Immobilized Lipases in the Synthesis of Short-Chain Esters: An Overview of Constraints and Perspectives

Abstract

Biocatalysis—specifically the use of immobilized lipases—has been proposed as a greener alternative for ester production. Several critical challenges, such as the high cost of biocatalysts, are delaying the industrial implementation of biocatalysis. Moreover, for short-chain ester synthesis, the strong inhibition/inactivation potential of short-chain acids and alcohols on lipases leads to long reaction cycles and/or the need to use organic solvents to overcome the limitations of solvent-free systems and, consequently, the decrease in product concentrations. This review presents an overview of the scientific developments in enzymatic short-chain ester synthesis, compiling the constraints on their syntheses from a process perspective, including insights about key performance indicators (KPI) and economic parameters.

1. Introduction

Ester synthesis has been intensively explored in the scientific literature throughout the past decades [1,2,3,4,5,6]. Biocatalytic routes have gained great relevance in this field due to their environmental benefits toward greener and sustainable processes in chemical industries [7,8,9,10,11]. The adoption of biocatalysis in ester syntheses using lipases (E.C. 3.1.1.3) in immobilized form is challenging considering the high costs involved in the process and other technical drawbacks related to their catalytic activity and/or operational stability [5,8,12]. These drawbacks are more significant when considering the synthesis of short-chain esters, due to the acidity induced by short-chain acids, their relative hydrophilic characteristics, and the enzyme inactivation potential of short-chain alcohols [13,14,15,16,17,18]. Short-chain esters, whose chain length range comprises molecules containing 2 to 10 total carbons, are well known for their high volatility and flavor/fragrance properties. These compounds are derived from short-chain carboxylic acids and alcohols usually found in the metabolism of living organisms. Accordingly, some short-chain esters are naturally occurring compounds that can be extracted from plants and microorganisms [1,3,19,20,21,22,23] but with high costs and low yields. Short-chain esters, such as methyl, ethyl, and butyl acetate, are also important solvents and plasticizers [2,21,23,24,25]; the esters of dicarboxylic acids such as acrylic and succinic acids are precursors of polymers like polybutylsuccinate and polymethylmethacrylate [2,23,26]. Acetates, propionates, butyrates, and valerates are useful as chemical building blocks to obtain more complex molecules like fragrances, in which chemical modifications are required to improve stability or compatibility in personal care product formulations or even polymers [2,21,23,27]. Short-chain esters are stable liquids at room temperature. Most of them show a relatively hydrophilic behavior (logP < 2.0), have moderate solubility in aqueous buffers, are miscible with hydrophilic/moderately hydrophobic solvents, and are generally less dense than water [28,29]. Most of them are non-toxic and biodegradable compounds, which are important features for food ingredients, cosmetics, and personal care products [27,30,31,32,33]. Industrially, due to the high cost of their extraction from natural resources, esters are produced via esterification between a short-chain carboxylic acid and a short-chain alcohol in the presence of a catalyst via nucleophilic attack, forming water as a by-product [2,16,23].

Esterification is a thermodynamically controlled process, which means that yields are marked exclusively by the thermodynamic constant of the reaction; the catalysts can only alter the reaction rates, and yields can be under the thermodynamic value if the biocatalyst suffers inhibition or inactivation, as opposed to transesterification reactions [16,34,35]. The use of an excess of one of the reagents (generally the alcohol) is a possibility to improve the yields regarding the minority substrate; another option is the removal of water (the reaction by-product) to shift the thermodynamic equilibrium toward the desired product [16,17,36,37,38,39,40,41,42,43,44]. Due to the lower volatility of the substrate alcohol, selective distillation of the ester may be complex, and extracting the water by evaporation is not straightforward, as the substrates and products may have a lower evaporation temperature [45,46,47,48,49].

In practical terms, however, both strategies are simultaneously adopted—the use of an excess of alcohol and the removal of the formed water (via adsorption on molecular sieves, evaporation whenever possible, etc.) [45,46,47,50,51,52,53,54]. Short-chain alcohol and short-chain acid molecules frequently have mutual solubility, simplifying the use of solvent-free systems [17,37,43,55,56,57]. In this case, the hydrophobicity of the media will increase along with the reaction progress (as the ester will be more hydrophobic than the acid and the alcohol) with possible partitioning of the most hydrophobic unreacted reagents from the formed water where the most hydrophilic reagents may be concentrated if a water phase is formed [17,40,43,58,59]. If the increase in the hydrophobicity of the reaction medium is high enough when the ester is produced, this partitioning may form an aqueous phase in the system at some point in the reaction course (Figure 1). As the biocatalyst particles will be initially filled with the acid and the alcohol, it is likely that the hydrophobic drops of the ester may be unable to penetrate inside the biocatalyst by capillary forces, forming a kind of biphasic system and enabling higher yields of possible esters, as the product will be extracted from the enzyme environment [58,60,61]. This may be similar to the use of standard biphasic systems to shift the equilibrium in thermodynamically controlled reactions [62,63,64,65] with the difference that this is an in situ formed biphasic system and the researcher has scarce control over the hydrophobic phase, that will ultimately be the reaction product. In the following sections, we will discuss the short-chain esterification reactions in the industrial context and perspectives for biocatalysis in this field.

2. Discussion

2.1. Industrial Relevance and Market Size

Accurate data about production volume and specific market sizes are scarce. Some market research companies claim to offer such estimations [17,66]. For instance, one of the main players in this market, IFF^®—International Flavors and Fragrances—which has many short-chain esters in its portfolio, reported sales of USD 11.7 billion in 2021 [67]. The demand for esters as crucial ingredients, functioning as emulsifiers or stabilizers, coupled with the ingredient biodegradability and environmentally friendly characteristics are expected to drive and boost esters demand in many industries [1,68]. The high interest rates applied to innovative ingredients and sustainable processes in some sectors can be attested in the fragrance/cosmetics sector, which promotes annual awards for new products and specific categories for these topics, such as “In-Cosmetics Global Awards”, “Pure Beauty Global Awards”, and “Barcelona Perfumery Congress” [69,70,71].

Considering the range of direct applications in dairy products, it is intuitive to think that high volumes of short-chain esters are produced per year. In the United States alone, the esters market was valued at USD 3.8 billion in 2019 and is projected to surge to USD 5 billion by 2025. In a worldwide scenario, the esters market size was projected to be worth USD 94.19 billion in 2023, and there are prospects that the global market for esters will reach a staggering USD 159.36 billion by 2033 [72,73]. A significant concentration of major industry players is observed in the United States (U.S.), highlighting the influential role these companies (will) play in shaping global trends, innovations, and standards within the esters market [73].

This aforementioned growth is chiefly propelled by the rising need for emulsifiers and stabilizers in the personal care and detergent sectors [1]. Ethyl acetate, which is used as a solvent, chemical building block, or plasticizer, dominates the commercial market due to its biodegradable and environmentally friendly attributes [24,25,30,31,74]. Ethyl acetate, isopropyl acetate, and n-butyl acetate are recommended in the CHEM21 selection guide of solvents, which includes safety, health, and environmental criteria [75]. Some recent news from great worldwide companies like BASF^®, Evonik^®, and OQ^® Chemicals indicate that the production of short esters is continuously growing, as well as the expansion and construction of new facilities dedicated to ester production in recent years [76,77,78,79,80].

The economic viability of producing short-chain esters hinges on the production scale and successful commercialization, given their relatively modest costs [8,66]. Nevertheless, the biocatalytic pathway encounters hurdles in this scenario, primarily attributed to the high costs of commercial biocatalysts, notably immobilized lipases.

2.2. Evolution in the Ester Production Processes Using Standard Catalysts

Generally, industrial production uses conventional catalysts to obtain esters such as strong acids or metallic compounds [1,2,68]. Strong acids (inorganic, such as sulfuric or hydrochloric acid, or organic ones, such as p-toluenesulphonic acid), ion exchange resins, metals, metal oxides, or derivatives are usual catalysts for esterification reactions [2,9,68,81,82,83,84,85,86,87,88]. Acid catalysts act as the proton donor to the carboxylic acid; then, a nucleophilic attack by the alcohol may occur [2,16,37,38]. Jyoti and co-workers (2018) have demonstrated that sulfuric acid is the best catalyst when compared to other inorganic acids or even other heterogeneous catalysts in the syntheses of acrylic esters [85]. Its strong acidity and dehydrating capacity promote a rapid reaction rate. However, sulfuric acid is miscible with many different organic media, leading to additional difficulties in post-reaction operations besides its corrosive characteristics [2,81,85].

Homogeneous catalysts in a liquid–liquid reaction in a batch configuration demands a proper separation between products, non-reacted reagents, and the catalyst [2,68,84,89,90,91]. On the other hand, heterogeneous catalysts enable catalyst recovery by simple filtration or decanting after each reaction cycle [10,21,49,90,91,92,93]. The formation of water is another problem in these systems due to its negative effect on the stability of the catalysts. Thus, water removal is the critical step for obtaining high yields from the thermodynamic and kinetic perspective [17,40,41,49].

Due to the lack of catalytic selectivity and harsh reaction conditions (temperatures close to or higher than 100 °C), careful downstream operations are required for product purification; side products (such as sulphonic acid esters) may bring undesirable color and odor to the obtained ester, and unsaturated carboxylic acids are susceptible to thermal degradation or polymerization [2,68,94,95]. This purification step is more exigent in personal care applications or food/beverage industries, requiring deodorizing and bleaching processes, among other operations [1,68]. The neutralization of waste acid catalysts generally requires large volumes of alkali and costly waste treatment for final disposal [2,9].

To solve these limitations, alternative catalysts have been studied, including copper, tungsten, and zirconium oxides, modified clays, and processed wastes [2,86,88,96,97,98], showing promising results. As sulfuric acid is a widespread low-cost bulk chemical, careful comparison with new catalysts and sulfuric acid is required for future implementation. Additional criteria should be included to shift the chemical technology trend toward greener processes, such as life cycle assessment, other environmental indicators, and labeling/certification of the chemicals as green or “natural” [9,66,99,100,101]. However, during the development of new processes, the economic aspects should be realistic, considering the low percentual contribution of conventional catalysts in operational costs of the current processes [8,12,66,102,103].

2.3. New Trends for Ester Production: Biocatalysis

Cosmetics, personal care, and food products are important markets in which consumers may push the transition of ester production to green routes [1,3,9,68,104]. These industrial sectors are dynamic, demanding frequent changes in formulations and new appeals to the final customers [68]. Enzyme selectivity ensures the lack of undesired side products and their capacity to work under mild conditions, prevent undesired modification of the products, and allows more energy-efficient processes [1,9,68,105]. Esters produced by biocatalysis can be labeled as “green” or “natural” in some specific markets, permitting them to reach higher added values [1,38,68,106,107,108]. Moreover, biocatalytic routes can address many of the principles of Green Chemistry [37,102,109,110] and reduce the carbon footprint and greenhouse gas emissions [9,111,112]. This set of environmental advantages may increase the attractiveness of developing and implementing enzymatic routes in short ester syntheses.

Enzymes are highly selective catalysts that operate under mild reaction conditions; their use in organic syntheses, consequently, leads to the production of purer products, as the selectivity prevents the production of undesired by-products, and the mild conditions prevent decomposition of the reactants [1,9,68,102]. Enzymes used as biocatalysts are considered safe materials, although they may be subjected to some level of legal regulation as preconized in the Regulation (EC) 1907/2006 of the European Parliament concerning the Registration, Evaluation, Authorization and Restriction of Chemicals (REACH) and European Union Classification, Labelling and Packaging Regulation 1272/2008 [113,114]. More specifically, Enzyme TEC 21/19—ERC, describes that the only safety concern is related to a potential respiratory allergy during enzyme handling [115]. The mild reaction conditions under which lipases operate bring forth an energy saving to the process: about 60% less than the conventional process using sulfuric acid [9,68]. Their enantioselectivity permits obtaining a unique enantiomer from a prochiral substrate. Enzymes are also specific catalysts useful for the resolution of racemic mixtures [5,111]. The minimal formation of by-products in an enzymatic reaction facilitates the purification steps of products and increases their quality [1,68,94,107].

Among the possible enzymes that may be useful as biocatalysts for ester production, lipases stand out. Lipases catalyze in vivo metabolism of glycerides to produce free fatty acids and glycerol, but in vitro they can catalyze a diversity of reactions. Besides esterification, they can be used as catalysts in reactions such as transesterifications [16,116,117,118,119,120,121,122], acidolysis [123,124,125,126], interesterifications [127,128,129,130], or amidations [131,132,133,134,135], together with promiscuous reactions [5,136,137]. One of the most remarkable features of lipases is the presence of a mobile polypeptide chain (lid or flat) [138,139,140], that allows two different lipase conformations. The internal face of the lid is quite hydrophobic and interacts with the hydrophobic surroundings of the active center in homogeneous aqueous media (close and usually inactive form). When it moves, a large hydrophobic pocket is exposed to the medium and the active center is also exposed to the medium (open and active form) [138,139,140]. In most instances, the lid fully covers the active site in the closed form. There are cases with some exceptions such as the lipase B from Candida antarctica [140]. The huge hydrophobic pocket of the lipase open form can interact with any hydrophobic surface (like drops of the substrate, such as triglycerides), fixing the open form of the lipase to the drops of the substrate. This phenomenon is known as interfacial activation [138,141,142,143,144], since when the substrate concentration exceeds the solubility concentration and drops of the substrate are formed, a leap in enzyme activity is detected [138,141,142,143,144]. However, this tendency of the lipases to become adsorbed on hydrophobic surfaces may be expanded to any other hydrophobic surface different from oil drops [145], such as gas bubbles [146], some hydrophobic proteins [147,148,149], the open form of another lipase molecule [150,151,152,153], or, as it will be discussed later, a hydrophobic matrix [143].

In this aforementioned context, immobilized lipases have been extensively explored in the past decades for esterification reactions [6,66,111,154]. Despite the potential advantages, the ability of immobilized lipases to catalyze esterification reactions has been neglected in large-scale contexts considering that few industrial companies are applying this route [1,10,11,66,68]. As far as we know, none of them are related to short-chain esters. The main reason is due to technical and economic constraints, particularly the higher costs of immobilized lipases (generally between USD 100–1000 kg⁻¹) compared to the costs of soluble enzymes [8,12,105,155]. Purification and immobilization of lipases contribute greatly to the final costs [8,156], and strategies to obtain cheaper biocatalysts are required for the development of this technology, such as immobilization using agro-industrial wastes or the use of dry fermented solids with enzymatic activity [10,154,157,158]. According to Tüvfesson and co-authors (2011) [8], the maximum allowable cost contribution for the biocatalyst should be EUR 0.05–0.25 kg⁻¹ and productivities should be 400–2000 kg product per one kg of biocatalyst and 2000–10,000 kg product per one kg of biocatalyst, respectively, for production of specialties and commodities. Elevated costs could be compensated if the biocatalysts demonstrate a significant improvement in performance and extended reuse in the reaction [12,159].

The use of high concentrations of acids, which is a requirement to have a high concentration of the ester, may have a negative effect on the lipase performance [13,15,37]. These constraints are usually overcome using hydrophobic solvents, as this reduces the concentration of reagents and the extent of their negative effects [14,36,111,160]. The use of solvents is undesirable in industrial processes for different reasons: (i) reduction of volumetric productivity; (ii) the need for solvent recovery steps; (iii) increase in operational costs, and (iv) potential occupational and environmental risks [14,17,43,144]. Therefore, short-chain ester syntheses using immobilized lipases have an additional challenge—the need for an organic solvent—to achieve feasibility in a large-scale context, apart from the other well-known constraints for biocatalytic routes on ester production. The adoption of green solvents or conventional solvents that allow easy and efficient recycling can be a convenient path when their use is crucially required [161,162]. Another option is the design of lipase biocatalysts able to withstand the presence of high short-chain acids and alcohol concentrations.

Thus, even the operational and environmental advantages of the biocatalytic route—mild reaction conditions, purity of obtained products, and simplification of downstream operations—are overshadowed by the costs of immobilized lipases in the context of obtaining products of low–moderate value. Simple, efficient, and lipase-improving features of lipase immobilization should be developed to enable the use of these biocatalysts for this specific application.

2.4. Immobilization of Enzymes

Enzyme immobilization was initially developed to solve the problem of simplifying the recovery of the initially extremely expensive enzymes, and if the enzyme remains active, its reuse over subsequent reaction cycles [163,164,165]. Moreover, the use of immobilized enzymes allows the user to benefit from the advantages of the application of a heterogeneous catalyst (simplifying reactor control and the downstream of the product) [163,164,165]. Nowadays, the production prices of enzymes, and specifically of lipases, are rapidly decreasing, reducing the necessity for enzyme immobilization. For instance, Eversa^® has been launched by Novozymes^® as a lipase for biodiesel production to be employed in soluble (non-immobilized) form, and it accounts for only 5% of the production costs [166].

However, researchers tried to couple immobilization, an initially almost compulsory step to use enzymes in industry, to the improvement of other enzyme features [154]. The first goal was to improve enzyme stability, and soon it was shown that if the enzyme was covalently attached to the support via multiple bonds, enzyme rigidity may increase along with enzyme stability [154,167,168]. Later, some other reasons for enzyme stabilization after immobilization were pointed out, as recently reviewed [5,143,169]. However, it should be clear that enzyme stabilization is not a direct consequence of any randomly designed immobilization protocol, but the result of an adequate design of the immobilization process [154]. Immobilization has proved to be a valuable tool in tuning enzyme selectivity, specificity, inhibition, etc., as the enzyme structure is distorted by interacting with the support and this alters their final properties [143,170,171]. Comparative studies between immobilized and soluble lipases in the same synthesis demonstrate that the immobilized enzymes enable to get higher ester yields [172,173,174,175], due to the increase in stability, and specificity, together with the prevention of enzyme aggregation, which avoids the generation of new mass transfer matters [175,176,177]. Moreover, immobilization following a proper protocol can permit it to be coupled to enzyme purification, saving costs and enabling highly mass-activity biocatalysts using crude enzyme preparations [178].

Usual protocols for lipase immobilization include adsorption [143,179,180,181,182], entrapment [183,184,185,186,187,188], encapsulation [189,190,191], covalent coupling [192,193,194], and formation of cross-linked aggregates or crystals [172,173,195,196]. A variety of materials have been used as immobilization matrix–polymers [53,176,178,182,197,198,199,200,201], resins [120,202,203,204,205], inorganic compounds [61,129,205,206], or even waste materials (as fibers, straws, cloth, rice husk ash, and other agricultural waste) [207,208,209,210,211,212]. However, the most popular protocol is lipase immobilization on hydrophobic supports via interfacial activation of lipases against the support surface [143,154]. The hydrophobic supports used for this immobilization strategy are very chemically stable supports (as they have no chemically reactive moieties that can be inactivated or modified during storage or the lipase immobilization process) [143]. This method is very rapid and simple. By just mixing the enzyme solution and the support, the lipase becomes rapidly immobilized [143]. As the lipase adsorption on the support may be performed at low ionic strength, only lipases are adsorbed on these materials, enabling couple immobilization to the purification of the lipases [143]. Moreover, as the immobilization involves the open form of the lipase, a hyperactivated lipase form is obtained [143,213]. This adsorbed form of the lipases is more stable than the lipase in conformational equilibrium. It even is more stable than multipoint covalently immobilized lipases [214,215,216]. The properties of the immobilized enzymes on hydrophobic supports may be tuned by altering the support features (internal morphology, hydrophobicity, texture) [217,218] or the immobilization conditions [219,220,221], as it has been shown that the strong adsorption is enough to preserve the conformational form of the lipase induced by the immobilization conditions [220,221]. This way, a certain modulation of the properties of the final biocatalysts may be achieved using this reversible immobilization strategy. Using just one support and one enzyme, the final biocatalyst features (stability, activity, specificity) become very different depending on the immobilization conditions [219,220,221]. Thus, the control of the immobilization conditions must be quite strict to permit reproducible results. Lipase immobilization via interfacial activation on hydrophobic supports is a reversible process, enabling the reuse of the support after the enzyme immobilization [143]. That way, the global costs of the immobilization step may be reduced: the simplicity, increase in activity, and the possibility of support reuse are advantages from an economic point of view [154].

The main drawback of this immobilization strategy is founded on the reversibility of the immobilization: under certain circumstances, there are risks of enzyme release from the support [154,222]. This occurs using high temperature, high concentrations of hydrophobic cosolvents [223], or detergent-like reactants [222]. Researchers have proposed different solutions to this drawback. Physical cross-linking with ionic polymers maintains reversibility and can greatly reduce enzyme leakage [224,225]. Using covalent reactive polymers, such as polyethylenimine or aldehyde-dextran, avoids undesired enzyme release and ensures biocatalyst support reuse [226,227,228,229,230,231]. This makes the immobilization process more complex, demanding an additional reagent (the polymer) and an additional step in the biocatalyst preparation. Another alternative to prevent lipase release during operation is the use of heterofunctional supports, that is, supports bearing several functionalities able to interact with the enzyme [223,232,233,234,235,236,237,238,239,240,241,242,243,244,245,246,247]. The combination of ionic and hydrophobic moieties on the support surface maintains the reversibility and reduces the risks of enzyme release [244]. This can make the support more expensive but maintains the simplicity of the immobilization protocol. The use of chemically reactive groups able to give enzyme-support covalent bonds can suppress enzyme release but also prevents support reuse [243,244,245]. Moreover, the immobilization protocol may be more complex, requiring more reagents and additional immobilization steps (e.g., some reagents and steps to suppress the chemical reactivity of the support after immobilization). However, this opens the opportunity for a final tailoring of the lipase features [235,236,238,243] and also for the co-immobilization of lipases bearing different stabilities with the possibility of reusing the most stable one [248,249,250].

An inadequate selection of the mechanical properties of the support used to immobilize the enzyme may have very negative effects on the operational stability of the biocatalysts. For example, the shear stress of stirrers may be able to break the support particles, and this has many negative effects on the operation performance [5,12,95]. The formation of fine solid particles leads to the blocking of the filters, making it necessary to discard the biocatalysts even when the enzyme is fully active. Another significant problem of using immobilized lipases is the possibility of the formation of a water layer inside the biocatalysts if the activity of the biocatalysts is high enough [59,250,251]. That way, even if some strategy is utilized to control the bulk water activity, and a system using very low water activity is employed, this water layer around the enzyme can lead to enzyme inactivation, for example, in continuous oil transesterification [252]. The alcohol and mainly the acid may become concentrated in this aqueous phase, reducing the pH value where the enzyme is located and exposing the enzyme to very high alcohol concentrations. Furthermore, short-chain acids and alcohols may inactivate the enzyme and are also strong inhibitors acting as competitive inhibitors [13,14,16,17,37,144,253,254]. This can be at least partially solved using very hydrophobic supports, where water will not become adsorbed, or using ultrasounds, able to stir inside the particles and preventing the water layer formation [179,180,252,255,256,257,258,259,260,261,262]. For example, the use of Novozym^® 435, a commercial immobilized preparation of the lipase B from Candida antarctica using the interfacial adsorption of the enzyme in a moderately hydrophobic acrylic resin, Lewatit^® VP OC 1600 (a macroporous support formed by poly(methyl methacrylate) cross-linked with divinylbenzene) and perhaps the most used immobilized lipase biocatalysts present all these limitations: the mechanical resistance is short, the support permits the formation of a hydrophobic layer compromising the enzyme stability, enzyme leakage can occur, the support can be even dissolved in some solvents due to a no proper cross-linking [5,263].

Therefore, not only a proper immobilization protocol must be adopted but many other aspects must be considered to prepare a suitable industrial lipase biocatalyst.

2.5. Current Status of Immobilized Lipases on Short-Chain Ester Syntheses

The literature describes many efforts to produce short-chain esters using immobilized lipases. Some of them are focused on the chemical reaction, its optimization, and the understanding of kinetics and thermodynamics aspects, but a considerable part of them is focused on the immobilization of lipases, whose efficacy is tested in short-chain ester syntheses. Generally, these last ones do not include assays on a wide range of reaction conditions. However, these studies are important in the general context of the enzymatic ester syntheses, due to the aforementioned drawbacks related to the biocatalytic route, and the relevance of immobilization of enzymes for industrial applications.

Table 1. Short-chain ester syntheses from aliphatic reagents using immobilized lipases.

Ester	Biocatalyst Biocat. Loading	Reaction Conditions	Acid: Alcohol Molar Ratio	Solvent	Conversion (%)	Concentration (g L−1)	Productivity (g L−1 h−1)	Ref.
Octyl Formate	Novozym® 435 15 g L−1	Orbital shaker, 150 rpm, 1 h, 40 °C	1:7	1,2-dichloro- ethane	96.5	7.6	7.6	[264]
Hexyl Formate	Novozym® 435 15 g L−1	Orbital shaker, 150 rpm, 1.5 h, 40 °C	1:5	1,2-dichloro- ethane	98.3	12.8	8.5	[265]
Ethyl Acetate	Novozym® 435 8 g L−1	Orbital shaker, 150 rpm, 40 °C	1:4.4	n-heptane solvent-free	Not ment- ioned (only reaction rate)	-	-	[13]
Ethyl Acetate	Novozym® 435 6% (w/v)	Ultrasound (150 W/28 kHz), ~3 h	1:4	n-hexane	~82.0	-	-	[24]
Butyl Acetate	Lipase B from Candida antarctica immobilized onto porous styrene-divinylben-zene beads 10.0% (w/w) Novozym® 435 10.0% (w/w)	Orbital shaker, 200 rpm, 2 h Orbital shaker, 200 rpm, 2 h	1:5 1:3	n-hexane n-hexane	~90.0 ~90.0	31.4 31.4	15.7 15.7	[182]
Butyl Acetate	Candida antarctica immobilized onto porous styrene-divinylben-zene beads 7.5% (w/w)	Ultrasound (220 W/40 kHz), ~49 °C, 1.5 h	1:3.46	n-hexane	90.0	31.4	20.9	[255]
Butyl Acetate	Rhizopus oryzae lipase immobilized onto Celite 545 500 U in 5 mL	Orbital shaker, 200 rpm, 37 °C, ~12 h [a]	1:1	solvent-free n-heptane n-hexane	60.0 80.0 76.7	334.0 18.6 17.6	27.8 1.6 1.5	[203]
Butyl Acetate	Novozym® 435 7.5% (w/w)	Orbital shaker, 200 rpm, 2.5 h, 40 °C	1:3	n-hexane	~90.0	31.4	15.7	[266]
Butyl Acetate	Novozym® 435 7.0% (w/w)	Ultrasound, 2.5 h, 46 °C	1:3.6	n-hexane	94.0	32.8	13.1	[260]
Isopentyl Acetate	Lipozyme® RM IM 9.7% (w/w)	Orbital shaker, 150 rpm, 72 h, 40 °C	1:4	n-heptane	95.3	150	2.1	[267]
Pentyl Acetate	Lipozyme® 435 10.0% (w/w acid)	Shaken glass reactor, 100 rpm, 8 h, 40 °C	1:2	solvent-free	89.0	398.3	49.8	[21]
Isopentyl Acetate	Novozym® 435 5.0% (w/w)	Orbital shaker, 150 rpm, 6 h, 30 °C	1:2	solvent-free	80.0	0.57 g g−1	-	[268]
Isopentyl Acetate	Staphylococcus simulans lipase immobilized onto CaCO3 60 U	Screw-capped tubes, 200 rpm, 8 h, 37 °C [a]	1:2	solvent-free	64.0	495.7	61.2	[61]
Isopentyl Acetate	Candida antarctica immobilized onto polyurethane 10% (w/w)	Mechanical agitation, 160 rpm, 6 h, 64 °C Ultrasonic power, 105 W, 1 h, 65 °C	1:7 1:9	solvent-free	94.4 95.4	158.2 125.7	26.3 125.7	[269]
Isopentyl Acetate	Bacillus aerus lipase immobilized onto silica gel cross-linked with glutaraldehyde 1% (w/w)	Constant shaking, 10 h, 55 °C	1:1	solvent-free	68.0	65.1	0.7	[270]
Isopentyl Acetate	Porcine pancreas lipase immobilized onto activated carbon 0.5 g	Orbital shaker, 200 rpm, 4 h, 40 °C	1:1	n-hexane	93.0	29.1	7.28	[271]
Isopentyl Acetate	Candida rugosa lipase immobilized onto silica support/sodium alginate/calcium chloride 4 mg protein	Orbital shaker, 200 rpm, 8 h, 50 °C	1:2.6	n-hexane	85.2	374.0	46.7	[272]
Hexyl Acetate	Novozym® 435 13.8 g mol−1	Orbital shaker, 200 rpm, 3 h, 40 °C	1:1	n-hexane	93.0	67.1	22.3	[273]
3-Hexen-1-yl Acetate	Novozym® 435 2% (w/w) 9% (w/w)	Round bottom flask, 250 rpm, ~8 h, 40 °C Round bottom flask, 250 rpm, 24 h, 40 °C	1:1	n-hexane solvent-free	94.0 70.0	100.2 566.5	12.5 23.6	[274]
Ethyl Propionate	Bacillus coagulants (MTCC-6375) lipase immobilized onto polyhydrogel 31.4% (w/w)	Water-batch-incub- ator shaker and molecular sieves, 160 rpm, 9 h, 65 °C	1:3	n-heptane	89.3	9.1	1.0	[197]
Ethyl Propionate	Pseudomonas aeruginosa (BTS-2) lipase immobilized onto polyhydrogel network 23.6% (w/w)	Orbital shaker, 160 rpm, 3 h, 65 °C	1:3	n-nonane	94.0	9.6	3.2	[53]
Butyl Propionate	Novozym® 435 2.35% (w/w acid)	Orbital shaker, 130 rpm, ~25 h, ~43 °C	1:9	solvent-free	93.7	132.5	5.3	[275]
Butyl Propionate	Novozym® 435 Lipozyme® RM IM Lipozyme® TL-IM 0.98% (w/w) each	Microwave, 50 W, 400 rpm, 8 h, 60 °C	1:12	solvent-free	92.0 14.0 9.0	102.2 15.6 10.0	12.8 1.9 1.1	[276]
Isobutyl Propionate	Fermase® CALB 1000 4% (w/w)	Ultrasound, 40 W, 25 kHz, 150 rpm, 3 h, 60 °C	1:3	solvent-free	95.1	354.5	118.1	[277]
Isobutyl Propionate	Novozym® 435 5% (w/w)	Orbital shaker, 300 rpm, 10 h, 40 °C	1:3	solvent-free	92.5	345.0	34.5	[37]
Hexyl Propionate	Novozym® 435 13.8 g mol−1	Orbital shaker, 200 rpm, 3 h, 40 °C	1:1	n-hexane	94.0	74.3	24.7	[273]
Ethyl Butyrate	Candida rugosa lipase immobilized onto cotton cloth 25.4% (w/w)	Orbital shaker, 220 rpm, 24 h, 25 °C	2.4:1	Cyclohexane	91.2	26.5	1.1	[207]
Ethyl Butyrate	Candida rugosa lipase immobilized into polyurethane foams 1560 mg	Continuous packed-bed reactor, 260 min, residence time, 36 h, 30 °C	1:1.5	n-hexane	80.7	32.7	0.7	[278]
Ethyl Butyrate	Novozym® 435 7% (w/w)	Orbital shaker, 150 rpm, 96 h, 34 °C	1:3	n-heptane	72.9	33.9	0.4	[279]
Ethyl Butyrate	Candida antarctica lipase B immobilized onto styrene beads 10% (w/w) Novozym® 435 7.5% (w/w)	Orbital shaker, 200 rpm, 1 h, 37.5 °C [a] Orbital shaker, 200 rpm, 1 h, 44 °C [a]	1:4	n-hexane n-hexane	86.2 87.7	69.9 70.7	69.9 70.7	[183]
Isobutyl Butyrate	Lipozyme® RM-IM 14% (w/w)	Ultrasound, 2.5 h, 45 °C	1:1	n-hexane	86.0	-	-	[259]
Butyl Butyrate	Novozym® 435 -	Shaken round bottom flask, ~10 h, 56 °C	1:3.93	solvent-free	99.6	-	-	[280]
Butyl Butyrate	Thermomyces lanuginosus lipase immobilized onto glyoxyl-agarose beads 10% (w/w)	Orbital shaker, 200 rpm, 24 h, 37 °C	1:1	heptane	85.0	-	-	[194]
Butyl Butyrate	Novozym® 435 12% (w/w)	Round-bottom flasks, 250 rpm, 0.5 h, 57 °C	1:1	diesel oil	90.0	129.8	259.6	[281]
Butyl Butyrate	Cercospora kikuchii lipase immobilized onto chitosan beads 42.0% (w/w)	45 °C	1:1.5	n-heptane	95.0	18.0	3.0	[282]
Butyl Butyrate	Candida rugosa lipase immobilized onto iron-alginate nanoparticles -	Orbital shaker, 100 rpm, 6 h, 50 °C	1:4	solvent-free	90.0	-	-	[283]
Butyl Butyrate	Porcine pancreas lipase immobilized onto activated carbon 0.5 g	Orbital shaker, 200 rpm, 4 h, 40 °C	1:2	n-hexane	82.0	14.2	3.5	[284]
Butyl Butyrate	Candida rugosa lipase immobilized onto multiwalled carbon nanotubes 11.6% (w/w)	Orbital shaker, 200 rpm, 4 h, 37 °C	1:3	n-hexane	91.5	13.2	3.3	[285]
Butyl Butyrate	Candida rugosa lipase immobilized onto multiwalled carbon nanotubes 4.2% (w/w)	Orbital shaker, 200 rpm, 4 h, 30 °C	1:1	n-hexane	92.7	13.4	3.3	[175]
Isopentyl Butyrate	Lipozyme® TL IM 14.7% (w/w)	Orbital shaker, 180 rpm, 6 h, 40 °C,	1:2	n-hexane	96.1	259.5	43.3	[286]
Isopentyl Butyrate	Lipozyme® IM-20 1.9% (w/w)	Orbital shaker, 150 rpm, 24 h, 40 °C	1:1	n-hexane	~92.0	36.3	1.51	[287]
Isopentyl Butyrate	Rhizopus oryzae lipase immobilized onto Purolite (PMMA with epoxide and octadecyl groups) 35 U	Orbital shaker, 1200 rpm, 12 h, 30 °C	1:2	cyclohexane	91.0	59.0	4.9	[108]
Hexyl Butyrate	Novozym® 435 13.8 g mol−1	Orbital shaker, 200 rpm, 3 h, 40 °C	1:1	n-hexane	93.0	80.1	26.7	[273]
Hexyl Butyrate	Candida rugosa lipase immobilized onto Diaion HP20 10.8% (w/w)	Orbital shaker, 200 rpm, 3 h, 40 °C	1:2	n-heptane	94.5	81.8	27.6	[288]
Ethyl Valerate	Candida rugosa lipase immobilized onto microemulsion-based organogels -	Orbital shaker, 150 rpm, 144 h, 40 °C	1:1.6	cyclohexane	99.9	13.0	0.1	[289]
Ethyl Valerate	Candida rugosa lipase immobilized onto app membrane 696.6 U in 20 mL	Screw-capped flasks, 150 rpm, 24 h, 40 °C [a]	2:1 1:1	solvent-free n-hexane	34.8 67.2	164.2 583.2	6.8 24.3	[178]
Ethyl Valerate	Novozym® 435 10.06 g L−1	Microwave reactor, 0.67 h, 50 °C	1:1	solvent-free	69.2	8.0	12.0	[290]
Ethyl Valerate	Candida antarctica lipase B 12% (w/v)	Orbital shaker, 1000 rpm, 2 h, 40 °C Microwave, 100 W, 1 h, 40 °C	1:2 1:2	solvent-free	90.0 82.0	518.1 472.1	259.1 472.1	[291]
Ethyl Valerate	Aspergillus oryzae immobilized onto an electrospun nanofibrous membrane 1.10 g L−1	Magnetical stirring in oil bath, 250 rpm, 24 h, 50 °C [a]	1:1	solvent-free	72.1	23.3	1.0	[292]
Ethyl Valerate	Burkholderia cepacian lipase immobilized onto sodium alginate 20% (w/v)	Orbital shaker, 150 rpm, 48 h, 37 °C	1:2	n-heptane	87.1	58.6	1.2	[293]
Ethyl Valerate	Candida rugosa lipase immobilized onto biogenic sílica/magnetite/ graphene oxide 2.1% (w/w)	Orbital shaker, 200 rpm, 3 h, 40 °C	1:2	n-heptane	90.4	87.5	29.1	[294]
Pentyl Valerate	Candida rugosa lipase immobilized onto nanocrystalline cellulose, silica, and polyethersulfone 3 mg mL−1	Orbital shaker, 200 rpm, 3 h, 50 °C	1:2	cyclohexane	91.3	-	-	[295]
Pentyl Valerate	Candida rugosa lipase immobilized onto microemulsion-based organogels 70 g L−1	Orbital shaker, 150 rpm, 199 h, 37 °C	2.9:2	cyclohexane/AOT	99.9	17.0	0.1	[109]
Ethyl Hexanoate	Antarctic pseudomonas (AMS8) immobillized onto chitosan 5% (w/w)	Orbital shaker, 200 rpm, 2 h, 20 °C	1:1	toluene solvent-free	69.0 47.0	2.5 372.8	1.3 93.2	[296]

[a] Studies that include water addition in the beginning of the reaction.

and

Table 2. Short-chain ester syntheses from aromatic reagents using immobilized lipases.

Ester	Biocatalyst Biocat. Loading	Reaction Conditions	Acid: Alcohol Molar Ratio	Solvent	Conversion (%)	Concentration (g L−1)	Productivity (g L−1 h−1)	Ref.
Phenethyl Formate	Novozym® 435 15 g L−1	Orbital shaker, 150 rpm, 4 h, 40 °C	1:5	1,2-dichloroethane	95.9	14.4	3.6	[297]
Phenethyl Formate	Novozym® 435 15 g L−1	Orbital shaker, 150 rpm, 30 min, 30 °C	1:1	n-hexane	69.2	-	-	[121]
Benzyl Propionate	Novozym® 435 10% (w/w)	Thermal bath with mechanical agitation, 24 h, 65 °C	1:5	solvent-free	44.0	127.6	5.3	[298]
Benzyl Propionate	Novozym® 435 10% (w/w)	Fed-batch system with molecular sieves, 150 rpm, 6 h, 50 °C	1:5	solvent-free	99.0	270.7	45.1	[299]
Benzyl Butyrate	Pseudomonas cepacian lipase immobilized onto polyvinyl alcohol and chitosan 8.6% (w/w)	Ultrasound, 100 W, 3 h, 52 °C	1:3	isooctane	40.0	-	-	[176]
Benzyl Butyrate	Novozym® 435 2% (w/w)	Round bottom flask with fish-clip spinner, 20 h, 52 °C; irreversible esterification reaction (chemical drying)	1:1	methyl tert-butyl ether	82.0	160.4	8.0	[300]
Methyl Cinnamate	Candida sp. lipase immobilized onto ordered mesoporous silicon C8 10 mg	Orbital shaker, 220 rpm, 2 h, 70 °C	1:3	isooctane	90.4	1.5	0.8	[301]

present a bibliographic survey on the main reaction parameters for the enzymatic syntheses of esters via direct esterification with chain length of up to 10 carbons in the final molecule.

2.6. Performance Indicators on Enzymatic Esterification Studies and Environmental Aspects

Few studies achieved productivities higher than 100 g L⁻¹ h⁻¹ in enzymatic short-chain ester syntheses. The highest result was obtained by Garcia-Cebrian et al. (2018) [291], 472.1 and 259.1 g L⁻¹ h⁻¹, in solvent-free ethyl valerate synthesis using microwave and conventional heating, respectively. Other studies also found productivities close to 100 g L⁻¹ h⁻¹, such as Jaiswal and Rathod (2018) [277] in isobutyl propionate manufacture, Musa et al. (2018) [296] in ethyl hexanoate synthesis, and Friedrich et al. (2012) [182] in ethyl butyrate production. The main features of these studies are the short reaction time (1–3 h) and high product concentrations obtained. In this sense, we emphasize two important aspects—at least one of the substrates has a chain length equal to or superior to 4, which means that one of them is not so strong an inhibitor/inactivator of the acting lipases, and high biocatalyst loadings (superior to 5% w/w) are utilized. For comparative purposes, the work of Jiang and co-authors used a copper salt as a heterogeneous catalyst in the synthesis of isoamyl acetate resulting in a productivity of 358.9 g L⁻¹ h⁻¹ [86]. The biocatalyst loading is directly correlated with reaction time, but high loadings have operational and economical constraints for the process, as previously discussed. The long reaction time (higher than 24 h) required for some syntheses [278,279,289,293] may be associated with enzymatic inhibition or inactivation caused by short-chain acids and alcohols even using solvents. Thus, the use of solvents in this reaction is justified to reduce the concentration of substrates as they are potential inhibitors/inactivating agents for lipases. These reagents are relatively hydrophilic, which means that they can interact with the solvation layer of the enzyme altering the three-dimensional enzyme structure [14,15,16,302,303,304,305,306]. This may affect the required flexibility of the enzymes, and consequently, their catalytic activity [60,303,307,308,309]. Therefore, the inhibition/inactivating potential and the hydrophilic characteristic of short-chain acids and alcohols are significant challenges for the optimum performance of immobilized lipases. Some authors evaluate the enzymatic activity assays using standard substrates for hydrolysis, that are also different from the ones in the esterification reaction (for instance, p-nitrophenyl derivatives, olive oil, or oleic acid esters); this hydrolytic activity may not be directly related to the esterification activity [178,293,296,310]. There are some possible approaches to circumvent these drawbacks: (i) the adoption of high biocatalyst loadings (>5% w/w), aiming to increase reaction rates; (ii) optimization of the immobilization protocol and a careful choice of immobilization support to improve enzyme activity and stability.

It is possible to observe from the data in Table 1 and Table 2 that concentrations of obtained esters below 50 g L⁻¹ are frequent, which is low for industrial applications. This result is directly associated with the use of the solvents. Even in fermentative processes, that are known for low product concentrations and the need for complex downstream operations to separate the main product and co-products of cell metabolism, concentrations of products superior to 50 g L⁻¹ are usually obtained [11,311]. Therefore, it is hard to justify in economic terms the adoption of expensive catalysts such as immobilized lipases for producing commodity/specialty chemicals in the presence of solvents and at low concentrations. On the other hand, concentrations as high as 100 g L⁻¹ using solvents were achieved by some authors [178,272,286,300]. An important aspect to highlight in these studies is the possibility of working with high concentrations of substrates. For instance, Anschau and co-workers (2021) [286] used Lipozyme^® TL-IM as a biocatalyst in the synthesis of isopentyl butyrate in n-hexane without any water removal method and using response surface methodologies (RSM) as an optimization strategy; relatively high acid concentrations were used (0.5 mol L⁻¹ of butyric acid), indicating the significant tolerance of this lipase for butyric acid [286]. However, this result was achieved with high biocatalyst loading (11.5 g L⁻¹ as reported, equivalent to 21.3% w/w total substrate mass) and a relatively long reaction time (6 h). Thus, it is possible to obtain high concentrations of the product even using solvents, since high substrate concentrations are tolerated by the immobilized lipase. Fed-batch strategies are also interesting alternatives to circumvent the effects on immobilized lipases of high concentrations of substrates.

Considering KPI—yield, concentration, and productivity—it is possible to suggest how far short-chain ester syntheses using biocatalysis are from industrial applications. Using the feasibility criteria proposed by Tüvfesson and co-workers (2011) [8] in a biocatalytic production—2000 kg product per one kg of immobilized biocatalyst for specialties chemicals—we can check that even high concentrations of products result in low kg product per kg biocatalyst, as the results reported by Musa et al. (2018) [296]—only 4.2 kg of product per kg of biocatalyst. Some works [21,108,194,278,279,280,293] argue that biocatalysis may bring advantages in terms of simplification of processes (and consequential reduction in costs) due to the specificity and selectivity of biocatalysts, but the need for solvent separation and recovery in a biocatalytic process will represent the opposite.

The acid-mediated syntheses (using H₂SO₄ or other inorganic ones) are generally carried out for short reaction times, varying from 1 to 8 h in many cases [2]; contrarily, a significant part of the studies shown in Table 1 and Table 2 present reaction times equal to or higher than 24 h, although many of them did not include a detailed evaluation of the reaction time; this permits to speculate that equilibrium may be achieved earlier than reported. The long reaction times bring economic constraints related to the increase in operational and labor costs, as well as a reduction in the productivity of the process. The lack of explicit information about the initial quantities of reagents used and/or the volume of the reaction media impairs a reliable calculation of concentrations of products and biocatalyst productivity. An interesting study carried out by Kovalenko et al. (2021) [312] synthesized many short-chain esters using in-house immobilized lipases, but this study did not give specific details on the yields for each product. A proper investigation of economic constraints on this topic depends on these measures. However, it is important to highlight that many studies described in Table 1 and Table 2 delve into exploring some scientific aspects, independently from a potential scaling-up and the development of a biocatalytic process.

The importance of a proper exploration of the economic aspects of biocatalytic production relies on the environmental advantages that can be obtained from enzymatic technology for ester production. Thum and Oxenbøll (2008) [9] present a comparison between esters obtained in chemical and biocatalytic routes from an environmental perspective—energy savings, reduction in greenhouse gas emissions, reduction in the generated wastewater, non-hazardous production (in the case of solvent-free systems), among other important findings. These environmental advantages associated with higher purity of obtained products, due to the high specificity of immobilized lipases as catalysts and mild reaction conditions, may lead to products with differentiated prices that reach specific high added-value markets. Furthermore, these advantages may be the drivers pushing chemical industries to increase their sustainability indicators. Biocatalytic routes to produce important compounds such as short-chain esters may have a range of positive market appeals as clean and sustainable production (related to the Sustainable Development Goals from the United Nations), Green Chemistry, Bioeconomy, and energy transition. In summary, it is worth further exploring the enzymatic short-chain ester syntheses aiming to overcome the technical and economic drawbacks.

2.7. Solvents and Solvent-Free Systems on Enzymatic Esterification Studies

The use of hydrophobic solvents is disseminated in most of the studies, particularly hydrocarbons. Hydrocarbons are frequently miscible with low concentrations of acids and alcohols and are highly volatile, favoring the purification of the products. Another important feature of the solvents in esterification reactions is to promote an increase in the pK_a of the acids, and only non-ionized forms are valid for the chemical equilibrium of esterification reactions [17,313,314]. On the other hand, hydrocarbon solvents are toxic and flammable bringing safety and health concerns, in addition to other disadvantages associated with the key performance indicators (KPI), such as the decreased concentration of the product and productivity. n-Hexane, cyclohexane, n-heptane, or toluene, are considered problematic or hazardous compounds by the CHEM21 selection guide of solvents [75]. When solvents are indispensable to perform a reaction, green solvents should be the selected alternative. Green solvents are characterized by their low environmental and safety impact during production, use, and final destination [315,316]. In this context, green solvents are compounds that conciliate solvent technical features (stability, inertness, viscosity, volatility) with a green/renewable obtaining process, non-toxicity, and biodegradability. Green solvents such as ionic liquids, deep-eutectic solvents, supercritical CO₂, and other bio-based compounds have been intensively studied in the past years [159,314,315,316]. A survey on the Web of Science database revealed 1084 scientific papers with “green solvent” as a keyword in the past 5 years. Furthermore, some short-chain esters like ethyl acetate, isopropyl acetate, butyl acetate, and amyl acetate, are also considered green solvents in CHEM21 classification [75]. Other esters, such as lactones, seem promising solvent alternatives for biocatalytic oxidation reactions [317]. However, we have found few enzymatic short-chain esters studies that use green solvents [254,273].

Even using solvents, a significant part of the studies reached the highest carboxylic acid transformation to ester when using a stoichiometric excess of the alcohol higher than 3, as may be seen in Table 1 and Table 2. This excess of alcohol also leads to low concentrations of products. Moreover, in some cases, the high stoichiometric excesses of alcohol may suggest that there is a combination of the solvent and the surplus alcohol. For example, Baek and co-workers (2020) [264] found that phenomenon when 0.9 mols L⁻¹ of n-octanol was used utilizing 1,2-dichloroethane as solvent. This solvent is only moderately hydrophobic (logP 1.48), but the overall hydrophobicity can be improved by the presence of the surplus n-octanol (logP 3.0).

In solvent-free systems (SFSs), there is a great potential increase in product concentration, as demonstrated by Musa et al. (2018) [296], even if lower conversions than in the presence of solvent are achieved (47.0% versus 69.0%) or by the study of Salah and co-workers (2007) [203], as can be seen in Table 1. Other solvent-free syntheses also obtained concentrations of products higher than 100 g L⁻¹ [37,178,275,277,291,298,299]. The syntheses carried out in SFSs permit different strategies to improve the process. The most obvious is the adoption of stoichiometric excesses of alcohol, taking as an example the study of Lorenzo et al. (2022) [21] in the synthesis of isopentyl acetate, and Dai et al. (2014) [275] in the synthesis of butyl propionate. Isopentanol and n-butanol, in these cases, act simultaneously as reagents and solvents. However, in this kind of approach, two important consequences should be evaluated—the formed water (if not removed from the media) is miscible with the surplus alcohol, and the surplus alcohol will increase the probability of enzyme inhibition. The occurrence of inhibition by excess alcohol will cause an increase in the reaction time, demanding high biocatalyst loadings to compensate for the reduction in reaction rates. This effect has a drastic impact on the productivity of the syntheses, as can be observed in some studies [37,175,275].

Some studies also consider the addition of water in the reaction media, in different proportions, as well as keeping the produced water in the reaction media. At first glance, these approaches seem counterintuitive considering that water accumulation will favor hydrolysis. However, as highly hydrophilic reagents may cause distortions in the hydration layer of lipases, the addition of water may compensate for these deleterious effects [17,37]. Thus, the presence of water causes simultaneously opposite impacts—a positive one, in kinetic aspects, and a negative one, from a thermodynamic perspective [18]. Another possible negative impact, discussed by Krishna and Karanth (2001) [318] and other authors is the deactivation (partial or total) of the lipases due to the acidification of the microaqueous environment of the enzyme, considering that short-chain acids have relatively low pK_a [17]. The impact of the addition of water on the yield of esters varies greatly, as can be seen in Table 1 [37,61,178,183,203,292]. There is a great dependence on the amount of water, the reaction media, and the immobilized lipase studied, considering that each immobilized lipase has an optimum water activity [111].

2.8. The Use of Ultrasound and Microwave on Enzymatic Esterification Studies

One way to improve solvent-free short-chain ester production is the use of microwaves or ultrasound [276]. As both technologies enhance the reaction rates, and the last one prevents the formation of a water layer inside the biocatalyst particles, the effects of reagents inhibition (or even partial inactivation) caused by the short-chain reagents on reaction rates become less pronounced [261,277]. These technologies, despite intense lab-scale exploration, are still far from application in the industrial context.

The use of ultrasound promotes a very effective agitation, even inside the particles (that way preventing the hydrophilic phase formation in the enzyme environment) and improve the mixing of phases, which is crucial to avoid any mass transfer limitations when immobilized enzymes are present [1,261]. Microwaves, on the other hand, act under kinetic reaction conditions due to the orientation of polar functional groups of the reagents and improve enzyme catalytic activity and stability [1,290]. Ultrasonically assisted synthesis, also known as sonochemistry, is an emerging technology that has been employed in the production of esters. This technique is based on the generation of high-frequency sound waves (>16 kHz) in a liquid medium. These sound waves induce vibrations that facilitate the movement of reagent molecules, thereby increasing interactions among them. Additionally, ultrasound can trigger cavitation, a phenomenon that creates localized supercritical conditions characterized by high temperatures and pressures [1,261]. Cavitation occurs when high-frequency sound waves generate vapor bubbles within the liquid. These bubbles grow and collapse rapidly, generating shear forces and impact that drive the progression of the enzymatic reaction and improve the solubility of the reagents. The intensity of ultrasound is a crucial factor that affects the efficacy of ester synthesis through this technique. Excessively high intensities can damage the enzyme structure, while excessively low intensities may not induce sufficient cavitation [277]. Several experimental studies have confirmed the effectiveness of ultrasound to improve ester synthesis, and here the work of Jaiswal and Rathod (2018) [277] is a successful example of its use, obtaining high ester concentration and productivity in the synthesis of isobutyl propionate in SFSs.

The other alternative to improve the reaction is the use of microwave irradiation. It covers a wide range of frequencies and wavelengths (1 mm to 1 m and 300 GHz to 300 MHz) [319]. This technology involves essential components such as the wave source, transmission lines, and applicator [297]. The effectiveness of microwave irradiation heating depends on the dielectric properties of the material, penetration depth, frequency, and mode of operation (monomode or multimode) [320,321,322]. Monomode reactors accurately direct radiation to the sample, maximizing energy efficiency, while multimode reactors distribute radiation chaotically, resulting in non-uniform heating. In addition to thermal effects, the scientific literature debates the possibility of non-thermal effects related to the direct interaction of the electric field with specific molecules [320]. Microwave technology offers several advantages, including ease of use, energy savings, reduced processing time, and a lower alcohol ratio to obtain similar yields in ester production. Some studies using conventional catalysts, such as that of Choedkiatsahul et al. (2015) [323], demonstrated a 99.4% conversion in just 1.75 min at 70 °C with the addition of 1% NaOH when using microwaves in the production of esters from palm oil. Similarly, Yu et al. (2010) [324] carried out a noteworthy study on the synthesis of methyl esters of fatty acids using microwaves, achieving a 94% conversion in 12 h, in contrast to the 24 h required in the conventional method. In the field of biocatalytic synthesis, where challenges like low reaction rates, long reaction periods, and low conversions are common, the application of microwaves stands out as an efficient solution. Microwave irradiation provides precise heating, substantially increasing the reaction rate and significantly reducing synthesis reaction time [325]. Furthermore, microwaves have a greater impact on molecules with higher dipole moments, such as lipases and substrates used in enzymatic synthesis with polar properties, making the process more effective by facilitating interaction between the substrate and the biocatalyst [325]. Some notable examples of enzymatic short-chain ester syntheses using microwaves are described in Table 1, in the works of Cebrian-Garcia (2018) [291] and Bhavsar and Yadav (2018) [276].

2.9. Biocatalysts on Enzymatic Esterifications and Reusability Studies

Different biocatalysts were used in the studies. Novozym^® 435, a widely studied commercial lipase from Novozymes^®, was used in many studies as can be seen in Table 1 and Table 2, even with the problems this biocatalyst presents. Its high catalytic activity and stability in different chemical environments and temperatures are crucial for synthetic applications; the main features, advantages, and disadvantages of Novozym^® 435 were in-depth reviewed in the work of Ortiz and co-authors (2019) [5]. Novozym^® 435 permits to reach high ester yields for the syntheses of octyl formate, hexyl formate, phenethyl formate [264,297], butyl propionate [275,276], butyl butyrate [278], and 3-hexen-1-yl acetate [274].

Novozym^® 435 shows better performance than other biocatalysts in ester syntheses, including comparisons with other immobilized forms of lipase B of Candida antarctica [12,42,183,276]. This is also the case for short-chain ester syntheses, as shown in Table 1 [182,183,276]. One of the main reasons is its stability and high catalytic activity in more hydrophilic and acidic environments. Hydrophobic solvents are generally adopted for short-chain ester syntheses, as already discussed. Still, Novozym^® 435 shows good performance in the presence of hydrophilic solvents, as demonstrated in the work of Hasegawa and co-authors (2008) in the synthesis of ethyl lactate in 1,4-dioxane [326]. Moreover, lactic acid is a relatively strong organic acid (pK_a equal to 3.86, lower than acetic acid, 4.76). Novozym^® 435 stability in such environments allows solvent-free short-chain ester syntheses [15,21,268,274,275,276,327].

The main limitation of Novozym^® 435 is relative to its immobilization support. This support is poly(methyl methacrylate) cross-linked with divinylbenzene and commercially denominated Lewatit^® VP OC 1600. Lewatit^® VP OC 1600 has a surface area of 130 m² g⁻¹, an average particle size of 315–1000 μm and pore diameter of 150 Å, (product information Lewatit^® VP OC 1600, Lanxess, edition: 13 October 2011). The support is moderately hydrophobic and permits enzyme immobilization via interfacial activation, with the advantages previously described. However, its only moderate hydrophobicity makes relatively simple the release of the enzyme during operation [328,329,330,331,332,333]. Furthermore, it facilitates the retention of water and hydrophilic substrates in the enzyme environment, which as stated above is a major reason for the enzyme inactivation in esterification processes [59,252,260,266,334,335]. The support presents relatively high mechanical fragility that causes problems for the industrial operation of the biocatalysts [5]. An even more serious problem is that the cross-linking seems to be not fully efficient, and the support can be dissolved in certain solvents, causing problems not only related to the biocatalyst operation stability, but also to the downstream [336,337]. Even with these problems, this biocatalyst remains the most used lipase biocatalyst in academic literature.

As a significant part of the studies are focused on the immobilization of lipases, and the short-chain ester synthesis is just one test to characterize the biocatalyst features, a variety of lipases and immobilization support materials are observed in Table 1 and Table 2. In short-chain ester syntheses there are hydrophilic environments and the formation of water; thus, highly hydrophobic support tends to have better performance. Highly hydrophilic supports tend to favor the water phase formation in the lipase environment, favoring the hydrolysis of the formed ester if water is not removed/absorbed, as in the case of ethyl hexanoate synthesis mediated by lipase immobilized onto chitosan [296]. Another possible effect of the support nature is the dissolution of highly hydrophilic support (and resultant enzyme release) in the reaction media if high temperatures are present [5,154]. A recent article from Bolivar and co-authors (2022) [154] argues that immobilization is not yet a mature discipline due to some misconceptions in the development of the support, the immobilization protocol adopted, and its application in a specific reaction media/reactor.

Another possible approach in enzymatic ester synthesis is the utilization of aggregated lipases (e.g., lyophilized or precipitated) as a method for enzyme immobilization. The use of powder enzymes is one of the oldest “immobilization” techniques, but it only works under conditions where enzyme solubilization is not possible, for example, in organic solvents, as the enzyme is not soluble in these media. As this immobilized lipase form is not supported in any material, this represents a lower cost in biocatalyst preparation [111,299]. Generally, these biocatalysts may present low enzymatic activity, but strategies such as salt-induced activation may improve their activity [111,338]. Meneses and co-authors found that lyophilized CALB had similar performance (>90% conversion) than Novozym^® 435 in the synthesis of benzyl propionate in a fed-batch system [298]; methyl benzoate was synthesized by Leszczak and Tran-Minh using lyophilized Candida rugosa lipase suspended in hexane-toluene [339]; ethyl butyrate, ethyl hexanoate, and ethyl caprylate were also synthesized by lyophilized Rhizopus chinensis lipase at 40 °C achieving conversions close to or higher than 90% after 20 h in the work of Sun and co-workers (2009) [340].

The biocatalyst loadings used for short-chain ester syntheses vary greatly between the different studies, including the way it is represented in the different publications (concentration, mass, activity, etc.), making their comparison a complex task in many instances. In this review, we have expressed the biocatalyst loading as the total mass (enzyme + immobilization support) for practical reasons; really, the total biocatalyst load influences the reactor and processes design. Another alternative, perhaps reflecting better the enzyme performance, would have been the use of the used amount of enzymes, but that was impossible to calculate in most of the publications, mainly if they used commercial biocatalysts. It was possible to observe high biocatalyst loadings in many studies (superior to 5% w/w or equivalent measures) from Table 1 and Table 2. High loadings are required in most cases due to accentuated inhibition and partial deactivation effects of short-chain reagents on the performance of immobilized lipases, and this brings two obstacles for scaling up of a process: (i) additional costs, and (ii) operational issues due to mixing and effective mass transfer in continuous stirred tank reactors or fluidized reactors [12,341]. Immobilized lipases are still a high-cost income and thus, high loadings mean a drastic increase in the operational costs of the process [8,12,66]. Furthermore, there is no proportional correlation between the increase in biocatalyst loading and reusability [12]. In terms of agitation, high loadings of catalysts tend to decrease the operational stability of immobilization supports, due to the potential increase in high-energy collisions between biocatalyst particles [5,12,37,342]. This leads to a decrease in the reusability of immobilized lipases with consequential impacts on operational costs. Moreover, if a high percentage of the reactor volume is the biocatalyst particles, a large amount of the final reaction mixture will be retained inside the particles and between particles, requiring extensive washing to recover it.

One of the advantages of the biocatalytic route using immobilized lipases is the possibility of reuse of the biocatalyst, due to the easy removal of a solid catalyst from a liquid reaction media. The maximum number of sequential cycles of reaction for a given biocatalyst has the utmost importance for the determination of the production cost per batch. Reusability is affected by the nature of the immobilization support, its resistance (chemical and mechanical), and the severity of the reaction media, which includes temperature, reaction time, and agitation intensity [5,12,159]. Few studies dedicated an in-depth evaluation of this crucial aspect. Generally, the topic is limited to checking how many reaction cycles (in optimized conditions) it is possible to carry out without significant loss of enzymatic activity. The number of reuses is frequently inferior to 10 cycles using different immobilized lipases in most studies, as shown in

Table 3. Reusability of different biocatalysts in short-chain ester syntheses.

Ester	Biocatalyst	Number of Sequential Reactions	Ref.
Butyl Propionate	Novozym® 435	20	[275]
Hexyl Formate	Novozym® 435	20	[265]
Ethyl Butyrate	Candida rugosa lipase immobilized into polyurethane foams	20	[278]
Butyl Acetate	Novozym® 435	14	[259]
Octyl Formate	Novozym® 435	10	[264]
Pentyl Acetate	Lipozyme® 435	10	[21]
Pentyl Valerate	Candida rugosa lipase immobilized onto microemulsion-based organogels	10	[109]
Ethyl Butyrate	Candida antarctica lipase B immobilized onto styrene beads	8	[183]
Isobutyl Propionate	Fermase® CALB 10000	7	[277]
Isobutyl Propionate	Novozym® 435	7	[37]
Butyl Acetate	Candida antarctica lipase B immobilized onto porous styrene-divinylbenzene beads	6	[182]
Butyl Butyrate	Thermomyces lanuginosus lipase immobilized onto glyoxyl-agarose beads	5	[194]
Butyl Acetate	Novozym® 435	5	[256]
Ethyl Valerate	Candida rugosa lipase immobilized onto amine polypropylene membrane	5	[178]
Ethyl Valerate	Aspergillus oryzae immobilized onto an electrospun nanofibrous membrane	5	[292]
Isopentyl Butyrate	Rhizopus oryzae lipase immobilized onto Purolite (PMMA with epoxide and octadecyl groups)	5	[108]
Isobutyl Butyrate	Lipozyme® RM-IM	4	[259]
Isopentyl Acetate	Staphylococcus simulans lipase immobilized onto CaCO3	4	[61]
Isopentyl Propionate	Novozym® 435	4	[254]
Butyl Propionate	Novozym® 435	3	[276]
Butyl Acetate	Rhyzopus oryzae lipase immobilized onto Celite 545	3	[210]
Ethyl Valerate	Novozym® 435	2	[290]
Isopentyl Acetate	Lipozyme® IM-20	2	[266]

.

There are a few exceptions to this general lack of long-time operational biocatalyst stability, such as the studies of Dai et al. [25], Seo et al. [267], Pires-Cabral et al. [278], and Martins et al. [259]. This means that, in general, we lack information on the immobilization support materials‘ behavior when submitted to standard mechanical/orbital agitation in the presence of organic solvents for long reaction times.

3. Conclusions

Short-chain esters have a huge relevance in the chemical industry context and technologies for their current production are well established. However, the market sizes and the different sectors involved (particularly the ones that deal with daily products) may be a driver for the gradual implementation of new technologies that reduce environmental impacts and greenhouse gas emissions. The scientific community has been proposing alternatives to short-chain esters production in a greener and more sustainable way. The adoption of immobilized lipases as biocatalysts in these processes may be a very suitable solution from these points of view, but it has technical and economic obstacles to be overcome, particularly the necessity of using organic solvents and the high inhibition potential of the substrates on the lipases, which lead to unfavorable process performance indicators. Further developments in enzymatic immobilization, the understanding of thermodynamics and kinetics aspects of the esterification processes, and the establishment of effective reaction setups are crucial to obtaining cost-effective and efficient biocatalysts and the full conversion of the substrates into products at high rates. Preliminary techno-economic analyses are scarce in the literature for biocatalytic ester production, and this information is of utmost importance to establish reliable quantitative metrics on the lab-scale to be used in the future scaling-up of the processes. This gap of information should be filled to take advantage of the high positive environmental indicators that biocatalytic routes present when compared to conventional chemical routes.