolic acid[84,85]. There is a dearth of scientific literature on key enzymes in the biosynthesis of pentacyclic triterpenoid saponins in the Lonicera genus.

    Scientific evidence by diverse research groups has linked members of the Lonicera genus to a wide range of pharmacological effects (Fig. 3). These pharmacological effects are elicited by different chemical constituents, much of the underlying mechanisms of which have been elucidated by the omics techniques. Here, we summarize the pharmacological effects and pharmacodynamics of the Lonicera genus in the last 6 years.

    Figure 3.  Schematic summary of four main pharmacological effects (anti-inflammatory, antimicrobial, anti-oxidative and hepatoprotective effects) of the Lonicera genus and the underlying mechanisms of actions.

    Bioactive compounds of plants in the Lonicera genus have demonstrated varying degrees of anti-inflammatory actions. In a recent study, Lv et al. showed that lonicerin inhibits the activation of NOD-like receptor thermal protein domain associated protein 3 (NLRP3) through regulating EZH2/AtG5-mediated autophagy in bone marrow-derived macrophages of C57BL/6 mice[86]. The polysaccharide extract of L. japonica reduces atopic dermatitis in mice by promoting Nrf2 activation and NLRP3 degradation through p62[87]. Several products of Lonicera have been reported to have ameliorative effects on DSS-induced colitis. Among them, flavonoids of L. rupicola can improve the ulcerative colitis of C57BL/6 mice by inhibiting PI3K/AKT, and pomace of L. japonica can improve the ulcerative colitis of C57BL/6 mice by improving the intestinal barrier and intestinal flora[88,89]. The flavonoids can also ameliorate ulcerative colitis induced by local enema of 2,4,6-trinitrobenzene sulfonic acid (TNBS) in Wistar rats by inhibiting NF-κB pathway[90]. Ethanol extract from L. Japonica has demonstrated the potential to inhibit the expressions of inflammatory cytokines in serum and macrophages of LPS-induced ICR mice[91]. The water extract of L. japonica and luteolin were found to exhibit their anti-inflammatory effects via the inhibition of the JAK/STAT1/3-dependent NF-κB pathway and induction of HO-1 expression in RAW263.7 cells induced by pseudorabies virus (PRV)[92].

    Existing scientific evidence indicates that the extracts of plants in the Lonicera genus exhibit strong inhibition against different pathogenic microorganisms. Phenolic compounds from L. japonica demonstrated a particularly significant inhibitory effect against Staphylococcus aureus and Escherichia coli, in vitro, making these compounds potential food preservatives[93]. Influenza A virus is a serious threat to human health. Recent research has found the ethanol extract of L. japonica to possess a strong inhibitory effect against H1N1 influenza virus-infected MDCK cells and ICR mice[94]. The incidence of the COVID-19 pandemic called to action various scientists in a bid to find safe and efficacious treatment[95]. Traditional Chinese medicines became an attractive alternative in this search. The water extract of the flower bud of L. japonica which has traditionally served as a good antipyretic and antitussive agent attracted the attention of researchers. Scientific evidences have confirmed that the water extract of L. japonica can induce let-7a expression in human rhabdomyosarcoma cells or neuronal cells and blood of lactating mice, inhibiting the entry and replication of the virus in vitro and in vivo[96]. In addition, the water extract of L. japonica also inhibits the fusion of human lung cancer cells Calu-3 expressing ACE2 receptor and BGK-21 cells transfected with SARS-CoV-2 spike protein, and up-regulates the expression of miR-148b and miR-146a[97].

    Oxidative stress has been implicated in the pathophysiology of many diseases, hence, amelioration of the same could be a good therapeutic approach[98,99]. In keeping with this therapeutic strategy, various compounds from the Lonicera genus have demonstrated the ability to relieve oxidative stress due to their pronounced antioxidant effects. For instance, the polyphenolic extract of L. caerulea berry was found to activate the expression of AMPK-PGC1α-NRF1-TFAM proteins in the skeletal muscle mitochondria, improve the activity of SOD, CAT and GSH-Px enzymes in blood and skeletal muscle, relieve exercise fatigue in mice by reducing oxidative stress in skeletal muscle, and enhance mitochondrial biosynthesis and cell proliferation[100]. The diverse health benefits of the anthocyanins from L. japonica have been mainly credited to their antioxidant and anti-inflammatory effects. The anthocyanin and cyanidin-3-o-glucoside have been reported to possess the potential to prolong life and delay senescence of Drosophila through the activation of the KEAP1/NRF2 signaling pathway[101].

    The liver is an essential organ that contributes to food digestion and detoxification of the body. These functions expose the liver to diverse toxins and metabolites. The Lonicera genus is rich in phytochemicals that confer protection on the liver against various toxins. The phenolic compound, 4, 5-di-O-Caffeoylquinic acid methyl ester was shown to be able to improve H2O2-induced liver oxidative damage in HepG2 cells by targeting the Keap1/Nrf2 pathway[102]. Hepatic fibrosis is a complex dynamic process, with the propensity to progress to liver cancer in severe cases. The L. japonicae flos water extract solution increased the cell viability of FL83B cells treated with thioacetamide (TAA), decreased the levels of serum alanine aminotransferase (ALT) and alkaline phosphatase (ALP), inhibited the transformation growth factor β1 (TGF-β1) and liver collagen deposition[103]. Sweroside, a secoiridoid glucoside isolate of L. japonica is known to protect the C57BL/6 mice liver from hepatic fibrosis by up-regulating miR-29a and inhibiting COL1 and TIMP1[104].

    Aside from the aforementioned, other pharmacological effects have been ascribed to the Lonicera genus. The ethanolic extract of L. caerulea has been reported to inhibit the proliferation of SMMC-7721 and H22 hepatoma cells, while its anthocyanins induced the apoptosis of tumor cells via the release of cytochrome C and activation of caspase[105]. AMPK/PPARα axes play an important role in lipid metabolism. A chlorogenic acid-rich extract of L. Japonica was found to significantly decrease the early onset of high-fat diet-induced diabetes in Sprague-Dawley rats via the CTRPs-AdipoRs-AMPK/PPARα axes[106]. In a high-fat diet-induced non-alcoholic fatty liver disease in C57BL/6 mice, treatment with L. caerulea polyphenol extract decreased serum inflammatory factors and endotoxin levels and the Firmicutes/Bacteroidetes ratio, an indication of its modulatory effect on the gut microbiota[107]. The iridoid-anthocyanin extract from L. caerulea berry contributed to alleviating the symptoms of intestinal infection with spirochaeta in mice[108].

    The traditional classification of the Lonicera genus based on the morphology of member plants is further categorized into two subgenera, Chamaecerasus and Periclymenum. The Chamaecerasus includes four categories, Coeloxylosteum, Isika, Isoxylosteum and Nintooa. The Periclymenum includes two categories, Subsect. Lonicera and Subsect. Phenianthi (Supplemental Table S1).

    High-throughput chloroplast genome sequencing of L. japonica found its length to be 155078 bp, which is similar to the structure of the typical angiosperm chloroplast genome. It contains a pair of inverted repeat regions (IRa and IRb, 23774 bp), a large single copy region (LSC, 88858 bp) and a small single copy area (SSC, 18672 bp)[109,110]. However, compared with chloroplast genomes of other plants, the chloroplast genome of L. japonica has a unique rearrangement between trnI-CAU and trnN-GUU[110]. Based on the phylogenetic analysis of the plastid genomes of seven plants in the Lonicera genus, 16 diverging hot spots were identified as potential molecular markers for the development of the Lonicera plants[111]. The phylogeny of Lonicera is rarely researched at the molecular level and the pattern of repetitive variation and adaptive evolution of the genome sequence is still unknown. Chloroplast genome sequences are highly conserved, but insertions and deletions, inversions, substitutions, genome rearrangements, and translocations also occur and have become powerful tools for studying plant phylogeny[112,113].

    We present here the phylogenetic tree of the Lonicera genus based on the published complete chloroplast genome sequences downloaded from the National Center for Biotechnology Information (NCBI) database using the Maximum likelihood method (Fig. 4). Based on our chloroplast phylogenies, we propose to merge L. harae into Sect. Isika and L. insularis into Chamaecerasus, but whether L. insularis belongs to Sect. Isika or Sect. Coeloxylosteum is uncertain. Based on protein-coding regions (CDS) of the chloroplast genome or complete chloroplast genomes, Liu et al. and Chen et al. supported the classification of the two subgenera in Lonicera[111,114]. Sun et al. and Srivastav et al. demonstrated a classification between the two subgenera with more species by using sequences of nuclear loci generated, chloroplast genome, and restriction site-associated DNA sequencing (RADSeq)[115,116]. However, our phylogenetic analysis and that of Sun et al. show relations within the subgenus Chamaecerasus are tanglesome in some respects[116]. Plant traits are affected by the environment to varying degrees. Since evidence of plant speciation is implicit in its genome sequence, comparative analysis at the molecular level provides a relatively accurate depiction of inherent changes that might have occurred over time. These findings suggest the need for more species of the Lonicera genus to be sequenced to provide a more accurate theoretical basis for the evolution of the Lonicera plants and a more effective revision in the classification of the Lonicera genus.

    Figure 4.  Phylogenetic tree of 42 species of the Lonicera genus based on complete chloroplast genome sequence data. The phylogenetic tree was constructed by the maximum likelihood method. Coeloxylosteum, Isika, Isoxylosteum, and Nintooa belong to Chamaecerasus and Subsect. Lonicera belongs to the Periclymenum. Chamaecerasus and Periclymenum are the two subgenera of Lonicera. 'Not retrieved' indicates that the species failed to retrieve a subordinate taxon in the Lonicera.

    The Lonicera genus is rich in diverse bioactive compounds with immeasurable prospects in many fields. Members of this genus have been used for thousands of years in traditional Chinese medicine for heat-clearing and detoxification. These plants generally have a good taste and form part of the ingredients of various fruit juices. In cosmetics, they are known to possess anti-aging and moisturizing functions. Plants of the Lonicera genus are also known for their good ecological adaptability and can be used to improve soil and ecological environment. Based on the value of the Lonicera genus, besides researching their use through molecular biological means, their efficient utilization can also be promoted in the following ways: (1) The stems and leaves of the plants could be developed for consumption and use since the chemical profiles of these parts do not differ significantly from the flowers. This way, the wastage of this scarce resource could be minimized or avoided. (2) Most of the Lonicera plants are vines or shrubs and their natural regeneration speed is slow, so the introduction and domestication of species could be strengthened to avoid overexploitation of wild resources.

    At present, only the research on the biosynthesis and efficacy of chlorogenic acid is quite comprehensive and has been used widely in various fields. There is limited research on various aspects of other bioactive compounds and should therefore be given priority in future research goals. Currently, the multi-omics analytical approach has gradually evolved as a reliable and helpful analytical platform. Hence, multi-omics research on the Lonicera genus could lead to discoveries in drug discovery and human health.

    The authors confirm contribution to the paper as follows: study conception and design, draft manuscript preparation: Yin X, Chen X, Li W, Tran LSP, Lu X; manuscript revision: Yin X, Chen X, Li W, Tran LSP, Lu X, Chen X, Yin X, Alolga RN; data/literature collection: Chen X, Yin X; figure preparation: Chen X, Yin X; figure revision: Alolga RN, Yin X, Chen X, Li W, Tran LSP, Lu X. All authors reviewed the results and approved the final version of the manuscript.

    All data generated or analyzed during this study are included in this published article and its supplementary information file.

    This work was partially supported by the National Natural Science Foundation of China (NSFC, Nos 82173918 and 82373983).

  • The authors declare that they have no conflict of interest. Xiaojian Yin is the Editorial Board member of Medicinal Plant Biology who was blinded from reviewing or making decisions on the manuscript. The article was subject to the journal's standard procedures, with peer-review handled independently of this Editorial Board member and the research groups.

  • Supplemental Fig. S1 lllustration of layer-by-layer electrostatic deposition for synthesis of secondary emulsion-2.5 % oil, OSA starch as primary emulsifier and chitosan as secondary emulsifier.
    Supplemental Fig. S2 lllustration of storage study at 37 °C where selected β-carotene loaded secondary emulsion along with bulk oil was exposed to light.
    Supplemental Fig. S3 Trend in β-carotene dissolved in bulk sunflower oil degradation during storage in dark, n ≥3.
    Supplemental Table S1 ANOVA table for storage parameters: For storage conditions, each data set had 3 source of variation (factors) and therefore, a 3-way ANOVA analysis was used to study the data. As indicated in the table, in all the data set studied, interactions were significant (green highlights) and therefore, single factor main effects are not to be interpreted.
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  • Cite this article

    Sivabalan S, Ross CF, Tang J, Sablani SS. 2024. Physical, thermal, and storage stability of multilayered emulsion loaded with β-carotene. Food Innovation and Advances 3(3): 244−255 doi: 10.48130/fia-0024-0022
    Sivabalan S, Ross CF, Tang J, Sablani SS. 2024. Physical, thermal, and storage stability of multilayered emulsion loaded with β-carotene. Food Innovation and Advances 3(3): 244−255 doi: 10.48130/fia-0024-0022

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Physical, thermal, and storage stability of multilayered emulsion loaded with β-carotene

Food Innovation and Advances  3 2024, 3(3): 244−255  |  Cite this article

Abstract: Carotenoids are colored bioactive substances increasingly used due to their antioxidant properties, vitamin A precursor role, and ability to function as a natural food color. Knowledge of carotenoid behavior during high-heat processing and subsequent storage in emulsified food matrix is essential to expand their application natural food colors and neutraceuticals. Firstly, the physical, thermal, and colloidal stability of emulsions constructed from octenyl succinic anhydride-modified starch (OSA starch)-chitosan multilayered interfaces were investigated. Results of charge reversal from −32.4 ± 1.9 mV to +38.0 ± 0.8 mV indicate that multilayered interfaces were formed in emulsions. As measured by Z-average size, the emulsions were stable after the thermal treatment at 121 °C for 60 min, thus demonstrating a novel heat-stable multilayered emulsion. Subsequently, a select multilayered emulsion was loaded with β-carotene, and its storage stability was assessed. The degradation of β-carotene in an oil-in-water emulsion was better described with zeroth order kinetics; β-carotene dissolved in bulk oil was better described using a second-order kinetic equation. The presence of an encapsulating material around the oil droplets loaded with β-carotene enhanced its stability, which makes it instrumental in extending shelf-life and maintaining a consistent appearance. The results can be used to predict the availability of β-carotene during storage.

    • Carotenoids are a group of colored substances commonly occurring in green and yellow leafy vegetables with their structure consisting of a 40 carbon polyene backbone[1]. Carotenoids are further classified into carotenes (β-carotene and lycopene) with the structural formula C40H56 and xanthophylls (lutein, zeaxanthin, cryptoxanthin, neoxanthin, and violaxanthin) that have oxygen in their molecular structures[2,3]. Carotenoids play a vital role in quenching reactive oxygen species and inhibiting tumor growth[2]. In addition to being a dietary source of retinoids, carotenoids are associated with several therapeutic functions. Protective effects against dangerous disorders such as degenerative eye diseases and cancer have been reported[3,4].

      Carotenoids exist in many different chemical forms. Among the various dietary carotenoids, provitamin A activity is expressed by those with at least one terminal β-ionyl ring. β-carotene, α-carotene and β-cryptoxanthin show provitamin A activity while others such as lycopene, lutein, and zeaxanthin do not convert into vitamin A in the human body[2]. Vitamin A, chemically, known as retinoids are absorbed in the intestines and stored in the liver. The conversion occurs in living systems with the aid of enzymes. The two carotenoid cleavage enzymes β-Carotene ′ oxygenase-1 and β-carotene 9′,10′ oxygenase-2 occurring in humans are known to aid in bioconversion of β-carotene to retinoids[2,5]. β-carotene is the most commercialized and well-established representative of lipophilic nutraceutical owing to its function as a natural coloring material, antioxidant activity, and dietary source of vitamin A. β-carotene is naturally present in trans isomeric form in foods while other cis isomers may also occur[4,6].

      The knowledge of the factors influencing the stability of naturally colored β-carotene emulsions in food systems is essential in developing their new applications. There is an increasing inclination for consumers to prefer natural colors over synthetic colorants in foods and beverages. With increasing consumer preference for natural sources, the use of colors such as carotenoids in food and beverages has increased[79]. However, natural alternatives are susceptible to environmental stresses encountered during processing including heat, light, pH, and oxygen. β-carotene is a bioactive substance that degrades with time, and factors such as high temperature, light exposure, and air contact expedite the degradation.

      Color is an essential marker for quality; thus there is an interest in understanding the stability of natural food colorants to degradation factors[8,10]. Oxidation and isomerization causes color loss of carotenoids and nutritive value[11]. Naturally occurring β-carotene is a favored means of enhancing food color[12]. An exceptional way to apply it to food is through encapsulation[12,13].

      Encapsulation encases a nutraceutical bioactive substance in a protective matrix with the aim of improving thermal endurance, solubility, chemical stability, bioavailability and flavor masking[14,15]. While bioactive substances are fortified, competent encapsulation structures are essential to preserve the active material and prevent degradation during processing[12]. Encapsulated structures are known to offer protection to the encased substance, enhance solubility in aqueous matrices, deliver to the target location and retard the rate of degradation reactions[3]. Encapsulation can be achieved by emulsification, spray drying, coacervation, freeze-drying, and extrusion. Various types of emulsions that can be utilized for encapsulation of bioactive materials include conventional emulsions, multilayered emulsions, protein-polysaccharide conjugate stabilized emulsions, Pickering emulsions and nanoemulsions[16].

      Oil-in-water (o/w) emulsions are popular for improving solubility of liposoluble nutraceuticals, triggered or controlled delivery, decelerating isomerization and degradation due to its simple and cost-effective preparation method[3,17]. Emulsifiers at the oil-water interface play a pivotal role in regulating performance and functionality[3]. Natural emulsifiers are preferred by consumers due to their perceived health benefits. In a shift from well-performing conventional surfactant-based emulsifiers towards more natural Generally Recognized as Safe (GRAS) certified food-grade emulsifiers, proteins, and polysaccharides serve as the two main classes to work with[18,19]. Macromolecules such as proteins and polysaccharides serve as naturally derived emulsifiers. Several proteins have been proven to produce small-sized emulsion droplets due to their amphiphilic nature[3,20]. However, proteins at elevated temperatures induce emulsion destabilization due to their instability arising from pH variation, temperature fluctuation, or high ionic strength[3,21]. Long-term emulsion stability is mainly offered by stabilizers[18]. Aqueous phase polysaccharides can function as stabilizers in emulsified systems, increasing viscosity and consequently preventing droplet coalescence[22]. Polysaccharides such as modified starches largely stabilize the emulsion interface by steric repulsion thereby making the emulsified system they stabilize less likely disturbed by stresses such as heat and ionic strength[21].

      This research was designed to explore the stability of multilayered emulsions made of octenyl succinic anhydride (OSA) modified starch and chitosan for enhanced stability of β-carotene in thermal treatments and during storage. Heat stability was specifically considered to expand application in thermal processing scenarios like sterilization. OSA starches are a type of modified starch and are derived from waxy maize, wheat, corn, or tapioca where the chemical modification confers unique functional properties, the most important of it being emulsification[23,24]. Chitosan is a polycationic linear heteropolysaccharide consisting of N-acetyl-2-amino-2-deoxy-ᴅ-glucopyranose and 2-amino-2-deoxy-ᴅ-glucopyranose bonded by β–(1→4) glycosidic linkages. Commercial chitosan is derived by alkaline treatment of chitin[2530].

      Studies to understand the carotenoid stability in processing and storage are essential. Many studies have explored the stability of emulsified β-carotene when subjected to heat[31,32], light[20,31,33] and in storage[3,3436]. There has been little published information on the stability of β-carotene incorporated emulsified and bulk oil systems within transparent polymeric pouches during exposure to light. Fortified food stored in transparent pouches or PET bottles may be exposed to light during transportation in the supply chain and shelf display in grocery stores. Ensuring the stability of natural pigments in a wide variety of conditions is essential to expand their application in the food industry[12,37]. The present research attempts to systematically study the stability of β-carotene to light exposure and in storage to elucidate the extent of protection offered by encapsulated matrices against the non-encased bulk matrices. β-carotene was mixed in bulk triglyceride oils as well as dispersed in emulsified droplets of the corresponding oils. In preliminary storage tests on the degradation of β-carotene with unsaturated fats, we observed no specific trend in its degradation. Therefore, two saturated fat materials were chosen, medium chain triglyceride (MCT oil) and glycerol trioctanoate (GTO), as lipophilic carriers for β-carotene. GTO is a pure triglyceride while MCT oil contains a mixture of capric, caprylic, and lauric fatty acids according to label specifications.

    • Synthetic β-carotene, GTO, Chitosan (degree of deacetylation ≥ 75%), and citric acid were procured from Millipore Sigma Inc. (St. Louis, MO, USA). MCT oil Spring Valley water was purchased from a local store, Walmart, (Pullman, WA, USA). OSA starch was kindly supplied by Ingredion Incorporated (Bridgewater, NJ, USA). Deionized water obtained from Milli-Q Reagent Water System, (Massachusetts, USA) was utilized throughout the study. High barrier multilayer polymeric film (Structure - Coated PET//PA//PP) with an oxygen transmission rate of 1.05 cm3 m−2 day−1 and water vapor transmission rate of 5.11 gm−2 day−1 was used to prepare small size (3 in × 2.25 in) pouches.

    • Oil-in-water emulsions with bilayer interface were constructed using two polysaccharides — OSA starch and chitosan. The multilayered emulsions were prepared by following layer-by-layer deposition of biopolymers to facilitate electrostatic interaction. First, an oil-in-water primary emulsion composed of OSA starch was prepared by a two-step homogenization process wherein an oil phase and a primary aqueous phase were homogenized using a rotor-stator homogenizer (Kinematica Polytron PT 2500E, Bohemia, NY, USA) at 7,000 rpm for 3 min. The primary course emulsions were further broken down by means of ultrasound treatment using a probe sonicator of 0.25 in diameter tip of titanium construction (Fisher Scientific Sonic Dismembrator Model 100 Hampton, NH, USA). The primary aqueous phase consisted of varying concentrations, 1.0%–2.5% w/v, of OSA starch dispersed in deionized water. Therefore, the primary emulsion had a final composition of 5% oil and varying concentrations of OSA starch as emulsifier.

      Secondary emulsions were formed by slow addition of primary emulsion to the secondary aqueous phase with continuous homogenization, thus promoting the deposition of chitosan on primary emulsion droplets (Supplemental Fig. S1). The secondary aqueous phase consisted of varying proportion of Chitosan (0.025%–0.80% w/v solution) dissolved in 1% citric acid solution and was added with the primary emulsion at the ratio of 1:1. Overnight stirring was allowed for the complete hydration of biopolymers. For β-carotene loaded lipid phase, β-carotene was mixed with 10 mL MCT oil or GTO in an ultrasonic bath for uniform dissolution. Hence, secondary emulsions would have 2.5% oil, 1% OSA starch as primary emulsifier and 0.0125%–0.40% chitosan as a secondary emulsifier.

    • Particle size distribution, Zeta (ζ) potential, and Z-average size were determined using a Zetasizer Nano ZS (Malvern Instruments, Malvern, UK). The instrument used dynamic light scattering to measure the Brownian movement of particles and correlate it to the size under the theory that smaller particles produce faster movement. In this measurement, a monochromatic light source, laser at 633 nm, is directly beamed on a suspension containing particles; the scattered light intensity is measured as a function of time. The scattered light was measured at 173° and refractive indices of 1.48 and 1.33 were specified for oil and water, respectively[38]. The emulsions were diluted 100 times before the measurements to prevent multiple scattering effect. ζ-potential reflects the charge possessed by emulsified droplets and samples were equilibrated at 25 °C before measurement.

    • Emulsions were treated to a temperature of 121 °C for 60 min to simulate the maximum sterilization condition using an oil bath and a metal cell assembly as described previously[32] wherein the emulsions were filled in PET-based pouches. Prepared 10 mL emulsions were filled in PET-based pouches and sealed. The pouches were placed within an air-tight cell and immersed in an oil bath (Thermo HAAKE W15, Waltham, MA, USA) filled with Fisher's Bath oil. Once the required heating time was completed, the cells were immediately cooled in an ice bath. The particle size distribution of the treated emulsions was measured to evaluate changes in droplet size due to heat treatment.

    • To understand the state transition of prepared emulsions during heat treatments, thermograms of multilayered emulsions made up of OSA starch and chitosan interfacial materials were obtained using differential scanning calorimeter (TA Instruments, DSC Q2000 V24.11 Build 124, Newcastle, DE, USA). About 6–10 mg prepared emulsions were measured into a Tzero aluminum pan and sealed hermetically. The emulsions were subjected to temperature extremes by first freezing it to –50 °C at the ramp rate of 5 °C/min and then controlled heating at 5 °C/min up to 121 °C under nitrogen atmosphere. Another sealed pan with air served as a reference pan.

    • Continuous light exposure was achieved using two 7 watts 12-inch LED under cabinet lights of color temp 5,000 K placed ~18 cm above the sample with the flat surface of pouch facing light. The level of light exposure of samples within the incubator was measured as 3,530 ± 200 Lux using Dr. Meter Digital Illuminance Light meter (Model LX 1330B, range 0.1–200,000 Lux). The temperature of samples was maintained at 37 °C using an incubator (Heratherm IGS60 ThermoScientific, Langenselbold, Germany). The samples that were treated in the dark were placed in the incubator covered in aluminum foil (Supplemental Fig. S2).

    • β-carotene was estimated by solvent extraction method[39] with slight modification. Briefly, a mixture of chloroform and methanol in the ratio 2:1 (v/v) was used as the extraction medium. Two hundred μL of emulsion or oil was withdrawn and 1.5 mL of organic solvent mixture was added, vortexed thrice, and centrifuged at 210 g for 5 min. The clear extract was carefully withdrawn, and absorbance was measured at 450 nm against a solvent blank. A standard curve was prepared with known quantities of β-carotene dissolved in the organic phase and used to estimate concentration from absorbance (R2 = 0.997).

    • Thiobarbituric acid reactive substance assay was used to estimate products of oxidation[40]. Preparation of TBA-BHT Reagent was made as follows: 2% v/v solution of Butylated Hydroxy Toluene (BHT) was prepared in ethanol—BHT solution, 75 g of trichloroacetic acid (TCA), 1.875 g of 2-thiobarbituric acid (TBA), 8.8 mL of 12 M HCL and 414.5 mL of Millipore water was dissolved together to form the TBA solution. TBA-BHT reagent was prepared by slowly mixing 500 mL of TBA solution and 15 mL of BHT solution. The reagent was stored in the dark until use. Preparation of Malondialdehyde (MDA) included the following steps: MDA standard curve was constructed using 1,1,3,3-Tetraethoxypropane (TEP). TEP dissolves in water and produces MDA. For the preparation of calibration standards, 1 μM of MDA stock solution was prepared by dissolving 25 μL TEP in 100 mL of 1% v/v H2SO4. Two mL of this stock solution was made up to 50 mL 1% v/v H2SO4 to yield 40 nM standard solution and incubated at room temperature for 2 h. Standard solution was further diluted to 5, 10, 15, and 20 nM using 1% v/v H2SO4. These solutions of known concentration underwent the same procedure as the sample and absorbance was measured at 532 nm. A straight line with R2 = 0.999 was obtained as standard curve. A blank with 1% v/v H2SO4 was used for spectrophotometer (UV160U, Shimadzu Corporation, Kyoto, Japan). 1.6 mL of the sample was vortexed for 30 s with 3.2 mL of TBA-BHT solution, followed by heating in a boiling water bath for 15 min. The mixture was cooled to room temperature and centrifuged (Ohaus Frontier Centrifuge FC5706, Parsippany NJ, USA) at 4,430 g for 10 min. The supernatant was carefully transferred into a transparent disposable cuvette to record its absorbance at 532 nm.

    • Sample color in reflectance mode with specular component excluded was recorded as L* (lightness) values ranging from 0–100, a* greenness (–) to redness (+) and b* blueness (–) to yellowness (+) respectively using a Spectrophotometer CM-5, Konica Minolta, New Jersey in CIELAB color space. The range for a* and b* are approximately 80 in positive and negative axis[41]. 2 mL samples were transferred to a clear petri dish of dimensions 35 mm diameter and 10 mm height. A measurement aperture of 8 mm , D65 illuminant, and 10° observation angle was used to record color values. L*, a*, and b* were used to estimate change in color ΔE using Eqn (1):

      ΔE=(LL0)2+(aa0)2+(bb0)2 (1)

      where, L*, a*, and b* are for sample stored at 37 °C for various time and L0*, a0*, and b0* are values at day 0[42].

    • Kinetic change in quality attribute C is expressed by Eqn (2)[42].

      dCdt=k[C]n (2)

      where, [C] denotes the concentration of substrate at a given time t, k is the rate constant of the degradation reaction, 'n' stands for order of reaction.

      Integrating Eqn (2) with an initial substrate concentration of [C0] for zeroth (Eqn 3), first (Eqn 4), and second (Eqn 5) order we get the following:

      [C]=[C0]kt (3)
      ln[C]=ln[C0]kt,[C]=[C0]ekt (4)
      1[C]=1[C0]kt,[C]=[C0]1+kt[C0] (5)
    • Analysis of the means and standard deviations was performed in Microsoft Excel (Office 365, Microsoft Corp., Redmond, WA, USA). Analysis of variance test (ANOVA) was performed using 'agricolae' package in R-Studio version 1.4.1717 following which, Tukey's HSD procedure was used to identify the means that differed significantly. The interaction effects between the independent variables carrier oil type (MCT and GTO), light/dark storage, and storage days were included in the model. The independent variables assessed include β-carotene concentration, color values (L*, a*, b*), and TBARS.

    • In this section, stable multilayered emulsions were identified and selected based on particle characteristics including Z-average size and ζ-potential as well changes due to heat treatment.

    • The Z-average size of OSA-starch primary emulsions was ~250 nm for all the concentrations studied. There was no significant difference in Z-average size or ζ-potential of emulsions (Table 1) stabilized by OSA starch concentrations of 1.0%, 1.3%, 1.7%, 2.0%, or 2.5% (p > 0.05). A range of biopolymer concentrations were attempted to ascertain that the interface was not short of complete coverage due to lack of emulsifier. There was sufficient biopolymer to coat the surface at 1% concentration itself. The constant ζ-potential value ~ −30 mV within the range of 1.0%–2.5% w/v OSA-starch adds to evidence that biopolymer coverage around native oil droplets is sufficient at 1% OSA-starch concentration. The value of ζ-potential is considered favorable as higher repulsive forces exist between droplets, thus preventing them from coming closer and coalescing.

      Table 1.  Z-average size and ζ-potential of primary emulsions prepared with varying concentrations of OSA starch (n ≥ 3).

      OSA-starch concentration
      (% w/v)
      Z-average size
      (nm)
      ζ-potential
      (mV)
      1.0 255 ± 28.3a −30.5 ± 1.3A
      1.3 249 ± 8.2a −30.0 ± 2.9A
      1.7 244 ± 1.5a −30.6 ± 2.4A
      2.0 258 ± 0.6a −31.6 ± 2.1A
      2.5 244 ± 1.4a −30.2 ± 1.9A
      Note: Means with same superscript are not significantly (p > 0.05) different.

      The ζ-potential obtained in the present study is in agreement with observations made by Paulo et al.[43], where ζ-potential for modified starch concentrations of 1.4% and 3.6% was recorded. It was hypothesized that a lower biopolymer concentration would be detrimental to emulsion stability upon excessive heating, and hence decided to go with the higher end of the concentration spectrum. A higher concentration was chosen with the premise that the unabsorbed polysaccharide would offer additional stability during heat treatment by the mechanism of steric stabilization. The adsorption of surface-active agent depends on its diffusion coefficient from bulk to the interface which in turn depends on the molecular weight of the emulsifier molecule. Modified starch falls on the higher end of molecular weight that stabilizes the interface (against proteins and surfactants) with a molecular weight of MW 8.6 × 106 g mol−1 and, therefore, requires larger quantities for emulsion stabilization. The OSA substitution of in a largely hydrophilic starch yields modified starch, thus offering the essential non-polar hydrophobic moiety[43].

      In some cases, emulsion stabilization can benefit from excess polysaccharides present in the aqueous phase. This is due to emulsion acting like a stabilizer of the aqueous phase by viscosity modification and thus preventing droplets from coming closer and coalescing into larger droplets[22]. In other cases, care must be taken to prevent excess polysaccharides as it may be detrimental to emulsion physical stability by promoting bridging flocculation[44]. Hence, it is essential to establish the amount of emulsifier that is necessary to fully coat the oil droplet surface area for multilayer emulsions.

    • The ζ-potential of emulsions exhibited a characteristic charge reversal with the layer-by-layer deposition of chitosan over primary droplets emulsified with OSA starch. The overall charge on primary OSA starch emulsions went from highly negative (−32.4 ± 1.9 mV) to highly positive charge (+38.0 ± 0.8 mV). This can be attributed to the electrostatic adsorption of cationic chitosan on anionic OSA starch-stabilized oil droplets. Such charge reversal on the droplet surface has been reported by several researchers[17,45,46] where layer-by-layer deposition of protein or polysaccharide was utilized. From Fig. 1, it can be observed that the magnitude of the positive charge increased with increasing concentration of chitosan. It is likely that with added chitosan (0–0.3% chitosan), the previously anionic droplet surface was gradually covered with chitosan and became progressively cationic. Eventually, the surface charge evened out around 0.3% w/v chitosan. Here the droplet surface was saturated with chitosan and any further addition would not alter the ζ-potential substantially. The large positive charge recorded by emulsions around 0.3% w/v chitosan could contribute considerably to droplet stability as the electrostatic repulsion between the oil particles would keep them apart and prevent them from coming closer and coalescing.

      Figure 1. 

      Charge reversal when secondary biopolymer chitosan was deposited over a primary emulsion interface, n = 3. Different letters represent significant difference at p < 0.05.

      Using OSA-starch emulsions with a net negative charge of −31.6 ± 2.1 mV and Z-average size of 257.8 ± 0.6 nm as a primary base, the bilayer interface was prepared by layer-by-layer electrostatic deposition of oppositely charged chitosan. The size of secondary OSA-starch–chitosan emulsions prepared with varying proportions of chitosan ranging from 0.0125%–0.4% w/v is plotted in Fig. 2 along with the size values obtained for those emulsions when treated to heat at 121 °C for 60 min. Secondary emulsions with varying concentrations of chitosan showed similar sizes in the range of 290.4–388.5 nm for all chitosan concentrations studied. When these emulsions were treated at 121 °C for 60 min, emulsions prepared with lower chitosan recorded large increases in size. From Fig. 2, the size of emulsions formulated with 0.0125% chitosan increased sharply, from 314 ± 6.3 nm to 1,521 ± 226 nm when subjected to heat, indicating emulsion destabilization by flocculation or agglomeration. In all the other concentrations of chitosan studied between 0.025% and 0.4% w/v chitosan, secondary emulsions exhibited a stable behavior with a small increase in size. Stability after heat treatment was additionally reflected by the lack of visible emulsion destabilization phenomena such as oiling off or coagulation. Except the lowest chitosan concentration studied (0.0125%), OSA-starch–Chitosan secondary emulsions were in a consistent range of 377 ± 2.9 nm to 472 ± 8.9 nm after treatment at 121 °C for 60 min. Therefore, Z-average size measurements recorded a slight increase before and after treatment at 121 °C for 60 min, which is an acceptable change given that they underwent extreme thermal treatment. OSA starch-chitosan emulsions exhibit exceptional physical stability which could be attributed partly to the steric stabilization function of secondary emulsifiers.

      Figure 2. 

      Change in Z-average size of emulsions before and after heat treatment at 121 °C for 60 min, n = 3. Error bars represent standard deviation.

      A DSC thermogram of emulsion subjected to temperature extremes is given in Fig. 3. The emulsion samples indicate an endothermic peak at about –10 °C on freezing from 25 to –50 °C, indicating freezing, and an exothermic peak on heating around 0 °C, indicating melting of emulsions. There are no degradation peaks observed during the rest of the heat ramp-up to 121 °C, indicating that the emulsions were stable to the heat treatment. Consistent thermograms were obtained for other multilayered emulsions with varying chitosan concentration (0.05%–0.4%) studied (Fig. 3b). A DSC study on hydrogels produced comparable patterns in crystallization and freezing[47] and can be considered characteristic of dispersions.

      Figure 3. 

      (a) DSC thermogram of emulsions (0.35% final chitosan concentration) showing characteristic peak for crystallization and melting of water. Inset represents part of the thermogram that was heated from –50 to 121 °C. (b) Stacked thermograms subjected to temperature extremes, ~ –50 to 121 °C, for comparison of varying concentration of emulsifier.

      Emulsions at 0.35% chitosan concentration were selected for further study as they formed droplets with large positive charge, monodisperse particle size distribution (Fig. 4) and negligible change upon heating indicating stable dispersions. Additionally, a higher concentration of interfacial material was found to be more effective at preventing β-carotene from degradation due to the formation of thicker interfaces around emulsified oil droplets[37].

      Figure 4. 

      Particle size distribution of (a) untreated and (b) heat treated emulsions prepared with the same concentration of chitosan (0.35%). Each chart displays distribution of three replicates (Series 1, 2 and 3).

    • This study aimed at understanding the rate of β-carotene degradation with and without light exposure at accelerated storage of 37 °C that could be translated as the shelf life of food material. The effect of light exposure on the destruction of β-carotene is illustrated in Fig. 5a for β-carotene dissolved in bulk oil and Fig. 5b β-carotene dispersed in emulsified form enclosed within polymeric pouches over 25 d after which β-carotene degraded in all the systems studied. β-carotene in bulk oil and emulsified form were parallelly stored under dark and light storage conditions to establish its storage stability on exposure to light.

      Figure 5. 

      β-carotene degradation in (a) bulk oils and (b) emulsions during storage at 37 °C, n = 4.

      In the case of bulk oil, both dark and bright storage conditions produced slightly different results. Up to 5 d, β-carotene degradation was similar for both dark and light storage conditions. After the initial period, storage in the dark recorded lower degradation than storage in the light. For instance, on day 15, the amount of β-carotene in bulk MCT oil was 44 μg/mL (14.7% retention) when stored in light while in the absence of light, a higher amount of 74 μg/mL (24.7% retention) of β-carotene was recorded. It was noted that up to day 20, the bulk oil stored in the dark had slightly better β-carotene retention than that exposed to light. The results are consistent with the results of Liang et al.[21] who stored β-carotene dissolved in bulk MCT oil under light and dark storage at 25 °C. Their results indicate that β-carotene under dark degraded completely in 10 d and under light in 7 d. In a pure state, β-carotene undergoes auto-oxidation in the presence of light and oxygen as it interacts with free radicals such as peroxyl radicals[2].

      For β-carotene in oils, a significant 3-way interaction was observed between carrier oil type, light /dark storage, and storage days (F = 9.77; p < 0.05). Significant 2-way interaction was observed between carrier oil type and storage days (F = 21.6; p < 0.05), light/dark storage and storage days (F = 55.6; p < 0.05) and carrier oil type and light/dark storage (F = 36.4; p < 0.05). Each factor effect cannot be interpreted as interaction effects are significant. For a* of emulsions, a significant 3-way interaction was observed between carrier oil type, light/dark storage and storage days (F = 42.8; p < 0.05). Significant 2-way interaction was observed between carrier oil type and storage days (F = 172; p < 0.05), light/dark storage, and storage days (F = 792; p < 0.05) and carrier oil type and light/dark storage (F = 104; p < 0.05). For b* of emulsions, significant 3-way interaction was observed between carrier oil type, light /dark storage, and storage days (F = 103; p < 0.05). Significant 2-way interaction was observed between carrier oil type, and storage days (F = 76.2; p < 0.05) and light/dark storage and storage days (F = 4,660; p < 0.05). No significant 2-way interaction was evaluated between carrier oil type, and light/dark storage (F = 0.04; p > 0.05) (Supplemental Table S1).

      It can be seen from Fig. 5 that the degradation of β-carotene occurred over a span of about 25 d under dark and lit storage at 37 °C. β-carotene loaded canola oil emulsions made from OSA-starches emulsifiers showed nearly complete degradation in 13 d at 55 °C storage[35]. Similarly, in the present study, β-carotene completely disappeared in 25 d whether stored in the dark or under light conditions. To extend the retention of β-carotene for a longer period in an emulsified system, multiple antioxidants may need to be used to produce a synergistic effect. For instance, Yi et al.[3] has shown the survivability of β-carotene for 30 d in 50 °C with the aid of emulsions made from whey protein isolate-dextran-resveratrol. Resveratrol is a polyphenol compound with a myriad of biological functions including antioxidant, anti-ageing, anti-obesity anti-viral, anti-inflammatory and antitumor activity[3,48]. The presence of additional antioxidant resveratrol at the interface, besides β-carotene could scavenge and chelate pro-oxidants that extended the period of retention of β-carotene within the oil phase[3]. In addition, proteins such as WPI could have additional antioxidant effect owing to their amino acid residues that have the potential to scavenge reactive oxygen species and chelate metal ions[49]. Therefore, the amount of protection offered by the interfacial layer to β-carotene stability is pronouncedly dependent on the antioxidant nature of emulsifiers.

      For β-carotene of emulsions, a significant 3-way interaction was observed between carrier oil type, light /dark storage and storage days (F = 18.2; p < 0.05). Significant 2-way interaction was observed between carrier oil type and storage days (F = 7.83; p < 0.05), light/dark storage and storage days (F = 97; p < 0.05) and carrier oil type and light/dark storage (F = 8.20; p < 0.05).

      Figure 5 indicates that for the first 5 d of storage, the presence of light did not affect β-carotene retention. After 5 d, the experiments indicated that among emulsified systems studied, those stored in lighted condition degraded faster as compared to dark storage (Fig. 5b). Irradiation of β-carotene loaded emulsions with UV light were found to remarkably diminish its concentration. Guo et al.[20] observed an 82% loss of β-carotene in emulsions crafted with High Methoxyl Pectin-Rhamnolipid-Pea Protein Isolate-Curcumin complex on exposure to UV light for only 5 h. Interestingly, β-carotene loaded emulsions synthesized with OSA-modified starches recorded no significant difference between those kept under light and dark storage conditions at 25 °C[21].

      The change in color of emulsions stored at 37 °C for 25 d in the presence and absence of light with respect to freshly prepared emulsions on day 0 is illustrated in Fig. 6. β-carotene degradation resulted from isomerization and oxidation leading to loss of pigmentation. Degradation of β-carotene is not only detrimental to the biological activity of the nutraceutical but also to perceived color[3]. Therefore, the amount of remaining in emulsions were determined colorimetrically. Both emulsion carrier oils, MCT and GTO produced similar color changes (Fig. 6a & b). The chain length of fatty acids in commercial MCT oil and analytical GTO is very similar. This may explain the similarity in the trends of both carriers. The presence or absence of light had a remarkable effect on ΔE of emulsions. During 25-d storage period, ΔE spaned over a small range of 16.2–24.0 for MCT emulsion and 15.7–22.4 for GTO emulsion in the dark. On the other hand, ΔE ranged from 19.1–43.1 for MCT emulsions and 19.3–37.7 for GTO emulsions. Exposure to light increased the lightness of emulsions and reduced the yellowness much faster than when stored in the dark and showed a greater degree of ΔE variation. Color change in oils was an assortment of fluctuating measurements and therefore was not considered for making reliable assessments. A perceptible color change is known to happen when ΔE is in the range of 3.5–5 and ΔE between 1–2 can be visualized by an experienced observer[50]. All the emulsions have shown ΔE that can be visualized.

      Figure 6. 

      Change in color of emulsions with (a) MCT carrier oil, and (b) GTO carrier oil subjected to storage under dark and light condition. Components (c) L*, (d) a*, and (e) b* of CIELAB color coordinates during storage at 37 °C, n = 3.

      The CIELAB color components measured for emulsions stored under dark and light conditions are illustrated in Fig. 6c, d & e. For emulsions stored at 37 °C lightness L* increased slightly while a* and b* associated with β-carotene pigmentation dropped substantially. The drop in a* was gradual throughout the storage period while the drop in yellowness of emulsions was more prominent during the first 5 d of storage. Yellowness disappeared at different rates for emulsions stored in the dark and those exposed to light. After the initial drop in b* (to day 5) emulsions stored in the dark, essentially showed no difference until the last day of study. On the other hand, the yellowness of emulsions exposed to light continued to deplete up to day 25. A similar change in yellowness was observed by Sweedman et al.[35] where a rapid drop in the first 24 h of storage was recorded, followed by an almost constant trend for a storage period of 8 d at 55 °C. It is worthwhile to note that the emulsions synthesized by Sweedman et al.[35] has OSA starch as an interfacial material as in the present study. Therefore, exposure to light has a pronounced effect on the yellowness of β-carotene emulsions.

      For L* of emulsions, significant 3-way interaction was observed between carrier oil type, light/dark storage, and storage days (F = 455; p < 0.05). Significant 2-way interaction was observed between carrier oil type and storage days (F = 454; p < 0.05), light/dark storage, and storage days (F = 908; p < 0.05) and carrier oil type and light/dark storage (F = 1,127; p < 0.05).

      In summary, for short-term storage, the presence of light did not affect β-carotene stability contained within transparent polymeric pouches — both for bulk oil and emulsified systems. However, for prolonged storage, it is advisable to store β-carotene-containing systems in the dark or in light-blocking packaging material.

    • The two lipids used, namely MCT and GTO, exhibited similar behavior under light and dark storage at 37 °C. MCT oil mainly consists of medium-chain triglycerides and is made of saturated fatty acids. GTO also has saturated fatty acids and is a pure substance consisting of C8:0. Our previous study utilized vegetable oil made up of a variety of unsaturated fatty acids and residual antioxidants such as α-tocopherol. The sunflower oil-in-water emulsions of chitosan self-aggregates during storage at 37 °C in the dark showed an almost constant level of β-carotene during the 13-d storage period[31]. The choice of lipid carrier in the present study was made such that we eliminate the interference from such antioxidants that may be present in vegetable oil carriers so that β-carotene is the sole antioxidant in the emulsion during storage. By using a pure saturated lipid, it is expected that the main mechanism of degradation is by β-carotene alone[51].

      As previously observed, the β-carotene loaded emulsions and bulk oils made from GTO and MCT carrier oils degraded in 25 d with the bulk oils exhibiting faster disappearance than emulsions. Similar behavior in MCT bulk oil incorporated with β-carotene was observed, where there was no detectable pigment after 14 d when stored at 20 °C[51]. In the case of β-carotene loaded emulsions, the degradation reported is dependent on the nature of the lipid carrier used. For instance, when vegetable oils such as corn oil or canola oil are used as carrier oil for β-carotene loading, a longer storage stability (> 30 d) of bioactive is reported[3,52]. A study comparing corn and MCT oils with sodium caseinate as interface stabilizing material found that at a storage of 25 °C, 75% of β-carotene was lost in MCT oil emulsions while only 22% β-carotene loss was reported for corn oil emulsions. Given the fact that storage conditions and emulsion interfacial composition were the same, the oil nature was the main factor that could have produced such a prominent difference. Vegetable oils may contain traces of antioxidants such as carotenoids and α-tocopherol. Though the amounts of such residual traces of α-tocopherol may be small compared to the fortification levels, it can still have a pronounced effect on the stability of carotenoids[36].

      The present study utilized carrier lipids in a liquid state. However, when solid lipids were used as a carrier medium, the rate of β-carotene was higher in the case of triglycerides such as tripalmitate and tristearate and lower in the case of monoglycerides. It is possible that co-crystallization of β-carotene and monoglycerides was favored while this combination was utilized. However, when solid triglycerides were used, the accelerated degradation suggested that β-carotene was not incorporated into the crystal structure of tristearete or tripalmitate[50]. The exclusion of β-carotene from the solidified lipid fraction was also recorded by Pan et al.[53] and hence liquid lipid carriers served as better carriers during β-carotene encapsulation. Therefore, the use of solid lipids for β-carotene loaded encapsulated system should be considered with caution and take precedence over the emulsified system only if β-carotene gets incorporated into the crystal lattice. Further research is needed to carefully study the incorporation of β-carotene within solid lipid carriers.

    • The effect of multilayered encapsulation structure around β-carotene loaded oil phase against β-carotene dispersed in bulk oil (encapsulation absent) within polymeric pouches during storage at 37 °C in dark and lighted conditions is illustrated in Fig. 5. β-carotene in bulk oil and emulsified form are experimented simultaneously to determine the level of protection encapsulation would offer β-carotene against oxidation. Emulsions offered protection against β-carotene degradation under storage. In our studies, the samples were stored at 37 °C to reflect accelerated storage. After 5 d of storage in the dark, β-carotene in GTO and MCT bulk oils showed rapid degradation recording a loss of 70% and 66%, respectively. When protected within bilayer emulsions under similar conditions of storage and time, β-carotene loss was negligible. Extending the assessment period up to 15 d of storage, the encapsulants offer significantly better protection to β-carotene against bulk oils. This protection wears off after 20 d of storage at 37 °C.

      The above results are in agreement with Li et al.[33] & Liang et al.[21]. Liang et al.[21] used OSA-modified starches of various molecular weights to prepare emulsion and compared it against bulk MCT oil. The carrier oil for emulsion synthesis was MCT and so was the bulk control, both loaded with β-carotene. Their results indicated that β-carotene dispersed in bulk MCT oil stored at 25 °C degraded completely in 7–10 d. The β-carotene loaded emulsions survived better with a retention of 51%–64%. Similar observations were made by Yi et al.[37] & Li et al.[33] where emulsions stabilized by WPI and chitosan hydrochloride/carboxymethyl starch complex offered significantly enhanced preservation than their respective bulk vegetable oils. A high internal phase emulsion of WPI nanoparticles was found to offer better protection to the internal oil phase consisting of β-carotene in corn oil against bulk corn oil control. Additional antioxidant effect from the protein WPI, which has the potential to scavenge free radicals and chelate metal pro-oxidants is also a possible reason for the enhanced stability of the emulsified system[37].

      Therefore, the emulsion interface with tight packing of multilayered OSA starch and chitosan layers could have shielded from degrading free radicals and slowed the diffusion of pro-oxidants towards the core of bioactive β-carotene. The barrier effect of emulsified structure enhanced the retention of β-carotene during storage when compared to bulk oils despite both the system packaged within polymeric pouches with sufficient barrier properties of its own.

      β-carotene loaded MCT and GTO bulk oils within polymeric pouches degraded more on exposure to light (Fig. 5a). Within 5 d of light exposure at 37 °C, a 74% loss in β-carotene was measured for bulk oils. On the other hand, β-carotene within emulsified oils under the same storage condition for 5 d measured significantly higher values. Furthermore, the β-carotene retention is higher in emulsions than their respective bulk oils up to 15 d at 37 °C after which, both the curves–bulk oils and emulsions, began to merge. Comparing the degradation of β-carotene due to radiation exposure from similar experiments from literature, emulsions were found to retain the bioactive over longer periods than bulk oils. For instance, a significantly higher retention of β-carotene was observed in genipin cross-linked chitosan emulsions in an 8 h UV exposure study assessing retention of β-carotene in bulk and emulsified dodecane[31]. Similarly, encapsulated β-carotene was retained better in a 7 h UV exposure study evaluating the preservation of β-carotene in bulk and emulsified corn oil systems[33]. Therefore, the present results are consistent with similar studies.

      Another interesting observation revolving around the storage stability of β-carotene dispersed in MCT bulk oil was made by Liang et al.[21]. They assessed β-carotene retention with and without nitrogen flushing at 4 °C and observed that the presence of nitrogen gas within glass vials significantly slowed down β-carotene degradation in bulk MCT oil. On the other hand, nitrogen flushing in β-carotene emulsions produced no such improvements. It is possible that shielding from oxygen was achieved by encapsulated structures, and therefore nitrogen flushing made no difference in emulsions. β-carotene in bulk MCT oil was offered lesser exposure to oxygen when nitrogen was flushed. Therefore, encapsulated structures could provide a protective effect against degradation factors such as oxygen and free radicals.

      There have been reports of different kinds of isomerization and degradation reactions when β-carotene in a food environment is subjected to heat, light, and storage[11]. Several pathways for isomerization and degradation of β-carotene are possible in experimental scenarios, including auto-oxidation, photo-oxidation, and thermal degradation. Over 20 oxidation products were observed for β-carotene's reaction to oxygen gas by auto-oxidation. β-carotene auto-oxidation was observed to be decelerated by the presence of radical scavengers such as BHT and α-tocopherol and accelerated by the presence of a free radical reaction initiator. These observations indicate that free radicals are involved in β-carotene auto-oxidation[54]. β-carotene under conditions of elevated temperatures undergo isomerization followed by degradation. Trans-cis isomerization of β-carotene diminishes its biological activity with cis isomers diminishing the bioavailability as vitamin A precursor, as well as antioxidant properties[55].

      The range of TBARS obtained for 25 d for GTO oil-in-water emulsions and MCT oil-in-water emulsions stored under dark and light conditions and bulk oils stored under similar conditions at 37 °C are shown in Fig. 7. Comparing the emulsions and oils, we notice that the range of TBARS in the studied period was higher for emulsions (2–7 mM) than for oils (1–3 mM). Sharif et al.[34] produced a similar TBARS chart to the present study with regard to TBARS between emulsions and oils. The range of TBARS obtained for emulsions was higher (up to 3.5 mM) than for bulk oil of flax seed oil (up to 0.8 mM) where the secondary products of oxidation were recorded for 4 weeks. Tan et al.[56] reported that lipid oxidation could proceed faster in emulsions as compared to bulk oil. Emulsified oils being in a diphasic system have a large oil-water interfacial area that exposes more of the oil surface to aqueous phase pro-oxidative factors. However, measurement of β-carotene in the previous sections has shown that the bioactive is better retained in encapsulated systems.

      Figure 7. 

      Secondary oxidation products TBARS measured for (a) emulsions and, (b) their respective bulk oils stored in light and dark storage at 37 °C, n = 4.

    • The degradation of β-carotene dissolved in oil and that dispersed in emulsified form was monitored in this study for a 25-d period with and without light exposure. The kinetic plots for zero, first and second order with corresponding R2 are presented in Tables 2, 3 & 4. From these tables, it can be seen that the best fit for β-carotene dissolved in bulk GTO and MCT oils is given by second-order kinetics. On the other hand, the best fit for β-carotene degradation dispersed emulsions is given by zero-order kinetics. The present results for dispersed systems are similar to those observed by Cornacchia & Roos[57] who observed β-carotene to exhibit zero-order kinetics in solid lipid particles and liquid lipid particles stabilized by whey protein isolate at 15 °C for 28 d.

      Table 2.  Zeroth order rate constants of β-carotene dissolved in oil and dispersed in emulsified form to storage in dark vs photodegradation.

      Dark Light
      R2 k (μg mL−1 day−1) R2 k (μg mL−1 day−1)
      Oil MCT 0.726 9.13 0.632 9.13
      GTO 0.675 8.18 0.648 8.47
      Emulsions MCT 0.925 13.1 0.930 14.0
      GTO 0.950 13.0 0.978 13.1
      All samples were stored at an accelerated storage of 37 °C.

      Table 3.  First order rate constants of β-carotene dissolved in oil and dispersed in emulsified form subjected to storage in dark vs photodegradation.


      Dark Light
      R2 k (day−1) R2 k (day−1)
      Oil MCT 0.833 0.106 0.916 0.109
      GTO 0.870 0.093 0.906 0.112
      Emulsions MCT 0.746 0.119 0.943 0.152
      GTO 0.800 0.102 0.904 0.143
      All samples were stored at an accelerated storage of 37 °C.

      Table 4.  Second order rate constants of β-carotene dissolved in oil and dispersed in emulsified form subjected to storage in dark vs photodegradation.

      Dark Light
      R2 k (mL μg−1 day−1) R2 k (mL μg−1 day−1)
      Oil MCT 0.933 0.0006 0.979 0.0014
      GTO 0.930 0.0008 0.985 0.0013
      Emulsions MCT 0.699 0.0004 0.738 0.0018
      GTO 0.768 0.0004 0.622 0.0018
      All samples were stored at an accelerated storage of 37 °C.

      The majority of the storage studies that measured β-carotene over storage periods in food systems or encapsulated systems reported first order kinetics[51,5860]. When the present results for bulk oils were fit to the first-order kinetic model, the fit was good but not the best. In the present trials with sunflower oil dissolved with β-carotene and stored in the dark at 37 °C, no specific trend was observed for bulk oil (Supplemental Fig. S3). Calligaris et al.[51] described degradation of β-carotene in bulk MCT oil kept under a storage condition of 20 °C for 14 d as pseudo-first order kinetics, after which β-carotene was not detected.

      The present study attempted to eliminate the effects of antioxidants by selecting saturated medium-chain triglycerides, it is possible that the effect of fatty acid chain length of carrier oil has some part to play in encapsulated systems. Further studies that eliminate residual antioxidants as well as use oils with long-chain fatty acids as carriers may prove to further understand storage stability of β-carotene in encapsulated emulsions.

    • Multilayered emulsions of OSA-starch and chitosan were prepared by layer-by-layer deposition method. The polysaccharide based multilayered emulsions were found to be physically stable in heat treatments up to 121 °C for 60 min. β-carotene enclosed within emulsions were retained better than β-carotene dissolved in bulk oil during storage at 37 °C. Emulsions functioned as encapsulating structures of the oil phase and offered protection to the bioactive material β-carotene enclosed. The extent of the protection depended largely on the nature of the interfacial material and the presence of additional antioxidants in the system. Emulsified and oil-based systems composed of β-carotene were only slightly affected by exposure to light. However, perceived loss in color was more prominent in emulsions stored under light than in the dark. The reaction kinetics of emulsified β-carotene best fit the zeroth-order model and oil solubilized β-carotene best fit the second-order kinetic model.

    • The authors confirm contribution to the paper as follows: conceptualization: Sivabalan S, Sablani SS; methodology: Sivabalan S, Sablani SS; data curation: Sivabalan S, Sablani SS; formal analysis and visualization: Sivabalan S, Sablani SS, Ross C, Tang, J; writing-original draft: Sivabalan S; writing-review & editing: Sivabalan S, Sablani SS; Ross C, Tang J; funding acquisition and supervision: Sablani SS. All authors reviewed the results and approved the final version of the manuscript.

    • All data generated or analyzed during this study are included in this published article and its supplementary information files.

      • This research was partially supported by the (USDA) National Institute of Food and Agriculture Research (Grant Nos 2016-67017-24597 and 2016-68003-24840), and Hatch project (#1016366). The authors thank Dr. Hanu Pappu and Dr. Sindhuja Sankaran for the use of the Ultrasonicator and Luxmeter.

      • The authors declare that they have no conflict of interest.

      • Supplemental Fig. S1 lllustration of layer-by-layer electrostatic deposition for synthesis of secondary emulsion-2.5 % oil, OSA starch as primary emulsifier and chitosan as secondary emulsifier.
      • Supplemental Fig. S2 lllustration of storage study at 37 °C where selected β-carotene loaded secondary emulsion along with bulk oil was exposed to light.
      • Supplemental Fig. S3 Trend in β-carotene dissolved in bulk sunflower oil degradation during storage in dark, n ≥3.
      • Supplemental Table S1 ANOVA table for storage parameters: For storage conditions, each data set had 3 source of variation (factors) and therefore, a 3-way ANOVA analysis was used to study the data. As indicated in the table, in all the data set studied, interactions were significant (green highlights) and therefore, single factor main effects are not to be interpreted.
      • Copyright: © 2024 by the author(s). Published by Maximum Academic Press on behalf of China Agricultural University, Zhejiang University and Shenyang Agricultural University. This article is an open access article distributed under Creative Commons Attribution License (CC BY 4.0), visit https://creativecommons.org/licenses/by/4.0/.
    Figure (7)  Table (4) References (60)
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    Sivabalan S, Ross CF, Tang J, Sablani SS. 2024. Physical, thermal, and storage stability of multilayered emulsion loaded with β-carotene. Food Innovation and Advances 3(3): 244−255 doi: 10.48130/fia-0024-0022
    Sivabalan S, Ross CF, Tang J, Sablani SS. 2024. Physical, thermal, and storage stability of multilayered emulsion loaded with β-carotene. Food Innovation and Advances 3(3): 244−255 doi: 10.48130/fia-0024-0022

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