Fatty Acid Hydratases: Versatile Catalysts to Access Hydroxy Fatty Acids in Eﬃcient Syntheses of Industrial Interest

catalysts

Review Fatty Acid Hydratases: Versatile Catalysts to Access Hydroxy Fatty Acids in Eﬃcient Syntheses of Industrial Interest

Jana Löwe and Harald Gröger *

Industrial Organic Chemistry and Biotechnology, Faculty of Chemistry, Bielefeld University, Universitätsstraße 25, 33615 Bielefeld, Germany; [email protected] * Correspondence: [email protected]; Tel.: +49-521-106-2057

Received: 21 December 2019; Accepted: 17 February 2020; Published: 3 March 2020

Abstract: The utilization of hydroxy fatty acids has gained more and more attention due to its applicability in many industrial building blocks that require it, for example, polymers or fragrances. Furthermore, hydroxy fatty acids are accessible from biorenewables, thus contributing to a more sustainable raw material basis for industrial chemicals. Therefore, a range of investigations were done on fatty acid hydratases (FAHs), since these enzymes catalyze the addition of water to an unsaturated fatty acid, thus providing an elegant route towards hydroxy-substituted fatty acids. Besides the discovery and characterization of fatty acid hydratases (FAHs), the design and optimization of syntheses with these enzymes, the implementation in elaborate cascades, and the improvement of these biocatalysts, by way of mutation in terms of the substrate scope, has been investigated. This mini-review focuses on the research done on process development using fatty acid hydratases as a catalyst. It is notable that biotransformations, running at impressive substrate loadings of up to 1 280 g L− , have been realized. A further topic of this mini-review is the implementation of fatty acid hydratases in cascade reactions. In such cascades, fatty acid hydratases were, in particular, combined with alcohol dehydrogenases (ADH), Baeyer-Villiger monooxygenases (BVMO), transaminases (TA) and hydrolases, thus enabling access to a broad variety of molecules that are of industrial interest.

Keywords: biocatalysis; cascades; enzymes; fatty acids; hydratases; hydroxy fatty acids

1. Introduction Hydroxy fatty acids are of the highest interest for industrial applications, since they serve as starting materials for polymers [1] or cyclic lactones [2], which are used in fragrances and antibiotics [3], as well as in plasticizers [4], surfactants [5], lubricants [6] and detergent formulations [7]. Besides being available in nature and accessible through plant oil hydrolysis (e.g., in the case of ricinoleic acid from castor oil [8]), chemical approaches towards hydroxy-functionalized fatty acids include oxygenation, epoxidation [9–11] and ozonolysis [12] as key reaction steps. Enzymatically, hydroxy fatty acids can be synthesized by means of oxidation reactions in the presence of 12-hydroxylases [13], cytochrome P450 monooxygenases, lipoxygenases, and peroxygenases [12]. An enzyme-class, which catalyzes the addition of water to non-activated C=C double bonds are hydratases. Representative examples for such enzymes are the linalool dehydratase-isomerase, which converts myrcene towards linool [14], or carotenoid 1,2-hydratase [15], which catalyzes the conversion of acyclic carotenes. However, one of the most common enzymes for the conversion of unsaturated, non-activated C=C-double bonds are fatty acid hydratases (FAHs) [16,17]. Hydratases can only be found in microorganisms [18]. After being reported for the ﬁrst time by Wallen et al. in 1962 [19], they gained high interest since [20,21]. Until now, no detailed statements can be made about the physiological role of fatty acid hydratases. Some

Catalysts 2020, 10, 287; doi:10.3390/catal10030287 www.mdpi.com/journal/catalysts Catalysts 2020, 10, 287 2 of 13 Catalysts 2019, 9, x FOR PEER REVIEW 2 of 12 possiblephysiological functions role are of thefatty detoxiﬁcation acid hydratases. of fatty Some acids possible [22] or changesfunctions in host–microbeare the detoxification interactions of fatty due toacids their [22] capacity or changes to enhance in host–microbe cell hydrophobicity interactions [23 due]. FAHs to their are capacity FAD-dependent to enhance enzymes cell hydrophobicity [24], in which the[23] FAD. FAHs seems are to FAD-dependent stabilize the active enzymes conformation. [24], inThe wh FAD,ich the which FAD is seems none-covalently to stabilize bound the toactive the conservedconformation. N-terminal The FAD, nucleotide which is binding none-covalently motif (GXGXXGX21E bound to the/D) conserved [25], is, however, N-terminal not involved nucleotide in thebinding reaction motif mechanism (GXGXXGX21E/D) for the hydration [25], is, however, of the unsaturated not involved fatty in acid the [reaction22,25]. Furthermore,mechanism for there the arehydration a range of structuralthe unsaturated prerequisites fatty foracid the [22,25]. substrates Furthermore, in order to bethere tolerated are a byrange this enzymeof structural class. Accordingly,prerequisites fatty for the acid substrates hydratases in require order to a carboxylicbe tolerated group by this in the enzyme substrate, class. a distanceAccordingly, of nine fatty carbons acid betweenhydratases the require double a bond carboxylic and the group acid group,in the substrat a minimume, a distance chain length of nine of C-14carbons and between a cis-conformation the double ofbond the and double the acid bond group, [26]. Fora minimum these substrates, chain length the of hydration C-14 and then a cis-conformation proceeds by the of simplethe double addition bond of[26]. water, For these which substrates, makes such the a hydration transformation then proceed a highlys by attractive the simple reaction, addition which of water, is also which the case makes for industrialsuch a transformation purposes. a highly attractive reaction, which is also the case for industrial purposes.

2. Application of Fatty Acid Hydratase in Synthesis and Process Development 2. Application of Fatty Acid Hydratase in Synthesis and Process Development The utilization of enzymes in the chemical industry has become indispensable, and today The utilization of enzymes in the chemical industry has become indispensable, and today numerous large-scale applications, based on the use of enzymes, are already known [27–33]. Industrial numerous large-scale applications, based on the use of enzymes, are already known [27–33]. processes that use enzymes as a catalyst often have excellent productivities. Well-known examples are Industrial processes that use enzymes as a catalyst often have excellent productivities. Well-known the production of acrylamide [34] in the presence of a nitrile hydratase or the use of lipases for the examples are the production of acrylamide [34] in the presence of a nitrile hydratase or the use of hydrolysis of milk fat, oils and fat [27]. In addition, the industry also makes use of the naturally high lipases for the hydrolysis of milk fat, oils and fat [27]. In addition, the industry also makes use of the stereoselectivity of enzymes, and representative examples in this field are the enzymatic production of naturally high stereoselectivity of enzymes, and representative examples in this field are the L-amino acids by means of an aminoacylase [35] or the use of a penicillin acylase for the production of enzymatic production of L-amino acids by means of an aminoacylase [35] or the use of a penicillin antibiotics [36]. acylase for the production of antibiotics [36]. A further enzyme class with broad potential applicability for industrial purpose are fatty acid A further enzyme class with broad potential applicability for industrial purpose are fatty acid hydratases, which are capable of hydrating unsaturated fatty acids, thus forming hydroxy-substituted hydratases, which are capable of hydrating unsaturated fatty acids, thus forming hydroxy- fatty acids [37]. This type of reaction only requires water as a reagent, which also makes this substituted fatty acids [37]. This type of reaction only requires water as a reagent, which also makes transformation attractive from the point of view of sustainability. Since fatty acids are key components this transformation attractive from the point of view of sustainability. Since fatty acids are key of fats and oils as renewable raw materials, this type of biotransformation is also of interest within the components of fats and oils as renewable raw materials, this type of biotransformation is also of current research efforts to develop processes for industrial chemicals based on biorenewable feedstocks interest within the current research efforts to develop processes for industrial chemicals based on instead of petrochemical sources. In Figure1, a representative example for such a transformation with biorenewable feedstocks instead of petrochemical sources. In Figure 1, a representative example for asuch fatty a acid transformation hydratase is shown,with a exemplifiedfatty acid byhydratase the synthesis is shown, of 10-hydroxyoctadecanoic exemplified by the acidsynthesis starting of from10-hydroxyoctadecanoic oleic acid. acid starting from oleic acid.

fatty acid hydratase O O OH OH OH H2O oleic acid 10-hydroxyoctadecanoic acid Figure 1. Reaction scheme for the synthesis of 10-hydroxyoctadecanoic acid starting from oleic acid as Figure 1. Reaction scheme for the synthesis of 10-hydroxyoctadecanoic acid starting from oleic acid a selected example for the synthesis of hydrated fatty acids [37]. as a selected example for the synthesis of hydrated fatty acids [37]. The characterization of fatty acid hydratases has been intensively studied, as well as the process developmentThe characterization of hydroxy fatty of fatty acid acid production hydratases by means has been of this intensively approach. studied, One of as the well ﬁrst as applications the process ofdevelopment hydroxy fatty of acidhydroxy hydratases fatty inacid the processproduction development by means for of the this synthesis approach. of hydroxy One of fatty the acids first isapplications the use of theof hydroxy strain Candida fatty acid tropicalis hydratasesDSM in 3152 the andprocess a mutant development M 25 in for the the transformation synthesis of hydroxy of oleic fatty acids is the use of9 the strain Candida tropicalis DSM 3152 and a mutant M 25 in the transformation1 acid into 3-hydroxy-∆ -cis-1,18-octadecenedioic acid, which gave the desired product with 19.4 g L− 9 afterof oleic 223 acid h in into the 3-hydroxy- fermentationΔ -cis-1,18-octadecenedioic process [38]. In addition, acid, a range which of gave other the comprehensive desired product studies, with −1 including19.4 g L otherafter 223 microbes, h in the which fermentation possess theprocess ability [38] to. convertIn addition, unsaturated a range fattyof other acids comprehensive into hydroxy fattystudies, acids, including were published, other microbes, such as which resting possessSaccharomyces the ability cerevisiae to convertcells unsaturated [39], ruminal fatty bacteria acids [into40] orhydroxy bacteria fatty isolated acids, from were compost published, or soil such [41 ,42as], resting to name Saccharomyces just a few, which cerevisiae converts cells oleic[39], acid ruminal into 10-hydroxyoctadecanoicbacteria [40] or bacteria isolated acid. In from 2009, compost one of or the soil most [41,42], frequently to name used just a fatty few, acid which hydratases converts oleic was acid into 10-hydroxyoctadecanoic acid. In 2009, one of the most frequently used fatty acid hydratases was described for the first time. In detail, the isolation and biochemical characterization of the oleate hydratase from Elizabethkingia meningoseptica (formerly known as Pseudomonas sp. strain 3266 and

Catalysts 2020, 10, 287 3 of 13 described for the first time. In detail, the isolation and biochemical characterization of the oleate hydratase from Elizabethkingia meningoseptica (formerly known as Pseudomonas sp. strain 3266 and widely found in nature, e.g., in soil and water) [43], was reported by Hagen et al. [44]. In 2011, Oh et al. described a fatty acid hydratase of Stenotrophomonas nitritireducens, the genus Stenotrophomonas lives in close association with plants [45], and by means of the corresponding whole cells containing this enzyme, with 31.5 g of 10-hydroxystearic in 4 h, were formed when starting from 30 g of oleic acid [46]. Since 2012, the interest in the development of fatty acid hydratases and its usage in elevated gram-scale applications further increased. In 2012, Oh et al. described a putative fatty acid hydratase, which was isolated from Stenotrophomonas maltophilia and was defined as an oleate hydratase. In the presence of this enzyme, 1 10-hydroxystearic acid was obtained with 49 g L− in 4 h, corresponding to a conversion of 98% [37]. The same group also reported a recombinant enzyme from Lysinibacillus fusiformis (organism isolated from soil) [47], which was isolated and defined as an oleate hydratase. Under optimized reaction 1 conditions, 40 g L− (133 mM) of the desired product 10-hydroxystearic acid was isolated, which corresponds to a yield of 94% [48]. Park et al. used the oleate hydratase from Stenotrophomonas maltophilia and recombinantly expressed the ohyA gene in E. coli. This biocatalyst was able to produce 1 10-hydroxystearic acid with a final concentration of 46 g L− in the culture medium [49]. Subsequently, Oh et al. described a process with an oleate hydratase from Lysinibacillus fusiformis, which converted 1 ricinoleic acid to 10,12-dihydroxystearic acid, which was obtained with 13.5 g L− [50]. Lactobacillus species are mostly found in dairy products, such as fermented milks and cheeses [51]. Additionally, Ogawa et al. classified genes that can be found in the gut bacterium Lactobacillus plantarum, encoding for enzymes, which are involved in the metabolism of polyunsaturated fatty acids [52]. Moreover, the LAH (Lactobacillus acidophilus hydratase) was described as a hydratase [53], whereas, in the same year, the crystal structure of this enzyme in the apo form was determined, showing a structural similarity to the 10,12-CLA-producing fatty acid double-bond isomerase from Propionibacterium acnes (PAI) and an amine oxidoreductase from Calloselasma rhodostoma. This crystallization gave an insight into the protein, which consist of two identical monomers, whereas an additional domain at the C-terminus acts as a lid, which covers the entrance of the substrate-binding channel. This lid probably opens through the recognition of the carboxylate residue on the surface of the enzyme. The opening of one channel led to the opening of the other protomer [54]. In 2014, Oh et al. described a fatty acid hydratase 1 1 from Stenotrophomonas nitritireducens, which converted 5.3 g L− linoleic acid and 0.93 g L− oleic acid [55]. The same group also utilized a linoleate 13-hydratase from Lactobacillus acidophilus to convert 1 linoleic acid to hydroxy fatty acid via the use of recombinant whole cells, which produced 79 g L− within 3 h [56]. In 2015, Ogawa et al. described a further gut bacterium (Lactobacillus acidophilus NTV001), which expresses a gene called FA-HY1 that is suitable to convert C18, as well as C20 and C22 unsaturated fatty acids [57]. Additionally, Oh et al. identified a double-bond hydratase that produces 13(S)-hydroxy-9(Z)-octadecenoic acid with high stereoselectivity [12]. Another milestone was the clarification of the crystal structure of Elizabethkingia meningoseptica, which also shows FAD as a bound cofactor. These findings gave a deep insight into the process of enzymatic water addition to C=C double bonds. The postulated mechanism starts with the protonation of the double bond by the means of a tyrosine. This positive charge is on the one hand stabilized by FAD and on the other hand by the means of an asparagine and a phenylalanine, which are localized at the C-terminus of a α-helix. A water-molecule, which is activated by a glutamate, then is able to attack on the partially charged double bond [25]. The production of a hydroxylated fatty acid at very high substrate loading could be achieved with the fatty acid hydratase of Lactobacillus plantarum, which was overexpressed in E. coli and used for the conversion of 280 g L-1 linoleic acid into (S)-10-hydoxy-cis-12-octadecenoic acid (HYA), leading to an enantiomeric excess (ee) of > 99% [58]. In addition, Oh et al. described an oleate hydratase from Stenotrophomonas nitritireducens, which converts plant oils to 10-hydroxy fatty acids. Hauer et al. summarized, in the Hydratase Engineering Database (HyED) 2046, putative oleate hydratases from Catalysts 2019, 9, x FOR PEER REVIEW 5 of 12

Stenotrophomona 89; 88 5.0, 0.85 102, 22 7.5 g/L s nitritireducens (exp. in E. coli) 10-hydroxy-12(Z)- [55] octadecenoic acid

10-hydroxysteric acid

Lactobacillus 79 79.0 631 100 g/L acidophilus (exp.

in E. coli) [56] 13-hydroxy-9(Z)-octadecenoic acid Catalysts 2020, 10, 287 4 of 13 Lactobacillus 98 280.0 552 90 g/L plantarum (exp. elevenin E. homologouscoli) families [59]. Moreover, Oh et al. used a 10-hydratase from Stenotrophomonas [58] (S)-10-hydoxy-cis-12-octadecenoic acid maltophilia DSM 29579 and 13-hydratase from Lactobacillus acidophilus ATCC 4796 for the conversion of 1 31.7Stenotrophomona and 15.6 g L− of unsaturated fatty acids in OA-rich oil65, [60 81]. All 21.7,hydroxy 13.3 fatty acids, - which 50 mL have beens maltophilia, synthesized from fatty acids at substrate loadings that are on a multi gram per liter scale,(reaction are listed Lactobacillus volume) in Table1, together with further10-monohydroxy information fatty onacids experimental data. Additionally, all characterized oleateacidophilus hydratases (exp. are categorized in two groups, namely OhyA1 or OhyA2. This categorization is basedin E. on coli the) [60] fourth conserved amino acid of the ﬂavin adenine dinucleotide (FAD)-binding motif [61]. Moreover, it is possible to use a plant oil as a cheap and renewable [62] starting material in combination 7,8-dihydroxy fatty acids with a hydroxy fatty acid hydratase and a lipase.

Furthermore, several examples of the synthesis of hydroxy fatty acids, starting from the cleavage Furthermore, several examples of the synthesis of hydroxy fatty acids, starting from the cleavage of the oils (Figure2), have been demonstrated, and in the following, some selected examples are given. of the oils (Figure 2), have been demonstrated, and in the following, some selected examples are Kim et al. used a lipase from Candida rugosa to hydrolyze olive and soybean oil, and the resulting given. Kim et al. used a lipase from Candida rugosa to hydrolyze olive and soybean oil, and the fatty acids were then further converted by means of the Flavobacterium sp. strain DS5, which was used resulting fatty acids were then further converted by means of the Flavobacterium sp. strain DS5, which to oxygenate fatty acids, thus leading to 10-ketostearic acid as major product and a minor amount was used to oxygenate fatty acids, thus leading to 10-ketostearic acid as major product and a minor of 10-hydroxystearic acid when starting from olive oil [63]. Candida rugosa can also be used for the amount of 10-hydroxystearic acid when starting from olive oil [63]. Candida rugosa can also be used hydrolysis of saﬄower oil [43], as well as perilla seed oil [55,64] and tree oils [65]. The lipase from for the hydrolysis of safflower oil [43], as well as perilla seed oil [55,64] and tree oils [65]. The lipase Thermomyces lanuginosus is able to hydrolyze olive and soybean oil [66]. from Thermomyces lanuginosus is able to hydrolyze olive and soybean oil [66].

Figure 2. Enzymatic hydrolysis of oils, thus releasing fatty acids and glycerol. Figure 2. Enzymatic hydrolysis of oils, thus releasing fatty acids and glycerol.

Catalysts 2019, 9, x FOR PEER REVIEW 4 of 12 CatalystsCatalysts 20192019,,, 99,,, xxx FORFORFOR PEERPEERPEER REVIEWREVIEWREVIEW 44 ofof 1212 are listed Catalystsin Table 2019 1,, 9together, x FOR PEER with REVIEW further information on experimental data. Additionally, all 4 of 12 areare listedlisted inin TableTable 1,1, togethertogether withwith furtherfurther informationinformation onon experimentalexperimental data.data. Additionally,Additionally, allall characterized oleate hydratases are categorized in two groups, namely OhyA1 or OhyA2. This characterizedcharacterizedare oleateoleatelisted hydratasesinhydratases Table 1, arearetogether categorizedcategorized with further inin twotwo information groups,groups, namelynamely on experimental OhyA1OhyA1 oror OhyA2. OhyA2.data. Additionally, ThisThis all categorization is based on the fourth conserved aminoino acidacid ofof thethe flavinflavin adenineadenine dinucleotidedinucleotide (FAD)-(FAD)- categorizationcategorizationcharacterized isis basedbased onon oleate thethe fourthfourth hydratases conservedconserved are amamcategorizedinoino acidacid ofof in thethe two flavinflavin groups, adenineadenine namely dinucleotidedinucleotide OhyA1 (FAD)-(FAD)- or OhyA2. This binding motif [61].. Moreover,Moreover, itit isis possiblepossible toto useuse aa plantplant oiloil asas aa cheapcheap andand renewablerenewable [62][62] startingstarting bindingbinding motifmotifcategorization [61][61].. Moreover,Moreover, is based itit isis on possiblepossible the fourth toto useuse conserved aa plantplant oilamoil asinoas aa acid cheapcheap of theandand flavin renewablerenewable adenine [62][62] dinucleotide startingstarting (FAD)- material in combination with a hydroxy fatty acid hydratase and a lipase. materialmaterial ininbinding combinationcombination motif with with[61] . aaMoreover, hydroxyhydroxy fatty fattyit is acidpossibleacid hydratasehydratase to use andanda plant aa lipase.lipase. oil as a cheap and renewable [62] starting Table material1. Summary in combinationof fatty acid hydratases with a hydroxy and resulting fatty acidproducts hydratase synthesized and a on lipase. multi gram-scale. TableTable 1.1. SummarySummary ofof fattyfatty acidacid hydrataseshydratases andand resultingresulting productsproducts synthesizedsynthesized onon multimulti gram-scale.gram-scale. Catalysts 2020, 10, 287 Exp. in E. coli: expressed in E. coli; —: no information was given. 5 of 13 Exp.Exp. inin E.E. colicoli::: expressedexpressed inin E.E. colicoli;;; —:—:—: nonono informationinformationinformation waswaswas given.given.given. Exp. in E. coliTable: expressed 1. Summary in E. coli of ;fatty —: no acid information hydratases was and given. resulting products synthesized on multi gram-scale. Exp. in E. coli: expressed in E. coli; —: no information was given. Table 1. Summary of fatty acid hydratases and resulting products synthesized on multi gram-scale. Exp. in E. coli: expressed in E. coli; —: no information was given.

Strain (Source Product Conv. Product- PROD Substrate StrainStrain (Source(Source ProductProProductduct Conv.Conv. Product-Product- PRODPROD SubstrateSubstrate of Fatty Acid /% Amount/g UCTIV Loading ofof FattyFatty AcidAcid /%/% Amount/gAmount/g UCTIVUCTIV LoadingLoading Strain (Source of Fatty Acid Product Conv./%−1 Product-Amount/ PRODUCTIVITY/ Substrate Hydratase)Strain (Source Product Conv.L−−11 Product-ITY/g PROD Substrate Hydratase)Hydratase) L−−11 ITY/gITY/g 1 1 1 Hydratase) LL −1 g−1 L− g L− h− Loading of Fatty Acid /% Amount/gL−−11 h−−11 UCTIV Loading LL−−11 hhh−−11 −1 −1 Candida tropicalisHydratase) - 19.4 L 0.8 70ITY/g mL d −−11 CandidaCandida tropicalistropicalis -- 19.4 19.4 0.8 0.8 70 70 mLmL dd−−11 DSM 3152 (wild- L−1 h−1 DSMDSM 31523152 (wild-(wild- Candida tropicalis Candida tropicalis - 19.4 0.8 70 mL d−1 1 DSMtype) 3152 [38] 9 - 19.4 0.8 70 mL d− type)type) [38][38] 3-hydroxy-Δ99-cis-1,18- (wild-type) [38] [38]DSM 3152 (wild- 3-hydroxy-3-hydroxy-Δ∆Δ99---cisciscis-1,18--1,18--1,18- octadecenedioic acid type) [38] octadecenedioicoctadecenedioic acid acid Stenotrophomona 3-hydroxy-Δ9-cis-1,18- - 31.5 7.9 15 g/L StenotrophomonaStenotrophomona -- 31.5 31.5 7.9 7.9 1515 g/Lg/L s nitritireducens octadecenedioic acid ss nitritireducensnitritireducens Stenotrophomona - 31.5 7.9 15 g/L Stenotrophomonas(wild-type)(wild-type) [46] - 31.5 7.9 15 g/L (wild-type)(wild-type) [46][46] 10-hydroxystearic acid nitritireducens (wild-type) [46s] nitritireducens 10-hydroxystearic10-hydroxystearic acid acid

Stenotrophomona(wild-type) 98 49.0 12.3 50 g/L StenotrophomonaStenotrophomona [46] 10-hydroxystearic acid 9898 49.049.0 12.312.3 5050 g/Lg/L ss maltophiliamaltophilia ss maltophiliamaltophilia Stenotrophomonas(exp.(exp. maltophilia inin E. coliStenotrophomona)) 9898 49.0 49.012.3 50 g/L 12.3 50 g/L (exp.(exp. inin E.E. colicoli)) 10-hydroxystearic acid (exp. in E. coli)[37[37]] s maltophilia 10-hydroxystearic10-hydroxystearic acid acid [37][37] Lysinibacillus(exp. in E. coli) 94 40.0 384 40 g/L LysinibacillusLysinibacillus 10-hydroxystearic acid 9494 40.040.0 384 384 4040 g/Lg/L fusiformisfusiformis (exp.(exp. [37] fusiformisfusiformis (exp.(exp. Lysinibacillus fusiformisin E. coli) [48]Lysinibacillus 9494 40.0 40.0 384 40 g/L 384 40 g/L ininin E.E. colicoli)) [48][48] 10-hydroxystearic acid (exp. in E. coli)[48] [48]fusiformis (exp. 10-hydroxystearic10-hydroxystearic acid acid

Stenotrophomonain E. coli) [48] 91 46.0 197 50 g/L StenotrophomonaStenotrophomona 10-hydroxystearic acid 9191 46.046.0 197 197 5050 g/Lg/L s maltophilia ss maltophiliamaltophilia Stenotrophomonas maltophiliaStenotrophomona 9191 46.0 46.0 197 50 g/L 197 50 g/L (exp.(exp. inin E. coli) (exp. in E. coli(exp.(exp.)[49 inin] E.E. colicoli)) 10-hydroxystearic acidacid [49] s maltophilia 10-hydroxystearic10-hydroxystearic acidacid [49][49] [49][49] (exp. in E. coli) Lysinibacillus 10-hydroxystearic acid 90 13.5 108 15 g/L LysinibacillusLysinibacillus [49] 9090 13.513.5 108 108 1515 g/Lg/L fusiformisfusiformis (exp.(exp. fusiformisfusiformis (exp.(exp. in E. coli) [50]Lysinibacillus 90 13.5 108 15 g/L ininin E.E. colicoli)) [50][50] 10,12-dihydroxystearic acid [50]fusiformis (exp.10,12-dihydroxystearic10,12-dihydroxystearic acidacid

in E. coli) [50] 10,12-dihydroxystearic acid

Catalysts 2019, 9, x FOR PEER REVIEW 4 of 12

are listed in Table 1, together with further information on experimental data. Additionally, all characterized oleate hydratases are categorized in two groups, namely OhyA1 or OhyA2. This categorization is based on the fourth conserved amino acid of the flavin adenine dinucleotide (FAD)- binding motif [61]. Moreover, it is possible to use a plant oil as a cheap and renewable [62] starting material in combination with a hydroxy fatty acid hydratase and a lipase.

Table 1. Summary of fatty acid hydratases and resulting products synthesized on multi gram-scale. Exp. in E. coli: expressed in E. coli; —: no information was given.

Strain (Source Product Conv. Product- PROD Substrate of Fatty Acid /% Amount/g UCTIV Loading Hydratase) L−1 ITY/g L−1 h−1 Candida tropicalis - 19.4 0.8 70 mL d−1 DSM 3152 (wild-

type) [38] 3-hydroxy-Δ9-cis-1,18- octadecenedioic acid Stenotrophomona - 31.5 7.9 15 g/L s nitritireducens

(wild-type) [46] 10-hydroxystearic acid Stenotrophomona 98 49.0 12.3 50 g/L s maltophilia (exp. in E. coli) 10-hydroxystearic acid [37] Lysinibacillus 94 40.0 384 40 g/L fusiformis (exp.

Catalysts 2020, 10, 287 in E. coli) 6 of 13 [48] 10-hydroxystearic acid

Stenotrophomona 91 46.0 197 50 g/L Table 1. Cont. s maltophilia

Strain (Source of(exp. Fatty in Acid E. coli) Product Conv./% Product-Amount/ PRODUCTIVITY/ Substrate 10-hydroxystearic acid 1 1 1 Hydratase) [49] g L− g L− h− Loading Catalysts 2019, 9, x FOR PEER REVIEW 5 of 12 CatalystsLysinibacillus 2019, 9, x FOR PEER REVIEW 90 13.5 108 15 g/L5 of 12 Catalysts 2019, 9, x FOR PEER REVIEW 5 of 12 Catalystsfusiformis 2019 ,(exp. 9, x FOR PEER REVIEW 5 of 12 LysinibacillusStenotrophomona fusiformis 89; 88 90 5.0, 0.85 102, 22 13.5 7.5 g/L 108 15 g/L CatalystsStenotrophomonain E. 2019coli), [50]9, x FOR PEER REVIEW 89; 88 5.0, 0.85 102, 22 7.5 g/L5 of 12 (exp. in E. coliStenotrophomonaSteno)[50trophomona] 10,12-dihydroxystearic10,12-dihydroxystearic acidacid 89; 88 5.0, 0.85 102, 22 7.5 g/L Stenotrophomonas nitritireducens 89; 88 5.0, 0.85 102, 22 7.5 g/L s nitritireducens Stenotrophomonas(exp. nitritireducens in E. coli) 89; 88 5.0, 0.85 102, 22 7.5 g/L s(exp. nitritireducens in E. coli) 10-hydroxy-12(Z)- s(exp. nitritireducens in[55] E. coli) 10-hydroxy-12(Z)- (exp. [55]in E. coli) 10-hydroxy-12(octadecenoic acidZ)- Stenotrophomonas(exp. [55]in E. coli) 10-hydroxy-12(octadecenoic acidZ)- 89; 88 5.0, 0.85 102, 22 7.5 g/L [55] 10-hydroxy-12(octadecenoic acidZ)- nitritireducens (exp. in E. coli) 10-hydroxy-12(octadecenoic acidZ)- [55] [55] octadecenoic acid

10-hydroxysteric acid 10-hydroxysteric acid 10-hydroxysteric acid 10-hydroxysteric acid Lactobacillus 10-hydroxysteric acid 79 79.0 631 100 g/L Lactobacillus 10-hydroxysteric acid 79 79.0 631 100 g/L Lactobacillus 79 79.0 631 100 g/L acidophilusLactobacillus (exp. 79 79.0 631 100 g/L acidophilus (exp. acidophilusinLactobacillus E. coli) (exp.[56] 79 79.0 631 100 g/L Lactobacillus acidophilusacidophilusin E. coli) [56](exp. 13-hydroxy-9(Z)-octadecenoic acid 79 79.0 631 100 g/L in E. coli) [56] 13-hydroxy-9(Z)-octadecenoic acid acidophilusin E. coli) [56](exp. 13-hydroxy-9(Z)-octadecenoic acid (exp. in E. coli)[56] 13-hydroxy-9(13-hydroxy-9(ZZ)-octadecenoic)-octadecenoic acidacid inLactobacillus E. coli) [56] 98 280.0 552 90 g/L Lactobacillus 13-hydroxy-9(Z)-octadecenoic acid 98 280.0 552 90 g/L Lactobacillus 98 280.0 552 90 g/L plantarumLactobacillus (exp. 98 280.0 552 90 g/L plantarum (exp. Lactobacillus plantarumplantaruminLactobacillus E. coli) (exp.[58] 98 280.0 552 90 g/L plantarumin E. (exp.coli) [58](exp. in (S)-10-hydoxy-cis-12-octadecenoic acid 98 280.0 552 90 g/L in E. coli) [58] (S)-10-hydoxy-cis-12-octadecenoic acid E. coli)[plantarum58in] E. coli) (exp.[58] (S)-10-hydoxy-cis-12-octadecenoic acidacid (S)-10-hydoxy-cis-12-octadecenoic acid Stenotrophomonain E. coli) [58] 65, 81 21.7, 13.3 - 50 mL Stenotrophomona (S)-10-hydoxy-cis-12-octadecenoic acid 65, 81 21.7, 13.3 - 50 mL Stenotrophomona 65, 81 21.7, 13.3 - 50 mL Stenotrophomonas maltophilia, 65, 81 21.7, 13.3 - (reaction 50 mL s maltophilia, (reaction StenotrophomonasStenotrophomonas maltophiliaLactobacillus maltophilia,, 10-monohydroxy fatty acids 65, 81 65, 21.7, 81 13.3 21.7,- 13.3(reactionvolume) 50 mL - 50 mL Lactobacilluss maltophilia, 10-monohydroxy fatty acids (reactionvolume) Lactobacillus acidophilusacidophilussLactobacillus maltophilia, (exp. 10-monohydroxy fatty acids (reactionvolume) (reaction volume) acidophilusLactobacillus (exp. 10-monohydroxy fatty acids volume) (exp. in E. coliacidophilus)[60] (exp. 10-monohydroxy fatty acids acidophilusinLactobacillus E. coli) [60](exp. volume) in E. coli) [60] 10-monohydroxy fatty acids acidophilusin E. coli) [60](exp. in E. coli) [60] 7,8-dihydroxy fatty acids in E. coli) [60] 7,8-dihydroxy fatty acids 7,8-dihydroxy fatty acids 7,8-dihydroxy fatty acids 7,8-dihydroxy fatty acids Furthermore, several examples7,8-dihydroxy of the fatty synthesis acids of hydroxy fatty acids, starting from the cleavage Furthermore,Furthermore, several examples of the synthesis of hydroxy fatty acids, starting from the cleavage of theFurthermore, oils (Figure several 2), have examples been demonstrated, of the synthesis and of inhydroxy the following, fatty acids, some starting selected from examples the cleavage are of the oils (Figure 2), have been demonstrated, and in the following, some selected examples are ofgiven. theFurthermore, oilsKim (Figure et al. usedseveral2), have a exampleslipase been fromdemonstrated, of theCandida synthesis rugosa and of inhydroxyto thehydrolyze following, fatty acids,olive some startingand selected soybean from examples theoil, cleavage and arethe given.of the oilsKim (Figure et al. used 2), have a lipase been from demonstrated, Candida rugosa and into thehydrolyze following, olive some and selected soybean examples oil, and theare given.ofresulting the oilsKim fatty (Figure et acidsal. used 2), were have a thenlipase been further from demonstrated, convertedCandida rugosa byand means intoto thehydrolyzehydrolyze of following, the Flavobacterium oliveolive some andand selected soybeansoybean sp. strain examples oil,oil, DS5, andand which thearethe resulting fatty acids were then further converted by means of the Flavobacterium sp. strain DS5, which resultinggiven.was used Kim fatty to etoxygenate acidsal. used were fattya thenlipase acids, further from thus convertedCandida leading rugosa toby 10-ketostearic means to hydrolyze of the Flavobacterium acid olive as major and soybean productsp. strainstrain andoil, DS5,DS5, anda whichwhichminor the was used to oxygenate fatty acids, thus leading to 10-ketostearic acid as major product and a minor wasresultingamount used of fattyto 10-hydroxystearic oxygenate acids were fatty then acids, acid further whthus enconverted leading starting to by from 10-ketostearic10-ketostearic means olive of oil the [63]. Flavobacteriumacidacid Candida asas majormajor rugosa productproductsp. strain can andandalso DS5, aabe whichminorminor used amount of 10-hydroxystearic acid when starting from olive oil [63]. Candida rugosa can also be used amountwasfor the used hydrolysis of to 10-hydroxystearic oxygenate of safflower fatty acids, acid oil [43], whthusen as leadingstarting well as to fromperilla 10-ketostearic olive seed oil oil [63]. [55,64]acid Candida as andmajor treerugosa product oils cancan [65]. alsoandalso The abebe minorlipase usedused for the hydrolysis of safflower oil [43], as well as perilla seed oil [55,64] and tree oils [65]. The lipase foramountforfrom thethe Thermomyces hydrolysishydrolysis of 10-hydroxystearic ofoflanuginosus safflowersafflower acid isoiloil able [43],[43],wh toen asas hydrolyze starting wellwell asas fromperillaperilla olive olive andseedseed oilsoybean oiloil [63]. [55,64][55,64] Candida oil andand [66]. treetreerugosa oilsoils can [65].[65]. also TheThe be lipaselipase used from Thermomyces lanuginosus is able to hydrolyze olive and soybean oil [66]. fromforfrom the Thermomyces hydrolysis oflanuginosus safflower is isoil ableable [43], toto as hydrolyzehydrolyze well as perilla oliveolive andandseed soybeansoybean oil [55,64] oiloil and [66].[66]. tree oils [65]. The lipase from Thermomyces lanuginosus is able to hydrolyze olive and soybean oil [66].

Figure 2. Enzymatic hydrolysis of oils, thus releasing fatty acids and glycerol. Figure 2. Enzymatic hydrolysis of oils, thus releasing fatty acids and glycerol. Figure 2. Enzymatic hydrolysis of oils, thus releasing fatty acids and glycerol. Figure 2. Enzymatic hydrolysis of oils, thus releasing fatty acids and glycerol. 3. Cascade ProcessesFigure 2.Involving Enzymatic Fatty hydrolysis Acid Hydrataseof oils, thus releasing fatty acids and glycerol. 3. Cascade Processes Involving Fatty Acid Hydratase 3. CascadeFrom theProcesses year 2013 Involving on, a range Fatty of enzymaticAcid Hydratase cascades combining fatty acid hydratases with other 3. CascadeFrom the Processes year 2013 Involving on, a range Fatty of enzymaticAcid Hydratase cascades combining fatty acid hydratases with other enzymesFrom have the year been 2013 reported on, a rangein the ofliterature. enzymatic In cascadessuch cascades, combining fatty acidfatty hydratases, acid hydratases in combination with other enzymes have been reported in the literature. In such cascades, fatty acid hydratases, in combination enzymeswithFrom other have the enzymes, yearbeen 2013 reported were on, aused inrange the to literature.ofbuild-up enzymatic Ina broad sucascadesch cascades, range combining of fattyindustrially acid fatty hydratases, acid relevant hydratases inproducts. combination with otherSuch withenzymes other have enzymes, been reported were used in the to literature.build-up aIn broad such cascades,range of industriallyfatty acid hydratases, relevant products.in combination Such withenzymescascades other havecombining enzymes, been reported variouswere used inenzymatic the to literature.build-up steps a Inare broad su ofch interest cascades,range ofalso fattyindustrially from acid an hydratases, industrial relevant products.perspectivesin combination Such as cascades combining various enzymatic steps are of interest also from an industrial perspectives as cascadeswithwork-steps other combining enzymes,of intermediates various were used canenzymatic beto avoided,build-up steps thus aare broad ofenabling interest range a ofalsobetter industrially from process an industrial economyrelevant perspectivesproducts.with decreased Such as work-steps of intermediates can be avoided, thus enabling a better process economy with decreased work-stepscascadessolvent consumption combining of intermediates various and solvent canenzymatic be avoided,waste steps formation. thus are enablingofenabling interest A key aa better alsobetterprerequisite from processprocess an industrial (andeconomyeconomy at the perspectiveswithwith same decreaseddecreased time as a solvent consumption and solvent waste formation. A key prerequisite (and at the same time a solventwork-stepschallenge) consumption forof intermediatesrealizing and such solvent can cascades be avoided,waste is toformation. acthushieve enabling compatibilityA key a betterprerequisite processbetween (and economy the atdifferent the with same enzymaticdecreased time a challenge) for realizing such cascades is to achieve compatibility between the different enzymatic challenge)solventreaction consumptionsteps. for realizing and such solvent cascades waste is toformation. achieve compatibility A key prerequisite between (and the atdifferent the same enzymatic time a reaction steps. reactionchallenge) steps. for realizing such cascades is to achieve compatibility between the different enzymatic reaction steps.

Catalysts 2020, 10, 287 7 of 13

3. Cascade Processes Involving Fatty Acid Hydratase From the year 2013 on, a range of enzymatic cascades combining fatty acid hydratases with other enzymes have been reported in the literature. In such cascades, fatty acid hydratases, in combination with other enzymes, were used to build-up a broad range of industrially relevant products. Such cascades combining various enzymatic steps are of interest also from an industrial perspectives as work-steps of intermediates can be avoided, thus enabling a better process economy with decreased Catalysts 2019, 9, x FOR PEER REVIEW 6 of 12 solvent consumption and solvent waste formation. A key prerequisite (and at the same time a challenge) for realizingExamples such of cascades the usage is of to fatty achieve acid compatibility hydratases in between biocatalytic the dicascadefferent processes enzymatic for reaction the synthesis steps. of long-chainExamples α of,ω the-dicarboxylic usage of fatty acids acid and hydratases ω-hydroxycarboxylic in biocatalytic acids cascade are summarized processes for in the Figure synthesis 3 and ofare long-chain describedα ,inω -dicarboxylicthe following. acids Park and et al.ω -hydroxycarboxylicdeveloped a cascade acids starting are summarized with a hydratase in Figure catalyzing3 and arethe described hydration in theof the following. fatty acid Park double et al. developedbond. Afterwards, a cascade the starting alcohol with is a oxidized hydratase by catalyzing means of the an hydrationalcohol dehydrogenase of the fatty acid (ADH) double and bond. a Baeyer-Villiger Afterwards, the monooxygenase alcohol is oxidized (BVMO). by means In the offinal an alcoholstep, the dehydrogenaseester is then hydrolysed (ADH) and towards a Baeyer-Villiger α,ω-dicarboxylic monooxygenase acids and (BVMO).ω-hydroxycarboxylic In the final step, acids the (Figure ester is4). thenAll these hydrolysed enzymes towards were combinedα,ω-dicarboxylic in one whole acids andcell, ωand-hydroxycarboxylic the process was conducted acids (Figure without4). All addition these enzymesof an external were combined cofactor. inSuch one products whole cell, are and of theparticular process interest was conducted for applications without additionin the field of an of externalpolymers cofactor. as these Such bifunctional products molecules are of particular can serv intereste as monomers for applications for the insynthesis the field of of corresponding polymers as thesepolyesters bifunctional [67]. molecules can serve as monomers for the synthesis of corresponding polyesters [67].

O R HO n m

O O O O NH2 OH HO HO OH HO n m n m n m polymer area, lubricants, polymer area, lubricants, polymer area plasticizers, perfumes plasticizers, perfumes

O O O O OH HO NH HO OH HO n m 2 n m n m polymer area flavors, resins, waxes, nylons, flavors, resins, waxes, nylons, plastics, lubricants, polymers plastics, lubricants, polymers

O O O HO n m O

flavors, resins, waxes, nylons, flavorin plastics, lubricants, polymers

FigureFigure 3. 3.Collection Collection of of di differentﬀerent molecules molecules that that were were synthesized synthesized in in a a cascade cascade that that involve involve fatty fatty acid acid ω α ω hydratases.hydratases. Listed Listed are are ω-hydroxycarboxylic-hydroxycarboxylic acids acids [ 66[66]] ,α,ω-dicarboxylic-dicarboxylic acids acids [ 68[68],], aminocarboxylic aminocarboxylic acidsacids [68 [68,69],,69], oxostearic oxostearic acid acid [70 [70]] and and lactones lactones [2 [2].].

Catalysts 2020, 10, 287 8 of 13 Catalysts 2019, 9, x FOR PEER REVIEW 7 of 12

Figure 4. Chosen examples for the enzymatic cascade reactions involving fatty acid hydratases. Figure 4. Chosen examples for the enzymatic cascade reactions involving fatty acid hydratases. Shown are the synthesis of ω-hydroxycarboxylic acid [67], ω-aminocarboxylic acids [68,69] and Shown are the synthesis of ω-hydroxycarboxylic acid [67], ω-aminocarboxylic acids [68,69] and γ-lactone [2]. NAD/NADH (Nicotinamide adenine dinucleotide), ADH (Alcohol dehydrogenase), γ-lactone [2]. NAD/NADH (Nicotinamide adenine dinucleotide), ADH (Alcohol dehydrogenase), BVMO (Bayer-Villiger monooxygenase), TA (Transaminase). BVMO (Bayer-Villiger monooxygenase), TA (Transaminase).

BesidesBesides being being interesting interesting building building blocks blocks as as startingstarting materialsmaterials in in the the polymer polymer field, field, these these products productscan be can also be utilizedalso utilized for the for preparation the preparation of lubricants, of lubricants, plasticizers plasticizers or perfumes or perfumes [70]. [70]. This This cascade cascadeprocess process was was then then further further expanded expanded by couplingby coupling the the cascade cascade with with an alcoholan alcohol dehydrogenase, dehydrogenase, which whichoxidized oxidized the theω -hydroxycarboxylicω-hydroxycarboxylic acid acid towards towards the the corresponding corresponding diacid diacid or or the the aldehyde aldehyde [[67].67]. The The latter oneone isis subsequentlysubsequently transformed transformedvia viaa a transaminase transaminase towards towards the the primary primary amine amine (Figure (Figure4)[ 4)67 ]. It ω [67].is It noteworthyis noteworthy that, that, by by means means of thisof this concept, concept, fatty fatty acids acids can can be convertedbe converted into intoω-substituted -substituted amino aminoacids, acids, which which are are of interestof interest as monomersas monomers for for polyamides. polyamides. Furthermore,Furthermore, this thiscascade cascade was wascombined combined with with a double a double bond bond hydratase hydratase to obtain to obtain unsaturated unsaturated α,ω-dicarboxylicα,ω-dicarboxylic acids acids and andω-hydroxycarboxylicω-hydroxycarboxylic acids acids [71,72]. [71 ,In72 ].addition, In addition, a related a related cascade cascade system system usingusing a whole a whole cell cellcatalyst, catalyst, which which overexpresses overexpresses a fatty a fatty acid aciddouble double bond bond hydratase hydratase (OhyA) (OhyA) of of StenotrophomonasStenotrophomonas maltophilia maltophilia, the, ADH the ADH of Micrococcus of Micrococcus luteus luteus, and, andthe engineered the engineered Baeyer-Villiger Baeyer-Villiger monooxygenasemonooxygenase (E6BVMO) (E6BVMO) of Pseudomonas of Pseudomonas putida putida KT2440,KT2440, was wasdeveloped. developed. Furthermore, Furthermore, the long the long chainchain fatty fatty acid acidtransporter transporter FadL FadL was wasoverexpressed, overexpressed, which which transports transports the fatty the fatty acids acids into intothe whole the whole cell. cell.This Thisoverexpression overexpression further further contributes contributes to th toe improvement the improvement of the of whole-cell the whole-cell biotransformation. biotransformation. It shouldIt should be added be added that this that cascade this cascade starts starts and ends and with ends a with hydrolysis. a hydrolysis. In the initial In the step, initial a step,plant aoil plant is hydrolyzed,oil is hydrolyzed, and the andfinal the step final consists step consistsof the hydrolysis of the hydrolysis of the ester of theby the ester means by the of meansa lipase of from a lipase Thermomycesfrom Thermomyces lanuginosus lanuginosus. The ω-hydroxycarboxylic. The ω-hydroxycarboxylic acid is acidthen isfurther then further derivatized derivatized by means by means of of chemicalchemical [64] [ 64or] orenzymatic enzymatic oxidation, oxidation, and and the resultingresulting productproduct can can then then further further be usedbe used as a monomeras a monomerfor the for production the production of polyesters of polyesters and polyamides and polyamides [73]. This [73]. synthetic This synthetic method method was further was further extended extendedtowards towards a process a process based based on the on usethe ofuse a of whole a whole cell cell catalyst, catalyst, which which converts converts long long chain chain fatty fatty acids acidslike like ricinoleic ricinoleic acid, acid, in in combination combination withwith anan addedadded aldehydealdehyde dehydrogenase,dehydrogenase, in order to obtain obtain the the correspondingcorresponding ωω-hydroxycarboxylic-hydroxycarboxylic acid acid (Figure (Figure 44))[ [74].74]. In furtherIn further studies studies by Xu by et Xual., the et al., synthesis the synthesis of 10-oxostearic of 10-oxostearic acid was acidreported, was reported,utilizing a utilizing novel a oleate hydratase from Paracoccus aminophilus with a high specific activity of 5.21 U mg−1. In the1 novel oleate hydratase from Paracoccus aminophilus with a high specific activity of 5.21 U mg− . In hydration step, 90 g L−1 of hydrated1 oleic acid was obtained. The formed 10-hydroxystearic was the hydration step, 90 g L− of hydrated oleic acid was obtained. The formed 10-hydroxystearic convertedwas converted afterwards afterwards by an alcohol by an alcoholdehydrogenase dehydrogenase from Micrococcus from Micrococcus luteus, which luteus, was which used was in used combinationin combination with a lactate with a dehydrogen lactate dehydrogenasease for co-factor for co-factor recycling. recycling. After 10 h After of reaction 10 h reaction at a scale time of at a 1 L, scalea conversion of 1 L, aof conversion 95% was reached of 95% [69]. was This reached concept [69]. also This shows concept the alsohigh shows suitability the highof combining suitability of a lyase (exemplified by a hydratase) and redox enzymes (exemplified by alcohol and lactate dehydrogenases).

Catalysts 2020, 10, 287 9 of 13 combining a lyase (exemplified by a hydratase) and redox enzymes (exemplified by alcohol and lactate dehydrogenases). Another building block that is of interest for a different industrial product segment was synthesized by Oh et al. in 2014, which also utilize fatty acid hydratases. In detail, they produced γ-dodecalactone from safflower oil, which is widely used in perfume industry or enhancing aroma or taste [55]. The cascade starts with the hydrolysis of safflower oil by means of lipase from Candida rugosa producing free fatty oleic and linoleic acid. Afterwards, the hydration was catalyzed by the FAH from Stenotrophomonas nitritireducens, followed by lactonization in the presence of these Candida boidinii whole cells [57]. Furthermore, Park et al. very recently published a cascade for the production of aliphatic amines from renewable fatty acids, which are interesting for applications in the polymer area such as for the synthesis of carbamates and polyurethanes [67]. This concept is based on the combined use of three types of enzymes, namely a fatty acid hydratase, an alcohol dehydrogenase and a transaminases [68].

4. Conclusions The importance of fatty acid hydratases in biotechnology has been demonstrated by the many achievements of different groups, including the development of suitable biocatalysts, microorganisms or isolated enzymes overexpressed in E. coli. The biocatalysts were further used for the process development of fatty acid hydration in huge scale applications, producing industrially relevant hydroxy-substituted fatty acids from renewable raw materials, such as, e.g., vegetable oils as raw material. Nevertheless, high cell or enzyme amounts were used, as well as no elaborate isolation methods being presented, which is a bottleneck for further utilization. Moreover, the combination of fatty acid hydration with other enzymes in complex cascade processes was intensively studied for the further production of industrially valuable molecules. However, up to now, only little is known about the catalytic mechanism, which is due to the fact that only a limited number of crystal structures are available [25,54,75,76]. The advanced knowledge could contribute to the future improvement of productivity and the design of novel biotransformation pathways. Other confines are the substrate scope, which is limited by the seven carbon atoms between carboxylic residue and double bond, as well as the length of the fatty acids and the carboxylic acid itself. These problems have already been addressed in some publications, like the hydration of unactivated olefins by using short chain acids as decoy molecules [77], the rational of engineering a fatty acid hydratase to enhance the substrate scope [78] and the hydration of oleic acid derivatives without a free carboxylate moiety [79]. Thus, the current status represents a highly promising basis for further studies in this field with the perspective of a range of industrial applications for a sustainable manufacture of industrial chemicals for various commercial segments, based on the utilization of biorenewable raw materials.

Author Contributions: Conceptualization, J.L. and H.G.; writing—original draft preparation, J.L.; writing—review and editing, J.L. and H.G.; supervision, H.G. All authors have read and agreed to the published version of the manuscript. Funding: This work was funded by the U. Windmöller Innovation GmbH & Co. KG. Acknowledgments: J.L. and H.G. thank the U. Windmöller Innovation GmbH & Co. KG for financial support. Conflicts of Interest: The authors declare no conflict of interest.

References

1. Liu, C.; Liu, F.; Cai, J.; Xie, U.W.; Long, T.E.; Turner, S.R.; Lyons, A.; Gross, R.A. Polymers from Fatty Acids: Poly (Co-Hydroxyl Tetradecanoic Acid) Synthesis and Physico-Mechanical Studies. ACS Symp. Ser. 2012, 1105, 131–150. [CrossRef] 2. Swizdor,´ A.; Panek, A.; Milecka-Tronina, N.; Kołek, T. Biotransformations Utilizing β-Oxidation Cycle Reactions in the Synthesis of Natural Compounds and Medicines. Int. J. Mol. Sci. 2012, 13, 16514–16543. [CrossRef][PubMed] Catalysts 2020, 10, 287 10 of 13

3. Schneider, S.; Wubbolts, M.G.; Sanglard, D.; Witholt, B. Production of Chiral Hydroxy Long Chain Fatty Acids by Whole Cell Biocatalysis of Pentadecanoic Acid with an E. Coli Recombinant Containing Cytochrome P450BM-3 Monooxygenase. Tetrahedron Asymmetry 2002, 9, 2832–2844. [CrossRef] 4. Ashby, R.D.; Solaiman, D.K.Y.; Liu, C.K.; Strahan, G.; Latona, N. Sophorolipid-Derived Unsaturated and Epoxy Fatty Acid Estolides as Plasticizers for Poly (3-Hydroxybutyrate). Am. Oil Chem. Soc. 2016, 93, 347–358. [CrossRef] 5. Hu, J.; Jin, Z.; Chen, T.Y.; Polley, J.D.; Cunningham, M.F.; Gross, R.A. Anionic Polymerizable Surfactants from Biobased ω-Hydroxy Fatty Acids. Macromolecules 2014, 47, 113–120. [CrossRef] 6. Mutlu, H.; Meier, M.A.R. Castor Oil as a Renewable Resource for the Chemical Industry. Eur. J. Lipid Sci. Technol. 2010, 112, 10–30. [CrossRef] 7. Hou, C.T. Biotechnology for Fats and Oils: New Oxygenated Fatty Acids. New Biotechnol. 2009, 26, 2–10. [CrossRef] 8. Patel, V.R.; Dumancas, G.G.; Viswanath, L.C.K.; Maples, R.; Subong, B.J.J. Castor Oil: Properties, Uses, and Optimization of Processing Parameters in Commercial Production. Lipid Insights 2016, LPI-S40233. [CrossRef] 9. Knothe, G.; Weisleder, D.; Bagby, M.O.; Peterson, R.E. Hydroxy Fatty Acids through Hydroxylation of Oleic Acid with Selenium Dioxide/Tert.-Butylhydroperoxide. J. Am. Oil Chem. Soc. 1993, 70, 401–404. [CrossRef] 10. Mountanea, O.G.; Limnios, D.; Kokotou, M.G.; Bourboula, A.; Kokotos, G. Asymmetric Synthesis of Saturated Hydroxy Fatty Acids and Fatty Acid Esters of Hydroxy Fatty Acids. Eur. J. Org. Chem. 2019, 10, 2010–2019. [CrossRef] 11. Köckritz, A.; Martin, A. Oxidation of Unsaturated Fatty Acid Derivatives and Vegetable Oils. Eur. J. Lipid Sci. Technol. 2008, 110, 812–824. [CrossRef] 12. Kim, K.R.; Oh, H.J.; Park, C.S.; Hong, S.H.; Park, J.Y.; Oh, D.K. Unveiling of Novel Regio-Selective Fatty Acid Double Bond Hydratases from Lactobacillus Acidophilus Involved in the Selective Oxyfunctionalization of Mono- and Di-Hydroxy Fatty Acids. Biotechnol. Bioeng. 2015, 112, 2206–2213. [CrossRef][PubMed] 13. Van de Loo, F.; Broun, P.; Turner, S.; Somervillet, C. An oleate 12-hydroxylase from Ricinus communis L. is a fatty acyl desaturase homolog. Proc. Natl. Acad. Sci. USA 1995, 92, 6743–6747. [CrossRef][PubMed] 14. Brodkorb, D.; Gottschall, M.; Marmulla, R.; Lüddeke, F.; Harder, J. Linalool Dehydratase-Isomerase, a Bifunctional Enzyme in the Anaerobic Degradation of Monoterpenes. J. Biol. Chem. 2010, 285, 30436–30442. [CrossRef][PubMed] 15. Steiger, S.; Mazet, A.; Sandmann, G. Heterologous Expression, Purification, and Enzymatic Characterization of the Acyclic Carotenoid 1, 2-Hydratase from Rubrivivax Gelatinosus. Arch. Biochem. Biophys. 2003, 414, 51–58. [CrossRef] 16. Bornscheuer, U.T. Enzymes in Lipid Modification. Annu. Rev. Food Sci. Technol. 2018, 9, 116–127. [CrossRef] 17. Engleder, M.; Pichler, H. On the Current Role of Hydratases in Biocatalysis. Appl. Microbiol. Biotechnol. 2018, 102, 5841–5858. [CrossRef] 18. Wallen, L.L.; Benedict, R.G.; Jackson, W.R. The Microbiological Production from Oleic of 10-Hydroxystearic Acid from Oleic Acid. Arch. Biochem. Biophys. 1962, 99, 249–253. [CrossRef] 19. Kim, K.-R.; Oh, D.-K. Production of hydroxy fatty acids by microbial fatty acid-hydroxylation enzymes. Biotechnol. Adv. 2013, 31, 1473–1485. [CrossRef] 20. Wallen, L.L.; Davis, E.N.; Wu, Y.V.; Rohwedder, W.K. Stereospecific Hydration of Unsaturated Fatty Acids by Bacteria. Lipids 1971, 6, 745–750. [CrossRef] 21. Niehaus, W.G.; Schroepfer, G.J. The Reversible Hydration of Oleic Acid to 10D-Hydroxystearic Acid. Biochem. Biophys. Res. Commun. 1965, 21, 271–275. [CrossRef] 22. Volkov, A.; Liavonchanka, A.; Kamneva, O.; Fiedler, T.; Goebel, C.; Kreikemeyer, B.; Feussner, I. Myosin Cross-Reactive Antigen of Streptococcus Pyogenes M49 Encodes a Fatty Acid Double Bond Hydratase That Plays a Role in Oleic Acid Detoxification and Bacterial Virulence. J. Biol. Chem. 2010, 285, 10353–10361. [CrossRef][PubMed] 23. Chen, Y.Y.; Liang, N.Y.; Curtis, J.M.; Gänzle, M.G. Characterization of Linoleate 10-Hydratase of Lactobacillus plantarum and Novel Antifungal Metabolites. Front. Microbiol. 2016, 7, 1561. [CrossRef][PubMed] Catalysts 2020, 10, 287 11 of 13

24. Rosberg-Cody, E.; Liavonchanka, A.; Göbel, C.; Ross, R.P.; O’Sullivan, O.; Fitzgerald, G.F.; Feussner, I.; Stanton, C. Myosin-Cross-Reactive Antigen (MCRA) Protein from Biﬁdobacterium Breve Is a FAD-Dependent Fatty Acid Hydratase Which Has a Function in Stress Protection. BMC Biochem. 2011, 12, 1–12. [CrossRef] [PubMed] 25. Engleder, M.; Pavkov-Keller, T.; Emmerstorfer, A.; Hromic, A.; Schrempf, S.; Steinkellner, G.; Wriessnegger, T.; Leitner, E.; Strohmeier, G.A.; Kaluzna, I.; et al. Structure-Based Mechanism of Oleate Hydratase from Elizabethkingia Meningoseptica. ChemBioChem 2015, 16, 1730–1734. [CrossRef][PubMed] 26. Hiseni, A.; Arends, I.W.C.E.; Otten, L.G. New Cofactor-Independent Hydration Biocatalysts: Structural, Biochemical, and Biocatalytic Characteristics of Carotenoid and Oleate Hydratases. ChemCatChem 2015, 7, 29–37. [CrossRef] 27. Villeneuve, P.; Muderhwa, J.M.; Graille, J.; Haas, M.J. Customizing Lipases for Biocatalysis: A Survey of Chemical, Physical and Molecular Biological Approaches. J. Mol. Catal. B Enzym. 2000, 9, 113–148. [CrossRef] 28. Goldberg, K.; Schroer, K.; Lütz, S.; Liese, A. Biocatalytic Ketone Reduction—A Powerful Tool for the Production of Chiral Alcohols-Part II: Whole-Cell Reductions. Appl. Microbiol. Biotechnol. 2007, 76, 249–255. [CrossRef] 29. Breuer, M.; Ditrich, K.; Habicher, T.; Hauer, B.; Keßeler, M.; Stürmer, R.; Zelinski, T. Industrial Methods for the Production of Optically Active Intermediates. Angew. Chem. Int. Ed. 2004, 43, 788–824. [CrossRef] 30. Strohmeier, G.A.; Pichler, H.; May, O.; Gruber-Khadjawi, M. Application of Designed Enzymes in Organic Synthesis. Chem. Rev. 2011, 111, 4141–4164. [CrossRef] 31. Abdelraheem, E.M.M.; Busch, H.; Hanefeld, U.; Tonin, F. Biocatalysis explained: From pharmaceutical to bulk chemical production. React. Chem. Eng. 2019, 4, 1878–1894. [CrossRef] 32. Jemli, S.; Ayadi-Zouari, D.; Hlima, H.B.; Bejar, S. Biocatalysts: Application and engineering for industrial purposes. Crit. Rev. Biotechnol. 2016, 36, 246–258. [CrossRef][PubMed] 33. Liese, A.; Seelbach, K.; Wandrey, C. Industrial Biotransformations; Wiley-VCH: Weinheim, Germany, 2006. [CrossRef] 34. Yamada, H.; Nagasawa, T. Production of Useful Amides by Enzymatic Hydration of Nitriles. Agric. Chem. 1990, 613, 142–154. [CrossRef] 35. Miyazawa, T. Enzymatic Resolution of Amino Acids via Ester Hydrolysis. Amino Acids 1999, 16, 191–213. [CrossRef][PubMed] 36. Brugging, A.; Roos, E.C.; De Vroom, E. Penicillin Acylase in the Industrial Production of β-Lactam Antibiotics. Org. Process Res. Dev. 1998, 2, 128–133. [CrossRef] 37. Joo, Y.C.; Seo, E.S.; Kim, Y.S.; Kim, K.R.; Park, J.B.; Oh, D.K. Production of 10-Hydroxystearic Acid from Oleic Acid by Whole Cells of Recombinant Escherichia Coli Containing Oleate Hydratase from Stenotrophomonas Maltophilia. J. Biotechnol. 2012, 158, 17–23. [CrossRef] 38. Fabritius, D.; Schäfer, H.J.; Steinbüchel, A. Identiﬁcation and Production of 3-Hydroxy-∆9-Cis-1, 18-Octadecenedioic Acid by Mutants of Candida Tropicalis. Appl. Microbiol. Biotechnol. 1996, 45, 342–348. [CrossRef] 39. El-Sharkawy, S.H.; Yang, W.; Dostal, L.; Rosazza, J.P.N. Microbial Oxidation of Oleic Acid. Appl. Environ. Microbiol. 1992, 58, 2116–2122. [CrossRef] 40. Hudson, J.A.; MacKenzie, C.A.M.; Joblin, K.N. Conversion of Oleic Acid to 10-Hydroxystearic Acid by Two Species of Ruminal Bacteria. Appl. Microbiol. Biotechnol. 1995, 44, 1–6. [CrossRef] 41. Kaneshiro, T.; Kuo, T.M.; Nakamura, L.K. Conversion of Unsaturated Fatty Acids by Bacteria Isolated from Compost. Curr. Microbiol. 1999, 38, 250–255. [CrossRef] 42. Koritala, S.; Hou, C.T.; Hesseltine, C.W.; Bagby, M.O. Microbial Conversion of Oleic Acid to 10-Hydroxystearic Acid. Appl. Microbiol. Biotechnol. 1989, 32, 299–304. [CrossRef] 43. Shinha, T.; Ahuja, R. Bacteremia due to Elizab. Meningoseptica 2015, 2, 13–15. [CrossRef] 44. Bevers, L.E.; Pinkse, M.W.H.; Verhaert, P.D.E.M.; Hagen, W.R. Oleate Hydratase Catalyzes the Hydration of a Nonactivated Carbon-Carbon Bond. J. Bacteriol. 2009, 191, 5010–5012. [CrossRef] 45. Ryan, R.P.; Monchy, S.; Cardinale, M.; Taghavi, S.; Crossman, L.; Avison, M.B.; Berg, G.; van der Lelie, D.; Dow, J.M. The versatility and adaptation of bacteria from the genus Stenotrophomonas. Nat. Rev. Microbiol. 2019, 7, 514–525. [CrossRef] 46. Kim, B.N.; Yeom, S.J.; Oh, D.K. Conversion of Oleic Acid to 10-Hydroxystearic Acid by Whole Cells of Stenotrophomonas Nitritireducens. Biotechnol. Lett. 2011, 33, 993–997. [CrossRef] Catalysts 2020, 10, 287 12 of 13

47. Gallegos-Monterrosa, R.; Maróti, G.; Bálint, B.; Kovács, A.T. Draft Genome Sequence of the Soil Isolate Lysinibacillus fusiformis M5, a Potential Hypoxanthine Producer. Genome Announc. 2016, 4, e01272-16. [CrossRef] 48. Kim, B.N.; Joo, Y.C.; Kim, Y.S.; Kim, K.R.; Oh, D.K. Production of 10-Hydroxystearic Acid from Oleic Acid and Olive Oil Hydrolyzate by an Oleate Hydratase from Lysinibacillus fusiformis. Appl. Microbiol. Biotechnol. 2012, 95, 929–937. [CrossRef] 49. Jeon, E.Y.; Lee, J.H.; Yang, K.M.; Joo, Y.C.; Oh, D.K.; Park, J.B. Bioprocess Engineering to Produce 10- Hydroxystearic Acid from Oleic Acid by Recombinant Escherichia Coli Expressing the Oleate Hydratase Gene of Stenotrophomonas Maltophilia. Process Biochem. 2012, 47, 941–947. [CrossRef] 50. Seo, M.H.; Kim, K.R.; Oh, D.K. Production of a Novel Compound, 10, 12-Dihydroxystearic Acid from Ricinoleic Acid by an Oleate Hydratase from Lysinibacillus fusiformis. Appl. Microbiol. Biotechnol. 2013, 97, 8987–8995. [CrossRef] 51. De Angelis, M.; Gobbetti, M. Lactobacillus spp.: General Characteristics. Ref. Modul. Food Sci. 2016.[CrossRef] 52. Kishino, S.; Takeuchi, M.; Park, S.B.; Hirata, A.; Kitamura, N.; Kunisawa, J.; Kiyono, H.; Iwamoto, R.; Isobe, Y.; Arita, M.; et al. Polyunsaturated Fatty Acid Saturation by Gut Lactic Acid Bacteria Affecting Host Lipid Composition. Proc. Natl. Acad. Sci. USA 2013, 110, 17808–17813. [CrossRef] 53. Yang, B.; Chen, H.; Song, Y.; Chen, Y.Q.; Zhang, H.; Chen, W. Myosin-Cross-Reactive Antigens from Four Different Lactic Acid Bacteria Are Fatty Acid Hydratases. Biotechnol. Lett. 2013, 35, 75–81. [CrossRef] 54. Volkov, A.; Khoshnevis, S.; Neumann, P.; Herrfurth, C.; Wohlwend, D.; Ficner, R.; Feussner, I. Crystal Structure Analysis of a Fatty Acid Double-Bond Hydratase from Lactobacillus Acidophilus. Acta Cryst. Sect. D Biol. Cryst. 2013, 69, 648–657. [CrossRef] 55. Jo, Y.S.; An, J.U.; Oh, D.K. γ-Dodecelactone Production from Safflower Oil via 10-Hydroxy-12 (z)-Octadecenoic Acid Intermediate by Whole Cells of Candida Boidinii and Stenotrophomonas Nitritireducens. J. Agric. Food Chem. 2014, 62, 6736–6745. [CrossRef][PubMed] 56. Park, J.Y.; Lee, S.H.; Kim, K.R.; Park, J.B.; Oh, D.K. Production of 13S-Hydroxy-9 (Z)-Octadecenoic Acid from Linoleic Acid by Whole Recombinant Cells Expressing Linoleate 13-Hydratase from Lactobacillus acidophilus. J. Biotechnol. 2015, 208, 1–10. [CrossRef] 57. Hirata, A.; Kishino, S.; Park, S.-B.; Takeuchi, M.; Kitamura, N.; Ogawa, J. A Novel Unsaturated Fatty Acid Hydratase toward C16 to C22 Fatty Acids from Lactobacillus acidophilus. J. Lipid Res. 2015, 56, 1340–1350. [CrossRef] 58. Takeuchi, M.; Kishino, S.; Park, S.B.; Hirata, A.; Kitamura, N.; Saika, A.; Ogawa, J. Efficient Enzymatic Production of Hydroxy Fatty Acids by Linoleic Acid ∆9 Hydratase from Lactobacillus Plantarum AKU 1009a. J. Appl. Microbiol. 2016, 120, 1282–1288. [CrossRef] 59. Schmid, J.; Steiner, L.; Fademrecht, S.; Pleiss, J.; Otte, K.B.; Hauer, B. Biocatalytic Study of Novel Oleate Hydratases. J. Mol. Catal. B Enzym. 2016, 133, 243–249. [CrossRef] 60. Choi, J.H.; Seo, M.J.; Lee, K.T.; Oh, D.K. Biotransformation of Fatty Acid-Rich Tree Oil Hydrolysates to Hydroxy Fatty Acid-Rich Hydrolysates by Hydroxylases and Their Feasibility as Biosurfactants. Biotechnol. Bioprocess Eng. 2017, 22, 709–716. [CrossRef] 61. Meier, M.A.R.; Metzger, J.O.; Schubert, U.S. Plant Oil Renewable Resources as Green Alternatives in Polymer Science. Chem. Soc. Rev. 2007, 36, 1788–1802. [CrossRef] 62. Heo, S.H.; Hou, C.T.; Kim, B.S. Production of Oxygenated Fatty Acids from Vegetable Oils by Flavobacterium sp. Strain DS5. New Biotechnol. 2009, 26, 105–108. [CrossRef] 63. Kang, W.R.; Seo, M.J.; Shin, K.C.; Park, J.B.; Oh, D.K. Gene Cloning of an Efficiency Oleate Hydratase from Stenotrophomonas Nitritireducens for Polyunsaturated Fatty Acids and Its Application in the Conversion of Plant Oils to 10-Hydroxy Fatty Acids. Biotechnol. Bioeng. 2017, 114, 74–82. [CrossRef] 64. Jeon, E.Y.; Seo, J.H.; Kang, W.R.; Kim, M.J.; Lee, J.H.; Oh, D.K.; Park, J.B. Simultaneous Enzyme/Whole-Cell Biotransformation of Plant Oils into C9 Carboxylic Acids. ACS Catal. 2016, 6, 7547–7553. [CrossRef] 65. Kim, S.K.; Park, Y.C. Biosynthesis of ω-Hydroxy Fatty Acids and Related Chemicals from Natural Fatty Acids by Recombinant Escherichia Coli. Appl. Microbiol. Biotechnol. 2019, 103, 191–199. [CrossRef] 66. Song, J.W.; Lee, J.H.; Bornscheuer, U.T.; Park, J.B. Microbial Synthesis of Medium-Chain α, ω-Dicarboxylic Acids and ω-Aminocarboxylic Acids from Renewable Long-Chain Fatty Acids. Adv. Synth. Catal. 2014, 356, 1782–1788. [CrossRef] Catalysts 2020, 10, 287 13 of 13

67. Oh, H.J.; Kim, S.U.; Song, J.W.; Lee, J.H.; Kang, W.R.; Jo, Y.S.; Kim, K.R.; Bornscheuer, U.T.; Oh, D.K.; Park, J.B. Biotransformation of Linoleic Acid into Hydroxy Fatty Acids and Carboxylic Acids Using a Linoleate Double Bond Hydratase as Key Enzyme. Adv. Synth. Catal. 2015, 357, 408–416. [CrossRef] 68. Lee, D.S.; Song, J.W.; Voß, M.; Schuiten, E.; Akula, R.K.; Kwon, Y.U.; Bornscheuer, U.; Park, J.B. Enzyme Cascade Reactions for the Biosynthesis of Long Chain Aliphatic Amines from Renewable Fatty Acids. Adv. Synth. Catal. 2019, 361, 1359–1367. [CrossRef] 69. Wu, Y.X.; Pan, J.; Yu, H.L.; Xu, J.H. Enzymatic Synthesis of 10-Oxostearic Acid in High Space-Time Yield via Cascade Reaction of a New Oleate Hydratase and an Alcohol Dehydrogenase. J. Biotechnol. X 2019. [CrossRef] 70. Song, J.W.; Jeon, E.Y.; Song, D.H.; Jang, H.Y.; Bornscheuer, U.T.; Oh, D.K.; Park, J.B. Multistep Enzymatic Synthesis of Long-Chain α, ω-Dicarboxylic and ω-Hydroxycarboxylic Acids from Renewable Fatty Acids and Plant Oils. Angew. Chem. Int. Ed. 2013, 52, 2534–2537. [CrossRef] 71. Kim, S.U.; Kim, K.R.; Kim, J.W.; Kim, S.; Kwon, Y.U.; Oh, D.K.; Park, J.B. Microbial Synthesis of Plant Oxylipins from γ-Linolenic Acid through Designed Biotransformation Pathways. J. Agric. Food Chem. 2015, 63, 2773–2781. [CrossRef] 72. Koppireddi, S.; Seo, J.H.; Jeon, E.Y.; Chowdhury, P.S.; Jang, H.Y.; Park, J.B.; Kwon, Y.U. Combined Biocatalytic and Chemical Transformations of Oleic Acid to ω-Hydroxynonanoic Acid and α, ω-Nonanedioic Acid. Adv. Synth. Catal. 2016, 358, 3084–3092. [CrossRef] 73. Cha, H.J.; Seo, E.J.; Song, J.W.; Jo, H.J.; Kumar, A.R.; Park, J.B. Simultaneous Enzyme/Whole-Cell Biotransformation of C18 Ricinoleic Acid into (R)-3-Hydroxynonanoic Acid, 9-Hydroxynonanoic Acid, and 1, 9-Nonanedioic Acid. Adv. Synth. Catal. 2018, 360, 696–703. [CrossRef] 74. Farbood, M.I.; Morris, J.A.; McLean, L.B. Fermentation Process for Preparing 10-Hydroxy-C18-Carboxylic Acid and Gamma-Dodecalactone Derivatives. EP078388A25, 1 December 1994. 75. Park, A.K.; Lee, G.H.; Kim, D.W.; Jang, E.H.; Kwon, H.T.; Chi, Y.M. Crystal structure of oleate hydratase from Stenotrophomonas sp. KCTC 12332 reveals conformational plasticity surrounding the FAD binding site. Biochem. Biophys. Res. Commun. 2018, 499, 772–776. [CrossRef] 76. Lorenzen, J.; Driller, R.; Waldow, A.; Qoura, F.; Loll, B.; Brück, T. Rhodococcus erythropolis Oleate Hydratase: A New Member in the Oleate Hydratase Family Tree—Biochemical and Structural Studies. ChemCatChem 2018, 10, 407–414. [CrossRef] 77. Demming, R.M.; Hammer, S.C.; Nestl, B.M.; Gergel, S.; Fademrecht, S.; Pleiss, J.; Hauer, B. Asymmetric Enzymatic Hydration of Unactivated, Aliphatic Alkenes. Angew. Chem. Int. Ed. 2019, 58, 173–177. [CrossRef] 78. Eser, B.E.; Poborsky, M.; Dai, R.; Kishino, S.; Ljubic, A.; Takeuchi, M.; Jacobsen, C.; Ogawa, J.; Kristensen, P.; Guo, Z. Rational Engineering of Hydratase from Lactobacillus Acidophilus Reveals Critical Residues Directing Substrate Speciﬁcity and Regioselectivity. ChemBioChem 2019.[CrossRef] 79. Engleder, M.; Strohmeier, G.A.; Weber, H.; Steinkellner, G.; Leitner, E.; Müller, M.; Mink, D.; Schürmann, M.; Gruber, K.; Pichler, H. Evolving the Promiscuity of Elizabethkingia Meningoseptica Oleate Hydratase for the Regio- and Stereoselective Hydration of Oleic Acid Derivatives. Angew. Chem. Int. Ed. 2019, 58, 7480–7484. [CrossRef]

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