Next-generation biocatalysis: Integrating enzyme engineering, synthetic biology, and information technologies for industrial applications

Introduction

Biocatalysis, or the use of natural and engineered enzymes to perform chemical transformations, has emerged as one of the cornerstones of biotechnology (Bornscheuer et al., 2012; Fryszkowska et al., 2020) [9,35] . Enzymes are proteinous biocatalyzers that control the rate of biological reactions as well as complex metabolic pathways and other cellular processes (McLure et al., 2022; Du et al., 2023; Brandenberg et al., 2019) [10,27,76] . It is well known that enzymes are highly effective and versatile biocatalysts. When enzymes are present, reactions happen far more quickly than when they are lacking (Dotsenko et al., 2023) [25] . These biological molecules are extremely specific in identifying the chemicals that need to be transformed. Enzymes have developed independently over time, resulting in novel biocatalysts with superior properties and a variety of biological uses (Musil et al., 2018; enzymes have developed in industrial biotechnology scope to include pharmaceuticals, food and beverages, agriculture, biofuels, textiles, waste, and remediation (Vidya et al., 2017; Woodley, 2018) [121,123] . Nevertheless, natural enzymes present substantial disadvantages that limit immediate application at scale. Natural produced enzymes generally lack stability against extreme process conditions, have limited substrate range, efficiency recovering and reusing the enzyme, and costly production, posing significant challenges for commercialization (Hailes et al., 2007) [43] . To mitigate these challenges, the biocatalysis field has transitioned swiftly from historical enzymes utilized at either natural or altered levels, moving toward next generation concepts, merging disciplines of protein engineering, synthetic biology, enzyme immobilization, and computational software (Khan et al., 2019; Zhao et al., 2018; Gao et al., 2019) [38,55,137] . Recent advancements in protein engineering have permitted the emergence of stable enzymes with improved thermostability, solvent tolerance, and increased substrate specificity. Directed evolution and rational design methods enable us to modify the structure of an enzyme to provide an industrial need. In addition, the creation of chimeric or bifunctional enzymes has generated multifunctional catalysts capable of complex reactions within one reaction system. Simultaneously, synthetic biology has sparked a revolution in enzyme production and pathway optimization. Through reprogramming microbial hosts and creating artificial metabolic circuits, synthetic biology offers scalable strategies for enzyme synthesis and utilization that are more sustainable (Gao et al., 2007) [39] . In conjunction with immobilization technologies that improve stability and reusability, these strategies are practically beneficial for continuous and large-scale bioprocessing. Enzymes are perfect candidates for pharmaceutical applications because of their inherent specificity and compatibility with mild reaction conditions (Yi et al., 2021) [128] . In addition to improving reaction selectivity, using enzymes in drug discovery and synthesis aligns pharmaceutical operations with green chemistry concepts, reducing their environmental impact (Bell et al., 2021) [4] . The use of natural or artificial enzymes to speed up chemical reactions is known as biocatalysis, and it has become a potent and sustainable substitute for conventional chemical catalysis. Protein engineering has enabled scientists to modify enzymes for certain processes and improve their stability through directed evolution and rational design, opening the door to new uses in pharmaceutical synthesis (McIntosh et al., 2021; Chen et al., 2020; Sheldon et al., 2017) [16,75,108] . The rational design of enzymes with increased catalytic activity has been made easier by computational biology, which has also improved our comprehension of enzymatic operations. Enzymes are very appealing in industrial biotechnology because of their special capacity to offer strong chemo-, regio-, and stereoselectivity. Enzymes normally function in mild reaction conditions, such as ambient temperature, air pressure, and aqueous solutions, in contrast to harsh chemical catalysts that frequently ask for high temperatures, extreme pressures, or hazardous solvents. In addition to lowering energy use and the production of hazardous waste, this is in line with the global movement toward sustainable manufacturing methods and green chemistry (Leresche et al., 2006; Pollard et al., 2007) [62,87] . Over the past thirty years, the use of enzymes in pharmaceuticals, agrochemicals, food and drink, detergents, textiles, bioenergy, and environmental bioremediation have grown dramatically. These days, industrial biocatalysis is essential to the synthesis of chiral intermediates, highfructose syrups, bioethanol, eco-friendly detergents, and active pharmaceutical ingredients (APIs). In order to provide a solid basis for their later incorporation into various industrial processes, early research focused on isolating these specialized enzymes and describing their enzymatic characteristics (Tatineni et al., 2008; Zhang et al., 2016) [113,134] . They are currently employed in many different industries. For example, in agriculture, keratin-rich waste is converted into protein-rich animal feed and nutrient-dense biofertilizers (Saeed et al., 2024; Xu et al., 2022) [99,127] , encouraging resource recycling and lowering pollution levels. Additionally, waste management systems use enzymatic recycling to decrease the environmental impact of conventional disposal methods like landfilling and incineration (Jagadeesan et al., 2020; Srivastava et al., 2020) [53,111] . The global enzyme business is still expanding because of advancements in enzyme engineering technology and rising demand for sustainable processes. Despite these developments, there are still certain limitations on the industrial use of enzymes. Therefore, next-generation biocatalysis is an amalgam of cutting-edge disciplines like computational sciences and synthetic biology with conventional enzymology. In addition to discussing the revolutionary significance of information technology, this study attempts to give a thorough summary of recent developments in enzyme stabilization, regeneration, protein engineering, immobilization, and synthetic biology-driven biocatalyst design. Particular attention is paid to their usefulness in a variety of industries, as well as the difficulties and potential paths for making biocatalysis the foundation of environmentally friendly and technologically advanced industrial processes.

Challenges of Native Enzymes in Industrial

Applications: The direct use of natural enzymes often fails to meet the rigorous demands of large-scale processes. Native enzymes are typically evolved to function under physiological conditions within living organisms, which are markedly different from industrial environments. Challenges include:  Limited stability: Most enzymes lose activity under high temperatures, extreme pH, or in the presence of organic solvents commonly used in industrial reactions.  Narrow substrate specificity: Natural enzymes often act on a limited range of substrates, restricting their applicability for diverse chemical transformations.  Cofactor dependence: Many industrially relevant enzymes require cofactors (e.g., NADH, NADPH, FAD), which are expensive and unstable, making continuous processes uneconomical.  Recovery and reuse difficulties: Enzymes used in homogeneous systems are difficult to separate and recycle, increasing production costs.  Short shelf-life: Storage and operational stability limit long-term industrial deployment.

These challenges have necessitated a shift from the direct use of natural enzymes toward next-generation biocatalyst systems, which combine molecular biology, synthetic biology, materials science, and computational approaches.

Evolution of Next-Generation Biocatalysis

Scope and Objectives of This Review

This review aims to provide a comprehensive overview of recent advances in enzyme-based biocatalysis, with a focus on their stabilization, engineering, and application in technology-driven processes. The objectives are: 1. To critically analyze the advantages and disadvantages of biocatalysis in industrial biotechnology. 2. To examine strategies for stabilization and regeneration of enzyme systems, including immobilization, cofactor recycling, and material-assisted stabilization. 3. To discuss advances in protein engineering, including rational design, directed evolution, chimeric and bifunctional enzyme construction, and semi-synthetic enzyme development. 4. To highlight the role of modern information technologies AI, machine learning, molecular modeling, and bioinformatics in accelerating enzyme discovery and design.

5. To explore practical industrial applications across pharmaceuticals, food, bioenergy, textiles, and environmental remediation. 6. To present future perspectives on integrating enzyme engineering with synthetic biology and computational platforms for sustainable industrial transformation.

Through this framework, the review positions nextgeneration biocatalysis as a driving force in shaping the future of industrial biotechnology. By emphasizing both technological innovation and practical significance, it bridges the gap between academic research and real-world applications, offering valuable insights for researchers, industry practitioners, and policymakers.

Enzyme Stabilization and Regeneration Strategies

The low stability of biocatalysts in chemical conditions is one of the main obstacles to their practical use (Woodley et al., 2019) [124] . According to Rigoldi et al. (2018) [95] , thermostability is a critical issue that is controlled by numerous parameters and nearly always necessitates modifications to the peptide chain of amino acid residues that make up the protein. We changed the potentially important serine residues found in structure and sequence comparisons with the thermophilic homologous CelA from Clostridium thermocellum in order to increase the thermostability of chitosanase-CsnTS. This engineered CsnTS can be used in the industrial scale-up production of chitooligosaccharides (Zhou et al., 2021) [138] . Stabilization and regeneration of enzymes are critical for industrial application of biocatalysis. While enzymes confer great specificity and efficiency, they evolved to function optimally at mild physiological conditions and when subjected to industrial conditions, especially extremes in temperature, pH, solvents, and mechanical stress. Enzymes tend to denature, aggregate or be inactivated very quickly. Thus, the economic value of enzyme-based processes rests heavily on the stabilization, reusability and operational lifespan of enzymes. Many physical, chemical', biological, and engineering-based strategies have been developed to overcome these challenges.

Enzyme Immobilization

To allow to improve the catalytic activity, stability, and reusability of soluble enzymes, enzyme immobilization is defined as limiting the mobility of the enzyme in a specific area of space (Kujawa et [8,58,96] . They can be retrieved, reused, and limited to a specific area of space with retained catalytic activity (Das et al., 2023) [21] . According to Zhang et al.

(2023) [133] , an insoluble enzyme is immobilized. This type of heterogeneous biocatalysis is distinct. An immobilized enzyme affects some features, including catalytic activity, thermal stability, and storage stability (Abdel-Hameed et al., 2022) [2] . [100] . The enzyme, a carrier material, and the attachment technique are the three primary parts of an immobilized enzyme system (Cowan et al., 2011) [20] . Because the immobilization procedure has a significant impact on the biocatalyst's characteristics, the immobilization technique will have an impact on the stability and catalytic activity of the biocatalyst (Tang et al., 2022) [112] .

Adsorption

Adsorption is the mechanism whereby an enzyme builds up on a solid support surface through intermolecular interactions (Ramalho et al., 2023; Kharazmi et al., 2023) [56, 92] . It also eliminates the need to alter the surface and linkers like glutaraldehyde and cysteine for cross-linking or 1-ethyl-3-(3-dimethylaminopropyl) carbodiimide for covalent bonding (Kumar et al., 2020; Smith et al., 2020) [60,109] . Hydrogen bonds and electrostatic interactions are involved in surface adsorption interactions with enzymes (Feng et al., 2023) [33] . The adsorption process involves the interaction of enzymes with support surfaces. Therefore, in order to ensure high and repeatable enzyme coverage on a support to accomplish typically mild, easy, economical, rapid, good performance, and good reusability, the polarity and charge of the enzyme will be crucial attributes (Aghabeigi et al., 2023) [3] . Nevertheless, adsorption-based enzyme immobilization has drawbacks, such as enzyme leaching and possibly lower enzyme binding to the support than covalent binding (Nageib et al., 2022) [81] .

Cross-Linking

The act of creating a covalent bond between two or more molecules is known as cross-linking (Moosavi et al., 2022; Paiva et al., 2024; Feng et al., 2022) [34,79,85] . One commonly used technique for immobilizing enzymes is the use of cross-linked enzyme aggregates. It frequently results from an activation or surface modification process (Narayanan et al., 2022; Hu et al., 2023; Jaafar et al., 2022) [48, 52, 82] . The most popular method of initial surface modification of inorganic substrates is silanization, which is the creation of organic functional groups on a surface using an organofunctional silane reagent. Surfaces having natural amino groups or those coated with p-nitrobenzoyl chloride can be treated to produce aldehyde or arylamine groups. (Shaarani and others, 2016) [104] . Purification from an aqueous buffer and cross-linking the resulting physical aggregate of enzyme molecules are the only steps in this straightforward process, which is easily optimized (Liu et al., 2021) [69] . Cross-linking produces a stable, immobilized enzyme that can be used as a biocatalyst. However, the main disadvantages of gel-like cross-linked enzymes are frequently poor activity retention, challenges with reuse, low mechanical stability, and general handling problems (Li et al., 2020) [63] .

Entrapment or Encapsulation

Enzymes that are caged with covalent or non-covalent bonds are said to be immobilized through entrapment. These matrices preserve the biocatalyst's characteristics while minimizing conformation change (Reshmy et al., 2021; Tavernini et al., 2020; Xing et al., 2022) [94,114,126] . The enzyme is retained while allowing substrates and products to move via a network of polymers (Abbasi et al., 2023) [1] . After entrapment, enzymes are not bound to the polymeric matrix, but their diffusion is constrained. Compared to an enzyme that is physically adsorbed, an entrapped enzyme is more stable. Although enzyme activity is either maintained or increased, entrapment immobilization is easier to achieve than covalent bonding (Rashid et al., 2022; Ganonyan et al., 2020; v et al., 2022) [37,93] . Enzyme immobilization through physical entrapment has many uses and might not interfere as much with the characteristics of the native enzyme (Rivero Berti et al., 2021) [5] . To guarantee that the substrate and product can diffuse into and out of the polymer matrices, encapsulation techniques must consider the chemical conditions of the polymerization matrices, the pore diameter and volume, and the compatibility of enzymes with the pore matrices. Since many enzyme encapsulations do not cause a major reduction in enzyme activity following immobilization, catalytic activity retention is rather common (Datta et al., 2021) [22] .

Covalent Bonding

The formation of covalent bonds between each molecule of an immobilized enzyme and the support creates a section of the enzyme where the residues attached to the support stay in place during any movement or dislocation of the enzyme molecule caused by hazardous conditions like heat, extremely high or low pH, and organic solvents (Meng et al., 2023) [77] . The highly rigidized region can be relatively large if the support's structure is sufficiently compatible with the enzyme's surface (for example, a flat support structure and a globular protein that is moderately flexible). This establishes the essential stabilization of the enzyme molecule as protection against any harmful conditions, such as extremely high acidic or alkaline conditions and temperature (Guisan et al., 2022) [42] . Parmegiani Marcucci et al. (2022) [73] used the SBA-15, which was modified with tin (SnS8 and SnS23) and had varying pore diameters (S8 with 9.1 nm and S23 with 23 nm), to study the immobilization of Burkholderia cepacia lipase via covalent bonding. The immobilized enzyme had the highest specific activity on the support with the largest pore size (25 nm). A strong link between lipases and modified silicas was demonstrated by the absence of lipase in the buffer solution used for lipase desorption during the leaching tests. The successful production of PNGase F with a glutamine tag in Escherichia coli and the site-specific covalent immobilization of PNGase F with this unusual tag via microbial transglutaminase (MTG) are reported by Zhang et al. (2023) [133] . Using MTG, PNGase F was immobilized from the glutamine tag and covalently and sitespecifically transformed into primary amine-containing magnetic particles. Immobilized PNGase F may deglycosylate substrates and exhibit strong thermal and reusability capacity with enzymatic efficiency similar to that of its soluble counterpart (Tvorynska et al., 2022) [118] .

Sampaio et al.,

2022 [100] Immobilisation of protease via covalent bonding on chitosan.

Chitosan as a carrier; glutaraldehyde and ethylenediamine as modification agents and crosslinkers.

The catalytic activity of immobilised enzymes is equivalent to that of free enzyme (pH 9 and 60 °C); however, the immobilisation procedure broadened the optimum temperature range of enzyme activity (50-70 °C).

After three cycles of use, the immobilised enzyme only maintained 47.08% of its initial activity.

Ramalho et al.,

2023 [92] Laccase was immobilised on a low-cost, nanosized magnetic biochar (L-MBC) via adsorption, precipitation, and cross-linking.

Bagasse biochar as a carrier; ammonium sulfate as a precipitation agent; glutaraldehyde as a cross-linker.

The magnetic biochar could immobilise a substantial quantity of enzymes with increased catalytic activity (2.251 U per mg MBC), better stability, improved storage stability, pH tolerance, and thermal stability over free laccase.

The catalytic activity of immobilised laccase decreased more than 50% of its initial activity at pH 5.5 and 6.

Zhang et al., 2020 [136] Encapsulation of laccase using alginate, alginate-silica and silica sol-gel.

Tetraethyl orthosilicate (TEOS) and alginate as carriers.

The experimental results showed that incorporating silica into alginate resulted in a better (70%) encapsulation efficiency (EE) for the laccase extract than for the alginate alone (59%). Furthermore, encapsulating the laccase extract in sol-gel resulted in an increase in its catalytic activity, as well as a 90% rise in the EE. The alginate and solgel matrices also improved laccase catalytic efficiency compared to free laccase, with kcat values of 89.9 (alginate), 63.7 (alginate-silica), and 56.9 min -1 (silica sol-gel), respectively.

After three reaction cycles, the catalytic activity of immobilised laccase with alginate-silica was decreased by 50%. Difficult to achieve desired results.

Gill et al., 2018

Guisan et al., 2022 [42] Fig 1: Advantages and challenges of enzyme immobilization (Mohidem et al., 2023) [78] .

Protein Engineering-Based Stabilization

Enzyme robustness can be directly increased through genetic manipulation. Examples of employing protein engineering to control plant immunology and energy conversion have been documented, but there are very few instances of secondary plant metabolic regulation based on protein engineering. It was established that the modified rice RGA5 could specifically recognize and bind the pathogen effector protein AVR-PikD, which is not recognized by the original RGA5, extending the crop's resistance to disease by engineering the heavy metal-associated (HMA) structural domain of rice RGA5 (Maqbool et al, 2015) [72] . RbcS is a tiny subunit of Rubisco. In rice Rubisco, eliminating the OsRbcS (rice RbcS) multigene family and substituting OsRbcS entirely with SbRbcS (sorghum RbcS) increased the rate of photosynthetic CO2 assimilation and photosynthetic efficiency (Matsumura et al., 2020) [74] , which can significantly increase plant growth and productivity. Protein engineering offers a foundation for the development of new enzymes that can create a quick system for protein screening and modification in vivo, which offers the potential to reshape the secondary metabolic network in plants. CRISPR/Cas9-assisted engineering now offers technical support for the replacement and removal of pathway enzymes.

Protein Engineering of Enzymes

Rational Design, Directed Evolution, Chimeric and Semi-Synthetic Enzymes Thus, enabling the development of customized catalysts with enhanced activity, stability, and substrate selectivity, enzyme engineering has transformed contemporary biotechnology. Even though natural enzymes are quite effective in physiological settings, they frequently exhibit limitations in industrial settings with high temperatures, severe pH, organic solvents, or high substrate concentrations. Protein engineering offers answers by altering the structures and sequences of enzymes to achieve the appropriate stability and catalytic properties. Protein engineering techniques can be broadly classified into two groups: directed evolution and rational design. Hybrid approaches, such as chimeric proteins and semi-synthetic enzymes, are becoming more and more significant.

Rational Design of Enzymes

After altering the natural metabolic network of plants, metabolic engineering seeks to improve plant resilience and produce a value-added chemical, among other beneficial functional traits (Lynch et al., 2021) [70] . The most successful method of metabolic engineering is now changing the synthetic pathways in plants using genetic engineering techniques (Fu et al., 2018) [36] . With improved stability, increased catalytic activity, and most importantly a broader range of protein applications, protein engineering methods and approaches have made it a reality quickly and easily, in contrast to genetic engineering techniques that obtain targets through mass disorderly screening (Cai et al., 2015) [12] . This offered a great target for modifying the structure and/or content of lignin in plants. A better phenotype can be obtained in plants by substituting enzymes with a higher catalytic rate for those with a low catalytic rate and weak affinity and specificity. The sensing of arbuscular mycorrhizal fungus in rice is mediated by the LysM receptor heteromer OsMYR1/OsLYK2 and OsCERK1. The recognition of nodulation factors in rice was greatly enhanced by substituting the ectodomains of OsMYR1 and OsCERK1 with those from the homologous Nod factor receptors MtNFP and MtLYK3 of Medicago truncatula. This also increased calcium oscillations in response to Nod factors (He et al., 2019) [45] .  Introduction of disulfide bonds, salt bridges, and hydrophobic residues improves thermostability.  Alteration of surface charges enhances solvent tolerance.  Computational modeling and molecular dynamics predict stabilizing mutations prior to laboratory testing.

Rational design is a knowledge-driven approach that relies on structural, biochemical, and computational data to predict beneficial mutations.

Principles of rational design  Based on three-dimensional structures obtained via Xray crystallography, cryo-EM, or homology modeling.  Involves identifying catalytic residues, flexible regions, and stability determinants.  Uses computational tools such as molecular dynamics simulations, docking, and energy minimization.

Strategies in rational design

 Active-site redesign: Altering residues around the catalytic pocket to expand substrate specificity (e.g., redesigning lipases for biodiesel synthesis).  Stability enhancement: Introducing disulfide bonds, salt bridges, or proline residues to increase rigidity.  Surface charge modification: Enhancing tolerance to pH or ionic strength by altering surface amino acids.  Cofactor switching: Engineering enzymes to use alternative, cheaper cofactors (e.g., switching from NADPH to NADH).

Advantages and limitations

 Precise and predictable when sufficient structural data is available.  However, limited by the accuracy of computational models and incomplete understanding of protein folding dynamics.

Directed Evolution of Enzymes

This is a potent method for creating efficient enzyme catalysts that can carry out a range of biocatalytic tasks in different sectors. In contrast to rational design, directed evolution allows for relatively quick enzyme modification and does not require in-depth knowledge of the structurefunction correlations. Additionally, it is similar to the process of natural evolution (Zeymer and Hilvert, 2018) [131] .

The general process of directed evolution is quite straightforward: a library of mutants is created, and those that possess the desired trait are identified using an appropriate selection or screening procedure. The biosynthetic machinery of the cell is then utilized in an in vitro approach or engineering in a heterologous or homologous host organism to produce the comparable enzyme changes. Until an enzyme with the desired characteristics is obtained, the process of mutant creation and selection is repeated multiple times. Although useful, thorough prior knowledge of the enzyme is not required because directed evolution does not require any foresight regarding the mutation locations, which is an advantage over rational design. Frances H. Arnold's guided evolution of enzymes earned her the 2018 Chemistry Nobel Prize. George P. Smith and Sir Gregory P. Winter split the prize for their phage display of peptides and antibodies. The directed evolution approach is a reliable protein engineering technology that produces random mutations in a target gene and then quickly selects protein variants based on advantageous characteristics for certain uses (Vidal et al., 2023) [120] . An experimental, iterative method that simulates natural selection in a lab setting is called directed evolution. It entails creating genetic diversity and screening for desired features rather than depending on past structural knowledge.

To create DNA libraries, mutations are made to the coding gene of the biocatalyst that needs to be modified. By the end of the procedure, enzymes with better qualities are found by screening and selection, and the polymerase chain reaction is then applied to the corresponding genes (Zeng et al., 2020) [130] .  Iterative random mutagenesis and selection have yielded enzymes that remain active at temperatures above 100 °C, tolerate organic solvents, or resist high substrate/product concentrations.  Industrial lipases, cellulases, and oxidoreductases have been optimized using this method.

Methodology

 Library generation: Random mutagenesis (error-prone PCR, chemical mutagens) or recombination (DNA shuffling).  Screening/selection: High-throughput assays identify improved variants.  Iteration: Multiple cycles lead to cumulative improvements.

Applications

 Development of thermostable cellulases for biofuel production.  Engineering of oxidoreductases for synthesis of pharmaceutical intermediates.  Evolution of esterases and proteases with resistance to organic solvents.

Advantages

 Powerful for improving properties without structural knowledge.  Requires robust screening platforms; may generate "hidden" beneficial mutations not predicted by rational design.  Often combined with computational methods for semirational approaches.

De-novo design: De novo enzyme design is the process of creating a catalytic activity based on the exact residues of amino acids necessary for the stability of the transition state, as determined by the score derived from the configuring atoms. Both the transition state activation energy and the biocatalysis activation energy are decreased by the denovo design. Active site design is the first step in the de novo design process. In order to create new active sites for a certain transition state in a reaction, all of these designs rely on complex computational calculations. Generally speaking, the initial step is to develop a catalytic mechanism for the desired reaction. For transition state stability, essential residues that constitute a minimum active site are then added (Quin and Schmidt-Dannert, 2011) [90] . The active site supporting protein backbones are then found using a similarity searching technique like the Rosetta Match hashing method (Zanghellini et al., 2006) [129] . The geometry of catalysis and the binding energy of the transition state are analyzed using in silico screening (Smith et al., 2014) [109] . Table 3 contains a list of computer-aided designed enzymes. Recently, Lapidoth et al. proposed an effective automated method for building enzymes by combinatorial backbone assembly. Although catalytic residues are conserved, this method uses a variety of homologous but structurally distinct enzymes, assembles novel protein backbone combinations, and uses Rosetta software to arrange and optimize the amino acid residues. This method was used to design 43 and 34 proteins, respectively, for two distinct families of enzymes with TIM-barrel folds: GH10 xylanases and phosphotriesterase-like lactonases. Seven PLL designs and 21 GH10 designs were quite active. Accordingly, combinatorial-backbone assembly may be a suitable option for creating stable and active enzymes for industrial processes (Lapidoth et al., 2018) [61] . Figure 2 illustrates the main approaches of enzyme engineering. [31] Serine esterase engineering Quantum mechanical calculation + point mutations Substantial lytic activity toward p-nitrophenyl acetate Li et al., 2020 [63] Fig 2: Strategies of enzyme engineering (Madhavan et al., 2021) [70] .

Chimeric Enzymes

Chimeric enzymes are generated by fusing domains from different proteins to create hybrid catalysts with novel functionalities. Development of chimeric proteins with subdomains of other proteins is another way to improve thermostability. In this manner, positive traits from various parents might be blended. For instance, subdomains from Baeyer-Villiger monooxygenases (BVMOs) with varying stabilities were combined to create chimeric versions of BVMOs. While maintaining or even enhancing the substrate selectivity of a thermolabile BVMO, certain chimeric enzymes were 10-15 °C more stable than their thermolabile parent enzymes (van Beek et al., 2012) [119] . A number of stabilizing chimeras between various xylanases and between xylanases and glucanases have been produced (Liu et al., 2012) [67] . A chimera produced by grafting loops from an archeal enzyme onto a human aldo-keto reductase demonstrated that advantageous characteristics of archeal and human proteins might be combined, in this case stability and activity (Campbell et al., 2013) [13] . By merging the domains of two mesostable polyol dehydrogenases, it is even possible to create such a stabilized chimera without utilizing a thermostable variation (Hulf et al., 2012). Compared to its parents, the resulting chimera was 12 °C more stable. Lastly, it was demonstrated that proteins with minimal sequence similarity (<30%) can also form chimeric enzymes by substituting portions of a (βα)8-barrel protein with portions of flavodoxin-like proteins (Shanmugaratnam et al., 2012) [106] . Because the structure of flavodoxin-like proteins resembled a portion of the (βα)8-barrel, the replacement was feasible.

Another study used an active learning algorithm in conjunction with the SCHEMA approach to predict a huge collection of stabilizing chimeras by analyzing data from a training set of chimeras (Romero et al., 2012) [98] . Enzymes with improved long-term stability were created using this technique, which showed a negative association between a chimeric enzyme's isoelectric point and its long-term stability. A chimeric P450 enzyme that was 14.5 °C more stable than its parent was recently developed (Hwang et al., 2013) [51] .

Design principles  Domain shuffling:

Combining catalytic domains from one enzyme with stability-enhancing domains from another.  Fusion with affinity tags: Enhancing immobilization or recovery (e.g., His-tags, cellulose-binding domains).  Multifunctional chimeras: Engineering bifunctional enzymes capable of catalyzing two sequential reactions.

Examples

 Fusion of cellulase catalytic domains with carbohydrate-binding modules (CBMs) for improved cellulose degradation.  Chimeric lipases fused with hydrophobins to increase activity at water-oil interfaces.  Enzyme fusions in metabolic engineering to create artificial metabolic pathways.

Semi-Synthetic Enzymes

The lack of an enzyme with the required catalytic activity or substrate selectivity, as well as their instability in biotechnological processes, frequently limit the growing demand for enzymes as highly selective, mild, and environmentally benign catalysts. These issues included various classes of enzyme mimics as well as genetically modified enzymes. Here, we outline chemical enzyme engineering's potential: Native enzymes can be altered using simple tools and chemical processes to create "semisynthetic" enzymes. Thus, the catalytic adaptability of a synthetic active site is paired with the great substrate selectivity of the enzymatic peptide framework. We use the idea of the semisynthetic peroxidase seleno-subtilisin to demonstrate the possibilities of chemically modified enzymes. Subtilisin, a serine endoprotease, was first crystallized and cross-linked with glutaraldehyde to produce cross-linked enzyme crystals that were shown to be extremely stable and insoluble in organic solvents or water (Feng et al., 2022) [34] . Second, an oxidized derivative of selenocystein (Enz-SeO2H) was produced chemically from serine 221 in the active site (Enz-OH). Consequently, the hydroperoxide reduction was catalyzed by the former proteolytic enzyme, which also acquired peroxidase activity.

Given the well-known selectivity of subtilisin, the substrate selectivity of seleno-subtilisin was anticipated because the binding sites of semisynthetic peroxidase and protease were identical (Häring et al., 1999) [44] .

Design Approaches 1. Unnatural amino acid incorporation:

Provides novel reactivity or enhanced stability. 2. Artificial metalloenzymes: Insertion of synthetic metal complexes into enzyme frameworks for unique catalytic reactions.

Polymer conjugation:

Attaching polymers or nanoparticles to enzymes for improved solubility, stability, and recyclability.

Industrial Applications

 Semi-synthetic peroxidases for oxidative degradation of pollutants.  Artificial metalloenzymes in enantioselective chemical synthesis.  Hybrid enzyme systems for green chemical manufacturing and bioremediation.  Artificial metalloenzymes used in enantioselective hydrogenation and C-C bond formation. [103,122] found the amino acid residues in the Rossmann fold that give glutathione reductase selectivity for NADP+ over NAD+. When Ehsani et al. (2009) [32] compared the structure and sequence of NADP (H)-dependent yeast alcohol dehydrogenase to that of NAD(H)-dependent Bdh1, they found three alterations in the amino acid sequence that resulted in a variant of Bdh1 with a shift toward NAPH cofactor specificity. Alcohol dehydrogenase D is a member of the aldo-keto reductase (AKR) protein family and is an extremely effective and temperature-tolerant alcohol dehydrogenase. Several researchers expanded the selectivity of aldo-keto reductase to cofactor by introducing mutations in the cofactor's binding pocket (Sanli, 2004; Campbell et al., 2010) [15,101] . This enzyme's steady state kinetic research provided fresh insights into the conformational shifts and ensuing biocatalyst performance. A biofuel cell was created using this enzyme-cofactor pair when the maximal dynamic mutant catalyst simultaneously gained activity with a nonnative cofactor, nicotinamide mononucleotide, or NMN(H) (Campbell et al., 2012) [14] . By adding the active site loop region from human aldose reductase, the thermostable AdhD enzyme's substrate selectivity was altered. Both substrate specificity and cofactor specificity were altered by this engineering strategy. Clostridium thermocellum is a powerful candidate microbe in integrated bioprocessing for lignocellulosic conversion to ethanol. It is an obligatory anaerobe and a highly thermophilic bacterium. The primary economic disadvantage is intolerance to ethanol. According to genome sequencing, this acteria's mutant bifunctional acetaldehyde-CoA/alcohol dehydrogenase gene (adhE) is responsible for its ethanol tolerance (Brown et al., 2011) [11] .

Application of Enzyme Engineering in Food, Pharmaceuticals, Biofuels, and Environmental Biotechnology: Enzyme engineering has transformed industrial biotechnology by enabling tailored biocatalysts with improved efficiency, stability, and substrate specificity. These engineered enzymes are now pivotal in sectors such as food processing, pharmaceuticals, biofuels, and environmental remediation, where they offer greener, safer, and more cost-effective alternatives to conventional chemical processes. Future developments are likely to integrate artificial intelligence (AI), machine learning, and synthetic biology to accelerate enzyme design. Computational prediction of stabilizing mutations combined with high-throughput directed evolution will lead to faster discovery of optimized biocatalysts. Furthermore, hybrid enzymes that merge natural biocatalysis with synthetic catalytic capabilities are expected to open entirely new avenues for green and sustainable chemistry.

Food Industry Applications

Starch and other classes of food are the chief source of energy for non-photosynthetic organisms and plants. Depending on the situation, these food classes could be processed chemically or enzymatically to create a variety of products for the food and animal feed industries. Because of their complexity, these molecules used as raw materials need a variety of enzymes to properly break them down into smaller components. By engineering these enzymes used in this process, it may be possible to improve their selectivity or specificity, push a reaction in the desired direction, and ultimately stabilize their activity toward reaction conditions necessary for large-scale synthesis or production. Duan et al. (2013) [29] modified pullulanase from Bacillus deramificans by changing Asp437 to His437 and Asp503 to Tyr503. This mutation increased the thermostability of the enzyme, which is used to make fructose and glucose syrup (Duan et al., 2013) [29] . However, Gao et al. transformed mesophilic mannose isolated from Bacillus subtilis into the thermophilic enzyme using a molecular cyclization method (Gao et al., 2019) [38] .

Medical and pharmaceutical applications

The production of the antidiabetic sitagliptin by Codexis and Merck is a very promising example of protein engineering (Choi et al., 2015; v et al., 2011; Savile et al., 2010) [18,102] . R-selective transaminase, an enzyme created through directed evolution, was used in the procedure. This enzyme produced sitagliptin with a 99.95% enantiopurity by asymmetrically aminating prositagliplitin ketone (Savile et al., 2010) [102] . A modified monoamine oxidase is used to create the chiral amine boceprevir. It is used to treat chronic hepatitis C. Researchers found that water consumption decreased by 40% and process yield rose by 150% (Li et al., 2012). The reduction of a ketone intermediate into a chiral alcohol is catalyzed by LactoBacillus kefir ketoreductases (KRED). Intermediates for a variety of medications, such as montelukast, atorvastatin, duloxetine, ezetimibe, crizotinib, and phenylephrine, are produced by ketoreductase-mediated mechanisms (Huisman et al., 2010; Huisman and Collier, 2013) [49,50] . Techniques like directed evolution and process optimization are examples of potential KRED improvements. With a value exceeding 99.55%, KRED exhibits remarkable enantioselectivity (Huisman et al., 2010) [50] . By saturating the active site of a KRED enzyme, Zhang et al. created two mutants (P170H/L174Y and P170R/L174Y) that made it easier to reduce 3-(dimethylamino)-1-phenylpropan-1-one and 3-(dimethylamino)-1-(2-thienyl)-propan-1-one to their (R)-γamino alcohols with a maximum enantiomeric excess of 95% (Zhang et al., 2015). Chiral γ-amino alcohols are essential for the synthesis of certain serotonin reuptake medications, such as fluoxetine and tomoxetine (Zhang et al., 2015) [132] . Because enantiomers of the same drug may exhibit different biological activities and toxicities, there is an increasing need for chiral compounds (Patel et al., 2018) [86] . Pharmaceutical and medical research is paying attention to the creation of diagnostic enzymes. An engineered fructosyl peptide oxidase (FPOX) was used by Shahbazmohammadi et al. as a diagnostic enzyme for diabetes mellitus (Shahbazmohammadi et al., 2019) [105] . [105] LactoBacillus kefir (Keto-reductases, KRED)

Quantum mechanical calculations and molecular dynamics simulations

Enantioselective reduction of 3-triacyclopentanone and 3-oxacyclopentone Enhanced enantioselectivity

Glu145 and Ala94Phe

Noey et al., 2015 [83] Sporobolomyces salmonicolor AKU4429 (Carbonyl reductases, KRED) Directed evolution β-Amino ketones enantioselective reduction

Enhanced activity and improved enantioselectivity

Pro170His-Leu174Tyr and Pro170Arg-Leu174Tyr

Zhang et al., 2015 [132] Herpes simplex virus type 1 (Thymidine kinase, TICKLE switch)

Directed evolution Cancer treatment Enhanced specificity HSV-TK/CH1 fusion Shelat et al., 2017 [107] Biofuel and Bioenergy Applications In recent years, the demand for energy has been rising significantly worldwide. Investigating more ecologically benign and potentially renewable alternative energy sources is essential as fossil fuels cannot be replenished. The production of biofuels is becoming more and more popular. Enzymes are useful for producing biofuel because they can break down a variety of substrates to produce ethanol and biodiesel. However, many enzymes do not possess the intrinsic robustness needed for commercial use (Prajapati et al., 2018; Otte and Hauer, 2015) [84,88] . Enzymes called xylanases are widely used in the generation of biofuel. The main component of plant biomass that causes hemicellulose degradation, xylan, can be broken down by these enzymes. Therefore, xylanases can use the agroindustrial wastes that the industry normally discards as substrates (Denisenko et al., 2017) [23] . [115] . The main byproduct of xylan breakdown, xylose, has inhibitory effects that limit the large-scale production of xylanase. Hegazy et al. used directed evolution with errorprone PCR to create genetically engineered xylanases that are xylose-tolerant in order to overcome this limitation (Hegazy et al., 2019). Three alterations were discovered in the variants: Met116Ile, Leu131Pro, and Leu133Val. The most effective mutation showed a 3.5-fold increase in the inhibition constant (ki) and a threefold increase in catalytic activity (Hegazy et al., 2019) [46] . Additionally, lipases help hydrolyze triacylglycerides into glycerol and free fatty acids. A class of enzymes known as lipases is widely used in the synthesis of biofuels. In the field of oleochemistry, fatty acids are very helpful chemicals, especially for uses like making biodiesel (Dror et al., 2014; Zorn et al., 2018) [26,139] . Dror et al. created mutations in a lipase generated from GeoBacillus stearothermophilus T6 using two techniques: structureguided consensus and error-prone PCR random mutagenesis (Dror et al., 2014) [26] . Their objective was to produce a lipase with increased stability, methanol tolerance, and transesterification activity all of which are essential characteristics in the manufacture of biodiesel. In a solution containing 70% methanol, both engineering approaches might produce enzymes with a longer half-life. [89,117] . For instance, laccases have been muddied to function in alkaline environments and are elective in degrading azo dyes commonly found in textile industry wastewater. enhanced stability and catalytic efficiency of these engineered enzymes allow them to reduce pollutant levels significantly, improving wastewater quality before discharge (Bhandari et al., 2021) [6] . Several successful applications of enzymes in bioremediation highlight their environmental impact. A notable case is the use of laccases for dye degradation in wastewater treatment. Engineered laccases that operate in alkaline conditions have been shown to break down synthetic dyes by cleaving azo bonds, reducing water toxicity by up to 90% and facilitating safer disposal (Bilal et al., 2018; Kumar et al., 2022) [7,59] . Another impactful case is the application of engineered cutinases in plastic degradation. these enzymes have been modified to hydrolyze polyethene terephthalate (PET), a major component of plastic waste. Engineered cutinases can depolymerize PET under industrial conditions, providing a potential solution for recycling plastic waste and reducing environmental pollution. Such examples underscore the value of enzyme engineering in addressing pressing environmental issues (. Radley et al., 2023) [91] .

Practical Significance, Challenges, and Future Perspectives of Engineered Industrial Enzymes Engineered enzymes have emerged as pivotal tools in industrial biotechnology, offering eco-friendly, efficient, and highly selective alternatives to conventional chemical catalysts. By combining advanced protein engineering, immobilization, semi-synthetic modifications, and modern information technologies, industries have been able to optimize reaction rates, improve product quality, and reduce environmental footprints. Despite these advancements, practical implementation still faces certain challenges that require careful consideration.

Practical Significance

Engineered enzymes play a vital role in industrial biotechnology by offering eco-friendly, highly specific, and energy-efficient alternatives to traditional chemical catalysts. Their applications span food processing, pharmaceuticals, biofuels, textiles, and environmental remediation, where they enhance productivity while reducing waste and hazardous by-products. The ability to tailor enzymes through protein engineering, immobilization, and synthetic biology has significantly improved catalytic efficiency, substrate specificity, and operational stability, making them indispensable for green chemistry and sustainable manufacturing.

Challenges

Despite their promise, several challenges limit the widespread adoption of engineered enzymes:  Stability Issues: Enzymes often lose activity under harsh industrial conditions such as extreme pH, temperature, or solvent environments.  High Production Costs: Large-scale enzyme production and purification remain expensive compared to chemical catalysts.  Scale-up Limitations: Laboratory successes may not always translate effectively into industrial-scale operations.  Regulatory and Safety Concerns: Particularly in pharmaceuticals and food applications, approval processes can be lengthy and complex.

Future Perspectives

The future of enzyme engineering is closely linked with artificial intelligence, machine learning, and synthetic biology, which will accelerate predictive design and optimization. Nanotechnology and smart immobilization matrices will enhance enzyme stability and reusability, while hybrid catalytic systems combining natural and synthetic elements will broaden reaction possibilities.

Furthermore, the development of dynamic enzyme cascades, artificial cofactors, and modular biocatalytic platforms is expected to open new frontiers in industrial biotechnology. Ultimately, these innovations will help overcome current limitations, leading to cost-effective, scalable, and sustainable enzyme-based solutions at the heart of the green industrial revolution.

Conclusions

Engineered enzymes have emerged as pivotal tools in shaping the future of industrial biotechnology, providing sustainable, selective, and efficient alternatives to traditional chemical processes. The convergence of protein engineering, immobilization strategies, synthetic biology, and advanced computational approaches has not only expanded their applicability but also addressed key challenges of performance and scalability. With the integration of modern information technologies such as AI, machine learning, and bioinformatics, enzyme engineering is shifting toward a predictive, rational, and high-throughput discipline. Looking ahead, advancements in nanotechnology, artificial cofactors, and hybrid catalytic systems are expected to further enhance efficiency and broaden industrial applications. Collectively, these developments position engineered enzymes as central drivers of the green industrial revolution, enabling ecofriendly, cost-effective, and innovative solutions across diverse sectors including food, pharmaceuticals, biofuels, and environmental sustainability.

Acknowledgement

AcknowledgmentsThe authors would like to thank a guide and organization for their guidance and support. Furthermore, we would like to sincerely thank our colleagues for their insightful discussions and constructive arguments, which were essential to the preparation of this evaluation.

References

1. Abbasi YF, Bera H, Cun D, Yang M. Recent advances in pH/enzyme-responsive polysaccharide-smallmolecule drug conjugates as nanotherapeutics. Carbohydr Polym. 2023;312:120797.
2. Abdel-Hameed SA, Ahmed SA, Mostafa FA, Almasarawi ON, Wahab WAA. Preparation and characterization of sugilite glass from basalt for αamylase immobilization, statistical optimization of the immobilization process and description of free and immobilized enzyme. Heliyon. 2022;8(7):e09975.
3. Aghabeigi F, Nikkhah H, Zilouei H, Bazarganipour M. Immobilization of lipase on graphene oxides magnetized with NiFe2O4 nanoparticles for biodiesel production from microalgae lipids. Process Biochem. 2023;126:171-185.
4. Bell EL, Finnigan W, France SP, Green AP, Hayes MA, Hepworth LJ, et al. Biocatalysis. Nat Rev Methods Primers. 2021;1(1):46.
5. Berti IR, Islan GA, Castro GR. Enzymes and biopolymers: the opportunity for the smart design of molecular delivery systems. Bioresour Technol. 2021;322:124546.
6. Bhandari S, Poudel DK, Marahatha R, Dawadi S, Khadayat K, Phuyal S, et al. Microbial enzymes used in bioremediation. J Chem. 2021;2021:8849512.
7. Bilal M, Iqbal HMN, Guo S, Hu H, Wang W, Zhang X. State-of-the-art protein engineering approaches using biological macromolecules: a review from immobilization to implementation viewpoint. Int J Biol Macromol. 2018;108:893-901.
8. Bolivar JM, Woodley JM, Fernandez-Lafuente R. Is enzyme immobilization a mature discipline? Some critical considerations to capitalize on the benefits of immobilization. Chem Soc Rev. 2022;51(15):62516290.
9. Bornscheuer UT, Huisman GW, Kazlauskas RJ, Lutz S, Moore JC, Robins K. Engineering the third wave of biocatalysis. Nature. 2012;485(7397):185-194.
10. Brandenberg OF, Chen K, Arnold FH. Directed evolution of a cytochrome P450 carbene transferase for selective functionalization of cyclic compounds. J Am Chem Soc. 2019;141(22):8989-8995.
11. Brown SD, Guss AM, Karpinets TV, Parks JM, Smolin N, Yang S, et al. Mutant alcohol dehydrogenase leads to improved ethanol tolerance in Clostridium thermocellum. Proc Natl Acad Sci U S A. 2011;108(33):13752-13757.
12. Cai Y, Bhuiya MW, Shanklin J, Liu CJ. Engineering a monolignol 4-O-methyltransferase with high selectivity for the condensed lignin precursor coniferyl alcohol. J Biol Chem. 2015;290(44):26715-26724.
13. Campbell E, Chuang S, Banta S. Modular exchange of substrate-binding loops alters both substrate and cofactor specificity in a member of the aldo-keto reductase superfamily. Protein Eng Des Sel. 2013;26(3):181-186.
14. Campbell E, Meredith M, Minteer SD, Banta S. Enzymatic biofuel cells utilizing a biomimetic cofactor. Chem Commun. 2012;48(13):1898-1900.
15. Campbell E, Wheeldon IR, Banta S. Broadening the cofactor specificity of a thermostable alcohol dehydrogenase using rational protein design introduces novel kinetic transient behavior. Biotechnol Bioeng. 2010;107(5):763-774.
16. Chen K, Arnold FH. Engineering new catalytic activities in enzymes. Nat Catal. 2020;3(3):203-213.
17. Chen X, Li W, Ji P, Zhao Y, Hua C, Han C. Engineering the conserved and noncatalytic residues of a thermostable β-1,4-endoglucanase to improve specific activity and thermostability. Sci Rep. 2018;8(1):2954.
18. Choi JM, Han SS, Kim HS. Industrial applications of enzyme biocatalysis: current status and future aspects. Biotechnol Adv. 2015;33(7):1443-1454.
19. Chowdhury R, Ren T, Shankla M, Decker K, Grisewood M, Prabhakar J, et al. PoreDesigner for tuning solute selectivity in a robust and highly permeable outer membrane pore. Nat Commun. 2018;9(1):3661.
20. Cowan DA, Fernandez-Lafuente R. Enhancing the functional properties of thermophilic enzymes by chemical modification. Enzyme Microb Technol. 2011;49(4-5):326-333.
21. Das J, Mishra HN. Electrochemical biosensor for monitoring fish spoilage based on nanocellulose as enzyme immobilization matrix. J Food Meas Charact. 2023;17(4):3827-3844.
22. Datta S, Veena R, Samuel MS, Selvarajan E. Immobilization of laccases and applications for the detection and remediation of pollutants: a review. Environ Chem Lett. 2021;19(1):521-538.
23. Denisenko YA, Gusakov AV, Rozhkova AM, Osipov DO, Zorov IN, Matys VY, et al. Site-directed mutagenesis of GH10 xylanase A from Penicillium canescens for determining factors affecting enzyme thermostability. Int J Biol Macromol. 2017;104:665671.
24. Desai AA. Sitagliptin manufacture: a compelling tale of green chemistry, process intensification, and industrial asymmetric catalysis. Angew Chem Int Ed. 2011;50(9):1974-1976.
25. Dotsenko A, Denisenko J, Osipov D, Rozhkova A, Zorov I, Sinitsyn A. Testing and improving the performance of protein thermostability predictors for the engineering of cellulases. J Bioinform Comput Biol. 2023;21(2):2330001.
26. Dror A, Shemesh E, Dayan N, Fishman A. Protein engineering by random mutagenesis and structureguided consensus of GeoBacillus stearothermophilus lipase T6 for enhanced stability in methanol. Appl Environ Microbiol. 2014;80(4):1515-1527.
27. Du C, Tan S, Liu L, Zhou Y, Wu P, Zhang G. Improving the specific activity and stability of alkaline pectinase PEL3 through SpyTag/SpyCatcher cyclization. Biotechnol Lett. 2023;45(7):847-859.
28. Duan F, Sun T, Zhang J, Wang K, Wen Y, Lu L. Recent innovations in immobilization of βgalactosidases for industrial and therapeutic applications. Biotechnol Adv. 2022;61:108053.
29. Duan X, Chen J, Wu J. Improving the thermostability and catalytic efficiency of Bacillus deramificans pullulanase by site-directed mutagenesis. Appl Environ Microbiol. 2013;79(13):4072-4077.
30. Edison LK, Satheeshkumar PK, Pradeep NS. Industrial production and purification of recombinant betaglucanases. In: Microbial beta glucanases: molecular structure, functions and applications. Singapore: Springer Nature; 2022. p. 171-185.
31. Ehren J, Govindarajan S, Morón B, Minshull J, Khosla C. Protein engineering of improved prolyl endopeptidases for celiac sprue therapy. Protein Eng Des Sel. 2008;21(12):699-707.
32. Ehsani M, Fernández MR, Biosca JA, Dequin S. Reversal of coenzyme specificity of 2,3-butanediol dehydrogenase from Saccharomyces cerevisiae and in vivo functional analysis. Biotechnol Bioeng. 2009;104(2):381-389.
33. Feng L, Han Y, Niu J, Guo J, Zhang J, Li H, et al. Synergistic removal performance and mechanism in denitrification system under phenol stress. J Environ Chem Eng. 2023;11(5):110767.
34. Feng Y, Xu Y, Liu S, Wu D, Su Z, Chen G, et al. Recent advances in enzyme immobilization based on novel porous framework materials and its applications in biosensing. Coord Chem Rev. 2022;459:214414.
35. Fryszkowska A, Devine PN. Biocatalysis in drug discovery and development. Curr Opin Chem Biol. 2020;55:151-160.
36. Fu R, Martin C, Zhang Y. Next-generation plant metabolic engineering, inspired by an ancient Chinese irrigation system. Mol Plant. 2018;11(1):47-57.
37. Ganonyan N, Benmelech N, Bar G, Gvishi R, Avnir D. Entrapment of enzymes in silica aerogels. Mater Today. 2020;33:24-35.
38. Gao DY, Sun XB, Liu MQ, Liu YN, Zhang HE, Shi XL, et al. Characterization of thermostable and chimeric enzymes via isopeptide bond-mediated molecular cyclization. J Agric Food Chem. 2019;67(24):6837-6846.
39. Gao L, Zhuang J, Nie L, Zhang J, Zhang Y, Gu N, et al. Intrinsic peroxidase-like activity of ferromagnetic nanoparticles. Nat Nanotechnol. 2007;2(9):577-583.
40. Gill J, Orsat V, Kermasha S. Optimization of encapsulation of a microbial laccase enzymatic extract using selected matrices. Process Biochem. 2018;65:5561.
41. Grisewood MJ, Hernández-Lozada NJ, Thoden JB, Gifford NP, Mendez-Perez D, Schoenberger HA, et al. Computational redesign of acyl-ACP thioesterase with improved selectivity toward medium-chain-length fatty acids. ACS Catal. 2017;7(6):3837-3849.
42. Guisan JM, Fernandez-Lorente G, Rocha-Martin J, Moreno-Gamero D. Enzyme immobilization strategies for the design of robust and efficient biocatalysts. Curr Opin Green Sustain Chem. 2022;35:100593.
43. Hailes HC, Dalby PA, Woodley JM. Integration of biocatalytic conversions into chemical syntheses. J Chem Technol Biotechnol. 2007;82(12):1063-1066.
44. Häring D, Schreier P. Chemical engineering of enzymes: altered catalytic activity, predictable selectivity and exceptional stability of the semisynthetic peroxidase seleno-subtilisin. Naturwissenschaften. 1999;86(7):307-312.
45. He J, Zhang C, Dai H, Liu H, Zhang X, Yang J, et al. A LysM receptor heteromer mediates perception of arbuscular mycorrhizal symbiotic signal in rice. Mol Plant. 2019;12(12):1561-1576.
46. Hegazy UM, El-Khonezy MI, Shokeer A, Abdel-Ghany SS, Bassuny RI, Barakat AZ, et al. revealing a novel xylose-binding site of GeoBacillus stearothermophilus xylanase by directed evolution. J Biochem. 2019;165(2):177-184.
47. Hernández-Lozada NJ, Lai RY, Simmons TR, Thomas KA, Chowdhury R, Maranas CD, et al. Highly active C8-acyl-ACP thioesterase variant isolated by a synthetic selection strategy. ACS Synth Biol. 2018;7(9):2205-2215.
48. Hu X, Hu WX, Lu HY, Liu S, Rao SQ, Yang ZQ, et al. Glycosylated cross-linked ovalbumin by transglutaminase in the presence of oligochitosan: effect of enzyme action time and enhanced functional properties. Food Hydrocoll. 2023;138:108462.
49. Huisman GW, Collier SJ. On the development of new biocatalytic processes for practical pharmaceutical synthesis. Curr Opin Chem Biol. 2013;17(2):284-292.
50. Huisman GW, Liang J, Krebber A. Practical chiral alcohol manufacture using ketoreductases. Curr Opin Chem Biol. 2010;14(2):122-129.
51. Hwang ET, Gu MB. Enzyme stabilization by nano/microsized hybrid materials. Eng Life Sci. 2013;13(1):49-61.
52. Jaafar NR, Jailani N, Rahman RA, Öner ET, Murad AMA, Illias RM. Protein surface engineering and interaction studies of maltogenic amylase towards International Journal of Advanced Biochemistry Research https://www.biochemjournal.com improved enzyme immobilisation. Int J Biol Macromol. 2022;213:70-82.
53. Agadeesan Y, Meenakshisundaram S, Saravanan V, Balaiah A. Sustainable production, biochemical and molecular characterization of thermo-and solventstable alkaline serine keratinase from novel Bacillus pumilus AR57 for poultry solid waste management. Int J Biol Macromol. 2020;163:135-146.
54. Jochens H, Bornscheuer UT. Natural diversity to guide focused directed evolution. ChemBioChem. 2010;11(13):1861-1866.
55. Khan MF, Kundu D, Hazra C, Patra S. A strategic approach of enzyme engineering by attribute ranking and enzyme immobilization on zinc oxide nanoparticles to attain thermostability in mesophilic Bacillus subtilis lipase for detergent formulation. Int J Biol Macromol. 2019;136:66-82.
56. Kharazmi S, Taheri-Kafrani A. Bi-enzymatic nanobiocatalyst based on immobilization of xylanase and pectinase onto functionalized magnetic nanoparticles for efficient fruit juice clarification. LWT. 2023;183:114914.
57. Kroll A, Rousset Y, Hu XP, Liebrand NA, Lercher MJ. Turnover number predictions for kinetically uncharacterized enzymes using machine and deep learning. Nat Commun. 2023;14(1):4139.
58. Kujawa J, Głodek M, Li G, Al-Gharabli S, Knozowska K, Kujawski W. Highly effective enzyme immobilization on ceramics: requirements for supports and enzymes. Sci Total Environ. 2021;801:149647.
59. Kumar A, Jangra A, Pramanik J. Application of enzymes in dairy processing industry: a review. Curr Nutr Food Sci. 2022;18(5):428-431.
60. Kumar N, Upadhyay LSB. Enzyme immobilization over polystyrene surface using cysteine functionalized copper nanoparticle as a linker molecule. Appl Biochem Biotechnol. 2020;191(3):1247-1257.
61. Lapidoth G, Khersonsky O, Lipsh R, Dym O, Albeck S, Rogotner S, et al. Highly active enzymes by automated combinatorial backbone assembly and sequence design. Nat Commun. 2018;9(1):2780.
62. Leresche JE, Meyer HP. Chemocatalysis and biocatalysis (biotransformation): some thoughts of a chemist and of a biotechnologist. Org Process Res Dev. 2006;10(3):572-580.
63. Li G, Xu L, Zhang H, Liu J, Yan J, Yan Y. A de novo designed esterase with p-nitrophenyl acetate hydrolysis activity. Molecules. 2020;25(20):4658.
64. Li T, Liang J, Ambrogelly A, Brennan T, Gloor G, Huisman G, et al. Efficient chemoenzymatic process for manufacture of the boceprevir bicyclic [3.1.0] proline intermediate based on amine oxidase-catalyzed desymmetrization. J Am Chem Soc. 2012;134(14):6467-6472.
65. Li W, Huo Z, Zhang X, Zhao H, Cui Z, Fu P, et al. New sight for in-situ monitoring of silica growth process: the incorporation of Stöber process and aggregationinduced emission technique. Dyes Pigments. 2020;182:108637.
66. Liang L, Zhang J, Lin Z. Altering coenzyme specificity of Pichia stipitis xylose reductase by the semi-rational approach CASTing. Microb Cell Fact. 2007;6(1):36.
67. Liu L, Wang L, Zhang Z, Guo X, Li X, Chen H. Domain-swapping of mesophilic xylanase with hyperthermophilic glucanase. BMC Biotechnol. 2012;12(1):28.
68. Liu L, Zeng L, Wang S, Cheng J, Li X, Song A, et al. Activity and thermostability increase of xylanase following transplantation with modules subdivided from hyper-thermophilic CBM9_1-2. Process Biochem. 2012;47(5):853-857.
69. Liu S, Bilal M, Rizwan K, Gul I, Rasheed T, Iqbal HMN. Smart chemistry of enzyme immobilization using various support matrices: a review. Int J Biol Macromol. 2021;190:396-408.
70. Lynch JH, Huang XQ, Dudareva N. Silent constraints: the hidden challenges faced in plant metabolic engineering. Curr Opin Biotechnol. 2021;69:112-117.
71. Madhavan A, Arun KB, Binod P, Sirohi R, Tarafdar A, Reshmy R, et al. Design of novel enzyme biocatalysts for industrial bioprocess: harnessing the power of protein engineering, high throughput screening and synthetic biology. Bioresour Technol. 2021;325:124617.
72. Maqbool AHMCHT, Saitoh H, Franceschetti M, Stevenson CEM, Uemura A, Kanzaki H, et al. Structural basis of pathogen recognition by an integrated HMA domain in a plant NLR immune receptor. eLife. 2015;4:e08709.
73. Marcucci SMP, Zanin GM, Arroyo PA. Synthesis of SBA-15 and pore-expanded SBA-15 and surface modification with tin for covalent lipase immobilization. Microporous Mesoporous Mater. 2022;337:111951.
74. Matsumura H, Shiomi K, Yamamoto A, Taketani Y, Kobayashi N, Yoshizawa T, et al. Hybrid Rubisco with complete replacement of rice Rubisco small subunits by sorghum counterparts confers C4 plant-like high catalytic activity. Mol Plant. 2020;13(11):1570-1581.
75. McIntosh JA, Owens AE. Enzyme engineering for biosynthetic cascades. Curr Opin Green Sustain Chem. 2021;29:100448.
76. McLure RJ, Radford SE, Brockwell DJ. Highthroughput directed evolution: a golden era for protein science. Trends Chem. 2022;4(5):378-391.
77. Meng X, O'Hare D, Ladame S. Surface immobilization strategies for the development of electrochemical nucleic acid sensors. Biosens Bioelectron. 2023;237:115440.
78. Mohidem NA, Mohamad M, Rashid MU, Norizan MN, Hamzah F, Mat HB. Recent advances in enzyme immobilisation strategies: an overview of techniques and composite carriers. J Compos Sci. 2023;7(12):488.
79. Moosavi F, Ahrari F, Ahmadian G, Mohammadi M. Sortase-mediated immobilization of Candida antarctica lipase B on graphene oxide: comparison with chemical approach. Biotechnol Rep. 2022;34:e00733.
80. Musil M, Konegger H, Hon J, Bednar D, Damborsky J. Computational design of stable and soluble biocatalysts. ACS Catal. 2018;9(2):1033-1054.
81. Nageib AM, Jameel AT, Ahmad FB. Immobilization of alpha-naphthyl acetate esterase on κ-carrageenan for potential biosensor application in pesticide detection. Mater Today Proc. 2022;57:1296-1300.
82. Narayanan KB, Han SS. Peptide ligases: a novel and potential enzyme toolbox for catalytic cross-linking of protein/peptide-based biomaterial scaffolds for tissue International Journal of Advanced Biochemistry Research https://www.biochemjournal.com engineering. Enzyme Microb Technol. 2022;155:109990.
83. Noey EL, Tibrewal N, Jiménez-Osés G, Osuna S, Park J, Bond CM, et al. Origins of stereoselectivity in evolved ketoreductases. Proc Natl Acad Sci U S A. 2015;112(51):E7065-E7072.
84. Otte KB, Hauer B. Enzyme engineering in the context of novel pathways and products. Curr Opin Biotechnol. 2015;35:16-22.
85. Paiva MT, Kishima JOF, Silva JB, Mantovan J, Colodi FG, Mali S. Crosslinking methods in polysaccharidebased hydrogels for drug delivery systems. Biomed Mater Devices. 2024;2(1):288-306.
86. Patel RN. Biocatalysis for synthesis of pharmaceuticals. Bioorg Med Chem. 2018;26(7):1252-1274.
87. Pollard DJ, Woodley JM. Biocatalysis for pharmaceutical intermediates: the future is now. Trends Biotechnol. 2007;25(2):66-73.
88. Prajapati AS, Panchal KJ, Pawar VA, Noronha MJ, Patel DH, Subramanian RB. Review on cellulase and xylanase engineering for biofuel production. Ind Biotechnol. 2018;14(1):38-44.
89. Prasad S, Roy I. Converting enzymes into tools of industrial importance. Recent Pat Biotechnol. 2018;12(1):33-56.
90. Quin MB, Schmidt-Dannert C. Engineering of biocatalysts: from evolution to creation. ACS Catal. 2011;1(9):1017-1021.
91. Radley E, Davidson J, Foster J, Obexer R, Bell EL, Green AP. Engineering enzymes for environmental sustainability. Angew Chem Int Ed. 2023;62(52):e202309305.
92. Ramalho EX, De Castro RJS. Covalent bonding immobilization of a Bacillus licheniformis protease on chitosan and its application in protein hydrolysis. Biocatal Agric Biotechnol. 2023;50:102713.
93. Rashid MU, Daud WMAW, Mohidem NA, Mohamad MB, Akhtar J, Azam M, et al. Methane dry reforming with CO₂ over ceria supported Ni catalyst prepared by reverse microemulsion synthesis. Fuel. 2022;317:123433.
94. Reshmy R, Philip E, Sirohi R, Tarafdar A, Arun KB, Madhavan A, et al. Nanobiocatalysts: advancements and applications in enzyme technology. Bioresour Technol. 2021;337:125491.
95. Rigoldi F, Donini S, Redaelli A, Parisini E, Gautieri A. Engineering of thermostable enzymes for industrial applications. APL Bioeng. 2018;2(1):011501.
96. Rodrigues RC, Berenguer-Murcia Á, Carballares D, Morellon-Sterling R, Fernandez-Lafuente R. Stabilization of enzymes via immobilization: multipoint covalent attachment and other stabilization strategies. Biotechnol Adv. 2021;52:107821.
97. Romero PA, Krause A, Arnold FH. Navigating the protein fitness landscape with Gaussian processes. Proc Natl Acad Sci U S A. 2013;110(3):E193-E201.
98. Romero PA, Stone E, Lamb C, Chantranupong L, Krause A, Miklos AE, et al. SCHEMA-designed variants of human arginase I and II reveal sequence elements important to stability and catalysis. ACS Synth Biol. 2012;1(6):221-228.
99. Saeed M, Yan M, Ni Z, Hussain N, Chen H. Molecular strategies to enhance the keratinase gene expression and its potential implications in poultry feed industry. Poult Sci. 2024;103(5):103606.
100. Sampaio CS, Angelotti JA, Fernandez-Lafuente R, Hirata DB. Lipase immobilization via cross-linked enzyme aggregates: problems and prospects a review. Int J Biol Macromol. 2022;215:434-449.
101. Sanli G, Banta S, Anderson S, Blaber M. Structural alteration of cofactor specificity in Corynebacterium 2,5-diketo-D-gluconic acid reductase. Protein Sci. 2004;13(2):504-512.
102. Savile CK, Janey JM, Mundorff EC, Moore JC, Tam S, Jarvis WR, et al. Biocatalytic asymmetric synthesis of chiral amines from ketones applied to sitagliptin manufacture. Science. 2010;329(5989):305-309.
103. Scrutton NS, Berry A, Perham RN. Redesign of the coenzyme specificity of a dehydrogenase by protein engineering. Nature. 1990;343(6253):38-43.
104. Shaarani SM, Jahim JM, Rahman RA, Idris A, Murad AMA, Illias RM. Silanized maghemite for cross-linked enzyme aggregates of recombinant xylanase from Trichoderma reesei. J Mol Catal B Enzym. 2016;133:65-76.
105. Shahbazmohammadi H, Sardari S, Lari A, Omidinia E. Engineering an efficient mutant of Eupenicillium terrenum fructosyl peptide oxidase for the specific determination of hemoglobin A1c. Appl Microbiol Biotechnol. 2019;103(4):1725-1735.
106. Shanmugaratnam S, Eisenbeis S, Höcker B. A highly stable protein chimera built from fragments of different folds. Protein Eng Des Sel. 2012;25(11):699-703.
107. Shelat NY, Parhi S, Ostermeier M. Development of a cancer-marker activated enzymatic switch from the herpes simplex virus thymidine kinase. Protein Eng Des Sel. 2017;30(2):95-103.
108. Sheldon RA, Pereira PC. Biocatalysis engineering: the big picture. Chem Soc Rev. 2017;46(10):2678-2691.
109. Smith MD, Zanghellini A, Grabs-Röthlisberger D. Computational design of novel enzymes without cofactors. In: Protein design: methods and applications. 2014. p. 197-210.
110. Smith S, Goodge K, Delaney M, Struzyk A, Tansey N, Frey M. A comprehensive review of the covalent immobilization of biomolecules onto electrospun nanofibers. Nanomaterials. 2020;10(11):2142.
111. Srivastava B, Khatri M, Singh G, Arya SK. Microbial keratinases: an overview of biochemical characterization and its eco-friendly approach for industrial applications. J Clean Prod. 2020;252:119847.
112. Tang KHD, Lock SSM, Yap PS, Cheah KW, Chan YH, Yiin CL, et al. Immobilized enzyme/microorganism complexes for degradation of microplastics: a review of recent advances, feasibility and future prospects. Sci Total Environ. 2022;832:154868.
113. Tatineni R, Doddapaneni KK, Potumarthi RC, Vellanki RN, Kandathil MT, Kolli N, et al. Purification and characterization of an alkaline keratinase from Streptomyces sp. Bioresour Technol. 2008;99(6):15961602.
114. Tavernini L, Ottone C, Illanes A, Wilson L. Entrapment of enzyme aggregates in chitosan beads for aroma release in white wines. Int J Biol Macromol. 2020;154:1082-1090.
115. Teng C, Jiang Y, Xu Y, Li Q, Li X, Fan G, et al. Improving the thermostability and catalytic efficiency of GH11 xylanase PjxA by adding disulfide bridges. Int J Biol Macromol. 2019;128:354-362.
116. Throckmorton K, Vinnik V, Chowdhury R, Cook T, Chevrette MG, Maranas C, et al. Directed evolution reveals the functional sequence space of an adenylation domain specificity code. ACS Chem Biol. 2019;14(9):2044-2054.
117. Tuan LQA. Rational protein design for enhancing thermal stability of industrial enzymes. Ho Chi Minh City Open Univ J Sci Eng Technol. 2018;8(1):3-17.
118. Tvorynska S, Barek J, Josypcuk B. Influence of different covalent immobilization protocols on electroanalytical performance of laccase-based biosensors. Bioelectrochemistry. 2022;148:108223. 119. van Beek HL, de Gonzalo G, Fraaije MW. Blending Baeyer-Villiger monooxygenases: using a robust BVMO as a scaffold for creating chimeric enzymes with novel catalytic properties. Chem Commun. 2012;48(27):3288-3290.
119. Vidal LS, Isalan M, Heap JT, Ledesma-Amaro R. A primer to directed evolution: current methodologies and future directions. RSC Chem Biol. 2023;4(4):271-291.
120. Vidya J, Sajitha S, Ushasree MV, Sindhu R, Binod P, Madhavan A, et al. RETRACTED: Genetic and metabolic engineering approaches for the production and delivery of L-asparaginases: an overview. Bioresour Technol. 2017;245:1775-1781.
121. Wang Y, San KY, Bennett GN. Cofactor engineering for advancing chemical biotechnology. Curr Opin Biotechnol. 2013;24(6):994-999.
122. Woodley JM. Integrating protein engineering with process design for biocatalysis. Philos Trans A Math Phys Eng Sci. 2018;376(2110):20170062.
123. Woodley JM. Accelerating the implementation of biocatalysis in industry. Appl Microbiol Biotechnol. 2019;103(12):4733-4739.
124. Wulf H, Mallin H, Bornscheuer UT. Protein engineering of a thermostable polyol dehydrogenase. Enzyme Microb Technol. 2012;51(4):217-224.
125. Xing M, Chen Y, Li B, Tian S. Highly efficient removal of patulin using immobilized enzymes of Pseudomonas aeruginosa TF-06 entrapped in calcium alginate beads. Food Chem. 2022;377:131973.
126. Xu KL, Gong GX, Liu M, Yang L, Xu ZJ, Gao S, et al. Keratinase improves the growth performance, meat quality and redox status of broiler chickens fed a diet containing feather meal. Poult Sci. 2022;101(6):101913.
127. Yi D, Bayer T, Badenhorst CP, Wu S, Doerr M, Höhne M, et al. Recent trends in biocatalysis. Chem Soc Rev. 2021;50(14):8003-8049.
128. Zanghellini A, Jiang L, Wollacott AM, Cheng G, Meiler J, Althoff EA, et al. New algorithms and an insilico benchmark for computational enzyme design. Protein Sci. 2006;15(12):2785-2794.
129. Zeng W, Guo L, Xu S, Chen J, Zhou J. Highthroughput screening technology in industrial biotechnology. Trends Biotechnol. 2020;38(8):888-906.
130. Zeymer C, Hilvert D. Directed evolution of protein catalysts. Annu Rev Biochem. 2018;87:131-157.
131. Zhang D, Chen X, Chi J, Feng J, Wu Q, Zhu D. Semirational engineering a carbonyl reductase for the enantioselective reduction of β-amino ketones. ACS Catal. 2015;5(4):2452-2457.
132. Zhang L, Wang W, Yang Y, Zhu W, Li P, Wang S, et al. Site-specific, covalent immobilization of PNGase F on magnetic particles mediated by microbial transglutaminase. Anal Chim Acta. 2023;1250:340972.
133. Zhang RX, Gong JS, Su C, Zhang DD, Tian H, Dou WF, et al. Biochemical characterization of a novel surfactant-stable serine keratinase with no collagenase activity from BreviBacillus parabrevis CGMCC 10798. Int J Biol Macromol. 2016;93:843-851.
134. Zhang X, Lin Z, Cai Y, Liu X, Niu L, Liu A. Selective and sensitive colorimetric sensing of iodine ions based on porous MoS₂ particles with excellent haloperoxidase-like activity. Sens Actuators B Chem. 2023;392:134127.
135. Zhang Y, Piao M, He L, Yao L, Piao T, Liu Z, et al. Immobilization of laccase on magnetically separable biochar for highly efficient removal of bisphenol A in water. RSC Adv. 2020;10(8):4795-4804.
136. Zhao G, Wang J, Tang Q, Lan D, Wang Y. Improving the catalytic activity and thermostability of MAS1 lipase by alanine substitution. Mol Biotechnol. 2018;60(4):319-328.
137. Zhou Z, Wang X. Improve thermostability of Bacillus sp. TS chitosanase through structure-based alignment. Sci Rep. 2021;11(1):15846.
138. Zorn K, Oroz-Guinea I, Brundiek H, Dörr M, Bornscheuer UT. Alteration of chain length selectivity of Candida antarctica lipase A by semi-rational design for the enrichment of erucic and gondoic fatty acids. Adv Synth Catal. 2018;360(21):4115-4131.