Abstract
The 2-haloacid dehalogenases (EC 3.8.1.X) are industrially important enzymes that catalyze the cleavage of carbon–halogen bonds in 2-haloalkanoic acids, releasing halogen ions and producing corresponding 2-hydroxyl acids. These enzymes are of particular interest in environmental remediation and environmentally friendly synthesis of optically pure chiral compounds due to their ability to degrade a wide range of halogenated compounds with astonishing efficiency for enantiomer resolution. The 2-haloacid dehalogenases have been extensively studied with regard to their biochemical characterization, protein crystal structures, and catalytic mechanisms. This paper comprehensively reviews the source of isolation, classification, protein structures, reaction mechanisms, biochemical properties, and application of 2-haloacid dehalogenases; current trends and avenues for further development have also been included.
Introduction
Halogenated organic compounds show excellent thermal conductivity, insulation, heat resistance, lipophilicity, and biological activity (Kim et al., 2020; Zakary et al., 2021). They are widely used in industrial, agricultural, medical, and military fields as cleaning agents, biocides, gasoline additives, solvents, degreasers, pesticides, and intermediates for chemical synthesis, yielding enormous economic and social benefits (Kurumbang et al., 2014; Zhang et al., 2019; Gul et al., 2020b; Ameen et al., 2021). However, increasing amounts of halogenated compounds are discharged into the environment due to overproduction and extensive use, which results in environmental contamination. These compounds spread in lakes, drinking water, groundwater, seawater, and soil. Unlike naturally occurring halogenated compounds, which can be used as antibiotics to treat bacterial infections, man-made halogenated compounds, which are used as degreasers, solvents, biocides, pharmaceuticals, cleaning agents, and in many other industrial applications, are dangerous when introduced to the environment (Wu et al., 2019; Kirkinci et al., 2021). This is because these compounds do not degrade easily in natural environments because of their chemical stability, resulting in their environmental accumulation. Moreover, these compounds can become concentrated and accumulate in organisms through the food chain, with carcinogenic, teratogenic, and mutagenic effects (Fan et al., 2020; Lou Y. Y. et al., 2021; Zhang C. et al., 2021). This poses a serious threat to human health and has become an issue of concern all over the world (Artabe et al., 2020; Lou Y. Y. et al., 2021).
As the main decomposers in nature, microorganisms convert complex organic compounds into simple compounds, thus maintaining the cycle of elements that are vital to life (Hellal et al., 2021; Kajla et al., 2021; Yu et al., 2021). Microorganisms growing in environments polluted by organic halogenated compounds have the potential to transform these compounds owing to the presence of enzymes that catalyze dehalogenation in their cells, called dehalogenases (Atashgahi et al., 2018; Oyewusi et al., 2020b, 2021b). Among them, 2-haloacid dehalogenases are a family of critical enzymes that hydrolytically catalyze the dehalogenation of 2-haloacids to form corresponding 2-hydroxy acids (Kurihara and Esaki, 2008; Adamu et al., 2020). They cannot only degrade toxic pollutants with low energy consumption but also have a wide substrate profile and high catalytic efficiency. They have highly chiral resolution properties, which may enable the production of optically pure 2-halogenated and 2-hydroxyl compounds (Oyewusi et al., 2020a). Hence, 2-haloacid dehalogenases are highly valuable in the field of environmental remediation and environmentally friendly manufacturing of chiral chemicals. Here, we review the isolation source, classification, molecular structure, catalytic mechanism, catalytic properties, and industrial applications of 2-haloacid dehalogenases. These will enrich the biocatalytic repertoire of haloacid dehalogenases and broaden their applications and developments in the future.
Isolation Sources and Classification of 2-Haloacid Dehalogenases
Microorganisms possessing 2-haloacid dehalogenase are widespread in nature, and have been explored since the beginning of the 20th century. So far, increasing numbers of bacterial and fungal species capable of degrading halogenated xenobiotic pollutants have been isolated (Table 1). Most of these microorganisms were isolated from terrestrial environments, with only a few from marine environments, including Burkholderia sp. I37C (Chiba et al., 2009), Rhodobacteraceae sp. (Novak et al., 2013a), Psychromonas ingrahamii (Novak et al., 2013b), Pseudomonas stutzeri DEH130 (Zhang et al., 2013), Paracoccus sp. DEH99 (Zhang et al., 2014), Lysinibacillus boronitolerans MH2 (Heidarrezaei et al., 2020), and Bacillus megaterium BHS1 (Wahhab et al., 2020). The marine environment is the primary and optimal sink for halogenated pollutants because of their natural release by marine macroalgae, bacteria, sponges, tunicates, corals, worms, phytoplankton, and other invertebrates (Bidleman et al., 2019). Additionally, marine environments are considered extreme owing to a combination of unique properties including high pressure, high salinity, low temperature, oligotrophy, and special lighting conditions (de Oliveira et al., 2020; Ameen et al., 2021; Zhang J. et al., 2021). Because of this, microorganisms living in this environment are diverse and specific in gene composition and ecological functions; the intracellular enzymes of these microorganisms are correspondingly diverse and specific, conferring physiological and biochemical characteristics such as barophilicity, salt tolerance, cold adaptability, hyperthermostability, chemoselectivity, stereoselectivity, and regioselectivity (Thippeswamy et al., 2021; Zhang J. et al., 2021). The marine environment is therefore expected to be an important source of novel enzymes.
Table 1
| Microorganisms | Genus | References |
|---|---|---|
| Bacteria | Agrobacterium | Köhler et al., 1998 |
| Alcaligenes | Hill et al., 1999 | |
| Ancylobacter | Kumar et al., 2016 | |
| Arthrobacter | Bagherbaigi et al., 2013 | |
| Azotobacter | Diez et al., 1996 | |
| Bacillus | Horisaki et al., 2011; Ratnaningsih and Idris, 2018; Oyewusi et al., 2021a | |
| Burkholderia | Edbeib et al., 2020 | |
| Klebsiella | Idris Ratnaningsih, 2015 | |
| Lysinibacillus | Heidarrezaei et al., 2020 | |
| Methylobacterium | Kurihara and Esaki, 2008 | |
| Mesorhizobium | Zakary et al., 2021 | |
| Moraxella | Kurihara et al., 2000 | |
| Paracoccidioides | Satpathy et al., 2015 | |
| Paracoccus | Zhang et al., 2014 | |
| Pseudoalteromonas | Liao et al., 2015 | |
| Pseudomonas | Hasan et al., 1994; Park et al., 2003; Schmidberger et al., 2008; Zhang et al., 2013 | |
| Psychromonas | Novak et al., 2013b | |
| Pyrococcus | Arai et al., 2006 | |
| Rhizobium | Adamu et al., 2016; Oyewusi et al., 2020b | |
| Rhodobacteraceae | Novak et al., 2013a | |
| Serratia | Rosland Abel et al., 2012 | |
| Sulfolobus | Xu et al., 2004 | |
| Xanthobacter | van der Ploeg et al., 1991 | |
| Fungi | Beauveria | Satpathy et al., 2016 |
| Botrytis | Bustillo et al., 2003 | |
| Candida | Polnisch et al., 1991 | |
| Dichomitus | Muzikár et al., 2011 | |
| Fusarium | Li et al., 2011 | |
| Metarhizium | Satpathy et al., 2016 | |
| Phanerochaete | Wang et al., 2009 | |
| Pycnoporus | Muzikár et al., 2011 | |
| Trichoderma | Bagherbaigi et al., 2013 |
The reported microorganisms degrading 2-haloalkanoic acids.
The 2-haloacid dehalogenases have been classified according to amino acid sequence conservation and substrate selectivity (Wang et al., 2018; Adamu et al., 2020). These enzymes are classified into four types according to their substrate specificities and product configurations: D-2-haloacid dehalogenase (D-DEX, EC 3.8.1.9), L-2-haloacid dehalogenase (L-DEX, EC 3.8.1.2), configuration-inverting DL-2-haloacid dehalogenase (DL-DEXi, EC 3.8.1.10), and configuration-retaining DL-2-haloacid dehalogenase (DL-DEXr, EC 3.8.1.11) (Zakary et al., 2021). D-DEX catalyzes the dehalogenation of D-2-haloalkanoic acids, whereas L-DEX specifically acts on L-2-haloalkanoic acids. DL-DEXi and DL-DEXr act on both enantiomers of substrates, but yield different product configurations. The 2-haloacid dehalogenases in general are divided into Group I and Group II enzymes according to the amino acid sequence homology; D-DEX and DL-DEX belong to Group I and L-DEX to Group II.
Structural and Catalytic Characteristics of 2-Haloacid Dehalogenases
The structural diversity of 2-haloacid dehalogenases determines their diversity of function. The different types of 2-haloacid dehalogenases have different structures and catalytic mechanisms; an overview of this is provided in this section.
L-DEX
Structural Characteristics and Catalytic Mechanism
L-DEX specifically acts on L-2-haloalkanoic acids to produce D-2-hydroxyalkanoic acids. These enzymes are widespread in nature and their biochemical characteristics and structures have been studied extensively (Satpathy et al., 2016; Wang et al., 2016; Adamu et al., 2020). So far, the three-dimensional (3D) structures of specific L-DEXs and their substrate complexes have been analyzed, including L-DEX YL from Pseudomonas sp. strain YL (Hisano et al., 1996), DhlB from Xanthobacter autotrophicus GJ10 (Ridder et al., 1997), PH0459 from Pyrococcus horikoshii OT3 (Arai et al., 2006), DehIVa from Burkholderia cepacia MBA4 (Schmidberger et al., 2007), DehSft from Sulfolobus tokodaii (Rye et al., 2009) and DehRhb from Rhodobacteraceae (Novak et al., 2013a).
L-DEX is an α/β type hydrolase consisting of a typical Rossman-fold-like core domain and subdomain, with the active site located between the two domains (Figure 1), apart from DhlB, which is composed of a core domain and two subdomains. Most L-DEX molecules are dimers consisting of two identical subunits, except for PH0459, which is a monomer according to its crystal structure (Arai et al., 2006). In a typical L-DEX structure, six-stranded parallel β-sheets (in order: β5-β4-β1-β6-β7-β8) are flanked on both sides by five α-helices, forming three layers of α/β fold units together constituting a sandwich domain (Hisano et al., 1996; Poelarends and Whitman, 2010; Zhang et al., 2018). Although the core domain of L-DEX has an α/β-type structure, it does not belong to the α/β hydrolase fold family, in which the typical domains are eight-stranded β-strands (in order: β1-β2-β4-β3-β5-β6-β7-β8) with the β2-strand antiparallel to the others. Two β-strands are separated by α-helix from the third strand, forming a β/α/β unit. The first α-helix and the last α-helix are located at one side of the β-sheet, and the remaining α-helices are at the other side (Janssen, 2004; Kunka et al., 2018; Babkova et al., 2020; Mazur et al., 2021).
Figure 1
L-DEX structures. (A) Structural superposition of DhlB (hotpink, PDB ID: 1qq5) and L-DEX YL (green, PDB ID: 1jud). (B) Structural superposition of DhlB (hotpink, PDB ID: 1qq5), L-DEX YL (green, PDB ID: 1jud), PH0459 (cyan, PDB ID: 1x42), DehIVa (magenta, PDB ID: 2no4), DehSft (yellow, PDB ID: 2w11) and DehRhb (salmon, PDB ID: 2yml).
The dehalogenation is catalyzed by L-DEX in an SN2 nucleophilic substitution reaction as confirmed by X-ray Crystallography, O18 isotope labeling, liquid chromatography–mass spectrometry (LC–MS), site-directed mutagenesis, and quantum mechanic/molecular mechanic (QM/MM) calculations (Adamu et al., 2017a,b). The carboxylic acid group of the aspartic acid residue acts as the nucleophile in the active center, which attacks the C2 atom of the L-2-haloalkanoic acid to form an ester intermediate (Schmidberger et al., 2007). This intermediate product is then hydrolyzed by a water molecule, activated by His/Glu (in DehRhb) or Asn/Ser (in DehIVa) or Lys (in L-DEX YL) (Figure 2). The resultant halide ions are stabilized with the assistance of Arg or Asn or Phe. Greater numbers of halide ion acceptors can cleave stronger C-X bonds (Kurihara, 2011; Kondo et al., 2014).
Figure 2
Reaction mechanism of L-DEX (Schmidberger et al., 2007).
Biochemical Properties
L-DEXs have been isolated from both terrestrial and marine environments. Some biochemical characteristics are shared between enzymes, and some differ. For example, L-DEX exhibits high catalytic activity on chlorinated and brominated substrates, but no such activity on D-2-haloalkanoic acids. Additionally, this enzyme cannot catalyze the dehalogenation of fluorinated and C3-substituted haloalkanoic acids. With the exception of L-DEX YL, L-DEXs only show high catalytic activity on haloalkanoic acids of two or three carbons in length, with low or no activity on haloalkanoic acids four or more carbons in length (van der Ploeg et al., 1991; Liu et al., 1994; Zhang et al., 2013, 2014).
L-DEX enzymes differ in substrate specificity; L-DEX YL is more specific to L-2-chloropropionic acid than chloroacetic acid, whereas the L-DEX from Bacillus strain I37C is more specific to chloroacetic acid than to 2-chloropropionic acid (Liu et al., 1994; Chiba et al., 2009). The optimal pH range for L-DEX reactions is 9–11 (alkaline). Subunit molecular weights range from 25 to 28 kDa. Natural L-DEXs exist as monomers, dimers, and tetramers (van der Ploeg et al., 1991; Liu et al., 1994; Zhang et al., 2013, 2014).
L-DEXs isolated from different bacterial species have different thermal stability: the optimum reaction temperature for L-DEX from the terrestrial Pseudomonas putida is 30°C−45°C, and it loses 50% activity after 15 min incubation at 55°C. Psychromonas. ingrahamii is isolated from the sea-ice interface (−10°C) and exhibits psychrophilic properties; the lowest temperature at which this strain is able to grow is −12°C. L-DEX Pin, from P. ingrahamii, has an optimum reaction temperature of 45°C, with a melting temperature of 85°C. L-DEX Pin possesses the characteristics of both psychrophilic and thermophilic enzymes. Structurally, compared with mesophilic enzymes, L-DEX Pin has more hydrophobic surfaces and more salt bridges (Novak et al., 2013b).
The optimum reaction temperature for DehRhb, isolated from marine Rhodobacteraceae, is 55°C. The activity of this enzyme remains at ~45% after incubation for 1 h at 60°C, indicating moderate thermal stability. Its key catalytic residues are His183 and Glu21, which are different from L-DEXs from terrestrial environments, suggesting that it may catalyze the dehalogenation with a novel catalytic mechanism (Novak and Littlechild, 2013). In summary, natural dehalogenases with novel properties may be more likely to be isolated from marine and other extreme environments; a greater understanding of their structures, catalytic mechanism and catalytic properties may provide theoretical guidance for determining the direct evolution of L-DEXs and other dehalogenases.
DL-DEX
Structural Characteristics and Catalytic Mechanism
DL-DEX enzymes, which include DL-DEXi and DL-DEXr, catalyze the hydrolytic dehalogenation of both enantiomers of 2-haloalkanoic acids to produce corresponding 2-hydroxyalkanoic acids.
For DL-DEXi, the configuration of the product is opposite to the substrate: the C2 atom of the substrate configuration is inverted during dehalogenation catalyzed by DL-DEXi. Six DL-DEXi enzymes have been reported so far, including DL-DEX YL from Pseudomonas putida YL (Hasan et al., 1994; Soda et al., 1996), DL-DEX 113 from Pseudomonas sp. 113 (Nardi-Dei et al., 1999; Park et al., 2003), DehI from Pseudomonas putida PP3 (Park et al., 2003; Schmidberger et al., 2008), DL-DEX Mb from Methylobacterium sp. CPA1 (Siwek et al., 2013), DehE from Rhizobium sp. RC1 (Hamid et al., 2011; Zainal Abidin et al., 2019), and DhIIV from Alcaligenes xylosoxidans ABIV (Brokamp et al., 1996; Hamid et al., 2011). The crystal structures of DehI and DL-DEX Mb have been studied, revealing that DL-DEXi is an α-helical hydrolase, with no structural homology to L-DEX and other fold superfamilies in the hydrolases (Schmidberger et al., 2008; Siwek et al., 2013).
As shown in Figure 3, DehI is a homodimer according to its crystallographic structure. The N-terminus (amino acid residues 1–130) and C-terminus (residues 166–296) share 16% sequence identity in monomers, which form a pseudo-dimer. The active site is located at the interface of the pseudo-dimer, which binds D- and L- substrates (Schmidberger et al., 2008). The catalytic mechanism of DL-DEXi is different to that of L-DEX: dehalogenation catalyzed by D-DEXi is directly mediated by an activated water molecule, without involving the formation of E-S ester intermediate (Figure 4) (Nardi-Dei et al., 1999). The nucleophilic water molecule is likely activated by the conserved Asp and Asn residues; however, there is no relevant experimental evidence for this.
Figure 3
Structural superposition of DehI (green, PDB ID: 3bjx) and DL-DEX Mb (cyan, PDB ID: 4n2x). The active pocket is shown as surface.
Figure 4
Reaction mechanism of DL-DEXi (Nardi-Dei et al., 1999).
The transformation of the C2-configuration of the substrate catalyzed by DL-DEXr is opposite to that of DL-DEXi. DL-DEXr catalyzes dehalogenation with retention of the C2-configuration of the substrate. Therefore, the substrate and product share the same configuration. DL-DEXr has so far only been reported in P. putida PP3 (Weightman et al., 1982; Park et al., 2003). Gene sequence information for this enzyme is still unknown, and the reaction mechanism has not been analyzed. It is proposed that dehalogenation involves a cysteine residue, as DL-DEXr is highly sensitive to sulfhydryl reagents such as N-ethylmaleimide and p-chloromercuribenzoic acid. The reaction is thought to proceed with double inversion of the C2-configuration of the substrate, resulting in the retention of the C2-configuration: the first C2-configuration inversion releases halogen ions and forms an E-S thioester intermediate; then, the intermediate is hydrolyzed under the attack of a water molecule, and the C2-configuration is reversed again. However, there is currently no direct experimental data to confirm this hypothesis (Figure 5) (Weightman et al., 1982).
Figure 5
Possible mechanism of DL-DEXr involving the retention of C2-configuration of the substrate (Weightman et al., 1982).
Biochemical Properties
In DL-DEXs, only DehI, DehE, DL-DEX 113 and DL-DEX ABIV have been characterized in terms of their enzymatic properties (Brokamp et al., 1996; Schmidberger et al., 2008). These enzymes have a greater specificity for L-2-haloalkanoic acids than D-2-haloalkanoic acids (Table 2). DL-DEXi can catalyze the dehalogenation of haloalkanoic acids with a carbon chain length of two to four, and catalyzes the formation of oxalate from trichloroacetate (Soda et al., 1996). Most DL-DEXi enzymes are homodimers, except for DL-DEX YL, which is a monomer. The subunit molecular weight ranges from 26 to 36 kDa (Kondo et al., 2014). DL-DEXi maximum activity levels occur at a pH of ~9.5. The optimum reaction temperature is between 30 and 40°C (Leigh et al., 1986; Park et al., 2003; Hamid et al., 2011). DL-DEXr is sensitive to SH-reagents; like DL-DEXi, it degrades haloalkanoic acids with a chain length of 2–4 (Weightman et al., 1982).
Table 2
| Enzymes | Strains | L/Da | References |
|---|---|---|---|
| DehI | P. putida PP3 | 1.2 | Park et al., 2003 |
| DehE | R. sp. RC1 | 1.6 | Hamid et al., 2011 |
| DL-DEX 113 | P. sp. 113 | 1.4 | Park et al., 2003 |
| DhIIV | A. xylosoxidans ABIV | 1.1 | Brokamp et al., 1996 |
| DL-DEX YL | P. putida YL | – | |
| DL-DEX Mb | M. sp. CPA1 | – |
Enatioselectivity of DL-DEXis from different strains.
L/D, the ratio of catalytic activity on L-2-chloropropionic acid and D-2-chloropropionic acid; -, no experimental data is available.
D-DEX
Structural Characteristics and Catalytic Mechanism
D-DEXs specifically catalyze the hydrolytic dehalogenation of D-2-haloalkanoic acids to produce L-2-hydroxyacids. So far, only four kinds of primary structure information are available for D-DEX, including DehD from Rhizobium sp. RC1 (Sudi et al., 2014), DehII from Agrobacterium sp. NHG3 (Higgins et al., 2005), HadD AJ1 from Pseudomonas putida AJ1 (Smith et al., 1990) and DehDIV-R from Pseudomonas sp. ZJU26 (Wang Y. et al., 2020). HadD AJ1 and DehDIV-R share the highest sequence homology (89%); HadD AJ1 and DehII NHG3 share 22.2% sequence homology, and HadD AJ1 and DehD share 32.6% sequence homology.
The author has extensively studied on the structure and catalytic mechanism of HadD AJ1. The crystal structure of HadD AJ1 is highly similar to that of DL-DEXi. Both types of enzymes are α-helical proteins, different from the α/β fold structure. HadD AJ1 is a homotetramer according to its crystallographic structure; each monomer comprises two repeats with 20% sequence identity (Figure 6). The two repeated folds are composed of N-terminal α-helices 1–6 and C-terminal α-helices 7–12, respectively, with a linker section containing 33 amino acids and a 310-helix η1 (Figure 6A). These two repeats are stabilized by van der Waals forces, salt bonds, hydrogen bonds, and hydrophobic interactions. As shown in Figure 6B, helix α4 and α10 are arranged in parallel with each other, and α6 and α12 cross each other at the bulge between them. Helices α6 and α12 mutually interlace at their bulges, located in the middle of the helices (Wang et al., 2018). This has been reported in many proteins with internal structural repeats, which are considered to result from genetic processes such as fusion and fission of domains and gene duplication during protein evolution (Longo et al., 2014; Berezovsky et al., 2017; Vrancken et al., 2020).
Figure 6
HadD AJ1 monomer (Wang et al., 2018). (A) Ribbon representation of monomeric HadD AJ1 composed of two repeats (repeat 1: red; repeat 2: yellow) and a linker (cyan). (B) 3D superposition of repeat 1 (red) and repeat 2 (yellow).
In HadD AJ1, Asp205 is the key catalytic residue, activating the water molecule with the assistance of Asn131. This was identified through an analysis of the complex structure of wildtype (WT) enzyme binding the product L-lactic acid (L-LA) and a D205N mutant binding the substrate D-2-chloropropionate (D-2-CPA) (Figure 7A). The dehalogenation catalyzed by D-DEX is directly mediated by activated water molecules, without forming an ester intermediate in the reaction process; this is the same process as DL-DEXi (Figure 7B). The activated water molecule attacks the C2 atom of the substrate from the opposite side of the halogen atom, breaking the C-X bond (Figure 7A). The halogen ion is released toward F281, and simultaneously, the hydroxyl group of the activated water molecule is bonded to the C2 atom of the substrate to form L-lactic acid (Wang et al., 2018).
Figure 7
The molecular mechanism of dehalogenation catalyzed by D-DEX. (A) Structural superimposition of WT/L-LA (green, PDB ID: 5gzy) and D205N/D-2-CPA (cyan, PDB ID: 5gzx) complex. (B) Reaction mechanism of D-DEX (Wang et al., 2018).
D-DEX and DL-DEXi share high amino acid sequence as well as structural homology. Moreover, both types of enzymes catalyze dehalogenation by the same mechanism, directly mediated by the nucleophilic water molecule; this differs from dehalogenation catalyzed by L-DEX, which is mediated by E-S ester intermediates. This suggests an evolutionarily close relationship between D-DEX and DL-DEXi.
Biochemical Properties
Currently, there are only a few studies on D-DEX enzymes, likely a result of the lack of microorganisms known to produce D-DEX. From analyses of DehD and HadD AJ1 biochemical properties, D-DEXs specifically catalyze dehalogenation of D-2-chlorinated and D-2-brominated acids with carbon chain lengths of 2–4. However, D-DEX has a higher catalytic activity on brominated than chlorinated substrates (Smith et al., 1990; Huyop and Sudi, 2012). Km values of DehD, HadD AJ1, and DehDIV-R are 0.06, 0.94, and 2.2 mmol/L, respectively, with D-2-CPA as the substrate (Smith et al., 1990; Huyop and Sudi, 2012; Wang Y. et al., 2020). Compared with HadD AJ1 and DehDIV-R, DehD has a stronger affinity for D-2-CPA.
The natural active states of D-DEXs are different: DehD exists is a homodimer, while HadD AJ1 is a homotetramer. The optimal reaction pH of D-DEXs ranges from 9.0 to 10.0. The enzyme activity decreases rapidly when the pH falls outside the range of 8.0–10.0; under these conditions, HadD AJ1 exhibits <50% catalytic activity (Smith et al., 1990). In comparison with L-DEXs, D-DEXs are mesophilic, with an optimal reaction temperature of 50°C−60°C; however, the enzyme molecules are relatively stable between 30 and 40°C, but rapidly lose activity in a reaction temperature higher than 40°C (Smith et al., 1990).
Application
The 2-haloacid dehalogenases can detoxify halogenated pollutants by hydrolysis without the addition of other reductive agents; for this reason, their potential application in bioremediation is particularly attractive (Behbahani et al., 2018; Oyewusi et al., 2020b, 2021b; Zakary et al., 2021). The 2-haloacid dehalogenases are also highly stereoselective, and they may therefore be valuable in fine chemistry synthesis applications (Chen and Ribeiro de Souza, 2019; Adamu et al., 2020; Wang S. et al., 2020). These enzymes can be used to obtain chiral hydroxy acids and haloalkanoic acids with low molecular weights; these small organic acids generally act as intermediates for synthesizing agrochemicals, medicines, and other important chemicals (Leemans Martin et al., 2020; Gurushankara, 2021). Hence, 2-haloacid dehalogenases are promising and potentially highly valuable for their application in environmental remediation and chemical synthesis (Bommarius, 2015; Tanokura et al., 2015; Zhang et al., 2018); here, we discuss the main fields in which they could be applied.
Environmental Bioremediation
Halogenated carboxylic acids such as 2-chloropropionic acids and 2,2-chloropropionic acids are widely used as an intermediate in the synthesis of pesticides and pharmaceuticals, especially the chirally pure 2-chloropropionic acid precursors for synthesizing many chiral drugs (Nguyen et al., 2021; Zhou et al., 2021). However, these haloacids produce chlorinated organic contaminants owing to extensive use and improper disposal. Haloacids are also intermediates in the degradation of some halogenated compounds, such as 1,2-dichloroalkane and hexachlorocyclohexane, which results in more haloacid contaminants in the environment (Hermon et al., 2018). The accumulation of these pollutants causes serious environmental problems and threats to human and other organisms' health. The 2-haloacid dehalogenase can catalyze the dehalogenation of 2-chloropropionic acids and 2,2-dichloropropionic acids to form non-toxic hydroxyl acids, which is a very promising potential tool for environmental bioremediation (Oyewusi et al., 2021b; Zakary et al., 2021). Dioxin compounds are carcinogenic byproducts originating from natural and anthropogenic sources such as herbicides, pesticides, and combustion processes; high levels of dioxin-contamination have been reported in food, soils, and blood samples of local residents in Southern Vietnam (Nguyen et al., 2021). Burkholderia cenocepacia strain 869T2 can degrade 0.2 mg L−1 of dioxin within 1 week under aerobic conditions, in which L-2-haloacid dehalogenase plays a crucial role (Nguyen et al., 2021). Haloacetic acids are the second most prominent class of disinfection by-products, and are frequently detected in surface and drinking water systems. These compounds have genotoxic, mutagenic, cytotoxic, and tumorigenic effects in humans (Kim et al., 2020; Long et al., 2021; Lou J. et al., 2021). In metabolically engineered Burkholderia species, the degradation activity of haloacetic acids can be increased by 4–8 times (Su et al., 2013). The bacterial degradation of haloacetic acids was found to be affected by water distribution system conditions, including pH, phosphate, total organic carbon and residual chlorine (Behbahani et al., 2018). The order of mean haloacetic acid degradation rates has been found to be di >mono >tri-halogenated acetic acids.
Phytoremediation has been attracting more attention as an environmentally friendly technology to clean up environmental contamination (Kurade et al., 2021); transgenic tobacco that produces haloalkane dehalogenase and haloacid dehalogenase, and which therefore contains a complete degradation pathway, has been reported to degrade 1,2-dichloroethane (Mena-Benitez et al., 2008).
Fine Chemical Synthesis
The growing interest in the use of 2-haloacid dehalogenases in fine chemical synthesis is due to their chiral selectivity. Optically pure compounds are generally synthesized using chemical methods; however, this is unpopular owing to the involvement of toxic reagents, as well as the low yield and low optical purity of products (Santi et al., 2021). Biocatalysis is considered as a more environmentally friendly and effective method because of the mild reaction conditions, and remarkable enantioselectivity (Novak et al., 2013b; Schober and Faber, 2013; Wang S. et al., 2020).
L-2-chloropropionic acid is an important precursor in the synthesis of herbicides and pesticides (Zhou et al., 2021). D-DEX specifically hydrolyzes D-2-chloropropionic acid in racemic 2-chloropropionic acid; L-2-chloropropane acid is therefore obtained with high enantiomeric purity by separation (Gong et al., 2018). To obtain optically active L-2-chloropropionic acid, Imperial Chemical Industries has already applied HadD AJ1 to the resolution of racemic 2-chloropropionic acid in an industrial setting, which has been the primary method for producing chiral chloropropionic acid (Taylor Stephen, 1985; Parker and Colby, 1995). It has also been used by AstraZeneca in the resolution of rac-2-CPA by D-DEX. This method is also suitable for the production of other short-chain chiral 2-halogenated acids, and the scale can be higher than 1,000 tons/year (Schober and Faber, 2013).
D-2-CPA is an important raw material for chemical synthesis that can be directly used to produce a variety of pharmaceutical intermediates, such as the nutritional medicine L-alanyl-L-glutamine and the anti-tuberculosis drug thiolactomycin. L-DEX can be used for the resolution of racemic 2-chloropropionic acids to obtain D-2-CPA with enantiomeric purity (Breuer et al., 2004).
Optically pure lactic acid is an important chiral intermediate in the synthesis of agrochemical, pharmaceutical, and chemical industries; it has been reported that L-lactic acid can be used to synthesize nanoparticles and nanofibers, which act as drug carriers (Chuan et al., 2020; Liu et al., 2021; Ma et al., 2021; Yavari Maroufi et al., 2021). D-lactic acid is also involved in the synthesis of important chiral drug intermediates, such as methyl D-lactate (Sengupta et al., 2020). Xie and colleagues studied the reaction conditions of L-DEX from thermophilic archaea Sulfolobus tokodaii in the catalytic conversion of racemic 2-chloropropionic acid to D-lactic acid (Rye et al., 2009; Xie et al., 2015); after optimizing reaction conditions with regard to substrate, buffer, and enzyme concentration, preparation of D-lactic acid was found to work best with 0.5 mol/L 2-chloropropionic acid.
D-2-bromobutyric acid is used as an intermediate for the synthesis of pharmaceuticals and agrochemicals. The fluoroacetate dehalogenase mutant H155V/W156R/Y219M is reported to catalyze the kinetic resolution of rac-2-bromobutyric acid, producing D-2-bromobutyric acid with an enantiomeric excess of 99.7% (Wang S. et al., 2020).
Agricultural Production
Herbicides with broad spectrum can effectively remove a variety of weeds, such as monochloroacetic acid, 2-chloropropionic acid, and 2-dichloropropionic acid. However, these herbicides can also damage economically valuable crops, resulting in significant losses in agricultural production. These losses can be avoided by developing herbicide-resistant crops, which requires the introduction of genes encoding dehalogenases into these crops. The dehD gene from Rhizobium sp. RCI, encoding D-2-haloacid dehalogenase, has been successfully introduced into tobacco as selective tag, constructing a transgenic variant of Nicotiana benthamiana with anti-monochloroacetic acid activity (Mohamed et al., 2020). This transgenic, herbicide-resistant tobacco is confirmed to be effective at various development stages, including seed germination and mature leaf stages. The dehalogenase gene is therefore likely to play an important role as a dominant, selectable marker gene for the construction of other crop species resistant to broad-spectrum halogenated compound herbicides.
Other Fields
Dehalogenases can also be used to construct biosensors for in situ detection of organic halogenated pollutants in the environment (Artabe et al., 2020; Gul et al., 2020a,b). By immobilizing halohydrin dehalogenase on a glass fiber membrane, detection limits of 0.06 mmol/L 1,3-dichloro-2-propanol and 0.09 mmol/L 2,3-dibromo-1-propanol have been achieved (Gul et al., 2020b). A detection limit of 1 mg/L dichlorethane has been achieved by immobilizing haloalkane dehalogenase on stacked chitosan films (Shahar et al., 2019a,b).
Dehalogenases act as tags when genetically fused to a protein of interest, termed HaloTag technology (England et al., 2015; Döbber and Pohl, 2017; Erdmann et al., 2019). This technology overcomes the current limitations of traditional protein tagging platforms, as it can be applied to protein isolation and purification, studies of protein synthesis and degradation, analyses of protein function, studies of protein–protein and protein–DNA interactions, and molecular and cellular imaging (Encell et al., 2012; Merrill et al., 2019; Cattoglio et al., 2020; Freitas et al., 2021; Minner-Meinen et al., 2021). Furthermore, novel technologies have been developed for tumor diagnosis and treatment involving the linkage of dehalogenase fused with cancer cell recognition peptides to multifunctional nanoparticles (Garbujo et al., 2020).
Discussion and Prospects
A variety of 2-haloacid dehalogenases have so far been isolated and identified. Although structural information and catalytic mechanisms for L-DEX, DL-DEXi, and D-DEX have generally been well-understood, very little information on the structure and catalytic mechanism of DL-DEXr is available. Therefore, further study is necessary to understand DL-DEXr.
Enzyme stereoselectivity has been attracting a great deal of attention for asymmetric synthesis and chiral resolution. The 2-haloacid dehalogenases show typical stereoselectivity; however, little is known about the stereoselective mechanism. The enantioselective mechanism of L-DEX has been studied using quantum mechanics/molecular mechanics (QM/MM) and fragment molecular orbital calculation (Kondo et al., 2015; Adamu et al., 2019), which have confirmed that the high activation energy barrier prevents this enzyme from acting on the D-substrate. However, it is still unclear how selectivity of enzymes on chiral substrates is regulated. The stereoselective mechanism of D-DEX has been studied, and enzymatic stereoselectivity was found to be controlled by the residue Leu288, which determines the entry of L-substrate into the active site of the enzyme with steric hindrance. The mutation of residue leucine to isoleucine enables the enzyme to catalyze the dehaologenation of the L-substrate, owing to the different rotation position of Ile288 compared with Leu288. In the mutant enzyme, Ile288 functions as a wing gatekeeper, interacting with the substrate by gate-flipping during dehalogenation, allowing the L-substrate to enter the active site. However, it is still unclear how DL-DEXr and DL-DEXi recognize and interact with chiral substrates. Stereoselective properties make biocatalysts valuable in the preparation of optically pure compounds, which is an important area of environmentally friendly chemistry. An ideal industrial biocatalyst should have both high catalytic activity and specific stereoselectivity; exploring the molecular regulatory mechanisms underlying these properties forms the basis of artificial customization of dehalogenases with these properties. Reactions can be controlled using direct regulation of enzyme selectivity, forming products with high optical purity and unique structures. Further study on the stereoselectivity of 2-haloacid dehalogenase is therefore required in order to successfully manipulate this property.
Most 2-haloacid dehalogenases have a high catalytic activity with short-chain halogenated acid substrates containing fewer than four carbon atoms, while they show weak or no catalytic activity for longer-chain halogenated acids. Additionally, the low tolerance of these enzymes to organic solvents limits the range of their substrate profile. In order to obtain enantiomerically pure chiral products, enzymatic catalysis is sometimes used in enantiomeric resolution by combining with chemical convergence (Clayton et al., 2020). However, the conditions of the enzymatic reaction are incompatible with the high temperature and extreme pH required for chemical hydrolysis in the downstream separation process. Therefore, it remains necessary to identify novel 2-haloacid dehalogenases with unique properties, allowing them to function in these more extreme conditions (Marshall et al., 2021). Marine microorganisms may be the primary source of novel enzymes with extraordinary properties owing to their previously established genetic and biochemical diversity.
The birth of protein engineering technology has opened up a new route for researchers to develop excellent biocatalysts by redesigning natural enzymes (Marshall et al., 2021; Watanabe et al., 2021; Xiong et al., 2021). Many enzyme engineering design strategies have emerged, such as directed evolution, rational, semi-rational, de novo, computer-assisted, and artificial intelligence (Bunzel et al., 2021; Narayanan et al., 2021; Tunyasuvunakool et al., 2021; Woolfson, 2021; Wu et al., 2021). These strategies have been used to improve enzyme stability, activity, and selectivity for substrates. However, so far, only L-2-haloacid dehalogenases have been engineered to alter their substrate specificity. The mutation of residue Ser188 to Val in the enzyme DehE enables it to act on 3-chloropropionic acid (Hamid et al., 2015). Recent developments in understanding the structural and catalytic properties of 2-haloacid dehalogenases will also likely enable these enzymes to be more easily modified for commercial uses alongside L-2-haloacid dehalogenases. Given this overall direction of research, an increasing variety of 2-haloacid dehalogenases will likely be modified through protein engineering techniques to improve their properties for biotechnological applications.
Funding
This work was supported by the National Natural Science Foundation of China (22078308), Innovation Leadership Program in Sciences and Technologies for Central Plains Talent Plan (214200510009), Key Scientific Research Projects in the Universities of Henan Province (20A180024), Research Project of Shangqiu Normal University (700144), Postdoctoral Science Foundation of Henan Province (50026003), Program for Science and Technology Innovative Research Team in University of Henan Province (21IRTSTHN025), and Innovation Leadership Program in Sciences and Technologies for Zhengzhou Talent Gathering Plan, Henan Academician Workstation for Industrial Technology of Dry Chilli.
Publisher's Note
All claims expressed in this article are solely those of the authors and do not necessarily represent those of their affiliated organizations, or those of the publisher, the editors and the reviewers. Any product that may be evaluated in this article, or claim that may be made by its manufacturer, is not guaranteed or endorsed by the publisher.
Statements
Author contributions
YW and QX handled the literature collection and literature research. YW prepared the original draft. QZ, JX, and DP modified the manuscript. All authors critically reviewed, contributed to, and approved the final manuscript.
Acknowledgments
We would like to thank the reviewers for their constructive comments.
Conflict of interest
JX is an employee of Zhengzhou Tuoyang Industrial Co., Ltd. The remaining authors declare that the research was conducted in the absence of any commercial or financial relationships that could be construed as a potential conflict of interest.
References
1
AdamuA.Abdul WahabR.AliyuF.Abdul RazakF. I.MiendaB. S.ShamsirM. S.et al. (2019). Theoretical analyses on enantiospecificity of L-2-haloacid dehalogenase (DehL) from Rhizobium sp. RC1 towards 2-chloropropionic acid. J. Mol. Graph Model.92, 131–139. 10.1016/j.jmgm.2019.07.012
2
AdamuA.ShamsirM. S.WahabR. A.ParvizpourS.HuyopF. (2017a). Multi-template homology-based structural model of L-2-haloacid dehalogenase (DehL) from Rhizobium sp. RC1. J. Biomol. Struct. Dyn.35, 3285–3296. 10.1080/07391102.2016.1254115
3
AdamuA.WahabR. A.AliyuF.AminuA. H.HamzaM. M.HuyopF. (2020). Haloacid dehalogenases of Rhizobium sp. and related enzymes: Catalytic properties and mechanistic analysis. Process Biochem.92, 437–446. 10.1016/j.procbio.2020.02.002
4
AdamuA.WahabR. A.HuyopF. (2016). L-2-Haloacid dehalogenase (DehL) from Rhizobium sp. RC1. SpringerPlus5:695. 10.1186/s40064-016-2328-9
5
AdamuA.WahabR. A.ShamsirM. S.AliyuF.HuyopF. (2017b). Deciphering the catalytic amino acid residues of L-2-haloacid dehalogenase (DehL) from Rhizobium sp. RC1: an in silico analysis. Comput. Biol. Chem.70, 125–132. 10.1016/j.compbiolchem.2017.08.007
6
AmeenF.AlNadhariS.Al-HomaidanA. A. (2021). Marine microorganisms as an untapped source of bioactive compounds. Saudi J. Biol. Sci.28, 224–231. 10.1016/j.sjbs.2020.09.052
7
AraiR.Kukimoto-NiinoM.KuroishiC.BesshoY.ShirouzuM.YokoyamaS. (2006). Crystal structure of the probable haloacid dehalogenase PH0459 from Pyrococcus horikoshii OT3. Protein Sci.15, 373–377. 10.1110/ps.051922406
8
ArtabeA. E.Cunha-SilvaH.BarrancoA. (2020). Enzymatic assays for the assessment of toxic effects of halogenated organic contaminants in water and food. A review. Food Chem. Toxicol.145, 111677–111697. 10.1016/j.fct.2020.111677
9
AtashgahiS.LiebensteinerM. G.JanssenD. B.SmidtH.StamsA. J. M.SipkemaD. (2018). Microbial synthesis and transformation of inorganic and organic chlorine compounds. Front. Microbiol.9:3079. 10.3389/fmicb.2018.03079
10
BabkovaP.DunajovaZ.ChaloupkovaR.DamborskyJ.BednarD.MarekM. (2020). Structures of hyperstable ancestral haloalkane dehalogenases show restricted conformational dynamics. Comput. Struct. Biotechnol. J.18, 1497–1508. 10.1016/j.csbj.2020.06.021
11
BagherbaigiS.GicanaR.LamisR. J.NematiM.HuyopF. (2013). Characterisation of Arthrobacter sp. S1 that can degrade α and β-haloalkanoic acids isolated from contaminated soil. Ann. Microbiol.63, 1363–1369. 10.1007/s13213-012-0595-4
12
BehbahaniM.LinB.PharesT. L.SeoY. (2018). Understanding the impact of water distribution system conditions on the biodegradation of haloacetic acids and expression of bacterial dehalogenase genes. J. Hazard. Mater.351, 293–300. 10.1016/j.jhazmat.2018.02.047
13
BerezovskyI. N.GuarneraE.ZhengZ. (2017). Basic units of protein structure, folding, and function. Prog. Biophys. Mol. Bio128, 85–99. 10.1016/j.pbiomolbio.2016.09.009
14
BidlemanT. F.AnderssonA.JantunenL. M.KucklickJ. R.KylinH.LetcherR. J.et al. (2019). A review of halogenated natural products in Arctic, Subarctic and Nordic ecosystems. Emerg. Contam.5, 89–115. 10.1016/j.emcon.2019.02.007
15
BommariusA. (2015). Biocatalysis: a status report. Annu. Rev. Chem.6, 319–345. 10.1146/annurev-chembioeng-061114-123415
16
BreuerM.DitrichK.HabicherT.HauerB.KeselerM. D.SturmerR.et al. (2004). Industrial methods for the production of optically active intermediates. Angew Chem. Int. Edit.43, 788–824. 10.1002/anie.200300599
17
BrokampA.HappeB.SchmidtF. R. (1996). Cloning and nucleotide sequence of a D,L-haloalkanoic acid dehalogenase encoding gene from Alcaligenes xylosoxidans ssp. denitrificans ABIV. Biodegradation7, 383–396. 10.1007/BF00056422
18
BunzelH. A.AndersonJ. L. R.MulhollandA. J. (2021). Designing better enzymes: insights from directed evolution. Curr. Opin. Struct. Biol.67, 212–218. 10.1016/j.sbi.2020.12.015
19
BustilloA.AleuJ.Hernández-GalánR.ColladoI. (2003). Biotransformation of the fungistatic compound (R)-(+)-1-(4′-chlorophenyl) propan-1-ol by Botrytis cinerea. J. Mol. Catal. B Enzym.21, 267–271. 10.1016/S1381-1177(02)00231-X
20
CattoglioC.DarzacqX.TjianR.HansenA. S. (2020). Estimating cellular abundances of halo-tagged proteins in live mammalian cells by flow cytometry. Bio Protoc.10, 3527–3544. 10.21769/BioProtoc.3527
21
ChenB. S.Ribeiro de SouzaF. Z. (2019). Enzymatic synthesis of enantiopure alcohols: current state and perspectives. RSC Adv.9, 2102–2115. 10.1039/C8RA09004A
22
ChibaY.YoshidaT.ItoN.NishimuraH.ImadaC.YasudaH.et al. (2009). Isolation of a bacterium possessing a haloacid dehalogenase from a marine sediment core. Microbes Environ.24, 276–279. 10.1264/jsme2.ME09123
23
ChuanD.FanR.WangY.RenY.WangC.DuY.et al. (2020). Stereocomplex poly(lactic acid)-based composite nanofiber membranes with highly dispersed hydroxyapatite for potential bone tissue engineering. Compos. Sci. Technol.192, 1–12. 10.1016/j.compscitech.2020.108107
24
ClaytonA. D.LabesR.BlackerA. J. (2020). Combination of chemocatalysis and biocatalysis in flow. Curr. Opin. Green Sustain. Chem.26:100378. 10.1016/j.cogsc.2020.100378
25
de OliveiraB. F. R.CarrC. M.DobsonA. D. W.LaportM. S. (2020). Harnessing the sponge microbiome for industrial biocatalysts. Appl. Microbiol. Biot.104, 8131–8154. 10.1007/s00253-020-10817-3
26
DiezA.PrietoM. I.AlvarezM. J.BautistaJ. M.GarridoA.PuyetA. (1996). Improved catalytic performance of a 2-haloacid dehalogenase from Azotobacter sp. by ion-exchange immobilisation. Biochem. Biophl. Res. Co220, 828–833. 10.1006/bbrc.1996.0489
27
DöbberJ.PohlM. (2017). HaloTag™: evaluation of a covalent one-step immobilization for biocatalysis. J. Biotechnol.241, 170–174. 10.1016/j.jbiotec.2016.12.004
28
EdbeibM.WahabR.HuyopF.AksoyH.KayaY. (2020). Further analysis of Burkholderia pseudomallei MF2 and identification of putative dehalogenase gene by PCR. Indones J. Chem.20, 386–394. 10.22146/ijc.43262
29
EncellL. P.Friedman OhanaR.ZimmermanK.OttoP.VidugirisG.WoodM. G.et al. (2012). Development of a dehalogenase-based protein fusion tag capable of rapid, selective, and covalent attachment to customizable ligands. Curr. Chem. Genomics6, 55–71. 10.2174/1875397301206010055
30
EnglandC. G.LuoH.CaiW. (2015). HaloTag technology: a versatile platform for biomedical applications. Bioconjug. Chem.26, 975–986. 10.1021/acs.bioconjchem.5b00191
31
ErdmannR. S.BaguleyS. W.RichensJ. H.WissnerR. F.XiZ.AllgeyerE. S.et al. (2019). Labeling strategies matter for super-resolution microscopy: a comparison between HaloTags and SNAP-tags. Cell Chem. Biol.26, 584–592. 10.1016/j.chembiol.2019.01.003
32
FanY.ChenS. J.LiQ. Q.ZengY.YanX.MaiB. X. (2020). Uptake of halogenated organic compounds (HOCs) into peanut and corn during the whole life cycle grown in an agricultural field. Envirorn. Pollut.263, 1–9. 10.1016/j.envpol.2020.114400
33
FreitasA. I.DominguesL.AguiarT. Q. (2021). Tag-mediated single-step purification and immobilization of recombinant proteins toward protein-engineered advanced materials. J. Adv. Res. 10.1016/j.jare.2021.06.010. [Epub ahead of print].
34
GarbujoS.GalbiatiE.SalvioniL.MazzucchelliM.FrascottiG.SunX.et al. (2020). Functionalization of colloidal nanoparticles with a discrete number of ligands based on a “HALO-bioclick” reaction. Chem. Commun.56, 11398–11401. 10.1039/D0CC04355A
35
GongY.MaS.WangY.XuY.SunA.ZhangY.et al. (2018). Characterization of a novel deep-sea microbial esterase EstC10 and its use in the generation of (R)-methyl-2-chloropropionate. J. Oceanol. Limnol.36, 473–482. 10.1007/s00343-018-6297-4
36
GulI.BogaleT. F.ChenY.YangX.FangR.FengJ.et al. (2020a). A paper-based whole-cell screening assay for directed evolution-driven enzyme engineering. Appl. Microbiol. Biot.104, 6013–6022. 10.1007/s00253-020-10615-x
37
GulI.WangQ.JiangQ.FangR.TangL. (2020b). Enzyme immobilization on glass fiber membrane for detection of halogenated compounds. Anal. Biochem.609, 1–8. 10.1016/j.ab.2020.113971
38
GurushankaraH. P. (2021). Chapter 13-Recent developments in biocatalysis and its influence on the pharmaceutical industry, in Recent Developments in Applied Microbiology and Biochemistry, ed ViswanathB. (Cambridge, MA: Academic Press), 127–139.
39
HamidA. A. A.HamidT. H. T. A.WahabR. A.OmarM. S. S.HuyopF. (2015). An S188V mutation alters substrate specificity of non-stereospecific α-haloalkanoic acid dehalogenase E (DehE). PLoS ONE10:e0121687. 10.1371/journal.pone.0121687
40
HamidT. H. T. A.HamidA. A. A.HuyopF. (2011). A review on non-stereospecific haloalkanoic acid dehalogenases. Afr. J. Biotechnol.10, 9725–9736. 10.5897/AJB11.934
41
HasanA. K. M. Q.TakadaH.KoshikawaH.LiuJ. Q.KuriharaT.EsakiN.et al. (1994). Two kinds of 2-halo acid dehalogenases from Pseudomonas sp. YL induced by 2-chloroacrylate and 2-chloropropionate. Biosci. Biotechnol. Biochem.58, 1599–1602. 10.1271/bbb.58.1599
42
HeidarrezaeiM.ShokraviH.HuyopF.Rahimian KoloorS. S.PetruM. (2020). Isolation and characterization of a novel bacterium from the marine environment for trichloroacetic acid bioremediation. Appl. Sci.10, 1–17. 10.3390/app10134593
43
HellalJ.JoulianC.UrienC.FerreiraS.DenonfouxJ.HermonL.et al. (2021). Chlorinated ethene biodegradation and associated bacterial taxa in multi-polluted groundwater: insights from biomolecular markers and stable isotope analysis. Sci. Total Environ.763, 1–10. 10.1016/j.scitotenv.2020.142950
44
HermonL.DenonfouxJ.HellalJ.JoulianC.FerreiraS.VuilleumierS.et al. (2018). Dichloromethane biodegradation in multi-contaminated groundwater: insights from biomolecular and compound-specific isotope analyses. Water Res.142, 217–226. 10.1016/j.watres.2018.05.057
45
HigginsT. P.HopeS. J.EffendiA. J.DawsonS.DancerB. N. (2005). Biochemical and molecular characterisation of the 2,3-dichloro-1-propanol dehalogenase and stereospecific haloalkanoic dehalogenases from a versatile Agrobacterium sp. Biodegradation16, 485–492. 10.1007/s10532-004-5670-5
46
HillK. E.MarchesiJ. R.WeightmanA. J. (1999). Investigation of two evolutionarily unrelated halocarboxylic acid dehalogenase gene families. J. Bacteriol.181, 2535–2547. 10.1128/JB.181.8.2535-2547.1999
47
HisanoT.HataY.FujiiT.LiuJ.-Q.KuriharaT.EsakiN.et al. (1996). Crystal structure of L-2-haloacid dehalogenase from Pseudomonas sp. YL: an α/β hydrolase structure that is different from the α/β hydrolase fold. J. Biol. Chem.271, 20322–20330. 10.1074/jbc.271.34.20322
48
HorisakiT.YoshidaE.SumiyaK.TakemuraT.YamaneH.NojiriH. (2011). Isolation and characterization of monochloroacetic acid-degrading bacteria. J. Gen. Appl. Microbiol.57, 277–284. 10.2323/jgam.57.277
49
HuyopF.SudiI. Y. (2012). D-specific dehalogenases, a review. Biotechnol. Biotec. Eq.26, 2817–2822. 10.5504/BBEQ.2011.0143
50
IdrisRatnaningsihE. (2015). Cloning of haloacid dehalogenase gene from Bacillus cereus local strain with the addition of restriction sites. Procedia Chem.16, 314–320. 10.1016/j.proche.2015.12.058
51
JanssenD. B. (2004). Evolving haloalkane dehalogenases. Curr. Opin. Chem. Biol.8, 150–159. 10.1016/j.cbpa.2004.02.012
52
KajlaS.NagiG. K.KumariR. (2021). Microorganisms employed in the removal of contaminants from wastewater of iron and steel industries. Rend Lincei Sci. Fis.32, 257–272. 10.1007/s12210-021-00982-6
53
KimD.-H.ParkC. G.KimY. J. (2020). Characterizing the potential estrogenic and androgenic activities of two disinfection byproducts, mono-haloacetic acids and haloacetamides, using in vitro bioassays. Chemosphere242, 1–6. 10.1016/j.chemosphere.2019.125198
54
KirkinciS. F.EdbeibM. F.AksoyH. M.MarakliS.KayaY. (2021). Identification of Dalapon degrading bacterial strain, Psychrobacter sp. TaeBurcu001 isolated from Antarctica. Polar Sci. 28, 1–9. 10.1016/j.polar.2021.100656
55
KöhlerR.BrokampA.SchwarzeR.ReitingR. H.SchmidtF. R. (1998). Characteristics and DNA-sequence of a cryptic haloalkanoic acid dehalogenase from Agrobacterium tumefaciens RS5. Curr. Microbiol.36, 96–101. 10.1007/s002849900286
56
KondoH.FujimotoK. J.TanakaS.DekiH.NakamuraT. (2015). Theoretical prediction and experimental verification on enantioselectivity of haloacid dehalogenase L-DEX YL with chloropropionate. Chem. Phys. Lett.623, 101–107. 10.1016/j.cplett.2015.01.053
57
KondoH.NakamuraT.TanakaS. (2014). A significant role of Arg41 residue in the enzymatic reaction of haloacid dehalogenase L-DEX YL studied by QM/MM method. J. Mol. Catal. B Enzym.110, 23–31. 10.1016/j.molcatb.2014.09.006
58
KumarA.PillayB.OlaniranA. O. (2016). L-2-haloacid dehalogenase from Ancylobacter aquaticus UV5: sequence determination and structure prediction. Int. J. Biol. Macromol.83, 216–225. 10.1016/j.ijbiomac.2015.11.066
59
KunkaA.DamborskyJ.ProkopZ. (2018). Chapter Seven - Haloalkane Dehalogenases From Marine Organisms, in Methods in Enzymology, ed MooreB. S. (Cambridge, MA: Academic Press), 203–251.
60
KuradeM. B.HaY. H.XiongJ. Q.GovindwarS. P.JangM.JeonB. H. (2021). Phytoremediation as a green biotechnology tool for emerging environmental pollution: a step forward towards sustainable rehabilitation of the environment. Chem. Eng. J.415, 1–19. 10.1016/j.cej.2021.129040
61
KuriharaT. (2011). A mechanistic analysis of enzymatic degradation of organohalogen compounds. Biosci. Biotechnol. Biochem.75, 189–198. 10.1271/bbb.100746
62
KuriharaT.EsakiN. (2008). Bacterial hydrolytic dehalogenases and related enzymes: occurrences, reaction mechanisms, and applications. Chem. Rec.8, 67–74. 10.1002/tcr.20141
63
KuriharaT.EsakiN.SodaK. (2000). Bacterial 2-haloacid dehalogenases: structures and reaction mechanisms. J. Mol. Catal. B Enzym.10, 57–65. 10.1016/S1381-1177(00)00108-9
64
KurumbangN. P.DvorakP.BendlJ.BrezovskyJ.ProkopZ.DamborskyJ. (2014). Computer-assisted engineering of the synthetic pathway for biodegradation of a toxic persistent pollutant. ACS Synth. Biol.3, 172–181. 10.1021/sb400147n
65
Leemans MartinL.PeschkeT.VenturoniF.MostardaS. (2020). Pharmaceutical industry perspectives on flow chemocatalysis and biocatalysis. Curr. Opin. Green Sustain25:100350. 10.1016/j.cogsc.2020.04.011
66
LeighJ. A.SkinnerA. J.CooperR. A. (1986). Isolation and partial characterisation of dehalogenase-deficient mutants of a Rhizobium sp. FEMS Microbiol. Lett.36, 163–166. 10.1111/j.1574-6968.1986.tb01688.x
67
LiJ.CaiW.ZhuL. (2011). The characteristics and enzyme activities of 4-chlorophenol biodegradation by Fusarium sp. Bioresource Technol.102, 2985–2989. 10.1016/j.biortech.2010.10.006
68
LiaoL.SunX.ZengY.LuoW.YuY.ChenB. (2015). A new L-haloacid dehalogenase from the Arctic psychrotrophic Pseudoalteromonas sp. BSW20308. Polar Biol.38, 1161–1169. 10.1007/s00300-015-1674-3
69
LiuJ.ZhaiH.SunY.WuS.ChenS. (2021). Developing high strength poly(L-lactic acid) nanofiber yarns for biomedical textile materials: a comparative study of novel nanofiber yarns and traditional microfiber yarns. Mater. Lett.300:130229. 10.1016/j.matlet.2021.130229
70
LiuJ. Q.KuriharaT.HasanA. K.Nardi-DeiV.KoshikawaH.EsakiN.et al. (1994). Purification and characterization of thermostable and nonthermostable 2-haloacid dehalogenases with different stereospecificities from Pseudomonas sp. strain YL. Appl. Environ. Microbiol.60, 2389–2393. 10.1128/aem.60.7.2389-2393.1994
71
LongK.ShaY.MoY.WeiS.WuH.LuD.et al. (2021). Androgenic and teratogenic effects of iodoacetic acid drinking water disinfection byproduct in vitro and in vivo. Environ. Sci. Technol.55, 3827–3835. 10.1021/acs.est.0c06620
72
LongoL. M.KumruO. S.MiddaughC. R.BlaberM. (2014). Evolution and design of protein structure by folding nucleus symmetric expansion. Structure22, 1377–1384. 10.1016/j.str.2014.08.008
73
LouJ.WangW.ZhuL. (2021). Transformation of emerging disinfection byproducts Halobenzoquinones to haloacetic acids during chlorination of drinking water. Chem. Eng. J.418, 129326. 10.1016/j.cej.2021.129326
74
LouY. Y.FontmorinJ.-M.AmraneA.FourcadeF.GenesteF. (2021). Metallic nanoparticles for electrocatalytic reduction of halogenated organic compounds: A review. Electrochim. Acta377, 138039. 10.1016/j.electacta.2021.138039
75
MaF.HeL.LindnerE.WuD. Y. (2021). Highly porous poly(L-lactic) acid nanofibers as a dual-signal paper-based bioassay platform for in vitro diagnostics. Appl. Surf. Sci.542:148732. 10.1016/j.apsusc.2020.148732
76
MarshallJ. R.Mangas-SanchezJ.TurnerN. J. (2021). Expanding the synthetic scope of biocatalysis by enzyme discovery and protein engineering. Tetrahedron82:131926. 10.1016/j.tet.2021.131926
77
MazurA.PrudnikovaT.GrinkevichP.MestersJ. R.MrazovaD.ChaloupkovaR.et al. (2021). The tetrameric structure of the novel haloalkane dehalogenase DpaA from Paraglaciecola agarilytica NO2. Acta Crystallogr. D Struct. Biol.77, 347–356. 10.1107/S2059798321000486
78
Mena-BenitezG. L.Gandia-HerreroF.GrahamS.LarsonT. R.McQueen-MasonS. J.FrenchC. E.et al. (2008). Engineering a catabolic pathway in plants for the degradation of 1,2-dichloroethane. Plant Physiol.147, 1192–1198. 10.1104/pp.108.119008
79
MerrillR. A.SongJ.KephartR. A.KlompA. J.NoackC. E.StrackS. (2019). A robust and economical pulse-chase protocol to measure the turnover of HaloTag fusion proteins. J. Biol. Chem.294, 16164–16171. 10.1074/jbc.RA119.010596
80
Minner-MeinenR.WeberJ. N.AlbrechtA.MatisR.BehneckeM.TietgeC.et al. (2021). Split-HaloTag® imaging assay for sophisticated microscopy of protein-protein interactions in planta. Plant Commun.2:100212. 10.1016/j.xplc.2021.100212
81
MohamedE.MohamedJ.HuyopF.WahabR.Abdul-HamidH. (2020). Developing herbicide tolerant transgenic plants for sustainable weed management. Int. J. Emerg. Trends Eng. Res.8, 60–66. 10.30534/ijeter/2020/0981.22020
82
MuzikárM.KresinováZ.SvobodováK.FilipováA.CvančarováM.CajthamlováK.et al. (2011). Biodegradation of chlorobenzoic acids by ligninolytic fungi. J. Hazard. Mater.196, 386–394. 10.1016/j.jhazmat.2011.09.041
83
NarayananH.DingfelderF.Butt,éA.LorenzenN.SokolovM.ArosioP. (2021). Machine Learning for biologics: opportunities for protein engineering, developability, and formulation. Trends Pharmacol. Sci.42, 151–165. 10.1016/j.tips.2020.12.004
84
Nardi-DeiV.KuriharaT.ParkC.MiyagiM.TsunasawaS.SodaK.et al. (1999). DL-2-Haloacid dehalogenase from Pseudomonas sp. 113 is a new class of dehalogenase catalyzing hydrolytic dehalogenation not involving enzyme-substrate ester intermediate. J. Biol. Chem.274, 20977–20981. 10.1074/jbc.274.30.20977
85
NguyenB. T.HsiehJ. L.LoS. C.WangS. Y.HungC. H.HuangE.et al. (2021). Biodegradation of dioxins by Burkholderia cenocepacia strain 869T2: role of 2-haloacid dehalogenase. J. Hazard. Mater.401:123347. 10.1016/j.jhazmat.2020.123347
86
NovakH.LittlechildJ. (2013). Marine enzymes with applications for biosynthesis of fine chemicals, in Marine Enzymes for Biocatalysis, ed TrinconeA. (Sawston: Woodhead Publishing), 89–106.
87
NovakH. R.SayerC.IsupovM. N.PaszkiewiczK.GotzD.SpraggA. M.et al. (2013a). Marine Rhodobacteraceae L-haloacid dehalogenase contains a novel His/Glu dyad that could activate the catalytic water. FEBS J.280, 1664–1680. 10.1111/febs.12177
88
NovakH. R.SayerC.PanningJ.LittlechildJ. A. (2013b). Characterisation of an L-haloacid dehalogenase from the marine psychrophile Psychromonas ingrahamii with potential industrial application. Mar. Biotechnol.15, 695–705. 10.1007/s10126-013-9522-3
89
OyewusiH. A.HuyopF.WahabR. A. (2020a). Molecular docking and molecular dynamics simulation of Bacillus thuringiensis dehalogenase against haloacids, haloacetates and chlorpyrifos. J. Biomol. Struct. Dyn.38, 1–16. 10.1080/07391102.2019.1580616
90
OyewusiH. A.HuyopF.WahabR. A.HamidA. A. A. (2021a). In silico assessment of dehalogenase from Bacillus thuringiensis H2 in relation to its salinity-stability and pollutants degradation. J. Biomol. Struct. Dyn.39, 1–15. 10.1080/07391102.2021.1927846
91
OyewusiH. A.WahabR. A.HuyopF. (2020b). Dehalogenase-producing halophiles and their potential role in bioremediation. Mar. Pollut. Bull.160:111603. 10.1016/j.marpolbul.2020.111603
92
OyewusiH. A.WahabR. A.HuyopF. (2021b). Whole genome strategies and bioremediation insight into dehalogenase-producing bacteria. Mol. Biol. Rep.48, 2687–2701. 10.1007/s11033-021-06239-7
93
ParkC.KuriharaT.YoshimuraT.SodaK.EsakiN. (2003). A new DL-2-haloacid dehalogenase acting on 2-haloacid amides: purification, characterization, and mechanism. J. Mol. Catal. B Enzym.23, 329–336. 10.1016/S1381-1177(03)00096-1
94
ParkerK.ColbyJ. (1995). Immobilisation of the D-2-haloacid dehalogenase from Pseudomonas putida strain AJ1/23. Biodegradation6, 191–201. 10.1007/BF00700457
95
PoelarendsG. J.WhitmanC. P. (2010). Mechanistic and structural studies of microbial dehalogenases: How nature cleaves a carbon–halogen bond, in Comprehensive Natural Products II, ed LiuH. W.ManderL. (Oxford: Elsevier), 89–123.
96
PolnischE.KneifelH.FranzkeH.HofmannK. H. (1991). Degradation and dehalogenation of monochlorophenols by the phenol-assimilating yeast Candida maltosa. Biodegradation2, 193–199. 10.1007/BF00124493
97
RatnaningsihE.IdrisI. (2018). Cloning and expression of haloacid dehalogenase gene from Bacillus cereus IndB1. Indones. J. Biotechnol.22, 55–60. 10.22146/ijbiotech.27338
98
RidderI. S.RozeboomH. J.KalkK. H.JanssenD. B.DijkstraB. W. (1997). Three-dimensional structure of L-2-haloacid dehalogenase from Xanthobacter autotrophicus GJ10 complexed with the substrate-analogue formate. J. Biol. Chem.272, 33015–33022. 10.1074/jbc.272.52.33015
99
Rosland AbelS.IbrahimN.HuyopF. (2012). Identification of Serratia marcescens SE1 and determination of its herbicide 2,2-dichloropropionate (2,2-DCP) degradation potential. Malays. J. Microbiol.8, 259–265. 10.21161/mjm.44412
100
RyeC. A.IsupovM. N.LebedevA. A.LittlechildJ. A. (2009). Biochemical and structural studies of a L-haloacid dehalogenase from the thermophilic archaeon Sulfolobus tokodaii. Extremophiles13, 179–190. 10.1007/s00792-008-0208-0
101
SantiM.SancinetoL.NascimentoV.Braun AzeredoJ.OrozcoE. V. M.AndradeL. H.et al. (2021). Flow biocatalysis: a challenging alternative for the synthesis of apis and natural compounds. Int. J. Mol. Sci.22:990. 10.3390/ijms22030990
102
SatpathyR.KonkimallaV. S. B.RathaJ. (2015). In-silico gene co-expression network analysis in Paracoccidioides brasiliensis with reference to haloacid dehalogenase superfamily hydrolase gene. J. Pharm. Bioallied Sci.7, 212–217. 10.4103/0975-7406.160023
103
SatpathyR.KonkimallaV. S. B.RathaJ. (2016). In silico phylogenetic analysis and molecular modelling study of 2-haloalkanoic acid dehalogenase enzymes from bacterial and fungal origin. Adv. Bioinform.2016, 8701201–8701201. 10.1155/2016/8701201
104
SchmidbergerJ. W.WilceJ. A.TsangJ. S. H.WilceM. C. J. (2007). Crystal structures of the substrate free-enzyme, and reaction intermediate of the HAD superfamily member, haloacid dehalogenase DehIVa from Burkholderia cepacia MBA4. J. Mol. Biol.368, 706–717. 10.1016/j.jmb.2007.02.015
105
SchmidbergerJ. W.WilceJ. A.WeightmanA. J.WhisstockJ. C.WilceM. C. (2008). The crystal structure of DehI reveals a new alpha-haloacid dehalogenase fold and active-site mechanism. J. Mol. Biol.378, 284–294. 10.1016/j.jmb.2008.02.035
106
SchoberM.FaberK. (2013). Inverting hydrolases and their use in enantioconvergent biotransformations. Trends Biotechnol.31, 468–478. 10.1016/j.tibtech.2013.05.005
107
SenguptaS.DasP.DattaS. (2020). Fermentative production of optically pure lactic acid from renewable materials, in Encyclopedia of Renewable and Sustainable Materials, ed HashmiS.ChoudhuryI. A. (Oxford: Elsevier), 447–453.
108
ShaharH.TanL. L.TaG. C.HengL. Y. (2019a). Detection of halogenated hydrocarbon pollutants using enzymatic reflectance biosensor. Sensors Actuat. B Chem.281, 80–89. 10.1016/j.snb.2018.10.076
109
ShaharH.TanL. L.TaG. C.HengL. Y. (2019b). Optical enzymatic biosensor membrane for rapid in situ detection of organohalide in water samples. Microchem. J.146, 41–48. 10.1016/j.microc.2018.12.052
110
SiwekA.OmiR.HirotsuK.JitsumoriK.EsakiN.KuriharaT.et al. (2013). Binding modes of DL-2-haloacid dehalogenase revealed by crystallography, modeling and isotope effects studies. Arch. Biochem. Biophys.540, 26–32. 10.1016/j.abb.2013.09.012
111
SmithJ. M.HarrisonK.ColbyJ. (1990). Purification and characterization of D-2-haloacid dehalogenase from Pseudomonas putida strain AJ1/23. J. Gen. Microbiol.136, 881–886. 10.1099/00221287-136-5-881
112
SodaK.KuriharaT.LiuJ. Q.NardiDeiV.ParkC.MiyagiM.et al. (1996). Bacterial 2-haloacid dehalogenases: structures and catalytic properties. Pure Appl. Chem.68, 2097–2103. 10.1351/pac199668112097
113
SuX.DengL.KongK. F.TsangJ. S. (2013). Enhanced degradation of haloacid by heterologous expression in related Burkholderia species. Biotechnol. Bioeng.110, 2687–2696. 10.1002/bit.24917
114
SudiI. Y.HamidA. A.ShamsirM. S.JamaluddinH.WahabR. A.HuyopF. (2014). Insights into the stereospecificity of the D-specific dehalogenase from Rhizobium sp. RC1 toward D- and L-2-chloropropionate. Biotechnol. Biotechnol. Equip.28, 608–615. 10.1080/13102818.2014.937907
115
TanokuraM.MiyakawaT.GuanL.HouF. (2015). Structural analysis of enzymes used for bioindustry and bioremediation. Biosci. Biotech. Biochem.79, 1391–1401. 10.1080/09168451.2015.1052770
116
Taylor StephenC. (1985). D-2-haloalkanoic acid halidohydrolase. US19850789765. U.S. Patent.
117
ThippeswamyM.RajasrerlathaV.ShubhaD.NivedithaB. T. (2021). Chapter 24- Metagenomics and future perspectives in discovering pollutant degrading enzymes from soil microbial communities, in Recent Developments in Applied Microbiology and Biochemistry, ed ViswanathB. (Cambridge, MA: Academic Press), 257–267.
118
TunyasuvunakoolK.AdlerJ.WuZ.GreenT.ZielinskiM.ŽídekA.et al. (2021). Highly accurate protein structure prediction for the human proteome. Nature596, 1–9. 10.1038/s41586-021-03828-1
119
van der PloegJ.van HallG.JanssenD. B. (1991). Characterization of the haloacid dehalogenase from Xanthobacter autotrophicus GJ10 and sequencing of the dhlB gene. J. Bacteriol.173, 7925–7933. 10.1128/jb.173.24.7925-7933.1991
120
VranckenJ. P. M.TameJ. R. H.VoetA. R. D. (2020). Development and applications of artificial symmetrical proteins. Comput. Struct. Biotechnol. J.18, 3959–3968. 10.1016/j.csbj.2020.10.040
121
WahhabB.AnuarN.WahabR.Al-NimerM.SamsulrizalN.Abdul HamidA.et al. (2020). Characterization of a 2,2-dichloropropionic acid (2,2-DCP) degrading alkalotorelant Bacillus megaterium strain BHS1 isolated from blue lake in turkey. J. Trop. Life Sci.10, 245–252. 10.11594/jtls.10.03.08
122
WangC.XiJ.HuH. Y.WenX. (2009). Biodegradation of gaseous chlorobenzene by white-rot fungus Phanerochaete chrysosporium. Biomed. Environ. Sci.21, 474–478. 10.1016/S0895-3988(09)60005-2
123
WangS.ChengZ.XuY.YangL.WangJ. B.TianZ.et al. (2020). Structure-guided protein design of fluoroacetate dehalogenase for kinetic resolution of rac-2-bromobutyric acid. Green Synth. Catal.1, 60–65. 10.1016/j.gresc.2020.05.004
124
WangY.CaoX.FengY.XueS. (2016). Environment-induced conformational and functional changes of L-2-haloacid dehalogenase. J. Biosci. Bioeng.121, 491–496. 10.1016/j.jbiosc.2015.09.008
125
WangY.FengY.CaoX.LiuY.XueS. (2018). Insights into the molecular mechanism of dehalogenation catalyzed by D-2-haloacid dehalogenase from crystal structures. Sci. Rep.8:1454. 10.1038/s41598-017-19050-x
126
WangY.XueS.ZhouQ.PeiD. (2020). Recent progress in 2-haloacid dehalogenases. Chin. J. Biotechnol.36, 868–878. 10.13345/j.cjb.190370
127
WatanabeS.ItoM.KigawaT. (2021). DiRect: Site-directed mutagenesis method for protein engineering by rational design. Biochem. Biophys. Res. Co551, 107–113. 10.1016/j.bbrc.2021.03.021
128
WeightmanA. J.WeightmanA. L.SlaterJ. H. (1982). Stereospecificity of 2-monochloropropionate dehalogenation by the two dehalogenases of Pseudomonas putida PP3: evidence for two different dehalogenation mechanisms. Microbiology128, 1755–1762. 10.1099/00221287-128-8-1755
129
WoolfsonD. N. (2021). A brief history of de novo protein design: minimal, rational, and computational. J. Mol. Biol. 10.1016/j.jmb.2021.167160. [Epub ahead of print].
130
WuL.QinL.NieY.XuY.ZhaoY.-L. (2021). Computer-aided understanding and engineering of enzymatic selectivity. Biotechnol. Adv.10.1016/j.biotechadv.2021.107793. [Epub ahead of print].
131
WuQ.BouwmanH.UrenR. C.van der LingenC. D.VetterW. (2019). Halogenated natural products and anthropogenic persistent organic pollutants in chokka squid (Loligo reynaudii) from three sites along the South Atlantic and Indian Ocean coasts of South Africa. Environ. Pollut.255:113282. 10.1016/j.envpol.2019.113282
132
XieG.PanD.HeW.GaoG. (2015). Application of L-2-haloacid dehalogenase from thermophilic Archaea Sulfolobus tokodaii in the production of D-lactic acid. Chem. J. Chinese Univ.36, 698–703. 10.7503/cjcu20140739
133
XiongW.LiuB.ShenY.JingK.SavageT. R. (2021). Protein engineering design from directed evolution to de novo synthesis. Biochem. Eng. J.174:108096. 10.1016/j.bej.2021.108096
134
XuS.QinS.PanX.-M. (2004). Thermal and conformational stability of Ssh10b protein from archaeon Sulfolobus shibattae. Biochem. J.382, 433–440. 10.1042/BJ20040191
135
Yavari MaroufiL.GhorbaniM.MohammadiM.PezeshkiA. (2021). Improvement of the physico-mechanical properties of antibacterial electrospun poly lactic acid nanofibers by incorporation of guar gum and thyme essential oil. Colloid Surface A622:126659. 10.1016/j.colsurfa.2021.126659
136
YuF.LiY.WangH.PengT.WuY. R.HuZ. (2021). Microbial debromination of hexabromocyclododecanes. Appl. Microbiol. Biot. 105, 4535–4550. 10.1007/s00253-021-11095-3
137
Zainal AbidinM. H.Abd HalimK. B.HuyopF.Tengku Abdul HamidT. H.Abdul WahabR.Abdul HamidA. A. (2019). The mechanistic role of active site residues in non-stereo haloacid dehalogenase E (DehE). J. Mol. Graph. Model.90, 219–225. 10.1016/j.jmgm.2019.05.003
138
ZakaryS.OyewusiH.HuyopF. (2021). Genomic analysis of mesorhizobium loti strain tono reveals dehalogenases for bioremediation. J. Trop. Life Sci.11, 67–77. 10.11594/jtls.11.01.09
139
ZhangC.AllenK. N.DunawaymarianoD. (2018). Mechanism of substrate recognition and catalysis of the haloalkanoic acid dehalogenase family member α-phosphoglucomutase. Biochemistry57, 4504–4517. 10.1021/acs.biochem.8b00396
140
ZhangC.YuanZ.LiuY.ZhangQ.ChenY.SaleemS.et al. (2021). The online detection of halogenated hydrocarbon in the atmosphere. Opt Laser Eng142:106586. 10.1016/j.optlaseng.2021.106586
141
ZhangJ.CaoX.XinY.XueS.ZhangW. (2013). Purification and characterization of a dehalogenase from Pseudomonas stutzeri DEH130 isolated from the marine sponge Hymeniacidon perlevis. World J. Microbiiol. Biotechnol.29, 1791–1799. 10.1007/s11274-013-1340-2
142
ZhangJ.JiangL.ChenX.LvK.BasionyM.ZhuG.et al. (2021). Recent advances in biotechnology for marine enzymes and molecules. Curr. Opin. Biotechnol.69, 308–315. 10.1016/j.copbio.2021.05.009
143
ZhangJ.XinY.CaoX.XueS.ZhangW. (2014). Purification and characterization of 2-haloacid dehalogenase from marine bacterium Paracoccus sp. DEH99, isolated from marine sponge Hymeniacidon perlevis. J. Ocean Univ. China13, 91–96. 10.1007/s11802-014-2357-3
144
ZhangM.ShiQ.SongX.WangH.BianZ. (2019). Recent electrochemical methods in electrochemical degradation of halogenated organics: a review. Environ. Sci. Pollut. Res. Int.26, 10457–10486. 10.1007/s11356-019-04533-3
145
ZhouH.LiY.JiangR.WangX.WangY.XueY.et al. (2021). An efficient route towards R-2-phenoxypropionic acid synthesis for biotransformative production of R-2-(4-hydroxyphenoxy) propionic acid. Chinese J. Chem. Eng.32, 315–323. 10.1016/j.cjche.2020.06.013
Summary
Keywords
2-haloacid dehalogenases, protein structure, catalytic mechanism, biochemical properties, application
Citation
Wang Y, Xiang Q, Zhou Q, Xu J and Pei D (2021) Mini Review: Advances in 2-Haloacid Dehalogenases. Front. Microbiol. 12:758886. doi: 10.3389/fmicb.2021.758886
Received
15 August 2021
Accepted
13 September 2021
Published
15 October 2021
Volume
12 - 2021
Edited by
Benwei Zhu, Nanjing Tech University, China
Reviewed by
Fahrul Huyop, Universiti Teknologi Malaysia, Malaysia; Dirk Tischler, Ruhr University Bochum, Germany
Updates
Copyright
© 2021 Wang, Xiang, Zhou, Xu and Pei.
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*Correspondence: Jingliang Xu xujl@zzu.edu.cnDongli Pei peidongli@126.com
This article was submitted to Microbiotechnology, a section of the journal Frontiers in Microbiology
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