catalysts Review Recent Advances in Lipases and Their Applications in the Food and Nutraceutical Industry Ana Laura Reyes-Reyes 1,† , Francisco Valero Barranco 2 and Georgina Sandoval 1, * 1 LIBBA Lab, Industrial Biotechnology Unit, Center for Research and Assistance in Technology and Design of the State of Jalisco, A.C. (CIATEJ), Guadalajara 44270, Jalisco, Mexico 2 Department of Chemical, Biological and Environmental Engineering, School of Engineering, Universitat Autònoma de Barcelona, Bellaterra, 08193 Barcelona, Spain * Correspondence: gsandoval@ciatej.mx or gsandoval@confluencia.net † Present Address: National Institute of Forestry, Agricultural and Livestock Research (INIFAP), Tuxtla Chico 30870, Chiapas, Mexico; reyes.ana@inifap.gob.mx. Abstract: Lipases are efficient enzymes with promising applications in the nutraceutical and food industry, as they can offer high yields, pure products under achievable reaction conditions, and are an environmentally friendly option. This review addresses the production of high-value-added compounds such as fatty acid esters, with the potential to be used as flavoring agents or antioxidant and antimicrobial agents, as well as structured lipids that offer specific functional properties that do not exist in nature, with important applications in different food products, and pharmaceuticals. In addition, the most recent successful cases of reactions with lipases to produce modified compounds for food and nutraceuticals are reported. Keywords: lipases; high-value compounds; antioxidant; flavors; structured lipids; food; nutraceutical industry Citation: Reyes-Reyes, A.L.; Valero Barranco, F.; Sandoval, G. Recent Advances in Lipases and Their Applications in the Food and 1. Bioactives, Nutraceuticals, and Functional Foods Update Nutraceutical Industry. Catalysts In recent years, consumers have been increasingly interested in so-called functional 2022, 12, 960. https://doi.org/ foods and nutraceuticals. The concept of functional food was first introduced in Japan and 10.3390/catal12090960 is claimed to promote health and well-being beyond its nutritive properties [1]. Similarly, Academic Editor: Andres R. “bioactives” are nutritive substances that have a favorable impact on human health [1,2]. Alcantara The bioactive compounds that are extracted from the original food and maintain their beneficial properties for health are called nutraceuticals [2]. In the nutraceutical market, Received: 29 July 2022 lipases represent a great tool, and their economic cost is relevant because with added use in Accepted: 22 August 2022 2020, they were valued at USD 585.56 million. It is expected that by 2028 it will reach USD Published: 29 August 2022 961.85 million, at a compound annual growth rate (CAGR) of 6.4% from 2021 to 2028 [3]. Publisher’s Note: MDPI stays neutral The definitions of “functional foods” vary from place to place. In the USA, the Institute with regard to jurisdictional claims in of Food Technologists [4] defined functional foods as foods and food components that published maps and institutional affil- provide a health benefit beyond basic nutrition for the intended population [5]. In Japan, the iations. term “Food with Health Claims” (FHC) is used instead and refers to foods that comply with the specifications and standards established by the Ministry of Health, Labor, and Welfare and are labeled with certain nutritional or health functions. In Europe, the European Commission Concerted Action on Functional Food Science in Europe (FUFOSE) published Copyright: © 2022 by the authors. Licensee MDPI, Basel, Switzerland. a consensus concept that states that a food is “functional” if it is satisfactorily demonstrated This article is an open access article to beneficially affect one or more target functions in the body, beyond adequate nutritional distributed under the terms and effects, in a way that is relevant to either an improved state of health, well-being or conditions of the Creative Commons reduction of disease risk [6]. Subsequently, the European Food Safety Authority (EFSA) Attribution (CC BY) license (https:// adopted regulations about health claims that state, suggest, or imply that a relationship creativecommons.org/licenses/by/ exists between a food category, a food, or one of its constituents [7]. 4.0/). Catalysts 2022, 12, 960. https://doi.org/10.3390/catal12090960 https://www.mdpi.com/journal/catalysts Safety Authority (EFSA) adopted regulations about health claims that state, suggest, or im- ply that a relationship exists between a food category, a food, or one of its constituents [7]. The current awareness of the population toward the consumption of functional foods Catalysts 2022, 12, 960 and nutraceutical products demands that they provide not only nutrition but2 also of 24 func- tional properties and benefits to health. 2. Lipases as Biocatalysts The current awarenessin ofthe Food and Nutraceutical the population Industryof functional foods toward the consumption andIn nutraceutical the searchproducts demands that for alternatives they provide to improve the not only nutrition production but also of food andfunctional nutraceutical properties and benefits to health. supplements, including omics, biotechnology has provided tools to achieve and cover the requirements 2. Lipases as demanded Biocatalysts by legislation in the Food andand consumersIndustry Nutraceutical [8]. One way to produce and im- prove such compounds is by using biocatalysts. Lipases are widely used in the food in- In the search for alternatives to improve the production of food and nutraceutical dustry [9,10]. Lipases supplements, (triacylglycerol including hydrolases omics, biotechnology EC 3.1. 1.3) has provided play tools to aachieve crucialand rolecover in numer- ous industrial food processes [11,12] because they participate in the requirements demanded by legislation and consumers [8]. One way to producereactions that improve product qualitysuch and improve andcompounds provide greater stability, is by using solubility, biocatalysts. durability, Lipases and better are widely organolep- used in the ticfood characteristics [10,13,14]. industry [9,10]. Lipases (triacylglycerol hydrolases EC 3.1. 1.3) play a crucial role in numerous industrial food processes [11,12] because they participate in reactions that improve 2.1. product quality and provide greater stability, solubility, durability, and better Lipase Characteristics organoleptic characteristics [10,13,14]. These enzymes can hydrolyze triglycerides to obtain free fatty acids, monoacylglyc- erols 2.1. (MAGs), diacylglycerols (DAGs), and glycerol; on the other hand, they can synthe- Lipase Characteristics size new products These enzymes incan organic media hydrolyze by esterification, triglycerides to obtaintransesterification, and aminolysis free fatty acids, monoacylglyc- mechanisms erols (MAGs), (Figure 1) [15,16]. diacylglycerols Lipases (DAGs), and have a highly glycerol; conserved on the other hand, catalytic they cantriad comprising synthesize serine as a nucleophile, new products in organican aspartate/glutamate media as an acidic residue, by esterification, transesterification, and histidine. and aminolysis mech-In their anisms (Figure 1) [15,16]. Lipases have a highly conserved catalytic triad active conformation, lipases present in their active center a group of hydrophobic residues comprising serine asaround arranged a nucleophile, an aspartate/glutamate the catalytic serine that constituteas an an acidic residue, and electrophilic histidine. region knownIn as an their active conformation, lipases present in their active center a group of hydrophobic oxyanion cavity. Lipases are also characterized by the presence of disulfide bridges that residues arranged around the catalytic serine that constitute an electrophilic region known give them stability and are critical for their catalytic activity [16]. Some lipases also have as an oxyanion cavity. Lipases are also characterized by the presence of disulfide bridges a structural that give themfeature covering stability thecritical and are active for site,their called the “lid,” catalytic that[16]. activity opensSome at hydrophobic/hy- lipases also drophilic interphases. Ancient classifications denoted esterases as lipolytic have a structural feature covering the active site, called the “lid,” that opens at hydropho- enzymes lack- ing a lid. However, bic/hydrophilic because Ancient interphases. some lipases, such asdenoted classifications Candida antarctica esterases lipase Benzymes as lipolytic (CALB), lack the lid, an alternative classification has been proposed [17]. lacking a lid. However, because some lipases, such as Candida antarctica lipase B (CALB), lack the lid, an alternative classification has been proposed [17]. Figure 1. Different reactions are catalyzed by lipases [15]. Figure 1. Different reactions are catalyzed by lipases [15]. Catalysts 2022, 12, 960 3 of 24 Catalysts 2022, 12, x FOR PEER REVIEW 4 of 26 Lipases are characterized by maintaining their activity and high production in non- Meyerozyma guilliermondii aqueous Wild mediatype [18], high production, Feedand stability at pH ranges and do[47] industry not require cofac- tors. According to their substrate specificity, lipases can be chemoselective, regioselective, orOther stereoselective. importantThe first lipase bottlenecks of type can enzymes the free selectivelyincatalyze general aand reaction. lipasesThe aresecond the lowtype catalyzes a reaction specifically with one of the triglyceride positions (sn-1,3 regioselective, operational stability in synthesis reactions using solvents and substrates such as alcohols andsn-2 regioselective, organic acids, theor nonregioselective). high Additionally, cost of the enzymes, and the the needthird type catalyzes to reuse reactions the biocatalyst selecting product minimizing only oneseparation. of the stereoisomers from a mixture of enantiomers [10,16]. Different approaches are being applied (Figure 2) to solve these drawbacks. The use of 2.2. Sources and Tools to Improve Lipase-Catalyzed Reactions enzyme immobilization methods normally increases biocatalyst stability, specificity and se- lectivity,Lipases are reutilization allows the ubiquitous enzymes produced of the enzyme, and by various downstream minimizes organisms, including processes,microor- and has been reflected in the number of articles and patents published in this field [48]. interest in ganisms, plants, and animals [12,19–24]. Because of the increased commercial these proteins Advances in in thethe foodof study and nutraceutical lipases industry,more seek to develop the use of recombinant efficient processes production and, for technology is critical. this purpose, their stability under certain temperatures, solvents, and pH conditions, The productivity of lipase production bioprocesses has been increasing, reducing the among others. The development of a specific reaction medium to increase the activity, cost of enzymes by using cell factories for the heterologous production of lipases. Between stability, and productivity of biocatalysts has been a recurring topic of research over the them, Komogataella phaffi (P. pastoris) is one of the most common cell factories used [25]. last three decades. The remarkable properties and useful applications of enzymes, partic- Lipases have been improved using natural evolution techniques, protein engineering, ularly lipases, have inspired various strategies to improve their performance in near-an- bioinformatics design, directed evolution, saturation mutagenesis, site-directed mutage- hydrous media. Therefore, medium engineering can be used to modulate the activity and nesis, and DNA shuffling [26]. However, in the food industry, the native form is often selectivity of lipase-catalyzed reactions [49]. preferred (Figure 2). Figure 2. Sources and use of lipases in the food industry. Figure 2. Sources and use of lipases in the food industry. Table 1 shows some microbial lipases that are commercially available and immobilized onIonic liquids different (ILs) aretomolten supports enhance salts thatefficiency their originate and fromreuse the association [27–30]. Mostof organic cati- commercially onsimportant and organic/inorganic anions. The use of ILs as solvents in biocatalysis lipase-producing yeasts belong to the class of ascomycetes, such as Candidaprocesses has sp. recently received increased attention, ® and substantial progress has ® and Rhizopus sp. Novozymes (Bagsværd, Denmark), DuPont (Wilmington, DE, USA), been made, particu- larly in lipase-catalyzed Roche reactions. ® (Basel, Switzerland), andILs have the Amano advantages (Yokohama, of low Japan) are volatility, low inflam-that the main companies mability, produce and a low and melting point commercialize [50]. [31]. lipases Deep eutectic solvents [51] are eutectic mixtures of salts and Other important bottlenecks ofsufficiently hydrogen bond donors with low melting the free enzymes points in general andto lipases act as solvents. are the low DESs were demonstrated to be a viable alternative to traditional organic operational stability in synthesis reactions using solvents and substrates such as solvents and ILs alcohols in many biocatalytic processes, particularly for lipases. DESs have additional and organic acids, the high cost of the enzymes, and the need to reuse the biocatalyst advantages over ILs in simple minimizing preparation product separation.and lower costs because of their renewable and readily available raw materials [52]. 3. Established Applications of Lipases in the Food and Nutraceutical Industry Lipases in the food industry and nutraceutical production can be used in aqueous extracts and purified, immobilized, or whole cells to exploit the available raw material and increase their economic and nutritional value. These enzymes can be used to modify Catalysts 2022, 12, 960 4 of 24 Table 1. Sources of lipases with applications in food and nutraceutical industry. Source/Commercial Name Type Application/Products Reference Candida antarctica lipase B Recombinant Flavor esters [32] (CALB)/Novozym 435/Lipozyme 435 Candida rugosa Wild type Glycerides, production flavor compounds [33,34] Termomyces lanuginosus/Lipozyme TL IM Engineered Food formulation, Interesterification of fats and oils [35,36] Aspergillus sp. Wild type Flavor and fragance [37] Aspergillus oryzae Wild type Interesterification of fats and oils [36] Geotrichum candidum Wild type Oil with increased unsaturation [36] Enhancing fruit fragrance [38] Modification of the amount and composition of Rhizomucor miehei/Lipozyme RM IM Recombinant [39] volatile components in bovine milk Ras Cheese Flavor Concentrate (RCFC) [40] Rhizopus oryzae Wild type Human Milk Fat Substitutes [41] Lactococcus chungangensis Wild type Flavoring in milk, cream cheese, yogurt and butter. [42] Lactobacillus plantarum Wild type Fermented food and cheese [43,44] Staphylococcus epidermidis Wild type Flavor-compound production [45] Ophiostoma piceae Wild type Flavor-compound production [46] Meyerozyma guilliermondii Wild type Feed industry [47] Different approaches are being applied (Figure 2) to solve these drawbacks. The use of enzyme immobilization methods normally increases biocatalyst stability, specificity and selectivity, allows the reutilization of the enzyme, and minimizes downstream processes, and has been reflected in the number of articles and patents published in this field [48]. Advances in the study of lipases seek to develop more efficient processes and, for this purpose, their stability under certain temperatures, solvents, and pH conditions, among others. The development of a specific reaction medium to increase the activity, stability, and productivity of biocatalysts has been a recurring topic of research over the last three decades. The remarkable properties and useful applications of enzymes, particularly lipases, have inspired various strategies to improve their performance in near-anhydrous media. Therefore, medium engineering can be used to modulate the activity and selectivity of lipase-catalyzed reactions [49]. Ionic liquids (ILs) are molten salts that originate from the association of organic cations and organic/inorganic anions. The use of ILs as solvents in biocatalysis processes has recently received increased attention, and substantial progress has been made, particularly in lipase-catalyzed reactions. ILs have the advantages of low volatility, low inflammability, and a low melting point [50]. Deep eutectic solvents [51] are eutectic mixtures of salts and hydrogen bond donors with sufficiently low melting points to act as solvents. DESs were demonstrated to be a viable alternative to traditional organic solvents and ILs in many biocatalytic processes, particularly for lipases. DESs have additional advantages over ILs in simple preparation and lower costs because of their renewable and readily available raw materials [52]. 3. Established Applications of Lipases in the Food and Nutraceutical Industry Lipases in the food industry and nutraceutical production can be used in aqueous extracts and purified, immobilized, or whole cells to exploit the available raw material and increase their economic and nutritional value. These enzymes can be used to modify fats and oils and synthesize structured lipids or antioxidants with increased antioxidant power or modified lipophilicity, flavors, and aromas [53,54]. Catalysts 2022, 12, 960 5 of 24 3.1. Fats and Oils Patent searches suggest that lipase has an impressive number of applications in the modification of fats and oils and enhancement of flavor in food products—e.g., cheese, butter, milk, and chocolate [55]. Some applications of lipases in dairy products and the synthesis of structured lipids are described in the following sections. 3.1.1. Dairy Products In the dairy sector, lipases are used to provide desirable aromatic characteristics to cheddar, provolone, and Romano cheeses conferred by these free short-chain fatty acids generated in the hydrolysis of fats [40,56]. Recent advances have allowed the biosynthesis of short-chain ethyl esters with fruity notes in whole milk by coupling ethanolic fermenta- tion with transesterification using the commercial lipase Palatase. For fermentation, the following microorganisms were used: Kluyveromyces marxianus, Lactobacillus fermentum, and L. Paracasei. Many esters were obtained in ethanolic fermentation using K. marxianus yeast and lipase. This method of milk fermentation and lipase addition represents a new alternative for flavoring milk [57]. 3.1.2. Structured Lipids In recent years, structured lipids have become a topic of great importance in the food and nutraceutical industry because technological advances allow a generation of products of better quality and that better meet consumer demands. Within this innovation in food processes, structured lipids (SLs) have been generated [58,59]. Structured lipids are fats and oils whose fatty acid composition has been modified for nutritional purposes to achieve greater bioavailability because they are not naturally occurring. In several cases, lipids have certain limitations of use in their original state because of the specific composition of their fatty acids [60]. In other cases, even when they are available as raw materials, they cannot always meet nutritional demand, e.g., restrictions on the daily intake of saturated fatty acids and trans fatty acids have been increased because they are related to cardiovascular diseases [61,62]. Another clear example would be access to cocoa butter; its availability may be limited by external factors such as climate change, fluctuating prices, and availability [63,64]. Therefore, the search for alternatives to address these major issues is justified. In principle, deciding which type of fatty acids to use and in which position of the molecule to restructure is possible by obtaining structured lipids [58]. For this procedure, the use of stereospecific enzymes allows new lipids with a stable structure to be obtained. Lipases can hydrolyze a triglyceride in an aqueous medium, but they also catalyze the bind- ing of a fatty acid to a glycerol molecule in an anhydrous reaction medium [65]. Recently, the use of immobilized biocatalysts has minimized the production costs of structured lipids through reusing them in successive batches [58]. With all the knowledge generated on the subject, we can generate these products for nutritional, pharmacological, or industrial use, such as breast milk substitutes, cocoa butter, and low-calorie or enriched triacylglycerols [41,58,66,67]. In the case of low-calorie triacylglycerols, the energy equivalents are reduced. Typi- cally, these compounds are used to control poor fatty acid absorption and other metabolic problems. They are characterized based on triglycerides (TAGs) of this nature containing short or medium-chain fatty acids in the sn-1,3 positions; in the sn-2 position, they have a long-chain fatty acid esterified; therefore, the absorption of external fatty acids is released and metabolized more rapidly [68]. They can be synthesized by acidolysis of a TAG or oil containing long-chain fatty acids with one of the medium-chain fatty acids or by interester- ification of a TAG or oil with methyl or ethyl esters. In the lipase-catalyzed synthesis of SL, ethyl esters are preferred as acyl donors because they avoid the presence of high amounts of free fatty acids (FFAs) in the reaction medium; in food production, methanol poses toxicity risks [69,70]. In this context, the production of low-calorie triacylglycerols was achieved using as raw material cheap oils extracted from agro-food residues, such as spent coffee Catalysts 2022, 12, 960 6 of 24 grounds and olive pomace, and as catalysts, the regioselective lipase sn-1,3 from R. oryzae, which was immobilized on magnetic nanoparticles. This enzyme was shown to be a highly promising biocatalyst to produce structured lipids for both oils, presenting even higher activity than the commercial immobilized T. lanuginosus lipase (Lipozyme TL IM). For the two oils evaluated, a preference for acidolysis was observed, in addition to high stability when reused in acidolysis and interesterification of olive pomace oil [67]. However, SLs are an option to generate products that partially or totally substitute human milk in certain cases because, in terms of the fatty acid composition and distribution, they are manufactured to improve fat and mineral absorption, promote softer stools, and reduce constipation in infants [58,71,72]. Human milk fat substitute (HMFS) is synthesized by enzymatic interesterification of vegetable oils, animal fats, or oil mixtures, commonly using an immobilized regioselective lipase in either solvent or solvent-free media [70,73]. A recently reported lipase/acyltransferase from C. parapsilosis was used as a biocatalyst to synthesize HMFS by interesterification of ethyl oleate with tripalmitin in solvent-free media representing a new alternative to commercial immobilized lipases [70]. Because human milk is one of the most complex mixtures of natural lipids, studies using this approach will continue to advance steadily. 3.2. Vitamin Esters Food contains components known as bioactive compounds that, when consumed, provide energy to the body, promote good health and minimize the risk of disease. The bioactive compounds that are extracted from the original food and maintain their bene- ficial properties for health are called nutraceuticals [2]. For consumption and consumer acceptance, the functionality of bioactive compounds, safety, and nontoxicity must be guaranteed beforehand [74]. Highlighting a representative example, antioxidants play a crucial role in the food industry because, during food processing, the matrices used mostly incorporate lipids as emulsifiers or additives, making lipid oxidation a challenge to consider [75–78]. Lipid oxidation involves the attack of molecular oxygen on unsaturated fat molecules, which can generate undesirable volatile flavoring compounds that contribute to rancidity [79]. Even when quality controls are followed during food product preparation and packaging, the rate of lipid oxidation is influenced by several endogenous and exoge- nous parameters, including oxygen, light metals, and polyunsaturated lipids, primarily because the latter are prone to oxidation [80,81]. Antioxidants are used to mitigate this effect, meaning molecules that reduce, neutralize, or deplete molecular oxygen, remove pro-oxidative metal ions, and scavenge reactive oxygen species (ROS), hydrogen peroxide or superoxide anion radicals [82–85]. Antioxidants occur naturally, and the best known are ascorbic acid (vitamin C), to- copherols (vitamin E), carotenoids, and thiols [86]. During their absorption in the body, they complement the defense action as they help to slow down the use of endogenous antioxidants and improve the body’s ability to avoid oxidative stress [82,87–90]. The lack of action of endogenous antioxidants, either by diminution or stress, is related to the modifi- cation of lipid membrane components [91], resulting in neurodegenerative, cardiovascular, inflammatory diseases, diabetes, male infertility, and cancers of the breast, lung, liver, colon, prostate, ovary and brain [87,92–97]. The excessive presence of reactive oxygen species promotes the expression of oncogenic genes [93]. Antioxidants, as nutraceuticals, play a key role in the nutritional base because of their close relationship with biological pro- cesses; thus, skin benefits are also attributed to them for delaying aging [98–100]. Although the concept of nutraceuticals is not new, the trend to use antioxidants with biochemi- cal properties of high stability and biocompatibility as a complementary ingredient has become interesting [84,101,102]. In this context, the synthesis of vitamin derivatives obtained with biocatalysts is a tool that seems to minimize the technical problems of chemical synthesis or extraction from natural sources and is an ecological and environmentally friendly alternative [74,103–105]. 3.2.1. Retinol (Vitamin A) Esters Retinol is a vitamin A derivative found in foods (fish, dairy products, and meats). This molecule has attracted increased attention in the food and cosmetic industry for all Catalysts 2022, 12, 960 the benefits of its consumption because it controls mitochondrial energy homeostasis 7 of 24 by functioning as an electron carrier, maintains visual health, minimizes skin aging, pro- motes bone growth, and strengthens the immune system; thus, vitamin A plays a crucial role in 3.2.1. Retinol the health of(Vitamin A) Esters the organism [106,107]. This process involves a series of several steps, which generateRetinol is a vitamin various A derivative byproducts; found in foods thus, achieving good(fish, dairy quality products, of the and meats). final product may This molecule has attracted increased attention in the food and cosmetic require extensive time, derive waste pollutants to the environment and make the disposal industry for all the benefits of its consumption because it controls mitochondrial energy homeostasis by of chemical catalysts difficult [108]. This process involves a series of several steps, which functioning as an electron carrier, maintains visual health, minimizes skin aging, promotes generate various byproducts; thus, achieving good quality of the final product may re- bone growth, and strengthens the immune system; thus, vitamin A plays a crucial role quire too much in the time, health deposit of the too many organism waste [106,107]. pollutants This in the environment, process involves and steps, a series of several create difficult disposal which of the generate chemical various catalysts byproducts; [109,110]. thus, achieving Recently, immobilized good quality of the finallipase product(Novo- may zym 435) was successfully employed as a biocatalyst to generate a retinol require extensive time, derive waste pollutants to the environment and make the disposalderivative (ret- inol laurate). The process of chemical catalysts was previously difficult optimized [108]. This using biochemical process involves process a series of several modeling steps, which generate various and prediction byproducts; tools (artificial thus, achieving neural network, goodANN) quality and of the final product ultrasonic systems. may require The reac- too much time, deposit too many waste pollutants in the environment, tion, compared with the traditional method, takes less time, and the final product shows and create difficult higher disposal stabilityoftothe chemical catalysts [109,110]. Recently, immobilized lipase (Novozym 435) oxidation; thus, it could be used as an additive in human food supple- was successfully employed as a biocatalyst to generate a retinol derivative (retinol laurate). ments [110]. The process was previously optimized using biochemical process modeling and prediction tools (artificial neural network, ANN) and ultrasonic systems. The reaction, compared 3.2.2. Fatty with Acid Esters ofmethod, the traditional L-Ascorbic takes Acid (Vitamin less time, and theC)final product shows higher stability to oxidation; Ascorbic thus, acid it couldC) (vitamin beisused as an additiveantioxidant a water-soluble in human food notsupplements produced by[110]. the human body. Among its health benefits, some are derived from its antioxidant properties, such 3.2.2. Fatty Acid Esters of L-Ascorbic Acid (Vitamin C) as preventing male infertility and neurodegenerative diseases, in addition to counteract- Ascorbic acid (vitamin C) is a water-soluble antioxidant not produced by the human ing the effects of solar radiation, smoke, and environmental pollution [111]. The antioxi- body. Among its health benefits, some are derived from its antioxidant properties, such as dant property of ascorbic acid is due to its ability to donate individual hydrogen atoms preventing male infertility and neurodegenerative diseases, in addition to counteracting and subsequent formation the effects of of monodehydroascorbate solar radiation, smoke, and environmental (Figure 3), which pollution [111].reacts more rap- The antioxidant idly with property of ascorbic acid is due to its ability to donate individual hydrogen atomsdraw- radicals than with fully reduced or fully oxidized compounds [112]. The and back ofsubsequent degrading formation drives theofuse of strategies such as the monodehydroascorbate addition (Figure of fatty 3), which acids reacts to improve more rapidly with radicals their stability and than with fully reduced physiological or fully oxidized and antioxidant compounds activity [112]. The [113]. These drawback are derivatives of degrading drives the use of strategies such as the addition of fatty acids formed by the addition of a donor acyl to the primary alcohol to produce the correspond- to improve their stability andester ing 6-O-ascorbyl physiological and antioxidant without interfering withactivity [113]. These its antioxidant derivatives capacity, are formed catalyzed by li- by the addition of a donor acyl to the primary alcohol to produce the corresponding 6-O- pases, and generating compounds such as ascorbyl palmitate and ascorbyl oleate (Figure 3). ascorbyl ester without interfering with its antioxidant capacity, catalyzed by lipases, and generating compounds such as ascorbyl palmitate and ascorbyl oleate (Figure 3). Figure 3. Scheme of monodehydroascorbate, ascorbyl palmitate and oleate palmitate synthesis from Figure 3. Scheme of monodehydroascorbate, ascorbyl palmitate and oleate palmitate synthesis from vitamin C [114]. vitamin C [114]. Another method to improve the stability and physicochemical properties of vitamin C involve using emulsions and microchannel emulsification, which promote less interaction of the compound with the oxygen present in the medium and provide greater solubility in fats [115,116]. However, ascorbyl esters of unsaturated fatty acids possess equally good properties, such as miscibility in hydrophobic media [117]. Obtaining ascorbic acid derivatives was originally achieved by chemical processes, which have been migrating to the use of lipases because different solvents can be used in regioselective reactions, achieving high yields and easier product isolation. Lipases have been successfully employed to catalyze the synthesis of ascorbyl esters from saturated and unsaturated free fatty acids, alkyl and vinyl esters, TAGs, and oils as acyl donors. The Catalysts 2022, 12, 960 8 of 24 experimental conditions and yields mentioned in the literature to obtain some ascorbic acid derivatives are shown in Table 2. This type of reaction can also be exploited to synthesize chalcogen-containing vitamin C derivatives and use CALB to obtain esters containing differently substituted selenium, sulfur, and tellurium, which are of interest for their anticancer, antibacterial, and enzyme inhibitory capabilities [118]. Table 2. Summary of enzymatic synthesis of fatty acid esters of vitamin C. Vitamin Reaction Conversion Acyl Donor Solvent Biocatalyst Reference Derivative Conditions (%) AA:PA molar ratio Indigenously 1:5; 20 mL L-ascorbyl Palmitic acid tert-butyl alcohol immobilized lipase tert-butyl alcohol, 50 palmitate PyCal (CALB) 0.6 g of biocatalyst, 60 ◦ C (batch) [119] AA:PA molar ratio 1:5; 20 mL L-ascorbyl Palmitic acid tert-butyl alcohol Novozym 435 tert-butyl alcohol, 50 palmitate 0.6 g of biocatalyst, 60 ◦ C (batch) AA:PA molar ratio L-ascorbyl 2-Methyl-2-butanol Palmitic acid Novozym 435 1:8; 12 g/L 81 [114] palmitate (2M2B) biocatalyst; 55 ◦ C One of the widely used lipases for the acylation of L-ascorbic acid is Candida antarctica lipase B (Novozym 435) because of its good yield and conversion rate (Table 2). However, the price of immobilized has decreased more than chemical reagents because the processes of recombinant enzyme production and subsequent processing increase their prices. Dur- ing the modification of vitamin C, the most used reaction solvents are tertiary alcohols (e.g., 2-methyl-2-butanol and tert-butyl alcohol). 3.2.3. Tocopherols (Vitamin E) Esters The term vitamin E refers to a group of fat-soluble compounds that includes α, β, γ, and δ-tocopherol and α, β, γ, and δ-tocotrienol (Figure 4), among which α-tocopherol has higher antioxidant activity. In contrast, γ and δ-tocopherols and tocotrienols have higher cancer preventive but lower systemic properties [120]. Additionally, its effects include protecting against reactive oxygen species (ROS), reactive nitrogen species (RNS), and polyunsaturated fatty acid (PUFA) oxidation in the membrane, as well as modulating signal transduction and enhancing the immune response [121]. Furthermore, vitamin E can be combined with vitamin C to enhance the immune system, restore the antioxidant functions of vitamin E, and provide other benefits together [122–124]. Catalysts 2022, 12, x FOR PEER REVIEW 9 of 26 Catalysts 2022, 12, 960 9 of 24 FigureFigure 4. Examples 4. Examples of derivates of derivates of vitamin of vitamin E (α-tocopherol) E (α-tocopherol) catalyzedcatalyzed by lipases.byVE: lipases. VE:E;Vitamin E; EPA: Vitamin EPA: eicosapentaenoic eicosapentaenoic acid; DHA: acid; DHA:docosahexaenoic docosahexaenoicacid [125–127]. acid [125–127]. VitaminVitamin E is naturally E is naturally synthesized synthesized only by photosynthetic only by photosynthetic organisms, so organisms, it is mainlyso it is mainly extracted from plants, extracted from nuts, plants,andnuts, seeds;and therefore, seeds;totherefore, improve thetoamount improve andthe specific pro- and specific amount duction of α-tocopherol, production genetic modifications of α-tocopherol, have been made genetic modifications haveinbeen crops, suchin made ascrops, overex-such as overex- pression of the γ-TMT gene that allows the conversion of γ-tocopherol to α-tocopherol pression of the γ-TMT gene that allows the conversion of γ-tocopherol to α-tocopherol [128]. [128].However, However, onceoncevitamin vitamin E has E hasbeen extracted been from from extracted crops,crops, it is enriched using vege- it is enriched using vegetable table oil deodorizing distillate processes where lipases such as Lipozyme IM50 (immobi- oil deodorizing distillate processes where lipases such as Lipozyme IM50 (immobilized lized sn-1,3-specific lipase from Rhizomucor miehei) are used, with which enrichments close sn-1,3-specific lipase from Rhizomucor miehei) are used, with which enrichments close to to 50% are achieved [129]. Vitamin E derivatives are a more stable form than their precur- 50% sor and canare beachieved synthesized[129]. Vitamin chemically E derivatives [130] are a more or enzymatically, stable the latter beingform than their precursor performed by lipases (see Table 2). Thus, in commercial products, we can find vitamin E esters in theperformed by and can be synthesized chemically [130] or enzymatically, the latter being form lipases (seeETable of vitamin 2). vitamin acetate, Thus, inEcommercial products, succinate, vitamin we canvitamin E ferulate, find vitamin E esters in the form E eicosapen- of acid taenoic vitamin ester,Eand acetate, vitaminvitamin E succinate, acid E docosahexaenoic vitamin ester E ferulate, (Figure vitamin 4). The E eicosapentaenoic synthesis of theseacid ester, vitamin and vitamin E derivatives E docosahexaenoic is affected by factors suchacid ester as the (Figure enzyme used,4). The synthesis reaction me- of these dium,vitamin water activity, acyl donors, E derivatives and acylby is affected acceptors factorsemployed such as the[125,126,131]. enzyme used, reaction medium, water activity, acyl donors, and acyl acceptors employed [125,126,131]. Among the characteristics of α-tocopherol, it has three stereogenic centers at carbons 2, 4”- and 8”- and the RRR-α-tocopherol form is considered the most bioactive of the eight existing ones (RRR-, RSR-, RRS-, RSS-, SRR-, SSR-, SRS-, and SSS-). However, commercially, it is found as a mixture of all its stereoisomers in the esterified form (all-rac-α-tocopheryl acetate), obtained chemically from soybean byproducts [132,133]. For this reason, when Catalysts 2022, 12, 960 10 of 24 manufacturing food products with the different stereoisomers of this compound, separa- tion strategies such as the use of chromatographic methods should be included [134,135]. Currently, some foods, such as meat, are supplemented with vitamin E to improve shelf life, quality, and nutritional experience [136,137]. 3.3. Bakery Products New requirements in bakery products make the development of new formulations that conform to what would be green or less harmful labels. In bakery products, lipases have been successfully applied to improve dough processing, strength, volume, structure, and softness, decrease stickiness, and increase the quality and shelf life [138,139]. With a focus on the intermediate product of bread, dough plays an indispensable role in becoming the final product because it is a semisolid foam that is converted into a solid cellular sponge upon baking so that the mixture of the lipid fraction of wheat, eggs, or baker’s fat exerts major roles in gas incorporation and its stabilization, which are necessary to achieve a fluffy product [140]. Although wheat flour contains low levels of lipids, they affect the quality of fresh bread because they are related to storage duration. Briefly, the studies are directed toward knowledge of the relationship of the flours or their reformulation by adding lipids from other sources and their effect on quality. Recently, lipases have been successfully applied to investigate how endogenous or ex- ogenous lipids affect bread making. Lipases hydrolyze galactolipids, and their presence in the dough improves bread volume. The flour was defatted and subsequently reconstituted by adding different fractions of these lipids to determine the relationship of endogenous lipids in wheat flour and their impact on bread volume. The hydrolysis of endogenous lipids and their enzymatically released products are responsible for the positive effects on bread [138,141]. To understand the role of endogenous wheat lipids on the evolution of bread crumb firmness during storage, three lipases—Lipopan F, Lecitase Ultra, and Lipolase—were evaluated, and sodium lactylate stearoyl surfactant (SSL) was used as a surfactant. By forming amylose-lipid (AM-L) inclusion complexes, the surfactants retarded bread crumb firming. Some endogenous wheat lipids have surfactant-like structures, so the use of enzymes in bread making would increase the level of free fatty acids that allow the for- mation of amylose-lipid complexes. The evaluation of three enzymes showed that lipases and SSL similarly affected the texture of breadcrumbs during storage. However, after seven days of storage, the sample containing Lipolase significantly reduced amylopectin retrogradation, evidencing the importance of the formation of amylose-lipid inclusion complexes. Therefore, lipases have been proposed as alternatives to surfactants because they produce molecules in situ that possess hydrophilic and hydrophobic structures like those of surfactants [141,142]. 3.4. Flavors and Fragances In the world market, a high demand exists for fragrance and flavor esters for different industries, including food, cosmetics, and pharma, as ingredients of many products (food, beverages, candies, jellies, jams, wines, dairy products, perfumes, body lotions, shampoos, and other toiletries) [143,144]. The flavor and fragrance market was valued at $28 billion in 2019 and is expected to expand at a compound annual growth rate (CAGR) of 4.7% to $35 billion from 2021 to 2027 [145]. Another characteristic is that many of these products are chiral [146]. This potential chiral product can be consulted in the database [147]. Many of these products are obtained after extraction from their natural sources (plants, fruits, and flowers). However, the low concentration of these products in their natu- ral sources, climatic dependence of the source, and low yield and high production cost of the extraction and purification phases make it challenging to assume an increased world demand [143]. Catalysts 2022, 12, 960 11 of 24 A wide range of flavors and fragrances can be obtained by chemical synthesis, solving the of raw material producing the same products at a lower cost. However, these products have not been labeled as natural according to European legislation (EC 1334/2008), and obtaining pure chiral compounds is challenging. In this context, the substitution of a chemical using biotechnology (microbial biosynthesis or applied biocatalysis) is being widely explored because the products can be labeled as natural if the employed reactants are labeled as natural. The resolution of chiral compounds is generally higher with no problems in selectivity, reaching higher yields and with an easier downstream due to the absence of undesirable side reactions. However, the operational conditions (P, T) are softer than those of the chemical approach. A marketplace of bioflavors is actually 100–500 $/kg, and more than 100 flavor products are commercialized [148]. In 2019, the global biotech flavor market was close to 0.5 billion US$, approximately 1.5% of the estimated global market in the same year and is expected to grow at a compound annual growth rate (CAGR) of 9.3% from 2020 to 2027. Similarly, biotech vanillin represents ca. 3% of the total vanillin market, and it is speculated to increase at a CAGR exceeding 13% by 2023 [149]. Thus, the significant demand for these esters has boosted the need for greener produc- tion routes and food safety aspects for human consumption, making enzymatic synthesis a favorable alternative to chemical catalysts [150,151]. Approximately 4000 enzymes are known, and close to 200 have been mainly commercialized for stereoselective organic synthesis and the biotechnological production of flavor compounds [148]. Between them, lipases are the most applied enzyme family to produce flavor and fragrances. Although their natural biocatalysis is the hydrolysis of lipids to produce free fatty acids, glycerol, or other alcohols, they also work in reactions of esterification and trans- and interester- ification and the transfer of acyl groups from esters to other nucleophiles (e.g., amines and thiols) [143,152]. 3.4.1. Short-Chain Fatty Acids and Isoamyl Alcohol Esters Among the critical fragrance compounds produced via the esterification of short-chain alcohols and short-chain fatty acids, isoamyl alcohol esters, such as isoamyl butyrate and acetate, can be found. These esters serve as flavoring agents in numerous industries because of their characteristic fruity banana and intense banana flavor, respectively [153–156]. However, the use of short-chain fatty acids, being more hydrophilic, lowers the pH of the microenvironment and may lead to enzyme inactivation, while the use of short-chain alcohols tends to strip the essential water from the enzyme and serves as a dead-end inhibitor, making the enzymatic synthesis of esters challenging. Additionally, the use of isoamyl alcohol, which has a branched structure, exerts a higher steric hindrance on enzyme activity. Therefore, isoamyl alcohol might serve as an interesting model for understanding esterification with such acids and alcohols [157,158]. Isoamyl acetate (IAAC) has been produced in batch and continuous packed bed columns using porcine pancreatic lipase and Candida rugose lipase immobilized on chitosan and Ca-Ag chitosan. Operational conditions were optimized, and the amount of IAAC was close to ten times higher in a batch than in a continuous reactor [159]. IAAC was also obtained from the acylation of isoamyl alcohol with acetic anhydride by Candida antarctica B (CALB; Novozym 435) in ionic liquids in a continuously operated miniaturized enzymatic packed bed reactor. Up to a 92% isoamyl alcohol conversion with a volumetric productivity of 61 mmol L−1 min−1 was obtained. Interestingly, no decrease in productivity was observed 14 days after the operation [160]. Applying the same reaction and immobilized CALB from Sigma-Aldrich, the reaction was also made successfully in a miniaturized intensified reactor, obtaining a concentration of IAAC close to 1.2 mol L−1 [161]. In addition to using biocatalysis as an environmentally friendly approach for ester production, the use of byproducts as substrates for flavor ester production has emerged as a relevant way to support circular economy principles and reduce waste generation [162]. During large-scale bioethanol synthesis for fuel or food production, fusel oil is gen- erated, a byproduct obtained in the fermentation and distillation steps, and removed Catalysts 2022, 12, 960 12 of 24 during alcohol rectification. This byproduct accounts for approximately 0.25% by volume of bioethanol [163]. The quality and amount of generated fusel oil are affected by the pro- cessing parameters, such as the mash preparation, fermentation conditions, and distillation process. Higher alcohols (e.g., isoamyl alcohol, isobutanol, and butanol), water, aldehydes, and esters are the main compounds found in fusel oil samples [4,164]. Due to its intense odor, the use of fusel oil as a solvent is limited, although it has been used as a foam coating or added into diesel or gasoline to increase the cetane index and octane number. However, because of its high alcohol content (specifically isoamyl alcohol), it is attractive as a low-cost substrate for esterification and the production of various aromatic esters [164,165]. IAAC was produced from fusel oil in supercritical carbon dioxide (SC-CO2 ) using immobilized CALB (Lipozyme 435), demonstrating that acetic anhydride was a better acyl donor than ethyl acetate and acetic acid [21] in terms of IAAC conversion and specific pro- ductivity [4]. The same authors tested the same strategy using CALB (Novozyme 435) in a continuous packed bed reactor, obtaining the highest conversion at the lowest substrate rate. Additionally, Novozyme 435 maintained its stability and activity during the bioprocess [4]. IAAC was also obtained from the transesterification reaction of isoamyl alcohol and ethyl acetate using Aspergillus oryzae lipase obtained by fermentation and immobilized on sodium alginate with in situ ethanol removal. Under optimal conditions, conversion of IAAC of 89.55% and a yield ethanol extraction of 69.60% were obtained [166]. The production of isoamyl butyrate (IABU) from fusel oil and butyric acid in hexane using Lipozyme TL IM was also optimized, obtaining a conversion close to 96% in 24 h, a concentration of IABU of 1.64 mol L−1 , and a productivity of 0.19 mmol ester g−1 mixture h−1 [164]. Another example of the production of IABU from fusel oil and butyric acid using cyclohexane as a solvent was described using a covalently immobilized heterologous Rhizo- pus oryzae lipase. The enzyme showed better performance (1.8 times higher yield) in the synthesis of IABU than IAAC. The results were scaled up to a 150 mL reactor, and no differ- ences were observed in the yield, initial reaction rate, operational stability, and productivity using commercial isoamyl alcohol and fusel oil. Additionally, the structural isomers of isoamyl alcohol were evaluated. In conclusion, isoamyl ester industrial production has been proposed [167]. The esterification of fusel oil alcohols by butyric acid has also been reported with pancreatic lipase and Candida rugosa lipase with high yield under the optimized conditions of temperature, time, and acid/alcohol ratio [168]. Lipases are also used in the biocatalytic production of natural Green Leaf Volatiles (GLVs), which are aroma compounds associated with the green note odor. GLVs are widely used as aromas and food additives in the cosmetics and perfumes industry as well as in the food industry [169]. These examples indicate the successful synthesis of IAAC from fusel oil in a two-phase system, demonstrating that it is a viable alternative to pure isoamyl alcohol and an example of a circular economy reducing the waste generated from ethanol production plants. The use of lipase-displaying microorganism whole-cell biocatalysts is a promising alternative to classical immobilization supports with the advantage of low-cost preparation and, in some cases, high enzymatic activity [170]. CALB-displaying K. phaffii cells have been tested in the synthesis of a set of flavors esters, between them IAAC and IABU, and scaled up to a 5 L batch enzymatic bioreactor with conversions higher than 95% after four hours of reaction, with excellent operational stability. After ten batches, only an activity loss of 10% was detected in the presence of solvents [170]. A novel lipase obtained from metagenomics studies [171] is another example of a successful approach using whole cells—in this case, in the cell factory E. coli—to produce isoamyl acetate in a fluidized bed reactor with a reutilization of at least five cycles [172]. Catalysts 2022, 12, 960 13 of 24 3.4.2. Other Flavors and Alternative Reaction Systems Other flavors, such as ethyl butyrate, are the major component of many fruit flavors, such as pineapple, passion fruit, and strawberry [144]. The esterification was successfully implemented using Rhizopus oryzae lipase as a biocatalyst immobilized onto different supports. The best results were obtained with an acid:alcohol ratio of 1.45, and the reaction rate increased with increasing butyric acid concentration [173,174]. However, too high concentrations of butyric acid deactivated the enzyme [175]. Another problem concerns the difference in solubility between the substrates. En- zymatic bioreaction is usually performed in an aqueous environment; however, many flavor precursors and flavor products are not well soluble in water. Biphasic and al- ternative systems (aqueous]/organic, solid/gas, supercritical fluids, and ILs) overcome solubility problems [148]. As an alternative to using an enzymatic approach, the novo synthesis can also produce a mixture of flavors using the whole metabolic pathways of the microorganism. Using genetic engineering techniques, encoding specific genes from other microorganisms in cell factories, such as E. coli, has led to increased production of these compounds [148]. Reviews in this field have been recently published [149,176]. These natural metabolic routes have also been implemented in an enzymatic cascade reaction to produce cinnamyl cinnamate via a three-enzyme cascade incorporating the lipase Novozym 435 and in situ cofactor regeneration [177]. Additionally, a three-enzyme system (including lipase) to degrade curcumin to natural vanillin has been proposed [178]. 4. Trends in the Use of Lipases in Food and Nutraceuticals 4.1. Phenolic Antioxidants Phenolic compounds are secondary metabolites mainly extracted from fruits, veg- etables, and cereals, to which they impart color, flavor, and fragrance [179–181]. These compounds are mainly classified into flavonoids, phenolic acids, stilbenes, and curcum- inoids. They help with the control of diseases such as diabetes, obesity, hypertension, hyperlipidemia, and hyperglycemia, as well as inhibit adipogenesis [182–184]. Despite the benefits of consuming these phenolic compounds, the dose required to obtain their antioxidant power is limited by their low bioavailability and physiological stability, explaining why the synthesis of phenol polymers using enzymes such as glucan- otransferases and lipases has been resorted to [185–187]. The antioxidant activity of phenolic compounds present in foods can occur under different mechanisms. In the first case, by transferring a hydrogen atom from part of the phenolic compound to free radicals to be neutralized by a mechanism of transfer of a single electron from the phenolic compound with which it is left with an odd number of electrons distributed in the aromatic ring, the free radical forms an energetically stable spice with an even number of electrons. In the second case, proton transfer occurs from the phenolic compound to form an anion that subsequently donates an electron to transform the free radical into a stable molecule, and finally, a third case occurs by transition metal chelation [188]. However, when phenolic compounds are added to foods, their antioxidant, antimicrobial, anticarcinogenic, anti-inflammatory, antidiabetic, and antiobesity capacities are altered, increasing or decreasing them, because of intermolecular interactions with the macronutrients present. Thus, understanding these changes would allow their ex- ploitation to improve the quality, stability, organoleptic properties, and shelf life of the final products [189,190]. Due to their valuable benefits, the industrial demand for antioxidants in different sectors induces the search for new alternatives that facilitate the availability of these compounds. Thus, the use of new bioinformatics, molecular, proteomic, and biocatalysis tools suggests new antioxidant molecules to enhance their nutraceutical benefits [191,192]. The following section discusses cases of enzymatic modification and the synthesis of antioxidants for their biochemical improvement and benefit using biocatalytic processes to generate new molecules with nutraceutical prospects. Catalysts 2022, 12, 960 14 of 24 As mentioned previously, using lipases to synthesize food additives is a valuable tool for modifying ester or carboxyl groups. The aim is to improve the organoleptic properties of the ingredients and their miscibility with lipids, for which a hydrophilic element is introduced into the ester molecule. With the intervention of lipases through the acylation process, the structural modification of some flavonoids has been achieved, enhancing their stability and antioxidant activity; however, recent work has focused on the synthesis of lipophilic antioxidants [193]. Thus, a new compound called cyanidin-3-O-(dodecanoyl6) galactoside was obtained, generated after acylating lauric acid with the compound cyanidin- 3-O-galactoside extracted from alpine bearberry (Arctostaphylos alpine L.) and biocatalyzed by the commercial immobilized enzyme Novozym 435. Modification of this anthocyanin significantly improved its lipophilicity and thermostability while retaining its original antioxidant properties [193]. Another case of success by biocatalysis was performed with a natural phenolic acid, where caffeic acid was esterified by the commercial enzyme Novozym 435 to generate glyceryl-1-caffeate from ethyl caffeate and glycerol. Esterification of this compound with decanoic acid using the immobilized enzyme from Thermomyces lanuginosus (TL IM) and Novozym 435 in the presence of propylene carbonate was selective to monoacylated and diacylated products, respectively. This ingenious process is based on the low volatility of the solvent allowing the reaction to be performed under a vacuum and does not require sieves to remove the water produced. Likewise, the glyceryl caffeate ester products had greater stability than α-tocopherol in avoiding the oxidation of bulk tuna oil [194]. The search for new antioxidants with high antibacterial power synthesized from poorly studied precursors generates high expectations in research. For example, the enzymatic acylation of umbelliferone with different vinyl esters catalyzed with the enzyme Novozym 435 has been investigated for the first time. This allowed the generation of umbelliferone esters that presented a minimum inhibitory concentration of 1 mM for strains of clinical interest, such as Staphylococcus aureus (resistant to methicillin and oxacillin) and Klebsiella pneumoniae; for the Pseudomonas aeruginosa strain, its inhibitory capacity was 0.5 mM [195]. Considering that these improved products present greater solubility in lipid sub- stances, the prospect of these compounds as additives, antioxidants, and antimicrobials in different industrial sectors proves to be a real and available alternative. To improve the bioavailability of a flavonoid, a whole-cell enzyme system, a cell-bound lipase, and an intracellular enzyme were employed, which, by acylation and hydrolysis, respectively, allowed the bioconversion of naringin into two lipophilic derivatives: naringin esters and naringenin. The high antioxidant power of naringin esters was superior to its precursor, in addition to showing markedly enhanced permeability in human intestinal Caco-2 cells. However, naringenin is a product that can reduce bitterness in food products; thus, it has expectations in the industrial sector [54]. Continuing with the acylation mechanism, the antioxidant activity of some pheno- lipids, such as alkyl ferulate esters generated by biotransformation with lipases, was improved [51]. This new product showed significant antibacterial properties against Listeria monocytogenes (0.1 mM inhibitory capacity) and biofilm formation. The possible mechanism of action of the improved compound was related to the permeability and integrity of cell envelopes because it caused leakage of some cellular components in the bacteria under study. The new compound could bind to membrane proteins to disrupt protein activity or inhibit their synthesis, as well as bind to bacterial DNA and form complexes that affect their activity. This new compound has prospects in the nutraceutical industry because it can be employed as an adjuvant to address foodborne infections and biofilms [196]. Modification of polyphenol stilbenes, such as resveratrol, is another success of lipase. An acylation study using lipases from Alcaligenes sp. obtained a yield of approximately 70% of a new product that, after being characterized, was identified as 3-O-acetyl-resveratrol. The generation of this compound was observed when enzymes from T. lanuginosus and Pseudomonas cepacia were used. During the kinetic study, formation of the compounds identified as 3,40 -diacetyl- resveratrol and 3,5,40 -triacetylated in which all phenols were substituted was observed. Catalysts 2022, 12, 960 15 of 24 During the development of this study, an increase in temperature from 40 ◦ C to 60 ◦ C had a positive effect on the reaction yield; however, catalyst inactivation also occurred. When evaluating different acyl donors, such as free fatty acids, ethyl esters, and triglycerides, significant effects were only found with saturated and unsaturated vinyl esters. Addition- ally, the incorporation of long chains such as stearate was analyzed, achieving yields of up to 55% of monoesterified compounds at long reaction times, and the formation of di- and triacetylated compounds was not achieved [197–199]. However, a recent study used lipase from Candida sp. immobilized on hydrophobic- modified hollow mesoporous silica spheres (HMSS-C8). The catalytic efficiency of the enzyme was 15 times higher than that of the non-immobilized lipase, and the bioconversion to resveratrol ester was 98.8%, which was achieved in 2 h, the fastest reaction time recorded to date. This new compound presented higher solubility in plant lipids and Catalysts 2022, 12, x FOR PEER REVIEW provided 16 of 26 greater stability to oxidation, demonstrating its potential as an oil-soluble antioxidant [200]. The reactions described above can be visualized in Figure 5. Figure 5. Examples of phenolic antioxidants whose acylation is catalyzed by some lipase [54,201,202]. Figure 5. Examples of phenolic antioxidants whose acylation is catalyzed by some lipase [54,201,202]. 4.2. Prebiotics and Biosurfactants Prebiotics are nutrients that are broken down by gut microbiota (probiotics). In recent years, their relationship with human health attracted growing interest [201]. During fer- Catalysts 2022, 12, 960 16 of 24 4.2. Prebiotics and Biosurfactants Prebiotics are nutrients that are broken down by gut microbiota (probiotics). In recent years, their relationship with human health attracted growing interest [201]. During fer- mentation, the metabolic activity of microorganisms can change the nutritive and bioactive properties of food matrices, making them beneficial during consumption. Some oligo and polysaccharides act as prebiotic substrates that are fermented by the intestinal microbiota into short-chain fatty acids that are resistant to intestinal digestion. Although more clinical studies demonstrating the full functionality of the new products and strategies to increase yields are lacking, the advances thus far are very promising. While prebiotic oligosaccharides are water soluble, lipase-catalyzed esterification of oligosaccharides produces functionalized amphoteric molecules with biosurfactant/emulsifying properties and increased prebiotic activity, in addition to other bioactive properties, such as antibacterial, anti-inflammatory, and cytotoxic properties [127,202–204]. The acylation process has also allowed the use of other raw materials, such as sucrose, for value-added purposes, such as sugar fatty acid esters (SFAEs), which are compounds that have a carbohydrate moiety and one or more fatty acids as lipophilic SFAEs are nonionic surfactants that can be synthesized in a single enzymatic reaction step using lipases [205]. These products are employed in the food industry because of their high biodegradability and safety. They are synthesized by chemical and enzymatic methods, the latter being the most recent and studied because the reactions are performed under milder conditions and give greater confidence to the consumer by removing the synthetic chemical label. In this context, the synthesis of sucrose monolaurate was achieved by transesterifica- tion mediated by T. lanuginosus lipase immobilized on silica gel (Lipozyme TL IM), which has an antibacterial effect. With this precedent, new uses for biosurfactants alone or as adjuvants to address foodborne infections are sought [202]. Additionally, during the process, several challenges are encountered, such as the selection of the appropriate solvent for the reactions, because some hydrophilic organic solvents can dissolve sugars and fatty acids; at relatively high concentrations, most enzymes are affected. This is where the alternative of ionic liquids seems to minimize these effects, promoting favorable catalysis conditions [206,207]. 5. Concluding Remarks In this work, we observed the fundamental role of lipases in generating new products with high added value in the food and nutraceutical industry. Thus, the demand for these proteins in their different presentations is increasing. Enzymes are an alternative tool to produce new esters. In recent years, the use of lipases at the industrial level has been limited by their availability and costs; however, today, with the presence of new enzymes with properties that confer greater robustness, high yields, and stability to be reused in various reactions, they are positioned as a viable, accessible and compatible tool with the ecosystem that also provides valuable benefits to the food and nutraceutical sector, positively impacting the health of consumers. In the future, new advances in using biocatalysts are expected to continue to reduce the process steps and reaction times required to generate new products. Author Contributions: A.L.R.-R. retrieved the literature and writing original draft preparation of manuscript, figures, and tables. F.V.B. interpreted the literature, conceptualized, and supervised the manuscript, as well as funding acquisition. G.S. writing, review, and editing conceptualized and supervised the manuscript. All authors have read and agreed to the published version of the manuscript. Funding: This research was funded by the Spanish Ministry of Science and Innovation (Project PID2019-104666GB-100) (AEI/10.13039/501100011033). Data Availability Statement: As this manuscript is a review, datasets are not applicable and further information can be retrieved from the cited references. Catalysts 2022, 12, 960 17 of 24 Acknowledgments: Authors are grateful to ENZNUT and BIOCATEM networks for successful scientific collaborations. Gratitude to the National Institute of Forestry, Agricultural and Livestock Research (INIFAP) for its valuable support and interest in the scientific training of the first author during her doctoral stay. Conflicts of Interest: The authors declare no conflict of interest. References 1. Martirosyan, D.M.; Singh, J. A new definition of functional food by FFC: What makes a new definition unique? Funct. Foods Health Dis. 2015, 5, 209–223. [CrossRef] 2. Trifković, K.; Benković, M. Chapter 1-Introduction to Nutraceuticals and Pharmaceuticals. In Nutraceuticals and Natural Product Pharmaceuticals; Galanakis, C.M., Ed.; Academic Press: Cambridge, MA, USA, 2019; pp. 1–31. 3. Enespa; Chandra, P.; Singh, D. Sources, purification, immobilization and industrial applications of microbial lipases: An overview. Crit. Rev. Food Sci. Nutr. 2022, 18, 1–34. [CrossRef] [PubMed] 4. Baião, D.; Tahmasb, H.; Martínez, J.; Ciftci, O. Biocatalytic production of isoamyl acetate from fusel oil in supercritical CO2 . J. Supercrit. Fluids 2020, 164, 104917. [CrossRef] 5. Sloan, E. Top 10 Functional Food Trends. 2022. Available online: https://www.ift.org/news-and-publications/food-technology- magazine/issues/2022/april/features/top-10-functional-food-trends (accessed on 6 June 2022). 6. Contor, L. Functional Food Science in Europe. Nutr. Metab. Cardiovasc. Dis. NMCD 2001, 11, 20–23. 7. EFSA. European Food Safety Authority. General Function Health Claims under Article 13. 2022. Available online: https: //www.efsa.europa.eu/en/topics/topic/general-function-health-claims-under-article-13 (accessed on 6 June 2022). 8. Helal, N.; Eassa, H.; Amer, A.; Eltokhy, M.; Edafiogho, I.; Nounou, M. Nutraceuticals’ Novel Formulations: The Good, the Bad, the Unknown and Patents Involved. Recent Pat. Drug Deliv. Formul. 2019, 13, 105–156. [CrossRef] 9. Borrelli, M.; Trono, D. Recombinant Lipases and Phospholipases and Their Use as Biocatalysts for Industrial Applications. Int. J. Mol. Sci. 2015, 16, 20774–20840. [CrossRef] [PubMed] 10. Yao, W.; Liu, K.; Liu, H.; Jiang, Y.; Wang, R.; Wang, W.; Wang, T. A Valuable Product of Microbial Cell Factories: Microbial Lipase. Front. Microbiol. 2021, 12, 743377. [CrossRef] [PubMed] 11. Khan, U.; Selamoglu, Z. Use of Enzymes in Dairy Industry: A Review of Current Progress. Arch. Razi Inst. 2020, 75, 131–136. [CrossRef] [PubMed] 12. Szymczak, T.; Cybulska, J.; Podleśny, M.; Frac,˛ M. Various Perspectives on Microbial Lipase Production Using Agri-Food Waste and Renewable Products. Agriculture 2021, 11, 540. [CrossRef] 13. Chandra, P.; Enespa; Singh, R.; Arora, P.K. Microbial lipases and their industrial applications: A comprehensive review. Microb. Cell Factories 2020, 19, 169. [CrossRef] 14. Bilal, M.; Fernandes, C.D.; Mehmood, T.; Nadeem, F.; Tabassam, Q.; Ferreira, L.F.R. Immobilized lipases-based nano-biocatalytic systems—A versatile platform with incredible biotechnological potential. Int. J. Biol. Macromol. 2021, 175, 108–122. [CrossRef] 15. Casas, L.; Duquesne, S.; Bordes, F.; Sandoval, G.; Marty, A. Lipases: An Overview. In Lipases and Phospholipases: Methods and Protocols; Sandoval, G., Ed.; Humana Press: Totowa, NJ, USA, 2012; pp. 3–30. [CrossRef] 16. Sandoval, G. Lipases and Phospholipases, 2nd ed.; Springer: Berlin, Germany, 2018; Volume 1835, p. 437. 17. Ali, Y.; Verger, R.; Abousalham, A. Lipases or esterases: Does it really matter? Toward a new bio-physico-chemical classification. Methods Mol. Biol. 2012, 861, 31–51. [CrossRef] 18. Priyanka, P.; Tan, Y.; Kinsella, G.; Henehan, G.; Ryan, B. Solvent stable microbial lipases: Current understanding and biotechno- logical applications. Biotechnol. Lett. 2019, 41, 203–220. [CrossRef] [PubMed] 19. Lee, H.; Park, O. Lipases associated with plant defense against pathogens. Plant Sci. 2019, 279, 51–58. [CrossRef] [PubMed] 20. Rivera, I.; Mateos, J.; Sandoval, G. Plant Lipases: Partial Purification of Carica papaya Lipase. In Lipases and Phospholipases: Methods and Protocols; Sandoval, G., Ed.; Humana Press: Totowa, NJ, USA, 2012; pp. 115–122. 21. Gao, M.; Yin, X.; Yang, W.; Lam, S.; Tong, X.; Liu, J.; Wang, X.; Li, Q.; Shui, G.; He, Z. GDSL lipases modulate immunity through lipid homeostasis in rice. PLoS Pathog. 2017, 13, e1006724. [CrossRef] [PubMed] 22. Rivera, I.; Gutiérrez, A.; Sandoval, G. Functional Expression of Plant Lipases: The Case of CpLip1 from Carica papaya. Methods Mol. Biol. 2018, 1835, 169–178. 23. Villeneuve, P. Plant lipases and their applications in oils and fats modification. Eur. J. Lipid Sci. Technol. 2003, 105, 308–317. [CrossRef] 24. Raveendran, S.; Parameswaran, B.; Ummalyma, S.; Abraham, A.; Mathew, A.; Madhavan, A.; Rebello, S.; Pandey, A. Applications of Microbial Enzymes in Food Industry. Food Technol. Biotechnol. 2018, 56, 16–30. [CrossRef] 25. Valero, F. Heterologous Expression Systems for Lipases: A Review. In Lipases and Phospholipases: Methods and Protocols; Sandoval, G., Ed.; Humana Press: Totowa, NJ, USA, 2012; pp. 161–178. 26. Hamdan, S.H.; Maiangwa, J.; Ali, M.S.M.; Normi, Y.M.; Sabri, S.; Leow, T.C. Thermostable lipases and their dynamics of improved enzymatic properties. Appl. Microbiol. Biotechnol. 2021, 105, 7069–7094. [CrossRef] Catalysts 2022, 12, 960 18 of 24 27. Zhang, J.; Gao, B.; Lv, K.; Kumissay, L.; Wu, B.; Chu, J.; He, B. Specific immobilization of lipase on functionalized 3D printing scaffolds via enhanced hydrophobic interaction for efficient resolution of racemic 1-indanol. Biochem. Biophys. Res. Commun. 2021, 546, 111–117. [CrossRef] 28. Monteiro, R.; Virgen, J.; Berenguer, Á.; da Rocha, T.; dos Santos, J.; Alcántara, A.; Fernandez, R. Biotechnological relevance of the lipase A from Candida antarctica. Catal. Today 2021, 362, 141–154. [CrossRef] 29. Javed, S.; Azeem, F.; Hussain, S.; Rasul, I.; Siddique, M.; Riaz, M.; Afzal, M.; Kouser, A.; Nadeem, H. Bacterial lipases: A review on purification and characterization. Prog. Biophys. Mol. Biol. 2018, 132, 23–34. [CrossRef] [PubMed] 30. Juturu, V.; Wu, J. Heterologous Protein Expression in Pichia pastoris: Latest Research Progress and Applications. ChemBioChem 2018, 19, 7–21. [CrossRef] [PubMed] 31. Hee, K.; Huang, B.X.; Spector, A.A. Phosphatidylserine in the brain: Metabolism and function. Prog. Lipid Res. 2014, 56, 1–18. [CrossRef] 32. Souza, M.; Santos, K.; Freire, R.; Barreto, A.; Fechine, P.; Gonçalves, L. Production of flavor esters catalyzed by lipase B from Candida antarctica immobilized on magnetic nanoparticles. Braz. J. Chem. Eng. 2017, 34, 681–690. [CrossRef] 33. Kurtovic, I.; Nalder, T.D.; Cleaver, H.; Marshall, S. Immobilisation of Candida rugosa lipase on a highly hydrophobic support: A stable immobilised lipase suitable for non-aqueous synthesis. Biotechnol. Rep. 2020, 28, e00535. [CrossRef] [PubMed] 34. Trbojević, J.; Veličković, D.; Dimitrijević, A.; Bezbradica, D.; Dragačević, V.; Gavrović, M.; Milosavić, N. Design of biocompatible immobilized Candida rugosa lipase with potential application in food industry. J. Sci. Food Agric. 2016, 96, 4281–4287. [CrossRef] 35. Monoj, G. Chapter 13-Trans Fat Alternatives and Challenges. In Practical Guide to Vegetable Oil Processing, 2nd ed.; Gupta, M.K., Ed.; AOCS Press: Ubana, IL, USA, 2017; pp. 341–374. 36. Dayton, C. 11-Enzymatic Interesterification. In Green Vegetable Oil Processing; Farr, W.E., Proctor, A., Eds.; AOCS Press: Ubana, IL, USA, 2014; pp. 205–224. 37. Gricajeva, A.; Kazlauskas, S.; Kalėdienė, L.; Bendikienė, V. Analysis of Aspergillus sp. lipase immobilization for the application in organic synthesis. Int. J. Biol. Macromol. 2018, 108, 1165–1175. [CrossRef] [PubMed] 38. Muniandy, M.; Lasekan, O.; Ghazali, M.H.; Rahman Mohd, B. Lipase-Catalyzed formation of pentyl nonanoate using screened immobilized from Rhizomucor meihei. Braz. J. Chem. Eng. Life Sci. 2019, 36, 1089–1097. [CrossRef] 39. Zhang, X.; Ai, N.; Wang, J.; Tong, L.; Zheng, F.; Sun, B. Lipase-catalyzed modification of the flavor profiles in recombined skim milk products by enriching the volatile components. J. Dairy Sci. 2016, 99, 8665–8679. [CrossRef] 40. Shaimaa, H.; Shaaban, H.; Mahmoud, H.; Abbas, K.; Farouk, A. Preparation of Ras Cheese Flavour Concentrate using Lipolyzed Cream and Skim Milk Curd. Int. J. Dairy Sci. 2017, 12, 275–281. [CrossRef] 41. Tecelão, C.; Rivera, I.; Sandoval, G.; Ferreira-Dias, S. Carica papaya latex: A low-cost biocatalyst for human milk fat substitutes production. Eur. J. Lipid Sci. Technol. 2012, 114, 266–276. [CrossRef] 42. Konkit, M.; Kim, W. Activities of amylase, proteinase, and lipase enzymes from Lactococcus chungangensis and its application in dairy products. J. Dairy Sci. 2016, 99, 4999–5007. [CrossRef] [PubMed] 43. Esteban, M.; Mancheño, J.; de las Rivas, B.; Muñoz, R. Characterization of a halotolerant lipase from the lactic acid bacteria Lactobacillus plantarum useful in food fermentations. LWT-Food Sci. Technol. 2015, 60, 246–252. [CrossRef] 44. Alvarez, Y.; Esteban, M.; Cortés, Á.; Gago, F.; Acebrón, I.; Benavente, R.; Mardo, K.; de las Rivas, B.; Muñoz, R.; Mancheño, J. Esterase LpEst1 from Lactobacillus plantarum: A Novel and Atypical Member of the αβ Hydrolase Superfamily of Enzymes. PLoS ONE 2014, 9, e92257. [CrossRef] 45. Liu, C.; Chen, Y.; Hou, M.; Hu, N.; Chen, C.; Shaw, J. Crystallographic analysis of the Staphylococcus epidermidis lipase involved in esterification in aqueous solution. Acta Crystallogr. Sect. F Struct. Biol. Commun. 2018, 74, 351–354. [CrossRef] 46. Molina, M.; Hakalin, N.; Rodríguez, L.; Alcaraz, L.; López, F.; Martínez, M.; Prieto, A. Effect of the Immobilization Strategy on the Efficiency and Recyclability of the Versatile Lipase from Ophiostoma piceae. Molecules 2019, 24, 1313. [CrossRef] 47. Knob, A.; Izidoro, S.; Lacerda, L.; Rodrigues, A.; de Lima, V. A novel lipolytic yeast Meyerozyma guilliermondii: Efficient and low-cost production of acid and promising feed lipase using cheese whey. Biocatal. Agric. Biotechnol. 2020, 24, 101565. [CrossRef] 48. Chaves, A.F.L.; Castro, J.M.P.; Medeiros, T.B.; Soares, F.M.B. Trends in lipase immobilization: Bibliometric review and patent analysis. Process Biochem. 2021, 110, 37–51. [CrossRef] 49. Castillo, E.; Casas, L.; Sandoval, G. Medium-engineering: A useful tool for modulating lipase activity and selectivity. Biocatalysis 2016, 1, 178–188. [CrossRef] 50. Itoh, T. Activation of Lipase-Catalyzed Reactions Using Ionic Liquids for Organic Synthesis. In Application of Ionic Liquids in Biotechnology; Itoh, T., Koo, Y.-M., Eds.; Springer International Publishing: Cham, Switzerland, 2019; pp. 79–104. 51. Sandoval, G.; Quintana, P.; Baldessari, A.; Ballesteros, A.; Plou, F. Lipase-catalyzed preparation of mono- and diesters of ferulic acid. Biocatal. Biotransformation 2015, 33, 89–97. [CrossRef] 52. Xu, P.; Zheng, G.; Zong, M.; Li, N.; Lou, W. Recent progress on deep eutectic solvents in biocatalysis. Bioresour. Bioprocess. 2017, 4, 34. [CrossRef] [PubMed] 53. Coelho, A.L.S.; Orlandelli, R.C. Immobilized microbial lipases in the food industry: A systematic literature review. Crit. Rev. Food Sci. Nutr. 2021, 61, 1689–1703. [CrossRef] [PubMed] 54. Li, X.; Zhao, Y.; Lai, X.; Nong, J.; Zhao, G.; Xiao, X. One-pot biocatalytic synthesis and antioxidant activities of highly lipophilic naringin derivatives by using bi-functional whole-cells. Food Res. Int. 2020, 136, 109291. [CrossRef] Catalysts 2022, 12, 960 19 of 24 55. Sharma, R.; Sharma, N. Microbial Lipase Mediated by Health Beneficial Modification of Cholesterol and Flavors in Food Products: A Review. Recent Pat. Biotechnol. 2018, 12, 81–91. [CrossRef] 56. Khanniri, E.; Bagheripoor, N.; Sohrabvandi, S.; Mortazavian, M.; Khosravi, K.; Mohammad, R. Application of Liposomes in Some Dairy Products. Crit. Rev. Food Sci. Nutr. 2016, 56, 484–493. [CrossRef] 57. Shojaei, M.; Golmakani, M.; Eskandari, M.; Toh, M.; Liu, S. Natural flavor biosynthesis by lipase in fermented milk using in situ produced ethanol. J. Food Sci. Technol. 2021, 58, 1858–1868. [CrossRef] 58. Ferreira, S.; Osório, N.; Tecelão, C. 9-Bioprocess technologies for production of structured lipids as nutraceuticals. In Current Developments in Biotechnology and Bioengineering; Rai, A.K., Singh, S.P., Pandey, A., Larroche, C., Soccol, C.R., Eds.; Elsevier: Amsterdam, The Netherlands, 2022; pp. 209–237. 59. Guo, Y.; Cai, Z.; Xie, Y.; Ma, A.; Zhang, H.; Rao, P.; Wang, Q. Synthesis, physicochemical properties, and health aspects of structured lipids: A review. Compr. Rev. Food Sci. Food Saf. 2020, 19, 759–800. [CrossRef] 60. Burdge, G.; Calder, P. Introduction to fatty acids and lipids. World Rev. Nutr. Diet. 2015, 112, 1–16. [CrossRef] 61. Astrup, A.; Magkos, F.; Bier, D.; Brenna, J.; de Oliveira Otto, M.; Hill, J.; King, J.; Mente, A.; Ordovas, J.; Volek, J.; et al. Saturated Fats and Health: A Reassessment and Proposal for Food-Based Recommendations: JACC State-of-the-Art Review. J. Am. Coll. Cardiol. 2020, 76, 844–857. [CrossRef] 62. Temkov, M.; Mures, an, V. Tailoring the Structure of Lipids, Oleogels and Fat Replacers by Different Approaches for Solving the Trans-Fat Issue-A Review. Foods 2021, 10, 1376. [CrossRef] [PubMed] 63. Bahari, A.; Akoh, C. Synthesis of a Cocoa Butter Equivalent by Enzymatic Interesterification of Illipe Butter and Palm Midfraction. J. Am. Oil Chem. Soc. 2018, 95, 547–555. [CrossRef] 64. Bahari, A.; Akoh, C. Texture, rheology and fat bloom study of ‘chocolates’ made from cocoa butter equivalent synthesized from illipe butter and palm mid-fraction. LWT 2018, 97, 349–354. [CrossRef] 65. Casas, L.; Gasteazoro, F.; Duquesne, S.; Bordes, F.; Marty, A.; Sandoval, G. Lipases: An Overview. In Lipases and Phospholipases: Methods and Protocols; Sandoval, G., Ed.; Springer New York: New York, NY, USA, 2018; pp. 3–38. [CrossRef] 66. Simões, T.; Valero, F.; Tecelão, C.; Ferreira, S. Production of Human Milk Fat Substitutes Catalyzed by a Heterologous Rhizopus oryzae Lipase and Commercial Lipases. J. Am. Oil Chem. Soc. 2014, 91, 411–419. [CrossRef] 67. Mota, D.; Rajan, D.; Heinzl, G.; Osório, N.; Gominho, J.; Krause, L.; Soares, C.; Nampoothiri, M.; Sukumaran, R.; Ferreira, S. Production of low-calorie structured lipids from spent coffee grounds or olive pomace crude oils catalyzed by immobilized lipase in magnetic nanoparticles. Bioresour. Technol. 2020, 307, 123223. [CrossRef] [PubMed] 68. Kim, B.H.; Akoh, C.C. Recent Research Trends on the Enzymatic Synthesis of Structured Lipids. J. Food Sci. 2015, 80, C1713–C1724. [CrossRef] 69. Zhang, Z.; Zhang, S.; Lee, W.J.; Lai, O.M.; Tan, C.P.; Wang, Y. Production of Structured Triacylglycerol via Enzymatic Interesterifi- cation of Medium-Chain Triacylglycerol and Soybean Oil Using a Pilot-Scale Solvent-Free Packed Bed Reactor. J. Am. Oil Chem. Soc. 2020, 97, 271–280. [CrossRef] 70. Tecelão, C.; Perrier, V.; Dubreucq, E.; Ferreira, S. Production of Human Milk Fat Substitutes by Interesterification of Tripalmitin with Ethyl Oleate Catalyzed by Candida parapsilosis Lipase/Acyltransferase. J. Am. Oil Chem. Soc. 2019, 96, 777–787. [CrossRef] 71. Jiang, X.; Zou, X.; Chao, Z.; Xu, X. Preparation of Human Milk Fat Substitutes: A Review. Life 2022, 12, 187. [CrossRef] 72. Wei, W.; Jin, Q.; Wang, X. Human milk fat substitutes: Past achievements and current trends. Prog. Lipid Res. 2019, 74, 69–86. [CrossRef] 73. Hasibuan, H.A.; Sitanggang, A.B.; Andarwulan, N.; Hariyadi, P. Enzymatic Synthesis of Human Milk Fat Substitute—A Review on Technological Approaches. Food Technol. Biotechnol. 2021, 59, 475–495. [CrossRef] [PubMed] 74. Sun, L.; Xin, F.; Alper, H. Bio-synthesis of food additives and colorants-a growing trend in future food. Biotechnol. Adv. 2021, 47, 107694. [CrossRef] [PubMed] 75. Tomasz, P.; Maciej, B. The study on the use of flavonodi- prhosphatidylcholilne coating in extending the oxidative stability of flaxseed oil during storage. Food Packag. Sheld Life 2021, 28, 5. [CrossRef] 76. Farooq, S.; Abdullah; Zhang, H.; Weiss, J. A comprehensive review on polarity, partitioning, and interactions of phenolic antioxidants at oil-water interface of food emulsions. Compr. Rev. Food Sci. Food Saf. 2021, 20, 4250–4277. [CrossRef] [PubMed] 77. Boutillier, S.; Fourmentin, S.; Laperche, B. Food additives and the future of health: An analysis of the ongoing controversy on titanium dioxide. Futures 2020, 122, 102598. [CrossRef] 78. de Carvalho, C.; Caramujo, M. The Various Roles of Fatty Acids. Molecules 2018, 23, 2583. [CrossRef] 79. Carocho, M.; Barreiro, M.; Morales, P.; Ferreira, I. Adding Molecules to Food, Pros and Cons: A Review on Synthetic and Natural Food Additives. Compr. Rev. Food Sci. Food Saf. 2014, 13, 377–399. [CrossRef] 80. Revilla, I.; González, M.; Vivar, A.; Blanco, M.; Lobos, I.; Hernández, J. Antioxidant capacity of different cheeses: Affectinf factors and prediction by near infrared spectroscopy. J. Dairy 2015, 99, 5074–5082. [CrossRef] 81. Gammone, M.; Riccioni, G.; Parrinello, G.; D’Orazio, N. Omega-3 Polyunsaturated Fatty Acids: Benefits and Endpoints in Sport. Nutrients 2018, 11, 46. [CrossRef] 82. Pop, P.A.M.a.A. The role of antioxidants in the chemistry of oxidative stress: A review. Eur. J. Med. Chem. 2015, 97, 55–74. [CrossRef] 83. Zieniuk, B.; Wołoszynowska, M.; Białecka, E.; Fabiszewska, A. Application of freeze-dried Yarrowia lipolytica biomass in the synthesis of lipophilic antioxidants. Biotechnol. Lett. 2021, 43, 601–612. [CrossRef] [PubMed] Catalysts 2022, 12, 960 20 of 24 84. Zieniuk, B.; Białecka, E.; Wierzchowska, K.; Fabiszewska, A. Recent advances in the enzymatic synthesis of lipophilic antioxidant and antimicrobial compounds. World J. Microbiol. Biotechnol. 2021, 38, 11. [CrossRef] [PubMed] 85. Oteng, A.; Kersten, S. Mechanisms of Action of trans Fatty Acids. Adv. Nutr. 2020, 11, 697–708. [CrossRef] [PubMed] 86. Sies, H.; Jones, D. Reactive oxygen species (ROS) as pleiotropic physiological signalling agents. Nat. Reviews. Mol. Cell Biol. 2020, 21, 363–383. [CrossRef] 87. Godic, A.; Poljšak, B.; Adamic, M.; Dahmane, R. The Role of Antioxidants in Skin Cancer Prevention and Treatment. Oxidative Med. Cell. Longev. 2014, 2014, 860479. [CrossRef] 88. Khalaf, A.T.; Wei, Y.; Alneamah, S.J.A.; Al-Shawi, S.G.; Kadir, S.Y.A.; Zainol, J.; Liu, X. What Is New in the Preventive and Therapeutic Role of Dairy Products as Nutraceuticals and Functional Foods? BioMed Res. Int. 2021, 2021, 8823222. [CrossRef] 89. Ceriello, A.; Esposito, K.; La Sala, L.; Pujadas, G.; De Nigris, V.; Testa, R.; Bucciarelli, L.; Rondinelli, M.; Genovese, S. The protective effect of the Mediterranean diet on endothelial resistance to GLP-1 in type 2 diabetes: A preliminary report. Cardiovasc. Diabetol. 2014, 13, 140. [CrossRef] 90. Tan, B.; Norhaizan, M.; Liew, W. Nutrients and Oxidative Stress: Friend or Foe? Oxidative Med. Cell Longev. 2018, 2018, 9719584. [CrossRef] 91. Diamanti, E.; Papalou, O.; Kandaraki, E.; Kassi, G. Mechanisms in endocrinology: Nutrition as a mediator of oxidative stress in metabolic and reproductive disorders in women. Eur. J. Endocrinol. 2017, 176, R79–R99. [CrossRef] 92. Klaunig, J. Oxidative Stress and Cancer. Curr. Pharm. Des. 2018, 24, 4771–4778. [CrossRef] 93. Saha, S.K.; Lee, S.B.; Won, J.; Choi, H.Y.; Kim, K.; Yang, G.M.; Dayem, A.A.; Cho, S.G. Correlation between Oxidative Stress, Nutrition, and Cancer Initiation. Int. J. Mol. Sci. 2017, 18, 1544. [CrossRef] 94. Lee, J.D.; Cai, Q.; Shu, X.O.; Nechuta, S.J. The Role of Biomarkers of Oxidative Stress in Breast Cancer Risk and Prognosis: A Systematic Review of the Epidemiologic Literature. J. Women’s Health (2002) 2017, 26, 467–482. [CrossRef] [PubMed] 95. Zhang, L.; Li, L.; Gao, G.; Wei, G.; Zheng, Y.; Wang, C.; Gao, N.; Zhao, Y.; Deng, J.; Chen, H.; et al. Elevation of GPRC5A expression in colorectal cancer promotes tumor progression through VNN-1 induced oxidative stress. Int. J. Cancer 2017, 140, 2734–2747. [CrossRef] [PubMed] 96. Bisht, S.; Faiq, M.; Tolahunase, M.; Dada, R. Oxidative stress and male infertility. Nat. Rev. Urol. 2017, 14, 470–485. [CrossRef] 97. Tangvarasittichai, S. Oxidative stress, insulin resistance, dyslipidemia and type 2 diabetes mellitus. World J. Diabetes 2015, 6, 456–480. [CrossRef] 98. Durazzo, A.; Lucarini, M.; Santini, A. Nutraceuticals in Human Health. Foods 2020, 9, 370. [CrossRef] 99. Sachdeva, V.; Roy, A.; Bharadvaja, N. Current Prospects of Nutraceuticals: A Review. Curr. Pharm. Biotechnol. 2020, 21, 884–896. [CrossRef] 100. Aydin, S.; Dalgic, S.; Karaman, M.; Kirlangic, F.; Yildirim, H. Effects of Fulvic Acid on Different Cancer Cell Lines. Proceedings 2017, 1, 1031. 101. Swat, M.; Rybicka, I.; Gliszczyńska, A. Characterization of Fulvic Acid Beverages by Mineral Profile and Antioxidant Capacity. Foods 2019, 8, 605. [CrossRef] 102. Khan, J.; Deb, P.K.; Priya, S.; Medina, K.D.; Devi, R.; Walode, S.G.; Rudrapal, M. Dietary Flavonoids: Cardioprotective Potential with Antioxidant Effects and Their Pharmacokinetic, Toxicological and Therapeutic Concerns. Molecules 2021, 26, 4021. [CrossRef] 103. Martinelli, A.; Giannini, L.; Branduardi, P. Enzymatic Modification of Cellulose To Unlock Its Exploitation in Advanced Materials. ChemBioChem 2021, 22, 974–981. [CrossRef] [PubMed] 104. Karaki, N.; Aljawish, A.; Humeau, C.; Muniglia, L.; Jasniewski, J. Enzymatic modification of polysaccharides: Mechanisms, properties, and potential applications: A review. Enzym. Microb. Technol. 2016, 90, 1–18. [CrossRef] 105. de Araújo, M.; Franco, Y.; Messias, M.; Longato, G.; Pamphile, J.; Carvalho, P. Biocatalytic Synthesis of Flavonoid Esters by Lipases and Their Biological Benefits. Planta Med. 2017, 83, 7–22. [CrossRef] [PubMed] 106. Hammerling, U. Retinol as electron carrier in redox signaling, a new frontier in vitamin A research. Hepatobiliary Surg. Nutr. 2016, 5, 15–28. [CrossRef] 107. O’Byrne, S.; Blaner, W. Retinol and retinyl esters: Biochemistry and physiology. J. Lipid Res. 2013, 54, 1731–1743. [CrossRef] 108. Athar, M.; Zaidi, S.; Hassan, S. Intensification and optimization of biodiesel production using microwave-assisted acid-organo catalyzed transesterification process. Sci. Rep. 2020, 10, 21239. [CrossRef] 109. Huang, K.; Li, Y.; Kuo, C.; Twu, Y.; Shieh, C. Highly Efficient Synthesis of an Emerging Lipophilic Antioxidant: 2-Ethylhexyl Ferulate. Molecules 2016, 21, 478. [CrossRef] 110. Huang, S.-M.; Li, H.-J.; Liu, Y.-C.; Kuo, C.-H.; Shieh, C.-J. An Efficient Approach for Lipase-Catalyzed Synthesis of Retinyl Laurate Nutraceutical by Combining Ultrasound Assistance and Artificial Neural Network Optimization. Molecules 2017, 22, 1972. [CrossRef] 111. Caritá, A.; Fonseca, B.; Shultz, J.; Michniak, B.; Chorilli, M.; Leonardi, G. Vitamin C: One compound, several uses. Advances for delivery, efficiency and stability. Nanomed. Nanotechnol. Biol. Med. 2020, 24, 102117. [CrossRef] 112. Njus, D.; Kelley, P.; Tu, Y.; Schlegel, H. Ascorbic acid: The chemistry underlying its antioxidant properties. Free Radic. Biol. Med. 2020, 159, 37–43. [CrossRef] 113. Taira, N.; Katsuyama, Y.; Yoshioka, M.; Muraoka, O.; Morikawa, T. Structural Requirements of Alkylglyceryl-l-Ascorbic Acid Derivatives for Melanogenesis Inhibitory Activity. Int. J. Mol. Sci. 2018, 19, 1144. [CrossRef] [PubMed] Catalysts 2022, 12, 960 21 of 24 114. Tufiño, C.; Bernal, C.; Ottone, C.; Romero, O.; Illanes, A.; Wilson, L. Synthesis with Immobilized Lipases and Downstream Processing of Ascorbyl Palmitate. Molecules 2019, 24, 3227. [CrossRef] [PubMed] 115. Kharat, M.; Skrzynski, M.; Decker, E.; McClements, D. Enhancement of chemical stability of curcumin-enriched oil-in-water emulsions: Impact of antioxidant type and concentration. Food Chem. 2020, 320, 126653. [CrossRef] 116. Khalid, N.; Kobayashi, I.; Neves, M.; Uemura, K.; Nakajima, M. Microchannel emulsification: A promising technique towards encapsulation of functional compounds. Crit. Rev. Food Sci. Nutr. 2018, 58, 2364–2385. [CrossRef] 117. Díaz, Z.; Menghi, K.; Guerrero, M.; Nocelli, N.; Fanani, M. l-Ascorbic acid alkyl esters action on stratum corneum model membranes: An insight into the mechanism for enhanced skin permeation. Colloids Surf. B Biointerfaces 2020, 185, 110621. [CrossRef] [PubMed] 118. Tanini, D.; Lupori, B.; Malevolti, G.; Ambrosi, M.; Nostro, P.; Capperucci, A. Direct biocatalysed synthesis of first sulfur-, selenium- and tellurium- containing l-ascorbyl hybrid derivatives with radical trapping and GPx-like properties. Chem. Commun. 2019, 55, 5705–5708. [CrossRef] 119. Yadav, M.; Kavadia, M.; Vadgama, R.; Odaneth, A.; Lali, A. Production of 6-O-l-Ascorbyl Palmitate by Immobilized Candida antarctica Lipase B. Appl. Biochem. Biotechnol. 2018, 184, 1168–1186. [CrossRef] [PubMed] 120. Yang, C.; Luo, P.; Zeng, Z.; Wang, H.; Malafa, M.; Suh, N. Vitamin E and cancer prevention: Studies with different forms of tocopherols and tocotrienols. Mol. Carcinog. 2020, 59, 365–389. [CrossRef] 121. Lee, G.; Han, S. The Role of Vitamin E in Immunity. Nutrients 2018, 10, 1614. [CrossRef] 122. De la Fuente, M.; Sánchez, C.; Vallejo, C.; Díaz, E.; Arnalich, F.; Hernanz, Á. Vitamin C and vitamin C plus E improve the immune function in the elderly. Exp. Gerontol. 2020, 142, 111118. [CrossRef] 123. Zhou, P.; Shao, R.; Wang, H.; Miao, J.; Wang, X. Dietary vitamin A, C, and E intake and subsequent fracture risk at various sites: A meta-analysis of prospective cohort studies. Medicine 2020, 99, e20841. [CrossRef] [PubMed] 124. Busso, D.; David, A.; Penailillo, R.; Echeverría, G.; Rigotti, A.; Kovalskys, I.; Gómez, G.; Cortés Sanabria, L.Y.; Yépez García, M.C.; Pareja, R.G.; et al. Intake of Vitamin E and C in Women of Reproductive Age: Results from the Latin American Study of Nutrition and Health. Nutrients 2021, 13, 1954. [CrossRef] [PubMed] 125. Xin, J.; Chen, L.; Zhang, Y.; Wen, R.; Zhao, D.; Xia, C. Lipase-Catalyzed Synthesis of α-Tocopheryl Ferulate. Food Biotechnol. 2011, 25, 43–57. [CrossRef] 126. Zu, G.; Zhang, L.; Wang, Y. Research on Synthesis of Polyunsaturated Fatty Acid in Fish Oil and Vitamin E. Dalian Inst. Light Ind. 1999, 18, 294–298. 127. Casas, L.; Arrizon, J.; Arrieta, D.; Plou, F.; Sandoval, G. Synthesis and emulsifying properties of carbohydrate fatty acid esters produced from Agave tequilana fructans by enzymatic acylation. Food Chem. 2016, 204, 437–443. [CrossRef] 128. Mène-Saffrané, L.; Pellaud, S. Current strategies for vitamin E biofortification of crops. Curr. Opin. Biotechnol. 2017, 44, 189–197. [CrossRef] 129. Giménez, T.; Mula, D.; Gea, S.; Martínez, M.; Martí, N.; Valero, M.; Saura, D. Lipase catalyzed deacidification of tocopherol-rich distillates obtained from natural Vitamin E sources. Process Biochem. 2019, 77, 70–76. [CrossRef] 130. Chen, D.; Li, B.; Li, B.; Zhang, X.; Wei, L.; Zheng, W. Synthesis of vitamin E succinate catalyzed by nano-SiO2 immobilized DMAP derivative in mixed solvent system. Green Process. Synth. 2019, 8, 667–676. [CrossRef] 131. Zou, Z.; Dai, L.; Liu, D.; Du, W. Research Progress in Enzymatic Synthesis of Vitamin E Ester Derivatives. Catalysts 2021, 11, 739. [CrossRef] 132. Bampidis, V.; Azimonti, G.; Bastos, M.L.; Christensen, H.; Dusemund, B.; Durjava, M.F.; Kouba, M.; López-Alonso, M.; López Puente, S.; Marcon, F.; et al. Assessment of a feed additive consisting of all-rac-alpha-tocopheryl acetate (vitamin E) for all animal species for the renewal of its authorisation (DSM). EFSA J. 2021, 19, e06533. [CrossRef] 133. EFSA Panel on Dietetic Products, Nutrition, and Allergies (NDA). Scientific Opinion on Dietary Reference Values for vitamin E as α-tocopherol. EFSA J. 2015, 13, 4149. [CrossRef] 134. Gill, B.; Indyk, H. Separation of RRR-α-Tocopherol by Chiral Chromatography. J. AOAC Int. 2020, 103, 1288–1292. [CrossRef] [PubMed] 135. Fu, J.; Htar, T.; De Silva, L.; Tan, D.; Chuah, L. Chromatographic Separation of Vitamin E Enantiomers. Molecules 2017, 22, 233. [CrossRef] 136. Leal, L.; Beltrán, J.; Marc, B.; Bello, J.; den Hartog, L.; Hendriks, W.; Martín, J. Supplementation of lamb diets with vitamin E and rosemary extracts on meat quality parameters. J. Sci. Food Agric. 2020, 100, 2922–2931. [CrossRef] 137. Shan, L.C.; De Brún, A.; Henchion, M.; Li, C.; Murrin, C.; Wall, P.G.; Monahan, F.J. Consumer evaluations of processed meat products reformulated to be healthier—A conjoint analysis study. Meat Sci. 2017, 131, 82–89. [CrossRef] 138. Melis, S.; Delcour, J. Impact of wheat endogenous lipids on the quality of fresh bread: Key terms, concepts, and underlying mechanisms. Compr. Rev. Food Sci. Food Saf. 2020, 19, 3715–3754. [CrossRef] 139. Dai, Y.; Tyl, C. A review on mechanistic aspects of individual versus combined uses of enzymes as clean label-friendly dough conditioners in breads. J. Food Sci. 2021, 86, 1583–1598. [CrossRef] 140. Lien, G.; Bram, P.; Karolien, D.; Jan, D. Lipases and Their Functionality in the Production of Wheat-Based Food Systems. Compr. Rev. Food Sci. Food Saf. 2014, 13, 978–989. [CrossRef] Catalysts 2022, 12, 960 22 of 24 141. Min, B.; Salt, L.; Wilde, P.; Kosik, O.; Hassall, K.; Przewieslik, A.; Burridge, A.; Poole, M.; Snape, J.; Wingen, L.; et al. Genetic variation in wheat grain quality is associated with differences in the galactolipid content of flour and the gas bubble properties of dough liquor. Food Chem. X 2020, 6, 100093. [CrossRef] 142. Lien, R.G.; Bram, P.; Hanne, G.M.; Jan, A.D. A lipase based approach to understand the role of wheat endogenous lipids in bread crumb firmness evolution during storage. LWT-Food Sci. Technol. 2015, 64, 874–880. [CrossRef] 143. Gomes, A.; Meneses, A.; Araújo, P.; Oliveira, D. A review on enzymatic synthesis of aromatic esters used as flavor ingredients for food, cosmetics and pharmaceuticals industries. Trends Food Sci. Technol. 2017, 69, 95–105. [CrossRef] 144. López, J.; Benaiges, M.; Valero, F. Rhizopus oryzae Lipase, a Promising Industrial Enzyme: Biochemical Characteristics, Production and Biocatalytic Applications. Catalysts 2020, 10, 1277. [CrossRef] 145. Nitesh, C.; Himanshu, V.; Roshan, D. Flavors and Fragrance Market by Type (Flavors and Fragrance), Nature (Natural and Synthetic), and Application (Food & Beverages, Cosmetics & Personal Care, Home Care and Fabric Care): Global Opportunity Analysis and Industry Forecast, 2021–2027. Available online: https://www.researchandmarkets.com/reports/5341604/flavors- and-fragrance-market-by-type-nature-and (accessed on 11 August 2021). 146. Heinsman, N.; Franssen, M.; van der Padt, A.; Boom, R.; van’t Riet, K. Lipase-mediated Resolution of Branched Chain Fatty Acids. Biocatal. Biotransformation 2002, 20, 297–309. [CrossRef] 147. Leffingwell, J. Chirality & Odour Perception. Available online: www.leffingwell.com/chirality/chirality.htm (accessed on 23 April 2022). 148. Akacha, N.B.; Gargouri, M. Microbial and enzymatic technologies used for the production of natural aroma compounds: Synthesis, recovery modeling, and bioprocesses. Food Bioprod. Process. 2015, 94, 675–706. [CrossRef] 149. Bruno Nicolau, P.; Adones, S.; Lorena, F.; Gláucia Maria, P.; Gustavo, M.; Juliano Lemos, B. Recent advances in the microbial and enzymatic production of aroma compounds. Curr. Opin. Food Sci. 2021, 37, 98–106. [CrossRef] 150. Bansode, S.; Rathod, V. Enzymatic sythesis of Isoamyl butyrate under microwave irradiation. Chem. Eng. Process.-Process Intensif. 2018, 129, 71–76. [CrossRef] 151. Bansode, S.; Hardikar, M.; Rathod, V. Evaluation of reaction parameters and kinetic modelling for Novozym 435 catalysed synthesis of isoamyl butyrate. J. Chem. Technol. Biotechnol. 2017, 92, 1306–1314. [CrossRef] 152. Ferreira, S.; Sandoval, G.; Plou, F.; Valero, F. The potential use of lipases in the production of fatty acid derivatives for the food and nutraceutical industries. Electron. J. Biotechnol. 2013, 16, 1–38. [CrossRef] 153. Mahdieh, Z.; Mohammad, G.; Mehrdad, N. Lipase synthesis of isoamyl acetate using different acyl donors: Comparison of novel esterification techniques. LWT 2019, 101, 214–219. [CrossRef] 154. Tohid, T.; Atefeh, A.; Amin, T.; Somayeh, M.; Alieh, A.; Hamid, F.; Sara, T.; Mohammad, F. Lipase@zeolitic imidazolate framework ZIF-90: A highly stable and recyclable biocatalyst for the synthesis of fruity banana flavour. Int. J. Biol. Macromol. 2021, 166, 1301–1311. [CrossRef] 155. Hanen, G.; Karra, M.; Bezzine, S.; Nabil, M.; Youssef, G. Production of isoamyl acetate with immobilized Staphylococcus simulans lipase in a solvent-free system. Enzym. Microb. Technol. 2006, 38, 788–794. [CrossRef] 156. Macedo, G.; Pastore, G.; Rodrigues, M. Optimising the synthesis of isoamyl butyrate using Rhizopus sp. lipase with a central composite rotatable design. Process Biochem. 2004, 39, 687–693. [CrossRef] 157. Hari, K.; Prapulla, S.; Karanth, N. Enzymatic synthesis of isoamyl butyrate using immobilized Rhizomucor miehei lipase in non-aqueous media. J. Ind. Microbiol. Biotechnol. 2000, 25, 147–154. [CrossRef] 158. Bansode, S.; Rathod, V. Ultrasound assisted lipase catalysed synthesis of isoamyl butyrate. Process Biochem. 2014, 49, 1297–1303. [CrossRef] 159. Ozyilmaz, G.; Yağız, E. Isoamylacetate production by entrapped and covalently bound Candida rugosa and porcine pancreatic lipases. Food Chem. 2012, 135, 2326–2332. [CrossRef] 160. Cvjetko, M.; Jasna, V.; Polona, Ž. Isoamyl acetate synthesis in imidazolium-based ionic liquids using packed bed enzyme microreactor. Process Biochem. 2012, 47, 1344–1350. [CrossRef] 161. Azudin, N.Y.; Sangaran, S.; Abd Shukor, S.R. Non-enzymatic synthesis route for production of isoamyl acetate in a solvent-free system using miniaturized intensified reactor. J. Environ. Chem. Eng. 2020, 8, 103186. [CrossRef] 162. Muhammad, H.; Daniele, C.; Lasse, R. Two-stage catalytic hydrotreatment of highly nitrogenous biocrude from continuous hydrothermal liquefaction: A rational design of the stabilization stage. Biomass Bioenergy 2020, 139, 105658. [CrossRef] 163. Thi, T.; Suwadee, K.; Surachai, K.; Prasert, R.; Guoqing, G.; Narong, C.; Chanatip, S. Selective production of green solvent (isoamyl acetate) from fusel oil using a sulfonic acid-functionalized KIT-6 catalyst. Mol. Catal. 2020, 484, 110724. [CrossRef] 164. Anschau, A.; Aragão, V.; Porciuncula, B.; Kalil, S.; Burkert, C. Enzymatic Synthesis Optimization of Isoamyl Butyrate. J. Braz. Chem. 2011, 22, 2148–2156. [CrossRef] 165. Romero, M.; Calvo, L.; Alba, C.; Daneshfar, A.; Ghaziaskar, H. Enzymatic synthesis of isoamyl acetate with immobilized Candida antarctica lipase in n-hexane. Enzym. Microb. Technol. 2005, 37, 42–48. [CrossRef] 166. Kirdi, R.; Ben Akacha, N.; Messaoudi, Y.; Gargouri, M. Enhanced synthesis of isoamyl acetate using liquid-gas biphasic system by the transesterification reaction of isoamyl alcohol obtained from fusel oil. Biotechnol. Bioprocess Eng. 2017, 22, 413–422. [CrossRef] 167. López, J.; Benaiges, M.; Sebastian, X.; Bueno, J.; Valero, F. Producing Natural Flavours from Isoamyl Alcohol and Fusel Oil by Using Immobilised Rhizopus oryzae Lipase. Catalysts 2022, 12, 639. [CrossRef] 168. Gamayurova, V.; Shnaider, K.; Jamai, M. Enzymatic synthesis of butyrates of fusel oil. Catal. Ind. 2017, 9, 85–90. [CrossRef] Catalysts 2022, 12, 960 23 of 24 169. Vincenti, S.; Mariani, M.; Alberti, J.-C.; Jacopini, S.; Brunini-Bronzini de Caraffa, V.; Berti, L.; Maury, J. Biocatalytic Synthesis of Natural Green Leaf Volatiles Using the Lipoxygenase Metabolic Pathway. Catalysts 2019, 9, 873. [CrossRef] 170. Jin, Z.; Ntwali, J.; Han, S.; Zheng, S.; Lin, Y. Production of flavor esters catalyzed by CALB-displaying Pichia pastoris whole-cells in a batch reactor. J. Biotechnol. 2012, 159, 108–114. [CrossRef] [PubMed] 171. Meilleur, C.; Hupé, J.-F.; Juteau, P.; Shareck, F. Isolation and characterization of a new alkali-thermostable lipase cloned from a metagenomic library. J. Ind. Microbiol. Biotechnol. 2009, 36, 853–861. [CrossRef] 172. Sender, R.; Fuchs, S.; Milo, R. Revised Estimates for the Number of Human and Bacteria Cells in the Body. PLoS Biol. 2016, 14, e1002533. [CrossRef] 173. Guillén, M.; Benaiges, M.; Valero, F. Biosynthesis of ethyl butyrate by immobilized recombinant Rhizopus oryzae lipase expressed in Pichia pastoris. Biochem. Eng. J. 2012, 65, 1–9. [CrossRef] 174. Guillén, M.; Benaiges, M.; Valero, F. Improved ethyl butyrate synthesis catalyzed by an immobilized recombinant Rhizopus oryzae lipase: A comprehensive statistical study by production, reaction rate and yield analysis. J. Mol. Catal. B Enzym. 2016, 133, S371–S376. [CrossRef] 175. Grosso, C.; Ferreira, S.; Pires, P. Modelling and optimization of ethyl butyrate production catalysed by Rhizopus oryzae lipase. J. Food Eng. 2013, 115, 475–480. [CrossRef] 176. Verma, D.K.; Al-Sahlany, S.T.G.; Niamah, A.K.; Thakur, M.; Shah, N.; Singh, S.; Baranwal, D.; Patel, A.R.; Uta-ma, G.L.; Aguilar, C.N. Recent trends in microbial flavour Compounds: A review on Chemistry, synthesis mechanism and their application in food. Saudi J. Biol. Sci. 2022, 29, 1565–1576. [CrossRef] [PubMed] 177. Engelmann, C.; Johannsen, J.; Waluga, T.; Fieg, G.; Liese, A.; Bubenheim, P. A Multi-Enzyme Cascade for the Production of High-Value Aromatic Compounds. Catalysts 2020, 10, 1216. [CrossRef] 178. Esparan, V.; Krings, U.; Struch, M.; Berger, R. A three-enzyme-system to degrade curcumin to natural vanillin. Molecules 2015, 20, 6640–6653. [CrossRef] [PubMed] 179. Rashmi, H.; Negi, P. Phenolic acids from vegetables: A review on processing stability and health benefits. Food Res. Int. 2020, 136, 109298. [CrossRef] [PubMed] 180. Gómez, A.; Escobedo, Z.; Welti, J. Phenolic Compounds in Mesoamerican Fruits-Characterization, Health Potential and Processing with Innovative Technologies. Int. J. Mol. Sci. 2020, 21, 8357. [CrossRef] 181. Van, P. Phenolic Compounds of Cereals and Their Antioxidant Capacity. Crit. Rev. Food Sci. Nutr. 2016, 56, 25–35. [CrossRef] 182. Wang, W.; Li, J.; Zhang, H.; Wang, X.; Fan, J.; Zhang, X. Phenolic compounds and bioactivity evaluation of aqueous and methanol extracts of Allium mongolicum Regel. Food Sci. Nutr. 2019, 7, 779–787. [CrossRef] 183. Hussain, S.; Rehman, A.; Luckett, D.; Blanchard, C.; Obied, H.; Strappe, P. Phenolic Compounds with Antioxidant Properties from Canola Meal Extracts Inhibit Adipogenesis. Int. J. Mol. Sci. 2019, 21, 1. [CrossRef] 184. Wu, D.; Nie, X.; Shen, D.; Li, H.; Zhao, L.; Zhang, Q.; Lin, D.; Qin, W. Phenolic Compounds, Antioxidant Activities, and Inhibitory Effects on Digestive Enzymes of Different Cultivars of Okra (Abelmoschus esculentus). Molecules 2020, 25, 1276. [CrossRef] 185. Nagarajan, S.; Nagarajan, R.; Kumar, J.; Salemme, A.; Togna, A.; Saso, L.; Bruno, F. Antioxidant Activity of Synthetic Polymers of Phenolic Compounds. Polymers 2020, 12, 1646. [CrossRef] [PubMed] 186. González, J.; Míguez, N.; Padilla, D.; Leemans, L.; Poveda, A.; Jiménez, J.; Ballesteros, A.; Sandoval, G.; Plou, F. Optimization of Regioselective α-Glucosylation of Hesperetin Catalyzed by Cyclodextrin Glucanotransferase. Molecules 2018, 23, 2885. [CrossRef] [PubMed] 187. Li, H.M.; Xu, T.T.; Peng, Q.X.; Chen, Y.S.; Zhou, H.; Lu, Y.Y.; Yan, R.A. Enzymatic acylation of rutin with benzoic acid ester and lipophilic, antiradical, and antiproliferative properties of the acylated derivatives. J. Food Sci. 2021, 86, 1714–1725. [CrossRef] [PubMed] 188. Zeb, A. Concept, mechanism, and applications of phenolic antioxidants in foods. J. Food Biochem. 2020, 44, e13394. [CrossRef] 189. Zhang, Q.; Cheng, Z.; Wang, Y.; Fu, L. Dietary protein-phenolic interactions: Characterization, biochemical-physiological consequences, and potential food applications. Crit. Rev. Food Sci. Nutr. 2021, 61, 3589–3615. [CrossRef] 190. Wenfei, T.; Gengjun, C.; Michael, T.; Yonghui, L. Changes in phenolic profiles and antioxidant activities during the whole wheat bread-making process. Food Chem. 2021, 345, 128851. [CrossRef] 191. Panche, A.; Diwan, A.; Chandra, S. Flavonoids: An overview. J. Nutr. Sci. 2016, 5, e47. [CrossRef] 192. Zieniuk, B.; Groborz, K.; Wołoszynowska, M.; Ratusz, K.; Białecka, E.; Fabiszewska, A. Enzymatic Synthesis of Lipophilic Esters of Phenolic Compounds, Evaluation of Their Antioxidant Activity and Effect on the Oxidative Stability of Selected Oils. Biomolecules 2021, 11, 314. [CrossRef] 193. Yang, W.; Kortesniemi, M.; Yang, B.; Zheng, J. Enzymatic Acylation of Anthocyanins Isolated from Alpine Bearberry (Arctostaphy- los alpina) and Lipophilic Properties, Thermostability, and Antioxidant Capacity of the Derivatives. J. Agric. Food Chem. 2018, 66, 2909–2916. [CrossRef] 194. Cumming, H.; Marshall, S.N. Lipase-catalysed synthesis of mono- and di-acyl esters of glyceryl caffeate in propylene carbonate and their antioxidant properties in tuna oil. J. Biotechnol. 2021, 325, 217–225. [CrossRef] 195. Soares, V.; Marini, M.; de Paula, L.; Gabry, P.; Amaral, A.; Malafaia, C.; Leal, I. Umbelliferone esters with antibacterial activity produced by lipase-mediated biocatalytic pathway. Biotechnol. Lett. 2021, 43, 469–477. [CrossRef] [PubMed] Catalysts 2022, 12, 960 24 of 24 196. Shi, Y.; Bian, L.; Zhu, Y.; Zhang, R.; Shao, S.; Wu, Y.; Chen, Y.; Dang, Y.; Ding, Y.; Sun, H. Multifunctional alkyl ferulate esters as potential food additives: Antibacterial activity and mode of action against Listeria monocytogenes and its application on American sturgeon caviar preservation. Food Control 2019, 96, 390–402. [CrossRef] 197. Torres, P.; Poveda, A.; Jimenez-Barbero, J.; Ballesteros, A.; Plou, F.J. Regioselective lipase-catalyzed synthesis of 3-o-acyl derivatives of resveratrol and study of their antioxidant properties. J. Agric. Food Chem. 2010, 58, 807–813. [CrossRef] 198. Torres, P.; Reyes, D.; Ballesteros, A.; Plou, F. Lipase-Catalyzed Modification of Phenolic Antioxidants. In Lipases and Phospholipases: Methods and Protocols; Sandoval, G., Ed.; Humana Press: Totowa, NJ, USA, 2012; pp. 435–443. 199. Torres, P.; Plou, F.; Ballesteros, A. Modificación de antioxidantes fenólicos mediante procesos enzimáticos. In Obtención Enzimática de Compuestos Bioactivos a Partir de Recursos Naturales Iberoamericanos; Plou, F.J., Sandoval, G., Eds.; CSIC: Madrid, Spain, 2012; pp. 151–163. 200. Xu, L.; Yang, T.; Wang, J.; Huang, F.; Zheng, M. Immobilized Lipase Based on Hollow Mesoporous Silicon Spheres for Efficient Enzymatic Synthesis of Resveratrol Ester Derivatives. J. Agric. Food Chem. 2021, 69, 9067–9075. [CrossRef] [PubMed] 201. Davani, D.; Negahdaripour, M.; Karimzadeh, I.; Seifan, M.; Mohkam, M.; Masoumi, S.; Berenjian, A.; Ghasemi, Y. Prebiotics: Definition, Types, Sources, Mechanisms, and Clinical Applications. Foods 2019, 8, 92. [CrossRef] [PubMed] 202. Shao, S.; Shi, Y.; Wu, Y.; Bian, L.; Zhu, Y.; Huang, X.; Pan, Y.; Zeng, L.; Zhang, R. Lipase-Catalyzed Synthesis of Sucrose Monolaurate and Its Antibacterial Property and Mode of Action against Four Pathogenic Bacteria. Molecules 2018, 23, 1118. [CrossRef] [PubMed] 203. Ferrer, M.; Cruces, M.A.; Bernabé, M.; Ballesteros, A.; Plou, F.J. Lipase-catalyzed regioselective acylation of sucrose in two-solvent mixtures. Biotechnol. Bioeng. 1999, 65, 10–16. [CrossRef] 204. Sandoval, G.; Arrizon, J.; Gonalez-Avila, M.; Padilla-Camberos, E.; Matinez-Velazquez, M.; Villanueva-Rodriguez, S.; Casas- Godoy, L. Bioconjugate Molecules with Biological and Techno-Functional Activity, Method for the Production Thereof and Use Thereof. Patent Number MX 358789, 18 December 2013. 205. Neta, N.S.; Teixeira, J.A.; Rodrigues, L.R. Sugar ester surfactants: Enzymatic synthesis and applications in food industry. Crit. Rev. Food Sci. Nutr. 2015, 55, 595–610. [CrossRef] 206. van Rantwijk, F.; Madeira, L.; Sheldon, R. Biocatalytic transformations in ionic liquids. Trends Biotechnol. 2003, 21, 131–138. [CrossRef] 207. Shi, Y.; Wu, Y.; Lu, X.; Ren, Y.; Wang, Q.; Zhu, C.; Yu, D.; Wang, H. Lipase-catalyzed esterification of ferulic acid with lauryl alcohol in ionic liquids and antibacterial properties in vitro against three food-related bacteria. Food Chem. 2017, 220, 249–256. [CrossRef]