Study Towards Carotenoid 1,2-Hydratase and Oleate Hydratase as Novel Biocatalysts

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(1)Study Towards Carotenoid 1,2-Hydratase and Oleate Hydratase as Novel Biocatalysts. Aida HISENI.

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(3) Study Towards Carotenoid 1,2-Hydratase and Oleate Hydratase as Novel Biocatalysts. PROEFSCHRIFT. ter verkrijging van de graad van doctor aan de Technische universiteit Delft, op gezag van de Rector Magnificus prof. ir. K.C.A.M Luyben, voorzitter van het College voor promoties, in het openbaar te verdedigen op dinsdag 22 april 2014 om 10:00 uur. door. Aida HISENI. Diplom-Biologin, Heinrich-Heine-Universität Düsseldorf geboren te Doboj, Bosnië en Hercegovina..

(4) Dit proefschrift is goedgekeurd door de promotor: Prof. dr. I.W.C.E Arends. Samenstelling promotiecommissie: Rector Magnificus. voorzitter. Prof. dr. I.W.C.E. Arends. Technische Universiteit Delft, promotor. Prof. dr. U. Hanefeld. Technische Universiteit Delft. Prof. dr. J.H. de Winde. Universiteit Leiden. Prof. dr. G. Muijzer. Universiteit van Amsterdam. Prof. dr. R. Wever. Universiteit van Amsterdam. Dr. L.G. Otten. Technische Universiteit Delft. Dr. P. Dominguez De Maria Sustainable Momentum. Prof. dr. S. de Vries. Technische Universiteit Delft, reservelid. This project is financially supported by The Netherlands Ministry of Economic Affairs and the B-Basic partner organizations (http://www.b-basic.nl) through B-Basic, a publicprivate NWO-ACTS programme [Advanced Chemical Technologies for Sustainability (ACTS)].. ISBN. Copyright © 2014 by Aida HISENI All rights reserved. No part of this publication may be reproduced or distributed in any form or by any means, or stored in a database or retrieval system, without any prior permission of the copyright owner..

(5) To my father Ismet Nukičić.

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(7) . Table of Contents . 1 . General introduction .................................................................................................. 1 1.1 . Enzymes and biocatalysis............................................................................................................... 2 . 1.2 . Enzymes as industrial biocatalysts................................................................................................. 3 . 1.3 . Enzyme engineering ....................................................................................................................... 6 . 1.4 Hydro-Lyases ................................................................................................................................. 8 1.4.1 Non-enzymatic water addition to a carbon-carbon double bond ............................................... 8 1.4.2 Enzymatic water addition to a carbon-carbon double bond ....................................................... 9 1.4.3 Carotenoid 1,2-hydratase ......................................................................................................... 11 1.4.4 Oleate hydratase ...................................................................................................................... 14 1.5 . Scope and objectives .................................................................................................................... 22 . 1.6 . Supplementary figures.................................................................................................................. 24 . 1.7 . References .................................................................................................................................... 26 . 2 Biochemical Characterization of the Carotenoid 1,2-Hydratases (CrtC) from Rubrivivax gelatinosus and Thiocapsa roseopersicina ................................................... 33 Abstract ..................................................................................................................................................... 34 2.1 . Introduction.................................................................................................................................. 35 . 2.2 Materials and methods ................................................................................................................. 36 2.2.1 Construction of pET15b_CrtCRg and pET15b_CrtCTr expression vectors ............................ 36 2.2.2 Expression and purification of recombinant proteins .............................................................. 37 2.2.3 Tandem MS analysis ............................................................................................................... 37 2.2.4 CrtC activity assay and analysis of the products ..................................................................... 38 2.2.5 Substrate specificity ................................................................................................................. 38 2.2.6 Effects of pH and temperature on CrtC activity ...................................................................... 39 2.2.7 Effects of inhibitors and metal ions on enzyme activity .......................................................... 39 2.2.8 Circular dichroism (CD) spectroscopy .................................................................................... 39 2.2.9 Metal analysis using USN-ICP-OES ....................................................................................... 40 2.3 Results .......................................................................................................................................... 40 2.3.1 Expression and purification of the carotenoid 1,2-hydratases ................................................. 40 2.3.2 Hydratase activity .................................................................................................................... 41 2.3.3 Enzyme kinetics ....................................................................................................................... 41 2.3.4 Substrate specificity ................................................................................................................. 43 2.3.5 Effect of pH and temperature on hydratase activity and stability ............................................ 43 2.4 . Discussion .................................................................................................................................... 46 . 2.5 . Acknowledgments ......................................................................................................................... 49 . 2.6 . Supplementary information .......................................................................................................... 50 . 2.7 . References .................................................................................................................................... 57 . 3 Structural Characterization of the Carotenoid 1,2-Hydratases (CrtC) from Rubrivivax gelatinosus and Thiocapsa roseopersicina ................................................... 59 Abstract ..................................................................................................................................................... 60 . vii.

(8) 3.1 . Introduction .................................................................................................................................. 61 . 3.2 Materials and methods ................................................................................................................. 63 3.2.1 In silico analysis ...................................................................................................................... 63 3.2.2 Cloning of carotenoid 1,2-hydratase genes .............................................................................. 63 3.2.3 Construction of CrtC mutants .................................................................................................. 64 3.2.3.1 Single point mutations .................................................................................................... 64 3.2.3.2 N-terminally truncated Rg- and TrCrtC’s ....................................................................... 65 3.2.4 Recombinant expression of CrtC’s .......................................................................................... 65 3.2.5 CrtC purification ...................................................................................................................... 66 3.2.6 Determination of enzyme activity ............................................................................................ 66 3.3 Results and discussion.................................................................................................................. 67 3.3.1 Comparative in silico analysis of crtC genes ........................................................................... 67 3.3.2 Production of recombinant wildtype and mutant CrtC’s and enzymatic activity .................... 72 3.4 . Conclusion ................................................................................................................................... 80 . 3.5 . Acknowledgements ....................................................................................................................... 81 . 3.6 . References .................................................................................................................................... 82 . 4 Oleate hydratase as model enzyme to design and evaluate high-throughput screening assay for alcohol detection ............................................................................. 85 Abstract ..................................................................................................................................................... 86 4.1 . Introduction .................................................................................................................................. 87 . 4.2 Materials and methods ................................................................................................................. 89 4.2.1 Standard curves and Z-factor determination ............................................................................ 89 4.2.2 Large scale production of 10-HSA .......................................................................................... 90 4.2.3 Growth conditions in 96-well deep well plates ........................................................................ 90 4.2.4 Liquid handling ........................................................................................................................ 91 4.2.5 Assay conditions ...................................................................................................................... 91 4.2.6 Preparation of ohyA mutant libraries ....................................................................................... 92 4.2.7 Expression of ohyA variants .................................................................................................... 93 4.2.8 Library screening ..................................................................................................................... 93 4.3 Results and discussion.................................................................................................................. 94 4.3.1 Method performance and linearity with small substrates......................................................... 94 4.3.2 Method performance for larger substrates and reaction simulation ......................................... 95 4.3.3 Precision and accuracy (Z-factor) ............................................................................................ 98 4.3.4 Optimization of protein expression conditions ...................................................................... 100 4.4 . References .................................................................................................................................. 106 . 5 Preparation and properties of immobilized oleate hydratase as a cross-linked enzyme aggregate (CLEA) ............................................................................................ 109 Abstract ................................................................................................................................................... 110 5.1 . Introduction ................................................................................................................................ 111 . 5.2 Materials and Methods .............................................................................................................. 113 5.2.1 Bacterial strain, growth conditions and cell disruption .......................................................... 113 5.2.2 Precipitation procedure .......................................................................................................... 113 5.2.3 Cross-linking procedure......................................................................................................... 113 5.2.4 Activity assay......................................................................................................................... 114 5.2.5 Storage stability ..................................................................................................................... 115 5.2.6 pH activity and temperature stability ..................................................................................... 115 5.2.7 Biocatalyst recovery .............................................................................................................. 115 . viii.

(9) 5.3 Results and discussion ............................................................................................................... 115 5.3.1 Selection of the best precipitating agent for CLEA preparation ............................................ 116 5.3.2 Cross-linking and the effect of glutaraldehyde concentration ............................................... 117 5.3.3 Thermal stability and pH profile of OHase CLEA’s ............................................................. 120 5.3.4 Storage stability of OHase CLEA’s ....................................................................................... 122 5.3.5 Recycling of OHase CLEA’s................................................................................................. 124 5.3.6 Space-time yields................................................................................................................... 125 . 6 . 5.4 . Conclusion ................................................................................................................................. 125 . 5.5 . Supplemental Information ......................................................................................................... 127 . 5.6 . References .................................................................................................................................. 128 . Conclusions and future prospects ......................................................................... 131 6.1 . Carotenoid 1,2-hydratase .......................................................................................................... 132 . 6.2 . Oleate hydratase ........................................................................................................................ 134 . 6.3 . High-throughput screening assay .............................................................................................. 135 . 6.4 . References .................................................................................................................................. 138 . Summary/Samenvatting ................................................................................................ 141 Summary ................................................................................................................................................. 142 Samenvatting ........................................................................................................................................... 145 . Acknowledgements ........................................................................................................ 149 Curriculum vitae ............................................................................................................ 151 . ix.

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(11) . Chapter 1. 1 General introduction. . .

(12) General introduction. 1.1 Enzymes and biocatalysis Enzymes play a pivotal role in the metabolism of all living organisms. Nearly all biochemical reactions are accomplished and controlled by enzymes. By lowering the activation energy required for a reaction to occur, enzymes are able to dramatically accelerate the reaction rate (up to 1012-fold [1]) for reactions that otherwise would proceed very slowly or not at all. In addition, enzymes are capable of accepting a wide array of complex substrates, are highly selective (enantio-, regio- and chemoselective) and usually operate under mild conditions [2]. Enzymes found in nature have been used since ancient times in the production of food, alcoholic beverages, and manufacturing of commodities such as leather and linen. Jöns Jakob Berzelius, a Swedish chemist, observed in the early nineteenth century that a chemical reaction could be accelerated in the presence of specific compounds. At that time he also coined the term ‘proteins’, without even being aware of the existence of enzymes [3]. Only in the twentieth century, the first enzyme (urease) could be isolated in pure form by James B. Sumner, an American chemist [4]. Since then, enzymes have captured special attention of many researchers. According to Wikipedia, ‘biocatalysis is the use of natural catalysts, such as protein enzymes, to perform chemical transformations on organic compounds. Both enzymes that have been more or less isolated and enzymes still residing inside living cells are employed for this task’. Historically, catalysis is divided into two categories: homogeneous and heterogeneous [5]. Enzymes, however, do not fit into the classical definitions of these two categories. They are usually regarded as a separate class. On the other hand, the development of the biomimetic organocatalysis is causing fading of the boundaries between the catalysis domains. For instance, chemical catalysts are produced, which mimic the natural features of enzymes and also, artificial enzymes are synthesized with specific properties optimized for the targeted application. Despite the early discovery of the catalytic nature of enzymes, their application in industrial processes was not always competitive with chemical catalysis or vice versa [5]. Better understanding of enzyme structure-function relationships and the possibility to tailor their properties have significantly decreased the gap between chemical and enzymatic catalysis [6].. 2.

(13) 1.2 Enzymes as industrial biocatalysts. 1.2 Enzymes as industrial biocatalysts The evolution of modern biotechnology over the last four decades and the emergence of a key technology – genetic engineering - have opened new horizons in the fields of biocatalysis and industrial biotechnology. Today’s novel techniques allow not only the manufacturing of enzymes as purified, well-characterized preparations even on a large scale, but they also make it possible to produce tailor-made enzymes that are designed for a specific and often non-natural application. The application of enzymes as biocatalysts is recognized as a significant complement to the use of chemical reagents. [7] Enzymes are increasingly being utilized for both environmental and economic reasons in a number of industries including agro-food, animal feed, detergent, textile and specialty chemical industry [8, 9] (Table 1.1). Table 1.1 Enzymes used in various industrial segments and their application, adapted from [10]. Industry Enzyme class Application Detergent. Protease. (laundry and dish wash) Amylase. Protein stain removal Starch stain removal. Lipase. Lipid stain removal. Cellulase. Cleaning, color clarification, anti-redeposition (cotton). Starch and fuel. Mannanase. Mannanan stain removal (reappearing stains). Amylase. Starch liquefaction and saccharification. Amyloglucosidase. Saccharification. Pullulanase. Saccharification. Glucose isomerase. Glucose to fructose conversion. Cyclodextrin-. Cyclodextrin production. glycosyltransferase Xylanase. Viscosity reduction (fuel and starch). Protease. Free amino nitrogen production (yeast nutrition fuel). Food. Protease. (including dairy). Milk clotting, infant formulas (low allergenic), flavor. Lipase. Cheese flavor. Lactase. Lactose removal (milk). Pectin methyl esterase. Firming fruit-based products. Pectinase. Fruit-based products. 3.

(14) General introduction. Baking. Transglutaminase. Modify visco-elastic properties. Amylase. Bread softness and volume, flour adjustment. Xylanase. Dough conditioning. (Phospho)Lipase. Dough stability and conditioning (in situ emulsifier). Animal feed. Beverage. Glucose oxidase. Dough strengthening. Lipoxygenase. Dough strengthening, bread whitening. Protease. Biscuits, cookies. Transglutaminase. Laminated dough strength. Phytase. Phytate digestibility - phosphorus release. Xylanase. Digestibility. β-Glucanase. Digestibility. Pectinase. De-pectinization, mashing. Amylase. Juice treatment, low calorie beer. β-Glucanase. Mashing. Acetolactate decarboxylase. Maturation (beer). Laccase. Clarification (juice), flavor (beer), cork stopper treatment. Textile. Pulp and paper. Cellulase. Denim finishing, cotton softening. Amylase. De-sizing. Pectate lyase. Scouring. Catalase. Bleach termination. Laccase. Bleaching. Peroxidase. Excess dye removal. Lipase. Pitch control, contaminant control. Protease. Biofilm removal. Amylase. Starch-coating, de-inking, drainage improvement. Xylanase. Bleach boosting. Cellulase. De-inking, drainage improvement, fiber modification. Fats and oils. Organic synthesis. Leather. 4. Lipase. Transesterification. Phospholipase. De-gumming, lyso-lecithin production. Lipase. Resolution of chiral alcohols and amines. Acylase. Synthesis of semi-synthetic penicillin. Nitrilase. Synthesis of enantiopure carboxylic acids. Nitrile hydratase. Synthesis of acrylamide. Fumarase. Synthesis of malate. Protease. Unhairing, bating.

(15) 1.2 Enzymes as industrial biocatalysts. Personal care. Lipase. De-pickling. Amyloglucosidase. Antimicrobial (combined with glucose oxidase). Glucose oxidase. Bleaching, antimicrobial. Peroxidase. Antimicrobial. The main benefits offered by enzymes are: (i) waste and energy reduction; enzymes usually work at mild conditions, thereby circumventing the need for harsh chemicals and extreme working conditions; (ii) cleaner products; enzymes are highly specific, resulting in less/no unwanted side reactions and byproducts; (iii) environmental sustainability; enzymes are biodegradable and thereby have no environmental footprint by definition. The synthesis of the drug cortisone is an excellent example of the possibilities of enzyme technology [11, 12]. Here, the number of process steps needed to produce the drug could significantly be reduced from 31 steps (chemical synthesis) to only 5 steps by utilizing enzymes (Figure 1.1).. Figure 1.1 Chemical (upper reaction) and biochemical (lower reaction) route to cortisone (adapted from [5]).. The interest in industrial biocatalysis has increased rapidly and it still continues to grow. It has been estimated that the global market for industrial enzymes is going to reach $6 billion by 2016. Consequently, much effort has been devoted to the development of cleaner alternative technologies where enzymes are utilized as biocatalysts.. 5.

(16) General introduction. 1.3 Enzyme engineering Despite the huge potential of enzymes in the field of biotechnology, their application is often limited by low stability and/or catalytic activity of these enzymes under process conditions. This is one of the reasons why the use of hydrolases on industrial scale prevails (Figure 1.2). For instance, lipases which act carboxylic ester bonds, are very versatile enzymes with, among others, broad substrate specificity and stability, and can therefore be utilized in many different industrial applications, such as food, detergent and pharmaceutics [13].. EC 4 (Lyases) 4%. EC 5 (Isomerases) 2%. EC 1 (Oxidoreductases) 14% EC 2 (Transferases) 5%. EC 3 (Hydrolases) 75%. Figure 1.2 Pie chart illustrating the utilization of different enzyme classes (EC) on industrial scale, based on Table 1.1.. Therefore, the utilization of enzymes as biocatalysts in industrial processes requires an intensive study and optimization of enzyme properties, such as stability, specific activity and selectivity, beforehand. The development of strategies to overcome the limitations of natural enzymes as biocatalysts has received an enormous boost during the last years [6]. Protein engineering techniques, among others, offer solutions to removing the impediments of widespread application of enzymes in industrial processes. These techniques include random mutagenesis and (semi)rational design/focused mutagenesis (Figure 1.3).. 6.

(17) 1.3 Enzyme engineering. Figure 1.3 Strategies in protein engineering and prerequisites in terms of structural information. Recent methods in diversity generation have been assigned to two categories: (semi)rational design and directed evolution.. The prerequisite for rational design is a detailed structural and mechanistic knowledge of the target enzyme. However, for many enzymes that have been discovered, the X-ray crystal structure or even a viable homology model is not available yet, so that rational mutagenesis of these enzymes is not an option. In contrast, for random mutagenesis, knowledge of the structure-function relationship is not required. In this case, libraries containing a large number of randomly generated mutants with potentially improved and/or novel properties can be produced in a short time. A crucial step in this so-called directed evolution approach is the development of a highthroughput screening (HTS) or a selection assay. The assay allows rapid identification of mutants with the desired properties from a large number of random mutants within a reasonable timeframe [14]. In general, it needs to be sensitive, easy to perform, robust and has to have high throughput. Preferably, screening assays are performed in plate readers, where colorimetric changes or fluorescence formation can be detected upon enzymatic activity. Selection only yields variants that have an advantage over the wild type enzyme in contrast to screens, where the activity of each variant is monitored [15]. In addition, screening methods allow the use of the actual substrate and desired reactions conditions. In the end, “you get what you screened for” [16]).. 7.

(18) General introduction. 1.4 Hydro-Lyases As indicated above (Figure 1.2), the main EC class that is successfully used in industrial biocatalytic processes is the class of hydrolases (EC 3.-.-.-). Lyases (EC 4.-.-.-), the subject of this thesis, on the other hand, are underrepresented and only a few group members are amenable to be used for industrial scale reactions, including nitrile hydratase (EC 4.2.1.84) for the production of acrylamide [9] and fumarase (EC 4.2.1.2) to produce malate [17]. Hydro-lyases (EC 4.2.1.-), also called hydratases, are a subclass of carbon-oxygen lyases (EC 4.2.-.-). As a lyase, they catalyze the non-hydrolytic and non-oxidative addition and/or removal of a group to a carbon-carbon double bond. The ‘hydro’ relates to the added or removed group, and is in this case a water molecule.. 1.4.1 Non-enzymatic water addition to a carbon-carbon double bond The addition of a water molecule to a non-activated carbon-carbon double bond to yield an alcohol is a very non-selective reaction that requires harsh conditions in traditional chemistry [18]. The non-enzymatic hydration reaction is usually performed by using strong acids, high temperatures and high pressures, or transition metals as a catalyst. Furthermore, the hydration often does not proceed with the desired positional specificity. The acidcatalyzed hydration of an alkene follows the Markovnikov’s rule [19]. It states that the acidic proton binds to the carbon with the greater number of hydrogen atoms, whereas the alcohol group prefers the carbon with the most carbon-carbon bonds (Figure 1.4). The basis of the reaction is the formation of the most stable carbocation, which is subsequently attacked by the nucleophilic water to form the oxonium-ion. Another water molecule takes up the extra proton from the attached oxygen and an alcohol is formed.. Figure 1.4 Acid-catalyzed alkene hydration. 8.

(19) 1.4 Hydro-Lyases Depending on the structure of the alkene used, unwanted side products and product rearrangements can occur, especially with unsymmetrical alkenes. The chemical reaction is therefore limited to alkenes that cannot undergo rearrangement upon hydration.. 1.4.2 Enzymatic water addition to a carbon-carbon double bond The enzymatic hydration of carbon-carbon double bonds is catalyzed by hydro-lyases [20]. The alcohol is produced under very mild conditions in a neutral aqueous environment. Due to the inherent high selectivity (enantio-, regio- and chemospecificity) of enzymes, alcohols can be obtained in high yields and without undesired side products. The enzyme database BRENDA [21] counts 153 hydro-lyases (as at December 5, 2013), which catalyze the (de)hydration of a large number of different substrates. Most of these hydro-lyases act on conjugated carbon-carbon double bonds [20, 22]. In contrast to the hydration of isolated carbon-carbon double bond, which is subject to hydronium-ion catalysis (Figure 1.4), a Michael-type hydration occurs for activated double bonds. In this case, the carbon-carbon bond is activated by an electron withdrawing group such as carboxylic acid, thioester or a phosphate group, making it more electrophilic for the nucleophilic addition by water (Figure 1.5).. Figure 1.5 Michael addition of a water molecule to an α,β-unsaturated carbonyl compound.. An excellent overview of these enzymes was recently presented in literature [20]. However, in this thesis, we are aiming for the so far underrepresented class of hydro-lyases acting on isolated carbon-carbon double bonds. Most of the (de)hydratases are cofactor dependent (Table 1.2). These cofactors have several functions in the (de)hydration mechanism of (de)hydratases. Next to the direct participation in substrate binding by, for instance, coordination (metal ions, iron-sulfur clusters), the co-factors can also be involved in the. 9.

(20) General introduction stabilization of the carbocation intermediate (e.g. pyridoxal phosphate) or are producers of radicals, as found in the dehydration mechanism of diol dehydratase [23]. Table 1.2 Reported activity requirements for hydratases/dehydratases. Activity requirement Metal ions. Enzyme name. Reference 2+. Carbonate dehydratase (Zn ). [24]. Phosphopyruvate hydratase (Mg2+). [25] 2+. 3-Dehydroshikimate dehydratase (Mn ) 2+. Coenzyme Iron-sulfur. CoA activated substrates Heme-thiolate +. NAD(P) / NAD(P)H FAD. [26] 2+. O-succinylbenzoate synthase (Mn or Mg ). [27]. 1,5-Anhydro-D-fructose dehydratase (Ca2+ or Na+ or Mg2+). [28]. Propanediol dehydratase (Cyanocobalamin). [29]. Tryptophan synthase (Pyridoxal 5'-phosphate). [30]. Aconitate hydratase. [31]. 2-Methylcitrate dehydratase. [32]. Methanogen homoaconitase. [33]. Fumarase (Class I). [34]. Enoyl-CoA hydratase. [35]. Long-chain-enoyl-CoA hydratase. [36]. Hydroperoxide dehydratase. [37]. Colneleate synthase. [38]. CDP-glucose 4,6-dehydratase. [39]. UDP-N-acetylglucosamine 4,6-dehydratase. [40]. 4-Hydroxybutanoyl-CoA dehydratase. [41]. Electron carriers such as NAD+ and NADP+ are usually involved in redox reactions. Therefore, one would not expect to find these cofactors in hydratases (Table 1.2), as the catalytic mechanism of hydratase does not involve a net oxidation or reduction. From a functional point of view, NAD+ behaves as a prosthetic group in hydratases rather than as coenzyme, when it is tightly bound to the enzyme, such as in CDP-glucose 4,6-dehydratase [39]. In this case it initiates the dehydration reaction by oxidation of the substrate and reduction once the substrate has been dehydrated by the enzyme. The catalytic reaction is independent of the NAD+/NADH ratio because of the non-dissociable character of the prosthetic group. The same has been reported for flavoproteins, which catalyze reactions with no net redox change [42]. From an industrial point of view, however, enzymes without the requirement of any cofactor are preferred. The reason is the simplification of the process and no need for the usually expensive cofactors or development of a cofactor regeneration system. 10.

(21) 1.4 Hydro-Lyases The following two paragraphs describe two newly discovered hydratases that are cofactor independent and act on isolated carbon-carbon double bonds. Therewith, these are potentially interesting biocatalysts for industrial applications.. 1.4.3 Carotenoid 1,2-hydratase Carotenoid 1,2-hydratase (also known as CrtC) is a member of hydro-lyase group EC 4.2.1.131 and occurs in the biosynthetic pathway of carotenoids [43]. From a chemical point of view, CrtC’s are able to perform a very challenging chemical reaction, namely the addition of water to an isolated carbon-carbon double bond [20]. The reaction proceeds with no assistance from electron withdrawing groups, or transition metal cations and does not occur at all under mild conditions in vitro [44]. Carotenoids, which represent one of the most abundant natural pigments with structural and protective properties [45], play an essential role in the photosynthetic machinery of phototrophic organisms such as purple bacteria [46] and higher plants [47]. However, they have also been identified in fungi and some non-photosynthetic bacteria [48]. Depending on the producing organism, carotenoids can be acyclic, monocyclic or bicyclic. CrtC introduces a tertiary hydroxyl group into a carotenoid molecule by addition of water to the carbon-carbon double bond at the C-1 position. The substrate specificity of CrtC’s varies between species. For example, the substrate specificity of the CrtC from Rhodobacter capsulatus is very limited and the enzyme accepts only acyclic carotenoids, which possess two -end groups (acyclic C9 end group according to nomenclature of carotenoids), such as neurosporene and lycopene (Figure 1.6). Once one of the two -end groups is hydrated, the enzyme is not able to use the monohydroxylated carotenoid as a substrate [49]. In contrast, the CrtC’s from Rubrivivax gelatinosus (Rg) and Thiocapsa roseopersicina (Tr) are able to also hydrate monohydroxylated acyclic carotenoids [50]. Next to acyclic carotenoids, the CrtC’s from the purple sulfur bacteria Thiodictyon sp. CAD16 [51] and from the green sulfur bacteria Chlorobium tepidum [52] showed activity towards cyclic carotenoids such as γ-carotene and chlorobactene (Figure 1.6). The substrate specificity of the CrtC and other enzymes involved in the biosynthetic pathway of carotenoids, determine the final structure of accumulated carotenoids in the organism. CrtC belongs to the Pfam family PF07143 that encompasses members from several photosynthetic bacteria. Up to now, several carotenoid 1,2-hydratases have been identified in photosynthetic [52-56] as well as in non-photosynthetic bacteria [57, 58]. Recently, carotenoid 1,2-hydratases have been identified in the non-photosynthetic bacterium 11.

(22) General introduction Deinococcus [58], which are able to hydrate γ-carotene, a mono-cyclic substrate, but no acyclic carotenoids.. Lycopene. OH. 1-HO-Lycopene Neurosporene. OH. 1-HO-Neurosporene. OH. Demethylspheroidene. OCH3. Spheroidene. OCH3. 1-CH3O-3,4-didehydrolycopene. OH. 1-HO-3,4-didehydrolycopene.  -Carotene. Chlorobactene HO. Geranylgeraniol (?). Figure 1.6 Substrates accepted by carotenoid 1,2-hydratases. The shaded portions of each structure are hydrated to yield a tertiary alcohol group. Next to differences in one end group of each carotenoid, the number of double bonds in the molecules differs as well (circled).. They are, however, evolutionary very distinct from the PF07143 members [58] and hence, they have been given the name CruF. Interestingly, cruF homologues are found in a wide variety of carotenoid-synthesizing bacteria that lack a crtC gene [59]. For example, it was found in cyanobacterium Synechococcus sp. [59] and in Herpetosiphon aurantiacus [60]. To our knowledge no published data exist on the catalytic mechanism of CrtC’s, nor has the 3D structure been elucidated yet. However, the 3D structure of the first representative of the Pfam family PF09410 (putative AttH) has been solved, a family which is distantly related to the CrtC family PF07143 [61]. The mechanism of lycopene hydration to hydroxyl compounds, which involves proton attack at C-2 and C-2′ with a carbocation intermediate and the introduction of the hydroxyl group at C-1 and C-1′, was established from 2H2Olabeling studies with intact cells [62, 63]. Until now no mutagenesis studies have been. 12.

(23) 1.4 Hydro-Lyases published on CrtC, so we identified and mutagenized potential key residues in RgCrtC and TrCrtC. The results of the mutagenesis study together with modeling of a 3D structure with putative AttH led to the hypothesis that the hydration of lycopene is initiated by an acidic residue, Asp268 in RgCrtC and Asp266 in TrCrtC, followed by quenching with solvent water molecules present in the close proximity. From these findings it becomes clear that the complete structure of the enzymes, through crystallization studies, will be pivotal to further unravel the mechanism for this intriguing enzyme. Nevertheless, the results of the study described in chapter 3 shed for the first time light on structure-activity relationships of carotenoid 1,2-hydratases. Whereas CrtC’s from photosynthetic bacteria act on acyclic carotenoids, the CruF’s from non-photosynthetic bacteria only catalyze the hydration of mono-cyclic carotenoids. Protein sequence alignment of CrtC from Rubrivivax gelatinosus and CruF from Deinococcus radiodurans R1 did not reveal any structural similarities (Supplementary figure 1.1). Moreover, they showed substantial differences in the secondary structure. While the CrtC mainly consists of β-strands, the CruF contains notably α-helices (Supplementary figure 1.2). The catalytic and structural features, that determine hydratase activity and specificity of these two distinct families remains hypothetic or unknown. Recently, we recombinantly expressed and characterized two representatives of the PF07143 family, the CrtC from purple non-sulfur Betaproteobacteria Rubrivivax gelatinosus and purple sulfur Gammaproteobacteria Thiocapsa roseopersicina [50]. Biochemical studies revealed that these enzymes are able to convert cofactor independently lycopene into 1-HO-lycopene and 1,1’-(HO)2-lycopene. In addition, they showed some activity towards the unnatural substrate geranylgeraniol, a C20 molecule that resembles the natural substrate lycopene. However, the obtained product could not yet be identified as a hydration product. Furthermore, Steiger et al. [49] have shown that the CrtC from R. gelatinosus also has the ability to hydrate neurosporene, 1-HO-neurosporene and a few other carotenoids [49]. Both CrtC’s are located in the membrane fraction after the heterologous expression in E. coli. The analysis of the amino acid sequence with transmembrane prediction program TMHMM [64] did not reveal any transmembrane segments. However, amino acid region from ca. 120 to 140 is largely hydrophobic in both CrtC’s, which suggest that the enzymes is rather bound to the membrane through an anchor so that a close distance to the substrate, which is synthesized in the cell membranes, is facilitated.. 13.

(24) General introduction. 1.4.4 Oleate hydratase Oleate hydratase (OHase) catalyzes the conversion of oleic acid (OA) into (R)-10hydroxystearic acid (10-HSA). The enzymatic hydration of OA into 10-HSA (Figure 1.7) was first described in a Pseudomonas strain [65].. Figure 1.7 Conversion of oleic acid into 10-hydroxystearic acid.. Since then reports followed for a series of different bacterial and eukaryotic microorganisms, such as Corynebacterium [66], Saccharomyces cerevisiae [67], Sphingobacterium thalpophilum [68] and Stenotrophomonas nitritireducens [69]. However, no enzyme responsible for this hydration reaction could be identified. Only recently, Bevers et al. [70] succeeded in finding the enzyme and isolating the corresponding gene sequence using the primer walking method. The enzyme was isolated from Elizabethkingia meningoseptica (formerly known as Pseudomonas sp. 3266), the same strain that Davis et al. [71] described 43 years ago. After the recombinant expression in E. coli the enzyme indeed was able to cofactor independently form 10-HSA by hydrating the substrate OA. It is a monomeric 70 kDa soluble enzyme containing one non-essential Ca2+ion. This hydratase, as well as the carotenoid 1,2-hydratase, represents a new type of hydrolyase as it is able to hydrate an isolated carbon-carbon double bond. Following its disclosure, OHase has become a favorite topic of many researchers. A number of putative enzymes have been recombinantly expressed, characterized and identified as oleic acid hydratase or fatty acid hydratase. So far, the enzyme has been cloned from Streptococcus pyogenes [72], Bifidobacterium breve [73], Lysinibacillus fusiformi [74, 75], Stenotrophomonas maltophilia [76, 77], Macrococcus caseolyticus [78], Lactobacillus rhamnosus LGG, Lactobacillus plantarum ST-III, Lactobacillus acidophilus NCFM and Bifidobacterium animalis subsp. lactis BB12 [79]. Table 1.3 shows an overview of all characterized OHases and the tested substrates. The results indicate that in all cases, (i) the. 14.

(25) 1.4 Hydro-Lyases carboxylic group, (ii) a minimum distance of nine carbons between the double bond and the acid group, (iii) a minimum chain length of C-14 and (iv) a cis-conformation, are required for conversion of the substrate. For instance, all tested OHases were able to convert the OA into the 10-HSA, while no product was detected when the trans-isomer was used. Furthermore, differences in specificity were observed for the M. casolyticus OHase. In contrast to other OHases, this enzyme introduced a second hydroxyl-group at the C-12 position next to the hydroxyl-group at the C-9 position of the substrate gamma-linolenic acid (C18:3, 6Z, 9Z, 12Z). This could either indicate a true difference in substrate specificity or insufficient incubation time for other OHases, which might have lower reaction rates with this particular substrate. The molecular weight of all reported OHases is ~67 kDa, with B. animalis OHase being the exception with a molecular weight of 82 kDa. OHases from L. fusiformi, S. maltophilia and M. caseolyticus were shown to consist of a dimeric conformational structure upon purification. Although, the OHase from E. meningoseptica is monomeric upon purification, it dimerizes after some time. Furthermore, in our lab it has been established now that the OHase from E. meningoseptica does contain a flavin adenine dinucleotide (FAD) cofactor (Figure 1.8), as well as has been demonstrated for all other oleate hydratases. With the multiple sequences alignment a motif indicative of FAD binding has been identified in all reported and characterized oleic acid hydratases, including that from E. meningoseptica (Figure. 1.9).. They. all. share. the. common. conserved. sequence. motif. GxGxxG(S/A/N)(x)15E(K/D)(x)5E(D/G/S) (where x denotes any amino acid) at the N-terminal part of the sequence, known to bind the FAD cofactor. The first part of the motif (containing the GxGxxG sequence) is the well-known Rossmann fold, a common fold in the FADcontaining glutathione reductase family (GR) [80].. Figure 1.8 Flavin adenine dinucleotide (FAD) cofactor.. 15.

(26) . Lauric acid (C12, no double bond) Myristic acid (C14, no double bond) Myristoleic acid (C14:1, 9Z) 10-Hydroxymyristic acid Palmitic acid (C16, no double bond) Palmitoleic acid (C16:1, 9Z) 10-Hydroxyhexadecanoic acid Stearic acid (C18, no double bond) Petroselinic acid (C18:1, 6Z) Elaidic acid (C18:1, 9E) Oleic acid (C18:1, 9Z) 10-Hydroxyoctadecanoic acid Ricinoleic acid (C18:1, 9Z, 12-OH) 10,12-Dihydroxystearic acid Vaccenic acid (C18:1, 11Z) conjugated-Linoleic acid (C18:2, 9E, 11E) conjugated-Linoleic acid (C18:2, 9Z, 11E) Linoleic acid (C18:2, 9Z, 12Z) 10-Hydroxy-12(Z)-octadecenoic acid 10,13-Dihydroxyoctadecanoic acid Linoleic acid methyl ester (C18:2, 9Z, 12Z) conjugated-Linoleic acid (C18:2, 10E, 12Z) gamma-Linolenic acid (C18:3, 6Z, 9Z, 12Z) 10-Hydroxy-6(Z),12(Z)-octadecadienoic acid 10,13-Dihydroxy-6(Z)-octadecenoic acid alpha-Linolenic acid (C18:3, 9Z, 12Z, 15Z) 10-Hydroxy-12(Z),15(Z)-octadecadienoic acid 10,13-Dihydroxy-15(Z)-octadecenoic acid Arachidic acid (C20, no double bond) Eicosatrienoic acid (C20:3, 3Z, 6Z, 9Z) Arachidonic acid (C20:4, 5Z, 8Z, 11Z, 14Z) Erucic acid (C22:1, 13Z) Nervonic acid (C22:1, 15Z) Dilinoleoylphosphatidylcholine Trilinoleylglycerol. Substrate Product. Bifidobacterium animalis subsp. Lactis BB12 [79]. Y Y Y N N -. Y Y N -. N -. -. Bifidobacterium breve [73]. N. -. -. Elizabethkingia meningoseptica [70] -. -. -. -. Y -. -. -. -. Lactobacillus acidophilus NCFM [79] -. -. -. Y N -. Y N -. -. Y -. N. -. -. Lactobacillus plantarum ST-III [79]. -. Y -. N. -. -. Lactobacillus rhamnosus LGG [79] -. -. Y N -. -. Y -. N. -. -. Lysinibacillus fusiformi [74,75]. Y Y -. Y Y. Y N Y N -. Y Y -. -. Y -. Y -. Y -. -. Y N -. Y -. Y. Y -. Y -. -. Macrococcus caseolyticus [78]. 16 Y N N N N -. Y N. Y N N. N N N. Y -. Y N N N. Y N. N N. Stenotrophomonas maltophilia [76, 77]. Table 1.3 Overview of the substrates tested with oleate hydratases recombinantly expressed E. coli. N, no product detected; Y, product detected, -, not determined. . Streptococcus pyogenes [72] N Y N N N N. Y N. Y N -. -. Y -. Y N. N -. -. General introduction.

(27) 1.4 Hydro-Lyases Although distinct conserved sequence motifs were identified in all four FAD families (GR, ferredoxin reductase (FR), p-cresol methylhydroxylase (PCMH) and pyruvate oxidase (PO)), the GxGxxG sequence is the most conserved one and is found in proteins across all four families. The importance of the glycine residues was described by Wierenga et al. [81]. In their study they have been able to derive an amino acid sequence fingerprint, which can be attributed to the so-called βαβ-unit with ADP-binding properties (Figure 1.10). The hydrophobic amino acids of the fingerprint sequence form the hydrophobic core between the β-strand and the α-helix, while the first and the second glycine residues allow a sharp turn and a close approach of the pyrophosphate of the FAD cofactor to the N-terminus of the α-helix, respectively. The acid side-chain at the end of the fingerprint sequence forms a hydrogen bond with the hydroxyl group of the adenine moiety. The amino acid sequence as found in oleate hydratase reveals a slightly different motif compared to the described βαβ-unit with ADP-binding properties (Figure 1.10). It comprises two instead of one acid side-chain. Joo et al. [78] demonstrated by mutagenesis studies the importance of the second acid side-chain in the GxGxxG(S/A/N)(x)15E(K/D)(x)5E(D/G/S) motif (acid side-chain underlined) for the catalytic activity of the oleate hydratase from M. caseolyticus. While a mutant, with the first acid side-chain being substituted by an alanine, retained 60-85% of the wild-type activity, the mutation of the second acid side-chain by an alanine resulted in a fully inactivated enzyme. As already pointed out, the presence of the highly conserved GxGxxG motif in all FAD protein families indicates a crucial role for the molecular recognition of the pyrophosphate moiety. In contrast, residues involved in the binding of the isoalloxazine- and adenine moiety are less conserved and show higher diversity between all the FAD- family members. The isoalloxazine ring structure of the FAD cofactor is involved in the catalytic function in enzymes which catalyze redox reactions. The absent conserved motif for the binding of this part of the FAD molecule within the reported oleate hydratase sequences is consistent with the known fact that the hydration mechanism of these enzyme does not involve any redox reactions [82]. The partly conserved FAD-binding motif and the experimental data on cofactor removal by heat precipitation [78] show that the cofactor in oleate hydratases is held together by weak non-covalent rather than covalent bonds.. 17.

(28) General introduction. * * * . *. 18. * .

(29) 1.4 Hydro-Lyases Figure 1.9 Multiple sequence alignment showing conserved amino acids of the oleate hydratase (OHase) protein sequences from various bacteria. Identical amino acids are highlighted in black. Sequences analyzed: Elizabethkingia meningoseptica (GI 380877058), Lysinibacillus fusiformis (GI 424736965), Macrococcus caseolyticus (GI 222150326), Lactobacillus acidophilus (GI 58336974), Stenotrophomonas maltophilia (GI 459793677), Streptococcus pyogenes (GI 383479572), Bifidobacterium breve (GI 290048343), Bifidobacterium animalis (GI 384190730), Lactobacillus plantarum (GI 308179305), Lactobacillus rhamnosus (GI 258507498). The predicted FAD binding residues (G70, G72, G75, K91 and E97 of oleate hydratase from Elizabethkingia meningoseptica) are indicated with an asterisk.. Furthermore, through mutagenesis studies of the glycine residues in the oleate hydratase from M. caseolyticus [78] the crucial role for the binding of the cofactor was demonstrated. The molecular interaction of the obtained mutants with FAD was significantly reduced and resulted in inactivation of the enzymatic activity.. Figure 1.10 Schematic drawing of the βαβ-fold from spiny dogfish lactate dehydrogenase with ADP-binding properties, adapted from [81]. The properties of the amino acid residues are indicated with symbols (triangle, hydrophilic or basic; square, hydrophobic and small).. The structural and mechanistic data of oleate hydratases were not available until only recently. Volkov et al. [83] succeeded in determining the crystal structure of oleate hydratase from L. acidophilus, which shares 40% and 57% amino acid sequence identity and similarity, respectively, with that from E. meningoseptica (the enzyme we use in this. 19.

(30) General introduction research). The enzyme has been crystallized in apo and product-bound (linoleic acid) form and is shown to consist of two stably bound monomers. Upon dimerization, 9.7% of the surface of each monomer becomes buried. Based on the structural similarity to other FADbinding proteins four different domains were identified (Figure 1.11). Domain 1 (D1) consists of four regions throughout the whole gene and resembles a variant of the Rossmann fold. Domain 2 (D2) is shown to contain the FAD-substrate binding sites in concert with D1. Domain 3 (D3) and domain 4 (D4) comprise only α-helices. The latter was shown to have structural similarity with the N-terminal lid domain of the long-chain acylglycerol lipase from Archaeoglobus fulgidus. Based on the comparison of the obtained apo- and product-bound OHase structures, a displacement of 2 α-helices at the C-terminal region in D4 is observed upon binding of the substrate linoleic acid. This led the authors to the hypothesis that a cavity forming the entrance to a channel is generated, which runs from the surface down to a cleft at the interface of D1 and D3.. D2 . D3 D4 . D1 . Figure 1.11 Crystal structure of Lactobacillus acidophilus hydratase, adapted from [83]. Domains 1, 2 and 3 are shown in marine, light green and red color, respectively. The ‘flexible’ domain 4 is depicted in yellow. Solvent molecules are shown in ball-and-stick representation and are colored cyan. Channel leading to a putative active site and a putative FAD-binding site are depicted as transparent and yellow surfaces, respectively.. The interior of the channel mainly contains hydrophobic side chains, which accommodate the long fatty acid chain, while positively charged residues at the entrance of the channel (D4) possibly facilitate the recruitment of fatty acids by making a salt bridge to its carboxyl. 20.

(31) 1.4 Hydro-Lyases group. The inability of oleate hydratases to convert substrates lacking the carboxylic group, such as methyl linoleate [72, 73], argue in favor of the substrate recognition at the entrance of the channel. Due to the fact that the crystallization of the hydratase with the bound cofactor FAD was not successful, the FAD-binding domains were only identified through structural similarities to other FAD-binding proteins. Residues involved in the binding of the cofactor are similarly arranged as depicted in Figure 1.10. While the first two glycines are located in the loop region between the first β-strand and α-helix, and help the positioning of the sugar group of the cofactor, the third glycine is positioned in the loop region where the isoalloxazine ring resides. The acid-side chain glutamate (corresponds to E97 in E. meningoseptica OHase), located right after the second β-strand, forms a hydrogen bond to the hydroxyl group of the sugar moiety of the FAD molecule. The here proposed architecture of the FAD binding site is in agreement with the above described. With the availability of the first crystal structure of an oleate hydratase, it was possible to use it as a template in order to generate a model for oleate hydratase from E. meningoseptica. As mentioned earlier, the amino acid similarity of these two hydratases is 57%, which should be sufficient for a reasonable alignment. A model has been generated with an estimated accuracy of 0.63 ± 0.14 (model with a score > 0.5 is considered a good model). Figure 1.12 is a 3D representation of the superimposed structures of oleate hydratases from L. acidophilus and E. meningoseptica and is based on sequence alignment. Overall, both enzymes seem to share similar topology, indicating that these structures are closely related. The topology of the C-terminal region (D4) and D2 region comprising the FAD-binding site, however, has diverged. Volkov et al. [83] proposed the D4 region, which consist of 2 α-helices, as the entrance of the substrate to the active site. In case of E. meningoseptica oleate hydratase one of the two helices is extended, which might indicate different substrate specificity.. 21.

(32) General introduction. D2 . D3 D4 . D1 . Figure 1.12 Superimposed 3D structures of oleate hydratases from L. acidophilus (cyan) and E. meningoseptica (green), which are based on sequence alignment.. 1.5 Scope and objectives “We do not inherit the earth from our ancestors; we borrow it from our children” (Native American proverb). This quote perfectly pictures the motivation of the research described in this thesis. The environmental change has long-reaching consequences and now, it has been recognized that the development of sustainable and green technologies is of vital importance for our future. As has been introduced in this chapter, enzymes have a great potential in the field of industrial biotechnology. Specifically, hydratases could make very valuable biocatalysts for the chemical industry. The aim of the research described in this thesis was to gain knowledge on structure-function relationship of two newly discovered hydratases, namely carotenoid 1,2-hydratase and oleate hydratase. Based on that, the objective was to map the potential of these two hydratases for their use as biocatalysts in industrial processes. Chapter 2 describes the characterization of carotenoid 1,2- hydratases from photosynthetic bacteria Rubrivivax gelatinosus and Thiocapsa roseopersicina. The biochemical properties. 22.

(33) 1.5 Scope and objectives of the recombinant enzymes and their substrate specificities were studied. In Chapter 3, the two hydratases described in chapter 2 were subjected to protein engineering techniques site-directed evolution and semi-rational mutagenesis in order to identify relevant amino acids in the active site and their contribution to enzymatic activity. Homology modeling together with mutagenesis study helped to gain insight into the enzymatic mechanism of these enzymes. Chapter 4 focuses on the development of a high-throughput screening assay for the detection of alcohols, products of hydrating enzymes such as carotenoid 1,2hydratase and oleate hydratase (OHase). For this study, OHase from Elizabethkingia meningoseptica was used as the model enzyme. The assay allows for screening of mutant libraries generated by directed evolution. A continuation of the characterization work of OHase that was performed by Loes Bevers, included the study of OHase immobilization as cross-linked enzyme aggregates (CLEA) with the goal to develop OHase into a useful and efficient biocatalyst for high-added value compounds (Chapter 5). In Chapter 6, the main findings described in this thesis are evaluated with the respect to the implications of the work to the fundamental knowledge on carotenoid 1,2-hydratases and oleate hydratases. Next to that, perspectives for future research are presented. Finally, the main findings of this thesis are summarized.. 23.

(34) General introduction. 1.6 Supplementary figures. Supplementary figure 1.1 Sequence alignment and secondary structure prediction (PRALINE software) of carotenoid 1,2-hydratase (CrtC) from Rubrivivax gelatinosus and the evolutionary distinct carotenoid 1,2hydratase (CruF) from Deinococcus radiodurans R1[46].. 24.

(35) 1.6 Supplementary figures. Supplementary figure 1.2 Sequence alignment and secondary structure prediction (PRALINE software) of carotenoid 1,2-hydratases from Rubrivivax gelatinosus and Thiocapsa roseopersicina.. 25.

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(43) . Chapter 2 2 Biochemical Characterization of the. Carotenoid 1,2-Hydratases (CrtC) from Rubrivivax gelatinosus and Thiocapsa roseopersicina Aida Hiseni, Isabel W.C.E. Arends and Linda G. Otten. Appl Microbiol Biotechnol. Aug 2011; 91(4): 1029–1036.