Study Towards Carotenoid 1,2-Hydratase and Oleate ...

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

Aida HISENI

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.

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 public-private 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.

To my father Ismet Nukičić

vii

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

viii

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

ix

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

5.4 Conclusion ................................................................................................................................. 125

5.5 Supplemental Information ......................................................................................................... 127

5.6 References .................................................................................................................................. 128

6 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

Chapter 1

1 General introduction

General introduction

2

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].

1.2 Enzymes as industrial biocatalysts

3


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

(laundry and dish wash)

Protease Protein stain removal

Amylase Starch stain removal

Lipase Lipid stain removal

Cellulase Cleaning, color clarification, anti-redeposition

(cotton)

Mannanase Mannanan stain removal (reappearing stains)

Starch and fuel Amylase Starch liquefaction and saccharification

Amyloglucosidase Saccharification

Pullulanase Saccharification

Glucose isomerase Glucose to fructose conversion

Cyclodextrin-

glycosyltransferase

Cyclodextrin production

Xylanase Viscosity reduction (fuel and starch)

Protease Free amino nitrogen production (yeast nutrition -

fuel)

Food

(including dairy)

Protease 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


4

Transglutaminase Modify visco-elastic properties

Baking Amylase Bread softness and volume, flour adjustment

Xylanase Dough conditioning

(Phospho)Lipase Dough stability and conditioning (in situ

emulsifier)

Glucose oxidase Dough strengthening

Lipoxygenase Dough strengthening, bread whitening

Protease Biscuits, cookies

Transglutaminase Laminated dough strength

Animal feed Phytase Phytate digestibility - phosphorus release

Xylanase Digestibility

β-Glucanase Digestibility

Beverage 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 Cellulase Denim finishing, cotton softening

Amylase De-sizing

Pectate lyase Scouring

Catalase Bleach termination

Laccase Bleaching

Peroxidase Excess dye removal

Pulp and paper 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 Lipase Transesterification

Phospholipase De-gumming, lyso-lecithin production

Organic synthesis 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

Leather Protease Unhairing, bating


5

Lipase De-pickling

Personal care 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.


6

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].

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).

EC 1 (Oxidoreductases)

14%EC 2

(Transferases)5%

EC 3 (Hydrolases)

75%

EC 4 (Lyases)4%

EC 5 (Isomerases)

2%

1.3 Enzyme engineering

7

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 high-

throughput 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]).


8

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 acid-

catalyzed 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

1.4 Hydro-Lyases

9

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


10

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 Enzyme name Reference

Metal ions Carbonate dehydratase (Zn2+)

Phosphopyruvate hydratase (Mg2+)

3-Dehydroshikimate dehydratase (Mn2+)

O-succinylbenzoate synthase (Mn2+ or Mg2+)

1,5-Anhydro-D-fructose dehydratase (Ca2+ or Na+ or Mg2+)

[24]

[25]

[26]

[27]

[28]

Coenzyme Propanediol dehydratase (Cyanocobalamin)

Tryptophan synthase (Pyridoxal 5'-phosphate)

[29]

[30]

Iron-sulfur Aconitate hydratase

2-Methylcitrate dehydratase

Methanogen homoaconitase

Fumarase (Class I)

[31]

[32]

[33]

[34]

CoA activated substrates Enoyl-CoA hydratase

Long-chain-enoyl-CoA hydratase

[35]

[36]

Heme-thiolate Hydroperoxide dehydratase

Colneleate synthase

[37]

[38]

NAD(P)+ / NAD(P)H CDP-glucose 4,6-dehydratase

UDP-N-acetylglucosamine 4,6-dehydratase

[39]

[40]

FAD 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.

1.4 Hydro-Lyases

11

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


12

Deinococcus [58], which are able to hydrate γ-carotene, a mono-cyclic substrate, but no

acyclic carotenoids.

OH

OH

OH

OCH3

OCH3

OH

Lycopene

1-HO-Lycopene

Neurosporene

1-HO-Neurosporene

Demethylspheroidene

Spheroidene

1-CH3O-3,4-didehydrolycopene

1-HO-3,4-didehydrolycopene

-Carotene

HOGeranylgeraniol (?)

Chlorobactene

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 2H2O-

labeling studies with intact cells [62, 63]. Until now no mutagenesis studies have been

1.4 Hydro-Lyases

13

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.


14

1.4.4 Oleate hydratase Oleate hydratase (OHase) catalyzes the conversion of oleic acid (OA) into (R)-10-

hydroxystearic 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 hydro-

lyase 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

1.4 Hydro-Lyases

15

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 FAD-

containing glutathione reductase family (GR) [80].

Figure 1.8 Flavin adenine dinucleotide (FAD) cofactor.


16

Sub

stra

te

Pro

duct

Bifidobacterium animalis subsp. Lactis BB12 [79]

Bifidobacterium breve [73]

Elizabethkingia meningoseptica [70]

Lactobacillus acidophilus NCFM [79]

Lactobacillus plantarum ST-III [79]

Lactobacillus rhamnosus LGG [79]

Lysinibacillus fusiformi [74,75]

Macrococcus caseolyticus [78]

Stenotrophomonas maltophilia [76, 77]

Streptococcus pyogenes [72]

Lau

ric

acid

(C

12, n

o do

uble

bon

d)

- -

- -

- -

- -

N

- M

yris

tic

acid

(C

14, n

o do

uble

bon

d)

- -

- -

- -

- -

N

- M

yris

tole

ic a

cid

(C14

:1, 9

Z)

10-H

ydro

xym

yris

tic

acid

-

N

- -

- -

Y

Y

Y

N

Pal

mit

ic a

cid

(C16

, no

doub

le b

ond)

-

- -

- -

- -

- N

-

Pal

mit

olei

c ac

id (

C16

:1, 9

Z)

10-H

ydro

xyhe

xade

cano

ic a

cid

- Y

-

- -

- Y

Y

Y

Y

S

tear

ic a

cid

(C18

, no

doub

le b

ond)

-

- -

- -

- -

- N

-

Pet

rose

lini

c ac

id (

C18

:1, 6

Z)

- -

- -

- -

- -

N

- E

laid

ic a

cid

(C18

:1, 9

E)

N

- -

N

N

N

- -

N

N

Ole

ic a

cid

(C18

:1, 9

Z)

10-H

ydro

xyoc

tade

cano

ic a

cid

Y

Y

Y

Y

Y

Y

Y

Y

Y

Y

Ric

inol

eic

acid

(C

18:1

, 9Z

, 12-

OH

) 10

,12-

Dih

ydro

xyst

eari

c ac

id

- -

- -

- -

Y

- -

-

Vac

ceni

c ac

id (

C18

:1, 1

1Z)

- -

- -

- -

- -

N

- co

njug

ated

-Lin

olei

c ac

id (

C18

:2, 9

E, 1

1E)

- -

- -

- -

- -

N

- co

njug

ated

-Lin

olei

c ac

id (

C18

:2, 9

Z, 1

1E)

- -

- -

- -

- -

N

- L

inol

eic

acid

(C

18:2

, 9Z

, 12Z

) 10

-Hyd

roxy

-12(

Z)-o

ctad

ecen

oic

acid

10

,13-

Dih

ydro

xyoc

tade

cano

ic a

cid

Y

N

Y

N

- Y

N

Y

N

Y

N

Y

N

Y

Y

Y

N

Y

N

L

inol

eic

acid

met

hyl e

ster

(C

18:2

, 9Z

, 12Z

) -

N

- -

- -

- -

- -

conj

ugat

ed-L

inol

eic

acid

(C

18:2

, 10E

, 12Z

)

- -

- -

- -

- -

N

- ga

mm

a-L

inol

enic

aci

d (C

18:3

, 6Z

, 9Z

, 12Z

) 10

-Hyd

roxy

-6(Z

),12

(Z)-

octa

deca

dien

oic

acid

10

,13-

Dih

ydro

xy-6

(Z)-

octa

dece

noic

aci

d

- -

- -

- -

Y

N

Y

Y

Y

N

Y

N

alph

a-L

inol

enic

aci

d (C

18:3

, 9Z

, 12Z

, 15Z

) 10

-Hyd

roxy

-12(

Z),1

5(Z)

-oct

adec

adie

noic

aci

d 10

,13-

Dih

ydro

xy-1

5(Z

)-oc

tade

ceno

ic a

cid

- -

- -

- -

Y

N

Y

Y

Y

N

N

Y

Ara

chid

ic a

cid

(C20

, no

doub

le b

ond)

-

- -

- -

- -

- N

-

Eic

osat

rien

oic

acid

(C

20:3

, 3Z

, 6Z

, 9Z

) -

- -

- -

- -

- -

N

Ara

chid

onic

aci

d (C

20:4

, 5Z

, 8Z

, 11Z

, 14Z

) -

- -

- -

- -

- -

N

Eru

cic

acid

(C

22:1

, 13Z

) -

- -

- -

- -

- N

-

Ner

voni

c ac

id (

C22

:1, 1

5Z)

- -

- -

- -

- -

N

- D

ilin

oleo

ylph

osph

atid

ylch

olin

e -

- -

- -

- -

- -

N

Tri

lino

leyl

glyc

erol

-

- -

- -

- -

- -

N

Tab

le 1

.3 O

verv

iew

of

the

subs

trat

es te

sted

with

ole

ate

hydr

atas

es r

ecom

bina

ntly

exp

ress

ed E

. col

i. N

, no

prod

uct d

etec

ted;

Y, p

rodu

ct d

etec

ted,

-, n

ot

dete

rmin

ed.

1.4 Hydro-Lyases

17

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.


18

* * *

* *

1.4 Hydro-Lyases

19

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


20

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 FAD-

binding 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.

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

D1

D2 D3

D4

1.4 Hydro-Lyases

21

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.


22

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

D1

D2 D3

D4

1.5 Scope and objectives

23

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 1,2-

hydratase 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.


24

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 1,2-

hydratase (CruF) from Deinococcus radiodurans R1[46].

1.6 Supplementary figures

25

Supplementary figure 1.2 Sequence alignment and secondary structure prediction (PRALINE software) of

carotenoid 1,2-hydratases from Rubrivivax gelatinosus and Thiocapsa roseopersicina.


26

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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

Biochemical Characterization of the Carotenoid 1,2-Hydratases (CrtC) from Rubrivivax gelatinosus and Thiocapsa roseopersicina

34

Abstract

Two carotenoid 1,2-hydratase (crtC) genes from the photosynthetic bacteria Rubrivivax

gelatinosus and Thiocapsa roseopersicina were cloned and expressed in Escherichia coli

in an active form and purified by affinity chromatography. The biochemical properties of

the recombinant enzymes and their substrate specificities were studied. The purified CrtC’s

catalyze cofactor independently the conversion of lycopene to 1-HO- and 1,1′-(HO)2-

lycopene. The optimal pH and temperature for hydratase activity was 8.0 and 30ºC,

respectively. The apparent Km and Vmax values obtained for the hydration of lycopene were

24 µM and 0.31 nmol h-1 mg-1 for RgCrtC and 9.5 µM and 0.15 nmol h-1 mg-1 for TrCrtC,

respectively. SDS-PAGE analysis revealed two protein bands of 44kDa and 38kDa for

TrCrtC, which indicate protein processing. Both hydratases are also able to convert the

unnatural substrate geranylgeraniol (C20 substrate), which functionally resembles the

natural substrate lycopene.

2.1 Introduction

35

2.1 Introduction

Optically pure tertiary alcohols are highly valuable building blocks for the synthesis of

several bioactive natural products and pharmaceuticals [1]. However, the synthesis of

optically pure tertiary alcohols in high yield without undesired side products is still a

challenging task in traditional chemical synthesis [2]. Much effort has therefore been

devoted to the development of cleaner alternative technologies. The application of

biocatalysts is recognized as a significant complement to the use of chemical reagents.

Biocatalysts such as enzymes and whole microbial cells 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. The market for enzymes has

increased in an almost exponential manner from 1960s to 2000 [3]. This is due to the well-

known benefits of enzymes. They are remarkable catalysts capable of accepting a wide

array of complex substrates, are highly selective (enantio-, regio- and chemoselective) and

operate efficiently under mild conditions.

The possibility of using enzymes for the production of tertiary alcohols has generated our

interest in the enzyme class of hydro-lyases (EC 4.2.1-), which catalyze the reversible

addition of water to a carbon-carbon double bond. Although more than 100 hydro-lyases

have been discovered to date, only a few examples have been used in industrial applications

[4, 5]. For example, for the production of R-γ-dodeca-lactone, an essential flavor in whisky,

oleate hydratase has been utilized, which catalyzes the conversion of oleic acid to form (R)-

γ-hydroxy-stearate, which again is converted to the end product by baker’s yeast [6-8].

Carotenoid 1,2-hydratase (CrtC), another member of the hydro-lyases group, occurs in the

biosynthetic pathway of different acyclic carotenoids in photosynthetic bacteria. CrtC

introduces a tertiary hydroxy group into a carotenoid molecule by addition of water to the

carbon-carbon double bond at the C-1 position. Several carotenoid 1,2-hydratases have

been identified in photosynthetic [9-13] as well as in non-photosynthetic bacteria [14, 15].

Recently, a novel carotenoid 1,2-hydratase (CruF) has been described in the non-

photosynthetic bacterium Deinococcus [15], which catalyzes C-1′,2′ hydration of γ-

carotene. This enzyme though, is evolutionarily distinct from the CrtC family in

photosynthetic bacteria.

The CrtC from the purple non-sulfur photosynthetic bacterium Rubrivivax gelatinosus has

been partially characterized and it was found to be a membrane-bound enzyme with a


36

molecular weight of 44 kDa [16]. In vitro assay showed that the enzyme was able to hydrate

the carbon-carbon double bond at the ψ-end group of several natural substrates such as

neurosporene and lycopene to the corresponding products 1-HO- and 1,1′-(HO)2-

neurosporene and 1-HO- and 1,1′-(HO)2-lycopene without the use of any cofactor. Through

genetic analysis and characterization of the pigment biosynthesis genes in the purple sulfur

photosynthetic bacterium Thiocapsa roseopersicina a putative protein was found that

showed high identity to CrtC from R. gelatinosus [11]. Gene cluster analysis of T.

roseopersicina (Gammaproteobacteria) revealed a significant identity (55 %) of the crtC

gene product to the CrtC from R. gelatinosus (Betaproteobacteria), although the

arrangement of the pigment biosynthesis gene cluster resembles more that of Rhodobacter

species (Alphaproteobacteria) [17]. However, so far the enzyme has not been isolated or

characterized in any detail, which makes it a potential candidate for a hydro-lyase with new

properties.

In order to make this group of enzymes more attractive for green hydration reactions in

industrial applications, we have investigated parameters that could be of major importance

to that field. Herein we report on the detailed biochemical characterization of the two

CrtC’s from R. gelatinosus and T. roseopersicina. This provides an insight into their

potential to be used as biocatalysts. The broad stability and activity profiles of both

enzymes are promising for industrial biocatalysis.

2.2 Materials and methods

2.2.1 Construction of pET15b_CrtCRg and pET15b_CrtCTr expression vectors

The crtCRg and crtCTr genes were amplified with primers Rg_fw/Rg_rv

(GGGAGTACCATATGCGAGCAGCGGAGTC and ATACACTCGAGATGTATACG

TCAAGCGCGG) and Tr_fw/Tr_rv (GGAGTAATCATATGCGAGCAGCGGGC and

CCCTCGAGAACTATGTCTTCT-CAGCCGCC), respectively, containing restriction

sites for NdeI (forward) and XhoI (reverse) (restriction sites are underlined). Amplification

reactions were done under standard PCR conditions using plasmids pPQE30crtCRg and

pTcrt3 respectively, as template (Supplementary table 2.1). Using NdeI/XhoI restriction

sites the digested and purified fragment was ligated into the same sites of the pET15b vector


37

and transformed into E. coli TOP10 competent cells. The insertion of the crtC gene was

verified by restriction analysis with NdeI/XhoI enzymes and DNA sequencing (BaseClear).

2.2.2 Expression and purification of recombinant proteins E. coli BL21 (DE3) was the host for the pET15_CrtC plasmids. Cultures were grown at

37°C in LB-broth with 100 µg ml-1 ampicillin until an OD600 value of 0.6-0.8 was reached.

Protein expression was induced by addition of isopropyl-β-D-thiogalactopyranoside

(IPTG) to a final concentration of 0.1 mM, followed by cultivation at 25°C overnight. The

induced cells were harvested by centrifugation at 10.000 rpm for 10 min at 4°C (Sorvall),

washed once with 50 mM Na2HPO4 buffer (pH 8.0) and suspended in the binding buffer

(50 mM Na2HPO4, 300 mM NaCl, 20 mM imidazole, pH 8.0). Cell-free extract (CFE) was

obtained after lysis of the cells with 1 mg ml-1 lysozyme for 1 h at 4ºC followed by cell

disruption at the pressure of 2.4 kBar (Constant systems, IUL instruments) and

centrifugation at 10.000 rpm for 20 min at 4°C. The separation of the CFE into membrane

fraction and supernatant was done by centrifugation at 45.000 rpm for 1 h at 4ºC.

CFE’s were filtered through 0.45 µm filter (Whatman, FP 30/0, 45 CA-S) and each extract

was applied separately onto Ni-NTA HisTrap HP column (1.6 x 2.5 cm, 5 ml, GE

Healthcare) previously equilibrated with binding buffer. The purification and the loading

of the samples onto the column were performed with the HPLC-system in conjunction with

the LCsolution software (Shimadzu). Unbound proteins were washed from the column with

a gradient of 50-75 mM imidazole in washing buffer (50 mM Na2HPO4, 300 mM NaCl,

pH 8.0). Then, the CrtC protein was eluted from the column with a gradient of 75-300 mM

imidazole in elution buffer (50 mM Na2HPO4, 300 mM NaCl, pH 8.0). Enzyme fractions

were separated by sodium dodecyl sulfate polyacrylamide gel electrophoresis (SDS-PAGE;

10% Bis-Tris, BioRad) and visualized by staining with SimplyBlue SafeStain (Invitrogen).

Fractions containing CrtC were combined and concentrated using Amicon Ultra-30 filters

(Millipore). The concentrated sample was applied onto a PD-10 desalting column (GE

Healthcare) previously equilibrated with 50 mM Na2HPO4 buffer (pH 8.0). The eluted

enzyme sample was frozen in liquid nitrogen and stored in aliquots at -80ºC.

2.2.3 Tandem MS analysis The concentrated CrtC sample was further purified by SDS-PAGE. The protein band was

excised from the gel and subjected to in-gel proteolytic digestion as previously described

[18].


38

2.2.4 CrtC activity assay and analysis of the products Enzyme activity was determined either with the purified enzyme or with the CFE. The

assay was performed with 50-100 µg enzyme in 200 µl 50 mM Na2HPO4 buffer (pH 8.0),

containing 10 mg ml-1 L-α-phosphatidylcholine (from egg yolk) and 20 µM lycopene

(Lanospharma Laboratories Co., Ltd) from a stock in acetone. After incubation at 28ºC and

shaking at 800 rpm in the dark the substrate and products were extracted from the aqueous

layer after a desired time interval. Prior to the extraction 50 µl of saturated NaCl solution

was added and the carotenoids extracted with one volume of dichloromethane. The

mixtures were shaken for 5 min at 1400 rpm, centrifuged for 1 min at 13.200 rpm

(Eppendorf) and 150 µl of the dichloromethane phase was dried with a Speed Vac

Concentrator (Thermo). The dried carotenoids were dissolved in 10 µl dichloromethane,

diluted 1:10 with 100% acetonitrile and analyzed with HPLC. Separation was performed

with a Merck 4.6x50 mm Chromolith TM SpeedROD RP-18e 2µm-column with acetonitrile

/ water (95:5, v/v) as the eluent. Lycopene and the corresponding products were detected

at 470 nm (SPD-20A, Shimadzu). Marker carotenoids were obtained as described by

Steiger et al. [19] and used for the identification of the reaction products.

The lycopene concentration in the assay was quantified from the calibration curve

constructed by diluting a stock of lycopene in dichloromethane with acetone. A second

calibration curve, which was used to quantify the reaction products, was constructed in the

same way as the standard assay, including the extraction step.

For the determination of enzyme kinetic parameters, the purified enzyme was incubated for

4 hours with different concentrations of lycopene (0.5-35 µM) in 50 mM Na2HPO4 buffer

(pH 8.0), containing 10 mg ml-1 L-α-phosphatidylcholine. Each reaction was performed in

duplicate. The affinity constant (Km) and the maximal velocity (Vmax) were calculated from

the experimental data points using OriginPro 8 SR1 software.

2.2.5 Substrate specificity Substrate specificity was assayed using the following acyclic alkenes: 2-methyl-2-butene

(79 mM), 2-methyl-2-pentene (68 mM), farnesol (33 mM) and geranylgeraniol (14.3 mM),

as substrates. Reactions were carried out using standard assay conditions. E. coli carrying

the empty pET15-b vector was used as negative control reaction. Substrates and products

were extracted from aqueous layer with one volume of ethyl acetate. The samples were

dried with Na2SO4 prior to their injection. Separation and identification of the components


39

was effected with a Shimadzu GC-MS coupled to a QP-2010S with a FactorFour VF-

WAXms column (length 30 m, diameter 0.25 mm, and film thickness 0.25 µm).

2.2.6 Effects of pH and temperature on CrtC activity In order to investigate the pH effect on the CrtC activity, the reactions were carried out in

buffers with varying pH values. The buffers used for pH test were sodium acetate (100 mM,

pH 3.0-6.0), potassium phosphate (100 mM, pH 6.0-8.0) and Tris-HCl (50 mM, pH 8.0-

9.0). The measurements were conducted at 28ºC and lycopene (20 µM) was used as

substrate, as described in the section “CrtC activity assay and analysis of the products”.

The pH stability of the enzyme was performed by measuring the remaining activity at pH

8.0 after the enzyme had been incubated in the corresponding buffers for 30 min.

The optimum temperature for CrtC activity was determined by testing enzyme activity at

temperatures ranging from 0 to 50ºC using the standard activity assay. The thermal stability

was investigated by pre-incubating the enzyme at various temperatures (5-50ºC) in the

absence of substrate for 30 min, cooling the enzyme on ice, and then measuring the residual

activity in a standard assay with lycopene as substrate.

2.2.7 Effects of inhibitors and metal ions on enzyme activity The inhibitory effects on enzyme activity were investigated by performing activity assay

under standard conditions in the presence of several metal ions (MgCl2, MnCl2, CoCl2,

ZnCl2, CaCl2 and CuSO4) and chemicals (NAD+, NADH, protease inhibitor “Complete”

(Roche)) with a final concentration of 1 mM. Lycopene (20 µM) was used as substrate and

the activity was measured as described above. Reaction mixture without any additive was

used as control reaction and was designated as 100% activity.

2.2.8 Circular dichroism (CD) spectroscopy The purified RgCrtC and TrCrtC were diluted to 0.04 and 0.03 mg ml-1, respectively, in 10

mM Na2HPO4, pH 8.0. Samples were incubated for 5 min at temperatures from 5 to 90ºC

(5°C steps) and after each incubation samples were scanned. CD spectra were collected

from 190 to 250 nm as an average of five spectra, with a data pitch of 1 nm. A band width

of 1 nm was used with a detector response time of 0.25 sec and scanning speed of 100 nm

min-1. CD spectra were recorded on a Jasco J-810 spectrometer equipped with a Peltier

temperature control unit in 0.1 cm path length cuvette [20].


40

2.2.9 Metal analysis using USN-ICP-OES The metal content from purified protein sample and the buffer solution was measured using

Perkin-Elmer 4300 dual view inductively coupled plasma (ICP) with optical emission

spectroscopy (OES) spectrometer, coupled with ultrasonic nebulizer (USN) U-6000 AT,

Cetac. Measurements were performed for different metals and at different wavelengths, as

following: Co (228 nm and 238 nm), Fe (238 nm and 239 nm), Mo (202 nm and 203 nm),

Ni (231 nm and 221 nm) and Zn (206 nm and 213 nm).

2.3 Results

2.3.1 Expression and purification of the carotenoid 1,2-hydratases

For biochemical characterization of the carotenoid 1,2-hydratases and comparison of their

catalytic activities, the two crtC genes from R. gelatinosus and T. roseopersicina were

cloned into the expression vector pET15-b. The constructed pET15b_CrtCRg and

pET15b_CrtCTr plasmids were sequenced and the results confirmed that the genes were

successfully inserted in frame with the N-terminal His6-tag. In order to express the

recombinant CrtC’s, E. coli BL21 (DE3) was transformed with the expression plasmids.

SDS/PAGE analysis revealed in both cases a 44 kDa band (Figure 2.1), which is consistent

with the value calculated from the deduced amino acid sequence.

Figure 2.1 SDS-PAGE (10%) analysis of expression and IMAC purification for RgCrtC (lane 1-3) and TrCrtC

(lane 4-6). M: precision plus protein standard; a: whole cells before induction; b: whole cells after induction

with 0.1 mM IPTG and expression overnight at 25ºC; c: purified CrtC’s.

The expression level of RgCrtC was around 2 times higher than that of TrCrtC expressed

under the same conditions. In the case of TrCrtC an additional faint band around 38 kDa

was detected after induction (Figure 2.1, lane 5), which is absent in the non-induced sample

(Figure 2.1, lane 4).

2.3 Results

41

CrtC’s were purified from CFE’s by a single step IMAC column and led to a nearly

homogenous band of 44 kDa in the case of RgCrtC and bands of 38 kDa and 44 kDa in the

case of TrCrtC (Figure 2.1, lanes 3 and 6). However, the larger band could not be detected

again, once the sample was stored at -20ºC for a few days (Figure 2.1, lane 6).

2.3.2 Hydratase activity Activity measurements of the purified enzymes with lycopene as substrate demonstrated

functional expression of the recombinant CrtC’s in E. coli. The purified enzymes catalyze

the conversion of lycopene into both 1-HO-lycopene and 1,1′-(HO)2-lycopene. For both

CrtC’s the conversion rate was 30% and the ratio between mono- and dihydroxylated

product was 2:1. USN-ICP-OES metal analysis showed that the protein samples did not

contain any significant amounts of iron, zinc, cobalt, nickel or molybdenum (data not

shown). Furthermore, it was observed that the addition of coenzymes NAD+/NADH or

protease inhibitors had no detectable influence on enzyme activity. Although the effect of

various metal ions on the hydratase activity was tested, no firm conclusion could be drawn

from these data as the metals have a degrading effect on the substrate lycopene [21].

2.3.3 Enzyme kinetics In order to compare the catalytic activities of the two expressed CrtC’s, in vitro activity

studies were performed. Since the conversion of lycopene to 1-HO-lycopene and 1,1′-

(HO)2-lycopene with isolated enzyme was very slow, the reactions were carried out with

CFE’s (Figure 2.2). Kinetic parameters Km, Vmax, Vmax/Km and kcat/Km were determined by

activity assay using lycopene as substrate at 28ºC (Table 2.1). The results are shown in a

Michaelis-Menten plot (Figure 2.3) as the reaction rate versus the substrate concentration.

Table 2.1 Kinetic parameters for recombinant Rubrivivax gelatinosus CrtC and Thiocapsa roseopersicina

CrtC

Name Vmax (nmol h-1 mg-1) Km (µM) Vmax/Km (x 102) kcat/Km (h-1 nmol-1)

RgCrtC 0.32 ± 0.08 24.7 ± 12.7 1.3 0.57

TrCrtC 0.15 ± 0.02 9.8 ± 4 1.6 0.71


42

Figure 2.2 Reaction catalyzed by Rubrivivax gelatinosus and Thiocapsa roseopersicina carotenoid 1,2-

hydratase; the conversion of lycopene into 1-HO-lycopene and 1,1′-(HO)2-lycopene (A). HPLC separation of

carotenoids formed in vitro by E.coli extract expressing the RgCrtC (solid line) and TrCrtC (dashed line).

Peak 1, 1,1′-(HO)2-lycopene; peak 2, 1-HO-lycopene; peak 3, lycopene (B).

The affinity constant (Km) for recombinant RgCrtC and TrCrtC was calculated as 24 and

9.5 µM, respectively, and Vmax was 0.31 and 0.15 nmol h-1 mg-1, respectively. The substrate

specificity values were calculated as Vmax/Km and the results show a slightly higher

specificity of TrCrtC with 1.6 x 102 compared to RgCrtC with 1.3 x 102 for lycopene.

Furthermore, the catalytic efficiency values for TrCrtC (0.71 h-1 nmol-1) and RgCrtC (0.57

h-1 nmol-1) revealed no significant difference for lycopene hydration.

0 2 4 6 8 10

0

1000

2000

3000

4000

5000

Inte

nsit

y [m

V]

Time [min]

RgCrtC TrCrtC

(A)

(B)

1

2

3

2.3 Results

43

Figure 2.3 Michaelis-Menten plot of recombinant RgCrtC (●) and TrCrtC (○). The cell-free extracts were

assayed with various lycopene concentrations (0.5-40 µM) in 50 mM Na2HPO4 sodium phosphate (pH 8.0)

at 28ºC for 4 h. The rates of product formation (1-HO-lycopene plus 1,1′-(OH)2-lycopene) are plotted against

varying substrate concentrations. Kinetic constants are listed in Table 2.1.

2.3.4 Substrate specificity Substrate specificity was tested with acyclic alkenes of different chain length, which

possess the same alkenyl functional group like lycopene, the natural substrate of CrtC

(Supplementary figure 2.1). No activity was detected for the C5, C6 and C15 substrate

using standard assay conditions. However, a product was detected with the C20 substrate

geranylgeraniol for both RgCrtC and TrCrtC, which was absent in the control experiment.

The conversion was very low, approximately 5% (Supplementary figure 2.2).

2.3.5 Effect of pH and temperature on hydratase activity and stability

The dependence of the activity of recombinant RgCrtC and TrCrtC at different pH values

and temperatures was investigated using lycopene as substrate. The optimum pH for

hydratase activity appeared to be pH 8.0 (Figure 2.4A). While RgCrtC has a broader pH

optimum ranging from pH 7.0-8.0, a significant decrease was observed for TrCrtC with

0 5 10 15 20 25 30 35 400,00

0,05

0,10

0,15

0,20

RgCrtC

V [

nmol

h-1

mg-1

]

Lycopene [M]

TrCrtC


44

only 50% activity at pH 7.0. No activity could be detected at pH 4.0-5.0 for both enzymes.

At higher pH values, both enzymes showed rapid decrease of activity, although 50% of the

relative activity was still detected at pH 9.0. Both enzymes retained much residual activity

after 30 min incubation at pH 4.0-8.0, indicating that the CrtC’s are stable in both slightly

alkaline and acid environments (Supplementary figure 2.3A). However, compared with

RgCrtC, the TrCrtC stability decreases in the range from pH 6.5-9.0 to only 56% of the

residual activity, whereas RgCrtC still remains 95% activity at pH 9.0. Despite no detected

activity at pH 4.0-5.0 (Figure 2.4A) both enzymes seem to be stable at that pH range and

still show 75%-80% of residual activity.

The effect of temperature on hydratase activity from 0 to 50ºC is depicted in Figure 2.4B.

The favorable temperature range was from 25 to 35ºC with an optimum at 30ºC. Enzyme

activity for RgCrtC and TrCrtC was significantly lower at 20ºC (55 and 42%, respectively)

and 40ºC (47 and 31%, respectively). A negligible activity was found at 5ºC (around 10%).

Thermal stability was investigated by pre-incubating hydratases for 30 min at different

temperatures and subsequently testing residual activity under standard assay conditions

(Supplementary figure 2.3B).

2.3 Results

45

Figure 2.4 Effect of pH (A) and temperature (B) on activity of RgCrtC (●) and TrCrtC (○). For pH effect

measurements were performed with lycopene under standard assay conditions using different buffers: 100

mM acetate (pH 3.6, 4.0 and 5.0), 100 mM potassium phosphate (pH 6.0, 7.0 and 8.0) and 50 mM Tris-HCl

(pH 8.6 and 9.0). For temperature effect activity assays were performed with lycopene at various temperatures

(1-50ºC) under standard assay conditions.

4 5 6 7 8 90,00

0,05

0,10

0,15

0,20 RgCrtC TrCrtC

Enz

yme

acti

vity

[nm

ol h

-1 m

g-1]

pH

0 10 20 30 40 50 600,00

0,05

0,10

0,15

0,20 RgCrtC

Enz

yme

acti

vity

[nm

ol h

-1 m

g-1]

TrCrtC

Temperature [ºC]

(A)

(B)


46

Maximum stability was recorded at 5ºC. The enzymes did not show significant decrease of

the activity up to 40ºC. When pre-incubated at 45ºC they still showed relative high activity

(around 50 to 60%). However, RgCrtC was extremely sensitive at 50ºC retaining only 6%

activity after 30 min pre-incubation, while TrCrtC showed 30% residual activity at that

temperature. Additionally, the temperature stability of RgCrtC and TrCrtC was studied

using CD spectroscopy. It was not possible to obtain a good qualitative representation of

the CD spectra from RgCrtC, in contrast to TrCrtC, which is shown in Figure 2.5.

Figure 2.5 Enzyme stability of recombinant TrCrtC by CD spectroscopy. The purified CrtC was diluted to

0.03 mg/ml with 10 mM sodium phosphate, pH 8.0, and incubated for 5 min from 5 to 90ºC. CD assay was

performed by wavelength scan from 190 to 250 nm.

TrCrtC was found to be relatively stable below 50ºC. Significant change in the secondary

structure was observed at temperatures above 50ºC, which corresponds with the results

obtained in the activity assay (Figure 2.4B).

2.4 Discussion

This study reports on the purification and biochemical characterization of two

heterologously expressed carotenoid 1,2-hydratases (CrtC) from photosynthetic bacteria,

200 205 210 215 220 225 230 235 240 245 250-20

-15

-10

-5

0

5ºC 50ºC 70ºC 90ºC

Wavelength [nm]

CD

/CD

[m

deg]

2.4 Discussion

47

which are potential biocatalysts in the green hydration of carotenoid-like substrates. The

two crtC genes from R. gelatinosus (1221 bp) and T. roseopersicina (1218 bp) were cloned,

sequenced, and successfully expressed in E. coli BL21 (DE3). Many attempts have been

made to optimize expression levels and to reduce formation of inclusion bodies (data not

shown), as these enzymes are detected in the membrane fraction. Hydropathy plots,

determined with Kyte-Doolittle [22], did not reveal any putative transmembrane domain in

the two hydratases (Supplementary figure 2.5). However, it was noticed that the first 45-55

amino acids of RgCrtC and TrCrtC showed a significantly higher percentage of proline (13

and 16 %, respectively) whereas the rest of the sequence has the usual proline amount of 9

and 8%, respectively. Ouchane and co-workers described this already for RgCrtC [23].

Proline rich regions in proteins are widely found in prokaryotes and eukaryotes [24]. A

non-repetitive (XP)n sequence like identified in RgCrtC (10x) and TrCrtC (9x) can have

different functions as for instance stabilizing the enzyme by binding noncovalently to other

proteins, binding to other hydrophobic structures like hydrophobic substrates or function

as a “molecular trigger” passing signals to the inner membrane. Based on these findings

one may suggest that the hydrophobic N-terminus of the RgCrtC and TrCrtC could play a

role in stabilizing the enzyme in the hydrophobic membrane area. This hypothesis is

strengthened by the following data. On the nucleotide level the two crtC sequences

presented a relatively high identity of 70 % [25]. However, a significant difference was

observed after the heterologous expression in E. coli. Although the gene sequence predicts

a protein of 44 kDa (Supplementary figure 2.4B), the SDS/PAGE analysis of the expressed

enzymes showed a second band of about 38 kDa for TrCrtC, which was absent in RgCrtC

as well as in the empty vector control. Furthermore, membrane fractions with only a visible

38 kDa band showed good activity (data not shown) indicating that the lower band is active.

MS data of this band revealed that the N-terminal proline rich part is missing, thereby

supporting the hypothesis that this part is not important for biological activity or substrate

binding but for membrane association. Moreover, analysis of amino acid sequence

similarities of various known and putative CrtC’s also shows that the first part of the

sequence is missing in a number of the analyzed sequences (data not shown). As this

phenomenon of protein processing is not known from literature to occur in the CrtC family,

more experiments were performed. One approach currently under study, that addresses our

hypothesis, involves construction of mutants, which lack the N-terminal part of the

sequence. First results showed that the truncated CrtC’s are fully functional and catalyze

the conversion of lycopene to the corresponding products without any loss of activity


48

(Supplementary Figure 2.7). This observation confirms our hypothesis that the N-terminal

part is not involved in catalytic reaction nor in substrate binding. Furthermore, this means

that the truncated enzyme could be used in an industrial setting.

The RgCrtC and TrCrtC catalyze the conversion of lycopene with a Vmax of 0.31 and 0.15

nmol h-1 mg-1, respectively, and a Km of 24 and 9.5 µM, respectively, without the need of

any cofactor. The lower value of Km observed for TrCrtC shows that this enzyme presents

higher affinity for the substrate lycopene than RgCrtC. However, the catalytic efficiency

values were similar for both enzymes, despite TrCrtC presenting twofold lower Vmax.

Maximum activity was detected for both enzymes at pH 8.0 and at the temperature of 30ºC.

Moreover, they also presented quite good activities at temperatures ranging from 25 to

35ºC. However, temperatures above 50ºC caused denaturation of the protein structure and

therefore inactivation, which was confirmed by CD spectroscopy. Although both enzymes

are rather similar in their pH and temperature profile, RgCrtC is more stable at higher pH’s,

while TrCrtC is more stable at higher temperatures. This could be of importance when

choosing the right enzyme for a biocatalytic process.

The substrate scope study of CrtC’s is an important aspect as no investigation has been

made in this direction to date. Next to the substrate lycopene, activity measurements were

reported in literature with two other natural substrates neurosporene and spheroidene, as

demonstrated by Steiger et al. [16]. It was concluded that spheroidene, which possess a

terminal methoxy functional group, serves as the best substrate for RgCrtC. Furthermore,

this enzyme was also able to use monohydroxy carotenoids as substrates, which could not

be observed for Rhodobacter capsulatus CrtC [16]. Our primary objective with the

substrate specificity experiment was to investigate the possibility of using CrtC with

unnatural substrates to produce highly valuable compounds for industry. Based on the

observed activity with geranylgeraniol we postulate that the minimum size of the substrates

for both RgCrtC and TrCrtC is C20 (twenty carbon) chain. However, the low conversion

of about 5% clearly indicates that their substrate spectrum is limited. Since the crystal

structure of CrtC has not yet been solved, one can only speculate about the size of the active

site and the mechanism that is involved in the hydration of the substrates. Further structural

and biochemical characterization is necessary to achieve a full understanding of this

enzyme and its reaction mechanism.

2.5 Acknowledgments

49

In conclusion, both CrtC’s are stable at a broad and suitable temperature and pH range and

hydrate several long aliphatic substrates to give tertiary alcohols. Future studies will be

directed at improving the activity of these hydratases.

2.5 Acknowledgments

We thank Prof. Dr. Gerhard Sandmann and Prof. Dr. Kornél L. Kovács for providing the

plasmids. This project is financially supported by the Netherlands Ministry of Economic

Affairs and the B-Basic partner organizations (www.b-basic.nl) through B-Basic, a public-

private NWO-ACTS programme (ACTS = Advanced Chemical Technologies for

Sustainability).


50

2.6 Supplementary information

Supplementary table 2.1 Bacterial strains and plasmids used in this study

Strain and plasmid Relevant trait(s) Source or reference

Strains

E. coli BL21 (DE3) F– ompT gal dcm lon hsdSB(rB- mB

-) λ(DE3) Novagen

E. coli TOP10 F- mcrA Δ(mrr-hsdRMS-mcrBC) φ80lacZΔM15

ΔlacX74 nupG recA1 araD139 Δ(ara-leu)7697

galE15 galK16 rpsL(StrR) endA1 λ-

Invitrogen

Plasmids

pPQE30crtCRg pPQ30; carries the BamHI / KpnI fragment with

crtC from Rvi. gelatinosus

[16]

pTcrt3 pBluescript SK(+); carries the wild-type BamHI-

SacI fragment of the crtDC operon of Tca.

roseopersicina

[11]

pET15-b E. coli general expression vector with N-

terminal His tag; Ampr

Novagen

pET15b_CrtCRg pET15-b with 1252-bp NdeI / XhoI fragment

from pPQE30crtCRg

this work

pET15b_CrtCTr pET15-b with 1249-bp NdeI / XhoI fragment

from pTcrt3

this work


51

Supplementary figure 2.1 Structure of substrates used for substrate specificity studies of Rvi. gelatinosus and

Tca. roseopersicina carotenoid 1,2-hydratase.

Supplementary figure 2.2 GC separation of products formed in vitro by Escherichia coli extract expressing

RgCrtC (pink line) and TrCrtC (blue line) using the substrate geranylgeraniol. Obtained product is indicated

with arrow (RT 41 min). Extract with empty plasmid pET15-b served as negative control (black line).

5.0 10.0 15.0 20.0 25.0 30.0 35.0 40.0 45.0 50.0 55.0 60.0 65.0 70.00.00

0.25

0.50

0.75

1.00

1.25

1.50

1.75

2.00

2.25

2.50

2.75

3.00

3.25

3.50

3.75

4.00

4.25

4.50

4.75

(x1,000,000)


52

Supplementary figure 2.3 Stability of RgCrtC (●) and TrCrtC (○) at different pH (A) and temperature (B)

values. The remaining activity was assayed under standard assay conditions after the cell-free extracts had

been incubated in the corresponding buffers (pH 3.6 to pH 9.0) or at the indicated temperature (5-50ºC) in 50

mM Na2HPO4 sodium phosphate (pH 8.0) for 30 min

3 4 5 6 7 8 9 100,00

0,01

0,02

0,03

0,04 RgCrtC

Enz

yme

activ

ity

[nm

ol h

-1 m

g-1]

pH

TrCrtC

0,00

0,05

0,10

0,15

0,20

0 10 20 30 40 50 60

RgCrtC

Enz

yme

acti

vity

[nm

ol h

-1 m

g-1]

TrCrtC

T [ºC]

(A)

(B)


53

Supplementary figure 2.4 Amino acid sequence alignment of RgCrtC and TrCrtC (Clone Manager 9

Professional Edition). Identical amino acids are highlighted in red.


54

Supplementary figure 2.5 Hydropathy plot of the RgCrtC (A) and TrCrtC (B) amino acid sequence.

Hydropathy scores [22] for a window of 19 residues were averaged and assigned to the first amino acid of

the window. A hydropathy score greater than 1.6 indicates transmembrane region.

(A)

(B)


55

Supplementary figure 2.6 DNA sequence alignment of RgCrtC and TrCrtC (Clone Manager 9 Professional

Edition). Identical bases are highlighted in red.


56

M 1a 1b 2a 2b 3a 3b 4a 4b 5a 5b

Supplementary figure 2.7 SDS-PAGE (10%) analysis (A) and amino acid sequence alignment of the wildtype

and the truncated CrtC (B). M: precision plus protein standard; a: whole cells before induction; b: whole cells

after induction with 0.1 mM IPTG and expression overnight at 25°C; 1: pET15-b; 2: TrCrtC wildtype; 3:

TrCrtC truncated; 4: RgCrtC wildtype; 5: RgCrtC truncated.

50

37

(A)

(B)

2.7 References

57

2.7 References

[1] R. Kourist, et al., "Enzymatic synthesis of optically active tertiary alcohols: Expanding the

biocatalysis toolbox," Chembiochem, vol. 9, pp. 491-498, Mar 3 2008.

[2] P. G. Cozzi, et al., "Enantioselective catalytic formation of quaternary stereogenic centers,"

European Journal of Organic Chemistry, pp. 5969-5994, Dec 2007.

[3] P. Fernandes, "Miniaturization in Biocatalysis," International Journal of Molecular Sciences, vol.

11, pp. 858-879, Mar 2010.

[4] D. Brady, et al., "Characterisation of nitrilase and nitrile hydratase biocatalytic systems," Applied

Microbiology and Biotechnology, vol. 64, pp. 76-85, Mar 2004.

[5] K. Rzeznicka, et al., "Cloning and functional expression of a nitrile hydratase (NHase) from

Rhodococcus equi TG328-2 in Escherichia coli, its purification and biochemical characterisation,"

Applied Microbiology and Biotechnology, vol. 85, pp. 1417-1425, Feb 2010.

[6] S. Gocho, et al., "BIOTRANSFORMATION OF OLEIC-ACID TO OPTICALLY-ACTIVE

GAMMA-DODECALACTONE," Bioscience Biotechnology and Biochemistry, vol. 59, pp. 1571-

1572, Aug 1995.

[7] A. Wanikawa, et al., "Detection of gamma-lactones in malt whisky," Journal of the Institute of

Brewing, vol. 106, pp. 39-43, Jan-Feb 2000.



[9] G. A. Armstrong, et al., "Nucleotide-sequence, organization, and nature of the protein products of

the carotenoid biosynthesis gene-cluster of Rhodobacter-capsulatus," Molecular & General

Genetics, vol. 216, pp. 254-268, Apr 1989.

[10] H. P. Lang, et al., "Complete DNA-sequence, specific Tn5 insertion map, and gene assignment of

the carotenoid biosynthesis pathway of Rhodobacter-sphaeroides," Journal of Bacteriology, vol.

177, pp. 2064-2073, Apr 1995.

[11] A. T. Kovacs, et al., "Genes involved in the biosynthesis of photosynthetic pigments in the purple

sulfur photosynthetic bacterium Thiocapsa roseopersicina," Applied and Environmental

Microbiology, vol. 69, pp. 3093-3102, Jun 2003.



[13] N. U. Frigaard, et al., "Genetic manipulation of carotenoid biosynthesis in the green sulfur bacterium

Chlorobium tepidum," Journal of Bacteriology, vol. 186, pp. 5210-5220, Aug 2004.

[14] J. A. Botella, et al., "A cluster of structural and regulatory genes for light-iduced carotenogenesis in

Myxococcus-xanthus," European Journal of Biochemistry, vol. 233, pp. 238-248, Oct 1995.




58

[16] S. Steiger, et al., "Heterologous expression, purification, and enzymatic characterization of the

acyclic carotenoid 1,2-hydratase from Rubrivivax gelatinosus," Archives of Biochemistry and

Biophysics, vol. 414, pp. 51-58, Jun 1 2003.

[17] N. Igarashi, et al., "Horizontal transfer of the photosynthesis gene cluster and operon rearrangement

in purple bacteria," Journal of Molecular Evolution, vol. 52, pp. 333-341, Apr 2001.

[18] A. M. Sevcenco, et al., "Development of a generic approach to native metalloproteomics: application

to the quantitative identification of soluble copper proteins in Escherichia coli," Journal of

Biological Inorganic Chemistry, vol. 14, pp. 631-640, May 2009.

[19] S. Steiger, et al., "Substrate specificity of the expressed carotenoid 3,4-desaturase from Rubrivivax

gelatinosus reveals the detailed reaction sequence to spheroidene and spirilloxanthin," Biochemical

Journal, vol. 349, pp. 635-640, Jul 15 2000.

[20] Y. H. Chen, et al., "Determination of secondary structures of proteins by Circular-Dichroism and

Optical Rotatory Dispersion," Biochemistry, vol. 11, pp. 4120-&, 1972.

[21] C. S. Boon, et al., "Role of Iron and Hydroperoxides in the Degradation of Lycopene in Oil-in-Water

Emulsions," Journal of Agricultural and Food Chemistry, vol. 57, pp. 2993-2998, Apr 2009.

[22] J. Kyte and R. F. Doolittle, "A Simple Method for Displaying the Hydropathic Character of a

Protein," Journal of Molecular Biology, vol. 157, pp. 105-132, 1982.

[23] S. Ouchane, et al., "Pleiotropic effects of puf interposon mutagenesis on carotenoid biosynthesis in

Rubrivivax gelatinosus - A new gene organization in purple bacteria," Journal of Biological

Chemistry, vol. 272, pp. 1670-1676, Jan 17 1997.

[24] M. P. Williamson, "The Structure and Function of Proline-Rich Regions in Proteins," Biochemical

Journal, vol. 297, pp. 249-260, Jan 15 1994.

[25] S. F. Altschul, et al., "Basic Local Alignment Search Tool," Journal of Molecular Biology, vol. 215,

pp. 403-410, Oct 5 1990.

Chapter 3

3 Structural Characterization of the Carotenoid 1,2-Hydratases (CrtC) from

Rubrivivax gelatinosus and Thiocapsa roseopersicina

Aida Hiseni, Linda G. Otten and Isabel W.C.E. Arends

Submitted

Structural Characterization of the Carotenoid 1,2-Hydratases (CrtC) from Rubrivivax gelatinosus and Thiocapsa roseopersicina

60

Abstract

Carotenoid 1,2-hydratases (CrtC) catalyze the selective addition of water to an isolated

carbon-carbon double bond. Although their involvement in the carotenoid biosynthetic

pathway is well understood, little is known about the mechanism by which these hydratases

transform carotenoids such as lycopene into corresponding hydroxyl compounds. To gain

insight into the enzymatic mechanism of CrtC’s point mutants of selected conserved amino

acids were generated. Rubrivivax gelatinosus CrtC point mutants in which each of the

amino acids His239, Trp241, Tyr266 and Asp268 were individually changed into Ala, and

the corresponding point mutants of Thiocapsa roseopersicina CrtC, were completely

inactive. This result suggests the identification of key residues which are directly involved

in the catalytic reaction. Furthermore, the analysis of a partial 3D structure of CrtC, which

was obtained by homology modeling with the putative AttH protein from Nitrosomonas

europaea, supported these results as all amino acids were in close distance to each other.

These results for the first time shed light on the potential catalytic mechanism of CrtC.

3.1 Introduction

61

3.1 Introduction

Carotenoids, which represent one of the most abundant natural pigments with structural

and protective properties [1], play an essential role in the photosynthetic machinery of

phototrophic organisms such as purple bacteria [2] and higher plants [3]. In addition, they

have been identified in fungi and some non-photosynthetic bacteria [4]. Carotenoid 1,2-

hydratase (also known as CrtC) is a member of hydro-lyases group EC 4.2.1.131. The

enzyme takes part in the biosynthetic pathway of carotenoids [5]. CrtC introduces a tertiary

hydroxyl group into an acyclic carotenoid molecule by addition of water to the carbon-

carbon double bond at the C-1 position. The enzyme belongs to the Pfam family PF07143

that encompasses members from several purple photosynthetic bacteria. On the other hand,

CrtC’s have been identified, which are able to hydrate mono-cyclic carotenoid gamma-

carotene. These are evolutionary very distinct from the PF07143 members [6] and they

have been given the name CruF.

Recently, two representatives of the PF07143 family, the CrtC’s from purple non-sulfur

Betaproteobacteria Rubrivivax gelatinosus and purple sulfur Gammaproteobacteria

Thiocapsa roseopersicina, respectively, were recombinantly expressed and characterized

[7]. Biochemical studies have revealed that these enzymes are able to convert cofactor

independently lycopene into 1-HO-lycopene and 1,1’-(HO)2-lycopene (Figure 3.1).

Figure 3.1 Reaction catalyzed by Rubrivivax gelatinosus and Thiocapsa roseopersicina carotenoid 1,2-

hydratase; the conversion of 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 (Figure 3.2).

Figure 3.2 Molecular structure of geranylgeraniol.


62

CrtC’s are appealing enzymes in the biotechnology field because they are able to make a

tertiary alcohol, a highly valuable building block for the synthesis of several bioactive

natural products and pharmaceuticals [8]. Furthermore, they possess an intrinsically high

stability at a wide pH and temperature range, which constitute useful properties for an

industrial application [7]. The subcellular location of this enzyme in the cell membrane

fraction (membrane-bound) allows for a straightforward isolation and simplified large scale

purification.

From a chemical point of view, CrtC’s are able to perform a challenging chemical reaction,

namely the selective addition of water to an isolated carbon-carbon double bond [9]. The

reaction proceeds without assistance of electron withdrawal groups, or transition metal

cations and in vitro requires harsh acidic conditions [10]. Furthermore, the CrtC’s from

photosynthetic bacteria act on acyclic carotenoids, whereas the CruF’s from non-

photosynthetic bacteria catalyze the hydration of mono-cyclic carotenoids. Nevertheless,

the catalytic and structural features, that determine hydratase activity and specificity of

these two distinct families remains unknown.

To our knowledge no published data exist on the catalytic mechanism of this group of

enzymes, 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 [11]. The mechanism of lycopene

hydration, which involves proton attack at C-2 and C-2′ with a carbonium ion intermediate

and the introduction of the hydroxyl group at C-1 and C-1′, was established from 2H2O-

labeling studies with intact cells [12, 13]. For a hydration reaction, it is likely to assume

that the first step in the reaction is protonation of the alkene, leading to an intermediate

carbocation. Quenching of the carbocation by water will lead to the alcohol as product. The

protonation of hydrophobic long-chain alkenes has also been described for the enzyme

class of cyclases, of which the full mechanism is known [14, 15].

The objective of this study was to provide an insight into the possible hydration mechanism

of CrtC’s. Based on the better knowledge of the mechanistic reaction, it might be possible

to improve enzyme activity or substrate scope by for instance directed evolution or

(semi)rational design. Through multi-sequence alignment of several CrtC homologues,

highly conserved amino acids were identified, which could be functionally or structurally

important. The corresponding alanine mutants of these amino acids were produced and in

this way their involvement in the hydratase activity could be evaluated. Furthermore, a


63

homology model of CrtC was obtained by using the putative AttH protein from

Nitrosomonas europaea as a template [11]. Following the identification of catalytically

active amino acid residues the aim was to propose a catalytic mechanism for CrtC catalyzed

water addition.


3.2.1 In silico analysis BLAST [16] was used to select carotenoid 1,2-hydratase homologues. In order to look for

identities/similarities between the CrtC homologues, nucleotide and amino acid sequences

were aligned with the BioEdit Sequence Alignment Editor v.7.1.3.0

(www.mbio.ncsu.edu/bioedit/bioedit.html). In addition, protein sequences were subjected

to protein functional analysis using Conserved Domain Search (CDD) [17] and Pfam search

[18], and a protein phylogenetic tree was constructed with Phylogeny.fr [19, 20]. The CrtC

secondary structure prediction was carried out with the program PolyView 2D [21]. The

SWISS-MODEL program was used to model the structure of CrtC [22].

3.2.2 Cloning of carotenoid 1,2-hydratase genes Plasmids pET15b_CrtCRg and pET15b_CrtCTr containing CrtC from R. gelatinosus (Rg)

and T. roseopersicina (Tr), respectively, were constructed in a previous study [7]. Two

fosmids with crtC genes from metagenomic samples DelRiverFos06H03 (Fos06) and

DelRiverFos13D03 (Fos13), respectively, were kindly provided by Dr. Kirchman [23]. The

cosmid encoding CrtC from Bradyrhizobium (Br) was received from Dr. Dreyfus [24]. In

order to get sufficient DNA material for further studies, the fosmid- and cosmid DNA were

amplified in E. coli TOP 10 cells. After DNA isolation with the QIAprep Spin Miniprep

Kit (Qiagen) from the cells sufficient DNA was obtained for further research. The crtC’s

from Rhodospirillum rubrum (Rr) and Rhodopseudomonas palustris (Rp) were amplified

from genomic DNA. For that, genomic DNA of R. rubrum (Rr) was kindly provided by

Prof. Roberts (NCBI Reference Sequence: NC_007641.1). R. palustris cells (DSM No.

123) were obtained from DSMZ (Deutsche Sammlung von Mikroorganismen und

Zellkulturen), enriched in appropriate medium according to DSMZ instructions and gDNA

isolated using the UltraClean Soil DNA Isolation Kit (Mobio). Subsequently, primers were

designed for the isolation of all crtC genes (Table 3.1), which carry two restriction sites for


64

subsequent cloning: NdeI (forward) and XhoI (reverse). For BrcrtC the XhoI-site was

replaced with BamHI, because the XhoI-site was present in the gene itself. Amplification

reactions were done using standard PCR reactions. Using appropriate restriction sites, the

digested and purified fragment was ligated into the same sites of the pET15-b vector and

transformed into E. coli TOP10 competent cells. The insertion of the crtC gene was verified

by restriction analysis with the corresponding restriction enzymes (New England Biolabs)

and DNA sequencing (BaseClear).

Table 3.1 Primers used in this study. The respective restriction sites are underlined.

Name Sequence (5'→3') Restriction site

DRF06 FW GGGAGTACCATATGAGTGATGATGGCCAAC NdeI

DRF06 RV ATCCGCTCGAGATAATCTCAAGCCCGCCTCG XhoI

DRF13 FW GGGAGTACATATGGATGGCGTGTCAGAC NdeI

DRF13 RV CCGCTCGAGTAATGCTTAGGGCCACTTGGC XhoI

Br FW CGGACATCATATGTGCCCGCCAG NdeI

Br RV ATCCAGGATCCATCGCGTGAACTTCACCACC BamHI

Rp FW CGGGACTTCCATATGTCAGGAGCTGAGTTG NdeI

Rp RV ACCGCTCGAGTAACGTTCAGCGGAACGC XhoI

Rr FW GGGAAATTCCATATGCACCGCCCGGAC NdeI

Rr RV GCTCGAGTTCAATTAGCCCTTAACCGCCGC XhoI

3.2.3 Construction of CrtC mutants

3.2.3.1 Single point mutations Single amino acid exchange within the crtC genes of Rg and Tr was done by the

megaprimer PCR method introduced by Kammann et al. [25] and later modified by Sarkar

and Sommer [26] and Landt et al. [27]. The mismatch primers are listed in Table 3.2. In

the first PCR reaction, performed under standard reaction conditions, the megaprimer was

produced using the corresponding forward primer containing the desired base substitution

(Table 3.2) in combination with the reverse primer Rg_rv [7] and Tr_rv [7], respectively.

Plasmids pET15b_CrtCRg and pET15b_CrtCTr [7] were used as template. The size and

purity of the megaprimer was verified by agarose gel electrophoresis. In order to produce

the full length gene, a second PCR reaction was performed with the corresponding

megaprimer and Rg_fw [7] or Tr_fw [7], respectively. Subsequent steps were performed

as described in previous section. The insertion of the crtC gene and the presence of the


65

desired mutation were verified by restriction analysis with NdeI/XhoI enzymes and DNA

sequencing (BaseClear).

Table 3.2 Primers for site directed mutagenesis. Mismatch points are underlined.

Amino acid

exchange Sequence (5'→3')

R. g

elat

inos

us

H239A AGCGGCGGACGCGCTCGCTG

W241A CATCGCGCGGGGCCGATCG

H264A CTGGAGCGGCGCCGCCTACC

Y266A GCCACGCCGCCCTCGACT

D268A CGCCTACCTCGCCTCGAACGAAG

T. r

oseo

pers

icin

a

H237A GATCCGGCGGAACGCGCAGTCTGGTGG

W239A CGCCATGTCGCGTGGCCGATC

H262A GCTGGAGCGGCGCTGGCTAT

D266A CATGGCTATCTCGCCTCAAA

S58V GCGTCCGTCGTCGCGCAGCA

S58Q GCGTCCCAGGTCGCGCAGCA

3.2.3.2 N-terminally truncated Rg- and TrCrtC’s Rg- and TrCrtC lacking the first 45 and 57 amino acids, respectively, were constructed

using primers Rg_45aa (Table 3.3)/Rg_rv [7] and Tr_57 (Table 3.3)/Tr_rv [7] under

standard PCR conditions. Plasmids pET15b_CrtCRg and pET15b_CrtCTr [7], respectively,

were used as template. Subsequent steps were performed as described in the section

“Cloning of carotenoid 1,2-hydratase genes”. The insertion of the crtC gene was verified

by restriction analysis with the corresponding restriction enzymes (New England Biolabs)

and DNA sequencing (BaseClear).

Table 3.3 Primers used for construction of truncated CrtC’s. The NdeI restriction sites are underlined.

Name Sequence (5'→3')

Rg_45aa AGTACCATATGGGCGACGCACGGCTGG

Tr_57aa AGTACCATATGTCCGTCGCGCAGCAAGG

3.2.4 Recombinant expression of CrtC’s E.coli BL21 (DE3) was the host for the pET15_CrtC plasmids. Cultures were grown at

37°C in Luria–Bertani broth with 100 μg ml−1 ampicillin until an OD600 value of 0.6–0.8

was reached. Unless otherwise stated, protein expression was induced by addition of

isopropyl-β-D-thiogalactopyranoside (IPTG) to a final concentration of 0.1 mM, followed


66

by cultivation at 25°C overnight. The cells were harvested by centrifugation at 10.000 rpm

for 10 min at 4°C (Sorvall), washed once with 50 mM Na2HPO4 buffer (pH 8.0), and

suspended in the same buffer. In case of subsequent purification, 10 mM imidazole was

added to the buffer. Crude extract (CE) from cultures >100 ml was prepared by adding 1

mg ml−1 lysozyme and incubating the cells for 1 h at 4°C, followed by cell disruption at the

pressure of 1.5 kBar (Constant systems, IUL instruments). For cultures <100 ml, the cells

were disrupted by sonication for 2 min while immersed in an ice-water bath using the

microtip probe of a sonicator (Branson Sonicator Cell Disruptor) set at 50% maximal

energy. In an effort to reduce the liquid viscosity caused by DNA molecules, 0.1 mg ml-1

of DNAse was added. With the subsequent centrifugation at 10.000 rpm for 20 min at 4°C,

cell-free extract (CFE) and pellet were separated. Protein content of the crude extract was

determined by BCA assay (Pierce) with bovine serum albumin as the reference protein.

3.2.5 CrtC purification Rg- and TrCrtC ‘active site’ point mutants were purified from the membrane fraction, while

the Tr ‘processing’ mutants (S58V, S58Q) were purified from the CFE. The membrane

fraction was obtained after the centrifugation of the CFE for 4 h at 13.200 rpm and 8°C.

Prior to the addition of Ni-NTA HisTrap HP (GE Healthcare) (previously equilibrated in

50 mM Na2HPO4 buffer, pH 8.0, with 300 mM NaCl and 10 mM imidazole), to the CFE

or membrane sample, the membranes were homogenized by ca. 20 passages through a 25G

needle. The mixtures were incubated for 1 h at RT, loaded into a polypropylene tube with

porous disc (GE Healthcare) and washed 3 times with washing buffer (50 mM Na2HPO4

buffer, pH 8.0, with 300 mM NaCl and 75 mM imidazole). Then, the CrtC protein was

eluted from the column with elution buffer containing 1 M imidazole (50 mM Na2HPO4

buffer, pH 8.0, with 300 mM NaCl). Enzyme fractions were separated by SDS-PAGE (10%

Bis-Tris, BioRad) and visualized by staining with SimplyBlue SafeStain (Invitrogen).

3.2.6 Determination of enzyme activity Enzymatic activities were determined with CE on lycopene and geranylgeraniol (GGOH)

according to the method described earlier [7], with few modifications. The assay was

carried out with 50 μl CE and 20 μM substrate, and 10 mg ml−1 L-α-phosphatidylcholine

in the case of lycopene, in a reaction volume of 200 μl. In addition to the GC-MS method,

the GGOH reaction products were also analyzed with a newly developed HPLC method.

Prior to the analysis, acetonitrile was added to the reaction mixture in a ratio of 60:40

3.3 Results and discussion

67

(ACN:H2O), the mixtures were shaken vigorously for 1 min and solids removed by

centrifugation for 1 min at 13.200 rpm. Separation of the reaction products was performed

with a Merck 4.6× 50 mm Chromolith TM SpeedROD RP-18e 2 μm column with

ACN/H2O (60:40, v/v) as the eluent. GGOH and the corresponding products were detected

at 214 nm (SPD-20A, Shimadzu).


3.3.1 Comparative in silico analysis of crtC genes The RgcrtC nucleotide sequence was subjected to a BLAST search in order to identify

sequence similarity in different databases. 184 hits were identified, of which 111 were

representatives of Proteobacteria. Although, R. gelatinosus belongs to the

Betaproteobacteria, more than 77% of the identified 111 hits were from

Alphaproteobacteria and only 11% from Betaproteobacteria. Similarly, Igarashi et al. [28]

observed that most of the photosynthesis gene products from R. gelatinosus showed high

sequence identities to the gene products of R. palustris, an Alphaproteobacteria member.

They explain this occurrence as a result of horizontal transfer of the photosynthesis gene

clusters from an ancestral species belonging to the Alphaproteobacteria to that of the

Betaproteobacteria.

The selection of RgCrtC homologues for this study was based on sequence identity and

availability of the corresponding gene construct. They originate from all three

Proteobacteria subclasses (Alpha, Beta, Gamma) with two additional constructs originating

from metagenomic samples from the Delaware River (USA). Figure 3.1 displays a

phylogenetic analysis constructed with protein sequences of the selected CrtC homologues.

TrCrtC shows the closest relationship to RgCrtC, followed by BrCrtC (55% and 47%

sequence identity, respectively). The combined results of Pfam- and Conserved Domain

Search showed that all 7 CrtC’s belong to the PF07143 family consisting of several purple

photosynthetic bacterial hydroxyneurosporene synthase (CrtC) proteins.


68

Figure 3.1 Rooted phylogenetic tree showing the evolutionary relationship between the selected carotenoid

1,2-hydratases. TrCrtC, Thiocapsa roseopersicina (GI 31621263), BrCrtC, Bradyrhizobium sp. BTAi1 (GI

146403799), F06CrtC, uncultured Proteobacterium DelRiverFos06H03 (GI 61653228), F13CrtC, uncultured

Proteobacterium DelRiverFos13D03 (GI 61653190), RpCrtC, Rhodopseudomonas palustris (GI

115515977), RrCrtC, Rhodospirillum rubrum (GI 83574254) and RgCrtC, Rubrivivax gelatinosus (GI

29893477).

CrtC sequences were aligned in order to investigate if there are any conserved group-

clusters present (Figure 3.2). Indeed, they showed highly conserved regions of in total 64

amino acids. The conserved amino acids are distributed along the sequence ranging from

amino acid residues ~115 to ~405. Interestingly, the N-terminal part of the sequence does

not contain any conserved amino acids, indicating that this region is probably not necessary

for CrtC activity. This could also be the explanation for the absence and the shorter DNA

sequences for Fos06, Fos13 and partly RpCrtC, when compared to RgCrtC or TrCrtC. In

addition, we found little amino acid variety in 50 positions, as indicated within the boxes

in Figure 3.2.

Residues involved in the catalysis tend to be highly conserved in a set of homologous

proteins that exhibit the same reaction. On the other hand, sequence insertion and sections

of low sequence similarity tend to occur in the less important loop regions [29]. The

recognition of conserved blocks in CrtC homologues led to the obvious hypothesis that

these regions contain the amino acid residues most important for the hydratase activity,

specifically those involved in catalysis and substrate binding.


69

Figure 3.2 Multiple sequence alignment showing conserved amino acids of the CrtC protein sequences from

various bacteria. Identical amino acids are highlighted in black. Positions with only two different amino acids

are surrounded by boxes.

The 3D structure of TrCrtC was built by homology modeling based on the only known 3D

structure, which showed some sequence identity (17%) to the CrtC (Figure 3.3). The

homologue is the putative AttH protein from Nitrosomonas europaea [11]. It belongs to a

protein family of unknown function (DUF2006), which has remote similarity to the family

PF07143 encompassing carotenoid 1,2-hydratases. The topology of the CrtC structure

shows similarity to lipocalins, proteins that bind and transport small hydrophobic molecules

[30]. Lipocalin fold is typically formed by a large, twisted beta-sheet that closes in the back

to form a central, internal, ligand-binding cavity. This folding motif is frequently found in

porins, transmembrane proteins or in general in proteins that bind hydrophobic

ligands/substrates [31]. Depending on the protein and the corresponding function, the

bound ligand will be entirely within the cavity or part of the ligand will protrude from the

cavity at the surface of the protein.


70

Figure 3.3 A homology model of CrtC from Thiocapsa roseopersicina based on the crystal structure of a

putative AttH (PDB id: 2ICH) from Nitrosomonas europaea. The ribbon diagrams depict front (left-hand

side) and back (right-hand side) view of CrtC. The structure is color-coded from the N-terminus (blue) to the

C-terminus (red).

In order to investigate if there is any relationship between the conserved regions and

specific locations in the homology model of CrtC, the conserved residues were visualized

(Figure 3.4). Six striking sequence motifs were selected (I – VI) and the results show that,

indeed, most of the conserved residues are located either at the bottom or upper part of the

cavity, with one motif (III) located outside the cavity. This order is very interesting and

might indicate regions in the CrtC structure, which are involved in the binding of the

substrates, while the other might contain amino acids that are directly involved in the

catalytic reaction. From a catalytic point of view, amino acids located in regions IV and V,

i.e. aspartic acid (D), tyrosine (Y) and histidine (H), are most probably involved in the

catalytic hydration, as these amino acids are commonly involved as active residues in acid-

base type catalyzed reactions in the active sites of enzymes (Figure 3.5) [32]. Furthermore,

they are all in close distance to each other, which is important for the contact with the

substrate. It would thus be interesting to analyze these positions by mutagenesis, in order

to confirm their involvement in the catalytic process of CrtC’s.


71

RgCrtC …............SDDG…….GSVFSP.Y……...........GPS........H.W..........H.Y.D.........................PFY

TrCrtC …............SDDG…….GSVFSP.Y……...........GPS........H.W..........H.Y.D.........................PFY

BrCrtC …............SDDG…….GSVFSP.Y……...........GPS........H.W..........D.Y.D.........................PFY

RrCrtC …............SDDG…….GSVFSP.Y……...........GPS........H.W..........H.Y.D.........................PFY

RpCrtC …............SDDG…….GSVFSP.Y……...........GPS........H.W..........H.Y.D.........................PFY

Fos13CrtC.............SDDG…….GSVFSP.Y……...........GPS........H.W..........H.Y.D.........................PFY

Fos06CrtC.............SDDG…….GSVFSP.Y……...........GPS........H.W..........H.Y.D.........................PFY

Figure 3.4 CrtC conserved residues. Ribbon diagram of TrCrtC with marked regions that contain highly

conserved amino acid residues. The sequence motifs, which correspond to those regions, are shown in boxes

(I – VI). N and C indicate N-and C-terminus, respectively.

I II III IV V IV VI

I

II

III

V

IV

VI

N

C


72

Figure 3.5 View of the potential active site of TrCrtC. The conserved residues H237 (yellow), W239 (red),

Y264 (blue) and D266 (green) are show as sticks, with hydrogen bond as yellow dots (region IV).

3.3.2 Production of recombinant wildtype and mutant CrtC’s and enzymatic activity

6 out of the 7 selected CrtC’s were overexpressed from pET15b in E. coli (Figure 3.6).

Bands with apparent molecular weight of 32 kDa (Fos13CrtC), 38 kDa (RpCrtC) and 44

kDa (Rr-, Br-, Tr- and RgCrtC) were visualized on SDS-PAGE and were consistent with

the values calculated from the deduced amino acid sequences. TrCrtC is expressed as 44-

and 38 kDa protein [7].

Figure 3.6 SDS-PAGE (10%) analysis of CrtC expression in E. coli BL21. M, Precision plus protein standard.

First lane (a) of each sample shows cells before induction with 0.6 mM IPTG and the second lane (b) shows

cells after 4 h expression at 37°C. 1, pET15-b control; 2, Fos06CrtC (32 kDa); 3, Fos13CrtC (32 kDa); 4,

RpCrtC (38 kDa); 5, RrCrtC (44 kDa); 6, BrCrtC (44 kDa); 7, TrCrtC (44 kDa); 8, RgCrtC (44 kDa). The

indicated molecular weights are deduced from amino acid sequences. CrtC expression bands are indicated by

arrows.

M 1a 1b 2a 2b 3a 3b 4a 4b 5a 5b 6a 6b 7a 7b 8a 8b M

50

37

25


73

No expression band could be identified for Fos06CrtC. Although, relatively good

expression was achieved for most of the CrtC’s, only two were active with lycopene as

substrate (data not shown). The fact that all CrtC’s share highly conserved regions in the

amino acid sequence (Figure 3.2) indicates that they are performing the same or similar

biochemistry. However, no activity whatsoever could be detected for 5 CrtC’s in the

standard lycopene hydration assay. At this point it is unclear whether this is due to reasons

of low activity in the cell-extract and/or substrate specificity.

In our previous study, the two active CrtC’s from R. gelatinosus and T. roseopersicina were

biochemically characterized and we showed that both CrtC’s have the ability to convert

acyclic carotenoid lycopene into hydroxyl derivatives [7]. Furthermore, we reported on the

activity of both CrtC’s with the substrate geranylgeraniol, a C20 acyclic alkene molecule

containing a hydroxyl group at one end and -group (acyclic C9 end group according to

nomenclature of carotenoids) at the other end of the chain [7]. Unfortunately, the product

could not be identified due to low yields.

In order to get more insight into the hydration mechanism of CrtC’s, selected amino acid

residues in regions IV and V (Figure 3.4) were substituted by the amino acid alanine. The

selection was based on the fact that amino acids such as aspartic acid and histidine occur

more frequently in enzyme active sites than others [32]. In addition, truncated (Tr- and

RgCrtC) and N-terminal point mutants (TrCrtC) were constructed and analyzed. In our

previous study we have shortly discussed the preliminary results on the importance of the

N-terminal part of CrtC for the catalytic activity. The activity of the truncated versions was

fully retained, thus indicating that this part of the enzyme is not essential for activity [7].

Furthermore, the observed cleavage of TrCrtC also supports this hypothesis. Despite the

still unknown reason for this occurrence, we were able to identify the cleavage site between

S57 and S58 by using MS analysis [7]. In order to exclude any protease background activity

from the expression host E. coli, the S58 position was modified by substitution with valine

(same size, different chemical features) and glutamine (different size, same chemical

features). All mutants (Table 3.2, 3.3) were successfully cloned and expressed in E. coli

BL21 (Figure 3.7A). However, clear difference in expression levels was observed.

Therefore, all mutants were purified from the membrane fraction in order to ensure that

CrtC was present in the cells. As can be seen in Figure 3.7B, all mutants could be purified

and showed a band at 38- or 44 kDa, which was absent in the control sample (pET15b).

The introduction of mutations and modification of the protein lengths clearly has an effect


74

on the expression. While the removal of the N-terminus resulted in an increased expression

level, all point mutations negatively influenced the expression of the protein. In the case of

the wild type TrCrtC, the purification usually has to be performed as soon as it is expressed,

preferably before the cleavage of the N-terminal part (including the His-Tag). This was not

the case here and therefore only a very weak protein band is detected after the purification.

It should be noted here that the truncated version of TrCrtC shows a larger molecular weight

(Figure 3.7, lane 9) compared to the ‘cleaved’ versions (Figure 3.7, lanes 10-15), which is

due to the chosen primer position and the attached His-Tag.

Figure 3.7 SDS-PAGE (10%) analysis of expression (A) and IMAC purification from membrane (B) of

CrtC’s from R. gelatinosus (RgCrtC) (lane 1-7) and T. roseopersicina (TrCrtC) (lane 8-15) wildtype and

mutants. M, Precision plus protein standard. C, pET15b control. (A) First lane of each sample shows cells

before induction with 0.1 mM IPTG and the second lane shows cells after overnight expression at 25°C. 1,

RgCrtC wildtype; 2, RgCrtC truncated; 3, RgCrtC H239A; 4, RgCrtC W241A; 5, RgCrtC H264A; 6, RgCrtC

Y266A; 7, RgCrtC D268A; 8, TrCrtC wildtype; 9, TrCrtC truncated; 10, TrCrtC S58V; 11, TrCrtC S58Q;

12, TrCrtC H237A; 13, TrCrtC W239A; 14, TrCrtC H262A; 15, TrCrtC D266A.

With regard to the N-terminal point mutations, it seems that the cleavage rate increased in

the order of wildtype < S58V < S58Q. This conclusion is based on the fact that in the

M 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 C M

M 8 9 10 11 12 13 14 15 M

50

37

37

50

M 1 2 3 4 5 6 7 C M

(A)

(B)


75

wildtype sample in Figure 3.7A mainly the 44 kDa band is visible (lane 8). In the S58V

mutant about the same amount of both, the 38- as well as the 44 kDa bands could be

detected (lane 10), while in the S58Q mutant mainly the 38kDa band could be identified

(lane 11). Nevertheless, additional experiments are needed to get more insight into this

phenomenon. For example, by expressing larger amounts of the corresponding proteins,

and by purifying them, one could follow the change of the protein size in time. With the

wildtype TrCrtC, we have already shown that even though the purified enzyme consisted

of both sizes proteins, after few days of storage, the 44 kDa could not be detected anymore

(data not shown).

Next to the analysis of the expression levels, the activities of all constructed mutants were

measured with lycopene as substrate (Figure 3.8).

Figure 3.8 Enzymatic activity of wildtype (wt) and mutant CrtC from R. gelatinosus (A) and T.

roseopersicina (B). Extracts from E.coli cells expressing the respective enzymes were assayed with 20 μM

lycopene in 50 mM Na2HPO4 sodium phosphate (pH 8.0) at 28ºC overnight. Trunc, variants with missing N-

terminal residues 1-45 (RgCrtC) and 1-57 (TrCrtC).

As the expression levels were very low for some of the mutants and the activity of CrtC in

general is very low, crude extracts were used for the activity assay. Consequently, the

results cannot be quantitatively compared. However, in combination with the expression

wt trunc H239A W241A H264A Y266A D268A0

1

2

3

4

wt trunc S58V S58Q H237A W239A H262A D266A0

1

2

3

4

Enz

yme

acti

vity

[nm

ol m

g-1]

(A)

(B)


76

levels as shown in Figure 3.7A, indicative conclusions can be drawn. As stated in our

previous study the N-terminus is not important for catalytic activity [7]. This again was

proven here, as the truncated Rg- as well as TrCrtC were active. Furthermore, the

introduction of N-terminal point mutations did not affect the activity of TrCrtC, although

it did affect the cleavage rate of the N-terminal part. This might be explained by the fact

that the substitution of an amino acid by a smaller or chemically different amino acid could

result in conformational changes which promote or prevent the processing activity, either

by host proteases or through self-cleavage.

On the other hand, it appears that four key residues were identified, which have a potentially

important role in the hydration mechanism. By replacing each of the amino acids H239,

W241, Y266 and D268 individually by an alanine in RgCrtC the activity is completely

destroyed. The same mutations of the corresponding amino acids in TrCrtC, i.e. H237,

W239 and D266, also resulted in CrtC inactivation. Unfortunately, the mutagenesis of

Y264 in TrCrtC was not successful, and therefore, could not be included in this study.

However, based on all the results, one could expect that the mutation of Y264 in TrCrtC

would lead to inactivation, as has been seen for RgCrtC. On the other hand, the less

conserved H264 in RgCrtC and the corresponding histidine residue in TrCrtC (H262), seem

not to have any functional role. The mutants fully retained activity, and even showed

slightly increased activity when the expression levels were considered. For instance, the

truncated TrCrtC and H262A mutant showed almost the same level of expression (Figure

3.7A, lanes 9 and 14) but the activity of H262A mutant was ~ 1.3-fold higher (Figure 3.8B).

The same was observed for RgCrtC, where the expression of the wildtype is much more

than that of the mutant H264A, but both showed approximately the same activity.

All created mutants were also tested for the activity towards geranylgeraniol. In Figure 3.9,

representative HPLC results are depicted. We could confirm activity for mutants that were

still catalytically active for lycopene. The formation of the product was followed for two

days and an increase was observed, which indicates the activity of CrtC (Table 3.4). In

addition, when twice the amount of CrtC was used, the relative amount of the product also

increased 2-fold (data not shown). As described in chapter 2, it was not possible to isolate

the obtained product in amounts which are necessary for further analysis. However, from

the HPLC results we can conclude that the formed product is more hydrophilic than the

substrate itself, as it eluted earlier from the reversed phase C-18 column. If our assumption,

that CrtC recognizes only a specific part, i.e. -end group, in the carotenoid substrate


77

molecule is correct, then one can imagine that CrtC would be able to hydrate the terminal

double bond of geranylgeraniol.

Figure 3.9 HPLC separation of diterpene alcohols formed in vitro by E.coli extract expressing the RgCrtC

(solid line) compared with the blank reaction (dotted line) and the pET15b-plasmid (dashed line). Crude

extracts were assayed with 20 μM geranylgeraniol in 50 mM Na2HPO4 sodium phosphate (pH 8.0) at 30°C

and 800 rpm. Peak 1, reaction product; Peak 2, geranylgeraniol.

In order to investigate how the newly identified key residues could be involved in the

catalytic hydration reaction, the modeled structure of CrtC (Figure 3.4) was re-analyzed.

We have assumed earlier that regions IV and V might contain potentially important

residues, and indeed, it appears that our assumption is correct with regard to region IV. The

identified key residues H239, W241, Y266 and D268 in RgCrtC and the corresponding

residues in TrCrtC are all in close distance to each other (Figure 3.5). These four residues,

which are conserved throughout the CrtC family, are also found in the active site of

squalene-hopene cyclase (SHC) [14]. SHC catalyzes the cyclization reaction of squalene

to hopene as a major product (Figure 3.10). Hopanol is also formed to a minor extent. The

proposed mechanism for cyclases is proton-triggerd polycyclization, whereby the

intermediate carbocation is stabilized by aromatic amino acids.

0 1 2 3 4 5 6 7 8 9 10

100000

200000

300000

400000

500000

600000

Inte

nsity

[m

V]

Time [min]

blank pET15b RgCrtC

1

2


78

Table 3.4 Relative amount of the reaction product obtained from the substrate geranylgeraniol. Reactions

were performed with Escherichia coli crude extracts expressing R. gelatinosus and T. roseopersicina CrtC

wildtype and mutants, respectively. The areas under the substrate (1) and the product (2) peaks (Figure 3.9)

were set as 100% and were used as the measure of the relative activity. N.d. not detected.

Next to the stabilization role of the aromatic amino acids, they also create hydrophobic

environment in order to prevent quenching of the cation by water. The cyclization cascade

is terminated by a well-positioned enzymatic base. The formation of the side alcohol

product suggests significant water accessibility at the termination region of the active site.

Sample Reaction product (%)

after 1 day after 2 days

RgCrtC wt 0.37 1.00

RgCrtC trunc 0.47 0.82

RgCrtC H239A <0.05 <0.05

RgCrtC W241A <0.05 <0.05

RgCrtC H264A 0.08 0.11

RgCrtC Y266A <0.05 <0.05

RgCrtC D268A <0.05 <0.05

TrCrtC wt 0.06 0.09

TrCrtC trunc 0.14 0.44

TrCrtC S58V 0.14 0.29

TrCrtC S58Q 0.15 0.21

TrCrtC H237A <0.05 <0.05

TrCrtC W239A <0.05 <0.05

TrCrtC H262A 0.16 0.25

TrCrtC D266A <0.05 <0.05


79

Figure 3.10 Enzyme catalyzed cyclisation of squalene to hopene and hopanol.

The acidic residue aspartate (D376), which is located in the center of the active site in SHC,

is the likely general acid responsible for protonating the C3 atom of the squalene substrate

[14]. The acidity of D376 is enhanced by a connection to the side chain of Y495 through a

water molecule. Because of the similarity of the initial protonation reactions of squalene

and lycopene, we assume that the residues involved in catalysis will be alike in SHC and

CrtC. This hypothesis is in agreement with our results obtained by mutagenesis study as

well as the structure-based analysis. Therefore, we propose the following mechanism for

CrtC. D268 is the catalytic acid that initiates the hydration of lycopene (Figure 3.11).

Figure 3.11 Proposed mechanism for the initial protonation during lycopene hydration.


80

Upon diffusion of lycopene into the active site, it is required that the C2 atom of the

substrate is positioned near the proton of D268 that putatively will be added to the substrate.

In order to enhance the acidity of the catalytic D268 for olefin protonation, the amino acid

is directly bonded to H239 and to Y266 through an ordered water molecule, similar to what

has been proposed for SHC [14]. Mutation of one of these three amino acids leads to

inactivation of enzymatic activity, thereby supporting our hypothesis that they are all

directly involved in the hydration reaction of lycopene. In contrast to SHC, where

premature quenching of the cationic intermediate by water or nucleophiles is prevented by

well positioned aromatic amino acids, a water molecule is added to lycopene to yield the

desired hydroxylated lycopene derivative. This suggests that the active site of CrtC has

more water molecules present, so that the interaction between the substrate and solvent

water molecules is more significant. The aromatic amino acid Trp266 may be involved in

the stabilization of the intermediate carbocation.

Interestingly, according to the model, the residues H264 (Rg) and H262 (Tr), respectively,

which seem not to have any functional property, are located away (region V) from the

potential active site residues. This observation further supports our hypothesis that the

region IV is the active site of CrtC. On the other hand, the three conserved hydrophobic

residues proline, phenylalanine and tyrosine in region VI might play the role of attracting

the hydrophobic substrate and placing it in the right position. The mainly hydrophobic

amino acids in region II, which are located close to potential active side residues in region

IV, might play a role in stabilization of the substrate during catalytic activity.

3.4 Conclusion

The main purpose of this study was to investigate whether it is possible to get more

understanding of the hydration mechanism of carotenoid 1,2-hydratases. The used

approach was modeling of the 3D structure with the closest homologous protein with

known 3D structure, and subsequently the generation of point mutants of potentially

important amino acid residues. Overall results indicate that the 3D structure consist of a

beta-barrel, which closes with itself to form a central cavity. The substrate binding site,

which consists mainly of hydrophobic amino acid residues, is located at the top of the

cavity, while at the bottom inside the cavity potentially catalytic residues H, D, Y and W

3.5 Acknowledgements

81

are located. The absence of activity upon individual substitution of these residues by an

alanine supports their roles in the initial protonation. Although, the model with only 17%

sequence identity to the template is not very reliable, it fits to the data obtained on the

activities of the mutants, reassuring that the model is probably correct.

From our 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 this study shed for the first time light on structure-

activity relationships and opens the field for the engineering of carotenoid 1,2-hydratase to

generate industrially relevant mutants.

3.5 Acknowledgements

We thank Prof. Dr. Jaap Jongejan for advice on the potentially important residues in

carotenoid 1,2-hydratases and Jan van Leeuwen for making the CrtC model.


82

3.6 References

[1] G. A. Armstrong and J. E. Hearst, "Carotenoids .2. Genetics and molecular biology of carotenoid

pigment biosynthesis," Faseb Journal, vol. 10, pp. 228-237, Feb 1996.

[2] S. L. Jensen, et al., "Biosynthesis of carotenoids in purple bacteria : A re-evaluation based on

considerations of chemical structure," Nature, vol. 192, pp. 1168-1172, 1961.

[3] C. I. Cazzonelli, "Carotenoids in nature: insights from plants and beyond," Functional Plant Biology,

vol. 38, pp. 833-847, 2011.

[4] G. A. Armstrong, "Genetics of eubacterial carotenoid biosynthesis: A colorful tale," Annual Review

of Microbiology, vol. 51, pp. 629-659, 1997.







vol. 91, pp. 1029-1036, 2011.

[8] R. Kourist, et al., "Enzymatic synthesis of optically active tertiary alcohols: Expanding the

biocatalysis toolbox," Chembiochem, vol. 9, pp. 491-498, Mar 3 2008.



[10] C. M. Evans and A. J. Kirby, "A model for olefin hydration: Intramolecular nucleophilic addition of

phenolate oxygen to the unactivated double bond," Journal of the Chemical Society, Perkin

Transactions 2, pp. 1259-1267, 1984.

[11] H. J. Chiu, et al., "Structure of the first representative of Pfam family PF09410 (DUF2006) reveals

a structural signature of the calycin superfamily that suggests a role in lipid metabolism," Acta

Crystallographica Section F: Structural Biology and Crystallization Communications, vol. 66, pp.

1153-1159, 2010.

[12] N. J. Patel, et al., "Use of deuterium labelling from deuterium oxide to demonstrate carotenoid

transformations in photosynthetic bacteria," BBA - General Subjects, vol. 760, pp. 92-96, 1983.

[13] A. A. Yeliseev and S. Kaplan, "Anaerobic carotenoid biosynthesis in Rhodobacter sphaeroides

2.4.1: H2O is a source of oxygen for the 1-methoxy group of spheroidene but not for the 2-oxo group

of spheroidenone," Febs Letters, vol. 403, pp. 10-14, 1997.

[14] K. U. Wendt, et al., "Enzyme Mechanisms for Polycyclic Triterpene Formation," Angewandte

Chemie International Edition, vol. 39, pp. 2812-2833, 2000.

[15] S. C. Hammer, et al., "Squalene hopene cyclases: highly promiscuous and evolvable catalysts for

stereoselective CC and CX bond formation," Current Opinion in Chemical Biology, vol. 17, pp. 293-

300, 2013.

3.6 References

83

[16] S. F. Altschul, et al., "Basic Local Alignment Search Tool," Journal of Molecular Biology, vol. 215,

pp. 403-410, Oct 5 1990.

[17] A. Marchler-Bauer, et al., "CDD: A Conserved Domain Database for the functional annotation of

proteins," Nucleic Acids Research, vol. 39, pp. D225-D229, 2011.

[18] R. D. Finn, et al., "The Pfam protein families database," Nucleic Acids Research, vol. 38, pp. D211-

D222, January 1, 2010 2010.

[19] A. Dereeper, et al., "Phylogeny.fr: robust phylogenetic analysis for the non-specialist," Nucleic

Acids Research, vol. 36, pp. W465-469, 2008.

[20] A. Dereeper, et al., "BLAST-EXPLORER helps you building datasets for phylogenetic analysis,"

BMC Evolutionary Biology, vol. 10, 2010.

[21] A. A. Porollo, et al., "POLYVIEW: A flexible visualization tool for structural and functional

annotations of proteins," Bioinformatics, vol. 20, pp. 2460-2462, 2004.

[22] K. Arnold, et al., "The SWISS-MODEL workspace: A web-based environment for protein structure

homology modelling," Bioinformatics, vol. 22, pp. 195-201, 2006.

[23] L. A. Waidner and D. L. Kirchman, "Aerobic anoxygenic photosynthesis genes and operons in

uncultured bacteria in the Delaware River," Environmental Microbiology, vol. 7, pp. 1896-1908,

Dec 2005.



[25] M. Kammann, et al., "Rapid insertional mutagenesis of DNA by polymerase chain reaction (PCR),"

Nucleic Acids Research, vol. 17, p. 5404, 1989.

[26] G. Sarkar and S. S. Sommer, "The 'megaprimer' method of site-directed mutagenesis,"

BioTechniques, vol. 8, pp. 404-407, 1990.

[27] O. Landt, et al., "A general method for rapid site-directed mutagenesis using the polymerase chain

reaction," Gene, vol. 96, pp. 125-128, 1990.

[28] N. Igarashi, et al., "Horizontal transfer of the photosynthesis gene cluster and operon rearrangement

in purple bacteria," Journal of Molecular Evolution, vol. 52, pp. 333-341, Apr 2001.

[29] M. J. Zvelebil, et al., "Prediction of protein secondary structure and active sites using the alignment

of homologous sequences," Journal of Molecular Biology, vol. 195, pp. 957-961, 1987.

[30] D. R. Flower, "The lipocalin protein family: Structure and function," Biochemical Journal, vol. 318,

pp. 1-14, 1996.

[31] J. M. LaLonde, et al., "The up-and-down beta-barrel proteins," The FASEB Journal, vol. 8, pp. 1240-

7, December 1, 1994 1994.

[32] G. J. Bartlett, et al., "Analysis of catalytic residues in enzyme active sites," Journal of Molecular

Biology, vol. 324, pp. 105-121, 2002.

Chapter 4

4 Oleate hydratase as model enzyme to design and evaluate high-throughput screening assay for alcohol detection

Aida Hiseni, Rosario Medici, Isabel W.C.E. Arends, Linda G. Otten

Biotechnol. J. Feb 2014; DOI: 10.1002/biot.201300412

Oleate hydratase as model enzyme to design and evaluate high-throughput screening assay for alcohol detection

86

Abstract

A novel high-throughput screening assay for the detection of alcohols is developed by using

oleate hydratase (OHase) from Elizabethkingia meningoseptica as the model enzyme. It

allows for screening of mutant libraries generated by directed evolution techniques or other

mutagenesis methods. The assay is based on the reaction between the alcohol and nitrous

acid to form the corresponding alkyl nitrite and is valid for a broad range of alcohols,

differing in size and solubility. Cyclic as well as acyclic unsaturated alkenes (substrates)

and the corresponding alcohols (products) were tested and they showed sufficient

discrimination for analysis. Lower detection limits were 1.5 to 3 mM with excellent Z-

factors ranging from 0.70 – 0.91. Precision, linearity and plate uniformity were estimated

with pure substrates, mixtures and enzymatic reactions.

4.1 Introduction

87

4.1 Introduction

Oleate hydratase (OHase) (EC 4.2.1.53) belongs to the group of hydro-lyase enzymes (EC

4.2.1), which cofactor independently catalyze the reversible addition of a water molecule

to a substrate possessing a carbon-carbon double bond [1]. The 73 kDa OHase from

Elizabethkingia meningoseptica has been characterized with respect to its biochemical

properties [2] and has recently been immobilized as CLEA (cross-linked enzyme

aggregate) [3]. The enzyme shows high amino acid sequence similarities with members of

the Streptococcal 67 kDa myosin-cross-reactive antigen like family [4, 5]. A few group

members of this family were shown to also exhibit fatty acid hydratase activity [5-7].

Knowledge of the hydratase could be of great importance for the industry, as the reaction

product 10-hydroxystearic acid (10-HSA) is a high-added-value compound and can be used

for the production of a large number of industrial products including resins, waxes, nylons,

plastics, cosmetics and coatings [3]. Compared to the traditional acid-catalyzed water

addition, the enzymatic reaction proceeds under very mild conditions and is stereo- and

regioselective. Recently, attempts have been made to increase microbial 10-HSA

production by using recombinantly expressed oleate hydratase from Stenotrophomonas

maltophilia [6, 8].

In industrial biocatalytic processes, hydro-lyases are underrepresented and only a few

group members are amenable to be used for industrial scale reactions, including nitrile

hydratase for the production of acrylamide [9] and fumarase to produce malate [10] . This

is mainly due to the low stability and/or catalytic activity of these enzymes. However,

utilization of this class of enzymes as biocatalysts needs intensive study and optimization

of enzyme properties, such as stability, specific activity and selectivity, beforehand.

A powerful tool to address these issues is directed evolution, which has been used in the

past decade to improve biocatalysts [11-14]. An advantage of this technique is that no need

for knowledge about the structure-function relationship is required. Moreover, a large

number of mutants with potentially improved and/or novel properties can be produced in a

short time. However, a crucial step in any directed evolution experiment is the development

of a high-throughput screening (HTS) assay, which allows rapid screening of a large

number of variants within a reasonable timeframe [15]. In general, the assay has to be

sensitive, easy to perform, robust and has to have high throughput.


88

So far, the quantification of enzymatic activity of OHase has usually been determined by

either GC [2, 5-8] or HPLC [3, 16]. The methods are based on derivatization of fatty acids

and are straightforward and accurate. Nevertheless, they are not suitable for high-

throughput screening.

Therefore, we have developed a spectrophotometric high-throughput screening method for

the detection of alcohols. The assay is based on the simple reaction between the alcohol

that is produced by the enzyme and nitrous acid to form the corresponding alkyl nitrite

(Figure 4.1) [17].

Figure 4.1 Simplified scheme for the conversion of an alcohol to the corresponding ester of nitrous acid (box).

Figure 4.2 Absorption spectra of 2-methylbutyl nitrite (primary), 3-methylbutan-2-yl nitrite (secondary) and

tert-pentyl nitrite (tertiary) obtained from 2-methyl-1-butanol, 3-methyl-2-butanol and 2-methyl-2-butanol,

respectively. 10 μl of the 150 mM stock in acetone was added to 90 μl of 20 mM Tris-HCl (pH 8.0) and the

nitrosation reaction performed under standard conditions (as described in Materials and Methods, section

“Assay conditions”).

330 340 350 360 370 380 390 400 410 420 430 440

0,00

0,02

0,04

0,06

0,08

0,10

Abs

orba

nce

Wavelength [nm]

primary secondary tertiary


89

Next to enzyme activity detection, this assay provides also information about the position

of the alcohol group in the molecule. For example, alkyl nitrites obtained from tertiary

alcohols have a maximum at 400 nm, which is absent in those obtained from primary and

secondary alcohols (Figure 4.2) [18].

With this valuable information the regioselectivity of an enzyme can be easily determined,

especially of importance if the substrate contains several double bonds that could be

converted by the same hydratase.

The capability of the developed method for enabling an automated set up has been

examined and characterized with OHase as the model enzyme. Next to precision of the

system, the linearity and quality have also been addressed by using simulated and

enzymatic reaction systems.


4.2.1 Standard curves and Z-factor determination In order to validate the assay for linearity, standard curves and reaction simulation curves

(substrate-product mixtures) were prepared using different alkenes and alcohols (Sigma-

Aldrich) in a concentration range of 1 – 15 mM in triplicate. A desired amount of the 150

mM stock in acetone was manually added to the plates pre-filled with 20 mM Tris-HCl (pH

8.0). Subsequent steps were performed using the automated liquid handling system as

described in “Assay conditions” section. In the case of fatty acids 12-hydroxystearic acid

(12-HAS) and oleic acid (OA) (Sigma-Aldrich) a stock in DMSO was used and dispensed

by Hamilton syringe for better accuracy.

For the determination of the Z′-factor, a parameter that indicates the suitability of the assay

to be used in high-throughput format, 15 mM alkene, alcohol and fatty acids, respectively,

were used. Therefore, 10 μl of the 150 mM stock was added to plates pre-filled with 20

mM Tris-HCl (pH 8.0) (n = 32), and the nitrosation assay was performed using the standard

conditions. The calculation of the Z′-factor was done using following equation:

(3 3 )1 alcohol alkene

alcohol alkene

SD SDZ

mean mean

(Eq. 4.1)

To test the real case situation, E. coli TOP10 cells with pBAD-HISA-OH (pBAD/HisA

vector containing ohyA gene) were used instead of the standards, whereby pBAD-HISA


90

(pBAD/HisA empty vector) served as the negative control (n = 48). Following equation

was used for the Z-factor calculation:

(3 3 )Z 1 pBAD HISA OH pBAD HISA

pBAD HISA OH pBAD HISA

SD SD

mean mean

(Eq. 4.2)

4.2.2 Large scale production of 10-HSA As the OHase reaction product, 10-hydroxystearic acid (10-HSA), is commercially not

available, we used 12-HSA instead, with the assumption that both have similar properties.

In order to test this hypothesis, large scale production of 10-HSA was performed using E.

coli TOP10 cells overexpressing OHase. The reaction mixture contained 1 ml of the cell-

free extract (24 mg ml-1), 0.6 % (v/v) oleic acid in a total volume of 25 ml in 20 mM Tris-

HCl (pH 8.0). After overnight incubation at 30°C and 200 rpm, a 500 μl sample was used

to confirm the product by HPLC. First, the pH of the sample was adjusted with 50 μl of 3N

HCl. Subsequently, 50 μl of saturated NaCl solution was added and fatty acids extracted

with one volume of dichloromethane. The mixture was shaken for 5 min at 1400 rpm,

centrifuged for 1 min at 13.200 rpm and 1 μl of the dichloromethane phase was transferred

into a new tube. After drying with a SpeedVac Concentrator (Thermo Scientific), fatty acids

were derivatized and analyzed as described earlier [3]. Once the reaction product was

confirmed as 10-HSA, it was isolated as described previously [16] with small modification.

Briefly, the rest of the mixture (24.5 ml) was filtered through Whatman filter-paper (Grade

1) and the filter with the residues dried overnight at 39°C. The dry solid was scraped off

and dissolved in 15 ml of EtOH. Unsoluble material was filtered off and the filtrate dried

using a SpeedVac Concentrator (Thermo). The isolated product was stored at 4°C until

further use.

4.2.3 Growth conditions in 96-well deep well plates Well separated E. coli TOP10 colonies containing the plasmid pBAD-HISA-OH [3] or

empty pBAD-HISA (control) were picked and transferred to individual wells in 96-well

microtiter plates containing 150 μl LB medium with 100 μg ml-1 ampicillin and 0.2 % L-

arabinose (w/v) followed by overnight incubation at 37°C and 150 rpm. Using a 96-pin

colony replicator, the cells were transferred into 96-well deep well plates (2 ml, V-bottom,

Greiner Bio-One), which were pre-filled with 1 ml of TB-medium and 100 μg ml-1

ampicillin. Prior to incubation for 24 h at 37°C and 150 rpm, the plates were covered with

gas-permeable seals (BreathSeal, Greiner Bio-One) and lids (Lid with condensation ring,


91

Greiner Bio-One). After harvesting by centrifugation for 35 min at 4000 rpm and 4°C

(5810R, Eppendorf), the resulting cells were washed ones with 20 mM Tris-HCl (pH 8.0)

and stored at -20°C overnight.

4.2.4 Liquid handling All liquid handling steps were performed using the JANUS® automated workstation

(Perkin Elmer). The system is equipped with an 8-tip pipetting arm and an arm with a 96-

channel dispense head.

4.2.5 Assay conditions Cell lysates were prepared by resuspending the cell pellets in 130 μl of 20 mM Tris-HCl

(pH 8.0) containing 1 mg ml-1 lysozyme and 0.1 mg ml-1 DNAse and subsequent incubation

at 37°C for 1 h. Cell debris and unlysed cells were removed by centrifugation for 35 min at

4000 rpm and 4°C. For enzymatic reactions, 95 μl of the cell-free extract was transferred

into a 96-well deep well assay plate (1 ml, U-bottom, Greiner Bio-One) pre-filled with 5 μl

of the substrate oleic acid (1 M stock in DMSO). The plates were covered and incubated

overnight at 30°C and 300 rpm.

For whole cell reactions, cell pellets were resuspended in 130 μl of 20 mM Tris-HCl (pH

8.0) and 95 μl of the cell suspension was directly transferred into the assay plate containing

the substrate. The reaction mixtures were incubated overnight at 30°C and 300 rpm.

Following steps were performed with the automated workstation (Figure 4.3).

Figure 4.3 Conceptualized 96-well high-throughput screening assay procedure using JANUS® automated

workstation. Steps after the performance of the enzymatic reaction are shown. Refer to text for a stepwise

description of the operation.


92

First, 150 μl of octane was added to the assay plate (Figure 4.3, a), followed by 50 μl of

10% HCl (aq., v/v) (Figure 4.3, b) and 80 μl 10% NaNO2 (aq., w/v) (Figure 4.3, c) with the

96-channel dispense head. The plate was manually capped (CapMat, Greiner Bio-One),

shaken vigorously for 3 min at 1500 rpm using a small shaker with microtiter attachment

(IKA) and centrifuged for 3 min at 4000 rpm and RT. The plate was returned to the

workstation and by using the 8-tip pipetting arm, 50 μl of the organic layer was transferred

to a clean 96-well microtiter plate containing 50 μl of octane (Figure 4.3, d). The pipetting

height was carefully chosen so that only organic layer was transferred. After a short

agitation, absorption spectrum was taken between 330 nm and 440 nm using a

spectrophotometer (Synergy 2, Bio Tek Instruments, Inc.). To calculate activity the

absorbance of the highest peak was taken; this is approximately 356 nm, 372 nm and 382

nm for 1°, 2° and 3° alcohols respectively.

4.2.6 Preparation of ohyA mutant libraries For the generation of randomly mutated ohyA variants, epPCR (error-prone PCR) was

performed across the entire coding sequence (1941 bp) using the Genemorph II random

mutagenesis kit (Stratagene). Two libraries were constructed with low (0 – 4.5

mutations/kb) and high (9 – 16 mutations/kb) mutation frequencies. This could be achieved

by adjusting the template DNA concentration. Amplification reactions were done under

standard PCR conditions using plasmid pBAD-HISA-OH [3] as template and flanking

primers pBAD_For (CTCTTCTCGCTAACCAAACC) and pBAD_Rev

(GGCGTTTCACTTCGGCATGG). Prior to the cloning of the variant genes into an

appropriate vector, the first PCR products were subjected to a second PCR reaction, where

restriction sites for XhoI (forward) and HindIII (reverse) were introduced through a second

set of primers (OH_F AATCTCGAGATGAACCCAATAACTTC; OH_R

ATTAAGCTTTTATCCTCTTATTCCTTTTAC) (restriction sites are underlined) and the

amount of the DNA was increased. Taq PCR master kit (Qiagen) was used following the

manufacturer’s instruction. Using XhoI / HindIII restriction sites, the digested and purified

fragments were ligated into the same sites of the pBAD-HISA vector and transformed into

electrocompetent E. coli TOP10 cells. The insertion of the genes was verified by restriction

analysis with XhoI / HindIII enzymes. From each library, a representative number of

randomly selected clones was analyzed by sequencing (BaseClear).


93

4.2.7 Expression of ohyA variants After the transformation of the recombinant plasmids into E. coli TOP10, the cells were

spread on a 200 ml LB agar plate containing 100 μg ml-1 ampicillin and the plates were

incubated at 37°C overnight. The volume of the cells used was adjusted so that between

1000 and 2000 clones would be present per plate. Individual colonies from mutant libraries

were inoculated into individual wells of 96-well microtiter plate containing 150 μl LB

medium supplemented with 100 μg ml-1 ampicillin with a VersArray Colony Picker

(Biorad). By removing six of the colony picking needles, the empty wells could be

manually inoculated with six wild type ohyA clones, which served as positive controls in

each plate. The plates were covered with gaspermeable seals (BreathSeal, Greiner Bio-

One) and incubated at 37°C overnight with shaking (150 rpm). Following this, the cells

were transferred into 96-well deep well plates (2 ml, V-bottom, Greiner Bio-One) pre-filled

with 1 ml of TB-medium and 100 μg ml-1 ampicillin using a colony copier. The rest of the

cells was mixed with 40 μl of 60% glycerol (reference plate), sealed with seals (SilverSeal,

Greiner Bio-One) and stored at -80°C. Prior to incubation for 24 h at 37°C and 150 rpm,

the plates were covered with gaspermeable seals and lids (Lid with condensation ring,

Greiner Bio-One). Next, protein expression was induced by adding 10 μl of a 20% L-

arabinose stock using the JANUS® automated workstation (Perkin Elmer) and the plates

incubated for another 24 h under the same conditions. After harvesting by centrifugation

for 35 min at 4000 rpm and 4°C, the resulting cells were washed ones with 20 mM Tris-

HCl (pH 8.0) and stored at -20°C overnight.

4.2.8 Library screening Cell pellets were resuspended in 130 μl of 20 mM Tris-HCl (pH 8.0). For enzymatic

reactions, 95 μl of the cell suspension was transferred into a 96-well deep well assay plate

(1 ml, U-bottom, Greiner Bio-One) pre-filled with 5 μl of the substrate oleic acid (1 M

stock in DMSO). The plates were covered and the reaction mixtures incubated overnight at

30°C and 300 rpm. Following steps were performed with the automated workstation as

described in the “Assay conditions” section


94


A high-throughput screening assay for hydro-lyases was developed. At first, a product with

the alcohol group is obtained by the enzymatic reaction. Subsequently, the nitrosation

reaction is performed and the resulting alkyl nitrite extracted using an immiscible organic

solvent. Direct measurements of the UV spectra were used for product detection. After

validation of the assay it was assessed using oleate hydratase from E. meningoseptica as

the model enzyme.

4.3.1 Method performance and linearity with small substrates In order to demonstrate that the assay is applicable to molecules with different sizes,

structures and substituents, and to test the precision and detection limits of the developed

method, different concentrations of unsaturated cyclic and acyclic substrates and the

corresponding alcohols were assessed under high-throughput assay conditions (100 μl).

Nitrosation of alcohols is a standard procedure in laboratories for the synthesis of alkyl

nitrites and some nitrosation reactions have also been adapted to the industrial scale [19].

However, our objective is the miniaturization of this reaction in order to be applicable to

the 96-well plate format. To accommodate good mixing of the reaction mixture, 96-well

deep well plates (1 ml) were used instead of 96-well microtiter plates (~0.3 ml).

Standard curves ranging from 1 to 15 mM were constructed and the results (Figure 4.4)

demonstrate that the discrimination between the reaction with the alkene (substrate) or the

alcohol (product) is sufficient for analysis. However, a slight background reaction of the

substrate was observed for the cyclic compounds. Lower limits of detection for each

compound were 1.5 mM, except for compounds 2-methyl-1-butanol and 2-methyl-2-

butanol, where the response below 3 mM was not linear or reliable with relative standard

deviation (RSD) exceeding 20% (calculated based on three replicates). Thus, this assay is

not applicable to reactions where low or new enzyme activities are to be discovered. On

the other hand, the linearity of the standard curves from 3 to 15 mM indicates that the

method can also be used for quantification. This is a remarkable result, since the assay has

many steps and additional liquid-liquid extraction, which in general give high errors in 100

μl format. Moreover, these results demonstrate the high robustness of the assay despite the

complex set-up.


95

0 2 4 6 8 10 12 14 16

0,00

0,02

0,04

0,06

0,08

0,10

Abs

orba

nce

356

nm

Conc. [mM]0 2 4 6 8 10 12 14 16

0,00

0,02

0,04

0,06

0,08

0,10

Abs

orba

nce

382

nm

Conc. [mM]0 2 4 6 8 10 12 14 16

0,00

0,02

0,04

0,06

0,08

0,10

Abs

orba

nce

372

nm

Conc. [mM]

0 2 4 6 8 10 12 14 16

0,00

0,02

0,04

0,06

0,08

0,10

Abs

orba

nce

356

nm

Conc. [mM]0 2 4 6 8 10 12 14 16

0,00

0,02

0,04

0,06

0,08

0,10

Abs

orba

nce

382

nm

Conc. [mM]0 2 4 6 8 10 12 14 16

0,00

0,02

0,04

0,06

0,08

0,10

Abs

orba

nce

370

nm

Conc. [mM]

0 2 4 6 8 10 12 14 16

0,00

0,02

0,04

0,06

0,08

0,10

Abs

orba

nce

372

nm

Conc. [mM]0 2 4 6 8 10 12 14 16

0,00

0,02

0,04

0,06

0,08

0,10

Abs

orba

nce

360

nm

Conc. [mM]

Figure 4.4 Analysis of small alkene/alcohol pairs using the proposed high throughput screening method.

Nitrosation reactions were performed with different concentrations of the standard (stock in acetone) in 20

mM Tris-HCl (pH 8.0). Absorbance of the highest peak in the alcohol spectrum was plotted.

One important parameter in the described method is the choice of the organic solvent,

which has to be screened depending on the expected reaction product. Several organic

solvents were tested including octane, decane, dodecane, tetradecane and hexadecane.

Octane showed the best results for compounds used in this study (data not shown).

4.3.2 Method performance for larger substrates and reaction simulation

Besides small cyclic and acyclic unsaturated substrates, the larger substrate oleic acid was

tested, differing significantly in size and solubility. As 10-HSA is not commercially

available, 12-HSA was used in this study to identify and evaluate the OHase reaction

product, which has the alcohol group at a different position (position 12 vs. 10). To

determine if both products would react similarly in the proposed assay, a larger scale

production of 10-HSA was performed with OHase. The product 10-HSA was isolated and

confirmed with HPLC as > 85% pure. Nitrosation experiments were performed with 5 mM

HO

HO

OH

HOHO

OH

OH

HO


96

10- and 12-HSA and the results are depicted in Figure 4.5. They clearly show that 12-HSA

can be used as a standard for 10-HSA, since the spectra are identical. Lower purity of 10-

HSA explains the slightly lower values in the absorption spectra with the same

concentration.

Figure 4.5 Absorption spectra of 10-(nitrosooxy)octadecanoic acid obtained from 10-HSA, 12-

(nitrosooxy)octadecanoic acid obtained from 12-HSA and control reaction with OA. 50 mM stock in DMSO

was diluted 10 x with 20 mM Tris-HCl (pH 8.0) and the nitrosation reaction performed using standard

conditions.

Nitrosation of substrate oleic acid and the (artificial) product 12-HSA were performed. As

shown in Figure 4.6, the obtained standard curve illustrates that even with very unsoluble

substrates / products this HTS method can be used. No background reaction has been

observed from the substrate oleic acid. Thus, the assay can be applied for the enzymatic

reaction with OHase.

Next, enzymatic reaction progress was simulated by using different mixtures of the

substrate oleic acid and the product 12-HSA. The performed experiment gives an indication

of the approach applicability to real situation, where different ratios of substrate and

product are present in the mixture at the same time. From the results presented in Figure

4.7, a clear overlap of the simulated and the standard curve was observed, showing that the

assay is not negatively influenced by the presence of the substrate during the assay.

330 340 350 360 370 380 390 400 410 420 430 440

0,000

0,005

0,010

0,015

0,020

0,025

0,030

Abs

orba

nce

Wavelength [nm]

10HSA 12HSA OA


97

Figure 4.6 Analysis of 12-HSA/OA pair using the proposed high throughput screening method. Nitrosation

reactions were performed with different concentrations of the standard (stock in DMSO) in 20 mM Tris-HCl

(pH 8.0).

Figure 4.7 Simulated reaction progress and standard curve of 12-(nitrosooxy)octadecanoic acid obtained from

12-HSA. Different ratios of OA and 12-HSA were prepared (total concentration 16 mM) to simulate

enzymatic reaction. In comparison, the standard curve of nitrosation of 12-HSA only, was plotted (Figure

4.6).

0 2 4 6 8 10 12 140,00

0,02

0,04

0,06

0,08

Abs

orba

nce

372

nm

Concentration [mM]

0 2 4 6 8 10 12 140,00

0,02

0,04

0,06

0,08

Reaction simulation 12-HSA

Abs

orpt

ion

372

nm

Concentration [mM]


98

4.3.3 Precision and accuracy (Z-factor) To develop a HTS method that can be used for the identification of ‘hits’, robustness and

reproducibility are of a big importance. Therefore, a simple parameter (Z-factor) was

defined by Zhang et al. [20], which can be used to evaluate the quality of the HTS method

and recently has been applied for the detection of decarboxylase activities [21]. As this

factor is dimensionless, it can be used to compare different HTS methods. The factor takes

into account standard deviations and mean values of sample and control measurements

(Eq. 4.1). In this study, first the Z′-factor was determined for all tested compounds to

evaluate the quality of the assay itself and a summary of the results is shown in Table 4.1.

Table 4.1 Z′-factor values of various alkene / alcohol pairs, indicating the screening assay quality. The Z′-

factor was calculated using the formula (Eq. 4.1): Z′-factor = 1 – 3 × (SDalcohol + SDalkene) / |meanalcohol –

meanalkene|

Alkene / Alcohol Z′-factor

2-methyl-1-butene / 2-methyl-1-butanol 0.82




2-methyl-1-pentene / 2-methyl-1-pentanol 0.91




1-methylcyclohexene / 2-methylcyclohexanol 0.72

3-methylcyclohexene / 3-methylcyclohexanol 0.78

oleic acid / 12-hydroxystearic acid 0.75

The Z′-factors are all >0.5, while most of them are even >0.7. Z′-factor values ranging from

0.5 to 1 demonstrate a good quality of the assay and can be used for identification of ‘hits’

[20]. Thus, the obtained values indicate that the HTS assay is an ‘excellent assay’ for all

alkene/alcohol pairs tested, which makes this assay applicable to distinguish all kind of

hydroxylated compounds from their unsaturated counterparts in only one step.

Another important parameter when developing a 96-well format assay is the reproducibility

and spatial plate uniformity. The design of the plates and the process can introduce new

problems, for example, temperature gradients and aeration differences, especially when

plates are stacked during the incubation time. The results can reveal patterns of drifts or

edge effects, which have to be diminished by design optimization [22]. For this reason, a


99

scatter plot was prepared by plotting the response of the standards OA and 12-HSA against

well number (n = 32), either by column or by row. The results (Figure 4.8A, B) show that

no significant drifts were observed for the fatty acid standards, neither when plotting by

column nor when plotting by row. This indicates that the process design of the assay is

appropriate for high throughput screening. From these results we also can conclude that the

automated liquid handling workstation works with high accuracy.

Figure 4.8 Scatter plot to assess the spatial uniformity. The response of nitrosated 15 mM 12-HAS and 15

mM OA is plotted against well number, ordered by row (A) and by column (B). The nitrosation reaction was

performed under standard conditions.

0 12 24 36 48 60 72 84 96

0,00

0,02

0,04

0,06

0,08 12-HSA OA

Abs

orba

nce

372

nm

Well number, by row

0 8 16 24 32 40 48 56 64 72 80 88 96

0,00

0,02

0,04

0,06

0,08 12-HSA OA

Abs

orba

nce

372

nm

Well number, cy column

(A)

(B)


100

4.3.4 Optimization of protein expression conditions In order to test the applicability of the assay with real samples, E. coli TOP10 cell-extracts

and whole cells with and without overexpressed OHase were used. This is especially

important, as cell-extracts or whole cells naturally contain a lot of alcohols, which can serve

as substrate for the nitrosation reaction and may result in increased background signal. In

an effort to reduce the background signal, initial optimization of the expression was

performed by using TB-medium (Terrific Broth) instead of the usual LB-medium for the

enzyme expression. As expected, 2-fold higher protein concentrations were obtained with

TB-medium compared to the LB-medium (data not shown). Furthermore, reaction with

whole cells showed higher response to that obtained from cell-free extracts under the same

reaction conditions and was chosen for further experiments to simplify the HTS process.

10-HSA production was performed in 96-well plates using E. coli TOP10 cells carrying

either empty an expression vector (negative control) or a vector with ohyA gene (positive

control). Under the chosen assay conditions (50 mM oleic acid, 30°C, overnight incubation)

around 30% of the oleic acid was converted to 10-HSA (data not shown). Only partial

conversion of the substrate is required in order to be able to detect potentially improved

activity as long as the amount produced is above the detection limit. The nitrosation of the

reaction products was conducted using the automated workstation. In this experiment, not

only the accuracy of the workstation was tested, but also the spatial uniformity of the cell

growth, substrate addition and temperature influence. The obtained results (data not shown)

revealed substantial deviations in the response. In order to identify factors, which are

affecting these results, further experiments were performed. At first, we have looked into

the growth behavior of bacteria in 96-well deep well plates as it can differ from that in

standard laboratory containers like Erlenmeyer flasks. The cell densities showed significant

deviations during the first 10 hours of the growth (Figure 4.9). Usually, the protein

expression is induced in the exponential phase of the growth, when cell densities reach a

value of ~0.4 (OD600). However, with the miniaturization of cell cultures like in the high-

throughput screening set-up, this would decrease the efficiency because of the added step

in the procedure. Hence, the inducer agent is added immediately at the beginning of the cell

growth. Since plates are inoculated with a colony replicator, cell densities are relatively

different at that point. Consequently, this results in different enzyme quantities in each well.

In order to have approximately the same amount of the enzyme in each well, the protocol


101

was adapted and the protein expression was induced after 25 h growth, where the standard

deviation showed an acceptable value of 6.3% (Figure 4.9).

Figure 4.9 Growth curve of E. coli TOP10 in 96-well deep well plates. Mean values and standard deviations

of the cell densities from 96 E. coli cultures are shown, which were grown at 150 rpm and 37°C over a period

of 25 h.

The enzymatic reaction was repeated with cells grown under optimized conditions. Despite

inducing OHase production later, a high standard deviation was obtained in the response.

In an attempt to decrease the error in the OHase response, the protein induction time was

varied between 16 and 25 h. As shown in Table 4.2, the Z-factor increased significantly

from -46.50 to -0.77 by increasing the induction time.

Table 4.2 Z-factor values of E. coli cultures containing pBAD-HISA or pBAD-HISA-OH plasmid, indicating

the total screening assay quality. The Z-factor was calculated using the formula (Eq. 4.2): Z-factor = 1 – 3 ×

(SDpBAD-HISA-OH + SDpBAD-HISA) / |meanpBAD-HISA-OH – meanpBAD-HISA|.

Experiment Induction time (h) Z-factor

1 16 -46.50

2 21 -1.0

3 25 -0.77

0 5 10 15 20 25 300,0

0,1

0,2

0,3

0,4

0,5

0,6

0,7

0,8 Cell density E. coli TOP10

OD

600

Time [h]

Time (h) Mean SD SD (%)

4 0,01 0,003 29,5

8 0,2 0,026 13,4

10 0,42 0,062 14,6

25 0,67 0,042 6,3


102

In addition, responses obtained were assessed for patterns of drift or edge effects, by

plotting against well number and ordered either by row first, then by column

(Figure 4.10A), or by column first, then by row (Figure 4.10B).

Figure 4.10 Scatter plot to assess the spatial uniformity. The response of nitrosated E. coli pBAD-HISA-OH

and E. coli pBAD-HISA (n=48) reaction products is plotted against well number, ordered by row (A) and by

column (B). The enzymatic and nitrosation reactions were performed under standard conditions.

0 8 16 24 32 40 48 56 64 72 80 88 96-0,01

0,00

0,01

0,02

0,03

0,04

0,05

0,06

0,07

0,08

0,09

0,10 E. coli pBAD-HISA-OH E. coli pBAD-HISA

Abs

orba

nce

370

nm

Well number, by column

0 12 24 36 48 60 72 84 96-0,01

0,00

0,01

0,02

0,03

0,04

0,05

0,06

0,07

0,08

0,09

0,10 E. coli pBAD-HISA-OH E. coli pBAD-HISA

Abs

orba

nce

370

nm

Well number, by row

(B)

(A)


103

There are no drift patterns or edge affects visible in these figures, indicating that bacterial

growth is not affected by the place it is situated. It is, however, obvious from the plots that

the negative and positive points overlap, which will result in more false positives or missing

of hits, depending on the threshold set. Although the deviation of the negative controls is

larger than in the assay alone (32% vs 26%) it is still reasonable, considering the extra steps

in this process. The RSD of 26% for the positive controls indicates a substantial variability

in the response, since the 12-HSA variation itself was only 3.3%. This is probably due to

variability in cell growth and protein production, despite the improvements already

implemented. Another point to investigate is the octane extraction step. Compared to the

used oleic acid and 12-HSA standards, whole cells consists of a complex mixture of

different compounds, which may influence the extraction and introduce deviations to the

assay. In order to be applicable for the detection of potential ‘hits’ from mutant library, the

cell growth and enzyme production as well as the extraction step need further

investigations. Adding extra steps will take more time and effort, but hopefully result in a

good assay. Improvements should be at the beginning of the process, for instance,

inoculation from pre-grown plates by the pipetting robot instead of the colony replicator

tool, or afterwards by improving the extraction step. Also changing host or expression

vector could improve enzyme variability. As changing the induction time already improved

the Z-factor 60-fold, it should be possible to reduce the standard deviation of the OHase

signal in the plate by another factor of 10, leading to a Z-factor >0.5, which would be an

ideal assay.

From the results described so far, it is clear that further improvements are needed in order

to implement the new developed assay in the directed/random evolution studies. However,

for the time being we applied the new high-throughput screening assay to identify an OHase

variant with improved activity towards oleic acid by using a threshold value of 2 standard

deviations of the wildtype OHase activity in each plate. The generation of the mutants

included random mutagenesis of the ohyA, subsequent cloning in pBAD-HISA vector and

expression in E.coli TOP10 cells. Two separate mutant libraries were prepared with either

low- or high mutation frequency. The analysis of 55 randomly picked clones revealed an

average mutation frequency of 5 mutations/kb, which is close to the expected range of 0 –

4.5 mutations/kb (data not shown). The average mutation frequency on amino acid

sequence level was 3 mutations/gene.

Approximately 1500 transformants were obtained in each library. In first instance, the low

mutation frequency library was tested and the absorbances at 372 nm analyzed. An example


104

of the screening results is shown in Figure 4.11. The wild type OHase (Figure 4.11, black

bars) was included in each plate to account for the variation of the activities.

Figure 4.11 Evaluation scheme of a screening process. To account for the variation of the activities of the

individual variants (gray bars), a threshold (dashed line) was defined, above which a variant was declared as

positive. The threshold was calculated for each plate using the data of the control (wildtype) activities (black

bars). The absorbance spectra of a positive mutant and a wildtype are displayed.

13 mutants exhibiting higher activity than the wild type were identified by means of the

colorimetric high-throughput screening method (Table 4.3).

Due to the restricted time, further analysis of the obtained mutants could not be executed.

However, in order to confirm the screening result, the obtained clones need to be re-

cultivated and re-tested for activity. Once the results are re-produced and confirmed, more

detailed analysis of the mutations in these particular mutants would be of high interest, as

they could give an indication of directions for further improvement of the activity.

340 360 380 400 420 4400,00

0,02

0,04

0,06

0,08

0,10

A1

A2

A3

A4

A5

A6

A7

A8

A9

A10

A11

A12 B1

B2

B3

B4

B5

B6

B7

B8

B9

B10

B11

B12 C1

C2

C3

C4

C5

C6

C7

C8

C9

C10

C11

C12 D1

D2

D3

D4

D5

D6

D7

D8

D9

D10

D11

D12 E1

E2

E3

E4

E5

E6

E7

E8

E9

E10

E11

E12 F1 F2 F3 F4 F5 F6 F7 F8 F9 F10

F11

F12 G1

G2

G3

G4

G5

G6

G7

G8

G9

G10

G11

G12 H1

H2

H3

H4

H5

H7

H8

H9

H10

H11

0,00

0,02

0,04

0,06

0,08

0,10

Abs

orba

nce

372

nmA

bsor

banc

e

Wavelength

G12 wildtype average


105

Table 4.3 Screening analysis of 1500 epPCR OHase variants. The normalized values at λmax = 372nm of the

OHase variants were compared to the values obtained for the wildtype OHase in the same plate. All showed

mutants were declared as positive using the corresponding threshold values.

In conclusion, a robust, high quality high-throughput screening method has been developed

for the detection of alcohols. In general, this assay can be used with any hydro-lyase

member, whose product can undergo a reaction with a nitrosating agent to form alkyl

nitrites. The assay is applicable to a broad range of compounds varying in size and

solubility, with good to excellent Z′-factors. Future studies will be directed at optimization

of the assay procedure for improvement of the plate uniformity of the enzyme concentration

and of the octane extraction.

96-well plate

number

Threshold

(wt average + 2SD)

Absorbance 372 nm

Wildtype

(average) Mut1 Mut2 Mut3 Mut4

01 0.051 0.033 0.054 0.064 0.052

02 0.061 0.045 0.072 0.069 0.069 0.063

07 0.082 0.062 0.090

09 0.045 0.023 0.048 0.050

15 0.089 0.050 0.092 0.096 0.100


106

4.4 References





[3] A. Hiseni, et al., "Preparation and properties of immobilized oleate hydratase as a cross-linked

enzyme aggregate (CLEA)," 2012.

[4] E. Rosberg-Cody, et al., "Myosin-cross-reactive antigen (MCRA) protein from Bifidobacterium

breve is a FAD-dependent fatty acid hydratase which has a function in stress protection," Bmc

Biochemistry, vol. 12, Feb 2011.




[6] Y.-C. Joo, et al., "Production of 10-hydroxystearic acid from oleic acid by whole cells of


Journal of Biotechnology.


Macrococcus caseolyticus," Biochimie.




[9] S. van Pelt, et al., "Nitrile hydratase CLEAs: The immobilization and stabilization of an industrially

important enzyme," Green Chemistry, vol. 10, pp. 395-400, 2008.

[10] A. S. Bommarius and B. R. Riebel, "Introduction to Biocatalysis," in Biocatalysis, ed: Wiley-VCH

Verlag GmbH & Co. KGaA, 2005, pp. 1-18.

[11] F. H. Arnold, "Design by directed evolution," Accounts of Chemical Research, vol. 31, pp. 125-131,

Mar 1998.

[12] U. T. Bornscheuer and M. Pohl, "Improved biocatalysts by directed evolution and rational protein

design," Current Opinion in Chemical Biology, vol. 5, pp. 137-143, Apr 2001.

[13] L. G. Otten and W. J. Quax, "Directed evolution: selecting today's biocatalysts," Biomolecular

Engineering, vol. 22, pp. 1-9, Jun 2005.

[14] L. G. Otten, et al., "Enzyme engineering for enantioselectivity: from trial-and-error to rational

design?," Trends in Biotechnology, vol. 28, pp. 46-54, Jan 2009.

[15] J.-L. Reymond, Enzyme assays: High-throughput screening, genetic selection and fingerprinting:

Whiley-VCH, 2006.

[16] J. A. Hudson, et al., "Conversion of oleic acid to 10-hydroxystearic acid by two species of ruminal

bacteria," Applied Microbiology and Biotechnology, vol. 44, pp. 1-6, Dec 1995.

4.4 References

107

[17] D. L. H. Williams, "O-Nitrosation," in Nitrosation Reactions and the Chemistry of Nitric Oxide, ed

Amsterdam: Elsevier Science, 2004, pp. 105-115.

[18] I. A. Leenson, "Identification of primary, secondary, and tertiary alcohols - An experiment in

spectrophotometry, organic chemistry, and analytical chemistry," Journal of Chemical Education,

vol. 74, pp. 424-425, Apr 1997.

[19] W. Lyn, "Introduction," in Nitrosation Reactions and the Chemistry of Nitric Oxide, ed Amsterdam:

Elsevier Science, 2004, pp. xi-xii.

[20] J. H. Zhang, et al., "A simple statistical parameter for use in evaluation and validation of high

throughput screening assays," Journal of Biomolecular Screening, vol. 4, pp. 67-73, Apr 1999.

[21] R. Médici, et al., "A high-throughput screening assay for amino acid decarboxylase activity,"

Advanced Synthesis and Catalysis, vol. 353, pp. 2369-2376, 2011.

[22] B. Eastwood, et al. (2009). Assay Guidance Manual, Version 6 [from internet]. Available:

http://assay.nih.gov/assay/index.php/Table_og_Contents

Chapter 5

5 Preparation and properties of immobilized oleate hydratase as a cross-

linked enzyme aggregate (CLEA)

Aida Hiseni, Maria del Rosario Franco Berriel, Isabel W.C.E. Arends and

Linda G. Otten

Manuscript in preparation

Preparation and properties of immobilized oleate hydratase as a cross-linked enzyme aggregate (CLEA)

110

Abstract

The immobilization of oleate hydratase (OHase) from Elizabethkingia meningoseptica as

cross-linked enzyme aggregates (CLEA) is described. CLEA’s were prepared by

precipitation of OHase in cell-free extracts and subsequent cross-linking with

glutaraldehyde. The effects of different precipitating agents and different concentrations of

the cross-linking agent glutaraldehyde were investigated. In an optimized procedure ninety

percent ammonium sulfate saturation and 0.3wt% glutaraldehyde were used. Activity

recovery of 26% was achieved after 21 h cross-linking at 4°C. OHase CLEA’s had

increased activity over a range of different temperatures compared to that of the OHase

both in cell-free extracts as well as in whole cells.

5.1 Introduction

111

5.1 Introduction

The possibility of using renewable, plant-based resources for the production of fine

chemicals has generated wide interest in the field of industrial biotechnology. Reasons such

as fluctuating supply and price of the finite resource petroleum, and considerable

environmental issues have led to the development of new processes in the industry, where

petroleum-based products are being replaced by products derived from alternative and

sustainable sources [1-4]. One of the cheapest and most abundant biological raw materials

is vegetable oil [5].

Hydroxy fatty acids (HFA) have specific physical and chemical properties i.e. high

viscosity and reactivity, which make them suitable for the production of a number of

products, including resins, nylons, plastics, waxes, cosmetics and coatings [6]. They can be

obtained by chemical modification of unsaturated fatty acids using strong acids such as

sulfuric acid, followed by subsequent hydrolysis [6]. However, the resulting mixture of

several HFA’s requires costly downstream processing. Furthermore, regio- and

enantioselectivity is difficult to achieve. Therefore, the use of isolated enzymes and/or

microbial systems will offer significant advantages: Both the problem of selectivity as well

as the requirement of strong acids can be overcome.

The enzymatic hydration of oleic acid (OA) into 10-hydroxystearic acid (10-HSA) was first

described in a Pseudomonas strain [7]. Since then reports followed for a series of different

bacterial and eukaryotic microorganisms, such as Sphingobacterium thalpophilum [8]

Corynebacterium [9], Saccharomyces cerevisiae [6] and Stenotrophomonas nitritireducens

[10] with space-time yields ranging from 0.001 to 16 g l-1 h-1 for 10-HSA (Supplementary

table 5.1). Although over the years much research has been devoted to the optimization of

the fermentation conditions so that high productivities of the enantiomerically pure 10-

HSA can be obtained, rather little attention has been paid to the enzyme responsible for this

hydration reaction. Only recently, Bevers et al. [11] were able to recombinantly express

and characterize oleate hydratase (OHase) from Elizabethkingia meningoseptica (formerly

known as Pseudomonas sp. 3266), the same strain that Davis et al. [12] described 43 years

ago. This hydratase represents a new type of hydro-lyase, which is able to hydrate an

isolated carbon-carbon double bond (Figure 5.1) and is a possible biocatalyst for the

production of several alcohols and alkenes [13, 14]. The growing interest in this type of

hydro-lyases has been shown by many recent studies that have focused on finding and


112

characterizing oleate hydratases from other microorganisms for the production of 10-HSA

[15-18].

Figure 5.1 Reaction catalyzed by oleate hydratase (OHase): conversion of oleic acid into 10-hydroxystearic

acid.

The use of isolated enzymes in biocatalytic transformations allows for higher product

concentrations, less side reactions and simplified down-stream processing compared to

processes catalyzed by whole cells. Furthermore, diffusion limitations do not occur. On the

other hand, isolated enzymes may show high sensitivity to industrial conditions, which

often involve organic solvents, extreme pH’s or elevated temperatures. One strategy to

improve operational performances of enzymes in industrial processes is immobilization as

a means of stabilization. Furthermore, immobilization may improve other enzyme

properties, including selectivity or specificity or reduce enzyme inhibition by, for instance,

substrate or product [19]. A considerable number of immobilization techniques, such as

binding to a carrier, encapsulation in a polymeric matrix or cross-linking of enzyme

aggregates, are known to date [20-22]. However, there is no universal protocol. For every

enzyme the best immobilization method needs to be investigated. Hanefeld et al. [23] and

Garcia-Galan et al. [24] have reviewed enzyme immobilization and they highlighted some

important parameters that have to be taken into account when choosing a suitable

immobilization technique.

To our knowledge, the immobilization of oleate hydratase has not been reported to date.

As a first and straightforward methodology to immobilize OHase cross-linked enzyme

aggregates (CLEA) were prepared. This method offers the advantage that it does not lead

to ‘dilution of activity’ by usage of an carrier, has lower production costs through exclusion

of an expensive carrier, and produces a catalyst with highly concentrated activity [25]. In

this study, the results obtained for the immobilization of the overexpressed non-purified

O

OHoleic acid

O

OH

OH

10-hydroxystearic acid

H2OOHase

5.2 Materials and Methods

113

OHase are described. For this purpose, recombinant OHase from E. coli cell-free extracts

was aggregated and cross-linked using a bifunctional cross-linker glutaraldehyde.

Biochemical and biophysical properties as well as the efficiency of the CLEA biocatalyst

were investigated.

5.2 Materials and Methods

5.2.1 Bacterial strain, growth conditions and cell disruption E. coli TOP10 cells containing the plasmid pBAD-HISA-OH [11] were grown at 37°C in

TB medium with 100 μg ml-1 ampicillin until an OD600 value of 0.6 – 0.8 was reached.

Protein expression was induced with 0.2% arabinose (final concentration), followed by

cultivation at 28°C overnight. Cells were harvested by centrifugation (10.000 rpm, 10 min,

4°C; Sorvall), washed once with 20 mM Tris-HCl pH 8.0 and lysed in the same buffer with

a cell disruptor at the pressure of 1.5 kBar (Constant systems, IUL instruments). Cell-free

extract (CFE) was separated from cell debris by centrifugation at 10.000 rpm for 20 min at

4°C and stored on ice until further use. For long term storage, aliquots of CFE were frozen

in liquid nitrogen and stored at -80°C.

5.2.2 Precipitation procedure An amount of 0.1 ml of CFE (protein concentration: 4 mg ml-1) was added drop-wise to 0.9

ml precipitant (acetone, acetonitrile (ACN), ethanol, 2-propanol, 1,2-dimethoxyethane

(DME) or saturated ammonium sulfate), at room temperature (RT) and 4°C, respectively.

The resulting mixture was shaken at 400 rpm (Eppendorf Thermomixer) for 1 h, after which

the precipitated protein was separated at 13.200 rpm for 20 min (Eppendorf centrifuge).

Subsequently, the pellets were resuspended in 0.5 ml 20 mM Tris-HCl pH 8.0, and assayed

for activity using oleic acid as substrate.

5.2.3 Cross-linking procedure After protein precipitation 12.5 – 200 μl of 25wt% glutaraldehyde was added drop-wise

into the same tube and the mixture shaken at 400 rpm for 1 – 21 h. When the protein

concentration was varied from 1 to 24 mg ml-1 0.3wt% of glutaraldehyde was used. The

suspended CLEA’s were centrifuged (13.200 rpm, 30 min) and the supernatant was

removed. In order to remove non-cross-linked protein and the remaining glutaraldehyde,


114

CLEA’s were washed three times with 0.5 ml 20 mM Tris-HCl pH 8.0 and stored in the

same buffer on ice until further use. Washing supernatants were assayed for activity to

determine enzyme leakage.

For the scale-up procedure, the above described protocol was used, with 50-fold increased

amount of OHase. Briefly, 5 ml of CFE (protein concentration: 4 mg ml-1) was added drop-

wise to 45 ml saturated ammonium sulfate solution and the mixture shaken at 40 rpm

(Incubator shaker, Innova 44) and 4°C for 1 h. Subsequently, 625 μl of 25wt%

glutaraldehyde (endconc. 0.3wt%) was added and the mixture shaken further for 21 h. The

CLEA’s were centrifuged for 30 min at 4000 rpm and the supernatant removed. After

washing three times with 1 ml 20 mM Tris-HCl pH 8.0, the CLEA’s were stored in the

same buffer on ice until further use.

5.2.4 Activity assay OHase activity was determined using oleic acid as substrate. Unless otherwise stated, a

standard assay was performed with 0.5 ml final volume in 20 mM Tris-HCl pH 8.0,

containing 6 mM oleic acid and 2 μl of cell-suspension (55 mg ml-1), 2 μl of CFE (24 mg

ml-1) or a certain amount of CLEA, respectively. The mixtures were incubated at 30°C and

1400 rpm. After a desired time interval the reaction was stopped by the addition of 50 μl

3N hydrochloric acid (HCl), and substrate and product were extracted from the aqueous

layer. Prior to the extraction stearic acid (dissolved in acetone) was added, which served as

external standard, followed by addition of 50 μl saturated NaCl solution and the extraction

with one volume of dichloromethane (DCM). The mixtures were shaken for 5 min at 1400

rpm, centrifuged for 1 min at 13.200 rpm and 50 μl of the DCM phase was transferred into

a new tube. After drying with a SpeedVac Concentrator (Thermo), fatty acids were

derivatized [26, 27]. To the dried extracted fatty acids 25 μl of 2-bromoacetophenone (10

mg ml-1 in acetone) and 25 μl of triethylamine (10 mg ml-1 in acetone) were added and the

mixtures heated at 96°C for 15 min. 3.5 μl of acetic acid was added and the mixtures heated

for a further 5 min. After evaporation to dryness the samples were reconstituted in 0.1 ml

of ACN for HPLC analysis. Separation was performed with a 4.6 x 50 Merck Chromolith

SpeedROD RP-18e, using H2O-ACN mobile phase gradient (A: H2O with 0.1%, v/v

trifluoroacetic acid; B: ACN). The gradient consisted of 50% B over 3 min, 50 – 80% B

over 2 min and isocratic elution (80% B) over 7 min, at 1 ml min-1 and at a column

temperature of 50°C. Derivatized substrate and product were detected at 242 nm (SPD20A,

Shimadzu).


115

For quantitative analysis a linear relationship was established for the peak area ratios of

product versus external standard stearic acid.

5.2.5 Storage stability Storage stabilities of cell-suspension (55 mg ml-1), CFE (24 mg ml-1) and CLEA (0.2 mg

ml-1) were tested by storing the enzyme in 20 mM Tris-HCl pH 8.0 at 4°C and RT (21°C),

respectively, for several days. At various time points activities were determined using oleic

acid as substrate under standard assay conditions. Stabilities were given as residual

activities, calculated by taking the initial activity of the enzyme as 100%.

5.2.6 pH activity and temperature stability In order to investigate the pH effect on the enzyme activity, standard assay conditions were

used in buffers with varying pH values (100 mM sodium acetate, pH 3.0 – 6.0; 100 mM

potassium phosphate, pH 6.0 – 8.0; 50 mM Tris-HCl pH 8.0 – 9.0). Enzyme activity,

determined in 20 mM Tris-HCl pH 8.0 under standard conditions was designated as 100%.

Thermal stability was investigated by pre-incubating the enzyme at temperatures ranging

from 20 to 50°C in the absence of substrate for 20 min, cooling the enzyme solution on ice,

and then measuring the residual activity using the standard assay. Residual activities were

calculated by taking the initial activity of the enzyme as 100%.

5.2.7 Biocatalyst recovery Biocatalyst operational stability was studied using standard assay containing 10 mg ml-1 of

CLEA. After 1 h of reaction, product extraction and derivatization were performed as

described in section “Activity assay”. CLEA was removed from the interphase after

extraction, washed three times with 0.5 ml 20 mM Tris-HCl pH 8.0, and resuspended in

fresh buffer to perform a new reaction.


Recombinantly overexpressed OHase from E. meningoseptica was immobilized by self-

aggregation into cross-linked enzyme aggregates (CLEA’s). The crude enzyme is first

aggregated by a precipitating agent and subsequently covalently cross-linked using a bi-

functional agent glutaraldehyde [20]. One advantage of this method is that this

immobilization procedure is carrier-free. Protein sequence analysis of OHase revealed 50


116

lysine residues, which are potential sites for cross-linking. Since the crystal structure of

OHase has not yet been solved, it is not known if these residues are exposed to the solvent

or/and are close to the active site. However, given the fact that cross-linking is observed

(vide infra) it is reasonable to assume that a certain percentage is located on the enzyme

surface.

5.3.1 Selection of the best precipitating agent for CLEA preparation

As a first step, several precipitating agents were screened, including organic solvents and

an ammonium salt in an enzyme/precipitating agent ratio of 1:9 (v/v), both at 4°C and RT.

To evaluate the different precipitating agents, the possibility to reactivate the aggregated

enzyme after treatment (by dissolving in 20 mM Tris-HCl pH 8.0) is determined. As shown

in Figure 5.2, the highest activity recovery was obtained when 2-propanol and ammonium

sulfate were used. Using ammonium sulfate at RT results in more active enzyme, while no

good activity recovery was observed for 2-propanol.

Figure 5.2 Activity recovery of redissolved OHase aggregates after precipitation of 4 mg ml-1 CFE with 90%

(v/v) of precipitation agent. Assays were performed using standard conditions in 20 mM Tris-HCl pH 8.0

with oleic acid as substrate. Enzyme activity of the free and soluble enzyme was designated as 100% activity.

This can be explained by the fact that hydrophilic solvents such as acetonitrile, ethanol or

Acetone Acetonitrile EtOH 2-Propanol DME (NH4)2SO40

10

20

30

40

50

60

70

80

90

100

Act

ivit

y re

cove

ry [

%]

4°C RT


117

acetone are able to take up infinite amounts of water, which in this case is stripped off from

the enzyme surface [28]. As seen in Figure 5.2, this effect is for all solvents more

pronounced at RT and is observed even for the less polar solvents such as 2-propanol and

DME. For further studies both 2-propanol and ammonium sulfate procedures were used.

5.3.2 Cross-linking and the effect of glutaraldehyde concentration

OHase CLEA’s were prepared from the precipitate with different concentrations of

glutaraldehyde and three different incubation times, namely 1, 3 and 21 h at 4°C. In this

step of CLEA preparation it is important to define proper reaction conditions in order to

avoid excessive cross-linking, which can increase the rigidity of the enzyme and therefore

negatively influence the performance of the enzyme. In contrast, using too low

concentrations of the cross-linker may cause the formation of a highly flexible and small

CLEA, which cannot be centrifuged and therefore is not suitable for reuse. From the results

that are presented in Figure 5.3, a significant difference in activity recoveries was observed

when ammonium sulfate (Figure 5.3A) and 2-propanol (Figure 5.3B) were used as

precipitating agent.

The overall activity recovery was 100-fold higher for ammonium sulfate than that for 2-

propanol. The precipitation agent causes the enzyme to "freeze" in a certain conformation,

which is subsequently covalently stabilized by glutaraldehyde. Despite the relatively high

activity recovery obtained after precipitation of OHase with 2-propanol (Figure 5.2), hardly

any activity recovery was observed once the precipitated OHase has been covalently cross-

linked. This can be due to the fact that the precipitated OHase consisted of a rather

unfavorable and inactive conformation, which could be reactivated after redissolving in

buffer. On the contrary, this unfavorable conformation was stabilized after cross-linking

with glutaraldehyde, thus resulting in inactive CLEA particles. Similar results were

reported previously for a laccase from Trametes villosa, where good activity recoveries

were obtained after precipitation with 2-propanol. However, once the enzyme was cross-

linked, the catalytic activity decreased drastically [29].


118

Figure 5.3 Preparation of OHase CLEA’s (4 mg ml-1) using ammonium sulfate (A) and 2-propanol (B) as

precipitating agent in combination with different concentrations of the cross-linker glutaraldehyde. Cross-

linking time used was 1, 3 and 21 h. CLEA activity recovery is determined by comparison with the total

amount of units in 100 μl cell-free extract that was used for CLEA preparation.

Precipitation of OHase with ammonium sulfate resulted in significantly higher activity

recoveries. The remaining activity in the CLEA was inversely proportional to the

concentration of glutaraldehyde. With increasing glutaraldehyde concentration the

quaternary protein structure probably deforms in such way that the enzyme loses its

activity. Moreover, due to the high amount of lysine residues, the enzyme might become

rigid and is not able to undergo conformational changes upon binding to the substrate.

Therefore we chose a low glutaraldehyde concentration. Under these conditions a long

cross-linking time gives better results. For further tests CLEA’s were prepared using

0.3wt% of glutaraldehyde and an incubation time of 21 h. No enzyme leakage was detected

after 3 washing cycles with buffer when CLEA’s were prepared under these conditions.

This indicates that despite the low glutaraldehyde concentration all enzyme molecules were

properly cross-linked.

0 1 2 3 4 502468

10121416

0 1 2 3 4 50,00

0,05

0,10

0,15

Act

ivit

y re

cove

ry [

%]

1h 3h 21h

Glutaraldehyde [wt%]

(A)

(B)


119

Preparation of immobilized catalysts can introduce a new problem, i.e. diffusional

limitations of the substrate and product due to the large catalyst particle size. Oleic acid is

a C18 molecule that needs to be accommodated in the right position in the active site of

OHase. However, if the catalyst forms large aggregates, the accessibility of the active site

is impeded. This problem can be overcome by either reducing the aggregate particle size

by mechanical stirring [30], or by reducing the amount of enzyme used in the CLEA

preparation. We have prepared CLEA’s using 1 to 24 mg ml-1 of the CFE and the results

are shown in Figure 5.4. The activity recovery was the highest for the smallest CFE

concentration used. Although visually it was observed that more CLEA was obtained using

24 mg ml-1 of the CFE, it did not result in higher activity recovery. In general, the activity

recovery decreases with increasing protein concentration. From all these experiments we

deduced that the best way to make OHase CLEA’s is to prepare them with 90% ammonium

sulfate, 0.3wt% glutaraldehyde and 1 mg protein ml-1 at 4°C using a cross-linking time of

21 h.

Figure 5.4 Effect of enzyme concentration of the cell-free extract used for CLEA preparation on relative

activity recovery of OHase CLEA. Cell-free extracts of 1 mg ml-1 (A), 4 mg ml-1 (B), 10 mg ml-1 (C) and 24

mg ml-1 (D) were used.

A B C D0

5

10

15

20

25

30

Act

ivit

y re

cove

ry [

%]


120

5.3.3 Thermal stability and pH profile of OHase CLEA’s The thermal stability of OHase as cell-suspension, in CFE and as CLEA was investigated

by incubating the samples for 20 min at temperatures between 20 and 50°C in the absence

of the substrate oleic acid and measuring their residual activity with the standard assay. The

highest residual activity was obtained with CLEA (Figure 5.5). At 30°C the residual activity

for the CLEA was significantly higher (81%) than that of the CFE (34%). When pre-

incubated at 50°C, the CLEA still showed relative high activity (around 20%) compared to

cell-suspension and CFE (5 and 3%, respectively). This phenomenon of increased

temperature stability of CLEA has been observed earlier [30, 31]. This is probably due to

enhancement of the structure stability through inter- and intramolecular covalent cross-

links, which results in a molecule more resistant to conformational changes.

Figure 5.5 Comparison of temperature stabilities of OHase as CLEA (□), in cell-free extract (○) and as cell-

suspension (∆). Residual activities were assayed under standard conditions after the enzyme samples had

been incubated at the indicated temperature (20 – 50°C) in 20 mM Tris-HCl pH 8.0 for 20 min. Initial activity

determined under standard assay conditions was taken as 100%. Values are means of at least two independent

measurements.

The pH dependency of OHase activity was investigated using oleic acid as substrate. From

the results shown in Figure 5.6, it is obvious that OHase in CFE shows a different activity

profile compared to CLEA or cell-suspension. The optimal pH for OHase as free soluble

20 25 30 35 40 45 50 550

20

40

60

80

100

120 CLEA

Res

idua

l act

ivit

y [%

]

Temperature [°C]

cell-free extracts cell suspension


121

enzyme is at pH 7.0, on a broad plateau from pH 6.0 - 8.0, which is in agreement with

previously reported results [11]. It is shifted to a smaller peak in more alkaline conditions

(pH 8.0) for CLEA and cell-suspension. Acidic conditions (pH values 4.0 – 6.0) result in

inactive CLEA and cell-suspension, while CFE shows about 57% of the relative activity at

pH 5.0.

Figure 5.6 Comparison of pH profiles of OHase as CLEA (□), in cell-free extract (○) and as cell-suspension

(∆). Relative activities were determined using oleic acid as substrate in buffers with varying pH (4.0 – 8.6)

and compared to the activity measured using standard assay conditions (20 mM Tris-HCl pH 8.0).

There are many reasons, which could explain this different behavior. One important

parameter in this study is the physical state of oleic acid in biological aqueous systems.

Cistola and co-workers [32] discussed this in detail and concluded that oleic acid exists in

three different states, which is dependent on the pH of the solution and on the ionic strength.

At pH <7 oleic acid is in form of a stable oil phase and the carboxylic groups are protonated.

From pH 7 - 9 the degree of ionization increases, which results in the formation of more

structured lamellar system or large vesicles. Furthermore, this increase of ionization leads

also to increased fluidity. If the concentration at pH >9 is above its critical micelle

concentration (CMC ~6 μM), oleic acid starts to form micelles. Therefore, the pH

dependency of enzymatic activities is not only the result of kinetic parameters but also

4 5 6 7 8 90

20

40

60

80

100

120

CLEA

Res

idua

l act

ivit

y [%

]

pH

cell-free extracts

cell suspension


122

depends on the nature of the substrates and the products. Firstly, the above mentioned

fluidity of oleic acid at lower pH’s results in limited diffusivity, which again has big effect

on OHase activity as cell-suspension or CLEA. Secondly, the apparent pKa of monomeric

oleic acid is reported to be ~4.8, whereas it is 7.5 when incorporated into phospholipids

bilayer of the cell membranes [32]. As CLEA is a densely packed hydrophobic

environment, it may resemble a bilayer and have similar effects on the pKa of oleic acid.

The charge variation of the substrate and the enzyme itself, and the resulting structural

alterations may influence the binding of the substrate and therefore the catalytic activity of

the enzyme.

5.3.4 Storage stability of OHase CLEA’s The storage stability of the OHase in CFE, as cell-suspension and as CLEA was determined

at 4°C (Figure 5.7A) and RT (Figure 5.7B). In general, the residual activities for OHase in

CFE and as CLEA were greater for samples stored at 4°C. Complete loss of activity was

observed after 7 days of storage at RT. In contrast to that, the decrease of OHase activity

over time as cell-suspension is significantly lower at RT than that at 4°C. While about 50%

of the initial activity was lost after 7 days of storage at 4°C, the enzyme still retained 95%

of its initial activity when stored at RT for the same period of time. The cross-linking of

OHase leads to a slightly better storage stability at 4°C compared to the CFE. Nevertheless,

after 3 days of storage the activity decreases at the same degree as OHase in CFE or as cell-

suspension.


123

Figure 5.7 Comparison of storage stabilities at 4°C (A) and RT (B) of OHase as CLEA (□), in cell-free extract

(○) and as cell-suspension (∆) in 20 mM Tris-HCl pH 8.0. Stabilities were given as residual activities, calculated

by taking the initial activity of the enzyme as 100% under standard assay conditions.

0 2 4 6 8 10 12 14 160

20

40

60

80

100

120

cell-free extract

Res

idua

l act

ivit

y [%

]

Time [days]

cell suspension CLEA

0 2 4 6 8 10 12 14 160

20

40

60

80

100

120 CLEA

Res

idua

l act

ivit

y [%

]

Time [days]

cell-free extract cell suspension

(A)

(B)


124

5.3.5 Recycling of OHase CLEA’s For the recyclability studies, the CLEA’s were recovered and washed three times with

buffer after each cycle of operation and the activity assay repeated with the same CLEA’s.

The obtained results, which are depicted in Figure 5.8, show that the activity decreases with

every cycle of operation. In order to eliminate the possible negative influence of HCl, which

is needed for the extraction of fatty acids, we performed the extraction step without the

addition of HCl and compared the results with those obtained for the standard procedure.

Indeed, slightly higher residual activities were obtained when HCl was not used for the

extraction, and were 2.7-fold higher after the second cycle and 6.5-fold higher after the

third cycle. However, the loss of 70 and 87% of the initial activities after two and three

cycles, respectively, clearly indicates low stabilities of the CLEA’s. This effect can be

explained by the fact, that the product 10-HSA, which is a solid and starts to precipitate

after a certain point of time, accumulates inside and outside the aggregated biocatalyst and

blocks the structure for the diffusion of the substrate and the product. The inside portion

will not be washed out under current washing conditions, resulting in new substrate not

able to enter the CLEA’s and reach the active site of the OHase.

Figure 5.8 Recycling stability of OHase CLEA. Standard activity assay was performed and after 3 x washing

of the CLEA with buffer, the assay was repeated for a second and third time. The extraction step was

performed with and without the addition of 3N HCl.

1 2 30

20

40

60

80

100

120

Res

idua

l act

ivit

y [%

]

Cycle number

+ HCl - HCl

5.4 Conclusion

125

The same problem was observed by Cao et al., where penicillin G acylase CLEA starts to

accumulate the product ampicilline inside the CLEA matrix [33]. Another immobilization

technique may be considered at this point, such as soluble-insoluble supports [34]. With

this technique it would be possible to retain the enzyme in solution while centrifuging off

the solid product. With the subsequent lowering of the pH, the enzyme could be recovered

in its insoluble form for the next cycle [34]. An additional option would be the preparation

of OHase combined with another enzyme, such as lipase, which can convert 10-HSA to the

soluble product lactone [35]. Preliminary results indicate that this procedure warrants

further study.

5.3.6 Space-time yields Space-time yields for the production of 10-HSA by cell-suspension, CFE and CLEA’s were

calculated and compared with previously reported values (Supplementary table 5.1). Cell-

suspension and CFE produced roughly the same amount of the product 10-HSA with

volumetric productivities of 0.26 and 0.33 g l-1 h-1, respectively. The space-time yields

achieved with OHase CLEA’s were 4.7- and 6-fold lower than those obtained with the cell-

suspension and CFE, respectively. Nonetheless, they were up to 55-fold higher than

microbial productions with, for instance, Flavobacterium sp.DS5

(Supplementary table 5.1), performed under similar assay conditions (30°C) and with

similar yield (~6%). Interestingly, in the listed studies (Supplementary table 5.1) all

reported productivities above 5 g l-1 h-1 were achieved under optimized conditions, where

surfactants or organic solvents were used to increase the solubility of the substrate and the

product in the aqueous phase. It should be noted, however, that all obtained productivities

in this study were performed under non-optimized conditions. Although, investigations of

OHase immobilization as CLEA might not necessarily contribute directly to better 10-HSA

productivities and certainly leave room for optimization, they might guide further

investigations to make a stable and efficient biocatalyst for the industry.

5.4 Conclusion

This research describes the first steps towards a preparation of a biocatalyst for the

production of 10-HSA. For the first time OHase from E. meningoseptica has been

immobilized as cross-linked enzyme aggregates (CLEA’s). This immobilization technique


126

can be used to improve the biocatalytic properties of OHase. In the synthesis of 10-HSA,

CLEA’s preparation of OHase led to a 2.4-fold increase of biocatalyst stability at elevated

temperatures and better storage stabilities at cold temperatures in comparison with the

soluble enzyme in cell-free extracts or as whole-cells. The perspectives that we give here

could contribute to the preparation of a more successful biocatalyst for the application in

this industrial process. [36-39]

5.5 Supplemental Information

127

5.5 Supplemental Information S

trai

n C

ulti

vati

on

med

ium

Gro

wth

tem

p.

[°C

]

Ass

ay

bu

ffer

Ass

ay

tem

p.

[°C

]

Ass

ay

pH

Ass

ay

shak

ing

[rp

m]

Su

bst

rate

[g/l

]

Pro

du

ct

[g/l

]

Yie

ld

[%]

ST

Y

[g/l/

h]

Au

thor

Y

ear

Ref

.

Ent

eroc

occu

s fa

ecal

is

Enr

ichm

ent m

ediu

m

37

Enr

ichm

ent m

ediu

m-a

naer

obic

37

N

R

star

t aft

er

24h

0.9

0.8

89

0.01

1 H

udso

n 19

95

[26]

Ent

eroc

occu

s ga

llin

arum

H

B b

roth

45

/ 15

H

B m

edii

um

39

~6.5

90

2

1.9

97

0.02

7 M

orva

n 19

99

[36]

Fla

voba

cter

ium

sp.

DS

5 m

ediu

im

30

0.05

M K

-pho

spha

te

30

7.5

200

5.4

0.34

6

0.00

1 H

eo a

nd

Kim

2009

[3

9]

Lac

toba

cill

us s

p.

HB

bro

th

45

HB

med

iium

39

~6

90

2

1.52

76

0.

021

Mor

van

1999

[3

6]

Noc

ardi

a ch

oles

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m

YM

A

NR

0.

05 M

K-p

hosp

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ic

40

6.5

150

17.8

16

90

2.

7 K

orit

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1989

[3

7]

N. P

araf

fina

e N

BY

30

0.

05 M

K-p

hosp

hate

-ana

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ic

25

6.8

180

18

8 44

1.

6 L

atra

sse

1997

[3

8]

P. a

cidi

lact

ici

HB

bro

th

45 /

15

HB

med

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39

~6

.5

90

2 1.

86

93

0.02

7 M

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n 19

99

[36]

Sel

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inan

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t med

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37

E

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-ana

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37

NR

st

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0.63

70

0.

009

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1995

[2

6]

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ingo

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m th

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WF

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WF6

Mn

28

~7

350

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7 40

0.

07

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20

06

[8]

Ste

notr

ophm

onas

nitr

itir

educ

ens

Gro

wth

med

ium

+ O

A 2

8 0.

05 M

Tri

s-H

Cl +

0.0

5 %

Tw

een

80-a

naer

obic

35

7.5

200

30

31.5

10

5 7.

9 K

im

2010

[1

0]

Ste

notr

ophm

onas

mal

toph

ilia

N

R

NR

C

itrat

e-ph

osph

ate

buff

er +

0.0

5 %

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een

40

35

6.5

NR

50

40

80

5

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2012

[1

7]

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cont

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Has

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from

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inib

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35

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[18]

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from

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-med

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C

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+ 0

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35

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2012

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from

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His

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(fro

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Supplementary table 5.1 Comparison of space-time yield for the production of 10-HAS by OHase.


128

5.6 References

[1] R. A. Sheldon, "Green solvents for sustainable organic synthesis: state of the art," Green Chemistry,

vol. 7, pp. 267-278, 2005.

[2] N. Ran, et al., "Recent applications of biocatalysis in developing green chemistry for chemical

synthesis at the industrial scale," Green Chemistry, vol. 10, pp. 361-372, 2008.

[3] D. M. Alonso, et al., "Catalytic conversion of biomass to biofuels," Green Chemistry, vol. 12, pp.

1493-1513, 2011.

[4] R. A. Sheldon, "Utilisation of biomass for sustainable fuels and chemicals: Molecules, methods and

metrics," Catalysis Today, vol. 167, pp. 3-13, 2011.

[5] A. S. Carlsson, "Plant oils as feedstock alternatives to petroleum - A short survey of potential oil

crop platforms," Biochimie, vol. 91, pp. 665-670, Jun 2009.

[6] S. H. Elsharkawy, et al., "Microbial oxidation of oleic-acid," Applied and Environmental

Microbiology, vol. 58, pp. 2116-2122, Jul 1992.

[7] L. L. Wallen, et al., "The microbiological production of 10-hydroxystearic acid from oleic acid,"

Archives of Biochemistry and Biophysics, vol. 99, pp. 249-253, 1962.

[8] T. M. Kuo and W. E. Levinson, "Biocatalytic production of 10-hydroxystearic acid, 10-ketostearic

acid, and their primary fatty amides," Journal of the American Oil Chemists Society, vol. 83, pp.

671-675, Aug 2006.

[9] C. W. Seo, et al., "Hydration of squalene and oleic-acid by Corynebacterium sp. S-401,"

Agricultural and Biological Chemistry, vol. 45, pp. 2025-2030, 1981.





[12] E. N. Davis, et al., "Microbial hydration of cis-9-alkenoic acids " Lipids, vol. 4, pp. 356-362, 1969.

[13] P. Marliere, "Method for producing an alkene comprising the step of converting an alcohol by an

enzymatic dehydration step," 2011.











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pp. 1-9, 2011.

[19] C. Mateo, et al., "Improvement of enzyme activity, stability and selectivity via immobilization

techniques," Enzyme and Microbial Technology, vol. 40, pp. 1451-1463, May 2007.

[20] R. A. Sheldon, "Cross-linked enzyme aggregates (CLEAs): stable and recyclable biocatalysts,"

Biochemical Society Transactions, vol. 35, pp. 1583-1587, Dec 2007.

[21] K. Hernandez and R. Fernandez-Lafuente, "Control of protein immobilization: Coupling

immobilization and site-directed mutagenesis to improve biocatalyst or biosensor performance,"

Enzyme and Microbial Technology, vol. 48, pp. 107-122, Feb 2011.

[22] R. A. Sheldon, "Cross-Linked Enzyme Aggregates as Industrial Biocatalysts," Organic Process

Research & Development, vol. 15, pp. 213-223, 2011.

[23] U. Hanefeld, et al., "Understanding enzyme immobilisation," Chemical Society Reviews, vol. 38,

pp. 453-468, 2009.

[24] C. Garcia-Galan, et al., "Potential of different enzyme immobilization strategies to improve enzyme

performance," Advanced Synthesis and Catalysis, vol. 353, pp. 2885-2904, 2011.

[25] R. A. Sheldon, "Enzyme immobilization: The quest for optimum performance," Advanced Synthesis

& Catalysis, vol. 349, pp. 1289-1307, Jun 2007.

[26] J. A. Hudson, et al., "Conversion of oleic acid to 10-hydroxystearic acid by two species of ruminal

bacteria," Applied Microbiology and Biotechnology, vol. 44, pp. 1-6, Dec 1995.

[27] A. Mehta, et al., "Rapid quantitation of free fatty acids in human plasma by high-performance liquid

chromatography," Journal of Chromatography B, vol. 719, pp. 9-23, Nov 1998.

[28] A. S. Bommarius and B. R. Riebel, "Biocatalysis in Non-conventional Media," in Biocatalysis, ed:

Wiley-VCH Verlag GmbH & Co. KGaA, 2005, pp. 339-372.

[29] I. Matijosyte, et al., "Preparation and use of cross-linked enzyme aggregates (CLEAs) of laccases,"

Journal of Molecular Catalysis B-Enzymatic, vol. 62, pp. 142-148, Feb 2010.

[30] B. S. Aytar and U. Bakir, "Preparation of cross-linked tyrosinase aggregates," Process Biochemistry,

vol. 43, pp. 125-131, Feb 2008.

[31] M. E. Ortiz-Soto, et al., "Evaluation of cross-linked aggregates from purified Bacillus subtilis

levansucrase mutants for transfructosylation reactions," Bmc Biotechnology, vol. 9, p. 68, 2009.

[32] D. P. Cistola, et al., "Ionization and phase behavior of fatty acids in water: application of the Gibbs

phase rule," Biochemistry, vol. 27, pp. 1881-1888, Mar 1988.

[33] L. Q. Cao, et al., "Cross-linked enzyme aggregates: A simple and effective method for the

immobilization of penicillin acylase," Organic Letters, vol. 2, pp. 1361-1364, May 2000.

[34] J. Zhou, "Immobilization of cellulase on a reversibly soluble-insoluble support: Properties and

application," Journal of Agricultural and Food Chemistry, vol. 58, pp. 6741-6746, 2010.



vol. 91, pp. 1029-1036, 2011.


130

[36] B. Morvan and K. N. Joblin, "Hydration of oleic acid by Enterococcus gallinarum, Pediococcus

acidilactici and Lactobacillus sp. isolated from the rumen," Anaerobe, vol. 5, pp. 605-611, 1999.

[37] S. Koritala, et al., "Microbial conversion of oleic acid to 10-hydroxystearic acid," Applied

Microbiology and Biotechnology, vol. 32, pp. 299-304, 1989.

[38] A. Latrasse, et al., "Conversion of oleic acid to 10-hydroxystearic acid by Nocardia paraffinae,"

Biotechnology Letters, vol. 19, pp. 715-718, 1997.

[39] S. H. Heo, et al., "Production of oxygenated fatty acids from vegetable oils by Flavobacterium sp.

strain DS5," New Biotechnology, vol. 26, pp. 105-108, 2009.

Chapter 6

6 Conclusions and future prospects

Conclusions and future prospects

132

The study described in this thesis was set out to explore the potential of newly discovered

hydratases, carotenoid 1,2-hydratases (CrtC) from photosynthetic bacteria Rubrivivax

gelatinousus and Thiocapsa roseopersicina and oleate hydratase (OHase) from

Elizabethkingia meningoseptica, for their use as biocatalysts in industrial processes. In

order to do so, it was important to gain more insight in the activity, stability substrate scope

and structure-function relationship of these enzymes. When the research described in this

thesis started, limited or no literature was available on structural and mechanistic properties

of these two groups of hydratases. The work presented in this thesis sought to broaden the

structural, mechanistic and sequence-based knowledge required to define future research.

The main findings of this thesis are presented in the ‘Summary’ section of this thesis. This

chapter will evaluate the original question, which was posed at the start of the research

project, namely to evaluate if carotenoid 1,2-hydratases and oleate hydratases have

potential as industrial biocatalysts.

6.1 Carotenoid 1,2-hydratase

A comparative study of homologous carotenoid 1,2-hydratases sequences was successfully

applied to identify conserved residues within selected regions across the gene. Next to

mutagenesis of selected key residues and biochemical methods, which have proven to be

effective for determining the potential catalytic mechanism, the elucidation of a three-

dimensional structure is also pivotal to unravel the structure-function relationship of this

group of enzymes. Although, we made a homology model, which appeared to be accurate

since it can explain the mutants we made, the availability of a three-dimensional structure

would allow the application of site-saturated mutagenesis in residues governing the

property of interest and the generation of synergistic effects of neighboring mutations. In

addition, docking models can be used to predict the preferred orientation of substrates and

residues involved.

The work presented in this thesis has helped to shed light on structural and mechanistic

properties of carotenoid 1,2-hydratases. But also, it has clearly shown that their low

activities at present might prevent their successful utilization in industrial processes. The

fact that they are membrane-bound may have some advantages as it can be regarded as

natural immobilization. On the other hand, most enzymes used in industrial settings are

extracellular enzymes, as they require less downstream processing prior to catalysis and

Carotenoid 1,2-hydratase

133

they are more stable to external environmental perturbations. The overproduction of

carotenoid 1,2-hydratases is primarily limited to the membrane capacity of the host

organism. Therefore, the development of an effective expression system is of a high

importance. For instance, the usage of a host that is less sensitive to the toxic effects of

overexpressed membrane or membrane-bound proteins, which have possibly formed

aggregates due to space constrains, will form more biomass that can produce more protein

[1-6].

Another limitation of carotenoid 1,2-hydratases is the very low specific activity. This might

be due to either non-optimal assay conditions or the intrinsic property of membrane-bound

enzymes. High specific activity ensures sufficient space-time yield and makes the process

and reactor costs low. Therefore, the catalytic activity of these hydratases needs to be

improved, either by random mutagenesis, followed by site-saturated mutagenesis as a

means to further improve beneficial mutations obtained through random mutagenesis, or

by immobilization. As demonstrated by Umeno et al. [7] the evolvability of carotenoid

biosynthetic enzymes is remarkable. Carotenoid biosynthesis pathway appears to consist

of two groups of enzymes: ‘gatekeeper’ enzymes, which are located at the earliest steps of

the pathway and/or at important branch points and enzymes that are ‘locally specific’, i.e.

they recognize a particular structural motif of a possible substrate. The former dictate the

product diversity by allowing only certain molecules to enter the pathway and are therefore

expected to be very specific. Several researches have demonstrated the ability of these

enzymes to acquire new specificities after a limited genetic change. In contrast to the high

specificity of the ‘gatekeeper’ enzymes, ‘locally specific’ enzyme do not require this

property as they are placed at several locations in the pathway and are supposed to convert

more than one substrate. In case of a change upstream, which would allow a new compound

to enter the pathway, downstream enzymes are expected to be able to also metabolize this

new compound. Carotenoid 1,2-hydratase is such an enzyme that is less specific and

appears to recognize only a particular motif of the structure (-end group according to

carotenoid nomenclature). Thus, this fact predicts a good prospect for this enzyme to

metabolize unnatural substrates as long as it retains the required motif.

Overall, if it would be possible to address these two main issues, i.e. overexpression and

catalytic activity, this would significantly enlarge the potential of carotenoid 1,2-

hydratases. Future research could be the focus on optimizing additional enzyme properties,

such as substrate promiscuity, thermostability, solvent tolerance and activity at high


134

substrate concentration. Nevertheless, the time that needs to be invested in order to make

this enzyme suitable for industrial process has to be balanced against the chances for

success in reasonable timeframe.

6.2 Oleate hydratase

Another hydratase with high potential for industrial biocatalysis is oleate hydratase. Since

the first paper was published on characterization of recombinantly overexpressed oleate

hydratase by Hagen et. al [8], the literature on oleate hydratase significantly expanded with

several papers, mostly with the work from the group of Oh [9-15] and Feussner [16, 17].

While the production of 10-hydroxystearic acid in high yields and biochemical

characterization of several recombinantly overexpressed oleate hydratases have been

extensively studied by Oh et al., the latter author succeeded very recently in elucidating the

first three-dimensional structure of the oleate hydratase from Lactobacillus acidophilus

(LAH). Despite the significant contribution of this work to the better understanding of

structural properties of LAH, the precise location of the active site and the involved

catalytic mechanism still remain unclear. The researchers were able to elucidate the crystal

structure of an apo-LAH and LAH co-crystallized with linoleic acid (LA-LHA), but both

without the co-factor FAD (lost during enzyme purification). In addition, the binding of the

substrate LA in the active site, was hindered by two MPD (2-methyl-2,4-pentanediol; used

as precipitating agent) molecules so that it stayed bound at the entrance of the substrate

channel. Nevertheless, given the fact that the LAH shares more than 50% amino acid

sequence similarity with the oleate hydratase from E. meningoseptica, which was studied

in this thesis work, it is likely to assume that these novel structural insights will partly help

to increase our understanding of the structural properties of oleate hydratases. Future

research of the study presented in this thesis, however, could attempt to crystallize oleate

hydratase from E. meningoseptica in the presence of a substrate analog in order to locate

the active site and amino acids necessary for binding of the substrate and catalysis. Once

the catalytic mechanism is unraveled and together with other newly gained structural data,

the analysis of the created mutants (Chapter 4) can be performed in more detail, which

again could give an indication of directions for further improvement of the catalytic activity

or substrate specificity.

High-throughput screening assay

135

With regard to formulating the enzyme by means of immobilization through cross-linking

(Chapter 5) in order to make it suitable for an economically feasible industrial process,

other methods than cross-linking need to be explored. As the product 10-hydroxystearic

acid is a solid, a method needs to be chosen which prevents the accumulation of the solid

product within the immobilized enzyme complex. This might be accomplished by

immediate removal of the product either by a physical isolation through a second phase or

by reacting the product further to produce a water soluble intermediate or a final product.

Overall, the good activity of OHase together with its straightforward expression system

makes this enzyme a promising candidate for future directed evolution studies.

6.3 High-throughput screening assay

A crucial part of any directed evolution study is a robust and selective high-throughput

screening assay, which can be used to isolate mutants with improved properties. The assay

allows testing of large number of mutants in a quick and efficient way. The developed

colorimetric high-throughput screening assay (Figure 6.1) with oleate hydratase as model

enzyme (Chapter 4) that can be used for the detection of primary, secondary and tertiary

alcohols, provides an adequate basis for screening a large amount of generated mutants.

Although the assay has been proven to be robust and applicable for many different

substrate/product pairs, the signal window needs further improvement.

The current status is that by using purified oleate hydratase instead of crude E. coli extracts,

the signal window could significantly be increased, allowing sufficient discrimination

between the noise and enzymatic activity. This result increases the chance of finding

mutants with improved properties, without having the risk of detecting false positive

mutants. Although enzyme purification can be performed in a high-throughput manner, it

would be of more interest to follow optimization approaches that prevent too high signal of

the noise or that increase the overexpression yield of the enzyme in a microtiter plate.

Besides the colorimetric assay developed by us, also calorimetric assays could contribute

to HTS of activity of these enzymes [18]. Calorimetry measures the absorbed or exerted

heat during bond breaking or making. This means that this assay is the universal assay for

enzymatic activity using real substrates instead of artificial ones. It is already shown in our

lab that the enthalpy of hydration is large enough to measure enzyme kinetics. Overall, this


136

will allow the rate evaluation of activity, stability and substrate scope of large libraries of

mutants.

Figure 6.1 Schematic overview of the high high-throughput screening assay. The assay was developed for

the detection of enzymatic hydration activity assessed by selective spectrophotometric detection of alcohols.

Next to OHase and CrtC, other hydratases that also operate on (non-activated) double bonds

are continuously discovered by microbiologists [19]. Glycerol dehydratases (EC 4.2.1.30)

might be of interest for the dehydration of glycerol (a huge by-product of biodiesel

production) to get the building block 3-hydroxypropanol [20]. Also the very fast enzyme

enoyl-CoA hydratase (EC 4.2.1.17) could be investigated for promiscuous activities [21].

Although the promiscuous activity of an enzyme is usually orders of magnitudes lower than

the activity with the natural substrate, this could still be acceptable because of the almost

diffusion limited rate of this enzyme. The linalool dehydratase-isomerase (EC 4.2.1.127)

might be an interesting enzyme when looking at fatty acids, since it has its function in the

production of myrcene from linalool, a C10-terpene [22]. Hydroxycinnamoyl-CoA

hydratase-lyase (EC 4.2.1.101) could be used in the biosynthesis of the flavour compound

vanillin [23]. A key step is the addition of water to the thioester of ferulic acid, catalysed

by hydroxycinnamoyl-CoA hydratase lyase (HCHL, formerly feruloyl-CoA hydratase).

The hydration is immediately followed by a retro-aldol reaction, releasing the vanillin and

acetyl-CoA [24].

ACTIVITY SOURCE

ACTIVITY/SCREENING

ASSAY

ABSORPTION

MEASUREMENT

PLATE PROCESSING

DATA PROCESSING

High-throughput screening assay

137

In conclusion, I believe there is a bright future for hydratase enzymes and that there are

enough leads for follow-up research on the enzymatic hydration of double bonds. Our assay

can serve to explore these novel enzymes for their potential.


138

6.4 References

[1] S. Wagner, et al., "Tuning Escherichia coli for membrane protein overexpression," Proceedings of

the National Academy of Sciences, vol. 105, pp. 14371-14376, September 23, 2008 2008.

[2] G. J. Gopal and A. Kumar, "Strategies for the production of recombinant protein in escherichia coli,"

Protein Journal, vol. 32, pp. 419-425, 2013.

[3] N. Gul, et al., "Evolved escherichia coli strains for amplified, functional expression of membrane

proteins," Journal of Molecular Biology, vol. 426, pp. 136-149, 2014.

[4] T. M. Lo, et al., "Microbial engineering strategies to improve cell viability for biochemical

production," Biotechnology Advances, vol. 31, pp. 903-914, 2013.

[5] D. M. Molina, et al., "Engineering membrane protein overproduction in Escherichia coli," Protein

Science, vol. 17, pp. 673-680, 2008.

[6] E. Gordon, et al., "Effective high-throughput overproduction of membrane proteins in Escherichia

coli," Protein Expression and Purification, vol. 62, pp. 1-8, 2008.







pp. 1-9, 2011.



vol. 95, pp. 929-937, 2012.



[12] J. S. Kim, et al., "Identification and characterization of a novel nitrilase from Pseudomonas

fluorescens Pf-5," Applied Microbiology and Biotechnology, vol. 83, pp. 273-283, May 2009.

[13] K. R. Kim and D. K. Oh, "Production of hydroxy fatty acids by microbial fatty acid-hydroxylation

enzymes," Biotechnology Advances, 2013.






[16] A. Volkov, et al., "Crystal structure analysis of a fatty acid double-bond hydratase from

Lactobacillus acidophilus," Acta Crystallographica Section D: Biological Crystallography, vol. 69,

pp. 648-657, 2013.

References

139




[18] M. J. Todd and J. Gomez, "Enzyme kinetics determined using calorimetry: A general assay for

enzyme activity?," Analytical Biochemistry, vol. 296, pp. 179-187, 2001.



[20] S. Kwak, et al., "Biosynthesis of 3-hydroxypropionic acid from glycerol in recombinant Escherichia

coli expressing Lactobacillus brevis dhaB and dhaR gene clusters and E. coli K-12 aldH,"

Bioresource Technology, vol. 135, pp. 432-439, 2013.

[21] G. Agnihotri and H. W. Liu, "Enoyl-CoA hydratase: Reaction, mechanism, and inhibition,"

Bioorganic and Medicinal Chemistry, vol. 11, pp. 9-20, 2003.

[22] D. Brodkorb, et al., "Linalool dehydratase-isomerase, a bifunctional enzyme in the anaerobic

degradation of monoterpenes," Journal of Biological Chemistry, vol. 285, pp. 30436-30442, 2010.

[23] P. M. Leonard, et al., "The 1.8 Å resolution structure of hydroxycinnamoyl-coenzyme a hydratase-

lyase (HCHL) from Pseudomonas fluorescens, an enzyme that catalyses the transformation of

feruloyl-coenzyme A to vanillin," Acta Crystallographica Section D: Biological Crystallography,

vol. 62, pp. 1494-1501, 2006.

[24] J. P. Bennett, et al., "A ternary complex of hydroxycinnamoyl-CoA hydratase-lyase (HCHL) with

acetyl-CoA and vanillin gives insights into substrate specificity and mechanism," Biochemical

Journal, vol. 414, pp. 281-289, 2008.

Summary/Samenvatting


142

Summary

The rapid development in the field of biotechnology over the last four decades, in addition

to an increasing recognition that we have limited resources and thus need to move to

renewable raw materials, have been drivers for the chemical industry to look at enzymes as

novel catalysts. In addition, enzymes are highly specific, thereby leading to high regio- and

chiral selectivities and less/no unwanted side reactions and byproducts. They generally

operate under mild conditions, resulting in energy savings. Overall, it is safe to state that

enzymes contribute to the environmentally sustainable processing.

Hydratases catalyze the non-hydrolytic and non-oxidative addition and/or removal of a

water molecule to a carbon-carbon double bond. From a chemical point of view, this

reaction is difficult to achieve and requires harsh conditions, such as high temperature and

low pH. In contrast, the enzymatic route proceeds under very mild conditions in a neutral

aqueous environment, yielding products in high yields and without undesired side

reactions. Therefore, there is significant interest in the application of hydratases as efficient,

selective and environmentally friendly biocatalyst. The research of this thesis focused on

two hydratases: carotenoid 1,2-hydratase (CrtC) and oleate hydratase (OHase).

CrtC is an enzyme found in the biosynthetic pathway of carotenoids. 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. Another hydratase that has raised the attention of

researchers is OHase. OHase catalyzes the conversion of oleic acid (OA) into (R)-10-

hydroxystearic acid (10-HSA), a high-added value product used for the production of a

number of products, including resins, nylons, plastics, waxes, cosmetics and coatings. This

hydratase, as well as the carotenoid 1,2-hydratase, represents a new type of hydro-lyase as

it is able to hydrate an isolated carbon-carbon double bond.

In literature, a limited amount of data was available on the biochemical, structural and

mechanistic properties of these two hydratases. Therefore, it was decided to study these

enzymes with a focus on their structure-function relationship, thus allowing the evaluation

of the potential of these hydratases as industrial biocatalysts.

In Chapter 1, a general overview is given on enzymes and their application as biocatalysts

in various industries. Also, protein engineering tools used to overcome the limitations of

natural enzymes as biocatalysts at typical operating industrial conditions, such as high

substrate and salt concentrations, use of organic solvents, etc., are introduced. Our present


143

knowledge on hydro-lyases and their utilization in industrial processes is highlighted.

Special attention is given to aspects of the structure-function relationship of the two studied

hydratases CrtC and OHase.

Chapter 2 describes the detailed biochemical characterization of two newly discovered

CrtC’s from photosynthetic bacteria Rubrivivax gelatinosus and Thiocapsa roseopersicina.

In order to investigate the biochemical properties, the enzymes were recombinantly

overexpressed and purified by affinity chromatography. It was demonstrated that both

CrtC’s were able to cofactor independently catalyze the conversion of the natural substrate

lycopene to 1-HO- and 1,1′-(HO)2-lycopene. In addition, low activity was detected with an

unnatural substrate geranylgeraniol (C20 substrate), which functionally resembles the

natural C40 substrate lycopene. Both CrtC’s are stable at a broad and suitable temperature

and pH range, which makes them attractive for green hydration reactions in industrial

applications. Although, the amino acid sequences of RgCrtC and TrCrtC differ by only one

amino acid (406 vs. 405), a structural difference has been observed by means of SDS-

PAGE and MS analysis. Whereas RgCrtC is expressed as a 44 kDa protein, TrCrtC exist

as a 38 kDa protein, most likely caused by autocatalytic processing.

In order to increase our understanding of the structure and mechanism of CrtC’s from

photosynthetic bacteria, protein engineering techniques site-directed evolution and semi-

rational mutagenesis were applied (Chapter 3). By generating alanine point-mutants of

selected amino acid positions, it was possible to elucidate the role of the amino acids

His239, Trp241, Tyr266 and Asp268 in RgCrtC (and the corresponding amino acids in

TrCrtC) and identify them as key residues, which are directly involved in the catalytic

reaction. By analyzing a partial 3D structure obtained by homology modeling with the

distantly related putative AttH protein from Nitrosomonas europaea it could be shown that

all identified amino acids were in close proximity to each other. All these results indicate

that the aforementioned amino acid residues are involved in the catalytic cycle.

When considering the generation of tailor-made biocatalysts for a successful utilization in

industrial processes, the availability of a suitable high-throughput screening or selection

method is a pre-requisite. The existence of such an assay will also determine the method of

choice for protein engineering. Due to limited existing structural and mechanistic

knowledge on hydratases studied in this thesis, a high-throughput screening assay for the

detection of alcohols, products of hydrating enzymes such as CrtC and OHase, was

developed (Chapter 4), which allows rapid screening of a large number of variants within

a reasonable timeframe. OHase from Elizabethkingia meningoseptica was used as the


144

model enzyme to examine and characterize the capability of the developed method for

enabling an automated set up. The assay was able to detect primary, secondary and tertiary

alcohols in the presence of fatty acids as well as small cyclic and acyclic unsaturated

alkenes as substrates. Besides protein engineering techniques to improve the operational

performance of enzymes (e.g. thermo-stability, activity and solvent tolerance)

immobilization is a convenient approach towards stabilization. In Chapter 5 we report on

the immobilization of OHase as cross-linked enzyme aggregates (CLEA). For this purpose,

recombinant OHase from E. coli cell-free extracts was aggregated and cross-linked using a

bifunctional cross-linker glutaraldehyde. With an activity recovery of 26% after 21h cross-

linking at 4°C, CLEA’s preparation of OHase led to a 2.4-fold increase of biocatalyst

stability at elevated temperatures and better storage stabilities at cold temperatures.

Furthermore, up to 55-fold higher space-time yields were achieved with OHase CLEA’s

compared to microbial productions.

Overall, the work presented in this thesis has contributed to an understanding of the

structure-function relationship of two newly discovered hydratases: carotenoid 1,2-

hydratase and oleate hydratase. Furthermore, it may contribute to the development of a

biocatalyst that can be used for the production of high-added value compounds in industrial

processes.


145

Samenvatting

De snelle ontwikkelingen op het gebied van de biotechnologie in de afgelopen vier

decennia, samen met de toegenomen bewustwording dat we over beperkte hoeveelheden

aan fossielen grondstoffen beschikken, en er een noodzaak bestaat om over te gaan naar

hernieuwbare grondstoffen, zijn aanleiding geweest voor de chemische industrie om

enzymen te ontwikkelen als nieuwe en hernieuwbare katalysatoren. De selectiviteit van

enzymen zorgt ervoor dat minder of geen ongewenste nevenreacties en bijproducten

worden geproduceerd. Daarnaast werken enzymen onder milde condities, wat bijdraagt aan

energiezuinige procescondities. Over het algemeen kan men dus stellen dat het gebruik van

enzymen als katalysatoren kan bijdragen aan duurzame processen.

Hydratases katalyseren de niet-hydrolytische en niet-oxidatieve additie en/of eliminatie van

een watermolecuul aan een koolstof-koolstof dubbele binding. Vanuit chemisch oogpunt is

deze reactie moeilijk uit te voeren en vereist extreme condities, zoals hoge temperaturen en

lage pH. Daarentegen verloopt de enzymatische route onder zeer milde omstandigheden in

een neutrale waterige omgeving, waardoor producten in hoge opbrengsten en zonder

ongewenste nevenreacties gemaakt kunnen worden. Daarom is het zeer relevant om

hydratases te bestuderen als efficiënte, selectieve en milieuvriendelijke biokatalysatoren.

Het onderzoek van dit proefschrift richt zich op twee enzymen: carotenoïd 1,2- hydratase

(CrtC) en oleaat hydratase (OHase).

CrtC is een enzym dat aanwezig is in de biosynthese route van carotenoïden. CrtC

introduceert een tertiaire hydroxyl groep in een carotenoïde molecuul door toevoeging van

water aan de koolstof-koolstof dubbele binding op de C-1 positie. Een ander hydratase dat

de aandacht van onderzoekers heeft getrokken is OHase. OHase katalyseert de omzetting

van oliezuur (OA) in (R)-10-hydroxystearinezuur (10-HSA), een product met hoge

toegevoegde waarde voor de productie van materialen zoals harsen, nylon, kunststoffen,

wassen, cosmetica en coating. Dit hydratase, evenals de carotenoïd 1,2-hydratase,

vertegenwoordigt een nieuw type hydro-lyase dat in staat is om een geïsoleerde koolstof-

koolstof dubbele binding te hydrateren.

In de literatuur is een beperkte hoeveelheid gegevens beschikbaar met betrekking tot de

biochemische, structurele en mechanistische eigenschappen van deze twee hydratases.

Daarom werd besloten om deze enzymen te bestuderen om meer inzicht te krijgen in de


146

structuur-functie relatie en zodoende het potentieel van hydratases als biokatalysatoren in

industriële processen in kaart te brengen.

In Hoofdstuk 1 wordt een algemeen overzicht gegeven van enzymen en hun toepassing als

biokatalysatoren in diverse industrieën. Ook ‘protein engineering’ technieken die worden

gebruikt om de beperkingen van de natuurlijke enzymen als biokatalysatoren bij typische

operationele industriële omstandigheden te overwinnen, zoals hoge substraat- en

zoutconcentraties, gebruik van organische oplosmiddelen, etc., worden geïntroduceerd.

Onze huidige kennis over hydro-lyasen en hun gebruik in industriële processen wordt

daarbij benadrukt. Speciale aandacht wordt besteed aan aspecten van de structuur-functie

relatie van de twee bestudeerde hydratases CrtC en OHase.

Hoofdstuk 2 beschrijft de gedetailleerde biochemische karakterisering van twee nieuw

ontdekte CrtC’s van de fotosynthetische bacteriën Rubrivivax gelatinosus en Thiocapsa

roseopersicina. Om de biochemische eigenschappen te onderzoeken, werden de enzymen

recombinant tot overexpressie gebracht en gezuiverd middels affiniteitschromatografie. Er

werd aangetoond dat beide CrtC’s zonder de hulp van een cofactor de omzetting van het

natuurlijke substraat lycopeen naar 1-HO- en 1,1'-(HO)2-lycopeen konden katalyseren.

Bovendien werd lage activiteit gedetecteerd met het niet-natuurlijke substraat

geranylgeraniol (C20 substraat), dat structureel lijkt op het natuurlijke substraat lycopeen.

Beide CrtC’s zijn stabiel in een breed en gemiddeld temperatuur- en pH-bereik, waardoor

ze aantrekkelijk worden voor groene hydratatie reacties in industriële toepassingen. Hoewel

de theoretische eiwitgrootte van RgCrtC en TrCrtC slechts in één aminozuur verschilt (406

versus 405) is een structureel verschil waargenomen door middel van SDS-PAGE en MS-

analyse. Terwijl RgCrtC als een 44 kDa eiwit tot expressie wordt gebracht, bestaat TrCrtC

als een 38 kDa eiwit, waarschijnlijk veroorzaakt door autokatalytische verwerking.

Om onze kennis van de structuur en het mechanisme van CrtC’s uit fotosynthetische

bacteriën te verhogen, werden protein engineering technieken semi-gerichte evolutie (in

het engels semi-directed evolution) en semi-rationale mutagenese toegepast (Hoofdstuk

3). Door het genereren van specifieke alanine punt-mutanten van geselecteerde

aminozuurposities, was het mogelijk om de rol van de aminozuren His239, Trp241, Tyr266

en Asp268 in RgCrtC (en de overeenkomstige aminozuren in TrCrtC) te verduidelijken en

te identificeren als belangrijke residuen die direct betrokken zijn bij de katalytische reactie.

Door het analyseren van een gedeelte van de 3D-structuur, die verkregen is door homologie

modellering met het verwante AttH eiwit van Nitrosomonas europaea, kon worden

aangetoond dat alle geïdentificeerde aminozuren zich in directe omgeving van elkaar


147

bevinden. Al deze resultaten zijn een eerste aanwijzing dat deze aminozuren betrokken zijn

bij de katalytische cyclus.

Voor onderzoek naar ‘op maat gemaakte biokatalysatoren’ is de beschikbaarheid van een

‘high-throughput screening’ of selectie methode een eerste vereiste. Het bestaan van een

dergelijke methode zal ook de keuze van de ‘protein engineering’ methode bepalen. Een

high-throughput screening test is ontwikkeld voor de detectie van alcoholen, de producten

van hydraterende enzymen zoals CrtC en OHase (Hoofdstuk 4). OHase van

Elizabethkingia meningoseptica werd als model enzym gebruikt om het vermogen van de

ontwikkelde methode voor het mogelijk maken van een geautomatiseerde opstelling te

onderzoeken en te karakteriseren. De test bleek in staat om primaire, secundaire en tertiaire

alcoholen te detecteren in de aanwezigheid van de start-verbinding: onverzadigde vetzuren

en kleine cyclische en niet-cyclische onverzadigde alkenen als substraten.

Naast ‘protein engineering’ technieken, die worden gebruikt om operationele prestaties van

enzymen (bijvoorbeeld thermostabiliteit, activiteit en oplosmiddel tolerantie) te verbeteren,

wordt ook immobilisatie toegepast voor stabilisatie van enzymen. In Hoofdstuk 5

beschrijven we de immobilisatie van OHase als verknoopte enzym-aggregaten (in het

Engels cross-linked enzyme aggregates (CLEA)). Hiervoor wordt het recombinante OHase

uit E. coli celvrije extracten geaggregeerd en verknoopt met een bi-functionele crosslinker

glutaaraldehyde. Met een activiteitsbehoud van 26% na 21 uur verknoping bij 4°C, leidde

de CLEA bereiding van OHase tot een 2,4-voudige toename van biokatalysator stabiliteit

bij verhoogde temperaturen en een betere opslagstabiliteit bij lage temperaturen. Bovendien

werden tot 55-voudig hogere ruimte-tijd rendement (in het Engels space-time yield) bereikt

met OHase CLEA’s ten opzichte van microbiële productie.

Samenvattend heeft het werk dat in dit proefschrift is uitgevoerd, bijgedragen aan een

begrip van de structuur-functie relatie van twee pas ontdekte hydratases: carotenoïd 1,2-

hydratase en oleaat hydratase. Bovendien kan het werk bijdragen tot de ontwikkeling van

een biokatalysator die kan worden gebruikt voor de productie van stoffen met hoge

toegevoegde waarde in industriële processen.

Acknowledgements

149

Acknowledgements The completion of this thesis has been a long journey, but I did it! At this point I would like

to thank all the important people who have contributed to this thesis in one way or another.

First, I am grateful to my promotor Prof. Isabel Arends for her guidance and encouragement

throughout these years. You are truly an inspiration for me.

Then, my deepest gratitude goes to my daily supervisor Dr. Linda Otten. Linda, your

absolutely supportive, positive attitude towards all aspects of my research was a great help.

Certainly, we had to find our way in the beginning as I was your first PhD student, but we

succeeded in finding a good way that worked for us (for sure, the fact that we were sitting

in the same office helped a lot). It was your support and encouragement, which helped me

to also manage all the difficult phases of my PhD time. You have been through a very

difficult phase yourself, but even then, you managed to always find time when I needed

advice, inspiration or critical comments. I am very glad that I have met you and I will

always be grateful for everything that you have done for me. You are a great person.

I would also like to thank Martin Gorseling for all the technical support. Martin, I am glad

that you have found the way to our group. You have made a very big, positive change in

our group and I am for sure not the only one who appreciates everything that you have

done. No matter what technical problem I had, you managed always to solve it very quickly.

Your knowledge about all apparatus is amazing and I learned a lot from you.

Now, I would like to take this opportunity to thank Mieke van der Kooij. Thank you for all

your help in administrative matters and beyond. It is amazing how you keep track on

everything and never forget to send a reminder if something is about to expire. Although,

we did not have many opportunities to talk, I always enjoyed our few short conversations

in your office.

Special thanks go to my paranymph Rosario Franco Berriel. You are an amazing person

and working with you was a lot of fun. I always enjoyed our lunches together and our

conversations about ‘Gott und die Welt’. Thank you for you friendship and I wish you and

your cute family the best for your future.

I extend my gratitude to the ‘other’ Rosario, Rosario Medici. I am amazed about your

knowledge and professionalism. Working hard and focused, helping each other and

performing experiments carefully and precisely, these all comes to my mind when I think

about you. I appreciate all the work that you have done in order to make the HTS-assay

Acknowledgements

150

publishable. I truly enjoyed working together with you and being friend with you. All the

best for your professional carrier and your lovely family.

Then, my colleagues and officemates from ENZ and BOC, whose longer or shorter

presence enriched the life at work. Thank you all for the nice time during my stay at the

TU-Delft and all the support. I was privileged to work in two groups and to learn from all

of you.

All the supports from my family and my family in-low are highly appreciated. I am indebted

to them.

Finally, my greatest thanks goes to my friend, my soul mate, my beloved husband Senad.

Thank you for all your love, support and understanding.

Curriculum vitae

151

Curriculum vitae Aida Hiseni was born on May 20, 1980, in Doboj (Bosnia and Hercegovina). She pursued

studies in Biology at the Heinrich-Heine-Universität, Düsseldorf, Germany, where she

received the Diplom degree in Biology in 2007. In the same year she moved to the

Netherlands and commenced her Ph.D. work in Biotechnology at the Delft University of

Technology. Since November 2011, Aida works as associate scientist at DSM in Delft.

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