Sources, purification, immobilization and industrial applications of microbial lipases: An overview

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Sources, purification, immobilization and industrial applications of microbial lipases: An overview

Enespa, Prem Chandra & Devendra Pratap Singh To cite this article: Enespa, Prem Chandra & Devendra Pratap Singh (2022): Sources, purification, immobilization and industrial applications of microbial lipases: An overview, Critical

Reviews in Food Science and Nutrition, DOI: 10.1080/10408398.2022.2038076

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Sources, purification, immobilization and industrial applications of microbial lipases: An overview

Enespaa , Prem Chandrab and Devendra Pratap Singhc aSchool for Agriculture, Sri Mahesh Prasad Post Graduate College, University of Lucknow, Lucknow, Uttar Pradesh, India; bFood Microbiology

& Toxicology Laboratory, Department of Microbiology, School for Environmental Sciences, Babasaheb Bhimrao Ambedkar University (A

Central) University, Lucknow, Uttar Pradesh, India; cDepartment of Environmental Science, School for Environmental Sciences, Babasaheb

Bhimrao Ambedkar University (A Central) University, Lucknow, Uttar Pradesh, India

ABSTRACT Microbial lipase is looking for better attention with the fast growth of enzyme proficiency and other benefits like easy, cost-effective, and reliable manufacturing. Immobilized enzymes can be used repetitively and are incapable to catalyze the reactions in the system continuously. Hydrophobic supports are utilized to immobilize enzymes when the ionic strength is low. This approach allows for the immobilization, purification, stability, and hyperactivation of lipases in a single step. The diffusion of the substrate is more advantageous on hydrophobic supports than on hydrophilic supports in the carrier. These approaches are critical to the immobilization performance of the enzyme. For enzyme immobilization, synthesis provides a higher pH value as well as greater heat stability. Using a mixture of immobilization methods, the binding force between enzymes and the support rises, reducing enzyme leakage. Lipase adsorption produces interfacial activation when it is immobilized on hydrophobic support. As a result, in the immobilization process, this procedure is primarily used for a variety of industrial applications. Microbial sources, immobilization techniques, and industrial applications in the fields of food, flavor, detergent, paper and pulp, pharmaceuticals, biodiesel, derivatives of esters and amino groups, agrochemicals, biosensor applications, cosmetics, perfumery, and bioremediation are all discussed in this review.

Introduction Lipases (triacylglycerol acylhydrolase, EC 3.1.1.3) are hydro- lases that operate on carboxylic ester linkages (Kurtovic et al. 2009). Triglycerides are hydrolyzed naturally by lipases into mono and di-glycerides, fatty acids, and glycerol (Houde, Kademi, and Leblanc 2004; Javed et al. 2018). It is known as a particular enzyme since it has a unique method of action called interfacial activation (Gonçalves, Silva, and

Guidini 2019). In homogeneous conditions, most lipase mol- ecules contain a polypeptide chain termed a lid covering their active center, which can separate it from the reaction medium (Yaacob et al. 2019). Fixing a new structure in the presence of a hydrophobic surface where the enzyme becomes adsorbed and the active center is fully exposed, allowing lipases to hydrolyze oil drops (Palomo et al. 2004).

This approach was utilized to selectively immobilize lipases by their open forms on a variety of hydrophobic supports (Sakai et  al. 2010; Bezerra et  al. 2017; Klein et  al. 1997).

Lipase molecules are stabilized in their open and active forms on the support surface due to high adsorption (Alnoch et  al. 2018). The anhydrous form of organic media used to accelerate biotransformation with methacrylate-like hydro- phobic supports and lipase derivatives are solvents and solvent-free systems (Fernandez-Lorente, Rocha-Martín, and

Guisan 2020).

The solubility of water enzymes was explained using immobilized techniques (Sheldon 2007; Zdarta et  al. 2018).

An encapsulation step is a valuable approach to improving enzyme functionality on an industrial scale. Immobilization occurs after other enzyme properties such as immobility, activity, specificity, selectivity, and resilience to inhibitors and chemical reagents have been developed (Bilal et  al.

2019; Fernandez-Lafuente 2009; Garcia-Galan et  al. 2011;

Iyer and Ananthanarayan 2008; Mateo et al. 2007; Rodrigues and Ayub 2011). Immobilization can be used in conjunction with any other technique for enhancing enzyme character- istics. As a result, enzyme immobilization has become a prominent step in the creation of a commercial enzyme biocatalyst. If appropriately engineered, it can be used to purify the enzyme throughout the immobilization process (Barbosa et al. 2011). Physical and chemical techniques are preferred in industries for enzyme immobilization because they are easy and cost-effective (Mohamed et  al. 2021). It’s critical to understand the drawbacks and benefits of immo- bilized enzymes (Table 1). Amylases and proteases are dras- tically reduced when immobilized due to diffusion constraints, and immobilized enzymes are less economical

© 2022 Taylor & Francis Group, LLC CONTACT Prem Chandra p.chandrabbau@gmail.com https://doi.org/10.1080/10408398.2022.2038076

KEYWORDS Enzyme-modified cheese; food industry; immobilization techniques;

Microbial lipases; pharmaceuticals; phospholipase 2

E. P. CHANDRA AND D. P. SINGH due to the quick kinetics of native enzymes (Basso and

Serban 2019).

The presence of lipase observed in bacterial species is

Bacillus prodigiosus, B. pyocyaneus, B. fluorescens, and

Staphylococcus pyogenesaucreus (Salwoom et  al. 2019;

Mobarak-Qamsari, Kasra-Kermanshahi, and Moosavi-Nejad

2011), and the other significant species are Bacillus,

Pseudomonas, Burkholderia; and fungal species are Aspergillus, Penicillium, Rhizopus, Candida, Hypocrea pseu- dokoningii and the species of yeasts are Zygosaccharomyces,

Pichia, Lachancea, Kluyveromyces, Saccharomyces, Candida, and Torulaspora, detected to harvest lipase enzyme (Pereira et  al. 2014; Bora, Gohain, and Das 2013; Alcazar-Valle et  al. 2019).

Microbial lipases are highly favored in the enzyme mar- ket, accounting for about 90% of the global lipase market from bacteria and fungi (Chandra et  al. 2020b). With the aggregate usage of lipase in several feeds for livestock in the worldwide market, the global lipase market was valued at USD 585.56 million in 2020 and is expected to reach

USD 961.85 million by 2028, rising at a CAGR of 6.4 per- cent from 2021 to 2028. In the animal feed and dairy indus- tries, there is an increasing demand for lipases to improve the value of meat and treated dairy commodities (Guerrand

2017). Microbial lipases are more reliable than vegetable and animal enzymes, and because of their high consistency, optimization, and process modification, they can be easily and cost-effectively produced through fermentation proce- dures with less time and space requirements (Andualema and Gessesse 2012; Raveendran et  al. 2018; Rosenthal and

Lütz 2018). It aids in the refinement of food value and improves the taste, texture, shelf life, and smoothness of some products in the food sector as a nutrient (Barrett,

Beaulieu, and Shewfelt 2010). Lipases have been employed on a huge scale to improve the health fitness of numerous animal feeds on the global market, and are mostly used in animal feeds, bakery, dairy, and confectionery sectors.

Because of their numerous applications in a wide range of food processing from microorganisms, significant advance- ment is envisaged soon (Amit et  al. 2017). The advantages of microbial lipases over animal and plant lipases are also supporting market growth (Bilal et  al. 2021). Other actions covered involve interesterification, esterification, aminolysis, and alcoholysis, as well as relevant industries (Lee et  al.

2001; Lima et  al. 2017; Ismail and Baek 2020). Lipase pro- duces esters from glycerol and long-chain fatty acids in a non-aqueous media. Flavor enhancement lipases are con- sidered a potential biocatalyst in a variety of biotechnological processes, particularly in dairy products such as cheese recovery (Dandavate, Keharia, and Madamwar 2011).

Increased consumption of enzyme-modified cheese (EMC) and enzyme-modified dairy ingredients (EMDI) as a result of a growing perception of animal strength and value has boosted the lipase market tremendously (Carneiro et  al.

2020). In oil chemistry, bio-detergents are manufactured to remove harsh chlorine bleach from fresh diminishing sewage and industrial pollution (Fukunaga et  al. 1998). Ibuprofen, naproxen, pregabalin, and ketoprofen, as well as (S)-Prosimpal, (S)-Piperoxan, (S, S)-Dibozane, and (S)-Doxazosin, are all manufactured in the pharmaceutical industry. RAC -ibuprofen (R, S)-2-(isobutylphenyl)-propionic acid) is a racemic mixture of ibuprofen (R, S)-2- (isobutylphenyl)-propionic acid (Adrio and Demain 2014;

Sanchez and Demain 2011). The usage of recombinant DNA technology is a very appealing characteristic that can be exploited to overcome the economic constraints of industrial lipase use (Kanmani, Aravind, et al. 2015). Novozymes intro- duced Lipolase, the first lipase made with recombinant DNA technology, to the market in 1988. This lipase was developed in Aspergillus oryzae and came from Thermomyces lanugi- nosus (Gerits et  al. 2014; Sarmah et  al. 2018; Boel et  al.

1988; Lin et al. 2016). Lipophilic extractives include alkanes, fatty alcohols, resin acids, fatty acids, conjugated sterols, certain terpenoids, waxes, and triglycerides, which are non-polar extractable fractions from wood and other ligno- cellulosic materials known as wood resin (Gutiérrez, Río, and Martínez 2009). Removing the esters from pulp lipase could be employed to improve manufacturing capacity and quality (Horchani et  al., 2012).

Palatase from Rhizomucor miehei (Handayani et al. 2016), a commercial lipase articulated in A. oryzae, was another commercial lipase articulated in A. oryzae (Ansorena et  al.

1998; Sankaran, Show, and Chang 2016). Because of their excellent wetting properties, wax esters are widely used in cosmetics, medicines, greases, and other biochemical prod- ucts (Zalacain et al. 1997; Zhang, Aryee, and Simpson 2020).

Immobilization, manufacturing, purification, categorization, and application of microbial lipases from a variety of sources were highlighted in this review. Immobilized enzymes include a variety of commercial lipases, as well as their immobilization methods.

Explanation of lipases Lipases naturally hydrolyze mono, di, triglycerides, fatty acids, and glycerol into mono, di, triglycerides, fatty acids, and glycerol (Schomburg, Chang, and Schomburg 2014;

Chahinian and Sarda 2009). Carboxylic esters linkages can

Table 1.  Advantages and disadvantages of immobilized enzymes in industrial processes.

Advantages Disadvantages • Easy separation of biocatalyst

• Compared to the native enzyme it has a slow enzyme activity

• In downstream processing costs reduced • For carriers and immobilization, additional costs required

• Several uses of biocatalyst (reutilizing) • Compared to native enzymes reaction rates slow

• Toward organic solvents and higher temperatures, it shows better stability

• Subject to fouling • Without the need of membrane to isolate enzyme from product use of the fixed bed or batch reactors

• Exhausted immobilized enzyme incinerated (disposed)

• With other enzymes co-immobilization is possible

Critical Reviews in Food Science and Nutrition 3 be degraded by enzymes other than lipases and esterases (Levisson, Oost, and Kengen 2009). For a long time, the distinction between lipases and esterases has been based on interfacial activation and the presence of a lid for the former enzyme (Schmid and Verger 1998; Brocca et  al.

2003). Lipase activity rises rapidly in the presence of an interface, which is known as interfacial activation, whereas an amphiphilic surface loop known as cover shields the active site of lipase in solution and travels away with the interface when the two come into contact (Eggert et  al.

2004). Despite the presence of a lid, Pseudomonas aerugi- nosa, Candida anatarctica B (Theil and Björkling 1993), and Burkholderia glumae lipases do not show interfacial activation (Li and Zhang 2016; Hotta et  al. 2002; Kovacic et al. 2019). The presence of a lid and interfacial activation can simply be explained as a carboxylesterase that catalyzes the hydrolysis and production of long-chain acylglycerols and cannot be classified as a real lipase (Lopes et al. 2011;

Chamorro et  al. 1998).

Extracellular lipase Extracellular lipases were created by microbes through solid-state or submerged fermentation (Sales et  al. 2020).

An enzyme’s biocatalytic activity is frequently regained by raising the degree of purity in the fermentation process and then purifying it (Borkar et  al. 2009). Extracellular lipase production is a complex process regulated by the origin and structure of the lipase (Palekar, Vasudevan, and Yan 2000;

Doolittle and Péterfy 2010). Extracellular lipases should be cost-effective, rapid, simple, and efficient when used in large-scale production (Ventura and Coutinho 2016;

Robinson 2015; Soleymani et al. 2017). In the vast majority of cases, extracellular immobilized lipases are commercially accessible (Ingenbosch et  al. 2019; Robles-Medina et  al.

2009). Candida antarctica, Rhizomucor miehei, and Thermomyces lanuginosus were used to identify and purify the immobilized lipases Lipozyme RM IM and Lipozyme

TL IM, Novozyme 435 (Hernández-Martín and Otero 2008; de Souza et  al. 2018; Bueso et  al. 2015; Kobayashi 2011;

López-Serrano et  al. 2002; Wang et  al. 2014). That is, the support may first immobilize through a one-point or mul- tipoint interaction (Sunitha et  al. 2007; Seitz 1974), and then, as with heterofunctional supports, it may continue to increase the number (or even the quality) of interactions, including new groups (Muralidhar et  al. 2002; Ban et  al. 2002).

Intracellular lipase.  The utilization of entire cells as biocatalysts is an alternate way for resolving the high cost of extracellular lipase purification. Intracellular lipase refers to the use of lipase that has been liberated from cells (Schoemaker, Mink, and Wubbolts 2003; Adamczak and Bednarski 2004). Intracellular lipase is lipase that is secreted from the cell’s outer surface. Because of certain supports, certain microorganisms can naturally immobilize castoff as a lipase source (Uthoff, Bröker, and

Steinbüchel 2009). It eliminates the requirement for an

Figure 1.  Reactions catalysis using lipase.

4 E. P. CHANDRA AND D. P. SINGH expensive purification step as well as the necessity for a lengthy immobilization technique, which is required with extracellular lipase (Ferreira-Dias et al. 2013; Adlercreutz

2013).

Reactions catalyzed by lipases Lipase catalyzes the hydrolysis of carboxylate ester bonds at the organic-aqueous interface in the presence of excess water, releasing organic alcohols and free fatty acids (FFAs) (El

Seoud, Baader, and Bastos 2016; Schoffelen and Hest 2013).

The equilibrium was evaluated under the limitation of the reverse reaction using the water activity (aw) of the reaction mixture among the forward and reverse reactions (Lewis et  al. 1965). Under low water activity, a variety of transes- terification processes can be carried out (aw) (Bovara et al.

1993). An ester and alcohol (alcoholysis), an ester and acid (acidolysis), an ester and amine (aminolysis), or an ester and two esters (interesterification) undergo transesterifica- tion (Figure 1) (Santilli et  al. 1987).

Lipases have numerous types of selectivity for their sub- strates, including chemo-specificity: which shows both fatty acid (FA) and lipid-class specificity when utilizing lipases (Raclot, Holm, and Langin 2001; Albayati et  al. 2020). FA specificity is connected to the length or degree of unsaturation that declines within specific kinds of FFAs (Lennen and

Pfleger 2012; Barros, Fleuri, and Macedo 2010). For lipid-class specificity, lipases catalyze the hydrolysis of mono, di, and triacylglycerol; (ii) Regio-specificity: General lipases catalyze the entire hydrolysis of TAGs into glycerol and FFAs in an unplanned manner, producing DAGs and MAGs as interme- diates (Chang and Lee 2021). Using 1, 3-specific lipases, the

TAGs were hydrolyzed solely at the sn-1 and sn-3 locations, yielding FFAs, 1, 2-DAGs or 2, 3-DAGs, and 2-MAGs (Frayn et al. 2003; Ishchenko et  al. 2017). Because 2, 3-DAGs, and

2-MAGs are highly unstable, they undergo acyl migration to create 1, 3-DAGs and 1-MAGs or 3-MAGs, respectively, (iii)

Enantio-selectivity: Lipases can tell the difference between the two enantiomers in a racemic combination (Jaeger and

Reetz 1998). This selectivity fluctuates with admiration for the substrate, as it is related to the nature of esters.

Production and purification of lipases In this section, we discuss the current advancements in lipase manufacture through host strain and metabolic engi- neering processes, as well as the selectivity, stability, and broad substrate specificity of microbial lipases most com- monly employed in industry (Anobom et  al. 2014).

Host strain selection The most capable option for lowering the price of lipases is to use heterologous technologies for the creation of func- tional lipases (Macrae 1983; Baneyx 1999). Several types of microorganisms have been industrialized for heterologous and homologous manifestation in adept lipase host strains since the ancient period summarized in the given Table 2 (Contesini et  al. 2020).

Bacteria E. coli is the most commonly used manifestation host for recombinant protein expression for a variety of reasons. For hereditary employment, E. coli is more adaptable, and for genetic employment, it has higher refurbishment proficiency and faster progress rates (Rosano and Ceccarelli 2014;

Ozturkoglu-Budak et  al. 2016). However, because of a lack of adequate folding machinery, the typical E. coli system leads to the intracellular formation of sluggish or intractable inclusion bodies (Blank et  al. 2006). Candida antarctica (CalB) is well-known in the biocatalysis industry for pro- ducing Lipase B, an active version of the enzyme that can be expressed in E. coli by changing the reaction standard or lipase transformation (Paraskevopoulou and Falcone 2018;

Kundys et  al. 2018; Wu, Yang, and Ge 2017). Lipases fused can increase the solubility of proteins in E. coli containing a polycationic amino acid tag. Furthermore, certain types of lipases require the formation of specific disulfide bonds for helpful proteins to be transported (Liebeton, Zacharias, and Jaeger 2001; de Marco 2009). This issue can be solved by using a certain E. coli Origami (DE3) strain or co-expression of a Dsb-family protein, such as DsbA, which is involved in disulfide bond formation (Urban et  al. 2001;

Vieira Gomes et  al. 2018).

Yeast As expression systems for complicated proteins, yeasts have various advantages, including strong growth capabilities, sanctioning disulfide bond formation, informal genetic manipulation, and post-translational protein processing (Lobstein et  al. 2012). Under the control of methanol-responsive alcohol oxidase promoters, K. phaffii has the capabilities to express and produce lipases. In the extracellular medium it secretes fewer quantities of proteins naturally, but using methanol as an inducer high level of recombinant lipase can be achieved. The lipase purification step is easier and cost-effective makes by these special fea- tures (Cregg et al. 2000). Saccharomyces cerevisiae has been designated as a nonpathogenic host for the synthesis of heterologous lipase for a limited time (Darvishi 2012).

Yarrowia lipolytica effectively transformed Lipase 2 (LIP2) gene into S. cerevisiae using PEX11 promoter (Shockey et al.

2011), and Lip2 lipase (Lip2p) was active in growth aug- mentation (Liu et  al. 2012).

However, the S. cerevisiae expression system sanctions elevated heterologous protein expression, and inherent manipulation has numerous weaknesses like poor plasmid strength, less release ability, striving in scale-up, and hyper-glycosylation. Pichia pastoris used as a host for the production lipases (Passolunghi et al. 2003). It presents some benefits, like a highly-regulated promoter of the alcohol oxidase (AOX) over other hosts (Zhang et al. 2010; Li et al.

2010). It can be grown up to exceedingly more mass of cells in eukaryotic minimal medium and proteasome secretion

Critical Reviews in Food Science and Nutrition 5 Table 2.  Heterologous and homologous production of lipase using the prokaryotic and eukaryotic system.

Enzyme source Enzyme Expression vector (Expression host)

Cloning vector (Recipient strain) Expression vector

Remarks References Acinetobacter haemolyticus Lipase KV1

E. coli BL21 (DE3) pGEM-T Easy (E. coli JM109) pET-30a (+)

Using Response Surface Methods (RSM) optimization of production conditions

Batumalaie et  al. 2018; Fabiano Jares Contesini et  al.

2020.

Bacillus amyloliquefaciens G7 Lipase BaG7Lip E. coli BL21-Star (DE3) p15TV-L (E. coli

BL21-Star (DE3)) p15TV-L Metagenomics libraries construction and using a boolean network analysis prediction of the best-producing conditions. Activity enhancement with acetone, glycerol, and K+ ions

Khan et  al. 2020.

Burkholderia contaminans LTEB11 Lipase LipBC (LipA)

+ foldase LifBC (LipB) E. coli BL21 (DE3) – pET28a (+) and pT7-7 for LipA and LipB, respectively

Lipase and chaperone genes co-expressed. Specific activity of

1426 U/mg (tributyrin); > Over five reaction cycles of 6 h at 45 °C, 80% conversion of ethyl-oleate in n-hexane.

Alnoch et  al. 2018.

Burkholderia stabilis FERMP-21014 Cholesterol esterase + foldase

E. coli BL21 (DE3), E. coli Rosetta (DE3) and B. stabilis

E. coli DH5α and JM109 pET26b(+), pET39b(+), pET40b(+), pBBR122

Promoters screening by RNA-Seq + co-expression of lipase and foldase + heterologous and homologous expression. Recombinant activity was 243 fold higher than the WT without oleic acid; B. stabilis system was more efficient to produce esterase.

Yoshida et  al. 2019.

Burkholderia territorii GP3 Lipase LipBT and foldase LifBT

E. coli DH5α, E. coli DH10β, E. coli BL21 (DE3) pLysS, E. coli

Origami B, E. coli Shuffle B, and E pGEM-T Easy (E. coli

DH5α), pET15b (E. coli DH10β) pGEM-T Easy and pET15b

Metagenomics for screening and Identification of lipolytic strains and meta genomic for screening and best expression system evaluation. Higher lipase activity in E. coli BL21 (DE3) pLysS

Putra et  al. 2019.

. coli SHuffle K (pET15b) (6.73 ± 0.24 U/mg); optimum substrate pNP-C10; activity enhancement in n-hexane, Triton X100, and Ca2+ and Mg2+ ions.

Clostridium acetobutylicum (ATCC 824) Lipase Ca-Est

E. coli BL21 (DE3) pMCSG7 (E. coli DH5α) pMCSG7 Rational design, docking analysis, site-directed mutagenesis. Activity enhancement with methanol; residues Ser89 and His224 are crucial for catalysis.

Nagaroor, V and Gummadi, 2020.

Candida antarctica Lipase CALB Corynebacterium glutamicum

MB001 pEC-CALB and pEC-H36-CALB (E. coli DH5α) pEC-CALB and pEC-H36-CALB

Using 10 mM MgSO4 31% activity was inhibited.

González et  al. 2019 Drosophila melanogaster Lipase Lip3

E. coli BL21 (DE3) - pETMCSIII Directed evolution, error-prone PCR, construction of variant libraries. R7_59A mutant activity was higher than the WT toward tributyrin, glyceryl trioctanoate, coconut oil, glyceryl trioleate, and pNP (pNP-C3, pNP-C8, pNP-C16, pNP-C18) substrates; 57_59A activity enhancement of 228 fold compared to the WT using pNP-C8 substrate.

Alfaro-Chávez et  al.

2019.

Geobacillus zalihae Lipase HT1-5M E. coli BL21 (DE3) pLysS pUC57 and pGEX-4T1 (E. coli TOP10) pLysS

Rational design, molecular dynamics (MD), site-directed mutagenesis. Activity enhancement with Ca2+ ions; more stable in DMSO, n-hexane, and n-heptane with Ca2+ ions

Ishak et  al. 2019.

Proteus sp. NH 2-2 Lipase LipPN1 E. coli BL21 (DE3)

- pET-28a (+) Site-directed mutagenesis. Highest activity toward pNP-butyrate (pH

9.0, 40 °C); activity enhancement with acetone and ions Ca 2+,

Mn 2+ and Mg 2+; rLipPN1 and Shao et  al. 2019. rLipPN1_C85S reached 1662 U/L and 1436 U/L, respectively; 91.5% of soybean oil was converted into biodiesel.

Pseudomonas fluorescens AMS8 G55Y, T52Y and AMS8 recombinant lipases

E. coli BL21 (DE3) - pET32b Rational design, site-directed mutagenesis. G55Y and T52Y lipases had lower affinity by pNP-palmitate, laurate and caprylate substrates compared to WT AMS8 lipase; efficiency of G55Y lipase was higher than T52Y for pNPL and pNPP substrates.

Yaacob et  al. 2019. (Continued) 6 E. P. CHANDRA AND D. P. SINGH

Pseudomonas sp.

LSK25 E Lipase LSK25 E. coli BL21 (DE3) pGEMT Easy (-) pET32b(+)

Activity enhancement with Ca2+ ions; activity boosted in toluene, xylene, n-hexane, n-heptane, and n-hexadecane; higher activities toward long-chain fatty acids contained in coconut oil and rice brain oil, and pNP-C12.

Salwoom et  al. 2019.

Heterologous production of lipases using the eukaryotic system

Marine Streptomyces sp. strain W007 Lipase MAS1 K. phaffii X-33 pPICZαA (E. coli DH5α) pPICZαA

PDI co-expression gives 1.7 fold increases lipase expression. The highest lipase production was achieved at pH 6.0 and 24° C with an activity of 440 U/mL.

Lan et  al. 2016.

Rhizopus chinensis r27RCL K. phaffii GS115 pPIC9K pPIC9K

PDI co-expression gives 1 fold increase. The highest lipase activity reached 355 U/mL with one copy of PDI and five copies of r27RCL gene.

Sha et  al. 2013.

Candida antarctica CALB K. phaffii X-33 and M12 (leu2)

E. coli Stellar™ and XL10-Gold® pBluescript II SK pPGKΔ3PRO_

LIPB Strain with three copies achieved 48.760 U/L enzyme yield, 2.3 fold higher than the one-copy strain.

Robert et  al. 2019.

Candida antarctica CALB K. phaffii GS115 pPICZαB (E. coli

TOP10F’) pPICZαB Combinatorial overexpression of Ydj1p-Ssa1p resulted in the highest fold increase, 2.5. Individual overexpression of Ydj1p, Ssa1p, and

Sec63p increased CALB expression level by 1.6, 1.4, and 1.4 fold, respectively. Co-expression of Ydjlp-Sec63p Kar2p-Ssalp,

Kar2p-Sec63p, resulted in 1.5, 1.5 and 1.5 fold increases, respectively. Kar2p-Ydj1p combination resulted in decreased

CALB secretion.

Samuel et  al. 2013.

Rhizopus oryzae ROL K. phaffii GS115 pPICZα and pAO815 (E. coli Top10 cells) pPICZα-ROL

Strain with five copies resulted in 8 fold increase in ROL expression.

Co-expression of both genes Ubc1 and Hrd1 resulted in.54.2% higher ROL activity, 4750 U/mL.

Jiao et  al. 2018 Fusarium solani FSL K. phaffii X-33 pPICZαA and pGAPZαA (E. coli DH5α) pPICZαA-FSL and pGAPZαA-FSL

Strain expressing pGAPZαA-FSL produced the highest specific lipase activity, 18.81 ± 1.98 U/mg, in 3 days of cultivation time. Lipase activity was enhanced by Mn2+, Ba2+, Li+, Ca2+, Ni2+, CHAPS, and Triton X-100 but was inhibited by Hg2+, Ag+, and SDS.

Wongwatanapaiboon et  al. 2016.

Rhizopus oryzae ROL K. phaffii X-33 – pPICZFLDαROL pFLD gave a 1.9 fold increase compared to pAOX1. The best ROL production strategy with the PFLD-based system is a fed-batch induction phase with a constant sorbitol excess.

Resina et  al. 2005.

Candida antarctica CALB K. phaffi GS115 and Y. lipolytica

RIY368 E. coli K. phaffii (pIB4, promoter pAOX1, HIS4

After 72 h cultivation, the lipase activity was 5540 U/mg dcw for Y. lipolytica strain RIY368 and K

Theron et  al. 2020 marker) and Y. lipolytica (JMP4266, promoter pEYK1-A3B,

URA3 marker) . phaffii strain RIY311 1066 U/mg dcw for Y. lipolytica, during the growth phase the lipase activity increased, whereas during growth and stationary phase in both conditions for P. pastoris it increased

A. niger Lipase lip T. reesei Tu6 strain pBluescript II SK(+) and pMD18-T Simple (E. coli DH5α) pSKpyr4 and pSK-lip

Higher levels of lipase all cph1 gene silencing transformants expressed and less cbh 1 transcript than the reference strain, approximately lower than 2%. The RNAi mediated gene silencing of cbh 1 did not negatively affect the lipase transcript abundance.

Qin et  al. 2012.

Thermomyces dupontii TDL K. phaffii X33 pPICZαA (E. coli Top

10) PICZαA–tdl-opt The highest TDL activity was achieved with pFLD1 (27.076 U/mL), of

15 methanol-inducible promoters, whereas of nine constitutive promoters, pGCW14 gave the highest activity (17.353 U/mL).

Wang et  al. 2019.

Meyerozyma guilliermondii strain SMB L2 lipase Komagataella phaffii

Bacillus  sp.  (pFLDhα) pFLDhα To express L2 lipase under the regulation of PFLD1, A new host-vector system was established as a platform.

Salleh et  al. 2020.

Table 2.  (Continued).

Enzyme source Enzyme Expression Vector (Expression Host)

Cloning Vector (Recipient Strain) Expression Vector

Remarks References Critical Reviews in Food Science and Nutrition

7 at low levels and post-translational alterations of proteins.

It can also secrete heterologous protein hosts with ease (Tokmakov et  al. 2012; Ahmad et  al. 2014). Rhizophus sp. lipase, for example, is not expressed in E. coli due to a lack of necessary proteases to process fungal maturation signals.

However, it can be successfully expressed in the P. pastoris host (Karbalaei, Rezaee, and Farsiani 2020; Contesini et  al.

2020). P. pastoris ability to release heterologous target pro- teins extracellularly with less contaminating proteins is often employed to reduce process costs (Löbs, Schwartz, and

Wheeldon 2017). P. pastoris has been identified as the most capable host for heterologous lipase construction among several yeast strains, particularly from eukaryotic sources (Karbalaei, Rezaee, and Farsiani 2020). Non-conventional yeasts such as Hansenula polymorpha and Yarrowia lipolytica have been proposed (Abdala et  al. 2017; Zhou et  al. 2019).

These strains operate differently depending on the type of heterologous protein used. Due to its increased lipase pro- duction (Madzak, Gaillardin, and Beckerich 2004; Gasmi et al. 2011) and adaptation for CalB synthesis between these non-conventional strains, Y. lipolytica appears to be an added potential replacement host (Valero 2012).

Fungi The big lipase-producing bases include Mucor, Rhizopus,

Geotrichum, Rhizomucor, Aspergillus, and Penicillium (Ayinla et al. 2021). As with yeasts and bacteria, filamentous fungus hosts are assessed as a supplement. Extracellular protein secretion in heterologous hosts filamentous fungi benefit from a variety of factors, including increased plasmid copy number, plasmid stability, and capability (Contesini et  al.

2010; Rantasalo et  al. 2019). Business bids for lipase pro- duction between rehabilitated fungal species frequently involve Aspergillus and Trichoderma sp. (Tamalampudi et al.

2007; Hwang et al. 2004). Like most alert filamentous fungi,

Aspergillus oryzae is the host; CalB with higher esterification activity has been heterologously fashioned by Aspergillus oryzae and immobilized for whole-cell biocatalyst for enzy- matic biofuel generation (Adachi et  al. 2013; Hwang et  al.

2014; Budhwani et  al. 2019). In recent years, Trichoderma reesei has been considered as an alternative host for recom- binant protein production using the cbh1 promoter and has been measured for recombinant lipase production (Qin et  al. 2018).

Host strain engineering Various methods are used to increase the production of massive lipase, which is necessary for the commercial appli- cation of lipases. To maximize lipase productivity, we inves- tigate a variety of techniques and metabolic engineering performances (Ward 2012).

Genetic manipulation of host strains Microorganisms produce a wide range of extracellular lipases, which are then processed into commercial lipases (Fickers, Destain, and Thonart 2009; Gupta, Gupta, and

Rathi 2004). Using the fed-batch fermentation procedure, a significant increase in lipase production has been achieved. Lip2 produced a higher yield of lipase when two distinct orders of Y. lipolytica mutant-strain LgX64.81 were used (Yu, Wen, and Tan 2010; Batumalaie et al. 2018). The most promising techniques to achieving large amounts of purified lipases to fulfill the standard of quantity and manipulation of industrial procedures are replicating and recombinant lipase gene expression (Farrokh, Yakhchali, and Karkhane 2014; Krzeslak et  al. 2008). In most cases, the castoff technique is promoter optimization, which sig- nificantly enhances lipase manufacture (Meunchan et  al.

2015). Strong constitutive marketers like XPR2, TEF, and

RPS7 (Trassaert et  al. 2017) and inducible supporters like

ICL1, POT1, and POX2 (Park, Do, and Jung 2013) have been promoted for Y. lipolytica lipase yield (Liu et al. 2013).

Y. lipolytica was not measured as a perfect mass because it lacked a "perfect" inducible promoter. Heterologous expression of the protein is identified in the E. coli system by the intracellular gathering of sluggish or convoluted inclusion forms (Hellwig et  al. 2004; Xu, Lewis, and Chou

2008). Pseudozyma antarctica lipase B was co-expressed in

E. coli with several periplasmic folding factors, including

DegP, FkpA, DsbA, and DsbC (Narayanan, Khan, and Chou

2010). PalB (inclusion bodies) that are uneven and sluggish can help in the presence of these folding features (Ujiie,

Nakano, and Iwasaki 2016; Dugourd et  al. 1999). As a result, both the cytoplasm and periplasmic material improved significantly in the functional PalB expression.

The ABC transporter protein is naturally employed for a wide range of substrate transformations, including ions, carbohydrates, and amino acids (García, Hoyos, and

Hernáiz 2018; Navarre and Schneewind 1999; Van Geest and Lolkema 2000).

An inner membrane protein with trans membrane seg- ments 6-8 and a C-terminal ATPase domain is composed of an N-terminal membrane area (Ahn, Pan, and Rhee

1999; Jaenicke and Böhm 1998). TliD, TliE, and TliF involv- ing ABC transporters have been removed from E. coli to speed up the release of a thermostable lipase (TliA) (White et  al. 2008). By co-expressing TliA with altered TliD, the secretion levels of TliA lipase were increased thrice while the expression level of transporter protein remained almost unaffected, indicating that designed transporter proteins can speed up the secretion of lipases (Baek et  al. 2010;

Eom et  al. 2005). The process of determining the target proteins related to an attaching theme on the outside of many host cells is known as cell surface demonstration (Xu and Lee 1999). E. coli OmpC created a cell-surface display system as a fastening idea to boost the expression of P. fluorescens SIKW1 lipase TliA (Choi and Lee 2004). Lipase cell surface display appears to stress the cell due to its added heterologous protein "load," however in prokaryotic and eukaryotic host cells; this approach can greatly enhance protein production (Meadows, Kang, and Lee 2018; Hwang et  al. 2014).

8 E. P. CHANDRA AND D. P. SINGH Using metabolic engineering lipase production improvement

Metabolic manufacturing has played a significant role in boosting biofuels production (Stephanopoulos 2007; Saeui et al. 2015). Beneficial protein output has also been increased through cultivation. Though metabolic manufacturing has been compared to lipase production (Son et  al. 2012), it has been comparatively threatened in lipase production. In recent work for extracellular lipase yield, the metabolic man- ufacturing of P. fluorescens has been mentioned as an excep- tion. This keeps a secretion system running, allowing a thermostable lipase enzyme to be excreted (Kanimozhi and

Perinbam 2015). The recombinant protein degradation pro- duced varied depending on the culture media types and air circulation, consuming the natural lipase (TilA) and protease genes SIK W1 of P. fluorescens using the directed gene knockout strategy. The deletion deformed P. fluorescens hid- den recombinant lipase (TilA) at high levels in a blending usage without degradation regardless of growing conditions (Saxena et  al. 2003). To investigate the co-production of

D-psicose and lipase, B. subtilis and E. coli were created utilizing a recombinant co-culture method.

Purifications of lipase Several novel purification approaches are available to acquire lipase homogeneity from a wide number of microorganisms (bacteria, fungus). Purification of lipases excavated on the clarity sought for foodstuff usage usually involves many processes. The fermentation approach is commonly employed for the removal of extracellular microbial lipases from the broth culture. The traditional purification procedures are ultrafiltration, precipitation, and affinity chromatography.

However, several of these procedures are difficult, time-consuming, and costly.

Novel purification apparatuses like (i) membrane sepa- ration processes, (ii) immunopurification, (iii) hydrophobic collaboration chromatography by epoxy-activated spacer arm as a ligand and polyethylene glycol immobilized on sephar- ose, (iv) polyvinyl alcohol polymers as column chromatog- raphy stationary phases, and (v) aqueous two-phase systems are employed commonly (Gopinath et al. 2003). Hydrophobic interaction chromatography is commonly used for enzyme recovery and fold purification. An acid-resistant lipase was isolated from crude commercial preparations utilizing A. niger and size exclusion on Bio-gel-p-100 and ion exchange on Mono-Q (Oliveira de Medeiros, Burkert, and Kalil 2012;

Sugo and Okuyama 2018). Lipase from R. japonicus NR400 was isolated to homogeneity using hydroxyapatite, octylsep- harose, and sephacryl S-20037 chromatography (Niu, Wang, and Tseng 2006; Kojima and Shimizu 2003). For lipases purification from various sources, several distillation pro- cedures are listed in Table 3.

Properties and characteristics of lipases Galvão et  al. (2018) identified lipases as one of the most important biocatalysts with established potential for contributing to bio-industry through lipid technology.

They’ve been employed both in-situ and ex-situ in lipid metabolism and a variety of industrial applications (Delorme et al. 2011). Serine, aspartate, glutamate, and histidine make up the active site of lipases (Pinheiro et  al. 2018), Lipases can be found in two distinct conformations, open (active) and closed (inactive), as a result of their interfacial activa- tion (inactive). Lipases in their open state are said to be more stable than those in their closed state (Willems et  al.

2017). In the presence of hydrophobic surfaces, the open form of lipases usually occurs with the movement of the lid, boosting enzyme activity (Stauch, Fisher, and Cianci

2015). The ultimate features of lipases, such as specificity and selectivity, are not considerably affected by this move- ment (Dobreva, Zhekova, and Dobrev 2019). Genetic manip- ulation or physio-chemical changes can easily change these traits (including immobilization).

Lipases are monomeric proteins with a molecular weight of 19-60 kDa (Sharma, Chisti, and Banerjee 2001). The phys- ical features of lipases are primarily determined by factors such as the position of fatty acids in the glycerol strength, the length of the fatty acid chain, and the degree of unsat- uration (Rengachari et  al. 2012; Adlercreutz 2013). These characteristics have an impact on a triglyceride’s nutritional and sensory value. Numerous lipases are active in organic solvents and catalyze esterification and a variety of other useful reactions (Urban et al. 2001). The amount of ammo- nium sulfate solution cast off during the refinement process determines the rise in lipase activity (Svendsen et  al. 1997;

Raftari et al. 2012; Borkar et al. 2009). C. rugosa lipases for butanol, pentanol, hexanol, propionic acid, and butyric acid,

M. miehei and R. arrhizus lipases for long-chain acids and acetates, and M. miehei and R. arrhizus lipases for long-chain acids and acetates exposed major marketable lipases (Silva and Jesus 2003; Abramić et  al. 1999; Paluzar, Tuncay, and

Aydogdu 2021). Lipase activity is affected by pH, and some lipases are stable across a wide range of pH standards (Bakir and Metin 2016). Most lipases are stable at neutral pH, but others can withstand pH values as high as 4.0 and 8.0 (Hasan, Shah, and Hameed 2009). At acidic pH,

Chromobacterium viscosum, A. niger, and Rhizophus species generated extracellular lipases (Woodcock et  al. 2008). P. nitroaeducens alkaline lipase was isolated and found to be active at pH 11.0 (Rajendran, Palanisamy, and Thangavelu

2009). Lipases are capable of reversing the reaction that leads to esterification and interesterification under some uncertain settings, such as when water lipases are absent (Alvarez and Stella 1989; Akimoto et  al. 2003; Nicholson and Marangoni 2021). Divalent cations, such as calcium, excite activity for the production of lipase action, and cofac- tors are usually not required (Lu et  al. 2013; Falcocchio et  al. 2006; El Khattabi et  al. 2003). Lipase activity was strongly decreased by Co 2+, Ni 2+, Hg 2+, and Sn 2+, while

Zn 2+, Mg 2+, EDTA, and SDS were only mildly inhibited (Winkler and Stuckmann 1979; Carrie, Delaquis, and

Mazza 1999).

When S. marcescens SM-6 was exposed to glycogen, hyaluronate, pectin B, or gum Arabic, lipase growth was considerably improved, and spontaneous and cyclic AMP

Critical Reviews in Food Science and Nutrition 9 production was stimulated (Kambourova et al. 2003; Krebs et  al. 2014). At 40-60 °C, thermophilic bacteriological lipases from Icelandic hot springs performed better. In the temperature range of 30-35 °C, Epipactis gigantean has a strong lipase activity (Augustyniak et al. 2012; Royter et al.

2009; Rogalska et  al. 1993). Lipases can be divided into two groups based on the region-specificity they show when using an acyl glycerol substrate (Wang et al. 2017). Lipases in the first category have no specificity for regions and release fatty acids from all three places of glycerols (Stadler et  al. 1995; Gonzalez-Baró, Lewin, and Coleman 2007).

The second assemblies of lipases issue fatty acids regio-specifically from the outer 1 and 3 locations of acyl- glycerols (Borrelli and Trono 2015). These lipases hydrolyze triacylglycerol to produce free fatty acids 2-monoacylglycerol and 1, 2-(2, 3)-diacylglycerol (Dinanta et al. 2019; Brindley

1991). A. arrhizus, R. delemar, C. cylindracea, and P. aeru- ginosa have all demonstrated fractional stereo-specificity in the hydrolysis of triacylglycerols (Macrae and Hammond

1985). These enzymes are employed to sequester optically pure esters and alcohols that are left behind (De Maria et  al. 2007; Stehr et  al. 2003; Balcão, Paiva, and

Malcata 1996).

Selection of supports Immobilizations and support strategies have been established to increase enzyme activity (Tischer and Wedekind 1999).

In the realm of hydrodynamic circumstances, the selection of support material can be a difficult subject depending on reaction media, settings, enzyme types, and safety policies (Spahn and Minteer 2008; Boller, Meier, and Menzler 2002).

Physical and chemical variables such as pore size, hydro- philic/hydrophobic balance, and surface chemistry parame- ters provide diversity for enzyme attachment (Longo and

Combes 1997). These changes in morphological and physical parameters can alter enzyme immobilization and catalytic activity (Ittrat et  al. 2014). Organic and inorganic supports can be classed as organic or inorganic, and natural and manufactured polymers can be subdivided based on their chemical makeup (Zdarta et  al. 2018).

For enzyme immobilization, several supports have been reported such as natural and synthetic polymers (Bezerra et al. 2015), gels (Meunier and Legge 2013), inorganic mate- rials (Zucca and Sanjust 2014), nanomaterials like glass beads, alginate beads (Datta, Christena, and Rajaram 2013), activated carbons (Cho and Bailey 1977), alumina

Table 3.  Purification methods for lipase enzymes.

Microorganism Purification Method References Pseudomonas spp.

P. fluorescens HU380 Superdex − 200 HR FPLC, Phenyl-toyoperal fractionation (batch-wise),

DEAE– sepharose column chromatography (Nakai et  al. 1968)

Pseudomonas sp.

Superose 12B chromatography, Biogel P-10 chromatography (Lund and Granum 1996; Hiol et  al.

2000) P. fluorescens strain 2 D Hydrophobic interaction chromatography, Ammonium sulfate precipitation (Woods and Que 1987)

P. aeruginosa Hydroxyapatite column chromatography, Ammonium sulfate precipitation (Gilbert, Cornish, and Jones 1991)

P. aeruginosa EF2 Gel filtration (superose) FPLC, Ultrafiltration, Anion exchange chromatography (Mono Q) (Ndaw et  al. 2002)

Bacillus spp.

Bacillus sp.

Ammonium sulfate, Acrinol treatment, DEAE-Sephadex A-50, Toyopearl

HW-55F, Butyl toyoperal 650 M. (Vale et  al. 1986)

Bacillus sp.

Acetone fractionation, Two acetone precipitations, Octyl-sepharose CL-4B, Q

Sepharose, Sepharose-12. (Li and Zong 2010) B. thermoatenulatus

Calcium soap, Hexane extraction, Methanol precipitation, Q-sepharose (ion exchange). (Sarkar et  al. 2012)

Staphylococcus sp.

S. aureus Hydrophobic chromatography on phenyl-Sepharose CL-4B (Sato et  al. 1999)

S. hyicus Sephadex G-100/G-25 column Taipa, Aires-Barros, and Cabral 1992)

Rhizopus spp.

R. japonicus KY 521 Sephadex G-100 gel filtration (Farag and Hassan 2004)

R. oryzae Acetone precipitation (80%), Sephadex G-100. (Nascimento et  al. 2012)

R. arrhizus Ammonium sulfate fractionation and Sephadex G-100 gel filtration. (Tahoun and Ali 1986)

R. delemar Ultrogel and Sephadex G-150 columns (Palissa et  al. 1989)

Penicillium spp.

P. chrysogenum Ultrafiltration, Phenyl-sepharose, Mono Q HR5/5, and PD-10 column on

Sephadex G-25. (Krieger et  al. 1997) P. citrinum Extraction and back-extraction using AOT reversed micelles in isooctane and phenyl- superose column. (Kimiyasu, Akiba, and Yamaguchi 1988)

Penicillium cyclopium M1 Ammonium sulfate, DEAE-cellulose, DEAE-Sepharose, hexyl-Sepharose, and hydroxyapatite chromatography and gel Filtration on Cellulofine GC-700. (De Jong et  al. 1992)

Penicillium simplicissimum Phenyl-Sepharose column. (Isobe et  al. 1992)

Penicillium camembertii U-150 Ammonium sulfate fractionation, Aminooctyl-Sepharose, Hydroxyapatite (Gargova et  al. 2006)

Aspergillus sp.

Aspergillus niger Hydrophobic interaction (butyl Toyopearl 650 M), Gel filtration (Sephadex

G-75), Anion-exchange chromatography (DEAE-Sepharose CL-6B) (Nagaoka and Yamada 1973)

Mucor lipases sp.

Mucor lipolyticus Aac-0102 CM-Sephadex column chromatography (Whitaker 1963)

Mucor javanicus Gel filtration on Sephadex G-200 (Ota and Yamada 1966)

Yeast Lipase Candida paralipolytica M-Sephadex C-50 and DEAE-Sephadex C-50. (Maruyama et  al. 2000)

10 E. P. CHANDRA AND D. P. SINGH (Sigurdardóttir et al. 2018), silica (Zdarta et al. 2018), celite (Sağiroğlu, Kilinç, and Elefoncu 2004), ceramic (Kujawa et  al. 2021), metal oxides (Zucca, Fernandez-Lafuente, and

Sanjust 2016), modified sepharose (Mohamed et  al. 2021), or treated porous glass which is an organic material (Lee,

Lin, and Mou 2009) and certain polymers are the most commonly used supports. One of the desired qualities of support matrix is mesoporous material, which has a higher number of pores and surface areas, resulting in increased enzyme loading per unit mass (Nematian et  al. 2020). A technical process is used to fix enzymes to or within solid supports, resulting in a heterogeneous immobilized enzyme system (Krajewska 2004). The enzymes’ structure is stabi- lized by solid support systems, and their activity is main- tained (DiCosimo et  al. 2013).

Strategies for immobilization Enzyme immobilization is an efficient approach for improv- ing the stability and reusability of enzymes even under adverse environmental conditions such as pH, temperature, and organic solvents, making enzymes more inexpensive and feasible in industrial production (Guzik, Hupert-Kocurek, and Wojcieszyńska 2014; de Souza et al. 2020). Because the elimination of lipases can increase the enzyme’s selectivity, repeatability, substrate range, and separation from the raw materials, immobilization of the lipase enzyme is critical for its economic application (Chapman, Ismail, and Dinu 2018).

Immobilization of lipase has several advantages, including better stability, lipase enzyme ejaculation halt, and greater lipase enzyme interaction with raw materials (Muniandy et  al. 2019; Gonçalves, Silva, and Guidini 2019). Lipase enzyme contains a small number of lysine residues, making immobilization challenging. The lysine structure’s functional groups play a significant role in matrix binding (Rafiee and

Rezaee 2021). Its good solid support for lipase immobiliza- tion since it has available hydroxyl (-OH) and amine (-NH2) functional groups: Enzyme immobilization is a straightfor- ward and repeatable procedure that does not require spe- cialized equipment (Zaitsev et al. 2019; Sharma et al. 2021).

Techniques for enzyme immobilization Physical adsorption, ionic bonding, and covalent bonds are all maintained by enzymes (Casas-Godoy et  al. 2012).

Physical methods (reversible) and chemical methods (irre- versible) are two types of enzyme immobilization methods (Pinheiro et  al. 2019). Enzymes can be removed from the supporting material using reversible immobilization under mild circumstances. When enzymes adhere to supporting materials in irreversible immobilization, they are unable to be withdrawn without compromising the support or the enzyme’s biological activity (Moreno-Pérez, Guisan, and

Fernandez‐Lorente 2014; Singh and Mukhopadhyay 2012;

Xavier Malcata et  al. 1990; Mead, Irvine, and Ramji 2002).

Adsorption is a physical mechanism while cross-linking, covalent bonding processes, and entrapment is chemical methods (Harini et al. 2021; Liang et al. 2020; Soares et al.

1999; Gupta, Gupta, and Rathi 2004). In enzyme immobi- lization processes, a combination of these strategies has recently been applied (Liu, Schmid, and Rusnak 2006; Larios et  al. 2004).

The kinetic characteristics of immobilized lipase improved significantly in terms of activity, specific activity, Km and

Vmax, optimal pH, and temperature (Becker et  al. 1997;

Pereira et  al. 2001; Gawas, Jadhav, and Rathod 2016). To examine the impact of immobilization on the enzyme’s cat- alytic effectiveness, two kinetic parameters, the Michaelis constant Km and the maximal reaction velocity Vmax, are determined for an immobilized enzyme when compared to its non-immobilized counterpart (Neira and Herr 2017;

Cooney 2017). Mechanical strength, chemical resistance, and microbial breakdown are all requirements for carrier archi- tectures. On the other hand, on the surface of carriers, a large number of reactive groups and a hydrophilic chain were

Figure 2.  Categorizations and representation of different enzyme immobilization procedures.

Critical Reviews in Food Science and Nutrition 11 required (Salvi, Kamble, and Yadav 2018). Other (desired or undesirable) events may emerge after immobilization, but the researcher must recognize them and devise instruments to regulate them. The diverse enzyme immobilization approaches and their categorizations are shown in Figure 2.

Physical methods or reversible Adsorption Organic polymers, charcoal, glass, mineral salts, metal oxides, or silica gel materials were adsorbed by enzymes.

Nelson was the first to use this strategy, although the mechanics involved in adsorption aren’t fully understood (Samiey, Cheng, and Wu 2014; Hartmann and Kostrov

2013; Sugahara and Varéa 2014). This method of regu- lating enzymes has the advantage of being both cost-effective and chemically modest; additionally, no chemicals are required, and the approach only involves a few triggering stages (Nguyen and Kim 2017).

Furthermore, when compared to biochemical approaches for immobilizing enzymes, the process of immobilization enzyme is a lesser amount expected for a period to be denatured (Sheldon and van Pelt 2013). To express the immobilization in adsorption, hydrogen bonds, coordina- tion bonds, and Van der Waal’s forces are intertwined (Wang et  al. 2009). However, some disadvantages are identified as a result of the binding’s weakness. Desorption of the bound enzyme can occur as a result of changes in temperature, pH, ionic strength, or the presence of substrate, for example (Engasser and Horvath 1975).

Furthermore, the sustenance does not selectively fix the enzyme; other irrelevant matter can also be adsorbed onto the environment, causing impediments such as denatur- ation of the enzyme to become inconsistent.

Chemical methods or irreversible Entrapment The enzymes can be surrounded with courses such as encap- sulating an enzyme with a polyacrylamide gel (Klibanov

1983). The free diffusion of low molecular weight substrates is dealt with in this approach of immobilizing enzymes, and the final products and enzymes, which are considerably larger particles, cannot escape out of the beads (Mallardi et  al. 2015). Several matrices are used to capture enzymes, including polyurethane foams, silastic gels, and starch gels (Jancsik, Beleznai, and Keleti 1982). There is no bond for- mation between the enzyme and the matrix, implying that the enzyme is exposed to a minimum of severe stresses.

Though a wide range of enzymes can be immobilized using entrapment methods, only low molecular weight substrates can be employed in such performances. Because food sys- tems include macromolecules, this could provide a check for their usage in food knowledge (Jun et  al. 2019; Kilara et  al. 1979). Academically, the pore size of the polymer should prevent enzyme leakage; nevertheless, in practice, a wide range of pore sizes are encountered, which can result in enzyme loss due to diffusion (Bayne, Ulijn, and Halling

2013; Califano and Costantini 2020).

Cross-linking Bifunctional reagents, such as glutaraldehyde, were used to join intermolecularly the molecules of enzymes. Bifunctional compounds can contain two functional groups that are the same or different, as well as groups of reactivities that are different (Kim and Herr 2013; Kluger and Alagic 2004).

The intermolecular associating enzyme molecules are inad- equate, as the enzyme’s support function, resulting in low events. Because enzymes are expensive biochemicals, inad- equate use of enzymes in this technique results in a price increase for immobilized enzymes (Bedran-Russo et al. 2014).

Covalent bonding The method of enzyme immobilization is based on the creation of strong covalent bonds between the functional groups on the carrier particle and the enzyme (Brena,

González-Pombo, and Batista-Viera 2013). The amino groups (NH2) of arginine or lysine, a carboxylic group (COOH) of glutamic acid or aspartic acid, a hydroxyl group (OH) of threonine or serine, and a sulfhydryl group (SH) of cysteine are all covered on the enzyme molecule’s surface (Sulaiman et  al. 2015).

Ionic bonding The standard of a protein-ligand interface is ionic bonding, which is the basis of enzyme immobilization used in chro- matography, such as -amylase on calcium phosphate (Ueland et al. 1993). This procedure is cost-effective since it is simple and affordable, and it results in minimal or no loss of enzyme or carrier molecule (Katchalski-Katzir and

Kraemer 2000).

Entrapment of enzyme in support The process of entrapment leads to enzyme inclusion within a polymeric structure. The enzyme is supported by allowing the substrate and yields to pass through (Rodriguez-Abetxuko et  al. 2020). The support acts as a barrier to mass transfer and serious reaction kinetics (Benzinger, Becker, and

Hüttinger 1996). There are numerous major approaches to entrapment, including ionotropic gelation of macromolecules with multivalent cations (alginate), temperature-induced gelation (agarose, gelatin), organic polymerization reaction by biochemical/photochemical (polyacrylamide), and pre- cipitation from an immiscible solvent (polystyrene). On a polyacrylamide gel, -amylase or amyloglucosidase are two examples (Pillay et  al. 1998; Bocchetta 2020).

Cross-linking of the enzyme with support In this approach of enzyme immobilization, chemical bond- ing is used to connect an enzyme molecule to the support, resulting in the use of a three-dimensional structure. As the most common crosslinking agents, glutaraldehyde

12 E. P. CHANDRA AND D. P. SINGH bonds via its amino group, whereas diamines crosslink via carboxyl groups (Weinhold 2012; Zhou, Fan, and Chen

2016). The enzymes are well attached, therefore there are few chances of enzyme desorption using this method.

Cross-linking, on the other hand, has the potential to create significant changes in the dynamic site of the enzyme molecule, which is a disadvantage of this method.

Using nanotechnology in immobilization For enzyme immobilization, nanoparticles, nanofibers, nano- tubes, nanopores, nanosheets, and nanocomposites are employed as nano scaffolds (Misson, Zhang, and Jin 2015).

Using nanomaterials for enzyme immobilization has various advantages, including high enzyme loading, low mass trans- fer resistance, high mechanical strength, quick separation process, and better enzyme stability and activity thanks to the vast surface area. Nanocarriers were exposed to chemical and thermal accepting nanocatalysts with high enzyme load- ing and increased catalytic activity (Bilal et  al. 2019).

Because of their high surface-to-volume ratio, minimal mass transfer confrontation caused by the outer magnetic field, and ease of separation, magnetic nanoparticles are the best standard carriers for enzyme immobilization among all nanocarriers for enzyme loading (Johnson, Park, and Driscoll

2011; Husain et  al. 2011).

Advantages of enzymes immobilization Immobilization improves lipase stability at high tempera- tures, and immobilized lipase is more active than free lipase. The optimal reaction temperature for immobilized lipase shifts from 37-50 °C to 37-50 °C for free lipase (Carvalho et  al. 2013). High temperatures have less of an impact on lipase activity, although they do improve thermal stability. Immobilized Candida rugosa lipase has an optimal temperature of 20-50 °C, while free Candida rugosa lipase has an optimum temperature of only 40-50 °C (Mokhtar et  al. 2020: Lee et  al. 2011). The electrostatic charges shifted after immobilization, causing the immobilized enzyme’s optimum pH to shift to a mildly alkaline range (Kahveci and Xu 2011). The number of acidic groups increases after immobilization, making the enzyme more polyanionic appealing (Guzik, Hupert-Kocurek, and

Wojcieszyńska 2014). In alkaline settings, immobilized lipase has higher activity than free lipase. Immobilization improves enzyme characteristics by enhancing enzyme stiff- ness and heat resistance (Matsumoto and Ohashi 2003). It also causes the enzyme to have limited structural flexibility, which is evidenced by an increase in the optimal tempera- ture and stability against inactivation. Several industrialized methods necessitate high temperatures and better thermal performance of immobilized enzymes, which serve as the foundation for enzyme applications (Sirisha, Jain, and

Jain 2016).

Potential applications of microbial lipases In the processing of food, fats, and oils, the synthesis of fine chemicals and pharmaceuticals, detergents and degreas- ing formulations, paper manufacture, pharmaceuticals, bio- sensors, pulp and paper, textile production of cosmetics, industry, etc. Lipases are widely used (Sharma, Chisti, and

Banerjee 2001). Major potential applications of microbial lipases are summarized in Table 4 (Aravindan et al. 2006).

Lipases in food industry Enzymes are found in a wide range of products used by humans and animals regularly. It can be utilized as a dietary additive by changing the location of fatty acid chains in the glyceride and substituting one or more fatty acids with new ones (Alves-Bezerra and Cohen 2017; Henry 2009). Lipase can alter the properties of lipids, boosting the flavor of some foods, improving their texture, prolonging the shelf life of others, and increasing the softness of others, as well as improving food quality (Stanhope 2012; Turan and

Erkoyuncu 2012).

In flavouring and aroma Microbial enzymes generate a wide range of aromatic and flavoring compounds used in the food industry. Salad dress- ings, dairy, confectionery, and bread products all use lipases

Table 4.  Potential applications of microbial lipases.

Industries Applications of Products Action Food Industry

Fats and oils Cocoa butter, margarine, fatty acids, glycerol, mono- and diglycerides

Transesterification, hydrolysis Meat and fish Meat and fish product, fat removal

Flavor development Health foods Health foods Transesterification

Food sauce Mayonnaise, condiment, and whipping cream

Quality improvement Beverages Alcoholic beverages, e.g., sake, wine

Improved aroma Bakery foods Shelf life propagation

Flavor improvement Dairy foods Development of flavoring agents in milk, cheese, and butter

Hydrolysis of milk fat, cheese ripening, modification of butter fat

Agrochemicals Herbicides such as phenoxypropionate

Esterification Fuel industries Biodiesel production

Transesterification Pollution control To remove hard stains and hydrolyze oil and grease

Hydrolysis and transesterification of oil and grease

Cosmetics Act as emulsifiers and moisturizers Synthesis

Pharmaceuticals Production of various intermediates used in the manufacture of medicine

Hydrolysis of expolyester alcohols Detergents industry

Removal of oils stains from fabrics Hydrolysis of fats

Critical Reviews in Food Science and Nutrition 13 to flavor and aromatize them. Lipases used in the tea indus- try for the production of the flavor, the concentration of polyunsaturated fatty acids increases while the lipid com- position decreases. It’s also used to provide a distinct fra- grance in alcohol manufacturing. Short-chain fatty acids were converted into esters by lipase enzymes, which gave food its flavor. Processing variables like pH, temperature, amount, and emulsion content all influence the level of flavor and scent (Rajendran, Palanisamy, and Thangavelu

2009). Lipase can be used to change the flavor of a cake, cookie, dough, cheese, or confectionery, among other things.

In human milk fat substitutes Several lipids are contained in human milk fat (HMF), including palmitic (20-30%), stearic acids (5.7-8%), linoleic (7-14%), and oleic (30-35%) (Sánchez-Hernández et  al.

2019). Unlike vegetable oils and cow’s milk fat, palmitic acid is the predominant saturated fatty acid in HMF and is esterified at the sn-2 position of the TAGs (Zou et  al.

2013; Sahin, Akoh, and Karaali 2005). Unsaturated fatty acids are found at the exterior positions (Bracco 1994). The profile of HMF fatty acid has a significant impact on its digestion and intestinal absorption in babies (Ghosh et  al.

2016). Human milk fat alternatives (HMFS) have been pro- duced using sn-1, 3 lipase-catalyzed acidolysis of tripalmitin, butterfat, palm oil, palm stearin, or lard (rich in palmitic acid in the sn-2 position) and free fatty acids (FFA) from various sources (Soumanou, Pérignon, and Villeneuve 2013).

The commercial Betapol® product is industrialized utilizing

IOI Loders Croklaan, by acidolysis between lard and soybean fatty acids, catalyzed by sn-1, 3 specific lipases from

Rhizomucor miehei (Lipozyme® RM) (Haraldsson et al. 1989;

Xu 2000).

Lipases in egg processing In the food sector, eggs provide functional ingredients with a variety of qualities, including foaming, gelation, emulsifi- cation in batters and mayonnaise, and enhanced texture of baked items (Foegeding, Luck, and Davis 2006). The emul- sifying capabilities of eggs are also due to their lipids (Anton

2013). Emulsification of egg yolks in dressings: The global production of emulsified dressings is projected to be 3 mil- lion metric tons per year, with around 15, 0000 metric tons of egg yolks consumed in the process (Alu’datt et  al. 2017;

Miranda et al. 2015). One-third of the market for emulsified dressings is centered in Russia and Eastern European nations. Nestlé, Kraft, and Unilever operate in a highly industrialized market with worldwide players (Ene-Obong and Carnovale 1992). The egg yolk was created as a com- plicated oil-water emulsion with 50 percent water, 32 percent

Figure 3.  Phospholipids chemical structure in egg yolk (Chandra et al. 2020b).

Figure 4.  Phospholipases action in A1 and A2 positions (Guerrand 2017).

14 E. P. CHANDRA AND D. P. SINGH lipids, and 16 percent protein (Mine 1998). Egg yolk con- tains phosphatidylethanolamine; roughly 1/3 of the lipids are phospholipids, with phosphatidyl-choline (PC) (Figure

3), accounting for about 80% of the total. The emulsion stability was improved due to the enzymatic conversion of egg yolk phospholipids into lysophospholipids (van Hoogevest and Wendel 2014; Pichot, Watson, and Norton 2013).

Performing at several positions enzyme constructers have industrialized dissimilar phospholipases (PL) (Figure 4).

Microorganisms (Maxapal A2, DSM, NL) utilized for egg processing, and lipases derived from hog pancreas (Lipomod

699, Biocatalysts, UK) (Bjurlin, Bloomer, and Haas 2001).

Using pork pancreas lipases, nearly 80% of phospholipids are converted to lysophospholipids after 1 hour. Although the enzyme is of animal origin (vegan consumers, Halal/

Kosher requirements), its use is restricted (Fuglsang et  al.

1995). The high conversion yield is also influenced by cloned and articulated Aspergillus niger lipases (such as DSM

Maxapal A2), although numerous industrial players are opposed to depleting enzymes produced by reformed microbes.

Lipases in bakery and confectioneries In most bakers’ goods, enzymes are the most important ele- ment. Due to the controlled use of chemical additions in the production of bread and other fermented foodstuffs (Numan and Bhosle 2006). Enzymes are predicted to play an increas- ingly bigger role in baking shortly. Wheat flour is the most important ingredient in the production of baked products such as bread, cake, pastries, biscuits, crackers, cookies, pies, and tortillas, as well as a vital source of enzyme substrates (Villarino et  al. 2016). The first generation is 1, 3-specific, preferring removing fatty acids from positions 1 and 3 in

TAG (Beisson et  al. 2000; Grönke et  al. 2005). Lipases (TAG lipases) improve dough rheology, increase dough strength and stability, and hence improve dough machinability. Lipases function simultaneously on TAG, diacylgalactolipids, and phos- pholipids to generate more polar lipids, higher dough stability, and a fine, homogeneous bread crumb structure (Huang et al.

2019). Lipase enhances the volume and crumb structure of high fiber white bread by increasing the expansion of the gluten network, wall thickness, and cell density.

Lipases in cocoa processing Cocoa butter has a melting point of 37 °C and contains palmitic and stearic acids. It melts in your mouth, giving you a cooling sensation (Bloomer, Adlercreutz, and

Mattiasson 1990). Unilever patented this invention in 1976 to create a cocoa butter substitute utilizing immobilized lipase (Owusu-Ansah 1994; Ewens, Metilli, and Simone

2021). Rhizomucor miehei commercializes the immobilized lipases which form the transesterification reaction. In this technology, palmitic acid will be replaced by stearic acid, resulting in stearic-oleic-stearic triglyceride.

Processing of edible fats and oils Immobilized lipase is widely utilized in the oil and fat busi- ness, and is primarily employed in the processing of fats and oils to enhance flavor (Murty, Bhat, and Muniswaran

2002). The immobilization technique improves the enzyme’s stability and activity; immobilized lipase is preferred over free enzyme (Facin et al. 2019; Liu et al. 2012). The immo- bilized version of the enzyme can be reprocessed. The trans- esterification reaction induced by R. miehei is carried out with immobilized lipase. Palmitic acid is substituted with stearic acid in this method (Sultan and Sokolove 2001).

Immobilized lipase is employed for the esterification of functionalized phenols and the production of antioxidants, and it is also castoff in sunflower oil due to its lipophilic nature (Zoumpanioti et al. 2010). Trans-fat-free oils are also made with immobilized lipases. Although soy oil has low oxidative stability, it does contain a significant amount of unsaturated fatty acids (Song et  al. 2015). As a result of partially hydrolyzed activity, trans-fat was formed, which harmed health (Klonoff 2007; Tyburczy, Mossoba, and Rader

2013). Enzymatic interesterification is possible at very low aqueous medium concentrations. The immobilized enzyme is also utilized to create zero or low trans-fat oil, shortening, and cooking oil using corn, sunflowers, and/or soy oil as a foundation (Farfán et al. 2015; Colón-Ramos, Monge-Rojas, and Campos 2014).

Lipases in the cheese industry Following the settling whey was a fractional or full mixture of coagulating milk, cream, skimmed milk, or partially skimmed milk, buttermilk, or its fresh and mature products known as cheese. The enzymatic process is the most prev- alent method of milk coagulation; there are two types of cheese manufacturing processes: acidification and enzymes that coagulate milk (Dalgleish 1993; McMahon, Brown, and

Ernstrom 1984). The creation and ripening of this sort of cheese are done by lipase-like enzymes, and the cured and matured cheese is not ready to eat right away. The texture and flavor of fat have an important impact on the cheese properties of numerous milk ingredients. Based on the com- position of the base of fat, cheeses can be classified as having (less than 25 percent) less fat, (25-45 percent) mod- erate fat, (45-60 percent) loaded fat, and (> 60 percent) more fat (Moatsou et  al. 2019).

Cheese flavor improves throughout maturation via pri- mary (lipolysis, proteolysis, metabolism of residual lactose, lactate, and citrate) and secondary (metabolism of fatty acids and amino acids) biochemical pathways due to the produc- tion of a wide spectrum of complexes (Forde and Fitzgerald

2000; McSweeney 2004). Mold-ripened cheeses such as

Camembert, Brie, Roquefort, and Stilton are distinguished by a wide range of lipolysis. In such cheeses, both proteolysis and lipolysis leave a distinct flavor and scent (Dugat-Bony et  al. 2015; Brennan et  al. 2004). Lipases derived from a variety of sources catalyzed cheese by lipolysis, primarily by enzymes found in rennet adhesive, in the creation of cheese varieties such as Provolone, old-fashioned Greek Feta, and

Critical Reviews in Food Science and Nutrition 15 Italian Pecorino (Rani and Jagtap 2019; Schirone et al. 2012).

Secondary reactions result from the degradation of fatty acids and amino acids in the production of volatile taste molecules (Ianni et  al. 2020; Wang et  al. 2020). Lipolysis has an important part in the production of large quantities of short-chain volatile fatty acids in EMC taste, which is linked to aroma intensity and flavor (Kilcawley et  al. 1998;

Mohebbi et  al. 2008; Lee et  al. 2007).

The EMC emulsification course, which includes substrate-assisted lipolysis, provides an ideal environment for lipases by increasing the oil-water contact (Li and

McClements 2011). Which has a long-term preference for triglycerides, particularly those found near an oil-water interface, over free solutions, which can work up to 1000 times quicker (Armand et  al. 1999; Golding et  al. 2011).

The significance of uncontrolled lipolysis is the increase of rancidity or soapiness due to an imbalance between short- and long-chain fatty acids (Sobczak, Blindauer, and Stewart

2019; Crielly, Logan, and Anderton 1994). Using the lipolytic process with esterase and lipase activities to prevent the off-flavor (Kilcawley, Wilkinson, and Fox 2001). A variety of EMC flavors are used to make blue, Swiss, Gouda,

Mozzarella, and Parmesan cheeses (Fox 1993). Cheddar EMC is particularly important because it can be found in a variety of supposed Cheddar flavors and intensities from various manufacturers. Bacterial enzymes play a key role in the formation of taste in surface-ripened cheeses; these smear bacteria hydrolytic activities are involved in the production of cheese flavor components (Williams, Beattie, and Banks

2004) and this is determined by on profile of enzyme of pressures elaborate in the starter culture (Stefanovic et  al.

2018; Bockelmann 2000; Rubio et  al. 2014).

Lipases in the fish and meat industry Lipase is also utilized in the meat and fish industries to generate lean meat. It is also used to improve its suitability for the fermentation of meat products to improve the supe- riority of fermented sausages. The aggregation of unsatu- rated fatty acids (n-3 PUFA) and the hydrolysis of fish oil lipases are employed (Yan et  al. 2011).

Applications of lipases in the oleo-chemical industry

In addition to their natural role of hydrolyzing carboxylic ester bonds, lipases can catalyze esterification, inter-esterification, and transesterification processes in non-aqueous conditions (Yadav and Devendran 2012; Nishio et  al. 1989). In the modern oleo chemical sector, the use of immobilized lipases to initiate diverse reactions such as hydrolysis, alcoholysis, and glycerolysis, as well as the usage of mixed substrates, has expanded significantly (Arana-Peña et  al. 2020). As a result of the application of immobilized enzymes, the practices of continuous running and high pro- duction are confirmed. As a result, fat splitting and alter- ation can be accomplished without a major investment in expensive equipment or the expenditure of enormous amounts of thermal energy (Chapman, Ismail, and Dinu

2018; Ansari and Husain 2012). Lipases have a wide range of applications because they save energy and reduce heat deterioration during hydrolysis, glycerolysis, and alcoholysis (Arbige and Pitcher 1989). Ethyl caprylate, a sweet fruity-flowery flavoring produced from Bacillus subtilis EH37 strain employing immobilized lipase, is a very valuable tool for food and beverage flavorings. Using lipase, Streptomyces thermocarboxydus ME168 strain was found to create sugar esters from a variety of substrates including glucose, fruc- tose, and different vinyl esters (Siebenhaller et  al. 2018).

Lipase in bioremediation Lipids and carbohydrates are the principal waste materials produced by the food industry, and the formation of oily coatings on aquatic surfaces poses a threat to the eco- system. Because of the emulsification with organic mate- rials, the oily coating clogs and prevents oxygen diffusion (Baloch and Hameed 2005). The cytotoxic potential of cooking oil waste can be reduced by enzymatic activity based on lipase, which has well-known environmental consequences (Okino-Delgado et  al. 2017). Lipases cata- lyze a wide range of lipid changes because of their broad substrate affinity and great temperature and solvent sta- bility (Kanmani, Aravind, et  al. 2015). The effect of immobilized lipases such as P. aeruginosa, Staphylococcus epidermidis, and Bacillus subtilis on the removal of stored lipids and grease from wastewater pipelines has been evaluated and is known (Ηatzisymeon, Kamenopoulos, and Tsoutsos 2019).

Filamentous fungus-like Penicillium chrysogenum, Penicillium cyclopium, Penicillium simplicissimum, Penicillium expansum, Candida rugosa, Aspergillus fumigatus,

Trichoderma sp. (Bancerz et  al. 2005; Nowak, Kucharska, and Kamiński 2019; Mehta, Bodh, and Gupta 2018), and bacterial strains like Bacillus subtilis, Pseudomonas sp. (Singh et al. 2018; Hu et al. 2018). The toxicity of jatropha seed cake (JSC) is produced in large quantities as a by-product in the biodiesel industry. The AAU2 lipase enzyme from Pseudomonas aeruginosa degraded and detox- ified phorbol ester (phorbol 12-myristate 13-acetate) using

JSC as a substrate (Muhamad et  al. 2019).

The disposal of grease waste poses a severe challenge and poses a hazard to the ecosystem because it is non-biodegradable.

Waste grease has been thrown in littered sites or sewages in numerous countries without any preparation (Jayathilakan et al. 2012; Toldrá, Mora, and Reig 2016; Kanmani, Aravind, et al. 2015). Grease wastes induce a decrease in cell-aqueous phase transfer rates, reduced sedimentation, the creation of floating sludge, clogging, and the appearance of foul aromas in effluents (Chandra and Singh 2014). The lipase-producing activity of P. chrysogenum SNP5 is employed for grease cleanup (Yeo, Nihira, and Yamada 1998).

P. putida GPo1 alkB; Acinetobacter spp. alkM; Rhodococcus spp. alkB1, Rhodococcus spp. alkB2, P. putida xylE, P. putida ndoB, and Mycobacterium sp. strain PYR-1 nidA are indig- enous cold-adapted microbes that degrade n-alkane (Hasan,

Shah, and Hameed 2006). Pseudomonas protegens BC2-12,

16 E. P. CHANDRA AND D. P. SINGH Pseudomonas protegens CHA0, Pseudomonas protegens Pf-5,

P. fluorescens A506, and Pf0-1, Pseudomonas chlororaphis lipase producing bacteria are used in the biodegradation of polyurethanes (PUR) (Wilkes and Aristilde 2017; Chandra,

Enespa, and Singh 2020a). P. aeruginosa has been found to produce large levels of extracellular lipase, which aids in the biodegradation of polyesteramides and aromatic-aliphatic polyesters. One of the first enzymes on PUR was PueB lipase, which was discovered in Pseudomonas chlororaphis (Wilhelm, Tommassen, and Jaeger 1999; Pathak and Navneet

2017). The industrial bacterial residue of Paracoccus sp. strain KDSPL-02 was used to biodegrade penicillin G in an extended bed adsorption bioreactor (EBAB) (Nerurkar et  al. 2013).

Lipase in detergent engineering Pseudomonas aeruginosa AAU2 lipase enzyme showed sig- nificant stability in the presence of commercial detergents such as Ezee® and Wheel®. Ohgiya et al. 1999, used a deter- gent made from Bacillus sonorensis and lipase to replace corn-oil stains on un-dyed cotton fabric. Cold-active lipases can be used as additions in detergent formulations and in an organic mixture with a chiral intermediate to wash gar- ments at low temperatures in the laundry (Unni et al. 2016).

An alkaline and thermotolerant lipase produced by Pseudomonas aeruginosa strain BUP2 (Olusesan et al. 2009) are employed in the detergent business with efficiency and high specific activity. Lipo Prime ® is a lipase-containing detergent also made by them (Hemlata, Uzma, and Tukaram

2016). Trichosporon asahii MSR 54 produced an alkaline lipase, which was used to develop a presoak formulation for the removal of oil stains at room temperature (Guncheva and Zhiryakova 2011).

Lipases in the medical sector Bacillus lipases can be employed in the pharmaceutical industry for the manufacture of enantiopure substances because of their characteristics. Staphylococcus lipase pro- duces antioxidants such as tyrosol acetate, propyl gallate, and eugenol benzoate (Matull, Pereira, and O’Donohue

2006). When tuberculosis (TB) is detected, lipase can be utilized as a diagnostic tool. The level of lipase in blood serum can be utilized to detect acute pancreatitis and asso- ciated wound (McCutcheon 2000). A bile duct obstruction or an overdose of alcohol can cause pancreatitis (Hazem

2009). Lipases are employed in the creation of hair waving as an ingredient of topical anti-obese creams and for the treatment of malignant tumors since they act as TNF acti- vators (Aghdassi et  al. 2008). The lovastatin medication, which is made by Candida rugosa lipase, lowers serum cholesterol levels. Diltiazem hydrochloride is made via asymmetric hydrolysis of a crucial intermediate

3-phenylglycidic acid ester from S. marcescens lipase and is commonly used for coronary vasodilation (Gácser et  al. 2007).

Lipases in pathogenicity mechanisms Lipase 8 (LIP8) genes play an important role in Candida albicans pathogenicity. The mutant’s deficit in the LIP8 gene was less virulent expressively in lipid-containing media low- ering LIP8 expression observed in lowered growth and a murine intravenous contagion way (Toth et al. 2017; Fourie et  al. 2018). Lip5 and Lip8 from Candida albicans promote host-pathogen contact (Chin et  al. 2016). Candida species produce hemolysins, which decrease hemoglobin and extract elemental iron from host cells (Mayer, Wilson, and Hube

2013). Hemolysins were chosen as essential virulence factors because they may aid the pathogen’s survival and persistence in the host. Candida albicans, regarded as a versatile patho- gen implicated in oral, vaginal, and systemic infections, is responsible for a greater percentage of death in the United

States due to infection in the circulation (Naglik, Richardson, and Moyes 2014; Chaffin 2008). Lip5, Lip6, Lip8, and Lip9 lipases were released by C. albicans, and they can allow the organism to attach and colonize in people after infection (Stehr et al. 2004; Park et al. 2019; Lemieux and Simard 1991).

Lipases mediated transesterification ergosterol manufacturing

Bacillus subtilis, Pseudomonas fluorescens, and Aeromonas hydrophilia are dairy food putrefaction bacteria that have produced unfavorable changes in the quality of food and cosmetics, as well as affecting the market (Machado et  al.

2017). Psychrophilic P. fluorescens and A. hydrophilia, as well as thermophilic Bacillus subtilis, change the texture, off-flavor, and cause bitter odor in a variety of dairy prod- ucts (Torbeck et al. 2011). Burkholderia cepacia, as a poten- tial pathogen, caused many skin disorders as a result of an infection in creamy cosmetic goods (Akinboyo et  al. 2018;

Tavares et al. 2020). Several nosocomial outbreaks for immu- nocompromised individuals with cystic fibrosis and chronic granulomatous illness in health care facilities have been linked to Burkholderia cepacia complex (Bcc) infection (McMenamin et al. 2000). Ergosterol is used as a biomaterial substance in food and cosmetics manufacture because it has no antagonistic clinical effects with cholesterol and has var- ious useful functional properties (Corrêa et  al. 2017).

However, because ergosterol is simply crystalline and has low oil solubility, it is difficult to employ in the business (Kuksis and Beveridge 1960). This difficulty can be remedied by combining fatty acids with ergosterol. Ergosterol oleate (EO), ergosterol linoleate (EL), and ergosterol linolenate (ELn) were produced with the help of Proteus vulgaris K80 lipase (He, Zhu, and Chen 2018; Park and Kim 2020). The microorganisms that cause putrefaction in dairy and cos- metics can be inhibited by these unsaturated fatty acid ergosterol esters (FAEEs) (Badellino et  al. 2005).

Lipases use in a diagnostic tool Lipases are also important therapeutic targets or enzyme markers in the medical field. Their presence or increasing

Critical Reviews in Food Science and Nutrition 17 levels can be utilized to identify specific contagions or dis- eases and can also be employed as diagnostic tools. In the enzymatic resolution of blood triglycerides, glycerol lipases are used, with an enzyme-linked colorimetric response that is sequentially resolute (Treacy et  al. 2001). The level of lipases in the blood serum can be used as an analytical method to detect situations such as pancreatic damage and acute pancreatitis illnesses (Kivanc, Yilmaz, and Demir

2011). A large number of virulence-related genes Aeromonas bacteria are found in drinking water, so it’s important to look at as many different samples as possible to better understand the health risks these bacteria may pose (Sen and Rodgers 2004). The virulence characteristics of

Aeromonas bacteria indicate that drinking water is a source of potentially harmful Aeromonas bacteria cleaned from municipal water. Rasmussen-Ivey et al. (2016) found that P. acnes, Corynebacterium acnes, and Staphylococcus aureus are pathogenic bacteria and that lipase from these bacteria influ- ences skin rash in acne patients (Moradali, Ghods, and

Rehm 2017). Many enzymes, including lipases, are discov- ered as biological materials subsidizing the pathogenicity between the virulence factors by an opportunistic bacterium known as P. aeruginosa to produce and secrete numerous different virulence factors (Rasamiravaka et  al. 2015;

Vasil 1986).

Lipases in pharmaceuticals Aryl aliphatic glycolipids, citronellol laurate from citronellol and lauric acid, and ethyl esterification of docosahexaenoic acid to ethyl docosahexaenoic acid to ethyl docosahexaenoic acid to ethyl docosahexaenoic acid are the cold-active lipases utilized in industrial (Ota, Sawamoto, and Hasuo 2000;

Kauffmann and Schmidt-Dannert 2001; Ugur and Boran

2014). The fatty acid chain length of an ester was selected by Bacillus lipases, although only a small number of enzymes showed positional specificity (Ranjan et  al. 2016; Horchani et  al. 2010). Bacillus lipases can be discarded in the phar- maceutical industry for the synthesis of enantiopure com- plexes because of these features (Contesini et  al. 2020).

Staphylococcus lipase is used to produce antioxidant qualities such as tyrosol acetate, propyl gallate, and eugenol benzoate (Lim and Lee 2012). In the case of tuberculosis (TB) detec- tion, lipase can be employed as a diagnostic tool.

Mycobacterium tuberculosis lipase is used to assess for infec- tion with good specificity and sensitivity (Nyendak,

Lewinsohn, and Lewinsohn 2009; Li and Lo 2015). They have immobilized because of their regio-, chemo-, and enan- tioselectivity. CalB is a catalyst and pharmacological active ingredient in the synthesis of Odanacatib (Bovara et a.,

1993). Chronic liver disease is caused by the hepatitis C virus (HCV). Sofosbuvir is used to treat hepatitis C, an infectious condition (Richmond, Dunning, and Desmond

2007; Krystallis et al. 2012). Cirrhosis of the liver manifested itself as liver failure, gastric varices, and, ultimately, malig- nancy. Racemic atenolol from Candida antarctica is kineti- cally resolved using lipase (Siebenhaller et  al. 2017). The atenolol of enantiomeric resolution was used as an acrylate agent and an organic solvent in a transesterification proce- dure that used vinyl acetate as the reaction medium (Faraldos et  al. 2011; Carvalho et  al. 2006). The lipase from an

Aspergillus niger strain produced favorable results in the enantiomeric resolution of racemic ibuprofen (da Silva et al.

2009; Lee and Shuler 2007).

When the enzyme catalyzes enantiomerical esterification of R-ibuprofen, the enantiomer results in a higher ratio of

S-ibuprofen (Ong, Kamaruddin, and Bhatia 2005; Zhangde,

Jianhe, and Jiang 2007). In addition, Serratia marcescens lipases produce chiral drug precursors that are hydrolyzed enantioselectivity (Zhao et  al. 2008; Zanger and Schwab

2013). Enzymatic acylation of amines has been abandoned for the training of pharmacologically interesting -substituted isopropyl amines (Nordwald, Armstrong, and Kaar 2014;

Andriianova, Biglova, and Mustafin 2020). The inclusion of methyl groups on the nitrogen atom or methoxy groups on the aromatic ring is known to increase the drug’s effects (Gibbons 2012; Abdelraheem et al. 2019). All of these types solidify in the sphere of organic separation and pharmaceu- tical commerce with the employment of lipases striking for biotechnological uses on the population’s strength with the prospect of a progressive effect (Salihu and Zahangir Alam

2015; Reetz 2002). To manufacture enantiomerically pure secondary alcohol, there is multiple instances structure that encloses numerous chiral pharmaceuticals or intermediates that may be accepted out for the separation of pharma medications by acylation of alcohol or hydrolysis of esters utilizing lipases in organic solvents (Kato et  al. 2007; Liu et  al. 2010; Weissfloch and Kazlauskas 1995). As a result,

Pseudomonas cepacia lipase (PSL) used enzymatic acylation of halogen alcohols or cianoalcohols to disclose a dynamic kinetic resolution of azidoalcohols for the creation of (S)

-propranolol (Fernandez and Khiar 2003). This mixture of substances has more interest as calcium antagonists and is usually employed in the pharmaceutical business as a target in the creation of optically pure 1, 4-dihydropyridine deriv- atives, with enantiomerically pure 2-methoxy-2-phenyl eth- anol, observed as a desirable chiral auxiliary. Candida antarctica lipase B (CAL-B) can be used to resolve this primary alcohol after an E of 47 using a variety of lipases through enzymatic acylation. Paroxetine hydrochloride is a selective serotonin (5-HT) reuptake inhibitor that is used as an antidepressant (Davis et  al. 2016).

Lipases used in biosensors The breakdown of fats, oils, and phospholipids is catalyzed by the lipase and phospholipase enzymes, respectively. They can be used to detect a few diseases or contagions, such as high triglycerides or pancreatitis, and their presence or rising levels can be employed as diagnostic tools (Edwards and Tchounwou 2005). Pesticide residues containing organo- phosphorus (OP), commonly known as p-nitrophenyl pes- ticides (methyl parathion), are neurotoxic and difficult to remove from contaminated water (Ma et  al. 2018; Cheng et al. 2020; Zehani et al. 2015). For the detection of methyl parathion direct and rapid lipase@ZIF-8/chitosan/glassy

18 E. P. CHANDRA AND D. P. SINGH carbon electrode (GCE) and lipase @amine-modified

ZIF-8(An-ZIF-8),/chitosan/GCE biosensors have been devel- oped (Senbua, Mearnchu, and Wichitwechkarn 2020; de

Moura Barboza et  al. 2019; Ribeiro et  al. 2011). Lipase@

ZIF-8/chitosan/GCE and lipase@An-ZIF-8/chitosan/GCE are methyl parathion modified biosensors with a wide linear range, good reproducibility, recovery, and a detection limit as low as 0.28 M (Coelho, Silva, and Orlandelli 2021; Yang et al. 2003). The carbendazim (CBZ) insecticide was deter- mined using lamellar zinc hydroxy nitrate adorned gold nanoparticles immobilized with Ceratobasidium sp.2 lipases and then mixed into carbon paste using multi-walled carbon nanotubes (CNTs) (Chen et  al. 2019). Potentiometric bio- sensors based on Candida rugosa lipase have been devel- oped for the analysis of organophosphorus pesticides such as methyl-parathion and tributyrin (Kartal, Ali Kilin, and

Timur 2007).

Urea in milk can also be detected with potentiometric biosensors (Trivedi et  al. 2009). The urease enzyme was immobilized utilizing a polymer matrix of polyethyleneimine and poly (carbamoyl) sulfonate to capture it on an ion-sensitive membrane. The detection limit and average slope of the biosensor in the linear range were 2.510 5 mol/L and 51.7 0.5 mV/decade, respectively. Allergens in foods like milk and dairy products have sparked worries about their safety. Eissa et al. (2012) developed a label-free electrochem- ical immunological biosensor with a detection limit of

0.85 pg/mL for the detection and quantification of -lacto- globulin, a milk allergy. A chemical process and an organic film (graphene-modified screen-printed electrode) were used to make the biosensor.

Another analyte to be concerned about in milk and dairy products is bisphenol A (BPA). BPA is commonly utilized as a monomer in the manufacture of polycarbonate plastics and epoxy resin, which are commonly used to construct a variety of food packaging materials such as water and dairy bottles, as well as coating material for processed food cans (Kang and Kondo 2003; Lim et  al. 2009; Goodson et  al.

2004). Alkasir, Ornatska, and Andreescu (2012) developed a BPA-detecting enzyme-based nanoparticle functionalized electrochemical biosensor.

Melamine is a high-nitrogen organic chemical that has been illegally added to milk and its products, causing sig- nificant poisoning in humans (kidney damage). A melamine biosensor based on ionic liquid/nanoparticles/chitosan and a modified gold electrode was developed for the determi- nation of melamine in milk and its products (Rovina,

Siddiquee, and Wong, 2015). Melamine may be detected using a melamine biosensor with an exposure limit of

9.6 × 10 ‾16 M and a concentration range of 9.6 × 10 ‾15 −

3.3 × 10 ‾3 M. As a result, during food safety inspections, it can be used to look for melamine in milk products (Alizadeh et  al. 2022).

Lipases in the pulp and paper industry The paper and pulp business processes a large amount of lignocellulosic biomass per year. Triglycerides and waxes, sometimes known as hydrophilic components of wood (Pitch), combine to produce some challenges in the pulp and paper manufacturing process (Gutiérrez, Río, and Martínez 2009). One of Japan’s largest paper mills,

Nippon, developed this technology to regulate impurity from the wood pitch by affecting their hydrolysis (90 percent) utilizing C. rugosa fungal lipase (Patel et  al.

2019). To avoid the detrimental effects of the negative characteristics on the operation of the papermaking equipment Anion residue and bitumen deposition can also be treated with lipase (Demuner et  al. 2011). To increase manufacturing capacity and quality (Horchani et al. 2012), it may be necessary to eliminate the esters from pulp lipase.

Conclusions and future perspective Lipases should be able to withstand multiple reaction cycles in biotechnological applications. This stability can be improved through immobilization methods and catalytic characteristics, resulting in increased catalyst efficiency.

Protein adulteration is reduced or eliminated when enzyme purification is used. Immobilized lipase enzymes help a lot in the industrialized techniques where they’re used currently.

Oversimplification of the process reduced environmental impact, and a unique ecological approach to biochemical separation are all well-known benefits of aid. In well-established production techniques such as cocoa butter analogues goods, amino acids, bakery and confectionary, medications, and pharmaceuticals, the food sector uses the most immobilized enzymes. Enzyme catalysis on a commer- cial scale has been used in a variety of industries, including medicines and food, with recent trends in biofuel production and natural gas conversion. Immobilized enzymes will be employed more widely in the future, bringing in the era of enzyme technology.

Acknowledgements The authors wish to thank and dedicate this review manuscript to

Late Prof (Dr.) Devendra P. Singh Department of Environment Science,

BBA University, Lucknow for their valuable suggestion during the preparation of this manuscript.

Disclosure statement No potential conflict of interest was reported by the author(s).

Funding The author(s) reported there is no funding associated with the work featured in this article.

Abbreviations:

CBZ Carbendazim EMC Enzyme-modified cheese EMDI Enzyme-modified dairy ingredients

Critical Reviews in Food Science and Nutrition 19 EL

Ergosterol linoleate ELn Ergosterol linolenate EO Ergosterol oleate

FFA Free Fatty Acids GCE Glassy carbon electrode HCV

Hepatitis C virus HMF Human Milk Fat OP Organophosphorus

PC Phosphatidyl-Choline PUR Polyurethane TB Tuberculosis

ORCID Enespa http://orcid.org/http://orcid.org/0000-0003-3673-1641

Prem Chandra http://orcid.org/http://orcid. org/0000-0003-0281-9178

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