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Review

Epoxide Hydrolases: Multipotential Biocatalysts

by
Marek Bučko
1,*,
Katarína Kaniaková
2,
Helena Hronská
2,
Peter Gemeiner
1 and
Michal Rosenberg
2
1
Department of Glycobiotechnology, Institute of Chemistry, Center for Glycomics, Slovak Academy of Sciences, Dúbravská cesta 9, 845 38 Bratislava, Slovakia
2
Institute of Biotechnology, Faculty of Chemical and Food Technology, Slovak University of Technology, Radlinského 9, 812 37 Bratislava, Slovakia
*
Author to whom correspondence should be addressed.
Int. J. Mol. Sci. 2023, 24(8), 7334; https://doi.org/10.3390/ijms24087334
Submission received: 27 March 2023 / Revised: 11 April 2023 / Accepted: 12 April 2023 / Published: 15 April 2023
(This article belongs to the Special Issue Biocatalysis and Bioactive Molecules: Future and Development)
Browse Figures
Versions Notes

Abstract

:
Epoxide hydrolases are attractive and industrially important biocatalysts. They can catalyze the enantioselective hydrolysis of epoxides to the corresponding diols as chiral building blocks for bioactive compounds and drugs. In this review article, we discuss the state of the art and development potential of epoxide hydrolases as biocatalysts based on the most recent approaches and techniques. The review covers new approaches to discover epoxide hydrolases using genome mining and enzyme metagenomics, as well as improving enzyme activity, enantioselectivity, enantioconvergence, and thermostability by directed evolution and a rational design. Further improvements in operational and storage stabilization, reusability, pH stabilization, and thermal stabilization by immobilization techniques are discussed in this study. New possibilities for expanding the synthetic capabilities of epoxide hydrolases by their involvement in non-natural enzyme cascade reactions are described.

Graphical Abstract

1. Introduction

Epoxide hydrolases (EHs) catalyze the hydrolysis of the oxirane ring by adding a water molecule to form the corresponding vicinal diol without requiring any cofactor [1]. Most EHs were members of the broad superfamily of hydrolases with an α/β-fold with the assigned EC number 3.3.2.3 in the BRENDA database. However, the classification was deleted from this database, and EHs were divided into two new enzyme groups: EC 3.3.2.9, microsomal EHs, and EC 3.3.2.10, soluble EHs. There are also EHs (e.g., limonene-1,2-hydrolase or leukotriene-A4 hydrolase) with completely different structures and catalytic mechanisms that are classified separately [2].
Epoxide hydrolases are found in various organisms, from prokaryotes to eukaryotes. They perform different functions depending on their site of localization and the origin of the organisms. Among eukaryotes, mammalian EHs are the most studied, mainly because of their role in xenobiotic metabolism and signaling processes [3,4]. Prokaryotic EHs are necessary for catabolic pathways in which specific aromatic compounds or alkenes are used as carbon sources [5].
Due to their broad substrate specificity and high stereospecificity, epoxide hydrolases have received attention as industrial biocatalysts. The EH-catalyzed enantioselective hydrolysis of epoxides was first discovered in mammalian cells, but the application of EHs from this source on an industrial scale was limited due to their low availability. Large quantities of EHs could be prepared after the discovery of microbial EH producers and their recombinant expression in host organisms. This opened the way to their commercial use in the industrial production of chiral compounds, especially in the synthesis of enantiopure drugs, chiral epoxides, and diols [6]. The enantioselectivity and stability of microbial EHs can also be improved by using organic solvents, detergents, ionic liquids, immobilization, and by innovative methods, such as enzyme engineering or direct evolution methods [7].
Other reviews published on EHs have focused on their sources, substrate scope, enantioselectivity, and application in organic chemistry [1,8,9,10]. In this review, we summarized and updated the most interesting applications of EHs associated with the formation of enantiopure epoxides or diols from chiral precursors, which are key intermediates for synthesizing various target products. We also presented the latest approaches for modifying these enzymes by enzyme engineering techniques used to produce tailored EHs. We provided examples of successful applications of immobilized EHs and the benefits of immobilization methods. Finally, we highlighted the recent successful involvement of EHs in enzymatic cascade reactions, confirming the multipotential use of EHs.

2. Epoxides and Diols as Chiral Precursors and Their Applications

Chirality is an important property of bioactive molecules, especially in pharmacology, where different stereoisomers can have different pharmacological properties. Many epoxides and diols are intermediates in the synthesis of many drugs, although epoxides are often produced as racemic mixtures. Approximately 57% of commercially available drugs and approximately 99% of purified natural products are chiral compounds [11]. Chiral chemicals are in high demand commercially, and the global market for chiral drugs is expected to grow in the future [12].
Chiral epoxides can serve as useful intermediates for synthesizing many chemicals with industrial applications. Optically pure epoxides can be prepared via biocatalysis using two major approaches. The first approach includes the direct epoxidation of alkenes and vicinal halohydrins using monooxygenases, chloroperoxidases, and haloalcohol dehalogenases. The second method involves the hydrolysis of racemic epoxides using EHs by kinetic resolution or enantioselective hydrolysis. Among the enzymes mentioned above, the use of enantioselective EHs for producing chiral epoxides has several advantages over other enzymes. The main advantages are that EHs do not require cofactors or additional nucleophiles for their function, they are ubiquitous in nature, and they can be easily cloned and produced in large quantities [13]. The first industrial application of EHs was described in 1969 for the production of L-tartaric acid and meso-tartaric acid using whole bacterial cells [6]. The microbial production of L-(+)-tartaric acid was successfully commercialized in the late 1990s, and microbial methods are now considered to be more economical for producing both optical isomers of this organic acid from cis-epoxysuccinic acid [14]. In the 1990s, new EHs capable of enantioselective and enantioconvergent hydrolysis of structurally diverse epoxides were also discovered, which attracted the interest of organic chemists [15,16,17].
Trans-vicinal diols, products of EH-catalyzed reactions, also have many interesting synthetic applications. Some of the chiral precursors that can be prepared using EHs and the products that can be synthesized from them are presented in Table 1.

3. Natural and Recombinant EHs

Epoxide hydrolases (EHs) are ubiquitous in nature, but concerning their application in the industry, microbial enzymes are better suited for mass production. Therefore, novel EHs are mainly searched for among microorganisms. Since the discovery of the first enantioselective microbial EH, many new EH-producing organisms have been identified [6]. In the 1990s and 2000s, many EHs were discovered by the enrichment screening of isolates [27,46,47] or screening strains from various collections [22,48,49,50]. Through extensive screening, EH activity in bacteria was found to be associated with the genera Rhodococcus, Nocardia, Mycobacterium, and Arthrobacter [51]. Although the screening of microbial isolates is very laborious, it is still widely performed to discover new EHs [52,53,54].
The classic screening method has many limitations. One of them is the screening for enzyme activity, where the reaction substrates and products are identified by GC or HPLC after the samples are extracted from culture or reaction media [48]. To overcome this problem, different spectrophotometric methods for rapid activity assays were developed and used to easily determine the substrate or product. Some of these assays are the 4-(p-nitrobenzyl) pyridine assay (blue assay) [55], adrenaline assay (red assay) [56], and sodium metaperiodate assay [57].
Besides traditional culture-based methods used for screening microorganisms for enzyme activity, two new approaches for discovering novel enzymes emerged: genome mining and metagenomics.
One of the new methods is genome mining. Advancements in genome sequencing, bioinformatics, and the large number of genome sequences deposited in public databases enabled the discovery of uncharacterized biocatalysts [58]. About one-fifth of the total microbial genome in the databases is predicted to contain one or more putative EHs. Van Loo et al. [59] also showed that genome databases could be used as a source of novel EHs [59]. Stojanovski et al. [60] identified 29 putative EHs from the genomic data of six soil bacteria using genome mining. Eight of them were recombinantly expressed in E. coli and used for activity studies, where five were identified as α/β-fold EHs, and three showed sequence similarity to the rare class of limonene epoxide hydrolases (LEHs) [60].
Another new approach used for searching for EHs is metagenomics, i.e., the direct extraction and cloning of DNA from natural environments without culturing isolated microorganisms [61]. This method is not only used to identify novel putative EHs [62] but also to recombinantly express these genes, characterize enzymes and use them for enantioselective and regioselective hydrolysis [63,64]. The metagenomic approach can also be useful for discovering enzymes with extremophilic properties. Two novel LEHs and two α/β-fold EHs from environmental DNA were obtained from hot spring environments. Although all DNA samples were collected at around neutral pH and at lower temperatures (from 46 to 65 °C) compared to other hot spring samples, all four novel EHs had higher thermal stability than any other EHs and LEHs isolated from natural environmental sources [65,66].
Advancements in genetic engineering allowed the recombinant expression of EHs in various hosts to produce larger quantities of EHs and for easier purification. Different types of expression vectors, including mammalian and insect cell lines, have been used to express EHs. However, EHs are generally expressed in microbial cells, most often in Escherichia coli [10]. Along with the discovery of novel EHs and advances in recombinant techniques, various techniques have been implemented to enhance their properties through enzyme engineering, immobilization, and optimization of the reaction medium.

4. Improvement of EHs by Enzyme Engineering

One of the main constraints in the application of enzymes in the industry is their insufficient enantioselectivity, narrow substrate specificity, low activity, and thermal stability [67]. This problem could be overcome by using enzyme engineering techniques. Enzyme engineering refers to the process of modifying the amino acid sequences of enzymes to change their properties, such as catalytic activity, thermostability, organic solvent tolerance, and substrate and reaction specificity [68].
The relatively good knowledge of amino acid sequences, reaction mechanisms, and structures of various EHs [69,70,71,72] allowed the application of various methods of enzyme engineering of EHs. After the initial application of directed evolution to prepare stereoselective lipase [73], Cedrone et al. [74] were the first to achieve EH engineering, where they performed error-prone PCR to prepare mutants of A. niger EH (AnEH) with a 3.3-fold increase in the catalytic efficiency toward 4-(p-nitrophenoxy)-1,2-epoxybutane [74]. Since then, many researchers have used various methods of enzyme engineering to obtain EH mutants with enhanced properties. Some of these methods include error-prone PCR [75] saturation mutagenesis [76], DNA shuffling [77], iterative saturation mutagenesis (ISM) [78,79,80], computational design [81,82], and machine learning [83].
Enhancing enzyme activity and broadening their substrate spectrum are the main objectives of enzyme engineering. The modification of selected amino acid (AA) residues in the substrate-binding pocket of EH from Bacillus megaterium ECU 1001 enhanced activity toward bulky racemic epoxides by 6 to 430 times. It helped perform the bioresolution of various racemic epoxides to prepare (S)-epoxides, which are the precursors to various β-blockers, on a preparative scale [84].
The enantioselectivity of EHs has attracted the interest of most researchers in these biocatalysts, but enzymes often lack sufficient enantioselectivity or the ability to perform enantioconvergent hydrolysis. Hence, many studies focused on increasing the enantioselectivity or improving the enantioconvergence of EHs using enzyme engineering methods (Table 2).
Along with studies on the improvement of enzyme activity, enantioselectivity, and enantioconvergence, studies on enzyme engineering have been conducted to produce EHs with higher thermostability [83,108,109]. Gumulya et al. [109] used several rounds of ISM to generate mutants with higher thermal robustness. Mutation sites in AnEH were selected based on the B-FIT approach, where the criterion for selecting AA residues was the highest B factors available from the X-ray crystallography data. The best variant showed a 21 °C increase in T6050, the temperature at which 50% of enzyme activity is lost following heat treatment for 60 min, which represents an 80-fold improvement in enzyme half-life at 60 °C [109].
A computational design was also used for generating thermostable mutants of EH. In this case, the Framework for Rapid Enzyme Stabilization by Computational Libraries (FRESCO) was applied. It is a promising computationally guided approach for protein thermostabilization that uses the Rosettaddg, FoldX, and Disulfide Discovery software packages. The apparent melting temperature of the two best multisite mutants of LEH from R. erythropolis with 10–12 point mutations increased from 50 °C to ~85 °C [108]. Eight AA residues, which were expected to be sensitive to changes in thermostability based on the previous study [108], and eight AA residues that were expected to be sensitive to changes in enantioselectivity selected in the TCSM ISM study [80] were selected for ISM to prepare LEH mutants with enhanced thermostability, activity, and opposite enantioselectivity against cyclohexene oxide [94].
From the perspective of industrial application, the enhancement of thermostability of cis-epoxysuccinate hydrolase (CESH) is also interesting. Using a semi-rational design, combining directed evolution, simulated mutagenesis, and saturation mutagenesis, the half-life of the best CESH mutant at 50 °C increased from 8.5 min to 293.2 min, and T1550 increased from 44 °C to 64.8 °C. Additionally, the effective working range of the pH of the mutant extended to 5.0–10.0 from 8.0–9.0 for the wild-type enzyme [110].
Enzyme engineering can also be applied to completely change the catalytic activity of the enzyme. Jochens et al. [111] converted the enzymatic activity within the α/β-fold hydrolase family. They reported the conversion of esterase to EH by site-directed mutagenesis, although the EH activity of modified esterase was 800-fold lower than that of the template EH from Agrobacterium radiobacter AD1 [111].

5. Immobilization of EHs

According to IUPAC, immobilization in biotechnology is the technique used for the physical or chemical fixation of cells, organelles, enzymes, or other proteins on a solid support, in a solid matrix, or retained by a membrane to increase their stability and facilitate their repeated or continued use [112]. Immobilization techniques belong to the main pillars of optimization procedures for biotransformations. The application of these techniques for EHs includes the most common immobilization principles, such as (1) adsorption, (2) covalent bonding, (3) crosslinking, (4) entrapment, and (5) encapsulation. Immobilization is also considered to be an important step in the commercialization of EHs as it provides reusability and commercial value to the process [7]. Even the first commercial preparation with EH activity was an immobilized enzyme preparation derived from Rhodococcus sp. called SP 409, developed by NOVO Industry. The product was originally designed as a biocatalyst for the hydrolysis of nitriles. The EH activity was later discovered by Hechtberger et al. [113], and it was used for the asymmetric hydrolysis of various racemic epoxides [113]. The commercial availability of EHs is limited, although EHs from Rhodococcus rhodochrous and Aspergillus niger are commercially available as lyophilized powder [114].
The analysis of the processes to immobilize EHs [7] showed that while adsorption and entrapment techniques were used for immobilizing whole cells with EHs (see also [115,116,117]), covalent bonding and cross-linking were used for EHs as isolated or partially purified enzymes. An exception is the encapsulation technique, which is universally suitable for stabilizing Nocardia tartaricans cells [116,118] using CESH and EH isolated from the Sphinogomonas strain [119].
As shown in Table 3, most of the immobilized epoxide hydrolases are used as purified or partially purified enzymes, immobilized on different supports by the covalent binding method. Due to the stability of EHs and their cofactor independence, the application of purified enzymes is advantageous. One exception is the low stability of CESH, which is therefore used as a whole-cell biocatalyst for enantioselective production of L-(+)-tartrate. To overcome the problems of low cell permeability, various surface displaying systems for the mentioned CESH have been developed [120]. In some cases, whole-cell immobilization is occasionally performed, where cells are immobilized by entrapment and encapsulation within porous or semipermeable microparticles with high water content.
The list of developed immobilization techniques presented in Table 3 indicates that immobilization for EHs is desirable, and its application is increasing. Several innovative methods and new materials for immobilizing EHs have been developed in the past five years. For example, CESH was immobilized by metal-ion affinity interaction with Ni-IDA agarose particles to improve enzyme thermostability and pH stability [134]. Additionally, commercial immobilization matrices were used for the covalent binding of epoxide hydrolase from Vigna radiata [19]. The thermostability and operational stability of the EH immobilized later during the production of β-blocker Nifenalol was improved. Some researchers have also synthesized amino-modified mesocellular silica activated by glutaraldehyde [131]. The later immobilization method enabled the improvement of operational stability and thermostability of EH from red mung beans and cutinase from Fusarium sp. ICT SAC1 during enantioselective and regioselective model biotransformations. The development of organic–inorganic hybrid epoxide hydrolase nanoflowers represents a promising novel immobilization concept [144]. The EH nanoflowers impart unique properties, such as a high EH concentration, an optimum conformation for EH, low mass transfer limitations, and a high surface area, which can increase enzyme activity and stability.
The advantages of immobilizing EHs, presented in Table 3, explain why immobilization techniques are still in high demand. Additionally, immobilization techniques might also significantly enhance the biocatalytic efficiency of EHs as a part of enzyme cascades, mentioned in the next chapter. Though the results achieved by enzyme cascades with EHs are limited, the immobilization of recombinant E. coli cells with an overproduced cascade of the enzyme halohydrin dehalogenase and epoxide hydrolase using an adsorption technique yielded promising results [139].
There are also variations in immobilization protocols and the level of their characterization. Directly comparing the biocatalytic efficiency and other properties of different immobilized preparations of EHs is challenging. The latter might be the reason why the choice of immobilization technique for EHs is difficult. There are no general recommendations for the use of immobilization techniques. Selecting a proper immobilization system requires the individual consideration of several parameters, including the type of applied mechanical forces in bioreactors (Figure 1A). The general properties of immobilization systems, which are frequently evaluated and considered to be important for the successful utilization of immobilized biocatalysts, are schematically represented in Figure 1B.

6. Whole-Cell Cascade Biotransformations Using Microbial Epoxide Hydrolases

Intensive research on the use of epoxide hydrolases (EHs) from microbial sources started in 1991 [149]. The advantage of EHs is that they do not require cofactors for the enantioselective hydrolysis of epoxides to the desired vicinal diols and enantiomerically pure epoxides. Hence, they can be used as fresh whole native cells, lyophilizates, and recombinant cells with overproduced EHs. Thus, the regeneration of cofactors and isolation of enzymes are not required; these processes are expensive and can reduce their stability [149]. The main benefit of using EHs is their ability to catalyze enantioconvergent reactions, which allow the economically efficient production of the desired substances [7]. Besides the hydrolysis of epoxides to vicinal diols, racemic mixtures of epoxides can be resolved into pure enantiomers. The synthetic possibilities of EHs were extended by the discovery of the acceptance of non-natural nucleophiles instead of water molecules in epoxide hydrolysis catalyzed by EHs, leading to the aminolysis and azidolysis of epoxides to form the corresponding amino- and azido- derivatives [150]. The need for enantioconvergent approaches for producing pure stereoisomers of vicinal diols from racemates of epoxides led to the application of the advantages of modern enzyme catalysis techniques, which resulted in the construction of artificial enzyme cascades [151]. The process involved EHs in a non-natural enzyme cascade reaction with at least one other enzyme, which facilitated the one-pot production of diols, amino alcohols, and other specialty chemicals. The importance of the two-step enzyme cascade for the biocatalytic production of chiral vicinal diol [152] as an intermediate for the chemoenzymatic synthesis of pharmaceutical (R)-fluoxetine for treating psychiatric and metabolic disorders is shown in Figure 2 [153].
By involving EHs in whole-cell cascade systems, the general advantages of enzyme cascades over single-step biotransformations can be used, which are as follows: (1) reaction intermediates do not have to be isolated, which makes the process cheaper and helps in performing reactions with unstable intermediates; (2) higher product yield; (3) saving resources; (4) reduction of waste production; (5) avoiding the use of toxic compounds, which are consumed immediately in situ; (6) solutions for possible enzyme inhibition issues. Additionally, the use of EH-triggered enzyme cascades can expand the catalytic capabilities of EHs, which are indicated by spontaneous cyclizations associated with the formation of new C-O bonds catalyzed by EHs [154]. In recent years, the application of EHs as biocatalysts for epoxide hydrolysis in cascade reactions has shown several possibilities and advantages, for example, reversing the enantioselectivity of the reaction by properly designing the enzyme cascade. The applications of EHs are summarized in Table 4.
As shown in Table 4, microbial EHs represent a powerful biocatalytic tool, the synthetic possibilities of which can be further expanded by involvement in cascades. The usefulness of EHs can be increased, for example, by incorporating concrete epoxide hydrolase from Sphingomonas sp. HXN-200 (SpEH) into five differently designed enzyme cascades shown in Figure 3 and listed in Table 4. In this review article [163], other recent studies on new cascades using a tandem of EHs with styrene monooxygenases (SMO) connected with other enzymes to form cascades were presented. This confirmed the multifunctionality of EHs for the production of important building blocks and other chemical specialties using enzyme cascades. An important step in further applying the potential of EHs involves the development of optimization procedures leading to industrial processes.

7. Conclusions

The research and development of epoxide hydrolases have advanced significantly, especially their application in biocatalysis. The trend involves the expansion of the biocatalytic repertoire of epoxide hydrolases by involving them in enzyme cascades. The development of new immobilization techniques led to the improvement of the functional properties of epoxide hydrolases, especially concerning their stabilization. Classical screening techniques for new epoxide hydrolases have been replaced by more efficient approaches, especially enzyme metagenomics. Similarly, the techniques for controlling the enantioselectivity and thermostability of epoxide hydrolases are characterized by a higher degree of specificity. The development of these techniques involves the transition from directed evolution to a semi-rational design and a rational design. The progress in the field of epoxide hydrolase development is a prerequisite for its more intensive use in the industrial production of chiral building blocks, which in turn can be used for synthesizing important drugs.

Author Contributions

Writing—original draft preparation, M.B., K.K. and H.H.; writing—review and editing, M.B., K.K., H.H., P.G. and M.R. All authors have read and agreed to the published version of the manuscript.

Funding

This work was supported by the Slovak Research and Development Agency under the Contract no. APVV-20-0272 and by the Slovak Grant Agency for Science VEGA 2/0130/20. This publication is the result of the project implementation CEMBAM—Centre for Medical Bio-Additive Manufacturing and Research, ITMS2014+: 313011V358 supported by the Operational Programme Integrated Infrastructure funded by the European Regional Development Fund.

Institutional Review Board Statement

Not applicable.

Informed Consent Statement

Not applicable.

Data Availability Statement

Data sharing not applicable.

Acknowledgments

Graphical abstract for this article was created with BioRender.com.

Conflicts of Interest

The authors declare no conflict of interest.

References

  1. Choi, W.J.; Choi, C.Y. Production of chiral epoxides: Epoxide hydrolase-catalyzed enantioselective hydrolysis. Biotechnol. Bioprocess. Eng. 2005, 10, 167. [Google Scholar] [CrossRef]
  2. BRENDA. Available online: www.brenda-enzymes.org (accessed on 20 March 2023).
  3. Morisseau, C.; Hammock, B.D. Gerry Brooks and epoxide hydrolases: Four decades to a pharmaceutical. Pest Manag. Sci. 2008, 64, 594–609. [Google Scholar] [CrossRef]
  4. Decker, M.; Arand, M.; Cronin, A. Mammalian epoxide hydrolases in xenobiotic metabolism and signalling. Arch. Toxicol. 2009, 83, 297–318. [Google Scholar] [CrossRef] [Green Version]
  5. Orru, R.V.A.; Archelas, A.; Furstoss, R.; Faber, K. Epoxide Hydrolases and Their Synthetic Applications. In Biotransformations; Faber, K., Ed.; Springer: Berlin/Heidelberg, Germany, 1999; pp. 145–167. [Google Scholar]
  6. Archelas, A.; Furstoss, R. Epoxide hydrolases: New tools for the synthesis of fine organic chemicals. Trends Biotechnol. 1998, 16, 108–116. [Google Scholar] [CrossRef]
  7. Saini, P.; Sareen, D. An Overview on the Enhancement of Enantioselectivity and Stability of Microbial Epoxide Hydrolases. Mol. Biotechnol. 2017, 59, 98–116. [Google Scholar] [CrossRef]
  8. Choi, W.J. Biotechnological production of enantiopure epoxides by enzymatic kinetic resolution. Appl. Microbiol. Biotechnol. 2009, 84, 239–247. [Google Scholar] [CrossRef] [PubMed]
  9. Bala, N.; Chimni, S.S. Recent developments in the asymmetric hydrolytic ring opening of epoxides catalysed by microbial epoxide hydrolase. Tetrahedron Asymmetry 2010, 21, 2879–2898. [Google Scholar] [CrossRef]
  10. Archelas, A.; Iacazio, G.; Kotik, M. Epoxide Hydrolases and their Application in Organic Synthesis. In Green Biocatalysis; Patel, R.N., Ed.; John Wiley & Sons, Inc.: Hoboken, NJ, USA, 2016; pp. 179–229. [Google Scholar]
  11. Jiang, W.; Fang, B. Synthesizing Chiral Drug Intermediates by Biocatalysis. Appl. Biochem. Biotechnol. 2020, 192, 146–179. [Google Scholar] [CrossRef] [PubMed]
  12. Research, V.M. Global Chiral Chemicals Market Size by Technology, by Application, by Geographic Scope and Forecast. Available online: https://www.verifiedmarketresearch.com/product/chiral-chemicals-market/ (accessed on 11 April 2023).
  13. Saini, P.; Sareen, D. Epoxide Hydrolase for the Synthesis of Chiral Drugs. In Nanoscience and Biotechnology for Environmental Applications, 1st ed.; Gothandam, K.M., Ranjan, S., Dasgupta, N., Lichtfouse, E., Eds.; Environmental Chemistry for a Sustainable World; Springer: Cham, Switzerland, 2019; Volume 22, pp. 141–198. [Google Scholar]
  14. Xuan, J.; Feng, Y. Enantiomeric Tartaric Acid Production Using cis-Epoxysuccinate Hydrolase: History and Perspectives. Molecules 2019, 24, 903. [Google Scholar] [CrossRef] [Green Version]
  15. Chen, X.J.; Archelas, A.; Furstoss, R. Microbiological transformations. 27. The first examples for preparative-scale enantioselective or diastereoselective epoxide hydrolyses using microorganisms. An unequivocal access to all four bisabolol stereoisomers. J. Org. Chem. 1993, 58, 5528–5532. [Google Scholar] [CrossRef]
  16. Jacobs, M.H.J.; van den Wijngaard, A.J.; Pentenga, M.; Janssen, D.B. Characterization of the epoxide hydrolase from an epichlorohydrin-degrading Pseudomonas sp. Eur. J. Biochem. 1991, 202, 1217–1222. [Google Scholar] [CrossRef] [PubMed]
  17. Weijers, C.A.G.M. Enantioselective hydrolysis of aryl, alicyclic and aliphatic epoxides by Rhodotorula glutinis. Tetrahedron Asymmetry 1997, 8, 639–647. [Google Scholar] [CrossRef]
  18. Chen, L.; Shen, H.; Wei, C.; Zhu, Q. Bioresolution of (R)-glycidyl azide by Aspergillus niger ZJUTZQ208: A new and concise synthon for chiral vicinal amino alcohols. Appl. Microbiol. Biotechnol. 2013, 97, 2609–2616. [Google Scholar] [CrossRef] [PubMed]
  19. Li, F.-L.; Zheng, Y.-C.; Li, H.; Chen, F.-F.; Yu, H.-L.; Xu, J.-H. Preparing β-blocker (R)-Nifenalol based on enantioconvergent synthesis of (R)-p-nitrophenylglycols in continuous packed bed reactor with epoxide hydrolase. Tetrahedron 2019, 75, 1706–1710. [Google Scholar] [CrossRef]
  20. Pedragosa-Moreau, S.; Morisseau, C.; Baratti, J.; Zylber, J.; Archelas, A.; Furstoss, R. Microbiological transformations 37. An enantioconvergent synthesis of the β-blocker (R)-Nifénalol® using a combined chemoenzymatic approach. Tetrahedron 1997, 53, 9707–9714. [Google Scholar] [CrossRef]
  21. Manoj, K.M.; Archelas, A.; Baratti, J.; Furstoss, R. Microbiological transformations. Part 45: A green chemistry preparative scale synthesis of enantiopure building blocks of Eliprodil: Elaboration of a high substrate concentration epoxide hydrolase-catalyzed hydrolytic kinetic resolution process. Tetrahedron 2001, 57, 695–701. [Google Scholar] [CrossRef]
  22. Zhang, J.; Reddy, J.; Roberge, C.; Senanayake, C.; Greasham, R.; Chartrain, M. Chiral bio-resolution of racemic indene oxide by fungal epoxide hydrolases. J. Ferment. Bioeng. 1995, 80, 244–246. [Google Scholar] [CrossRef]
  23. Cleij, M.; Archelas, A.; Furstoss, R. Microbiological Transformations 43. Epoxide Hydrolases as Tools for the Synthesis of Enantiopure α-Methylstyrene Oxides:  A New and Efficient Synthesis of (S)-Ibuprofen. J. Org. Chem. 1999, 64, 5029–5035. [Google Scholar] [CrossRef]
  24. Li, C.; Liu, Q.; Song, X.; Ding, D.; Ji, A.; Qu, Y. Epoxide hydrolase-catalyzed resolution of ethyl 3-phenylglycidate using whole cells of Pseudomonas sp. Biotechnol. Lett. 2003, 25, 2113–2116. [Google Scholar] [CrossRef] [PubMed]
  25. Ueberbacher, B.J.; Osprian, I.; Mayer, S.F.; Faber, K. A Chemoenzymatic, Enantioconvergent, Asymmetric Total Synthesis of(R)-Fridamycin E. Eur. J. Org. Chem. 2005, 2005, 1266–1270. [Google Scholar] [CrossRef]
  26. Kong, X.-D.; Yuan, S.; Li, L.; Chen, S.; Xu, J.-H.; Zhou, J. Engineering of an epoxide hydrolase for efficient bioresolution of bulky pharmaco substrates. Proc. Natl. Acad. Sci. USA 2014, 111, 15717–15722. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  27. Choi, W.J.; Puah, S.M.; Tan, L.L.; Ng, S.S. Production of (R)-ethyl-3,4-epoxybutyrate by newly isolated Acinetobacter baumannii containing epoxide hydrolase. Appl. Microbiol. Biotechnol. 2008, 79, 61–67. [Google Scholar] [CrossRef] [PubMed]
  28. Jia, X.; Wang, Z.; Li, Z. Preparation of (S)-2-, 3-, and 4-chlorostyrene oxides with the epoxide hydrolase from Sphingomonas sp. HXN-200. Tetrahedron Asymmetry 2008, 19, 407–415. [Google Scholar] [CrossRef]
  29. Monterde, M.I.; Lombard, M.; Archelas, A.; Cronin, A.; Arand, M.; Furstoss, R. Enzymatic transformations. Part 58: Enantioconvergent biohydrolysis of styrene oxide derivatives catalysed by the Solanum tuberosum epoxide hydrolase. Tetrahedron Asymmetry 2004, 15, 2801–2805. [Google Scholar] [CrossRef]
  30. Monfort, N.; Archelas, A.; Furstoss, R. Enzymatic transformations. Part 53: Epoxide hydrolase-catalysed resolution of key synthons for azole antifungal agents. Tetrahedron Asymmetry 2002, 13, 2399–2401. [Google Scholar] [CrossRef]
  31. Monfort, N.; Archelas, A.; Furstoss, R. Enzymatic transformations. Part 55: Highly productive epoxide hydrolase catalysed resolution of an azole antifungal key synthon. Tetrahedron 2004, 60, 601–605. [Google Scholar] [CrossRef]
  32. Fujino, A.; Asano, M.; Yamaguchi, H.; Shirasaka, N.; Sakoda, A.; Ikunaka, M.; Obata, R.; Nishiyama, S.; Sugai, T. Bacillus subtilis epoxide hydrolase-catalyzed preparation of enantiopure 2-methylpropane-1,2,3-triol monobenzyl ether and its application to expeditious synthesis of (R)-bicalutamide. Tetrahedron Lett. 2007, 48, 979–983. [Google Scholar] [CrossRef]
  33. Roiban, G.-D.; Sutton, P.W.; Splain, R.; Morgan, C.; Fosberry, A.; Honicker, K.; Homes, P.; Boudet, C.; Dann, A.; Guo, J.; et al. Development of an Enzymatic Process for the Production of (R)-2-Butyl-2-ethyloxirane. Org. Process Res. Dev. 2017, 21, 1302–1310. [Google Scholar] [CrossRef]
  34. Bottalla, A.-L.; Ibrahim-Ouali, M.; Santelli, M.; Furstoss, R.; Archelas, A. Epoxide Hydrolase-Catalysed Kinetic Resolution of a Spiroepoxide, a Key Building Block of Various 11-Heterosteroids. Adv. Synth. Catal. 2007, 349, 1102–1110. [Google Scholar] [CrossRef]
  35. Goswami, A.; Totleben, M.J.; Singh, A.K.; Patel, R.N. Stereospecific enzymatic hydrolysis of racemic epoxide: A process for making chiral epoxide. Tetrahedron Asymmetry 1999, 10, 3167–3175. [Google Scholar] [CrossRef]
  36. Patel, R.N. Chapter 11—Applications of Biocatalysis for Pharmaceuticals and Chemicals. In Organic Synthesis Using Biocatalysis; Goswami, A., Stewart, J.D., Eds.; Academic Press: Cambridge, MA, USA, 2016; pp. 339–411. [Google Scholar]
  37. Pienaar, D.P.; Mitra, R.K.; Deventer, T.I.v.; Botes, A.L. Synthesis of a variety of optically active hydroxylated heterocyclic compounds using epoxide hydrolase technology. Tetrahedron Lett. 2008, 49, 6752–6755. [Google Scholar] [CrossRef]
  38. Halama, A.; Kruliš, R.; Rymeš, J. A Convenient Synthesis of Rivaroxaban from (S)-Epichlorohydrin. Org. Prep. Proced. Int. 2020, 52, 201–211. [Google Scholar] [CrossRef]
  39. Halama, A.; Krulis, R.; Dammer, O.; Kalasek, S. A Process for the Preparation of Rivaroxaban Based on the Use of (S)-Epichlorohydrin. W.O. Patent No. 2013/120465, 22 August 2013. [Google Scholar]
  40. Kasai, N.; Suzuki, T.; Furukawa, Y. Chiral C3 epoxides and halohydrins: Their preparation and synthetic application. J. Mol. Catal. B Enzym. 1998, 4, 237–252. [Google Scholar] [CrossRef]
  41. Gu, S.; Wang, X.; Li, Q. A Preparation Method of high-Purity L-Carnitine. W.O. Patent No. 2010/043110, 22 April 2010. [Google Scholar]
  42. Chang, J.-Y.; Lee, H.S.; Kim, D.J.; Ko, M.-S.; Kim, N.D.; Chang, Y.K.; Kim, M.S. Method for Preparing Sitagliptin and Amine Salt Intermediates Used Therein. W.O. Patent No. 2011/102640, 25 August 2011. [Google Scholar]
  43. Kitaori, K.; Takehira, Y.; Furukawa, Y.; Yoshimoto, H.; Otera, J. A Pratical Synthesis of Optically Active Atenolol from Chiral Epichlorohydrin. Chem. Pharm. Bull. 1997, 45, 412–414. [Google Scholar] [CrossRef] [Green Version]
  44. Narina, S.V.; Sudalai, A. Enantioselective synthesis of (S)-timolol via kinetic resolution of terminal epoxides and dihydroxylation of allylamines. Tetrahedron 2007, 63, 3026–3030. [Google Scholar] [CrossRef]
  45. Ling, X.; Lu, D.; Wang, J.; Chen, J.; Ding, L.; Chen, J.; Chai, H.; Ouyang, P. Kinetic investigation on enantioselective hydrolytic resolution of epichlorohydrin by crude epoxide hydrolase from domestic duck liver. Afr. J. Biotechnol. 2011, 10, 3436–3443. [Google Scholar] [CrossRef] [Green Version]
  46. Zhang, Z.; Sheng, Y.; Jiang, K.; Wang, Z.; Zheng, Y.; Zhu, Q. Bio-resolution of glycidyl (o, m, p)-methylphenyl ethers by Bacillus megaterium. Biotechnol. Lett. 2010, 32, 513–516. [Google Scholar] [CrossRef]
  47. Kotik, M.; Brichac, J.; Kyslík, P. Novel microbial epoxide hydrolases for biohydrolysis of glycidyl derivatives. J. Biotechnol. 2005, 120, 364–375. [Google Scholar] [CrossRef]
  48. Zocher, F.; Enzelberger, M.M.; Bornscheuer, U.T.; Hauer, B.; Wohlleben, W.; Schmid, R.D. Epoxide hydrolase activity of Streptomyces strains. J. Biotechnol. 2000, 77, 287–292. [Google Scholar] [CrossRef] [PubMed]
  49. Moussou, P.; Archelas, A.; Furstoss, R. Microbiological transformations 41.: Screening for novel fungal epoxide hydrolases. J. Mol. Catal. B Enzym. 1998, 5, 447–458. [Google Scholar] [CrossRef]
  50. Yeates, C.A.; van Dyk, M.S.; Botes, A.L.; Breytenbach, J.C.; Krieg, H.M. Biocatalysis of nitro substituted styrene oxides by non-conventional yeasts. Biotechnol. Lett. 2003, 25, 675–680. [Google Scholar] [CrossRef] [PubMed]
  51. Osprian, I.; Kroutil, W.; Mischitz, M.; Faber, K. Biocatalytic resolution of 2-methyl-2-(aryl)alkyloxiranes using novel bacterial epoxide hydrolases. Tetrahedron Asymmetry 1997, 8, 65–71. [Google Scholar] [CrossRef]
  52. Woo, J.-H.; Kang, K.-M.; Kwon, T.-H.; Park, N.-H.; Lee, E.Y. Isolation, identification and characterization of marine bacteria exhibiting complementary enantioselective epoxide hydrolase activity for preparing chiral chlorinated styrene oxide derivatives. J. Ind. Eng. Chem. 2015, 28, 225–228. [Google Scholar] [CrossRef]
  53. Woo, J.-H.; Kwon, T.-H.; Kim, J.-T.; Kim, C.-G.; Lee, E.Y. Identification and characterization of epoxide hydrolase activity of polycyclic aromatic hydrocarbon-degrading bacteria for biocatalytic resolution of racemic styrene oxide and styrene oxide derivatives. Biotechnol. Lett. 2013, 35, 599–606. [Google Scholar] [CrossRef]
  54. Woo, J.-H.; Kim, H.-S.; Park, N.-H.; Suk, H.Y. Isolation of a novel strain, Sphingorhabdus sp. YGSMI21 and characterization of its enantioselective epoxide hydrolase activity. J. Microbiol. 2021, 59, 675–680. [Google Scholar] [CrossRef] [PubMed]
  55. Zocher, F.; Enzelberger, M.M.; Bornscheuer, U.T.; Hauer, B.; Schmid, R.D. A colorimetric assay suitable for screening epoxide hydrolase activity. Anal. Chim. Acta 1999, 391, 345–351. [Google Scholar] [CrossRef]
  56. Wahler, D.; Reymond, J.-L. The Adrenaline Test for Enzymes. Angew. Chem. Int. Ed. 2002, 41, 1229–1232. [Google Scholar] [CrossRef]
  57. Mateo, C.; Archelas, A.; Furstoss, R. A spectrophotometric assay for measuring and detecting an epoxide hydrolase activity. Anal. Biochem. 2003, 314, 135–141. [Google Scholar] [CrossRef]
  58. Saini, P.; Wani, S.I.; Kumar, R.; Chhabra, R.; Chimni, S.S.; Sareen, D. Trigger factor assisted folding of the recombinant epoxide hydrolases identified from C. pelagibacter and S. nassauensis. Protein Expr. Purif. 2014, 104, 71–84. [Google Scholar] [CrossRef]
  59. van Loo, B.; Kingma, J.; Arand, M.; Wubbolts, M.G.; Janssen, D.B. Diversity and Biocatalytic Potential of Epoxide Hydrolases Identified by Genome Analysis. Appl. Environ. Microbiol. 2006, 72, 2905–2917. [Google Scholar] [CrossRef] [Green Version]
  60. Stojanovski, G.; Dobrijevic, D.; Hailes, H.C.; Ward, J.M. Identification and catalytic properties of new epoxide hydrolases from the genomic data of soil bacteria. Enzym. Microb. Technol. 2020, 139, 109592. [Google Scholar] [CrossRef] [PubMed]
  61. Robinson, S.L.; Piel, J.; Sunagawa, S. A roadmap for metagenomic enzyme discovery. Nat. Prod. Rep. 2021, 38, 1994–2023. [Google Scholar] [CrossRef] [PubMed]
  62. Jiménez, D.J.; Dini-Andreote, F.; Ottoni, J.R.; de Oliveira, V.M.; van Elsas, J.D.; Andreote, F.D. Compositional profile of α/β-hydrolase fold proteins in mangrove soil metagenomes: Prevalence of epoxide hydrolases and haloalkane dehalogenases in oil-contaminated sites. Microb. Biotechnol. 2015, 8, 604–613. [Google Scholar] [CrossRef] [PubMed]
  63. Kotik, M.; Štěpánek, V.; Marešová, H.; Kyslík, P.; Archelas, A. Environmental DNA as a source of a novel epoxide hydrolase reacting with aliphatic terminal epoxides. J. Mol. Catal. B Enzym. 2009, 56, 288–293. [Google Scholar] [CrossRef]
  64. Kotik, M.; Štěpánek, V.; Grulich, M.; Kyslík, P.; Archelas, A. Access to enantiopure aromatic epoxides and diols using epoxide hydrolases derived from total biofilter DNA. J. Mol. Catal. B Enzym. 2010, 65, 41–48. [Google Scholar] [CrossRef]
  65. Ferrandi, E.E.; Sayer, C.; Isupov, M.N.; Annovazzi, C.; Marchesi, C.; Iacobone, G.; Peng, X.; Bonch-Osmolovskaya, E.; Wohlgemuth, R.; Littlechild, J.A.; et al. Discovery and characterization of thermophilic limonene-1,2-epoxide hydrolases from hot spring metagenomic libraries. FEBS J. 2015, 282, 2879–2894. [Google Scholar] [CrossRef]
  66. Ferrandi, E.E.; Sayer, C.; De Rose, S.A.; Guazzelli, E.; Marchesi, C.; Saneei, V.; Isupov, M.N.; Littlechild, J.A.; Monti, D. New Thermophilic α/β Class Epoxide Hydrolases Found in Metagenomes from Hot Environments. Front. Bioeng. Biotechnol. 2018, 6, 144. [Google Scholar] [CrossRef] [Green Version]
  67. Reetz, M.T. Laboratory Evolution of Stereoselective Enzymes: A Prolific Source of Catalysts for Asymmetric Reactions. Angew. Chem. Int. Ed. 2011, 50, 138–174. [Google Scholar] [CrossRef]
  68. Basheer, S.M.; Chellappan, S. Enzyme Engineering. In Bioresources and Bioprocess in Biotechnology: Volume 2: Exploring Potential Biomolecules; Sugathan, S., Pradeep, N.S., Abdulhameed, S., Eds.; Springer: Singapore, 2017; pp. 151–168. [Google Scholar]
  69. Nardini, M.; Rink, R.; Janssen, D.B.; Dijkstra, B.W. Structure and mechanism of the epoxide hydrolase from Agrobacterium radiobacter AD1. J. Mol. Catal. B Enzym. 2001, 11, 1035–1042. [Google Scholar] [CrossRef]
  70. Zou, J.; Hallberg, B.M.; Bergfors, T.; Oesch, F.; Arand, M.; Mowbray, S.L.; Jones, T.A. Structure of Aspergillus niger epoxide hydrolase at 1.8 Å resolution: Implications for the structure and function of the mammalian microsomal class of epoxide hydrolases. Structure 2000, 8, 111–122. [Google Scholar] [CrossRef] [Green Version]
  71. Arand, M.; Hallberg, B.M.; Zou, J.; Bergfors, T.; Oesch, F.; van der Werf, M.J.; de Bont, J.A.; Jones, T.A.; Mowbray, S.L. Structure of Rhodococcus erythropolis limonene-1,2-epoxide hydrolase reveals a novel active site. EMBO J. 2003, 22, 2583–2592. [Google Scholar] [CrossRef] [Green Version]
  72. Johansson, P.; Unge, T.; Cronin, A.; Arand, M.; Bergfors, T.; Jones, T.A.; Mowbray, S.L. Structure of an atypical epoxide hydrolase from Mycobacterium tuberculosis gives insights into its function. J. Mol. Biol. 2005, 351, 1048–1056. [Google Scholar] [CrossRef]
  73. Reetz, M.T.; Zonta, A.; Schimossek, K.; Jaeger, K.-E.; Liebeton, K. Creation of Enantioselective Biocatalysts for Organic Chemistry by in vitro Evolution. Angew. Chem. Int. Ed. Engl. 1997, 36, 2830–2832. [Google Scholar] [CrossRef]
  74. Cedrone, F.; Niel, S.; Roca, S.; Bhatnagar, T.; Ait-abdelkader, N.; Torre, C.; Krumm, H.; Maichele, A.; Reetz, M.T.; Baratti, J.C. Directed Evolution of the Epoxide Hydrolase from Aspergillus niger. Biocatal. Biotransf. 2003, 21, 357–364. [Google Scholar] [CrossRef]
  75. Reetz, M.T.; Torre, C.; Eipper, A.; Lohmer, R.; Hermes, M.; Brunner, B.; Maichele, A.; Bocola, M.; Arand, M.; Cronin, A.; et al. Enhancing the Enantioselectivity of an Epoxide Hydrolase by Directed Evolution. Org. Lett. 2004, 6, 177–180. [Google Scholar] [CrossRef]
  76. Rui, L.; Cao, L.; Chen, W.; Reardon, K.F.; Wood, T.K. Active Site Engineering of the Epoxide Hydrolase from Agrobacterium radiobacter AD1 to Enhance Aerobic Mineralization of cis-1,2-Dichloroethylene in Cells Expressing an Evolved Toluene ortho-Monooxygenase. J. Biol. Chem. 2004, 279, 46810–46817. [Google Scholar] [CrossRef] [Green Version]
  77. Rui, L.; Cao, L.; Chen, W.; Reardon, K.F.; Wood, T.K. Protein Engineering of Epoxide Hydrolase from Agrobacterium radiobacter AD1 for Enhanced Activity and Enantioselective Production of (R)-1-Phenylethane-1,2-Diol. Appl. Environ. Microbiol. 2005, 71, 3995–4003. [Google Scholar] [CrossRef] [Green Version]
  78. Sun, Z.; Lonsdale, R.; Kong, X.-D.; Xu, J.-H.; Zhou, J.; Reetz, M.T. Reshaping an Enzyme Binding Pocket for Enhanced and Inverted Stereoselectivity: Use of Smallest Amino Acid Alphabets in Directed Evolution. Angew. Chem. Int. Ed. 2015, 54, 12410–12415. [Google Scholar] [CrossRef]
  79. Sun, Z.; Lonsdale, R.; Li, G.; Reetz, M.T. Comparing Different Strategies in Directed Evolution of Enzyme Stereoselectivity: Single- versus Double-Code Saturation Mutagenesis. ChemBioChem 2016, 17, 1865–1872. [Google Scholar] [CrossRef] [PubMed]
  80. Sun, Z.; Lonsdale, R.; Wu, L.; Li, G.; Li, A.; Wang, J.; Zhou, J.; Reetz, M.T. Structure-Guided Triple-Code Saturation Mutagenesis: Efficient Tuning of the Stereoselectivity of an Epoxide Hydrolase. ACS Catal. 2016, 6, 1590–1597. [Google Scholar] [CrossRef]
  81. Arabnejad, H.; Bombino, E.; Colpa, D.I.; Jekel, P.A.; Trajkovic, M.; Wijma, H.J.; Janssen, D.B. Computational Design of Enantiocomplementary Epoxide Hydrolases for Asymmetric Synthesis of Aliphatic and Aromatic Diols. ChemBioChem 2020, 21, 1893–1904. [Google Scholar] [CrossRef] [PubMed]
  82. Hu, D.; Zong, X.-C.; Xue, F.; Li, C.; Hu, B.-C.; Wu, M.-C. Manipulating regioselectivity of an epoxide hydrolase for single enzymatic synthesis of (R)-1,2-diols from racemic epoxides. Chem. Commun. 2020, 56, 2799–2802. [Google Scholar] [CrossRef]
  83. Li, G.; Qin, Y.; Fontaine, N.T.; Ng Fuk Chong, M.; Maria-Solano, M.A.; Feixas, F.; Cadet, X.F.; Pandjaitan, R.; Garcia-Borràs, M.; Cadet, F.; et al. Machine Learning Enables Selection of Epistatic Enzyme Mutants for Stability against Unfolding and Detrimental Aggregation. ChemBioChem 2021, 22, 904–914. [Google Scholar] [CrossRef] [PubMed]
  84. Kong, X.-D.; Ma, Q.; Zhou, J.; Zeng, B.-B.; Xu, J.-H. A Smart Library of Epoxide Hydrolase Variants and the Top Hits for Synthesis of (S)-β-Blocker Precursors. Angew. Chem. Int. Ed. 2014, 53, 6641–6644. [Google Scholar] [CrossRef] [PubMed]
  85. van Loo, B.; Spelberg, J.H.L.; Kingma, J.; Sonke, T.; Wubbolts, M.G.; Janssen, D.B. Directed Evolution of Epoxide Hydrolase from A. radiobacter toward Higher Enantioselectivity by Error-Prone PCR and DNA Shuffling. Chem. Biol. 2004, 11, 981–990. [Google Scholar] [CrossRef] [Green Version]
  86. van Loo, B.; Kingma, J.; Heyman, G.; Wittenaar, A.; Lutje Spelberg, J.H.; Sonke, T.; Janssen, D.B. Improved enantioselective conversion of styrene epoxides and meso-epoxides through epoxide hydrolases with a mutated nucleophile-flanking residue. Enzym. Microb. Technol. 2009, 44, 145–153. [Google Scholar] [CrossRef]
  87. Xue, F.; Liu, Z.-Q.; Wan, N.-W.; Zhu, H.-Q.; Zheng, Y.-G. Engineering the epoxide hydrolase from Agromyces mediolanus for enhanced enantioselectivity and activity in the kinetic resolution of racemic epichlorohydrin. RSC Adv. 2015, 5, 31525–31532. [Google Scholar] [CrossRef]
  88. Reetz, M.T.; Wang, L.-W.; Bocola, M. Directed Evolution of Enantioselective Enzymes: Iterative Cycles of CASTing for Probing Protein-Sequence Space. Angew. Chem. Int. Ed. 2006, 45, 1236–1241. [Google Scholar] [CrossRef]
  89. Reetz, M.T.; Zheng, H. Manipulating the Expression Rate and Enantioselectivity of an Epoxide Hydrolase by Using Directed Evolution. ChemBioChem 2011, 12, 1529–1535. [Google Scholar] [CrossRef]
  90. Gumulya, Y.; Sanchis, J.; Reetz, M.T. Many Pathways in Laboratory Evolution Can Lead to Improved Enzymes: How to Escape from Local Minima. ChemBioChem 2012, 13, 1060–1066. [Google Scholar] [CrossRef]
  91. Hu, D.; Hu, B.-C.; Hou, X.-D.; Zhang, D.; Lei, Y.-Q.; Rao, Y.-J.; Wu, M.-C. Structure-Guided Regulation in the Enantioselectivity of an Epoxide Hydrolase to Produce Enantiomeric Monosubstituted Epoxides and Vicinal Diols via Kinetic Resolution. Org. Lett. 2022, 24, 1757–1761. [Google Scholar] [CrossRef]
  92. Zheng, H.; Reetz, M.T. Manipulating the Stereoselectivity of Limonene Epoxide Hydrolase by Directed Evolution Based on Iterative Saturation Mutagenesis. J. Am. Chem. Soc. 2010, 132, 15744–15751. [Google Scholar] [CrossRef] [PubMed]
  93. Sun, Z.; Salas, P.T.; Siirola, E.; Lonsdale, R.; Reetz, M.T. Exploring productive sequence space in directed evolution using binary patterning versus conventional mutagenesis strategies. Bioresour. Bioprocess 2016, 3, 44. [Google Scholar] [CrossRef] [Green Version]
  94. Li, G.; Zhang, H.; Sun, Z.; Liu, X.; Reetz, M.T. Multiparameter Optimization in Directed Evolution: Engineering Thermostability, Enantioselectivity, and Activity of an Epoxide Hydrolase. ACS Catal. 2016, 6, 3679–3687. [Google Scholar] [CrossRef]
  95. Li, A.; Acevedo-Rocha, C.G.; Sun, Z.; Cox, T.; Xu, J.L.; Reetz, M.T. Beating Bias in the Directed Evolution of Proteins: Combining High-Fidelity on-Chip Solid-Phase Gene Synthesis with Efficient Gene Assembly for Combinatorial Library Construction. ChemBioChem 2018, 19, 221–228. [Google Scholar] [CrossRef] [PubMed]
  96. Wijma, H.J.; Floor, R.J.; Bjelic, S.; Marrink, S.J.; Baker, D.; Janssen, D.B. Enantioselective Enzymes by Computational Design and in silico Screening. Angew. Chem. Int. Ed. 2015, 54, 3726–3730. [Google Scholar] [CrossRef] [PubMed]
  97. Gurell, A.; Widersten, M. Modification of Substrate Specificity Resulting in an Epoxide Hydrolase with Shifted Enantiopreference for (2,3-Epoxypropyl)benzene. ChemBioChem 2010, 11, 1422–1429. [Google Scholar] [CrossRef]
  98. Carlsson, Å.J.; Bauer, P.; Ma, H.; Widersten, M. Obtaining Optical Purity for Product Diols in Enzyme-Catalyzed Epoxide Hydrolysis: Contributions from Changes in both Enantio- and Regioselectivity. Biochemistry 2012, 51, 7627–7637. [Google Scholar] [CrossRef]
  99. Li, Y.; Ou, X.; Guo, Z.; Zong, M.; Lou, W. Using multiple site-directed modification of epoxide hydrolase to significantly improve its enantioselectivity in hydrolysis of rac-glycidyl phenyl ether. Chin. J. Chem. Eng. 2020, 28, 2181–2189. [Google Scholar] [CrossRef]
  100. Kotik, M.; Zhao, W.; Iacazio, G.; Archelas, A. Directed evolution of metagenome-derived epoxide hydrolase for improved enantioselectivity and enantioconvergence. J. Mol. Catal. B Enzym. 2013, 91, 44–51. [Google Scholar] [CrossRef]
  101. Kotik, M.; Archelas, A.; Faměrová, V.; Oubrechtová, P.; Křen, V. Laboratory evolution of an epoxide hydrolase—Towards an enantioconvergent biocatalyst. J. Biotechnol. 2011, 156, 1–10. [Google Scholar] [CrossRef]
  102. Liu, Y.-Y.; Wu, M.-D.; Zhu, X.-X.; Zhang, X.-D.; Zhang, C.; Xu, Y.-H.; Wu, M.-C. Remarkable improvement in the regiocomplementarity of a Glycine max epoxide hydrolase by reshaping its substrate-binding pocket for the enantioconvergent preparation of (R)-hexane-1,2-diol. Mol. Catal. 2021, 514, 111851. [Google Scholar] [CrossRef]
  103. Ye, H.-H.; Hu, D.; Shi, X.-L.; Wu, M.-C.; Deng, C.; Li, J.-F. Directed modification of a novel epoxide hydrolase from Phaseolus vulgaris to improve its enantioconvergence towards styrene epoxides. Catal. Commun. 2016, 87, 32–35. [Google Scholar] [CrossRef]
  104. Zong, X.-C.; Li, C.; Xu, Y.-H.; Hu, D.; Hu, B.-C.; Zang, J.; Wu, M.-C. Substantially improving the enantioconvergence of PvEH1, a Phaseolus vulgaris epoxide hydrolase, towards m-chlorostyrene oxide by laboratory evolution. Microb. Cell Fact. 2019, 18, 202. [Google Scholar] [CrossRef] [Green Version]
  105. Zhu, X.-X.; Hu, B.-C.; Lin, W.-Q.; Zhang, D.; Zhao, J.; Wu, M.-C. Engineering the regiocomplementarity of an epoxide hydrolase from Rhodotorula paludigena by means of computer-aided design for the scale-up enantioconvergent hydrolysis of racemic m-nitrostyrene oxide. Biochem. Eng. J. 2022, 180, 108359. [Google Scholar] [CrossRef]
  106. Li, F.-L.; Kong, X.-D.; Chen, Q.; Zheng, Y.-C.; Xu, Q.; Chen, F.-F.; Fan, L.-Q.; Lin, G.-Q.; Zhou, J.; Yu, H.-L.; et al. Regioselectivity Engineering of Epoxide Hydrolase: Near-Perfect Enantioconvergence through a Single Site Mutation. ACS Catal. 2018, 8, 8314–8317. [Google Scholar] [CrossRef]
  107. Li, F.-L.; Qiu, Y.-Y.; Zheng, Y.-C.; Chen, F.-F.; Kong, X.D.; Xu, J.-H.; Yu, H.-L. Reprogramming Epoxide Hydrolase to Improve Enantioconvergence in Hydrolysis of Styrene Oxide Scaffolds. Adv. Synth. Catal. 2020, 362, 4699–4706. [Google Scholar] [CrossRef]
  108. Wijma, H.J.; Floor, R.J.; Jekel, P.A.; Baker, D.; Marrink, S.J.; Janssen, D.B. Computationally designed libraries for rapid enzyme stabilization. Protein Eng. Des. Sel. 2014, 27, 49–58. [Google Scholar] [CrossRef] [Green Version]
  109. Gumulya, Y.; Reetz, M.T. Enhancing the Thermal Robustness of an Enzyme by Directed Evolution: Least Favorable Starting Points and Inferior Mutants Can Map Superior Evolutionary Pathways. ChemBioChem 2011, 12, 2502–2510. [Google Scholar] [CrossRef]
  110. Qiao, P.; Wu, M.; Zhu, L.; Zhang, Y.; Yang, L.; Fei, H.; Lin, J. Enhancing the thermal tolerance of a cis-epoxysuccinate hydrolase via combining directed evolution with various semi-rational redesign methods. J. Mol. Catal. B Enzym. 2015, 121, 96–103. [Google Scholar] [CrossRef]
  111. Jochens, H.; Stiba, K.; Savile, C.; Fujii, R.; Yu, J.-G.; Gerassenkov, T.; Kazlauskas, R.J.; Bornscheuer, U.T. Converting an Esterase into an Epoxide Hydrolase. Angew. Chem. Int. Ed. 2009, 48, 3532–3535. [Google Scholar] [CrossRef] [PubMed]
  112. Nagel, B.; Dellweg, H.; Gierasch, L.M. Glossary for chemists of terms used in biotechnology (IUPAC Recommendations 1992). Pure Appl. Chem. 1992, 64, 143–168. [Google Scholar] [CrossRef] [Green Version]
  113. Hechtberger, P.; Wirnsberger, G.; Mischitz, M.; Klempier, N.; Faber, K. Asymmetric hydrolysis of epoxides using an immobilized enzyme preparation from Rhodococcus sp. Tetrahedron Asymmetry 1993, 4, 1161–1164. [Google Scholar] [CrossRef]
  114. Lin, H.; Liu, J.-Y.; Wang, H.-B.; Ahmed, A.A.Q.; Wu, Z.-L. Biocatalysis as an alternative for the production of chiral epoxides: A comparative review. J. Mol. Catal. B Enzym. 2011, 72, 77–89. [Google Scholar] [CrossRef]
  115. Vikartovská, A.; Bučko, M.; Gemeiner, P.; Nahálka, J.; Pätoprstý, V.; Hrabárová, E. Flow Calorimetry—A Useful Tool for Determination of Immobilized cis-Epoxysuccinate Hydrolase Activity from Nocardia tartaricans. Artif. Cells Blood Substit. Biotechnol. 2004, 32, 77–89. [Google Scholar] [CrossRef]
  116. Bučko, M.; Vikartovská, A.; Lacík, I.; Kolláriková, G.; Gemeiner, P.; Pätoprstý, V.; Brygin, M. Immobilization of a whole-cell epoxide-hydrolyzing biocatalyst in sodium alginate-cellulose sulfate-poly(methylene-co-guanidine) capsules using a controlled encapsulation process. Enzym. Microb. Technol. 2005, 36, 118–126. [Google Scholar] [CrossRef]
  117. Rosenberg, M.; Miková, H.; Krištofíková, L. Production of L-tartaric acid by immobilized bacterial cells Nocardia tartaricans. Biotechnol. Lett. 1999, 21, 491–495. [Google Scholar] [CrossRef]
  118. Bučko, M.; Vikartovská, A.; Gemeiner, P.; Lacík, I.; Kolláriková, G.; Marison, I.W. Nocardia tartaricans cells immobilized in sodium alginate–cellulose sulfate–poly (methylene-co-guanidine)capsules: Mechanical resistance and operational stability. J. Chem. Technol. Biotechnol. 2006, 81, 500–504. [Google Scholar] [CrossRef]
  119. Huang, R.; Wu, M.; Goldman, M.J.; Li, Z. Encapsulation of enzyme via one-step template-free formation of stable organic–inorganic capsules: A simple and efficient method for immobilizing enzyme with high activity and recyclability. Biotechnol. Bioeng. 2015, 112, 1092–1101. [Google Scholar] [CrossRef]
  120. Zhou, R.; Dong, S.; Feng, Y.; Cui, Q.; Xuan, J. Development of highly efficient whole-cell catalysts of cis-epoxysuccinic acid hydrolase by surface display. Bioresour. Bioprocess 2022, 9, 92. [Google Scholar] [CrossRef]
  121. Zou, S.-P.; Wang, Z.-C.; Qin, C.; Zheng, Y.-G. Covalent immobilization of Agrobacterium radiobacter epoxide hydrolase on ethylenediamine functionalised epoxy supports for biocatalytical synthesis of (R)-epichlorohydrin. Biotechnol. Lett. 2016, 38, 1579–1585. [Google Scholar] [CrossRef] [PubMed]
  122. Zou, S.-P.; Xuan, X.-L.; Wang, Z.-J.; Zheng, Y.-G. Conjugation of Agrobacterium radiobacter epoxide hydrolase with ficoll: Catalytic, kinetic and thermodynamic analysis. Int. J. Biol. Macromol. 2018, 119, 1098–1105. [Google Scholar] [CrossRef] [PubMed]
  123. Grulich, M.; Maršálek, J.; Kyslík, P.; Štěpánek, V.; Kotik, M. Production, enrichment and immobilization of a metagenome-derived epoxide hydrolase. Process Biochem. 2011, 46, 526–532. [Google Scholar] [CrossRef]
  124. Mateo, C.; Archelas, A.; Fernandez-Lafuente, R.; Manuel Guisan, J.; Furstoss, R. Enzymatic transformations. Immobilized A. niger epoxide hydrolase as a novel biocatalytic tool for repeated-batch hydrolytic kinetic resolution of epoxides. Org. Biomol. Chem. 2003, 1, 2739–2743. [Google Scholar] [CrossRef]
  125. Yildirim, D.; Tükel, S.S.; Alagöz, D.; Alptekin, Ö. Preparative-scale kinetic resolution of racemic styrene oxide by immobilized epoxide hydrolase. Enzym. Microb. Technol. 2011, 49, 555–559. [Google Scholar] [CrossRef] [PubMed]
  126. Yildirim, D.; Tükel, S.S.; Alptekin, Ö.; Alagöz, D. Immobilized Aspergillus niger epoxide hydrolases: Cost-effective biocatalysts for the prepation of enantiopure styrene oxide, propylene oxide and epichlorohydrin. J. Mol. Catal. B Enzym. 2013, 88, 84–90. [Google Scholar] [CrossRef]
  127. Petri, A.; Marconcini, P.; Salvadori, P. Efficient immobilization of epoxide hydrolase onto silica gel and use in the enantioselective hydrolysis of racemic para-nitrostyrene oxide. J. Mol. Catal. B Enzym. 2005, 32, 219–224. [Google Scholar] [CrossRef]
  128. Mateo, C.; Fernandez-Lafuente, R.; Archelas, A.; Guisan, J.M.; Furstoss, R. Preparation of a very stable immobilized Solanum tuberosum epoxide hydrolase. Tetrahedron Asymmetry 2007, 18, 1233–1238. [Google Scholar] [CrossRef]
  129. Ibrahim, M.; Hubert, P.; Dellacherie, E.; Magdalou, J.; Siest, G. Immobilization of epoxide hydrolase purified from rat liver microsomes. Biotechnol. Lett. 1984, 6, 771–776. [Google Scholar] [CrossRef]
  130. Ibrahim, M.; Hubert, P.; Dellacherie, E.; Magdalou, J.; Muller, J.; Siest, G. Covalent attachment of epoxide hydrolase to dextran. Enzym. Microb. Technol. 1985, 7, 66–72. [Google Scholar] [CrossRef]
  131. Kamble, M.; Salvi, H.; Yadav, G.D. Preparation of amino-functionalized silica supports for immobilization of epoxide hydrolase and cutinase: Characterization and applications. J. Porous Mater. 2020, 27, 1559–1567. [Google Scholar] [CrossRef]
  132. Cassimjee, K.E.; Kourist, R.; Lindberg, D.; Wittrup Larsen, M.; Thanh, N.H.; Widersten, M.; Bornscheuer, U.T.; Berglund, P. One-step enzyme extraction and immobilization for biocatalysis applications. Biotechnol. J. 2011, 6, 463–469. [Google Scholar] [CrossRef] [PubMed]
  133. Choi, S.H.; Kim, H.S.; Lee, I.S.; Lee, E.Y. Functional expression and magnetic nanoparticle-based Immobilization of a protein-engineered marine fish epoxide hydrolase of Mugil cephalus for enantioselective hydrolysis of racemic styrene oxide. Biotechnol. Lett. 2010, 32, 1685–1691. [Google Scholar] [CrossRef]
  134. Wang, Z.; Su, M.; Li, Y.; Wang, Y.; Su, Z. Production of tartaric acid using immobilized recominant cis-epoxysuccinate hydrolase. Biotechnol. Lett. 2017, 39, 1859–1863. [Google Scholar] [CrossRef]
  135. Karboune, S.; Archelas, A.; Baratti, J.C. Free and immobilized Aspergillus niger epoxide hydrolase-catalyzed hydrolytic kinetic resolution of racemic p-chlorostyrene oxide in a neat organic solvent medium. Process Biochem. 2010, 45, 210–216. [Google Scholar] [CrossRef]
  136. Karboune, S.; Archelas, A.; Furstoss, R.; Baratti, J. Immobilization of epoxide hydrolase from Aspergillus niger onto DEAE-cellulose: Enzymatic properties and application for the enantioselective resolution of a racemic epoxide. J. Mol. Catal. B Enzym. 2005, 32, 175–183. [Google Scholar] [CrossRef]
  137. Karboune, S.; Amourache, L.; Nellaiah, H.; Morisseau, C.; Baratti, J. Immobilization of the epoxide hydrolase from Aspergillus niger. Biotechnol. Lett. 2001, 23, 1633–1639. [Google Scholar] [CrossRef]
  138. Kroutil, W.; Genzel, Y.; Pietzsch, M.; Syldatk, C.; Faber, K. Purification and characterization of a highly selective epoxide hydrolase from Nocardia sp. EH1. J. Biotechnol. 1998, 61, 143–150. [Google Scholar] [CrossRef]
  139. Jin, H.-X.; Liu, Z.-Q.; Hu, Z.-C.; Zheng, Y.-G. Production of (R)-epichlorohydrin from 1,3-dichloro-2-propanol by two-step biocatalysis using haloalcohol dehalogenase and epoxide hydrolase in two-phase system. Biochem. Eng. J. 2013, 74, 1–7. [Google Scholar] [CrossRef]
  140. Kim, Y.H.; Lee, I.; Choi, S.H.; Lee, O.K.; Shim, J.; Lee, J.; Kim, J.; Lee, E.Y. Enhanced stability and reusability of marine epoxide hydrolase using ship-in-a-bottle approach with magnetically-separable mesoporous silica. J. Mol. Catal. B Enzym. 2013, 89, 48–51. [Google Scholar] [CrossRef]
  141. Yu, C.-Y.; Li, X.-F.; Lou, W.-Y.; Zong, M.-H. Cross-linked enzyme aggregates of Mung bean epoxide hydrolases: A highly active, stable and recyclable biocatalyst for asymmetric hydrolysis of epoxides. J. Biotechnol. 2013, 166, 12–19. [Google Scholar] [CrossRef] [PubMed]
  142. Yu, C.-Y.; Wei, P.; Li, X.-F.; Zong, M.-H.; Lou, W.-Y. Using Ionic Liquid in a Biphasic System to Improve Asymmetric Hydrolysis of Styrene Oxide Catalyzed by cross-Linked Enzyme Aggregates (CLEAs) of Mung Bean Epoxide Hydrolases. Ind. Eng. Chem. Res. 2014, 53, 7923–7930. [Google Scholar] [CrossRef]
  143. Kronenburg, N.A.E.; de Bont, J.A.M.; Fischer, L. Improvement of enantioselectivity by immobilized imprinting of epoxide hydrolase from Rhodotorula glutinis. J. Mol. Catal. B Enzym. 2001, 16, 121–129. [Google Scholar] [CrossRef]
  144. Salvi, H.M.; Yadav, G.D. Organic-inorganic epoxide hydrolase hybrid nanoflowers with enhanced catalytic activity: Hydrolysis of styrene oxide to 1-phenyl-1,2-ethanediol. J. Biotechnol. 2021, 341, 113–120. [Google Scholar] [CrossRef]
  145. Cao, S.-L.; Yue, D.-M.; Li, X.-H.; Smith, T.J.; Li, N.; Zong, M.-H.; Wu, H.; Ma, Y.-Z.; Lou, W.-Y. Novel Nano-/Micro-Biocatalyst: Soybean Epoxide Hydrolase Immobilized on UiO-66-NH2 MOF for Efficient Biosynthesis of Enantiopure (R)-1, 2-Octanediol in Deep Eutectic Solvents. ACS Sustain. Chem. Eng. 2016, 4, 3586–3595. [Google Scholar] [CrossRef]
  146. Onur, H.; Tülek, A.; Aslan, E.S.; Binay, B.; Yildirim, D. A new highly enantioselective stable epoxide hydrolase from Hypsibius dujardini: Expression in Pichia pastoris and immobilization in ZIF-8 for asymmetric hydrolysis of racemic styrene oxide. Biochem. Eng. J. 2022, 189, 108726. [Google Scholar] [CrossRef]
  147. Maritz, J.; Krieg, H.M.; Yeates, C.A.; Botes, A.L.; Breytenbach, J.C. Calcium alginate entrapment of the yeast Rhodosporidium toruloides for the kinetic resolution of 1,2-epoxyoctane. Biotechnol. Lett. 2003, 25, 1775–1781. [Google Scholar] [CrossRef]
  148. Bao, W.; Pan, H.; Zhang, Z.; Cheng, Y.; Xie, Z.; Zhang, J. Isolation of the stable strain Labrys sp. BK-8 for l(+)-tartaric acid production. J. Biosci. Bioeng. 2015, 119, 538–542. [Google Scholar] [CrossRef] [PubMed]
  149. Buchholz, K.; Kasche, V.; Bornscheuer, U.T. Enzymes in Organic Chemistry. In Biocatalysts and Enzyme Technology, 1st ed.; WILEY-VCH Verlag GmbH & Co. KGaA: Weinheim, Germany, 2005; pp. 141–145. [Google Scholar]
  150. Faber, K.; Mischitz, M.; Kroutil, W. Microbial Epoxide Hydrolases. Acta Chem. Scand. 1996, 50, 249–258. [Google Scholar] [CrossRef] [Green Version]
  151. Mayer, S.F.; Steinreiber, A.; Orru, R.V.A.; Faber, K. Enzyme-Triggered Enantioconvergent Transformation of Haloalkyl Epoxides. Eur. J. Org. Chem. 2001, 2001, 4537–4542. [Google Scholar] [CrossRef]
  152. Xu, Y.; Jia, X.; Panke, S.; Li, Z. Asymmetric dihydroxylation of arylolefins by sequential enantioselective epoxidation and regioselective hydrolysis with tandem biocatalysts. Chem. Commun. 2009, 12, 1481–1483. [Google Scholar] [CrossRef]
  153. Pandey, R.K.; Fernandes, R.A.; Kumar, P. An asymmetric dihydroxylation route to enantiomerically pure norfluoxetine and fluoxetine. Tetrahedron Lett. 2002, 43, 4425–4426. [Google Scholar] [CrossRef]
  154. Schrittwieser, J.H.; Velikogne, S.; Hall, M.; Kroutil, W. Artificial Biocatalytic Linear Cascades for Preparation of Organic Molecules. Chem. Rev. 2018, 118, 270–348. [Google Scholar] [CrossRef] [Green Version]
  155. Zhang, J.; Wu, S.; Wu, J.; Li, Z. Enantioselective Cascade Biocatalysis via Epoxide Hydrolysis and Alcohol Oxidation: One-Pot Synthesis of (R)-α-Hydroxy Ketones from Meso- or Racemic Epoxides. ACS Catal. 2015, 5, 51–58. [Google Scholar] [CrossRef]
  156. Wu, S.; Chen, Y.; Xu, Y.; Li, A.; Xu, Q.; Glieder, A.; Li, Z. Enantioselective trans-Dihydroxylation of Aryl Olefins by Cascade Biocatalysis with Recombinant Escherichia coli Coexpressing Monooxygenase and Epoxide Hydrolase. ACS Catal. 2014, 4, 409–420. [Google Scholar] [CrossRef]
  157. Xue, F.; Liu, Z.-Q.; Wang, Y.-J.; Zhu, H.-Q.; Wan, N.-W.; Zheng, Y.-G. Efficient synthesis of (S)-epichlorohydrin in high yield by cascade biocatalysis with halohydrin dehalogenase and epoxide hydrolase mutants. Catal. Commun. 2015, 72, 147–149. [Google Scholar] [CrossRef]
  158. Gao, P.; Wu, S.; Praveen, P.; Loh, K.-C.; Li, Z. Enhancing productivity for cascade biotransformation of styrene to (S)-vicinal diol with biphasic system in hollow fiber membrane bioreactor. Appl. Microbiol. Biotechnol. 2017, 101, 1857–1868. [Google Scholar] [CrossRef] [PubMed]
  159. Wang, L.; Parnell, A.; Williams, C.; Bakar, N.A.; Challand, M.R.; van der Kamp, M.W.; Simpson, T.J.; Race, P.R.; Crump, M.P.; Willis, C.L. A Rieske oxygenase/epoxide hydrolase-catalysed reaction cascade creates oxygen heterocycles in mupirocin biosynthesis. Nat. Catal. 2018, 1, 968–976. [Google Scholar] [CrossRef] [Green Version]
  160. Liu, S.; Zhang, X.; Liu, F.; Xu, M.; Yang, T.; Long, M.; Zhou, J.; Osire, T.; Yang, S.; Rao, Z. Designing of a Cofactor Self-Sufficient Whole-Cell Biocatalyst System for Production of 1,2-Amino Alcohols from Epoxides. ACS Synth. Biol. 2019, 8, 734–743. [Google Scholar] [CrossRef] [PubMed]
  161. Zhang, J.-D.; Yang, X.-X.; Jia, Q.; Zhao, J.-W.; Gao, L.-L.; Gao, W.-C.; Chang, H.-H.; Wei, W.-L.; Xu, J.-H. Asymmetric ring opening of racemic epoxides for enantioselective synthesis of (S)-β-amino alcohols by a cofactor self-sufficient cascade biocatalysis system. Catal. Sci. Technol. 2019, 9, 70–74. [Google Scholar] [CrossRef]
  162. Zhang, J.; Yang, X.; Dong, R.; Gao, L.; Li, J.; Li, X.; Huang, S.; Zhang, C.; Chang, H. Cascade Biocatalysis for Regio- and Stereoselective Aminohydroxylation of Styrenyl Olefins to Enantiopure Arylglycinols. ACS Sustain. Chem. Eng. 2020, 8, 18277–18285. [Google Scholar] [CrossRef]
  163. See, W.W.L.; Choo, J.P.S.; Jung, D.-Y.; Zhi, L. Recent developments in oxidative biocatalytic cascades. Curr. Opin. Green Sustain. Chem. 2022, 38, 100700. [Google Scholar] [CrossRef]
Ijms 24 07334 g001 550
Figure 1. (A) Differences between immobilization techniques based on covalent bonding and adsorption (left) or entrapment and encapsulation (right) from the perspective of applied mechanical forces (M) in the bioreactor represented by a rotating impeller. Mechanical abrasion was applied directly on the adsorbed or covalently bond biocatalyst (left). Entrapment and encapsulation matrices protected biocatalysts from direct mechanical forces (right). The contact of immobilized biocatalysts with substrates (S) and products (P) was direct, using covalent bonding and adsorption as immobilization principles. Entrapment and encapsulation require diffusion of S and P through the immobilization matrix or the semipermeable membrane. (B) The five most important aspects for choosing the immobilization technique for biocatalysts.
Figure 1. (A) Differences between immobilization techniques based on covalent bonding and adsorption (left) or entrapment and encapsulation (right) from the perspective of applied mechanical forces (M) in the bioreactor represented by a rotating impeller. Mechanical abrasion was applied directly on the adsorbed or covalently bond biocatalyst (left). Entrapment and encapsulation matrices protected biocatalysts from direct mechanical forces (right). The contact of immobilized biocatalysts with substrates (S) and products (P) was direct, using covalent bonding and adsorption as immobilization principles. Entrapment and encapsulation require diffusion of S and P through the immobilization matrix or the semipermeable membrane. (B) The five most important aspects for choosing the immobilization technique for biocatalysts.
Ijms 24 07334 g001
Ijms 24 07334 g002 550
Figure 2. An example of a non-natural enzyme cascade consisting of two steps, including styrene monooxygenase (SMO) in the form of resting E. coli cells and epoxide hydrolase (SpEH) as a cell-free extract for the production of (S)-1-phenyl-1,2,ethanediol [152] as an intermediate for synthesizing the pharmaceutical (R)-fluoxetine [153].
Figure 2. An example of a non-natural enzyme cascade consisting of two steps, including styrene monooxygenase (SMO) in the form of resting E. coli cells and epoxide hydrolase (SpEH) as a cell-free extract for the production of (S)-1-phenyl-1,2,ethanediol [152] as an intermediate for synthesizing the pharmaceutical (R)-fluoxetine [153].
Ijms 24 07334 g002
Ijms 24 07334 g003 550
Figure 3. The importance and schematic incorporation of epoxide hydrolase SpEH from Sphingomonas sp. HXN-200 into five independent, non-natural enzyme cascades consisting of two, three, and four enzymes, co-expressed and used in the form of five whole-cell systems, as mentioned in Table 4; S: substrate, P: product, CR: cofactor regeneration, CsR: co-substrate regeneration, NOX: NADH oxidase, and ADH: alcohol dehydrogenase [155,156,158,160,161,162].
Figure 3. The importance and schematic incorporation of epoxide hydrolase SpEH from Sphingomonas sp. HXN-200 into five independent, non-natural enzyme cascades consisting of two, three, and four enzymes, co-expressed and used in the form of five whole-cell systems, as mentioned in Table 4; S: substrate, P: product, CR: cofactor regeneration, CsR: co-substrate regeneration, NOX: NADH oxidase, and ADH: alcohol dehydrogenase [155,156,158,160,161,162].
Ijms 24 07334 g003
Table
Table 1. Chiral precursors that can be prepared by EHs and the products that can be synthesized from them.
Table 1. Chiral precursors that can be prepared by EHs and the products that can be synthesized from them.
Chiral PrecursorFinal ProductApplication of Final ProductReaction CommentRef.
Ijms 24 07334 i001Ijms 24 07334 i002Synthetic oxazolidinone antibiotic effective against Gram positive bacteriaSelective hydrolysis of (S)-enantiomer[18]
Ijms 24 07334 i003Cardio selective β-blocker for treatment of high blood pressure and heart associated chest pain
Ijms 24 07334 i004Dietary supplement, involved in long-chain fatty acid transport in cells
Ijms 24 07334 i005Ijms 24 07334 i006β-adrenergic blocker with antianginal and antiarrhythmic properties.Chemo-enzymatic enantioconvergent synthesis[19,20]
Ijms 24 07334 i007Ijms 24 07334 i008Neuroprotective agent (aspartate receptor antagonist)Sequential bi-enzymatic hydrolysis using 2 enantiocomplementary EHs[21]
Ijms 24 07334 i009Ijms 24 07334 i010HIV protease inhibitor MK 639Selective hydrolysis of 1(R),2(S)-enantiomer[22]
Ijms 24 07334 i011Ijms 24 07334 i012Non-steroidal anti-inflammatory drugSelective hydrolysis of (R)-enantiomer[23]
Ijms 24 07334 i013Ijms 24 07334 i014Calcium channel blockerKinetic resolution[24]
Ijms 24 07334 i015Ijms 24 07334 i016Anthracycline antibiotic with chemotherapeutic propertiesEnzymatic deracemization for production of (S)-diol used for chemical synthesis[25]
Ijms 24 07334 i017Ijms 24 07334 i018Major constituent of Matricaria chamomilla essential oil; ingredient for skin creams, lotions, ointments with anti-inflammatory, bactericidal and antimycotic propertiesChemo-enzymatic process for production of all 4 stereoisomers of bisabolol[15]
Ijms 24 07334 i019Ijms 24 07334 i020β-adrenergic receptor blocking drugsSelective hydrolysis of (R)-enantiomer[26]
Ijms 24 07334 i021Ijms 24 07334 i022
Ijms 24 07334 i023Ijms 24 07334 i024Neuromediator with antiepileptic and antihypertensive activitiesSelective hydrolysis of (S)-enantiomer[27]
Ijms 24 07334 i025Dietary supplement, involved in long-chain fatty acid transport in cells
Ijms 24 07334 i026Ijms 24 07334 i027IGF-1R kinase inhibitorSelective hydrolysis of (R)-enantiomer[28]
Ijms 24 07334 i028β3-adrenergic receptor agonistsEnantioconvergent hydrolysis of racemic epoxide[29]
Ijms 24 07334 i029
Ijms 24 07334 i030Ijms 24 07334 i031Antifungal triazole drugProduction of optically pure epoxide and diol that can be used for chemical synthesis of optically pure triazole derivatives[30,31]
Ijms 24 07334 i032Ijms 24 07334 i033Non-steroidal antiandrogen drug used for treatment of prostate cancerChemo-enzymatic synthesis of optically pure diol[32]
Ijms 24 07334 i034Ijms 24 07334 i035Ileal bile acid transport (iBAT) inhibitor indicated for diabetes type IIKinetic resolution[33]
Ijms 24 07334 i036Ijms 24 07334 i037Chiral precursors for synthesis of various steroidal compoundsKinetic resolution to produce both enantiomers of spiroepoxide, using 2 different EHs[34]
Ijms 24 07334 i038Ijms 24 07334 i039Melatonin receptor agonist used for treatment of sleep disordersSelective hydrolysis of (R)-enantiomer[35,36]
Ijms 24 07334 i040Ijms 24 07334 i041Calcium channel blocker used for treatment of hypertensionSelective hydrolysis of (R)-enantiomer[37]
Ijms 24 07334 i042Chiral chemical building block with broad applications in chemical, pharmaceutical, food industriesAsymmetric hydrolysis to produce optically pure diol[14]
Ijms 24 07334 i043Ijms 24 07334 i044Anticoagulant; direct factor Xa inhibitor developed by Bayer and marketed as Xarelto- 1[38,39]
Ijms 24 07334 i045Dietary supplement, involved in long-chain fatty acid transport in cells-[40,41]
Ijms 24 07334 i046Antidiabetic drug-[42]
Ijms 24 07334 i047β-adrenergic receptor blocking drug-[43]
Ijms 24 07334 i048β-adrenergic antagonist drug-[44,45]
1 Only potential application.
Table
Table 2. The summary of EHs engineered for higher enantioselectivity and improved enantioconvergence.
Table 2. The summary of EHs engineered for higher enantioselectivity and improved enantioconvergence.
Modified PropertySource of EH Used as
Template for Mutagenesis		Enzyme Engineering MethodMutantSubstrateE (-)eeP (%)C (%)Ref.
MutantWTMutantWTMutantWT
EnantioselectivityAgrobacterium radiobacter AD1Directed evolution—Error-prone PCR and DNA shufflingF108I/P205H/Y215H/E271Vstyrene oxide>5016NI 1NI[85]
p-nitrostyrene oxide8156NINI
p-nitrophenyl glycidyl ether323.4NINI
epichlorohydrin40<2NINI
1,2-epoxyhexane273.6NINI
Directed evolution—DNA shuffling and site-saturation mutagenesisI219Fstyrene oxide91 (R)17 (R)NINI[77]
Rational design—Site-saturation mutagenesisF108Ip-nitrophenyl glycidyl ether20 (S)3.4 (S)NINI[86]
F108T22 (S)3.4 (S)NINI
Agromyces mediolanus ZJB120203Rational design—Structure-based site saturation and site-directed mutagenesisW182F/S207V/N240Depichlorohydrin90.0 (R)12.9 (R)NINI[87]
Aspergillus niger LCP 521Directed evolution—one round of error-prone PCRA217V/K332E/A390Ephenyl glycidyl ether10.8 (S)4.6 (S)74 (S)56 (S)3933[75]
Semi-rational design —Directed evolution using ISM with combinatorial active site saturation (CASTing)L215F/A217N/R219S/L249Y/T317W/T318V/M329P/L330Y/C350Vphenyl glycidyl ether115 (S)4.6 (S)95 (S)56 (S)4833[88]
Semi-rational design—Directed evolution using ISM and optimalization of expression in recombinant cellsP221S/F244C/L249F/L215F/T317F/T318V(ProThrAlaSerAlaProHisThrTyrArgGluPheIle)-L349V 2phenyl glycidyl ether160 (S)4.6 (S)97 (S)56 (S)4533[89]
Semi-rational design—Directed evolution using all 24 possible pathways using 4 randomization sites for ISML215F/R219V/L249F/T317F/T318C/L349D/C350Yphenyl glycidyl ether158 (S)4 (S)98 (S)61 (S)3028[90]
Aspergillus usamii (AuEH2)Semi-rational design—Microtuning of the substrate-binding pocketA214C/A250Istyrene oxide20216>99 (R)NI50.240[91]
A250Io-nitrostyrene oxide3419698.0 (R)NI50.5NI
A250Wisopropyl glycidyl ether2046.880.2 (S)NI55.2NI
Rhodococcus erythropolis DCL14Directed evolution—ISM using NDT codon degeneracyM32C/I80F/L114C/I116Vcyclopentene oxideNI93 (S,S)13 (R,R)6572[92]
cyclohexene oxideNI97 (S,S)4 (S,S)9484
cycloheptene oxideNI98 (S,S)17 (S,S)7574
cis-2,3-butene oxideNI93 (R,R)5 (S,S)NI
phenyl glycidyl ether32 (R)2.6 (R)92 (R)37 (R)3133
styrene oxide44 (S)2.8 (R)91 (S)40 (R)4336
M32C/L74I/M78F/I80C/V83Icyclopentene oxideNI80 (R,R)13 (R,R)8172
cyclohexene oxideNI90 (R,R)4 (S,S)7484
cycloheptene oxideNI77 (R,R)17 (S,S)8474
cis-2,3-butene oxideNI83 (R,R)5 (S,S)NI
styrene oxide36 (R)2.8 (R)91 (R)40 (R)3036
Directed evolution—ISM using a single-code saturation mutagenesis (SCSM)L74F/M78F/L103V/L114V/I116V/F139V/L147Vcyclohexene oxideNI92 (S,S)4(S,S)>9984[78]
cycloheptene oxideNI94 (S,S)17 (S,S)5297
L74F/M78F/I80V/L114Fcyclohexene oxideNI96 (R,R)4 (S,S)8384
cycloheptene oxideNI94 (R,R)17 (S,S)6697
Directed evolution—ISM using double-code saturation mutagenesis (DCSM)L74F/M78F/I80F/L114V/I116V/F138Vcyclopentene oxideNI85 (S,S)13 (R,R)1384[79]
cyclohexene oxideNI97 (S,S)4 (S,S)98>99
cycloheptene oxideNI97 (S,S)17 (S,S)7397
M78V/I80V/L114Fcyclohexene oxideNI92 (R,R)13 (R,R)99>99
cycloheptene oxideNI85 (R,R)4 (S,S)4097
styrene oxideNI57 (S)21 (R)746
Directed evolution—ISM using triple-code saturation mutagenesis (TCSM)I80Y/L114V/I116Vcyclohexene oxideNI99 (S,S)4 (S,S)97>99[80]
cycloheptene oxideNI98 (S,S)17 (S,S)8197
styrene oxide28.01.892 (S)26 (R)1517
M32V/M78V/I80V/L114Fcyclohexene oxideNI97 (R,R)4 (S,S)>99>99
cycloheptene oxideNI94 (R,R)17 (S,S)8397
Semi-rational design—Directed evolution using ISM with reduced AA alphabets using binary pattern based on choosing hydrophobic and hydrophilic
amino acids		I80F/V83I/L114 V/I116Vcyclopentene oxideNI94 (S,S)7 (R,R)3469[93]
cyclohexene oxideNI97 (S,S)3 (S,S)9387
cycloheptene oxideNI97 (S,S)22 (S,S)9699
I80V/V83I/L114 Vcyclopentene oxideNI51 (R,R)7 (R,R)4869
cyclohexene oxideNI79 (R,R)3 (S,S)9787
cycloheptene oxideNI53 (R,R)22 (S,S)9999
Semi-rational design—Directed evolution using ISM with the aim to improve thermostability, enantioselectivity and activityT76K/L114V/I116V/N92K/F139V/L147F/S15D/A19K/L74F/M78F/E45Dcyclohexene oxideNI94 (S,S)2 (S,S)100100[94]
S15P/M78F/N92K/F139V/T76K/T85K/E45D/I80V/E124Dcyclohexene oxideNI80 (R,R)2 (S,S)100100
Semi-rational design—Directed evolution using saturation mutagenesis, mutants were prepared by high-fidelity solid-phase chemical gene synthesis on silicon chips followed by efficient gene assembly instead of PCR to overcome AA biasM78F/I80Y/L114V/I116Vcyclohexene oxideNI>98 (S,S)NI>98NI[95]
R. erythropolis DCL14 (mutant LEH-P) 3Rational design—Computational design of mutant library using CASCO strategyM32L/L74I/I80V/L103F/F139Lcyclopentene oxideNI85.5 (R,R)23.9 (R,R)NI[96]
M32L/L35W/L74F/M78F/I80A/I116V/F139LNI90.2 (S,S)23.9 (R,R)NI
Rational design—Use of Rosetta enzyme design to computationally predict enantioselective mutants and high-throughput-multiple independent molecular docking simulations for in silico screening of the generated mutant librariesM32A/M78I/I80F/L103I/I116V/F139Lcyclopentene oxideNI85 (S,S)14 (R,R)NI[81]
L35W/L74F/I80G/I116V/F139Lcis-2,3-butane oxideNI82 (S,S)2 (S,S)NI
M32L/L35G/I80W/L103V/F139Lcis-stilbene oxideNI>99 (R,R)92 (R,R)98NI
M32L/L35M/L103I/L114M/I116F/F139LNI88 (S,S)92 (R,R)63NI
Solanum tuberosum (StEH1)Semi-rational design—Directed evolution—ISM targeting AA residues around active site of enzymeW106L/L109Y/V141K/I155V(2,3-epoxypropyl)benzene15 (R)0.4 (R)60 (R)32 (S)NI[97]
Semi-rational design—Directed evolution with 2 rounds of iterative saturation mutagenesisW106L/L109Y/V141K/I155W/F189Cstyrene oxide5800 (S)69 (S)NINI[98]
trans-2-methylstyrene oxide770 (S)84 (S)NINI
Sphingomonas sp. HXN-200Semi-rational design—Site-directed mutagenesis of selected AA residues in active site based on homology modellingV196A/N226A/M332Aphenyl glycidyl ether21.2 (R)2.2 (R)79.2 (S)61.9 (S)5050[99]
metagenomic DNA (Kau2EH)Semi-rational design—Directed evolution by randomizing selected sites within substrate binding pocketV290Yp-chlorostyrene oxide130NI97 (R)NI50NI[100]
EnantioconvergenceA. niger M200Semi-rational design—ISM, mutated sites were chosen on structural similarity with EH from A. niger LCP 521L349V/C350W/T317W/T318V/M218W/R219E/L215M/A217G/M245Astyrene oxide221070.1 (R)3.0 (R)100100[101]
p-chlorostyrene oxide204070.5 (R)4.4 (R)100100
Glycine max (GmEH3)Semi-rational design —Site-saturation and combinatorial mutagenesis used for reshaping substrate-binding pocketW102V/P187F1,2-epoxyhexaneNI83.8 (R)47.2 (R)>99>99[102]
Phaseolus vulgaris (PvEH1)Rational design—Site-directed mutagenesis based on molecular docking simulations and multiple alignmentL105I/M160A/M175Istyrene oxide3.61.587.8 (R)33.6 (R)NI[103]
m-chlorostyrene oxideNI69.7 (R)1.0 (R)NI
p-nitrostyrene oxideNI64.7 (R)50.3 (R)NI
m-nitrostyrene oxideNI52.3 (R)14.7 (R)NI
p-chlorostyrene oxideNI70.9 (R)51.4 (R)NI
Rational design—Leucine scanning used for identification of AA residues at sites lining the enzyme’s binding pocket responsible for enantioconvergence and subsequent saturation mutagenesisL105I/M160A/M175I/Y149L/P184Wm-chlorostyrene oxideNI96.1 (R)1.0 (R)>99>99[104]
Rational design—Reshaping of substrate binding pocketL105I/V106I/M160A/M175I/S178T/P184Wstyrene oxideNI90.3 (R,R)33.6 (R,R)>99.999.1[82]
p-nitrostyrene oxideNI86.7 (R,R)50.3 (R,R)84.299.3
m-nitrostyrene oxideNI85.1 (R,R)14.7 (R,R)>99.999.7
p-fluorostyrene oxideNI90.6 (R,R)13.6 (R,R)>99.998.7
m-chlorostyrene oxide6296.2 (R,R)1.0 (R,R)99.299.9
Rhodotorula paludigena JNU001Rational design—Microtuning substrate-binding pocket of EH by computer-aided design using valine scanning mutagenesisL360Cm-nitrostyrene oxideNI93.4 (R)85.7 (R)99>99[105]
Vigna radiata (VrEH2)Rational design—Creation of smart library by site-directed mutagenesis using reduced AA alphabet to prepare enantioconvergent EHM263Np-nitrostyrene oxideNI98 (R)84 (R)99.5NI[106]
m-nitrostyrene oxideNI90 (R)20 (R)>99>99
Rational design—Creation of smart library by site-directed mutagenesis using reduced AA alphabet to prepare enantioconvergent EHM263Qm-chlorostyrene oxideNI90 (R)20 (R)NI[107]
M263V2-naphthyloxiraneNI9060NI
metagenomic DNA (Kau2EH)Semi-rational design —Directed evolution by randomizing selected sites within substrate binding pocketW110L/F113L/F161Y/P193G/V290Wp-chlorostyrene oxide172393 (R)84 (R)100100[100]
1 No information. 2 Amino acids in parentheses were randomly inserted into the protein sequence during error-prone PCR. 3 Thermostable mutant LEH-P of LEH from Rhodococcus erythropolis DCL14 [108] was used as the template for mutagenesis.
Table
Table 3. Immobilization techniques, materials, and their benefits for EHs.
Table 3. Immobilization techniques, materials, and their benefits for EHs.
Immobilization TechniqueEH (Source)Immobilized BiocatalystSupportBenefit of ImmobilizationRef.
Covalent
bond		ArEH (Agrobacterium radiobacter AD1)Crude enzyme extractLX-1000EP modified by EDA
LX-1000EP		Operational stability, reusability, increased thermal stability as compared to free enzyme[121]
Purified enzymeDextran activated with NaIO4 and ethylene glycol
Ficoll activated with NaIO4 and ethylene glycol
Amylopectin activated with NaIO4 and ethylene glycol
Carboxymethyl cellulose activated with NaIO4 and ethylene glycol		Improved tolerance to the inhibitory effects of Co2+, Fe3+ and EDTA[122]
Kau2EH (metagenomic
DNA)		Purified enzymeEupergit C 250L
Eupergit C
Eupergit C modified by IDA and CuSO4
Sepabeads EC-EP
Sepabeads EC-EP modified by IDA and CuSO4		Significantly higher thermal stability as compared to free enzyme[123]
VrEH2M263N (Vigna radiata)Purified enzymeECR8205F (Epoxy)
ECR4204F (Epoxy)
ECR8215F (Epoxy)
ES-103 (Epoxy)
ESR-1 (Amino)
ESQ-1 (Amino)
ECR8405F (Amino)		Improvement of thermal and operational stability as compared to free enzyme[19]
AnEH (Aspergillus niger LCP 521)Purified
enzyme
(lyophilized powder)		Eupergit C
Eupergit C 250L
Eupergit C 250L modified by EDA
Eupergit C modified by IDA and CuSO4		Improvement of enzyme stability and enantioselectivity[124]
Eupergit C 250L modified by EDA and glutaraldehydeImprovement of enzyme storage and thermal stability and enantioselectivity[125]
Eupergit C modified by EDA and glutaraldehyde; Florisil® silanized with 3-APTES and activated with glutaraldehydeImprovement of enzyme reusability and enantioselectivity[126]
Epoxide-derived silica gelEnhancement of enzyme stability in the presence of DMSO[127]
StEH (Solanum tuberosum)Crude enzyme extract (lyophilized powder)Sepabeads EP—Epoxy modified by IDA and CuSO4
Glyoxyl–agarose (agarose modified by glycidol and oxidized by NaIO4)		Stabilization of enzyme[128]
mEH (rat liver)Purified enzymeSephadex G-150 activated by 1,1′-carbonyldiimidazoleEnhancement of stability and repeated use of the enzyme[129]
Dextran activated by 1,1′-carbonyldiimidazoleIncreasement of enzyme stability[130]
VaEH (Vigna angularis)Partially
purified
enzyme		Mesocellular foam silica (MCF) amino modified and activated by glutaraldehyde; Santa Barbara Amorphous (SBA-15) amino modified and activated by glutaraldehydeEnhancement of enzyme operational stability and thermal stability[131]
Ionic bond/Affinity bond
(His-tag)		StEH (Solanum tuberosum)Crude
enzyme extract		Silica oxide powder modified by resacetophenone and Co2+Observation of enzyme activity in organic solvents[132]
mMcEH (triple mutant) (Mugil cephalus)Purified
enzyme		NiO presenting magnetic nanoparticlesReusability of enzyme[133]
CESH (Nocardia tartaricans CAS-52)Purified
enzyme		Metal ion affinity chromatography media Ni-IDA QZT 6FFEnhancement of enzyme activity[134]
AdsorptionAnEH (Aspergillus niger LCP 521)Purified
enzyme		Accurel EP 100 (polypropylene resin)Enhancement of enzyme operational stability using nonporous DEAE-cellulose[135]
DEAE cellulose (ionic bond)Reusability of enzyme[135,136]
Porous polypropyleneImmobilized for preparative purposes (reuse, continuous reactor)[137]
Lewatit® VP OC 1600Enzyme reusability, enhancement of enantioselectivity[126]
Nsp.EH (Nocardia sp. EH1)Partially
purified
enzyme		DEAE cellulose (ionic bond)Enzyme stabilization[138]
ArEH (Agrobacterium radiobacter AD1 expressed in E. coli)Whole cellsPerliteImmobilized for preparative purposes[139]
McEH (Mugil cephalus)Purified
enzyme		Magnetically separable silica Mag-MSU-F (adsorption) + cross-linking with glutaraldehydeEnhancement of enzyme stability and reusability[140]
CLEAVrEH (Vigna radiata)Partially
purified
enzyme extract		Cross-linker: glutaraldehydeEnhancement of catalytic efficiency, enantioselectivity and product yield[141]
Enhancement of initial reaction rate, product yield, enantioselectivity, operational stability[142]
Co-polymerizationRgEH (Rhodotorula glutinis CIMW 147 (ATCC 201718))Partially purified enzymeAcylation of enzyme by itaconic acid, bio-imprinted with substrate and copolymerized with ethylene glycol dimethacrylateEnzyme stabilization, reusability and product separation, improvement of enantioselectivity[143]
NanoflowersGmEH (Glycine max)Purified enzymeOrganic–inorganic nanoflowers formed with Ca2+ ionsHigh catalytic activity and stability[144]
Metal–organic framework (MOFs)GmEH (Glycine max)Crude enzyme preparation (extract)UiO-66-NH2 metal−organic framework (MOF) cross-linked with glutaraldehydeHigher enzyme pH stability, thermostability and tolerance to organic solvents as compared to free enzyme[145]
HdEH (Hypsibius dujardini)Purified enzymeZeolitic imidazole frameworks (ZIF-8)
Zeolitic imidazole frameworks treated with glutaraldehyde (Glu/ZIF-8)		Enhancement of stability, enantioselectivity, reusability of enzyme[146]
EncapsulationCESH (Nocardia tartaricans ATCC 31191)Whole cellsPolyelectrolyte complex microcapsules from sodium alginate−cellulose sulfate−poly(methylene-co-guanidine)Enhanced enzyme activity, storage stability and decreased reaction time using immobilized whole cells as compared to free cells[116]
Enhancement of operational stability[118]
EntrapmentRtEH (Rhodosporidium toruloides UOFS Y-0471)Whole cellsCalcium alginateStabilization of cells[147]
CESH (Labrys sp. BK-8)Whole cellsκ-carrageenanStabilization of cells[148]
not mentionedNOVO SP409 (Rhodococcus sp. commercial preparation)Crude enzymeNot mentionedPreparative purposes[113]
Table
Table 4. Examples of epoxide hydrolases in the whole-cell enzyme cascades and their role in the cascades.
Table 4. Examples of epoxide hydrolases in the whole-cell enzyme cascades and their role in the cascades.
Enzymes in the Cascade including EH and Enzyme Source/GMO CellsSubstrate(s)Product(s)Note to the Role of EH in the CascadeRef.
Epoxide hydrolase SpEH from Sphingomonas sp. HXN-200 and butanediol dehydrogenase BDHA from Bacillus subtilis BGSC1A1 and NADH oxidase NOX from Lactobacillus brevis DSM 20054/
  • E. coli expressing separately SpEH and E. coli co-expressing BDHA-NOX
  • E. coli co-expressing SpEH-BDHA-NOX
Meso- or racemic epoxidesR-(α)-hydroxyketonesNo significant influence of using separately expressed vs. co-expressed enzymes of the cascade on ee and conversion[155]
Epoxide hydrolase SpEH from Sphingomonas sp. HXN-200 or Epoxide hydrolase StEH from Solanum tuberosum and styrene monooxygenase SMO/
  • E. coli co-expressing SpEH-SMO
  • E. coli co-expressing StEH-SMO
Aryl olefinsChiral vicinal diolsThe first enzyme cascade which enabled reversing enantioselectivity of dihydroxylation using StEH instead of SpEH[156]
Epoxide hydrolase AmEH from Agromyces mediolanus and halohydrin dehalogenase HheC from Agrobacterium radiobacter AD1/
  • E. coli expressing separately HheC and AmEH
  • E. coli co-expressing HheC-AmEH
1,3-dichloro-2-propanolChiral epichlorohydrinEffect of co-expressed vs. separately expressed enzymes on the enantioselectivity of the cascade[157]
Epoxide hydrolase SpEH from Sphingomonas sp. and styrene monooxygenase SMO from Pseudomonas sp./
  • E. coli co-expressing SpEH-SMO
Styrene(S)-1-phenyl-1,2-ethanediolAqueous/organic biphasic reaction system was used for the first time for cascade biotransformation to enhance productivity[158]
Epoxide hydrolase MupZ from Pseudomonas fluorescens NCIMB 10586 and Rieske non-heme oxygenase MupW Pseudomonas fluorescens NCIMB 10586/
  • E. coli expressing MupW
  • E. coli co-expressing MupW-MupZ
MupirocinsHydroxylated tetrahydropyrans and tetrahydrofuransCascade containing epoxide hydrolase and Rieske non-heme oxygenase enabled formation of heterocyclic THP ring, which is difficult to achieve biosynthetically[159]
Epoxide hydrolase SpEH from Sphingomonas sp. HXN-200, alcohol dehydrogenase MnADH from Mycobacterium neoaurum VKM AC-1815D, ω-transaminase PAKω-TA from Pseudomonas aeruginosa and glutamate dehydrogenase GluDH from E. coli BL21/
E. coli BL21 (SGMP) co-expressing 4-enzyme self-sufficient cascade system SpEH-MnADH-PAKω-TA-GluDH 		(S)-epoxidesChiral 1,2-aminoalcoholsThe first one-step synthesis of optically pure 1,2-amino alcohols from (S)-epoxides employing a synthetic redox-self-sufficient enzyme cascade in recombinant cells[160]
Epoxide hydrolase SpEH from Sphingomonas sp. HXN-200, 2,3-butanediol dehydrogenase BDHA from Bacillus subtilis, polyol dehydrogenase GoSCR from Gluconobacter oxydans, (R)-ω-transaminase MVTA from Mycobacterium vanbaalenii/
  • E. coli expressing separatelly SpEH, BDHA, GoSCR, MVTA
  • E. coli co-expressing SpEH-BDHA-GoSCR-MVTA
Racemic epoxidesEnantiopure β-amino alcoholsGeneral access to variety of chiral β-amino alcohols starting from inexpensive racemic epoxides using designed enzyme cascade process in recombinant cells[161]
Styrene monooxygenase SMO from Pseudomonas sp., epoxide hydrolase SpEH from Sphingomonas sp. HXN-200, polyol dehydrogenase GoSCR from Gluconobacter oxydans, (R)-ω-transaminase MVTA from Mycobacterium vanbaalenii or transaminase BMTA from Bacillus megaterium SC6394/
  • E. coli CGS-DEM co-expressing GoSCR-SMO-SpEH-MVTA
  • E. coli CGS-DEB co-expressing GoSCR-SMO-SpEH-BMTA
Styrenyl olefins2-amino-2-phenyl ethanolsChallenging direct regio- and stereoselective aminohydroxylation of olefins to unprotected enantioenriched β-amino alcohols was enabled by novel one-pot four-enzyme biocatalytic cascade in good yields and excellent enantioselectivity[162]
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Bučko, M.; Kaniaková, K.; Hronská, H.; Gemeiner, P.; Rosenberg, M. Epoxide Hydrolases: Multipotential Biocatalysts. Int. J. Mol. Sci. 2023, 24, 7334. https://doi.org/10.3390/ijms24087334

AMA Style

Bučko M, Kaniaková K, Hronská H, Gemeiner P, Rosenberg M. Epoxide Hydrolases: Multipotential Biocatalysts. International Journal of Molecular Sciences. 2023; 24(8):7334. https://doi.org/10.3390/ijms24087334

Chicago/Turabian Style

Bučko, Marek, Katarína Kaniaková, Helena Hronská, Peter Gemeiner, and Michal Rosenberg. 2023. "Epoxide Hydrolases: Multipotential Biocatalysts" International Journal of Molecular Sciences 24, no. 8: 7334. https://doi.org/10.3390/ijms24087334

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