Research Collection
Doctoral Thesis
Directed evolution of 2-hydroxybiphenyl 3-monooxygenase from Pseudomonas azelaica HBP1
Author(s): Meyer, Andreas Jörg
Publication Date: 2002
Permanent Link: https://doi.org/10.3929/ethz-a-004455847
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ETH Library Diss. ETH No. 14735
DIRECTED EVOLUTION OF
2-HYDROXYBIPHENYL 3-MONOOXYGENASE FROM
PSEUDOMONAS AZELAICA HBP1
A dissertation submitted to the
SWISS FEDERAL INSTITUTE OF TECHNOLOGY ZURICH for the degree of DOCTOR OF NATURAL SCIENCES
Presented by
ANDREAS JÖRG MEYER Dipl. Nat. Sci., Swiss Federal Institute of Technology Zurich, Switzerland born July 23, 1971 citizen of Frauenfeld (TG), Switzerland
Accepted on the recommendation of
Prof. Dr. B. Witholt, examiner Prof. Dr. D. Hilvert, co-examiner Dr. H.-P. E. Kohler, co-examiner Dr. A. Schmid, co-examiner
2002
THEY SAY THAT DEATH KILLS YOU, BUT DEATH DOESN’T KILL YOU.
BOREDOM AND INDIFFERENCE KILL YOU.
I NEED MORE, IGGY POP
ACKNOWLEDGEMENTS
It is entirely impossible to write a dissertation without the help of many other people. A big “THANK YOU” to all these people, especially the IBT family!
Like every family, also the IBT family has a Dad: Bernard, my `Doktorvater`, I am deeply grateful that you gave me the opportunity to carry out my PhD thesis at your institute. I loved the great atmosphere at the IBT and I especially enjoyed the academic freedom you gave me. The person who initially appointed me to this project was Hanspi; thank you for the confidence you put on me and for always being a very deliberate adviser, not only concerning scientific questions. I also owe my acknowledgements to Andreas Schmid for his willingness to help wherever he could. Furthermore I would like to thank Prof. Dr. D. Hilvert for acting as a co-examiner and for critical reading of the manuscript.
Special thanks to all the people who gave their ideas, patience, support, and encouragement to make this project possible. Thanks to Andrew, Biggi, Jan, Marcel, Martin, Qun, Sven, Vali, and not to forget the Diploma students Dani, Michi, and Sandra who contributed a lot to this thesis and with whom I had a very joyful time in and outside the lab. I am also indebted to all the people who are not members of the IBT and who helped me during my work in a very uncomplicated way. Thanks to Willem, Adrie, and Robert from the Biochemistry Department in Wageningen, and to Prof. Dr. T. J. Richmond, Dave, and Eric from the Biophysics Institute of the ETH Zurich.
I am also very grateful to Helena who rendered every assistance in filling in all kind of forms and masterminded all administrative stuff in an impressive way. Thanks to Petra and Rahel for perfectly managing my sometimes unusual orders, to Monica and Helen for providing clean glassware and coffee cups, to Colette for excellent IT support, and to Peter Koller for repairing everything that needed to be fixed. During my thesis I had the pleasure to share the lab with some extraordinary persons with unique personalities. Thank you Martin for all the serious discussions about science, the meaning of life, books, music, movies, billiard, card-games, German beer, etc. and your company in exercising these topics. Special thanks also to Jochen who was always extremely helpful and took care of a reasonable sound level of our lab-radio. Thanks to Zhi and Poete who were very enjoyable lab-mates, unfortunately just for a short time.
Daily life at the IBT was fun. Sometimes it was more fun and sometimes less fun. Thanks to the people who have contributed to the great atmosphere at the IBT, in particular to Andreas B., Barbara, Bruno, Frank, Karin, Kuno, Martina, Peter R., René, and Winni. Thank you for being extremely agreeable colleagues at work and for being very enjoyable fellows during spare time. Furthermore, special thanks to my friends who were always ready to accompany me when a beer had to be drunken and to all those I did not mentioned here but who also contributed to this project.
Many thanks to my dear parents and my sister for their unlimited support in everything I did, regardless of their opinion whether it made sense or not. Last but not least sincere thanks are given to the person who brings so much happiness into my life, for her love and her company in good times and bad.
TABLE OF CONTENTS
TABLE OF CONTENTS
SUMMARY 3
ZUSAMMENFASSUNG 5
I. INTRODUCTION 7 GENERAL INTRODUCTION 8 ENZYMATIC HYDROXYLATIONS 9 PROTEIN ENGINEERING 16 ACTIVATION OF MOLECULAR OXYGEN BY ENZYMES 22 FLAVOPROTEINS 26 2-HYDROXYBIPHENYL 3-MONOOXYGENASE FROM PSEUDOMONAS AZELAICA HBP1 30 CATECHOLS 34 GOAL OF THIS RESEARCH 40 SCOPE OF THIS THESIS 40
II. CHANGING THE SUBSTRATE REACTIVITY OF 2-HYDROXYBIPHENYL 3-MONOOXYGENASE FROM PSEUDOMONAS AZELAICA HBP1 BY DIRECTED EVOLUTION 43
III. SYNTHESIS OF 3-TERT-BUTYLCATECHOL BY AN ENGINEERED MONOOXYGENASE 69
IV. HYDROXYLATION OF INDOLE BY LABORATORY EVOLVED 2-HYDROXYBIPHENYL 3-MONOOXYGENASE 83
V. ASPARTATE 222 IN 2-HYDROXYBIPHENYL 3-MONOOXYGENASE: A NEGATIVE CHARGE ESSENTIAL FOR EFFICIENT CATALYSIS 105
VI. CRYSTALLIZATION AND PRELIMINARY X-RAY ANALYSIS OF NATIVE AND SELENOMETHIONINE 2-HYDROXYBIPHENYL 3-MONOOXYGENASE 121
VII. CONCLUDING REMARKS AND OUTLOOK 129
VIII. REFERENCES 133
CURRICULUM VITAE 163
1
SUMMARY
SUMMARY
A major goal in enzymatic organic synthesis is to develop protein catalysts with tailored activities and selectivities. Despite a significant amount of work directed towards the understanding of protein folding and structure-function relationships, there is still little predictive understanding of how the protein primary sequence translates into its catalytically active tertiary structure. Therefore rational enzyme design for specific applications is difficult. In contrast, random approaches such as directed enzyme evolution have been shown to provide a very powerful alternative to engineer proteins with desired properties. The substrate reactivity of 2-hydroxybiphenyl 3-monooxygenase (HbpA) from Pseudomonas azelaica HBP1 was changed by directed enzyme evolution. We used random mutagenesis to generate HbpA variants and screened for improved monooxygenase activity on various 2-substituted phenols by the formation of colors indicative of the autooxidation of the reaction products. Doing so, mutants with increased activities as well as improved coupling efficiency could be
isolated. One of these mutants, which we denoted HbpAT2, was used for the biocatalytic synthesis of 3-tert-butylcatechol. In a lab-scale whole cell biotransformation with a recombinant E. coli JM101 strain a productivity of 63 mg L-1 h-1 was obtained, which is only about six fold lower than the optimized synthesis of 3-phenylcatechol with the wild-type enzyme. As a side product in the engineering of HbpA for extended biocatalytic
applications, we obtained a variant (HbpAind) that promotes the hydroxylation of indole. This compound is not a substrate for the wild-type enzyme. Investigation
of the reaction products showed that HbpAind hydroxylates indole exclusively at the carbons of the pyrrole ring and that the subsequent condensation and autooxidation then leads to the formation of indigo and indirubin. The formation of indirubin is of particular interest because it has been shown to possess
antileukaemic characteristics. Since the catalytic properties of HbpAind differ significantly from that of the wild-type enzyme, this variant was characterized
3 SUMMARY
biochemically in more detail. These investigations revealed that the negative charge of Asp222 is essential for efficient catalysis in HbpA. When this residue was exchanged with an uncharged amino acid, uncoupling of NADH oxidation from product formation was substantially increased. Evidence suggests that Asp222 is critically involved in substrate activation in HbpA. Structural studies were initiated to further assess the sequence-structure-function relationships of HbpA. Native and selenomethionine HbpA crystals were grown and high-resolution diffraction data were collected. We expect that the elucidation of the three- dimensional structure will allow a better understanding of the effects of the amino acid substitutions in the HbpA-variants, thus enabling the rational design of new and better biocatalysts.
4 ZUSAMMENFASSUNG
ZUSAMMENFASSUNG
Ein Schwerpunkt in der enzymatischen organischen Synthese ist die Entwicklung von Proteinkatalysatoren mit bestimmten Aktivitäten und hoher Selektivität. Trotz vielfältiger Bemühungen die Zusammenhänge zwischen Struktur und Funktion sowie die Faltungsmechanismen von Proteinen zu verstehen sind die Erkenntnisse auf diesem Gebiet nach wie vor äusserst unzureichend. Deshalb ist die zielgerichtete Entwicklung von Enzymen für bestimmte Anwendungen sehr schwierig. Eine vielversprechende Alternative bieten Methoden, die auf zufälligen Veränderungen der Primärsequenz basieren, wie zum Beispiel Directed Enzyme Evolution. Aus einer grossen Anzahl von zufällig hergestellten Varianten wird der Biokatalysator mit den gewünschten Eigenschaften selektioniert. Das Substrat Spektrum der 2-Hydroxybiphenyl 3-Monooxygenase (HbpA) von Pseudomonas azelaica HBP1 wurde mit Hilfe von Directed Enzyme Evolution verändert. Wir verwendeten Zufallsmutagenese für die Herstellung von HbpA- Mutanten mit höheren Aktivitäten gegenüber verschiedenen 2-substituierten Phenolen. Die veränderten Proteine wurden anhand von Farbbildung, welche das Resultat der Autooxidation der Reaktionsprodukte war, identifiziert. Die Anwendung dieser Methode führte zu Proteinvarianten mit höherer Aktivität und verbesserter Effizienz gegenüber HbpA. Eine dieser Mutanten, welche wir
HbpAT2 benannten, wurde für die biokatalytische Synthese von 3-tert-Butylbrenzkatechin ausgewählt. In einem Experiment im Labormassstab wurde mit einem rekombinanten E. coli JM101 Stamm eine Produktivität von 63 mg L-1 h-1 erreicht. Verglichen mit einem optimierten Prozess mit wildtyp-HbpA zur Herstellung von 3-Phenylbrenzkatechin ist diese Produktivität nur etwa sechs mal kleiner. Ein Nebenprodukt der Directed Evolution Experimente für die biokatalytische
Anwendung von HbpA war eine HbpA-Variante (HbpAind), welche Indol hydroxyliert. Diese Reaktion wird vom wildtyp-Enzym nicht katalysiert.
5 ZUSAMMENFASSUNG
Untersuchungen der Reaktionsprodukte von HbpAind mit Indol zeigten, dass die Hydroxylierung ausschliesslich am Pyrrol-Ring des Substrates stattfindet und dass anschliessende Autooxidation zur Bildung von Indigo und Indirubin führt. Hierbei ist speziell das gebildete Indirubin interessant, da es gegen Leukämie Wirksamkeit gezeigt hat. Die deutlichen Unterschiede der katalytischen
Eigenschaften von HbpAind gegenüber denen des wildtyp-Proteins veranlasste eine genauere biochemische Charakterisierung dieses Enzyms. Demzufolge ist die negative Ladung von Asp222 für eine effiziente Katalyse unerlässlich. Der Austausch von Asp222 gegen einen ungeladenen Aminosäurerest resultierte in einer signifikanten Erhöhung der Entkopplung der NADH Oxidation von der Produktbildung. Die erhaltenen Resultate deuten darauf hin, dass Asp222 für die Aktivierung des Substrats in HbpA verantwortlich ist. Erste strukturelle Studien um die Sequenz-Struktur-Funktion-Beziehung von HbpA weiter zu untersuchen wurden begonnen. Native- und Selenomethionin-HbpA-Kristalle wurden hergestellt und hochauflösende Brechungsdaten gesammelt. Wir erwarten, dass die Auflösung der dreidimensionalen Struktur von HbpA die Folgen des Austausches der Aminosäuren in den HbpA-Varianten verdeutlicht und so erlaubt bessere Biokatalysatoren durch zielgerichtete Methoden zu erschaffen.
6 CHAPTER I
CHAPTER I:
INTRODUCTION
ANDREAS MEYER
7 INTRODUCTION
GENERAL INTRODUCTION
In the last century synthetic organic chemists have managed to synthesize a very large number of complex molecules. Today, two major challenges have come into focus: i) the preparation of complex, water-soluble molecules and ii) the development of environmentally sustainable synthetic processes that are also economically acceptable (Faber, 1997; Wong & Whitesides, 1994). The application of enzymes as catalysts in synthetic organic chemistry is one possibility for the resolution of these issues, since enzymes are highly selective and function under natural conditions. Enzymes are proteins and the catalysts of living systems. Their ability to perform absolutely chemo-, regio-, and stereoselective reactions allows the synthesis of complex molecules that are not easily accessible by other chemical routes. This potential is used industrially for the production of different chiral compounds with lipases (Berglund & Hult, 2000; Liese et al., 2000). Lipases are cofactor-independent enzymes and generally stable, which allows their in vitro use in immobilized form for extended time periods with simple reaction set-ups. On the other hand, oxygenases, whose prowess at performing complex chemistry with exceptional selectivity has captured the attention of bioorganic chemists, have seen limited practical applications because of their complexity, poor stabilities, and often low catalytic rates (Cirino & Arnold, 2002). As a consequence, special process designs are required to resolve these difficulties. One possibility to improve the economic efficiency of processes based on oxygenases is the use of protein engineering techniques, which have been useful in increasing catalytic activity, altering substrate reactivity, or enhancing enzyme stability. This introduction deals with the fundamentals of biohydroxylations by oxygenase enzymes, with respect not only to process development but also to the biochemistry of oxygen activating enzymes, with a focus on
8 CHAPTER I
monooxygenases and their applications, and special reference to the flavoprotein 2-hydroxybiphenyl 3-monooxygenase.
ENZYMATIC HYDROXYLATIONS
Hydroxylation reactions activate carbon atoms by the conversion of a carbon-hydrogen bond into a carbon-oxygen bond. Enzymes that perform hydroxylations, called hydroxylases, are widely distributed in all forms of life from single cell organisms to humans because these reactions are key parts of the oxidative metabolism of many organic compounds. In the last decade, the application of hydroxylases in biocatalytic organic synthesis has become a prominent research area. For example, enzymatic hydroxylations for the production of chiral synthons and pharmaceuticals has been studied intensively (Ahmed et al., 1999; Hemenway & Olivo, 1999; Holland, 2000; Li et al., 2001b; Li et al., 1999; Li et al., 2002; Lister et al., 1999). The synthetic value of hydroxylases in such reactions mainly lies in the generally higher enantioselectivity achieved compared to chemical counterparts. However, not only the enantioselectivity but also the regioselectivity of hydroxylating enzymes is extremely valuable for organic synthesis, because no byproducts are produced. The formation of byproducts directly reduces the reaction yield and usually makes product purification more difficult. As a consequence, the profitability as well as the environmental sustainability of a process decreases.
Considerations in applying hydroxylases Three major issues have to be addressed if hydroxylases are applied in biocatalytic organic synthesis: i) The dependency of most hydroxylases on NAD(P)H, ii) the oxygen-transfer rates in reactor systems, and iii) the effects of substrate and/or product on the biocatalyst.
NAD(P)H dependency of hydroxylases For catalytic activity most hydroxylases need an external electron donor, usually NADH or NADPH, which has to be recycled during the biocatalytic process. One possibility to do this is the in vivo application of whole cells as
9 INTRODUCTION
biocatalysts (Duetz et al., 2001b). In doing so, the cofactor is regenerated by the cell, which in turn must remain metabolically active. This may require appropriate process design if substrate and/or product exhibit bactericidal properties (Lye & Woodley, 1999). An alternative approach is the in vitro application of isolated enzymes or cell-free preparations thereof (Pasta et al., 1996; Schmid et al., 2001). In this case, the cofactor has to be regenerated either enzymatically or electrochemically (Hollmann et al., 2001; Kula & Wandrey, 1987). This approach demands the isolation and purification of the applied enzymes, which are significant cost-factors for such a process. In addition, many hydroxylases are multicomponent, membrane-bound proteins that show only limited in vitro activity and stability (Duetz et al., 2001b). Therefore hydroxylases are typically applied in in vivo rather than in in vitro processes.
Another feature of NAD(P)H-dependent hydroxylases is a relatively low kcat
-1 value in the range of 1 to 100 s compared to hydrolases, which often have kcat values exceeding 100’000 s-1 (Duetz et al., 2001b). This has been attributed to the comparatively slow electron transfer from NAD(P)H to the enzyme (Nesheim & Lipscomb, 1996). This suggestion is supported by the observation that mono- and dioxygenases which do not involve NAD(P)H in their catalytic mechanism in
general have a 10 to 100-fold higher kcat value compared to NAD(P)H-dependent oxygenases (Fujisawa & Hayaishi, 1968; Gorlatova et al., 1998; Harpel & Lipscomb, 1990). Due to this low activity, NAD(P)H-dependent oxygenases have a theoretical upper limit for specific activity in whole cell biotransformations of approximately 4500 U (g CDW)-1 (Duetz et al., 2001b).
Oxygen-transfer rate in reactor systems In in vivo processes, the relevant hydroxylases have to compete with the
enzymes of the host organism for oxygen. The Km value for molecular oxygen of most oxygenases lies in the range of 10-60 µM (Shaler & Klecka, 1986), and is significantly higher than that of the respiratory systems of microorganisms, which generally have values smaller than 1 µM (Longmuir, 1954). Hence, the
10 CHAPTER I
bacterial cell density has to be chosen such that the dissolved oxygen concentration in the bioreactor still allows the hydroxylation reaction. For example, in a large-scale stirred tank reactor (>10 m3) the oxygen-transfer rate is about 100 mmol L-1 h-1, which corresponds to 1500 U L-1. To allow efficient hydroxylation, the dissolved oxygen concentration should be higher than 100 µM. Assuming an endogenous respiration activity of 100 U (g CDW)-1 the optimal cell density is 10 g CDW L-1 for a biohydroxylation process with an oxygenase (Duetz et al., 2001b). In combination with the maximal activity of such a biocatalyst the resulting optimal productivity would be about 5 kg m-3 h-1 for a product with a molecular mass of 150 Da. Thus, when whole cell hydroxylations with biocatalysts exhibiting >100 U (g CDW)-1 are used, the process is limited by the bioreactor setup rather than the activity of the biocatalyst (Duetz et al., 2001b). Strategies to increase the oxygen-transfer rate into reactors may result from aeration of the reactor with oxygen-enriched air, an increased internal pressure, the use of a second dispersed organic phase, or the introduction of molecular oxygen by hydrogen peroxide in combination with catalase. For comparison, two- to five-fold higher oxygen transfer rates are obtained in two phase gas/liquid membrane reactors or in membrane-aerated biofilm reactors than in stirred tank reactors (Casey et al., 1999; Léonard et al., 1998). However, the elaborate equipment required is difficult to apply on larger scale. A major advantage of in vitro applications of enzymes is the generally higher volumetric productivity compared to whole cell biocatalysts, because the processes are run at higher enzyme concentrations (Faber, 1997). When using this in vitro approach with hydroxylases, this advantage cannot be fully exploited. As in the case of in vivo processes, the oxygen transfer rate may be the limiting factor for biohydroxylation. Nevertheless, all the oxygen transferred to the system can be used for hydroxylation and is not consumed by the respiration system, which uses about two thirds of the oxygen in whole cell biotransformations (Duetz et al., 2001b).
11 INTRODUCTION
Effects of substrate and/or product Substrate and/or product at concentrations in the micro- or millimolar range inhibit many oxygenases. On the other hand, high product titers are required for efficient down-stream processing and high volumetric productivities. One way to handle this problem is to apply a controlled substrate feed and continuously remove the product in situ (Lye & Woodley, 1999). This is achieved by adsorption of the product on a solid matrix or through the use of a second organic phase in which the product but not the biocatalyst is extracted (Held et al., 1999; Schmid et al., 1998b). When using whole cell biocatalysts for in vivo biohydroxylations, substrate and/or product can be bactericidal. Here, as well, in situ product removal techniques can be applied. In addition, the use of host strains with increased tolerance towards the product can be considered. For example, when the product has a disruptive effect on the membrane, strains with efflux systems or cis-trans isomerases may yield a solution (Kieboom et al., 1998; Pedrotta & Witholt, 1999).
In vitro vs. in vivo application of hydroxylases The question whether enzymes should be used in isolated form (in vitro) or within a host cell (in vivo) continues to be discussed. In general, in vitro application is feasible when four major prerequisites are met (Kula & Wandrey, 1987): i) The number of reaction steps is limited, ii) the enzyme involved is highly stable, iii) a system for overproduction and isolation of the enzyme is available, and iv) the reaction is cofactor-independent or cofactor-regeneration can be integrated into the process. If these requirements are fulfilled, advantages, such as for example high volumetric productivity due to high enzyme concentration, the ability to handle bactericidal substrates and products, and individual optimization of enzyme and product synthesis, can be realized (Faber, 1997). In vitro applications of cofactor-independent hydrolases but also cofactor-dependent dehydrogenases have been successfully implemented in industry.
12 CHAPTER I
However, it is difficult to extrapolate the model built for hydrolases and dehydrogenases to the in vitro use of oxygenases. The main difference is that oxygenases show only limited stability in vitro and are frequently multi- component enzymes, which require the proper assembly of different proteins or subunits (Staijen et al., 2000). In addition, advantages of in vitro biotransformations, for example high enzyme concentrations, are useless due to the limited oxygen transfer rate into reactors. Therefore, biocatalytic routes that employ oxygenases are performed in vivo rather than in vitro. The main pros and cons concerning in vitro versus in vivo applications of hydroxylases are summarized in Table 1.
Table 1: Pros and cons for the in vitro and in vivo application of hydroxylases.
In vitro application In vivo application
Advantages Advantages • All oxygen is used for hydroxylation • Cofactor is regenerated by the cell • Simplified down-stream processinga • Hydroxylases are usually more stable in • Use of bactericidal compounds is possibleb vivoc • Side reactions are minimized • Membrane bound hydroxylases can be used • No substrate for cell maintenance needed • Application of cascade of enzymes possible
Disadvantages Disadvantages • Cofactor-regeneration system required • Approximately 2/3 of oxygen is used for • Limited stability of hydroxylase in vitroc endogenous respirationc • Enzymes have to be isolated • Sensitivity of cells towards bactericidal • Many hydroxylases are multi-component compoundsb enzymes: difficult to handle and tuned • Product separation from culture may be • Many hydroxylases are membrane bound laboriousa enzymes: low activity without membranec • Mass transfer limitation for diffusion of compounds through bacterial membrane
a (Schmidt et al., 1986), b (Kragl et al., 1993), c (Duetz et al., 2001b), d (Vasic-Racki et al., 1989)
13 INTRODUCTION
Examples of in vivo biohydroxylations The synthesis of several industrially relevant compounds by regio- and/or stereoselective biohydroxylation has been addressed (Table 2). The productivities are of the expected magnitude. The only processes that exceed the theoretical value of 5 g L-1 h-1 are those for the production of 6-hydroxynicotinic acid and toluene cis-glycol. This can be explained by the fact that in the first process the introduced oxygen derives from water and not from molecular oxygen. Thus, the low concentration of molecular oxygen in the bioreactor has no effect on the biotransformation. In the latter synthesis, a second dispersed organic phase was used, which results in an increased oxygen transfer rate into the reactor. The principle of not using molecular oxygen as co-substrate can be used as a general strategy to increase the productivities of biohydroxylations. In addition to enzymes that use water as an oxygen donor, peroxidases, which incorporate oxygen from hydrogen peroxide (van Deurzen et al., 1997), are also of special interest for biohydroxylations.
14 CHAPTER I
Table 2: Selected examples of in vivo biohydroxylations.
Microorganism Reference Substrate Product Productivity Remarks
Alcaligens (Glöckler & Roduit, 1996) Fed batch process in CSTRa, faecalis productivity >98 g L-1 in 42 h, N COOH HO N COOH 2.3 g L-1 h-1 inserted oxygen is obtained from a water molecule
(Glöckler & Roduit, 1996) COOH COOH Alcaligens Fed-batch process in CSTR, xylosoxydans productivity >100 g L-1 in 12 h, N HO N -1 -1 inserted oxygen is obtained from 8.3 g L h a water molecule
(Lilly & Woodley, 1996) F F Pseudomonas Resting cells, scale 2.5 L, in situ OH putida ML2 product removal, activity -1 -1 -1 23 U (g CDW) , OH 1.1 g L h productivity 15 g L-1 in 14 h
Ph Ph (Held et al., 1998) Recombinant Fed-batch mode in CSTR, scale OH OH E. coli 2 L, in situ product removal, -1 -1 -1 activity 5.4 U (g CDW) OH 0.4 g L h
(Lilly & Woodley, 1996) H OH Pseudomonas Two liquid phase process in putida UV4 CSTR, scale 1.7 L, activity -1 -1 -1 120 U (g CDW) , OH 9.5 g L h -1 H productivity 57 g L in 6 h
(Lee et al., 1996) COOH COOH Candida Fed-batch cultivation, scale 2 L, H H rugosa productivity 100 g L-1 in 120 h OH 0.8 g L-1 h-1
OH (Duetz et al., 2001a) Rhodococcus Resting cells, scale 5 mL, activity opacus PWD4 15 U (g CDW)-1 n.d.b
a CSTR, continuously stirred tank reactor, b not determined
15 INTRODUCTION
PROTEIN ENGINEERING
The efficient application of biocatalysts requires enzymes with high activity and stability under process conditions, desired substrate specificity, and high enantio- and regioselectivity. In contrast, in nature proteins evolve in response to different requirements, such as low substrate concentration, the prevention of accumulation of toxic intermediates, or a broad substrate range. The optimization of enzymes to the conditions in industrial processes is therefore of major interest (Bornscheuer & Pohl, 2001). Two rather different approaches can be used to create these tailor-made biocatalysts: rational and random protein design.
Rational protein design For rational protein design the availability of a three-dimensional structure of the enzyme or a homologue is a minimal prerequisite. In addition, knowledge about the sequence-structure-function relationship is usually also required. Thus, much information must be acquired to enable the rational design of a new biocatalyst. It is noteworthy that the techniques to accelerate rational protein engineering have undergone immense progress. For enzymes smaller than 30 kDa, structure determination can be performed by NMR spectroscopy instead of X-ray diffraction, which makes the time-consuming preparation of suitable protein crystals obsolete (Wüthrich, 1995). In addition, the greatly increasing number of publicly accessible structures and sequences simplifies the investigation of sequence-structure-function relationships. Molecular modeling then makes it possible to predict how to increase enzyme selectivity or activity based on the structure of a related enzyme. Using rational protein engineering the stereospecificity of vanillyl-alcohol oxidase from Penicillium simplicissimum was inverted (de Jong et al., 1992; van den Heuvel et al., 2000a). While the wild type enzyme hydroxylated 4-ethylphenol stereospecifically to (R)-1-(4’-hydroxyphenyl)ethanol with an enantiomeric excess of 94%, a rationally designed mutant had a strong preference for the (S)-enantiomer (e.e. 80%). The substrate specificity of
16 CHAPTER I
cytochrome P450 BM-3 from Bacillus megaterium was also changed by a rational approach (Boddupalli et al., 1990). P450 BM-3 hydroxylates long chain fatty acids and their amide or alcohol analogues (Capdevila et al., 1996). The wild type protein does not hydroxylate fatty acids with a chain length shorter than 12 carbons or 10 carbons for the analogues. By changing active site residues of P450 BM-3, a variant was obtained which accepts ω-p-nitrophenoxyoctanoic acid, a substrate that is not hydroxylated by the wild- type enzyme (Li et al., 2001a).
Random protein engineering Random protein design, also called directed or molecular evolution (Arnold, 1998a; Arnold, 1998b; Arnold & Volkov, 1999; Bornscheuer, 1998; Farinas et al., 2001; Kuchner & Arnold, 1997; MacBeath et al., 1998; Schmidt-Dannert & Arnold, 1998; Sutherland, 2000), is based on random mutagenesis of the gene encoding the protein of interest followed by screening or selection (Scheme 1). Diversification of the gene of interest can either be done by error-prone PCR, recombination of gene fragments, saturation mutagenesis, or by using mutator strains (Beckmann et al., 1985; Bornscheuer et al., 1999; Leung et al., 1989; Stemmer, 1994; Zhao et al., 1998). Subsequently, variants with the desired properties have to be selected from a large number of mutants, which is often the limiting step in directed evolution experiments (Zhao & Arnold, 1997). However, with an appropriate selection or screening procedure a broad range of enzyme properties can be altered, even though these are not required in nature (Taylor et al., 2001). Sequential cycles of random mutagenesis and screening of a para-nitrobenzyl esterase from Bacillus subtilis lead to enzymes with a 60 to 100-fold increased activity in aqueous-organic solvents (Moore & Arnold, 1996; Moore et al., 1997). These proteins were tailored to catalyze the deprotection of cephalosporin- derived antibiotics during chemical synthesis (Brannon et al., 1976). Due to the low water-solubility of the substrate the increased activity in this unnatural
17 INTRODUCTION
environment was required to compete with the conventional deprotection by zinc, which can be performed in organic solvents.
gene of interest
digestion
error prone PCR random mutagenesis
recombination
pool of mutant genes
introduction into host
mutant library
screening/selection
genes of improved clones
selection of most selection of several improved gene from further amplification rounds improved genes from selected clones selected clones
protein with desired properties
Scheme 1: Principle of a directed evolution experiment. The principle of the two most used methods, error-prone PCR (left) and DNA shuffling (right), are shown.
Enzymes for organic synthesis have also been changed, e.g. a biocatalyst for the enantioselective hydrolysis of racemic p-nitrophenyl 2-methyldecanoate. The (S)-configured free acid belongs to a class of chiral compounds which are useful as chiral building blocks (Sheldon, 1993). For this reaction the enantioselectivity of a lipase from Pseudomonas aeruginosa PAO1 was increased in only 4
18 CHAPTER I
generations from 2% e.e. to 81% e.e. yielding the (S)-isomer (Reetz et al., 1997). Table 3 shows a selection of enzymes that were successfully modified by directed evolution. The properties listed are the major altered characteristics of the most improved enzyme, usually from several obtained variants.
19 iyrxbpey ixgns -odAt3clrctco,atrdsbtaeseiiiyePR(Riegert 20 epPCR 5-fold Act 3-chlorocatechol, altered substrate specificity Dihydroxybiphenyl dioxygenase yohoeP5 M3Aiiyt yrxlt noeSatMut Ability to hydroxylate indole Cytochrome P450 BM-3 oun ixgns .-odAtpcln,12fl c ntleeePR&Stu (Sakamoto epPCR & SatMut 5.6-fold Act picoline, 1.2-fold on toluene Toluene dioxygenase ihnldoyeaeAttwrsmncci rmtchdoabn Ns (Suenaga DNAsh Act towards monocyclic aromatic hydrocarbons Biphenyl dioxygenase yohoeP5 M35fl c caeePR(Farinas epPCR (Delagrave epPCR (Li SatMut 5-fold Act octane Up to 5-fold Act towards different carboxylic acids 16-fold Act methylgalactose Cytochrome P450 BM-3 Cytochrome P450 BM-3 Galactose oxidase ihnldoyeae18fl c ihnlad44-ihooihnlDAh(Kumamaru (Brühlman & Chen, 1999) epPCR & DNAsh DNAsh Reference 1.8-fold Act biphenyl and 4,4’-dichlorobiphenyl KDPG Act towards polychlorinated biphenyls Lyases Method Biphenyl dioxygenase Biphenyl dioxygenase DNAsh Active in the absence of fructose-1,6-bisphosphate (effector) Altered properties 7.5-fold Act Lactate dehydrogenase Isopropylmalate dehydrogenase Oxidoreductases Enzyme Table 3: Selected examples of successful directed evolution experiments. Toluene aats xds Higher protein level in Galactose oxidase hteedstrs rdc pcfct epPCR & SdM Functional expression in Product specificity Horseradish peroxidase Phytoene desaturase
i adls 2-fold Act -aldolase ortho mnoyeae6fl c ahhln,18fl c rclrehln Ns (Canada DNAsh 6-fold Act naphthalene, 1.8-fold trichloroethylene -monooxygenase D a 2kt--exguoaeePR&DAh(Fong epPCR & DNAsh -2-keto-3-deoxygluconate
a 0 epPCR at 40°
E. coli E. coli
, 2-fold increased
k cat
/
K M ePR&Rc(Sun epPCR & Rec epPCR & Rec f c b e d
(Li (Lin (Wang & Liao, 2001) (Allen & Holbrook, 2000) (Suzuki
et al. et al.
et al.
et al.
et al.
et al.
et al. , 2001a) , 2000) et al. et al.
, 1999) et al.
, 2001)
et al. et al.
, 2000) et al. , 2001) , 2001) , 2001b) , 2002) , 2001) , 2001) , 2001) , 1998)
Staphylococcus aureus P. aeruginosa P. fluorescens yatiae5fl c n neso featoeetvt pC aMt(May epPCR & SatMut 5-fold Act and inversion of enantioselectivity p Hydantoinase α hmdn iaeIncreased Thymidine kinase Aspartate aminotransferase D-lcs yohshrls pt 0fl c ifrn usrts lee fetrseiiiyDAh(Salamone a Reference DNAsh Up to 30-fold Act different substrates, altered effector specificity Method ADP-glucose pyrophosphorylase Transferases GSSM Altered properties 30’000-fold half-life time at 55°C Haloalkane dehalogenase Hydrolases Enzyme utlsnEAti M pC (Chen & Arnold, 1993) epPCR 200-fold half-life time at 65°C, T Act in DMF Subtilisin E Subtilisin E Aspartate aminotransferase β β β saturation mutagenesis;
Ntoezletrs 2fl c n3%DFePR&Rc(Moore & Arnold, 1996) epPCR & Rec 32-fold Act in 30% DMF -Nitrobenzyl esterase -Galactosidase -Lactamase -Lactamase -Lytic protease Act, increased activity; iaeIcesdeatoeetvt rm2 ..t 1 ..ePR(Reetz epPCR Increased enantioselectivity from 2% e.e. to 81% lipase seae2fl nrae nnislciiyEpPCR & MuStr 2-fold increased enantioselectivity esterase iae1.-odAtpopoiisePR&DAh(van Kampen & Egmond, 2000) epPCR & DNAsh 11.6-fold Act phospholipids lipase g GSSM, gene site saturation mutagenesis; b
epPCR, error-prone PCR; c n rmr pcfcte aMt(Graham SatMut Act and primary specificities 2.1 10 pt 6fl c oad ifrn uoyaoie Ns (Zhang DNAsh Up to 66-fold Act towards different fucopyranosides 6fl c lvlnt aMt(Oliphant & Struhl, 1989) (Stemmer, 1994) SatMut DNAsh 16-fold Act clavulanate 32’000-fold Act cefotaxime 5 -fold increased × 10 6
-fold increased
k cat
/
K M frzdvdn Ns (Christians DNAsh for zidovudine
c k
DNAsh, DNA shuffling; cat
/
K
M k
for cat
/ K
β M -branched amino- and oxo-acids frvln SdM & DNAsh for valine h opt MuStr, mutator strain; 1° nrae pC e (Zhao & Arnold, 1999) epPCR & Rec 17°C increased d
SdM, site directed mutagenesis i KDPG, D-2-keto-3-deoxy-6-phosphogluconate. Ns (Yano DNAsh g &Rc(Gray & Rec e
Rec, recombination; h
(Oue (Henke & Bornscheuer, 1999)
et al. et al.
et al.
et al.
et al.
et al. et al.
, 2000) , 1999) et al. et al. , 2001) , 1998) , 1997) , 1997) , 1993) , 1999) f , 2002)
SatMut, 21 INTRODUCTION
ACTIVATION OF MOLECULAR OXYGEN BY ENZYMES
Respiration is one of the most fundamental activities of life. In a bacterial cell molecular oxygen has mainly two functions: i) Molecular oxygen serves as ultimate hydrogen acceptor in the process of biological oxidation, being reduced to water or hydrogen peroxide, and ii) molecular oxygen is used to transform carbohydrates into cellular constituents and other biologically important substances. Because molecular oxygen is kinetically inert it has to be activated by enzymes. Enzymes involved in the first process are referred to as oxidases, whereas those involved in the latter process are referred to as mono- or dioxygenases (Nozaki, 1978).
Oxidases Oxidases transfer electrons from an organic substrate to molecular oxygen,
thereby forming H2O2 or H2O.
In oxidases that reduce molecular oxygen to H2O2, metal ions are not directly involved in catalysis. Many of these oxidases are simple flavoproteins, in which the organic substrate is oxidized by reacting with the flavin. The reduced flavin then combines with molecular oxygen, forming a C(4a)-flavoperoxide, which is
split heterolytically with the formation of H2O2. Examples of this class of oxidases are e.g. glucose oxidase or lactate oxidase (Su & Klinman, 1999; Yorita et al., 2000). Other oxidases of this class contain metal ions, but in these
enzymes the metals are not directly responsible for activation of O2. The role of the metal ions is rather the activation of the organic substrate, which thereafter
forms H2O2 by direct reaction with molecular oxygen (Ito et al., 1991).
In contrast, in oxidases that reduce O2 to two molecules of H2O, metal ions play an important role in utilizing the entire oxidizing capacity of molecular oxygen. These enzymes have bimetallic reduction sites, which readily react with
O2 after having received two electrons. The peroxide intermediate formed remains bound to the bimetallic centers and is further reduced in two
22 CHAPTER I
successive one-electron reactions. The most prominent member of this oxidase class is cytochrome c oxidase, which is responsible for more than 90% of the
O2 consumption by life on this planet (Slater et al., 1965). The biotechnological application of oxidases is rather versatile. In research glucose oxidase is used to maintain anaerobic conditions during investigation of the redox properties of enzymes (Suske et al., 1999). Xanthine oxidase is applied for the determination of redox potentials of prosthetic groups in proteins (Massey, 1991). But oxidases may also have synthetic value. For example, vanillyl alcohol oxidase was used for the enantioselective hydroxylation of 4-substituted alkylphenols (Drijfhout et al., 1998).
Dioxygenases
Dioxygenases catalyze the incorporation of both oxygen atoms from O2 into a substrate. In general, one single substrate molecule acts as oxygen acceptor (intramolecular dioxygenases). Some intramolecular dioxygenases, like the multicomponent Rieske dioxygenases (Coulter & Ballou, 1999), require one or two-electron transfer proteins to shuttle electrons from NADH to the site of oxygenation. However, many enzymes of this class do not need any organic cofactor or additional proteins for activity. The only requirement is the presence of a metallic prosthetic group. This is also true for the most prominent intramolecular dioxygenases, the catechol-cleaving dioxygenases (Broderick, 1999). In some dioxygenase reactions, one atom each of molecular oxygen is introduced into two different substrate molecules (intermolecular dioxygenases) (Lindstedt & Lindstedt, 1970). Intermolecular dioxygenases, such as iron- dependent clavaminate synthase (Salowe et al., 1990), deacetoxycephalosporin C synthase (Lee et al., 2001), and thymine hydroxylase (Thornburg & Stubbe, 1993), generally require α-ketoglutarate as co-substrate, which is converted to succinate by ascorbate-dependent oxidative decarboxylation.
23 INTRODUCTION
Monooxygenases
Monooxygenases incorporate one oxygen atom from O2 into the organic substrate, whereas the other is reduced to water. The required electrons are either donated from the substrate itself (internal monooxygenases), or from an external electron donor (external monooxygenases), mostly NAD(P)H. According to their prosthetic group, they can be divided into several different classes: Heme, non-heme iron, quinone-forming, and flavin monooxygenases.
Heme monooxygenases In heme monooxygenases the catalytic iron is bound to a protoporphyrin IX tetrapyrrole system. Here, four nitrogen atoms coordinate the metal, one from each of the pyrrole rings. Atoms from the protein environment, water, or during catalysis, oxygen, occupy the two remaining coordination sites. The P450 enzymes, also known as cytochromes P450, constitute a large superfamily of heme-thiolate proteins and are widely distributed in bacteria, fungi, plants, and animals (Nelson et al., 1996). Although much work has been done to resolve the catalytic mechanism (Bell-Parikh & Guengerich, 1999; Modi et al., 1996; Poulos, 1995; Vidakovic et al., 1998), the way in which P450 activates oxygen is not known in detail. It is clear that substrate binding changes the redox properties of the enzyme and the first electron is subsequently transferred (Sligar & Gunsalus, 1976). Then oxygen binds to the ferrous P450 and the second electron is ceded (Brewer & Peterson, 1988). There is strong evidence that proton transfer is the critical step in the final activation of molecular oxygen (Vidakovic et al., 1998). After the second electron transfer and oxygen bond cleavage, a highly reactive iron-oxo complex is believed to be formed that serves as the oxygenating species.
Non-heme monooxygenases Non-heme iron monooxygenases show a variety of coordination structures for metal binding and can catalyze monooxygenations as well as dioxygenations (Lange & Que, 1998). Among non-heme iron enzymes there are two different classes of monooxygenases: cytoplasmic (soluble) and membrane
24 CHAPTER I
bound (particulate) proteins (Patel et al., 1982; Staijen et al., 1997). Both enzyme types activate oxygen by a catalytic diiron center (Moënne-Loccoz et al., 1998; Shanklin et al., 1997), in contrast to the dioxygenases that in general have only one metal involved in catalysis. However, not all non-heme monooxygenases contain iron for oxygen activation. Dopamine-ß-mono- oxygenases that play an important role in the catecholamine biosynthesis contain two copper atoms per subunit, which are essential for catalysis (Stewart & Klinman, 1987). In addition, in these enzymes ascorbate serves as electron donor rather than NAD(P)H (Diliberto & Allen, 1980). Due to the great diversity among non-heme monooxygenases a general mechanism of oxygen activation by this class of proteins does not exist.
Quinone-forming monooxygenases Quinone-forming monooxygenases constitute a unique group of enzymes that do not require a cofactor for catalysis (Fetzner, 2002). In these proteins, molecular oxygen and/or substrate activation has to be performed exclusively by amino acid residues. The only two enzymes known from this family are the tetracenomycin F1 monooxygenase from Streptomyces glaucescens (Shen & Hutchinson, 1993) and the ActVA-orf6 protein from S. coelicolor A3 (Caballero et al., 1991). Evidence suggests that a conserved histidine is required for efficient catalysis, but this residue is not absolutely essential. So far, two possible mechanisms for oxygen activation by quinone-forming monooxygenases have been proposed (Fetzner, 2002): i) The formation of a protein radical intermediate, and ii) direct electron transfer from the (deprotonated) substrate to molecular oxygen. However, detailed characterization of the catalytic mechanism still remains to be done.
Flavoprotein monooxygenases Activation of molecular oxygen by flavoprotein monooxygenases is discussed in the next section.
25 INTRODUCTION
FLAVOPROTEINS
Flavoproteins are involved in a variety of biological processes. They participate in a wide range of reactions, such as dehydrogenation of metabolites, one- and two-electron transfer from and to redox centers, in light emission, and in the activation of oxygen for oxidation and hydroxylation reactions (Massey, 1995). Despite the diversity of their catalytic function, flavoproteins can be grouped into a relatively small number of classes (Table 4). Within a specific class the proteins share many common properties and clearly allow classification, which is based on the catalyzed reaction, the ability to use molecular oxygen as acceptor, and the nature of auxiliary redox centers (Fraaije & Mattevi, 2000; Massey, 1995).
26 CHAPTER I
Table 4: Classification of flavoproteins.
Enzyme class Examples Reference
Simple flavoproteins
Oxidases D-amino acid oxidase (Ronchi et al., 1982) Glucose oxidase (Gibson et al., 1964) Glycin oxidase (Job et al., 2002) Vallinyl-alcohol oxidase (de Jong et al., 1992)
Electron transferases Flavodoxin (Mayhew & Massey, 1969) Ferredoxin-NADP reductase (Karplus et al., 1990)
Monooxygenases Phenol hydroxylase (Neujahr & Gaal, 1973) p-hydroxybenzoate hydroxylase (Hosokawa & Stanier, 1966) Salicylate hydroxylase (White-Stevens & Kamin, 1972) Bacterial luciferase (Hastings & Presswood, 1978) Lactate 2-monooxygenase (Sullivan, 1968)
Flavoprotein-disulfide Lipoamide dehydrogenase (Williams Jr., 1992) oxidoreductases Glutathione reductase (Williams Jr., 1992)
Flavoproteins with auxiliary redox centers
Metal-containing Xanthine oxidase (Olson et al., 1974) flavoproteins Succinate dehydrogenase (Reddy & Weber, 1986) Naphthalene dioxygenase (Haigler & Gibson, 1990)
Flavocytochromes Cellobiose dehydrogenase (Li et al., 1996) Lactate dehydrogenase (Lederer, 1991)
(flavocytochrome b2)
27 INTRODUCTION
Flavoprotein monooxygenases Due to the spectral properties of the flavin prosthetic group, the catalytic mechanism of flavoprotein monooxygenases has been resolved in detail. Flavoprotein monooxygenases have three substrates: NAD(P)H to reduce the enzyme bound flavin, the substrate to be oxygenated, and molecular oxygen (Massey, 1994). Flavin reduction by NAD(P)H and reaction with molecular oxygen leads to the formation of a readily detectable flavin C(4a)-hydroperoxide (Palfey et al., 1994a; van Berkel & Müller, 1991). In the absence of substrate,
the hydroperoxide decays nonproductively to H2O2 and oxidized flavin. If substrate is present, it can react with the flavin hydroperoxide. The transient stabilization of this intermediate is characteristic for this class of flavoproteins and is not observed for dehydrogenases or oxidases with a flavin group. By splitting the peroxide O-O bond the terminal oxygen is inserted into the substrate. The remaining C(4a)-hydroxyflavin undergoes dehydration and the flavin returns to its oxidized state for the next catalytic cycle. Based on sequence alignments, flavoprotein monooxygenases are divided into three subclasses: i) Single component aromatic hydroxylases (Eppink et al., 1997), ii) two-component aromatic hydroxylases (Becker et al., 1997), and iii) ketone monooxygenases and N-hydroxylating enzymes (Stehr et al., 1998). Flavoprotein monooxygenases generally exhibit high selectivity. This property has been exploited for chiral organic synthesis by the application of ketone monooxygenases and for the regioselective hydroxylation by aromatic hydroxylases.
Ketone monooxygenases (Baeyer-Villiger monooxygenases) Ketone monooxygenases, more commonly called Baeyer-Villiger monooxygenases, exhibit the rare characteristic of being able to catalyze two mechanistically different types of biochemical reactions. On one hand, they can catalyze the nucleophilic oxygenation of ketones and aldehydes to lactones or esters. This proceeds in a manner analogous to the established peracid- catalyzed organic chemical reaction from which they take their name (Baeyer &
28 CHAPTER I
Villiger, 1899). On the other hand, Baeyer-Villiger monooxygenases can also catalyze the electrophilic oxygenation of various heteroatoms, as illustrated by their ability to form sulfoxides from organosulfides (Latham & Walsh, 1987). Because they generally perform these reactions highly selectively they are valuable biocatalysts for the synthesis of chiral compounds. The best-studied member of this enzyme family is the cyclohexanone monooxygenase (Ryerson et al., 1982; Sheng et al., 2001). This enzyme has been used for the asymmetrization of prochiral 4-methylcyclohexanone to the homochiral lactone product with an e.e. >98% (Taschner et al., 1992). An even higher enantioselectivity was observed when the enzyme performed the electrophilic oxygenation of a sulfur atom: Synthesis of (R)-methyl phenyl sulfoxide from the equivalent sulfide proceeded with an e.e. >99% (Carrea et al., 1992).
Aromatic hydroxylases Flavoprotein aromatic hydroxylases catalyze the oxygenation of aromatic rings, which are activated by a substituent such as a hydroxy or amino group (Massey, 1994). In contrast to Baeyer-Villiger monooxygenases, the flavin C(4a)-peroxide attacks the substrate exclusively by an electrophilic mechanism (Moran et al., 1997). The catalytic mechanism of flavoprotein aromatic hydroxylases has been studied in detail using p-hydroxybenzoate hydroxylase as model enzyme. Because the reactive species of flavin monooxygenases, the flavin C(4a)-peroxide, is unstable in protic solvents it has to be sequestered from water. In general, as has been shown for different proteins, this is often achieved by induced fit rearrangements of protein loops or domains (Bennett & Steitz, 1978; Joseph et al., 1990). In contrast, p-hydroxybenzoate hydroxylase, and probably all flavoprotein aromatic hydroxylases, have developed an alternative mechanism to address this issue. In these proteins the flavin group can adopt two conformations (Gatti et al., 1994; Schreuder et al., 1994). In the absence of substrate, the prosthetic group is in the so-called ‘in’ conformation
29 INTRODUCTION
and not accessible for reduction by NADPH. When substrate binding and ionization occurs, the flavin moves to the ‘out’ position where reduction takes place (Frederick et al., 2001; Moran et al., 1996; Palfey et al., 1999). Substrate hydroxylation proceeds in the active site of the protein protected from water. The synthetic value of flavoprotein aromatic hydroxylases lies in their absolute regioselectivity. This property has been used for the synthesis of different 3-substituted catechols by 2-hydroxybiphenyl 3-monooxygenase from Pseudomonas azelaica HBP1 (Held et al., 1998), which is discussed below.
2-HYDROXYBIPHENYL 3-MONOOXYGENASE FROM PSEUDOMONAS AZELAICA HBP1
2-Hydroxybiphenyl 2-Hydroxybiphenyl (o-phenylphenol) is abundant in the environment due to its extensive use as post-harvest fungicide to delay rotting and fouling of fruits (Eckert, 1977). In addition, it is a dead-end product of microbial desulfurization of petroleum and coal (Gallagher et al., 1993; Rhee et al., 1998). 2-Hydroxybiphenyl has deleterious effects on various prokaryotic and eukaryotic cells. For instance, it efficiently inhibits the essential NADH oxidase and NADH:cytochrome c reductase of the photosynthetically active bacterium Rhodospirillum rubrum, both of which are essential for the transformation of NADH to ATP (Maudinas et al., 1973; Oelze & Kamen, 1975). Investigations in vertebrates have shown that in the liver 2-hydroxybiphenyl is hydroxylated to 2,5-dihydroxybiphenyl by the microsomal cytochrome P450 system (Nakagawa & Moore, 1995; Tayama & Nakagawa, 1994). This reaction product is readily autooxidized to the corresponding quinone and semiquinone, which have cytotoxic and carcinogenic properties.
2-Hydroxybiphenyl degradation by Pseudomonas azelaica HBP1 The soil bacterium Pseudomonas azelaica HBP1 was isolated in 1988 from sewage sludge and is able to grow on 2-hydroxybiphenyl as sole source of carbon and energy (Kohler et al., 1988). In contrast to other microorganisms, it
30 CHAPTER I
tolerates 2-hydroxybiphenyl concentrations up to its water-solubility limit of 0.7 g L-1. The first enzyme of the degradation pathway in P. azelaica HBP1 is the 2-hydroxybiphenyl 3-monooxygenase (HbpA, EC 1.14.13.44) (Scheme 2) (Kohler et al., 1988).
123
O2 NADH + H+ NAD+
O OH OH COOH O2 H2O OH OH
H O2 O 2 COOH O OH NAD+ NADH + H+ OH COOH + COOH OH OH NAD+ NADH + H+ CO2 7 6 5 4
Scheme 2: Proposed pathway for 2-hydroxybiphenyl degradation by Pseudomonas azelaica HBP1. 1, 2-hydroxybiphenyl; 2, 2,3-dihydroxybiphenyl; 3, 2-hydroxy-6-phenyl-6-oxo- 2,4-hexa-dienoic acid; 4, 2-hydroxy-2,4-pentadienoic acid; 5, benzoic acid; 6, catechol; 7, 2-hydroxymuconic semialdehyde.
HbpA catalyzes the initial, highly regioselective ortho-hydroxylation of 2-hydroxybiphenyl to form 2,3-dihydroxybiphenyl, which is further converted to 2-hydroxy-6-phenyl-6-oxo-2,4-hexadienoic acid (HOPDA) by a meta ring cleavage dioxygenase (HbpC). HOPDA is then hydrolyzed to benzoate and 2-hydroxy-2,4-pentadienoic acid by HOPDA hydrolase (HbpD) (Kohler et al., 1988; Kohler et al., 1993a; Schmid, 1997). These two compounds are further metabolized via intermediates also formed in the analogous biphenyl degradation pathway (Catelani et al., 1973; Gibson et al., 1973).
31 INTRODUCTION
General characteristics of 2-hydroxybiphenyl 3-monooxygenase 2-Hydroxybiphenyl 3-monooxygenase is a homotetrameric flavoprotein with a molecular mass of 256 kDa (Suske et al., 1997). Each subunit contains a noncovalently bound FAD and the corresponding conserved sequence motif characteristic for flavin binding, according to which it was classified as flavoprotein aromatic hydroxylase (Eppink et al., 1997). HbpA needs three substrates for catalytic activity: NADH as electron donor, molecular oxygen, and the substrate to be hydroxylated. The catalytic mechanism of flavoprotein aromatic hydroxylases, hence also of HbpA, can be divided into a reductive and an oxidative half reaction (Scheme 2) (van Berkel et al., 1997). In the reductive half reaction substrate binds to the enzyme and the flavin is reduced by NAD(P)H. In the oxidative part, molecular oxygen reacts with the reduced flavin and forms a flavin (C4a)-hydroperoxide. Subsequently the terminal oxygen is transferred to the substrate. After product and water release the enzyme can undergo another catalytic cycle.
Substrate NAD(P)H NAD(P) + EFlox EFlred-S R R - N N O N N O
NH NH N N H O O Substrate
H2O H O + 2 2 O2 + H Product
R R N N O N N O NH N NH H O N O H O HO H O EFlHOH-P EFlHOOH-S
Scheme 3: Reaction cycle of flavoprotein aromatic hydroxylases (van Berkel et al.,
1997). EFlox, enzyme containing oxidized flavin; EFlred-S, reduced flavin enzyme-substrate complex; EFlHOOH-S, flavin C(4a)-hydroperoxide enzyme-substrate complex; EFlHOH-P, flavin C(4a)-hydroxide enzyme-product complex.
32 CHAPTER I
The reductive and oxidative half-reactions of HbpA have been studied in detail (Suske et al., 1999). As observed for other flavoprotein aromatic hydroxylases, substrate binding stimulated the anaerobic reduction of the flavin. When the reduced flavin was oxidized in the presence of substrate, two consecutive spectral intermediates were observed, which were assigned to the flavin C(4a)-hydroperoxide and flavin C(4a)-hydroxide. As a result, the conversion of the flavin peroxide to the hydroxide intermediate reflects the transfer of the terminal oxygen to 2-hydroxybiphenyl. Substrate hydroxylation was much faster than the decay of the flavin C(4a)-hydroxide. Thus, the resulting elimination of water is presumably the limiting step in overall catalysis.
Application of 2-hydroxybiphenyl 3-monooxygenase 2-Hydroxybiphenyl 3-monooxygenase has three main features which are especially helpful for biotechnological applications. First, HbpA has a broad substrate range and catalyzes the ortho-hydroxylation of different 2-substituted phenols (Kohler et al., 1988). Thus the synthesis of different products with a single process is possible (Held, 2000). Second, the hbpA gene could be cloned into E. coli and HbpA levels of up to 30% of total protein were reached in this host (Schmid, 1997). Such a recombinant strain was used for the in vivo gram scale synthesis of different 3-substituted catechols, and 3-phenylcatechol was even produced in kilogram scale (Held, 2000). These processes were designed such that the bactericidal properties of the substrate and the instability of the formed products could be handled (Held et al., 1999). Third, HbpA can easily be purified and is stable over 8 hours at optimal reaction conditions or several months in lyophilized form. This allows the uncoupling of biocatalyst production from biotechnological organic synthesis and consequently simplifies both processes. In doing so, HbpA could be used like a conventional chemical catalyst in a two liquid phase system (Schmid et al., 2001). The required cofactor was successfully regenerated either enzymatically by formate dehydrogenase from Candida boidinii or
33 INTRODUCTION
electrochemically using a rhodium complex (Hollmann et al., 2001; Schmid et al., 2001).
CATECHOLS
Catechols (1,2-dihydroxybenzenes) and products derived from them are omnipresent in our daily life. For example, catechol is used as a reagent for photography, for coloring fur, for the production of rubber and plastic, and in the pharmaceutical industry (Merck, 1989). In nature, catechols occur as intermediary compounds in the degradation of aromatics and lignin by microorganisms (Crawford, 1981; van der Meer et al., 1992), as metabolites during degradation of benzene or estrogens in humans and mammals (Bolton et al., 1998; Porteous & Williams, 1948), and as endogenous compounds, such as neurotransmitters and their precursors (Andén, 1979).
Chemical properties of catechols Catechols can undergo a variety of chemical reactions. They can form stable complexes with different di- and trivalent metal ions (Martell & Smith, 1989). These complexes are so stable that catecholic nuclei can sequester metals from other complexes, thus preventing metals from undergoing redox reactions (Avdeef et al., 1978). In the presence of heavy metals, such as copper, catechols can be oxidized to semiquinone radicals and benzoquinones (Irons & Sawahata, 1985). As a result cycling reactions between catechols in different oxidation states occurs. Catechol oxidation does not take place in the case of di-coordinated iron (III) complexes, whereas mono-coordinated complexes allow iron to be involved in electron transfer reactions (Avdeef et al., 1978).
Biological activity of catechols The biological activity of catechols is as diverse as its chemical properties. For example, catechols can act both as antioxidant, preventing lipid peroxidation, and as pro-oxidant damaging macromolecules such as DNA and proteins (Schweigert et al., 2001b). A short summary of the biological effects of catechols is given below.
34 CHAPTER I
Inhibition of lipid peroxidation Lipid peroxidation (LPO), the oxidative modification of polyunsaturated fatty acids, is a chain reaction, which can be initiated by a variety of oxidizing compounds. LPO results in membrane damage and is the cause of a range of pathological processes (Dix & Aikens, 1993). Catechols and catechol derivatives, such as adrenaline, dopamine, and catechol estrogens, have been shown to have antioxidative properties and strongly inhibit LPO, most likely by acting as scavengers for free radicals (Liu & Mori, 1993; Schweigert et al., 2001b).
Damaging of DNA Catechols as such do not cause oxidative DNA damage in vitro. However, in the presence of heavy metals and molecular oxygen, reactive oxygen species are formed and DNA strand breaks can be observed (Li & Trush, 1994). Furthermore, catechols can cause chromatid strand breaks, chromosome aberrations in metaphase, and chromatid exchanges (Hecht et al., 1981; Stich et al., 1981). Semiquinones and/or quinones formed from catechol estrogens can also damage DNA directly by covalently binding to the DNA (Dwivedy et al., 1992). Inhibitory effects on DNA synthesis in the mouse lymphoma cell line L5178YS have been attributed to DNA alkylation and oxidative DNA damage by catechol (Pellack-Walker et al., 1985). However, similar effects in human T lymphoblasts have been explained by enzyme inhibition through the formation of iron-catechol complexes (Li et al., 1997).
Inactivation of proteins Oxidized catechols can react with sulfhydryl groups of proteins and glutathione. As a consequence, catechols can directly inhibit proteins by covalently binding to amino acid residues. Alternatively they can produce protein and glutathione radicals, which might subsequently lead to protein cross-linking and glutathione dimer formation (Stoyanovsky et al., 1995). As glutathione is an important reductant, dimer formation changes the redox status of the cell and induces oxidative stress (Halliwell & Gutteridge, 1989). The
35 INTRODUCTION
formation of catechol-metal complexes may also result in protein inactivation, since metal ions are required by many proteins (Li et al., 1997).
Destruction of membrane potentials Catechols are able to destroy biological membranes, not by reactive oxygen species or reaction with lipid molecules, but by destruction of the energetic membrane potential. In membrane vesicles millimolar concentrations of catechol and 4-chlorocatechol were shown to uncouple electron transport from ATP synthesis by perturbing the lipid bilayer (Schweigert et al., 2001a). In contrast, higher chlorinated catechols destroy the membrane potential by actively transporting electrons across the membrane (Schweigert et al., 2001a).
Catechols in the pharmaceutical industry Because catechols exhibit a variety of molecular modes of action in biological systems, they have attracted the attention of the pharmaceutical industry. For example, the observation that catechols can damage DNA was used in the development of anti-cancer drugs, such as VP-16-213, a synthetic molecule with two catechol functionalities, that induces both single- and double-strand DNA breaks in tumor cells (Kalyanaraman et al., 1989). The in vivo chelating properties of catechols are used in drugs against uranium oxide accumulation in humans (Durbin et al., 1997). That catechols can act as antioxidants explains their application in cancer prevention (Ito & Hirose, 1989). In addition to these designed synthetic compounds, naturally occurring molecules containing catechol functionalities are of interest for therapeutic applications. For example, the diterpene barbatusol is known to lower blood pressure in mice, which makes it potentially valuable for the treatment of hypertension (Majetich et al., 1993). Another large family of pharmaceutically active catechol derivatives is that of the adrenergenic catechols, such as dopamine, dobutamine, and isoproterenol (Kirk & Creveling, 1986; Miura et al., 1998). In addition to these molecules containing catechol functionalities, catechols are valuable precursors for pharmaceuticals (Ennis & Ghazal, 1992; Hartog & Wouters, 1988). An example is taxodione, a compound that showed anti-tumor
36 CHAPTER I
properties in animal trials, and whose synthesis involves 3-i-propylcatechol (Stevens & Bisacchi, 1982a).
Synthesis of 3-substituted catechols Catechols can be obtained from different natural sources. For example, catechol can be isolated by dry distillation of Mimosa catechu (Neumüller, 1979). Substituted catechols, especially chlorinated and methylated derivatives, are by-products in pulp and oil mills (Capasso et al., 1995; Neilson et al., 1991). 3-Substituted catechols mainly occur as metabolites during the degradation of benzoic compounds by microorganisms and are not directly accessible for isolation.
Chemical synthesis of 3-substituted catechols Chemical synthesis of 3-substituted catechols is laborious and involves many reaction steps, resulting in low overall product yields. In general catechol serves as starting material and the substituent is introduced by electrophilic aromatic substitution. Therefore, blocking of the hydroxyl groups of catechol by strong protecting groups is required (Stern et al., 1957). Nevertheless, the main reaction product of electrophilic aromatic substitution on protected catechol is the 4-substituted isomer. The 3-substituted catechol has to be isolated from its isomer and the protected groups have to be removed (Stern et al., 1957). Due to the importance of 3-substituted catechols as building blocks, many alternative routes to these compounds have been investigated. For example, synthesis of 3-alkylated catechols is achieved by deprotonation of veratrole (1,2-dimethoxybenzene) with an organometallic reagent followed by electrophilic substitution of the metal ion. After deprotection of the resulting 3-substituted veratrole the corresponding catechol is obtained (Majetich & Liu, 1993). An even more promising approach for the synthesis of 3-alkylated catechols is the intramolecular cyclization of alkylated fufurals, which proceeds at overall yields between 20 and 50% (Miyakoshi & Togashi, 1990). Unfortunately, this procedure is only applicable for the production of 3-n-alkylcatechols. The use of heterogeneous catalysts has also been
37 INTRODUCTION
considered. However, alkylation of catechol over acidic zeolites or in the presence of silica gels mainly yielded the 4-substituted isomer (Kamitori et al., 1984; Yoo et al., 1999). 3-Alkyl- and 3-arylcatechols can also be synthesized from the bisdimethylacetal of ortho-benzoquinone. Electrophilic substitution of a hydride ion vicinal to the acetal functionality by organometallic reagents results in the formation of the corresponding 3-substituted veratrole (Kikuchi et al., 1982), from which the catechol can be obtained by deprotection. A route for the synthesis of 3-bromoveratrole is the incubation of veratrole with bromine in the presence of organometallic reagents (Stevens & Bisacchi, 1982b). Subsequent deprotection by trichloride in chlorobenzene is not efficient and results in low yields (Mason, 1947). 3-Chlorocatechol can be obtained in good yields from 3-aminoveratrole, but preparation of this starting material is troublesome (Hornbaker & Burger, 1955). Synthesis of 3-fluoroveratrole starts from 3-fluoroanisole and also requires metal-organic reagents (Ladd & Weinstock, 1981). However, yields are moderate and scale-up difficult (Ladd et al., 1985). In summary, many different approaches for the synthesis of 3-substituted catechols have been undertaken. Yields are generally low. In addition, most synthetic routes include organometallic reagents, which leads to problems with handling, safety, and waste disposal.
Biocatalytic synthesis of 3-substituted catechols Catechols are key metabolites of microbial degradation of aromatic hydrocarbons (Harayama & Timmis, 1987). Thus, enzymes that form catechols are widely distributed. In microorganisms catechols are mineralized further by ring-cleaving dioxygenases which open the aromatic ring. This has to be considered for the biotechnological production of catechols, because the cells degrade the product formed. Several approaches have been successfully undertaken to solve this problem. The first biocatalytic synthesis of 3-substituted catechols was performed with a Pseudomonas putida wild type strain grown on toluene (Gibson et al., 1968).
38 CHAPTER I
Feeding of halogenated benzenes resulted in the accumulation of the corresponding 3-substituted catechols because the ring-cleaving catechol-2,3-dioxygenase is inhibited by halocatechols. This inhibition was also used for the production of analytical amounts of ten different alkylated, halogenated, and aryl-substituted catechols by Pseudomonas sp. T12, which were co-synthesized with the catechols derived from benzonitrile or fluorobenzene (Johnston & Renganathan, 1987). Another strategy to prevent the further degradation of the initially formed catechols is the use of knockout mutants, which do not have enzymes with catechol ring-cleaving activity. Such a strain, P. putida 2312, was used for the synthesis of catechol from benzene and 3-methylcatechol from toluene (Robinson et al., 1992). In these processes the starting materials were fed through the vapor phase and the highly bactericidal 3-methylcatechol continuously removed by adsorption on activated charcoal. In situ product recovery was also used for the synthesis of 3-fluorocatechol with resting cells of P. putida ML2, an elaborate approach to biotechnological synthesis of 3-substituted catechols (Lilly & Woodley, 1996). The process was divided into three compartments: The bioreactor, an external loop with activated charcoal for product adsorption, and an external membrane oxygenator to increase the oxygen transfer into the culture broth. Using this set- up, a total of more than 35 g of 3-fluorocatechol was produced within 14 hours in a reaction volume of 2.5 L (Lilly & Woodley, 1996). Another method to prevent the degradation of the products is to perform the biotransformation in a recombinant organism such as E. coli, which can not metabolize catechols. For example, this was done with 2-hydroxybiphenyl 3-monooxygenase from P. azelaica HBP1, which enabled the synthesis of different substituted catechols (Held et al., 1998; Schmid et al., 1998a). Because E. coli is sensitive towards 3-substituted catechols, a process with in situ product removal had to be applied. In contrast to previous approaches where adsorption of catechols on charcoal was used, the
39 INTRODUCTION
hydrophobic resin Amberlite XAD-4 was applied for product removal (Held et al., 1999).
GOAL OF THIS RESEARCH
There are three main possibilities to obtain a biocatalyst for the synthesis of a specific compound: i) testing commercially available enzymes, ii) screening of organisms with the desired enzymatic activity from natural sources, and iii) altering the substrate specificity of a biocatalyst, which has already proven its potency in a biotechnological process. In recent years the third approach has become most popular and novel techniques like directed enzyme evolution have been introduced. The research done in this thesis was focused on the modification of 2-hydroxybiphenyl 3-monooxygenase (HbpA) from Pseudomonas azelaica HBP1 for the synthesis of 3-substituted catechols that were not accessible by the wild-type enzyme. HbpA has been successfully applied to the synthesis of different 3-substituted catechols both in vitro and in vivo. Therefore, HbpA variants with altered substrate specificities can be introduced directly in existing biotechnological processes. Another goal of this work was to obtain information about the structure-function relationships in HbpA. Such information could give insight into the evolutionary relationship of flavoprotein aromatic hydroxylases and allow the engineering of new biocatalysts by rational design.
SCOPE OF THIS THESIS
In Chapter II we describe the directed evolution experiment that enabled us to change the substrate specificity of 2-hydroxybiphenyl 3-monooxygenase and the biochemical characterization of several variants obtained. The application of one of these mutant monooxygenases in the biocatalytic synthesis of 3-tert-butylcatechol is illustrated in Chapter III. Chapter IV outlines the hydroxylation of indole by a HbpA variant, which was obtained as a side product of a directed evolution experiment. The effects of the amino acid substitutions in this enzyme are initially described and further examined in Chapter V. In
40 CHAPTER I
Chapter VI the crystallization and preliminary X-ray analysis of wild-type and the selenomethionine labeled HbpA, which serve as the basis for future structural analysis, is described. Finally, general conclusions are drawn in Chapter VII.
41
CHAPTER II
CHAPTER II:
CHANGING THE SUBSTRATE REACTIVITY OF
2-HYDROXYBIPHENYL 3-MONOOXYGENASE FROM
PSEUDOMONAS AZELAICA HBP1 BY DIRECTED EVOLUTION
ANDREAS MEYER, ANDREAS SCHMID, MARTIN HELD, ADRIE H. WESTPHAL, MARTINA
RÖTHLISBERGER, HANS-PETER E. KOHLER, WILLEM J. H. VAN BERKEL,
AND BERNARD WITHOLT
JOURNAL OF BIOLOGICAL CHEMISTRY, 2002, 277(7): 5575-5582
43 DIRECTED EVOLUTION OF 2-HYDROXYBIPHENYL 3-MONOOXYGENASE
SUMMARY
The substrate reactivity of the flavoenzyme 2-hydroxybiphenyl 3-monooxygenase (EC 1.14.13.44, HbpA) was changed by directed evolution using error prone PCR. In situ screening of mutant libraries resulted in the identification of proteins with increased activity towards 2-tert-butylphenol and guaiacol (2-methoxyphenol). One enzyme variant contained amino acid substitutions V368A/L417F, which were inserted by two rounds of mutagenesis. The double replacement improved the efficiency of substrate hydroxylation by reducing the uncoupled oxidation of
NADH. With guaiacol as substrate, the two substitutions increased Vmax from
-1 0.22 to 0.43 U mg protein and decreased the K'm from 588 to 143 µM
improving k'cat/K'm by a factor of 8.2. With 2-tert-butylphenol as the substrate,
k'cat was increased more than five fold. Another selected enzyme variant contained amino acid substitution I244V and had a 30% higher specific activity with 2-sec-butylphenol, guaiacol and the ‘natural’ substrate 2-hydroxybiphenyl.
The K'm for guaiacol decreased with this mutant but the K'm for 2-hydroxybiphenyl increased. The primary structure of HbpA shares 20.1% sequence identity with phenol 2-monooxygenase from Trichosporon cutaneum. Structure homology modeling with this three-domain enzyme suggests that Ile244 of HbpA is located in the substrate binding pocket and is involved in accommodating the phenyl substituent of the phenol. In contrast, Val368 and Leu417 are not close to the active site and would not have been obvious candidates for modification by rational design.
44 CHAPTER II
INTRODUCTION
2-Hydroxybiphenyl 3-monooxygenase (EC 1.14.13.44, HbpA) belongs to the family of flavoprotein hydroxylases (Eppink et al., 1997; Suske et al., 1997; Suske et al., 1999). These enzymes are involved in many important biological processes, such as the biosynthesis of cholesterol or the degradation of xenobiotics in mammals and in nature (Hines et al., 1994; Laden et al., 2000; van Berkel & Müller, 1991). HbpA was first found in Pseudomonas azelaica HBP1, a soil bacterium which is able to grow on the fungicide 2-hydroxybiphenyl as sole source of carbon and energy (Kohler et al., 1988). HbpA catalyses the ortho-hydroxylation of 2-hydroxybiphenyl to 2,3-dihydroxybiphenyl, which is then converted to 2-hydroxy-6-phenyl-6-oxo-2,4-hexadienoic acid (HOPDA) by a meta ring cleavage dioxygenase (HbpC). HOPDA is hydrolyzed by HbpD to benzoate and 2-hydroxy-2,4-pentadienoic acid (Kohler et al., 1988; Kohler et al., 1993b), which are further metabolized via intermediates also formed in the analogous biphenyl degradation pathway (Catelani et al., 1973; Gibson et al., 1973). HbpA has a broad substrate spectrum, catalyzing the regioselective ortho- hydroxylation of a wide range of 2-substituted phenols to the corresponding catechols (Fig. 1) (Kohler et al., 1988; Kohler et al., 1993a).
+ + NADH + O2 + H NAD + H2O OH OH OH
R HbpA R
R = phenyl, 2-OH-phenyl, methyl, ethyl, propyl, i -propyl, butyl, sec -butyl, fluoro, chloro, bromo, or iodo.
Fig. 1: Reaction catalyzed by wild-type HbpA. The substrate spectrum of wild-type HbpA is shown
45 DIRECTED EVOLUTION OF 2-HYDROXYBIPHENYL 3-MONOOXYGENASE
Recently, the hbpA gene was cloned into Escherichia coli and this recombinant biocatalyst has been used for the production of different 3-substituted catechols (Held et al., 1999; Held et al., 1998). One of these, 3-phenylcatechol, was produced in kilogram scale, showing that the biocatalytic production of 3-substituted catechols is a possible alternative to chemical synthesis routes (Held, 2000). HbpA mutants with an altered substrate reactivity should allow the synthesis of catechols which are not synthesized by wild-type HbpA. Rational protein design based on a known three-dimensional structure has been used for such purposes (Graf et al., 1987; Nickerson et al., 1997; Pikus et al., 1997; van den Heuvel et al., 2000a), but random approaches have lately become more popular (Schmidt-Dannert & Arnold, 1998). Directed enzyme evolution (Arnold, 1998a; Arnold & Volkov, 1999; Moore & Arnold, 1996; Stemmer, 1994), the most often used strategy, has been applied to improve substrate specificity, activity, enantioselectivity, or thermostability (Giver et al., 1998; Kuchner & Arnold, 1997; May et al., 2000; Moore & Arnold, 1996; Reetz et al., 1997). Here we report on the use of directed evolution to change the substrate reactivity of HbpA. We increased the specific activity of HbpA towards 2-hydroxybiphenyl, 2-sec-butylphenol, guaiacol (2-methoxyphenol), and 2-tert-butylphenol. Moreover, a significant increase in the efficiency of NADH utilization was achieved with one mutant monooxygenase. These results are interpreted at the structural level with the help of a three-dimensional model of HbpA.
MATERIALS AND METHODS
Chemicals, bacterial strains and plasmids E. coli JM101 and plasmid pAA1 were used for cloning and gene expression. Plasmid pAA1 is a pUC18 (Yanish-Perron et al., 1985) derivative harboring the hbpA gene as a SalI/NsiI fragment (Schmid, 1997) cloned into the SalI/PstI sites of the pUC18 polylinker.
46 CHAPTER II
Commercially available chemicals were purchased from Fluka AG (Buchs, Switzerland). Catalase from beef liver was obtained from Boehringer (Mannheim, Germany). Taq DNA polymerase, restriction enzymes, and T4 DNA ligase were purchased from Roche Molecular Biochemicals (Basel, Switzerland). 2,3-Dihydroxybiphenyl and 3-sec-butylcatechol were prepared by whole-cell biotransformations, using a recombinant E. coli JM101 containing the hbpA gene (Held, 2000).
Random mutagenesis The hbpA gene (1758 bp) in pAA1 was amplified using in vitro manganese mutagenesis (Beckmann et al., 1985). For the PCR reaction the M13/pUC-40 primers (MWG-Biotech GmbH, Münchenstein, Switzerland) were used, each of which complements a 23 bp region of the cloning vector. A 100 µL volume
containing 50 mM KCl, 10 mM Tris-HCl (pH 9), 6.5 mM MgCl2, 0.1% Triton X-
100, 10 µL DMSO, 0.5 mM MnCl2, 1 mM dNTPs, 15 pmol of each primer, 20 ng of template DNA, and 2.5 U of Taq DNA polymerase (Promega, Madison, WI, USA) was placed in a Perking Elmer (Emeryville, CA, USA) thermal cycler well. After 5 min at 95°C, the thermal cycler performed 25 cycles of the following steps: 1 min at 95°C, 1 min at 55°C, 2 min at 72°C. Prior to restriction the amplified DNA was purified with a DNA clean-up kit (Genomed GmbH, Bad Oeyenhausen, Germany, or Macherey-Nagel AG, Oensingen, Switzerland).
Cloning procedures
Competent E. coli JM101 cells were prepared using a modified CaCl2 based method (Sambrook et al., 1989). A 5 mL culture of E. coli JM101 was grown overnight in LB. The cells were diluted 1:100 into fresh LB and grown until they
reached an OD600 of 0.5±0.1. After centrifugation (5500 x g, 4°C, 8 min) the supernatant was discarded and the pellet resuspended in 0.2 volumes of
10 mM sodium acetate (pH 5.8), 50 mM MnCl2, and 5 mM NaCl. The suspension was put on ice for 30 min and recentrifuged as before. The pellet
was dissolved in 0.1 volumes of 10 mM sodium acetate (pH 5.8), 70 mM CaCl2,
47 DIRECTED EVOLUTION OF 2-HYDROXYBIPHENYL 3-MONOOXYGENASE
5 mM MnCl2, and 5% glycerol and stored in aliquots of 100 µL at -70°C until use. Ligation mixtures containing pUC18 (cut BamHI/SphI), amplified hbpA gene (cut BamHI/SphI), 0.5 U T4 DNA ligase (Boehringer Mannheim, Mannheim, Germany), and 10x ligation buffer were incubated overnight at 4°C. The ligation mixture was directly used for transformation. For this, an aliquot of competent E. coli JM101 was thawed on ice. The cells were mixed with 1 µL of ligation mixture and placed on ice for 30 min. The heat pulse was performed at 42°C for 90 sec and followed by incubation on ice for 1 min. After the addition of 1 mL LB the cells were incubated at 37°C for one hour. The complete transformation mixture was transferred onto selective LB plates and incubated overnight at 30°C.
Screening The screening procedure for the desired modified HbpA was based on the instability of the reaction products. At neutral pH catechols autooxidize to quinones and semiquinones, which readily form reddish or brownish, undefined, high molecular weight compounds (Ziechmann, 1980). After transformation, E. coli JM101 transformants were transferred directly onto LB plates containing 150 µg mL-1 ampicillin, 0.04‰ (w/v) Xgal, 200 µM IPTG, and 0.1-0.5 mM of the 2-substituted phenol to be screened for. Incubation at 30°C resulted in three colony types: blue colonies, which did not contain the amplified hbpA gene; white colonies, which contained an hbpA gene that encoded for an inactive enzyme towards the aromatic test substrate, or an inactive lacZ gene due to frame shifted ligation; and reddish brown colonies, which contained an enzyme with activity towards the added 2-substituted phenol. The time dependent intensity of color formation in the latter colonies could be used to distinguish between different activities. For substrates which were only poorly transformed to the corresponding catechol, the color formation could be intensified by the addition of 1.5 mM ferric chloride and 50 µg mL-1 p-toluidine (Parke, 1992). If
48 CHAPTER II
positive clones were detected, the plasmids containing the amplified hbpA genes were isolated and retransformed into E. coli JM101.
Preparation of cell extracts Cells from a 5 mL LB culture were spun down at 5000 x g for 15 min and resuspended in 800 µL 50 mM phosphate buffer (pH 7.2). This suspension was transferred to a 1.5 mL Eppendorf tube containing 1.2 g glass beads (Ø 0.1 - 0.2 mm) and the cells were disrupted in a Retsch mill (Retsch GmbH, Hann, Germany) for 10 min at 90% power. The cell extracts were separated from the glass beads by centrifugation (15000 x g, 15 min) and supplemented with FAD to a final concentration of 50 µM. Cell extracts could be stored at -20°C for 2-3 weeks without significant loss of HbpA activity.
Enzyme purification The purification of HbpA and its mutants was based on a simplified version of a procedure described earlier (Suske et al., 1997). 6 g (cell wet weight) of frozen E. coli JM101, harboring a pUC18 derivative encoding either the wild- type or the amplified hbpA gene, were suspended in 25 mL phosphate buffer (10 mM, pH 7.5). Cell extract was prepared by twice passing the suspension through a French pressure cell (20 K SIM AMINCO) at 70 bar, followed by ultracentrifugation at 4°C (Beckmann L8-60M, 40000 x g, 30 min). The clarified cell extract was diluted 1:1 with triethanolamine-HCl buffer (10 mM, pH 7.5) and loaded directly onto an anion exchange column (1.5 x 15 cm; Fractogel EMD DMAE-650 (S); Merck, Darmstadt, Germany) equilibrated with 10 mM triethanolamine-HCl buffer (pH 7.5). Elution was carried out with a linear gradient from 0 to 1 M NaCl in starting buffer. Fractions containing HbpA activity were pooled, supplemented with 0.9 M ammonium sulfate and loaded onto a hydrophobic interaction chromatography column (1 x 8 cm; Butyl Sepharose 4 Fast Flow; Pharmacia Biotech, Uppsala, Sweden) equilibrated with 0.75 M ammonium sulfate in 100 mM sodium phosphate buffer (pH 7.0). Elution was carried out with a linear gradient from 0.75 to 0 M ammonium sulfate in 100 mM sodium phosphate buffer (pH 7.0).
49 DIRECTED EVOLUTION OF 2-HYDROXYBIPHENYL 3-MONOOXYGENASE
Fractions containing HbpA were pooled and concentrated in an Ultrafree-15 centrifugal filter device (Biomax-50K; Millipore Corporation, Bedford, MA). Concentrated enzyme (0.3 mL) was supplemented with 0.3 mM FAD and passed through a Superdex 200 gel filtration column (1.6 x 60 cm; Pharmacia Biotech AB, Uppsala, Sweden) equilibrated with 50 mM sodium phosphate buffer (pH 7.5). Enzyme purity was assessed with SDS-PAGE (12% polyacrylamide), followed by staining with coomassie brilliant blue.
Analytical methods As is also the case for other flavin containing oxygenases, NADH oxidation is partially uncoupled from substrate hydroxylation in HbpA (Suske et al., 1997). Therefore specific activities were determined both for NADH oxidation and substrate consumption/product formation. NADH oxidation was followed spectrophotometrically at 340 nm or polarographically by monitoring oxygen consumption with an oxygen electrode (Kohler et al., 1988). The assay contained 0.2-1 mM substrate, 0.3 mM NADH, 20 mM air saturated phosphate buffer (pH 7.5), and 10-20 µL cell extract or purified protein in a total volume of 1 mL. To determine substrate utilization and product formation, the enzymatic reaction was stopped by the addition of perchloric acid and the resulting precipitate was removed by centrifugation (15000 x g, 10 min). The samples were diluted 1:1 with MeOH/0.1% phosphoric acid and analyzed with a Hypersil ODS column (5 µm, 4.5 x 125 mm) using a Hewlett Packard HP 1050 Ti HPLC coupled to a diode array detector (HP DAD 1040M). The elution was carried out
under isocratic conditions with MeOH/H2O (0.1% phosphoric acid) as mobile phase. Steady-state kinetic parameters were calculated by weighted non-linear regression analysis (Enzfitter, Elsevier-Biosoft, UK). Uncoupling of product formation from oxygen consumption was resolved by performing HbpA activity assays in the absence and in the presence of catalase (Entsch et al., 1976). The catalase recycles 50% of the oxygen which is used for hydrogen peroxide formation. By determining initial reaction rates in the absence and presence of
50 CHAPTER II
catalase the substrate related uncoupling of product formation from oxygen consumption could be calculated.
HbpA modeling The sequence of HbpA was aligned to the recently corrected sequence of phenol 2-monooxygenase from Trichosporon cutaneum (Xu et al., 2001) with
ClustalX (Thompson et al., 1997) using the PAM 350 matrix. Model building of
HbpA was performed with MODELLER (Sali & Blundell, 1993) using the CVFF forcefield (Dauber-Osguthorpe et al., 1988). The closed form of the phenol 2-monooxygenase structure (PDB entry: 1foh) was used as a template. The model was verified after several rounds of energy minimization. The
stereochemical quality of the homology model was verified by PROCHECK
(Laskowski et al., 1993), and the protein folding was assessed with PROSAII (Sippl, 1993), which evaluates the compatibility of each individual residue with its environment. The FAD and the substrate 2-hydroxybiphenyl were placed in identical positions and orientations as the FAD and phenol in the template structure.
Nomenclature Subscript letters indicate the substrate on which the mutant was screened (G, guaiacol; T, 2-tert-butylphenol); numbers indicate the round of error prone PCR. HbpA* means HbpA-variants in general.
RESULTS
Directed evolution of HbpA 2-Hydroxybiphenyl 3-monooxygenase (HbpA) was subjected to in vitro manganese mutagenesis with error prone PCR (Leung et al., 1989) and subsequent in situ screening for enzymes with an altered substrate reactivity (HbpA*). The base substitution rate in the error prone PCR was tuned to an exchange rate of 1-3 per hbpA gene, to produce an average of one amino acid substitution per HbpA* (Kuchner & Arnold, 1997). The Mn2+ concentration was adapted to template composition, template length, dNTP concentration, and
51 DIRECTED EVOLUTION OF 2-HYDROXYBIPHENYL 3-MONOOXYGENASE
polymerase type. We tested different Mn2+ concentrations in a range of 0.1- 1 mM. At a concentration of 0.5 mM Mn2+ 60-70% of the amplified hbpA genes encoded for active HbpA. Sequencing of randomly picked active or inactive clones showed that on the average there were 1.2 amino acid substitutions per HbpA*. With respect to the base substitutions, transitions exceeded transversions by a factor of 2. The mutant library was plated on substrate containing medium, where active HbpA produces aromatic polymers. The improvement of enzyme activity is
generally associated with a decrease of the Km towards the substrate (Kuchner & Arnold, 1997). We used aromatic substrate concentrations that were lower
than the Km of the parent enzyme. The color of the polymer formed depended on the screening substrate used; the polymer formed from 3-methoxycatechol was brownish, while the 3-tert-butylcatechol polymer was reddish. Figure 2 shows E. coli JM101 growing on solidified LB medium containing 0.5 mM
2-tert-butylphenol and expressing either wild-type hbpA or hbpAT2.
Fig. 2: E. coli synthesizing HbpA and HbpAT2 on LB medium containing 0.5 mM 2-tert-butylphenol. 2-tert-Butylphenol was directly added to the medium, and cells were allowed to grow overnight at 30°C. Time and intensity of the reddish color formation, which results from 3-tert-butylcatechol polymerization, was used to select enzymes with different substrate reactivities.
After the first round of mutagenesis active clones were screened for increased activity on guaiacol and 2-tert-butylphenol. From each experiment (500 clones) we took 6-8 clones which showed increased color formation, and determined NADH oxidation in crude cell extracts as a function of the test substrate and the physiological substrate 2-hydroxybiphenyl. In cases where the activity towards the test-substrate or the ratio between the activities for the test-substrate and 2-hydroxybiphenyl was higher than for the parent enzyme,
52 CHAPTER II
product formation was analyzed by reverse phase HPLC. The levels of HbpA and HbpA* were checked with SDS-PAGE to correct for different expression levels. Eight clones were initially selected following in situ screening on guaiacol. In five cell extracts the recombinant protein level was increased but the specific HbpA* activity remained constant. Two cell extracts contained HbpA* with a lower in vitro activity than the wild type enzyme. Sequencing revealed that both enzymes contained one amino acid substitution, which probably decreased the enzyme stability in vitro but not in vivo. One mutant monooxygenase was found to have an increased specific activity towards guaiacol, and was named
HbpAG1. Six clones were initially selected following in situ screening on 2-tert-butylphenol. Only one clone harbored an hbpA* gene with a base substitution that led to an amino acid exchange and a higher activity towards
this substrate. This gene (hbpAT1) was used for another round of error prone PCR and in situ screening. From two selected clones, we obtained one mutant monooxygenase with a higher activity towards 2-tert-butylphenol. This variant
was named HbpAT2.
Purification and characterization of mutant enzymes
The hbpAT1 and the hbpAG1 genes each contained one and the hbpAT2 gene contained two base substitutions which led to an amino acid change (Table 1).
In addition, hbpAT1 and hbpAT2 each carried one silent mutation and hbpAG1 carried 2 base substitutions which did not result in an amino acid change. Wild-type and mutant HbpA were purified from recombinant E. coli JM101 harboring a pUC18 derivative carrying the corresponding hbpA gene. Using this system, HbpA and HbpA* could be overexpressed to about 20% of total cell protein. The enzymes were purified as tetramers to homogeneity with yields around 30%. Purity was confirmed by SDS-PAGE (Fig. 3), which showed only HbpA or HbpA* monomers.
53 DIRECTED EVOLUTION OF 2-HYDROXYBIPHENYL 3-MONOOXYGENASE
Table 1: Amino acid substitutions in HbpA mutants Amino Acid Position Enzyme 244 368 417 HbpA Ile Val Leu → HbpAG1 Ile Val Val Leu → HbpAT1 Ile Val Ala Leu → → HbpAT2 Ile Val Ala Leu Phe
Fig. 3: SDS-PAGE of purified HbpA and mutants. Coomassie blue stained SDS-polyacrylamide gel containing 4 µg protein per lane. Proteins were purified from a recombinant E. coli JM101, which expressed either the hbpA or hbpA* gene. Lanes: M, marker; A,
HbpA; G1, HbpAG1; T1, HbpAT1; T2, HbpAT2.
The mutant enzymes followed Michaelis-Menten kinetics with all substrates
* tested as does wild-type HbpA. For K'm determinations, HbpA and HbpA activities were measured by NADH oxidation at different substrate
concentrations. All mutants showed an increased K'm towards the natural
substrate 2-hydroxybiphenyl, whereas K'm remained unchanged for
2-sec-butylphenol. The K'm towards guaiacol was significantly decreased for all mutants (Table 2). Because HbpA and HbpA* showed uncoupling of NADH oxidation from substrate hydroxylation for all substrates tested, apparent turnover rates were determined by measuring product formation and/or substrate consumption with
reverse phase HPLC (Table 3). Using this method, HbpAG1 showed a 30% increased specific activity towards 2-hydroxybiphenyl, 2-sec-butylphenol, and
guaiacol. Compared to HbpA, HbpAT2 showed half the activity towards 2-hydroxybiphenyl and a 12% lower activity towards 2-sec-butylphenol. At the same time, it revealed a five fold increase in activity towards 2-tert-butylphenol,
54 CHAPTER II
the substrate on which the enzyme was screened, and twice the activity of HbpA towards guaiacol and salicylaldehyde.
Table 2: Apparent Km values of HbpA and mutants towards different 2-substituted
phenols. Apparent Km values were determined by spectrophotometrically monitoring NADH consumption. The assays were performed at 30°C in 20 mM phosphate buffer (pH 7.5) with 0.3 mM NADH and following substrate concentrations: 2-Hydroxybiphenyl and 2-sec-butylphenol: 2, 3, 4, 5, 10, 15, 20, and 25 µM; guaiacol: 0.025, 0.05, 0.1, 0.2, 0.3, 0.4, 0.5, and 0.8 mM.
HbpA HbpAG1 HbpAT1 HbpAT2 a b b b b R Km Km Km Km
µM µM µM µM Phenyl 2.6±0.1 5.7±0.1 13±1.6 16±1.4 sec-Butyl 8.7±0.9 9.5±1.0 n.d.c 10.0±0.2 Methoxyd 588±13 222±13 337±58 143±19
a Substituent ortho to the phenolic hydroxy group. b Best fit parameters obtained from non-linear least square fits to the Michaelis-Menten model. c Not determined d Resulting phenol: guaiacol
Uncoupling of NADH oxidation and substrate hydroxylation The NADH oxidase activity of the mutant enzymes was determined spectrophotometrically in the absence of the aromatic substrate. At saturating concentrations of the coenzyme (6 mM), HbpA showed a NADH oxidase activity of 0.11 U mg-1 protein. The activity of the mutants was significantly higher and
-1 -1 found to be 0.19 U mg protein for HbpAG1 and 0.29 U mg protein for HbpAT2, respectively. In the presence of the aromatic substrate, the rate of NADH oxidation by HbpA or HbpA* generally exceeds the rate of substrate consumption, due to uncoupling of NADH oxidation from substrate hydroxylation (Table 4), thus lowering the hydroxylation efficiencies of these enzymes. Interestingly, while the
hydroxylation efficiencies of HbpA and HbpAG1 were similar for each of the
substrates tested, it was considerably higher for HbpAT1 and HbpAT2.
55 DIRECTED EVOLUTION OF 2-HYDROXYBIPHENYL 3-MONOOXYGENASE
Table 3: k cat and kcat/Km values of HbpA and HbpA mutants towards different
2-substituted phenols. The kcat values were determined for the tetrameric enzyme. The assays were performed at 30°C in 20 mM phosphate buffer (pH 7.5) containing 0.3 mM NADH.
Substrate concentrations used for the determination of kcat were 0.1 mM for 2-hydroxybiphenyl, 2-sec-butylphenol; 1 mM for salicylaldehyde, guaiacol, 2-tert-butylphenol.
HbpA HbpAG1 HbpAT2
a R kcat kcat/Km kcat kcat/Km kcat kcat/Km s-1 s-1µM-1 s-1 s-1µM-1 s-1 s-1µM-1 Phenylb 11.9 4.6 16.2 2.8 6.4 0.4 sec-Butylb 14.5 1.7 19.1 2.0 12.8 1.3 Methoxyb 0.95 1.6*10-3 1.28 5.8*10-3 1.83 1.3*10-2 Formylb 0.5 n.d.d 0.4 n.d. 0.9 n.d. tert-Butylc <0.1 n.d. <0.1 n.d. 0.5 n.d.
a Substituent ortho to the phenolic hydroxy group. b Activities determined by measuring product formation and substrate consumption with reverse phase HPLC. c Activities determined by measuring substrate consumption with reverse phase HPLC. d Not determined.
The uncoupling of substrate hydroxylation from NADH oxidation can have different origins (Suske et al., 1997; Suske et al., 1999). One possibility is that substrate is bound to the enzyme but not hydroxylated due to inefficient oxygen transfer from the flavin (C4a)-hydroperoxide. Alternatively, product remains bound to the enzyme and can not be hydroxylated but can induce the elimination of hydrogen peroxide (Fig. 4). To distinguish between substrate and
product related uncoupling for HbpA and HbpAG1, oxygen consumption was monitored in activity assays in the absence and in the presence of catalase. Whereas for 2-sec-butylphenol and guaiacol most uncoupling could be ascribed to the substrates, for 2-hydroxybiphenyl more than half of the uncoupling could be attributed to the product 2,3-dihydroxybiphenyl. This was confirmed by incubating the enzymes with the reaction product 2,3-dihydroxybiphenyl, which resulted in a NADH oxidase activity of 1.9 U mg-1 protein for HbpA and
-1 2.0 U mg protein for HbpAG1. With both enzyme variants, 2,3-dihydroxybiphenyl acted as a true non-substrate effector since no product
56 CHAPTER II
formation could be detected. In contrast, 3-sec-butylcatechol and 3-methoxycatechol hardly stimulated NADH oxidation in wild-type HbpA or any of the mutants.
Table 4: Uncoupling of product formation from NADH oxidation. The assays were performed at 30°C in 20 mM phosphate buffer (pH 7.5) containing 0.3 mM NADH. Substrate concentrations used: 2-hydroxybiphenyl, 2-sec-butylphenol, 0.2 mM; guaiacol, 1 mM.
Specific monooxygenase activity NADH oxidationa Product formationb Uncoupling µmol min-1 mg protein-1 µmol min-1 mg protein-1 % 2-Hydroxybiphenyl HbpA 3.65 2.89 21
HbpAG1 5.41 3.86 29
HbpAT1 3.80 3.70 3
HbpAT2 1.61 1.55 4 2-sec-Butylphenol HbpA 4.15 3.39 18
HbpAG1 5.77 4.51 22
HbpAT2 3.18 2.99 6 Guaiacol HbpA 0.98 0.22 78
HbpAG1 1.84 0.30 84
HbpAT1 0.66 0.31 53
HbpAT2 0.88 0.43 51 a All values were corrected for the endogenous NADH oxidation and have a standard error ≤10%. b Determined by measuring substrate consumption and product formation with reverse phase HPLC.
57 DIRECTED EVOLUTION OF 2-HYDROXYBIPHENYL 3-MONOOXYGENASE
Structure homology modeling To assess the effects of the amino acid replacements in the mutant enzymes, a sequence alignment between HbpA (586 residues) and phenol 2-monooxygenase from Trichosporon cutaneum (PHHY, 664 residues), the most closely related enzyme with known three-dimensional structure (Enroth et al., 1998), was performed. Figure 5 shows the alignment with the three conserved sequence motifs with a putative dual function in FAD/NAD(P)H binding (Eppink et al., 1997). Most sequence homology between HbpA and PHHY was found in the N-terminal part of the proteins, which consists of the FAD-binding and substrate-binding domains and constitutes the enzyme active site. A sequence identity of 24.4% was calculated when only these parts of HbpA and PHHY were taken into account. There was less homology in the C- terminal part of the proteins, reducing the overall sequence identity to 20.1%. The only known function of the C-terminal domain of PHHY is its participation in subunit association (Enroth et al., 1998).
Substrate NAD(P)H NAD(P) + EFlox EFlred-S R R - N N O N N O
NH NH N N H O O Substrate
H2O H O + 2 2 O2 + H Product
R R N N O N N O NH N NH H O N O H O HO H O EFlHOH-P EFlHOOH-S
Fig. 4: Reaction cycle of flavoprotein aromatic hydroxylases. Reaction cycle of
flavoprotein aromatic hydroxylases adapted from (van Berkel et al., 1997). EFlox, enzyme
containing oxidized flavin; EFlred-S, reduced flavin enzyme-substrate complex; EFlHOOH-S, flavin C(4a)-hydroperoxide enzyme-substrate complex; EFlHOH-P, flavin C(4a)-hydroxide enzyme-product complex.
58 CHAPTER II
HbpA 1 MSNSAETDVLIVGAGPAGAMSATLLASLG-----IRSLMINRWRSTSPGPRSHIINQRTMEILRDIGLEESAKSLAVPKEYMGEH 80 + DVLIVGAGPAG M+A +L+ ++ +I++ + ++ + RT+E L+++GL + S A + + PHHY 1 TKYSESYCDVLIVGAGPAGLMAARVLSEYVRQKPDLKVRIIDKRSTKVYNGQADGLQCRTLESLKNLGLADKILSEANDMSTIALY 86 < βA > <βB> < α1 > <βA> < α2 > < α3 > < βC VhhhGsGhhGhhhs FAD fingerprint (1)
HbpA 81 VYATSLAGEEFGRIPAWAS------HPQAHAEHELASPSRYCDLPQLYFEPMVVSEAALRG------ADVRFLTEY 144 RIP H + L S + D E P++ + +R+++E+ PHHY 87 NPDENGHIRRTDRIPDTLPGISRYHQVVLHQGRIERRILDSIAEISDTRIKVERPLIPEKMEIDSSKAEDPEAYPVTMTLRYMSED 172 > < βC > < α4 > < βA ><α5> < βA > <α
HbpA 145 LG------HVEDQDGVTARLLD-HVSGAEYEVRAKYIIGADGAHSLVAQNAGLPFEGQMGIGDSGSINIEFSAD 211 ++++ RL + +G V KY+IG DG HS V + G G+ G ++ +++ PHHY 173 ESTPLQFGHKTENGLFRSNLQTQEEEDANYRLPEGKEAGEIETVHCKYVIGCDGGHSWVRRTLGFEMIGEQTDYIWGVLDAVPASN 258 6> < α7 > < βA > <βB> < α8 > <βD> < βC ><βE> chhhssDGxcSxhR conserved motif
HbpA 212 LSSLCEHRKGDMYWMFRAGSGINGVGVAALRMIRPWNKWICV-WGYEKSKGTPEITKEEAKKIIHEIIGTDEIPVEVGPISTWTIN 296 + + ++ I +R +++K TPE+ +AKKI H + + + I PHHY 259 FPDIRS--RCAIHSAESGSIMIIPRENNLVRFYVQLQARAEKGGRVDRTKFTPEVVIANAKKIFHPYT---FDVQQLDWFTAYHIG 339 > <βC > < βC > < βC > < α9 > <βE> < βC ><
HbpA 297 QQYAVRNTSG-RVFCMGDAVHRHTPMGGLGLNTSVQDAYNLAWKLALVLKGTAAPTLLDSYDAERSPVAKQIVERAFKS------374 Q+ + + RVF GDA H H+P G G NTS+ D YNL WKL LVL G A +L +Y+ ER+P A +++ PHHY 340 QRVTEKFSKDERVFIAGDACHTHSPKAGQGMNTSMMDTYNLGWKLGLVLTGRAKRDILKTYEEERQPFAQALIDFDHQFSRLFSGR 425 βD> <βB><α10> < α11 > < α12 > GxxhhLhGDAAHxxxPxxGxGxNxsxxDsxxL FAD fingerprint (2)
HbpA 375 ------LSTFPPVFEALSLPPAPTESEMAEALVRLKDASEEGAKRRAALRKAMDAT-IIGLGGGHGVELNQRYVSR---- 443 + F F + T + E LV K +S + + + + + G + R V+ PHHY 426 PAKDVADEMGVSMDVFKEAFVKGNEFASGTAINYDENLVTDKKSSKQELAKNCVVGTRFKSQPVVRHSEGLWMHFGDRLVTDGRFR 511 < α13 > <α14> <βF> <βF><α15> <
HbpA 444 -AVFP----DGTPDPGFVRDQEFFYQASTRPGAHLPHVWLTENQRRISTLDLCGKGRFTLLTGLS---GAAWKH-----EAEQVSQ 516 VF D T + + + + P + + + + T+ C + + + W + + + PHHY 512 IIVFAGKATDATQMSRIKKFAAYLDSENSVISRYTPKGADRNSRIDVITIHSCHRDDIEMHDFPAPALHPKWQYDFIYADCDSWHH 597 βG > < α16 > <α17> < βG > <α18> <βG>
HbpA 517 SLGIELKVCVIGPGQEFVDTYGEYAKISEIGESGALLVRPDMFIAFRAKDASREGLEQLNVAVKSILGRA 586 + V S + + F + G + KS PHHY 598 PHPKSYQAWGVDETKGAVVVVRPDGYTSLVTDLEGTAEIDRYFSGILVEPKEKSGAQTEADWTKSTA 664 <α19> <βG> <βG > < α20 >
Fig. 5: Alignment of 2-hydroxybiphenyl 3-monooxygenase (HbpA) from Pseudomonas azelaica HBP1 and phenol 2-monooxygenase (PHHY) from Trichosporon cutaneum containing the conserved sequence motifs of flavoprotein aromatic hydroxylases. Letters and symbols between the two sequences represent identical (letters) and similar (+) residues. The consensus profiles according to Eppink et al. (Eppink et al., 1997) shown underneath the alignment include strictly conserved residues in bold letters. Uppercase letters are amino acid residues, lowercase letters and symbols are: h, hydrophobic residues; s, small residues; c, charged residues; x, all residues; -, gap. The shaded boxes mark the location of the amino acid substitutions in the HbpA mutants.
59 DIRECTED EVOLUTION OF 2-HYDROXYBIPHENYL 3-MONOOXYGENASE
Structure homology modeling with PHHY confirmed that HbpA consists of three domains. The FAD-binding and substrate-binding domains of both enzymes are structurally conserved, but the structure of the C-terminal domain of HbpA is more uncertain. Like p-hydroxybenzoate hydroxylase, which contains no extra domain (Mattevi, 1998; Schreuder et al., 1989), bacterial HbpA contains considerably fewer surface loops than eukaryotic PHHY. Figure 6 shows a model of the HbpA subunit, as obtained by using the ‘closed’ subunit of PHHY (PDB entry 1foh) as the template file. For PHHY it was reported that the two subunits in the homodimer do not have an identical conformation and that the largest difference involves a loop (residues 170-210, PHHY numbering) which can act as a lid that opens and closes the active site (Enroth et al., 1998). In HbpA, this active site loop (residues 142-164) is much shorter, but the model suggests that it is still able to cover the active site.
Fig. 6: Three-dimensional model of a subunit of HbpA. The ‘closed’ subunit conformation of phenol 2-monooxygenase from Trichosporon cutaneum served as template. The substrate binding domain is drawn in blue, the FAD binding domain in magenta and the C-terminal domain in green. Highlighted are the substrate 2-hydroxybiphenyl, the FAD prosthetic group, and the amino acids that were changed during the directed evolution experiment.
60 CHAPTER II
Further examination of the three-dimensional model of HbpA revealed that the amino acids Ile244, Val368 and Leu417 that were changed in the variants are spread throughout the structure. Ile244 is located in the substrate binding pocket and is rather close to the flavin ring. Interestingly, Ile244 corresponds with Tyr289 of PHHY. This tyrosine is believed to play an important role in catalysis by positioning the aromatic substrate for attack at the ortho position (Enroth et al., 1998). Val368 and Leu417 are both located near the protein surface of the HbpA subunit and Leu417 is positioned far away from the substrate binding site. Val368 corresponds with Ile414 in PHHY and is part of a conserved helix, whereas Leu417 corresponds with Val480 in PHHY and is located in the beginning of the C-terminal domain. Although Ile414 and Val480 are far away from the dimer interface of PHHY, it cannot be excluded that in HbpA, Val368 and Leu417 play a role in tetramer formation or tetramer stabilization.
DISCUSSION
Most members of the family of flavoprotein hydroxylases are involved in the degradation of aromatic compounds by soil microorganisms (van Berkel et al., 1997). However, the application of these redox enzymes is not restricted to the metabolism of pollutants in our environment. Due to their high regioselectivity flavoprotein hydroxylases also have considerable potential in the synthesis of new fine chemicals.
Directed evolution of HbpA Assuming an electrophilic aromatic substitution reaction mechanism (Suske et al., 1999), we chose 2-tert-butylphenol and guaiacol (2-methoxyphenol) as model substrates for directed evolution of HbpA. These substrates differ significantly from the natural substrate: i) the bulky side chain of 2-tert-butylphenol requires more room in the enzyme active site, and ii) the methoxy group of guaiacol is more polar and withdraws more charge from the aromatic system (+M, -I compared to +M, +I)(Allinger et al., 1976). Furthermore,
61 DIRECTED EVOLUTION OF 2-HYDROXYBIPHENYL 3-MONOOXYGENASE
so far no activity for the ortho-hydroxylation of guaiacol and 2-tert-butylphenol has been described. Microbial degradation of guaiacol proceeds only via demethylation (Boyd et al., 1983; Dardas et al., 1985; Sauret-Ignazi et al., 1988) whereas chlorinated guaiacols can also be degraded via para-hydroxylation (Gonzales et al., 1993; Haggblom et al., 1988). Thus, HbpA variants with an increased catalytic activity towards guaiacol and 2-tert-butylphenol will allow the biotechnological production of the corresponding catechols and may yield information about structure-function relationships in HbpA. The key factor for a successful directed evolution experiment is an effective screening or selection procedure (Kuchner & Arnold, 1997; Zhao & Arnold, 1997). With HbpA, the instability of 3-substituted catechols offered a good basis for the development of a suitable in situ screening procedure. The time dependent color-formation could be efficiently used for qualitative estimation of enzyme activity directly after construction of the mutant library. The high reliability of our in situ screening procedure was illustrated by the fact that only 2-8 clones per round of mutagenesis had to be selected to obtain a successfully modified enzyme. Furthermore, the in situ screening was not restricted to the directed evolution of HbpA towards 2-tert-butylphenol. A second mutant library was screened on guaiacol and enzymes with an increased activity towards this substrate could also be selected. This demonstrates that in situ screening provides an easy and rapid method for the detection of specific enzyme features if the corresponding assay can be applied on solid media.
Enzyme kinetics The mutant proteins, which were selected for higher activity towards 2-tert- butylphenol and guaiacol, were characterized with respect to their catalytic properties and substrate specificity. Mutations V368A/L417F changed the substrate reactivity of HbpA for the hydroxylation of different 2-substituted phenols, whereas mutation I244V reduced the substrate spectrum but
increased the turnover rate. Interestingly, HbpAG1 (I244V) showed an increased
activity with guaiacol but not towards salicylaldehyde, whereas HbpAT2
62 CHAPTER II
(V368A/L417F) showed doubled activity towards both phenols. This suggests that the substrate side chain causes mostly steric rather than inductive effects. This conclusion is supported by the results obtained with 2-sec-butylphenol. This substrate has a flexible side chain, which can move freely in several
directions. This results in an approximately equal activity and K'm values for the wild-type enzyme and all mutants.
Most enzymes in nature do not function under Vmax conditions, but catalyze [ ] reactions at S /Km ratios between 0.01 and 0.1 (Stryer, 1981). The catalytic
efficiency under these conditions is described by the ratio kcat/Km. This ratio was
determined for HbpA and its variants. K'm values for all mutant enzymes were decreased compared to wild-type HbpA towards guaiacol but increased for the
physiological substrate 2-hydroxybiphenyl. This was even the case for HbpAG1, which has an increased activity towards 2-hydroxybiphenyl. This suggests that
evolutionary advantage can be more easily achieved with a low Km rather than a high activity, probably because environmental substrate concentrations are low.
A low Km for the substrate of interest is also important for the biotechnological production of 3-substituted catechols with HbpA variants. Substituted phenols as well as the catechols are highly bactericidal and inactivate the whole cell biocatalyst. However, this problem can be solved by processes with integrated in situ product recovery or two-liquid-phase bioconversions (Held et al., 1999; Lye & Woodley, 1999; Witholt et al., 1990). These processes are based on the principle that both substrate and product are present at low concentrations in the aqueous phase of the bioreactor. Therefore, a desired enzyme feature for these processes is a high activity at the lowest possible substrate concentration,
i.e., a low Km.
Location of mutations in the HbpA model Lacking information on the three-dimensional structure of HbpA, an interpretation of the structural effects of the amino acid substitutions obained in the mutagenesis experiments is speculative. However, structure homology modeling with PHHY allows some statements on the effects of the modifications
63 DIRECTED EVOLUTION OF 2-HYDROXYBIPHENYL 3-MONOOXYGENASE
in the HbpA variants. Interestingly, Ile244 of HbpA corresponds with Tyr289 of PHHY (Enroth et al., 1998) and Tyr222 of p-hydroxybenzoate hydroxylase (Gatti et al., 1994; Schreuder et al., 1994). In p-hydroxybenzoate hydroxylase, Tyr222 interacts with the carboxylic moiety of the substrate and is critically involved in closing the active site allowing efficient substrate hydroxylation (for a review see (Entsch & van Berkel, 1995)). In PHHY, Tyr289 has been shown to play an important role in orienting the substrate for attack at the ortho position by forming a hydrogen bond with the hydroxyl moiety of the phenol. Moreover, Tyr289 favors the flavin out rather than the flavin in position through formation of a hydrogen bond between the N-3 of the isoalloxazine ring and the phenolic hydroxyl group (Xu et al., 2001). In HbpA, I244 clearly cannot fulfill a similar function. However, its position close to the side chain of the phenolic substrate suggests a direct influence on the shape of the substrate binding pocket (Fig. 7). From the above considerations and the catalytic properties of the I244V variant we suggest that substitution of Ile244 by Val in HbpA has a small but significant effect on substrate binding due to the reduced size of the side chain.
Fig. 7: Model of the substrate binding pocket of HbpA. Highlighted are the FAD prosthetic group, the substrate 2-hydroxybiphenyl, and amino acids His48 and Ile244.
64 CHAPTER II
Although the mode of binding of 2-hydroxybiphenyl in HbpA is unknown, it is reasonable to assume that it resembles the mode of binding of phenol in PHHY. This assumption is based on the conserved mode of binding of the FAD, the similar hydrophobic nature of the substrate binding pockets and the fact that the active site base Asp54 of PHHY is replaced by His48 in HbpA (Fig. 7). This histidine is likely to play an essential role in activating the 2-hydroxybiphenyl molecule prior to the regiospecific electrophilic attack by the flavin (C4a)-hydroperoxide at the 3-position of the phenolic ring.
Uncoupling of NADH oxidation from substrate hydroxylation A common feature among flavoprotein aromatic hydroxylases is the uncoupling of substrate hydroxylation from NADH oxidation with the concomitant formation of hydrogen peroxide (Massey & Hemmerich, 1975). This is also observed during hydroxylation of 2-hydroxybiphenyl with HbpA
(Suske et al., 1997). Mutation I244V in HbpAG1 had no significant influence on the degree of uncoupling compared to the wild-type enzyme, but the mutations
V368A/L417V in HbpAT2 decreased the uncoupling with all substrates. This is a remarkable result because both these substitutions are located far away from
the substrate binding site. From the properties of the single mutant HbpAT1 it can be concluded that the improvement of the efficiency of hydroxylation is related to the V368A substitution. This amino acid is located in the FAD binding domain which suggests an effect on the mobility of the flavin ring. Whether this results in a stabilization and/or improved positioning of the flavin (C4a)-hydroperoxide towards the substrate remains to be investigated. The mutation L417V is located in the third domain which in the case of PHHY is thought to be involved in subunit interactions (Enroth et al., 1998). The increased hydroxylation efficiency due to substitution V368A remained, but the activity towards 2-hydroxybiphenyl decreased significantly. In combination with the increased
K'm and higher k'cat for guaiacol we conclude that the altered catalytic properties of L417V are due mainly to altered substrate binding. Detailed structural information will be necessary to understand how the substitutions effect this
65 DIRECTED EVOLUTION OF 2-HYDROXYBIPHENYL 3-MONOOXYGENASE
change. Clearly, given their location in HbpA, substitutions V368A and L417V would not have been obvious targets for rational protein design.
For wild-type HbpA and HbpAG1 with 2-hydroxybiphenyl as the substrate, more than 50% of uncoupling could be ascribed to the reaction product 2,3-dihydroxybiphenyl. This suggests that 2,3-dihydroxybiphenyl competes with 2-hydroxybiphenyl for binding to the reduced enzyme and induces the non- productive heterolytic cleavage of the flavin (C4a)-hydroperoxide. This interpretation is supported by results from rapid reaction kinetics studies (Suske et al., 1999). In contrast, 3-sec-butylcatechol and 3-methoxycatechol have only
minor effects on the total uncoupling of wild-type HbpA and HbpAG1, indicating that these aromatic products do not interact strongly with the reduced enzyme. Flavoprotein aromatic hydroxylases such as p-hydroxybenzoate hydroxylase have a mechanism to decrease the rate of flavin reduction by several orders of magnitude in the absence of an aromatic substrate, thereby preventing the wasteful consumption of NAD(P)H (Howell et al., 1972; Husain & Massey, 1979; Massey, 1994). Other flavin enzymes such as 4-hydroxyphenylacetate 3-hydroxylase, PHHY and HbpA are less efficient in this respect and show some residual NAD(P)H oxidation (Arunachalam et al., 1994; Detmer & Massey, 1984; Suske et al., 1997). For HbpA it has been shown by stopped- flow absorption spectroscopy that flavin reduction is the rate-limiting step in this NADH oxidation (Suske et al., 1999). An increased NADH oxidase activity of the substrate-free enzyme, as determined for all HbpA-mutants, is therefore most likely related to an increase in the rate of flavin reduction. In conclusion, we have shown in this paper that the catalytic properties and substrate reactivity of HbpA can be improved by random mutagenesis. This is the first successful modification of a flavin-dependent monooxygenase by molecular evolution. We expect the mutants to be useful in new biocatalytic processes and the insights obtained to be helpful in further investigations of structure-function relationships in flavin monooxygenases.
66 CHAPTER II
ACKNOWLEDGEMENTS
This work was supported by the Swiss National Science Foundation grant number 5002-046098.
67
CHAPTER III
CHAPTER III:
SYNTHESIS OF 3-TERT-BUTYLCATECHOL BY AN
ENGINEERED MONOOXYGENASE
ANDREAS MEYER, MARTIN HELD, ANDREAS SCHMID, HANS-PETER E. KOHLER,
AND BERNARD WITHOLT
BIOTECHNOLOGY AND BIOENGINEERING, 2002, IN PRESS
69 BIOCATALYTIC SYNTHESIS OF 3-TERT-BUTYLCATECHOL
SUMMARY
Recombinant Escherichia coli JM101 was used for the in vivo biocatalytic synthesis of 3-tert-butylcatechol. The bacterial strain synthesized the laboratory-
evolved variant HbpAT2 of 2-hydroxybiphenyl 3-monooxygenase (HbpA, EC
1.14.13.44) from Pseudomonas azelaica HBP1. The mutant enzyme HbpAT2 is able to hydroxylate 2-tert-butylphenol to the corresponding catechol, a reaction which is not catalyzed by the wild type enzyme. The biotransformation was performed in a 3-L bioreactor for 24 hours. To mitigate the toxicity of the starting material 2-tert-butylphenol we applied a limited substrate feed. Continuous in situ product removal with the hydrophobic resin Amberlite™ XAD-4 was used to separate the product from culture broth. In addition, binding to the resin stabilized the product, which was important because 3-tert-butylcatechol is very labile in aqueous solution. The productivity of the process was 63 mg L-1 h-1 so that after 24 hours 3.0 g 3-tert-butylcatechol were isolated. Down-stream processing consisted of two steps. First, bound 2-tert-butylphenol and 3-tert-butylcatechol were eluted from Amberlite™ XAD-4 with methanol. Second, the two compounds were separated over neutral aluminum oxide, which selectively binds the catechol but not the phenol substrate. The final purity of 3-tert-butylcatechol was greater than 98%.
70 CHAPTER III
INTRODUCTION
Alkylated catechols are used as raw materials for the synthesis of pharmaceuticals, agricultural chemicals, dye developers, and polymerization inhibitors. The industrial route to these compounds proceeds via electrophilic
substitution, where the metal halides AlCl3, FeCl3, and ZnCl2 serve as Lewis acid catalysts (Wade, 1995). The problems of safety and waste disposal associated with these processes have stimulated the development of alternative routes. Newer approaches include the use of heterogeneous catalysts such as silica gel or acidic zeolites (Kamitori et al., 1984; Yoo et al., 1999). However, these chemical reactions preferentially yield the 4-substituted or polyalkylated isomers; 3-alkylcatechols are notoriously difficult to obtain. An effective alternate route to 3-n-alkylcatechols proceeds via intramolecular cyclization of 2-alkanoyl-2,5-dimethoxytetrahydrofurans with aqueous acid (Miyakoshi & Togashi, 1990), but it is not applicable to the synthesis of 3-sec- or 3-tert-alkylcatechols. Due to their high selectivity, biological catalysts offer a potent and environmentally friendly alternative to the chemical production of 3-substituted catechols (Held et al., 1999; Robinson et al., 1992). The corresponding benzenes or 2-substituted phenols, which are readily available, serve as starting materials and are transformed to the desired products by regioselective oxidation with a mono- or dioxygenase. Although this approach has been successful for the synthesis of different alkylated, halogenated and aryl- substituted catechols (Held et al., 1998; Johnston & Renganathan, 1987; Robinson et al., 1992; Schmid et al., 1998a), it has not been useful for the synthesis of 3-tert-butylcatechol. The reason for this is the lack of available enzymes with corresponding activities. Tuning the activity of a biocatalyst to a selected activity is a major topic in molecular biology. Both rational design by site-directed mutagenesis and random approaches, such as DNA shuffling or error prone PCR followed by screening for appropriate modifications (Moore & Arnold, 1996; Stemmer,
71 BIOCATALYTIC SYNTHESIS OF 3-TERT-BUTYLCATECHOL
1994), have been used to change and tune the substrate specificity of biocatalysts (Graf et al., 1987; Schmidt-Dannert & Arnold, 1998; Shao & Arnold, 1996; van den Heuvel et al., 2000a; Zhao & Arnold, 1999). Since we have been interested in developing biocatalysts for the production of various interesting and synthetically challenging target catechols by biooxidation of the corresponding phenols, we have used directed enzyme evolution to modify 2-hydroxybiphenyl 3-monooxygenase (HbpA, EC 1.14.13.44) (Meyer et al., 2002) of Pseudomonas azelaica HBP1 (Kohler et al., 1988). HbpA is a homotetrameric, NADH-dependent flavoprotein aromatic hydroxylase (Suske et al., 1997). We used in vitro manganese mutagenesis to generate HbpA variants and screened for improved monooxygenase activity on various 2-substituted phenols by observing the formation of colors indicative of the autooxidation of the reaction products (Meyer et al., 2002). We characterized several such
mutants. One of these, which we denoted HbpAT2, contained the two amino acid
-1 substitutions Val368Ala and Leu417Phe. HbpAT2 has a kcat of 0.5 s with
-1 2-tert-butylphenol as the substrate, whereas kcat <0.1 s for the wild-type protein
(Meyer et al., 2002). Here we report the application of HbpAT2 for the synthesis of 3-tert-butylcatechol using a recombinant whole cell biocatalyst. Due to the bactericidal properties of both substrate and product, an integrated process with limited starting material feed and in situ product removal was used (Held et al., 1999; Held et al., 1998; Lye & Woodley, 1999).
MATERIALS AND METHODS
Chemicals and media components 2-tert-Butylphenol, Amberlite™ XAD-4, aluminum oxide and all other chemicals were purchased from Fluka AG (Buchs, Switzerland). Solvents were obtained from Biosolve Ltd (Valkenswaard, The Netherlands) and components for complex media were obtained from Difco Laboratories (Detroit, USA).
72 CHAPTER III
Strains and plasmids Escherichia coli JM101 (F’, traD36, lacIq, ∆(lacZ)M15, proAB/supE, thi, ∆(lac- proAB)) (Yanish-Perron et al., 1985) containing plasmid pAMT2 was used as a biocatalyst. pAMT2 is a pUC18 (Yanish-Perron et al., 1985) derivative harboring
the hbpAT2 gene as a SalI/NsiI fragment under control of the lacZ promoter (Meyer et al., 2002b).
Media and growth conditions E. coli JM101 or recombinants thereof were usually stored as stab cultures and initially grown as 5 mL LB preculture (Sambrook et al., 1989). For assays or biotransformations such cultures were inoculated 1:100 in M9 medium (Sambrook et al., 1989), supplemented with 0.001% (w/v) thiamin, 0.1% (v/v)
MT trace element solution (Lageveen et al., 1988), 0.1 mM CaCl2, 2 mM
MgSO4, and 0.5 % (w/v) glucose or 0.5% (v/v) glycerol as carbon source. For recombinants 150 mg L-1 ampicillin was added to ensure pAMT2 maintenance. Cultivations were carried out at 30°C and biomass concentration was determined at 450 nm (Witholt, 1972).
Analytical methods 2-tert-Butylphenol and 3-tert-butylcatechol were quantified with reverse phase HPLC (Hewlett Packard HP 1050 Ti HPLC equipped with a diode array detector DAD 1040 M) using a Hypersil ODS column (5 µm, 4.5 x 125 mm), as described elsewhere (Held et al., 1999). Samples, which were freed from solids by centrifugation, were diluted 1:1 with methanol that was acidified with 0.1% (v/v) phosphoric acid. Elution was carried out with a flow rate of 1.5 mL min-1 under isocratic conditions with a methanol to water ratio of 40:60, also
containing 0.1% (v/v) H3PO4.
Biotransformation The biotransformation experiment was performed in a 3-L bioreactor from Bioengineering AG (Wald, Switzerland) connected to an external loop for in situ product extraction (Fig. 1). Cell growth and biotransformation were time-
73 BIOCATALYTIC SYNTHESIS OF 3-TERT-BUTYLCATECHOL
separated to optimize cultivation and production independently (Held et al., 1999). Cell cultivation. The preculture was grown overnight (o/n) in M9 medium (see growth conditions) with 0.5% (w/v) glucose as carbon source. Thereafter, the bioreactor containing 2 L M9 medium and 1% (w/v) glucose was inoculated with 50 mL of preculture and the cells were grown o/n with a stirrer speed of 1500 rpm and aeration by pressurized air (1 L min-1). Two hours prior to biotransformation experiments the culture was pulsed with 0.5% (v/v) glycerol and 150 mg L-1 ampicillin and the stirrer speed was increased to 3000 rpm. Preparation of product extraction module. Due to the bactericidal properties of substrate and product and the instability of the formed 3-tert-butylcatechol, the biotransformation was carried out with limited substrate feed and in situ product removal (Held et al., 1999). The extraction module consisted of a cylindrical external loop (diameter 8 cm, length 18 cm) filled with 390 g Amberlite™ XAD-4, a polystyrene-based hydrophobic resin. The loop and the tubing were sterilized by flushing with 70% ethanol for 24 hours. After washing with sterile water the module was filled with sterile M9 medium and connected to the reactor. Biotransformation. 1 hour before starting the biotransformation the content of the bioreactor was circulated (Fig. 1, P2) through to the external loop with a flux of 400 mL min-1. Thereafter, 2-tert-butylphenol was fed (Fig. 1, P1) at a rate of 0.15 g L-1 h-1 from a 2 M stock solution in methanol. When the dissolved oxygen concentration in the bioreactor increased above 80% DOT, indicating carbon source depletion, 50 mL of 30% (v/v) glycerol were added.
Fig. 1: Schematic representation of the experimental set-up. A product recombinant E. coli JM101 recovery loop was filled with product (pAMT2) extraction Amberlite™ XAD-4 and connected to a module 3-L bioreactor. Substrate was added by
Amberlite XAD-4 starting material feed 0.15 g L-1 h-1 a high-precision pump (P1) and the P1 P2 fermentation broth circulated (P2)
0.4 L min-1 through the extraction module. 74 CHAPTER III
Product purification Elution from Amberlite™ XAD-4. After biotransformation the extraction loop was removed from the reactor and the Amberlite™ XAD-4 washed with distilled water. Subsequently, the resin was transferred to a glass column and bound compounds were eluted with methanol (0.1% HCl) and collected in 20 fractions of 100 mL. Acidification to pH 2 was necessary to stabilize the catechol product (Held et al., 1999). After HPLC analysis, fractions containing product were pooled and filtered through a 2 µm nitrocellulose filter. Finally, the methanol was removed in a rotary evaporator. Reverse phase chromatography. Reverse phase chromatography was performed by preparative HPLC (Hewlett Packard HP 1050Ti) with C8-silica as solid phase and 40% MeOH (0.1% trifluoroacetic acid) as mobile phase. Eluted compounds were detected with a diode array detector (Hewlett Packard HP DAD 1040M). Aluminum oxide column. The fact that aluminum oxide efficiently binds catechols but not phenols (Pras et al., 1990; Schmid et al., 2001) was used to separate 3-tert-butylcatechol and 2-tert-butylphenol. The binding capacity was evaluated with two model compounds. Per gram of aluminum oxide 91 µmol of
catechol (Mw 110) and 107 µmol 2,3-dihydroxybiphenyl (Mw 186) were bound. The capacity for 3-tert-butylcatechol was therefore assumed to be approximately 100 µmol g-1. A glass column was filled with activated aluminum oxide (Type 507 C neutral, size: 100-125 mesh) and washed with hexane. An aliquot of the compounds eluted from XAD-4 was loaded onto the column bed and unbound substances were washed out with methanol. 3-tert-Butylcatechol was eluted with 10% 5 M HCl in methanol. Fractions of 15 mL were collected and analyzed with reverse phase HPLC.
75 BIOCATALYTIC SYNTHESIS OF 3-TERT-BUTYLCATECHOL
RESULTS
Growth inhibition of E. coli JM101 by 2-tert-butylphenol Substituted phenols generally have antimicrobial properties (Davidson & Brandon, 1981). The effect of 2-tert-butylphenol on the growth of E. coli JM101 (pUC18) was determined in 250 mL shaking flasks filled with 50 mL M9 medium and 0.5% (w/v) glucose as carbon source. The cells were inoculated to a biomass concentration of 20 mg L-1. After 6 hours the cultures had reached the early exponential phase and 2-tert-butylphenol was added in concentrations ranging from 0 to 1.5 mM. Significant growth inhibition was observed at all concentrations above 0.5 mM (Fig. 2). When the added 2-tert-butylphenol exceeded 1 mM partial cell lysis could be observed and substantially lower cell concentrations were reached after incubation overnight. At concentrations above 1.5 mM, no further growth was observed.
10 Fig. 2: Growth of E. coli JM101
2-TBP [mM] (pUC18) in the presence of
L L 0 different concentrations of 1 L L 0.1 2-tert-butylphenol. The arrow L 0.5 L 1.0 indicates the addition of
N CDW [g/L] 0,1 2-tert-butylphenol (2TBP) in L 1.5 concentrations of 0 (), 0.1 (L), 0.5 (G), 1 (I), and 1.5 mM (N). 0,01 CDW, cell dry weight. 0 200 400 600 800 1000 time [min]
HbpA activity in the presence of 2-tert-butylphenol The formation of 3-tert-butylcatechol by E. coli JM101 (pAMT2) was determined in 250 mL shaking flasks with 50 mL M9 medium and 0.5% (w/v)
glucose as carbon source. HbpAT2 synthesis was induced in the early exponential phase by the addition of 0.2 mM IPTG. 2 hours later 2-tert-butylphenol was added in concentrations of 0 to 1.5 mM and the cultures were incubated overnight at 30°C. The instability of 3-tert-butylcatechol, which forms a dark polymer in aqueous solution at neutral pH, was used to identify
76 CHAPTER III
active cell cultures. Product formation activity could be observed up to concentrations of 1 mM (Fig. 3).
Biotransformation of 2-tert-butylphenol to 3-tert-butylcatechol E. coli JM101 (pAMT2) was grown overnight on M9 medium with 1% glucose as carbon source as described in Materials and Methods. This led to a final
-1 biomass concentration of 3.2 g CDW L and no significant hbpAT2 expression. After pulsing with 0.5% glycerol the dissolved oxygen tension (DOT) immediately decreased, indicating a viable and growing cell culture. In addition,
changing the carbon source induced hbpAT2 expression indicating that catabolite repression of the lacZ promoter was reduced or eliminated. The content of the reactor was pumped through the extraction module and 1 hour later the substrate feed was switched on. The amount of 2-tert-butylphenol and 3-tert-butylcatechol in the reaction vessel was analyzed by HPLC and never exceeded 6 mg L-1 for the substrate and 3 mg L-1 for the product. Neither substrate nor product was detected in the reflux from the external loop, indicating complete extraction of these compounds. After 24 hours a total of 7.5 g 2-tert-butylphenol had been added and the substrate feed was switched off. The culture broth was circulated through the extraction loop for another 10 min. At the end of the biotransformation the biomass concentration was 7.6 g CDW L-1.
3-tert-Butylcatechol recovery from XAD-4 Amberlite™ XAD-4 has no effect on the growth of recombinant E. coli JM101
or on HbpAT2 activity and can be used efficiently for the adsorption of hydrophobic compounds from culture broth (Held, 2000). After biotransformation the extraction loop was disconnected from the reactor, the resin washed with water and the bound compounds eluted with 2 L methanol (0.1% HCl). A total of 4.6 g 2-tert-butylphenol and 3.0 g 3-tert-butylcatechol were collected, which corresponds to a conversion yield of 36.8% and a product space time yield of 0.063 g L-1 h-1. Substrate and product were present in similar ratios in all fractions (Fig. 4). The mass balance revealed a molar recovery of
77 BIOCATALYTIC SYNTHESIS OF 3-TERT-BUTYLCATECHOL
97% (substrate plus product), which indicated that no significant product polymerization took place. After filtration and methanol removal 45 mL of brownish oil remained.
Fig. 3: Cultures of E. coli JM101 (pAMT2) grown in the presence of different concentrations of 2-tert-butylcatechol. The dark color derives from the polymerization of 3-tert-butylcatechol, which is formed by active cultures.
25 Fig. 4: Elution of 2-tert-butylphenol and 3TBC 3-tert-butylcatechol from XAD-4. After 20 2TBP biotransformation the formed 3-tert-butylcatechol 15 and the remaining substrate were eluted from the
10 hydrophobic resin XAD-4 (390 g) and collected in % eluted 20 fractions of 100 mL. 2TBP, 2-tert-butylphenol; 5 3TBC, 3-tert-butylcatechol.
0 1 35791113151719 fraction
Separation of 3-tert-butylcatechol from 2-tert-butylphenol To separate remaining substrate and product, two different approaches were used: reverse phase chromatography and separation on neutral aluminum oxide (pH 7). Reverse phase chromatography. A 1 mL sample containing 40 g L-1 3-tert-butylcatechol and 61 g L-1 2-tert-butylphenol was loaded and eluted under
78 CHAPTER III
isocratic conditions. Retention times were 18-25 min for the catechol and more than 28 min for the phenol. All the 3-tert-butylcatechol was collected in a volume of 35 mL. Separation on aluminum oxide. A 2 mL sample containing 80 g L-1 3-tert-butylcatechol and 122 g L-1 2-tert-butylphenol was loaded onto neutral aluminum oxide. The column was flushed with methanol and fractions of 15 ml were collected. 2-tert-Butylphenol eluted in fractions 2 to 5 (Fig. 5). When no more phenol was detected in the effluent, the solvent was changed to 10% 5 M HCl in methanol. The 3-tert-butylcatechol that subsequently came off the column was collected in fractions 7 to 11 (Fig. 5). The mass balance revealed a 3-tert-butylcatechol yield of 84%. Purity was checked with reverse phase HPLC and determined to be greater than 98%.
100 Fig. 5: Separation of 2-tert-butyl-phenol 3TBC 80 2TBP and 3-tert-butylcatechol over neutral aluminum oxide. Neutral aluminum oxide 60 was used to separate the formed 3-tert-
40 butyl-catechol from remaining substrate. % eluted 2TBP, 2-tert-butylphenol; 3TBC, 3-tert- 20 butylcatechol.
0 1234567891011 fraction 3-tert-Butylcatechol authentication For verification of the authenticity of 3-tert-butylcatechol (Fig. 5) 1H- and 13C-NMR spectra of the product purified by HPLC were recorded. 1H-NMR
(CDCl3): 6.81 (d, Cf-H); 6.98 (dd, Ce-H); 6.82 (d, Cd-H); 1.52 (s, Ch-H); 5.01 (br,
13 1 OH); 5.70 (br, 1 OH). C-NMR (CDCl3): 137.1 (Ca); 143.2 (Cb); 143.8 (Cc);
113.4 (Cd); 119.5 (Ce); 119.8 (Cf); 35.0 (Cg); 29.9 (Ch). The measured spectra are in agreement with earlier data (Kamitori et al., 1984; Yoo et al., 1999).
h H3C h H3C CH3 Fig. 6: 3-tert-Butylcatechol. Chemical structure of h g a 3-tert-butylcatechol with assigned nomenclature for H f OH b 1H-NMR- and 13C-NMR-spectra. e c H d OH 79 H BIOCATALYTIC SYNTHESIS OF 3-TERT-BUTYLCATECHOL
DISCUSSION
Advances in protein engineering, especially approaches such as directed enzyme evolution, significantly enlarge the scope of biocatalytic synthesis. In this study, we described the gram scale production of 3-tert-butylcatechol by a laboratory-evolved monooxygenase.
Experimental set-up Phenolic compounds permeabilize bacterial membranes explaining the antimicrobial properties of these compounds (Davidson & Brandon, 1981). The addition of 2-tert-butylphenol in concentrations of 1.5 mM was toxic for E. coli JM101 cells and led to complete cell lysis. A limited substrate feed was therefore necessary for bioconversion of this substrate. An alternative would have been the application of cell free preparations of the mutant monooxygenase with an enzymatic or electrochemical regeneration system for NADH (Hollmann et al., 2001; Kragl et al., 1996; Kula & Wandrey, 1987). Such approaches have been used successfully using dehydrogenases in enzyme membrane reactors. However, the application of oxygenases, which constitute a more complex class of enzymes, has led to only few practical examples due to the limited stability of the enzymes under in vitro process conditions (Duetz et
al., 2001b). In contrast, recombinant E. coli JM101 synthesizing HbpAT2 showed in vivo activity for up to 24 hours without any loss. The instability of 3-tert-butylcatechol at neutral pH required the removal of the product from culture broth. This was achieved with Amberlite™ XAD-4, which has already been used for in situ product recovery of several substituted catechols (Held et al., 1999; Held et al., 1998).
3-tert-Butylcatechol synthesis The productivity of the process with the mutant 2-hydroxybiphenyl 3-monooxygenase was only 6 fold lower than the corresponding optimized biotransformation of 2-hydroxybiphenyl with the wild-type enzyme (Held et al., 1999). Considering that the specific activity towards 2-tert-butylphenol determined for the purified enzyme is only 4% of the value for HbpA and the
80 CHAPTER III
natural substrate, the observed productivity is unexpectedly high and significantly higher than for chemical catalysis (Kamitori et al., 1984; Yoo et al., 1999). We expect that either biocatalyst engineering or process optimization could increase the productivity further. The 2-tert-butylphenol conversion yield was determined to be 36.8% and could easily be raised by simply reducing the substrate feed rate. For synthesis of 2,3-dihydroxybiphenyl, yields up to 97% could be reached in this way. Yields for chemical catalysis were considerably lower. Synthesis with silica gel as the catalyst reached conversions of 8% (Kamitori et al., 1984), whereas in the best reaction set-up with acidic zeolites as the catalyst, yields of 3.1% were obtained (Yoo et al., 1999). A reason for the low chemical synthesis yields is the formation of side products, since the main product of chemical catechol alkylation is the 4-substituted isomer (Kamitori et al., 1984; Yoo et al., 1999). Side product formation was not observed for the biocatalytic production of
3-tert-butylcatechol. As is true for the wild type enzyme, HbpAT2 hydroxylates 2-substituted phenols in an absolutely regiospecific manner. This fact was also useful for the development of an easy down-stream processing based on the catechol functionality of the synthesized product.
Product purification Continuous extraction of 3-tert-butylcatechol formed in the reactor with the hydrophobic resin Amberlite™ XAD-4 was the first step of the downstream processing. Both separation and stabilization were in fact achieved and, after elution, more than 94% of the 3-tert-butylcatechol was recovered in 1 liter of methanol. In comparison to a two liquid phase process of the same scale, in which the second phase serves as the extraction medium, the product containing volumes to be processed are comparable. However, stabilization of catechol occurred only when it was immobilized on a solid matrix. The separation of 3-tert-butylcatechol from 2-tert-butylphenol by reverse phase HPLC led to a highly pure product preparation, as could be seen from the 1H- and 13C-NMR spectra. However, HPLC methods are of limited capacity and
81 BIOCATALYTIC SYNTHESIS OF 3-TERT-BUTYLCATECHOL
therefore difficult to apply on larger scales. A better option is the separation of the product on aluminum oxide, which selectively binds catechols but not phenols. The capacity of the matrix is determined by catechol functionalities rather than by molecular weight as shown by immobilization experiments with model compounds. Because the amount absorbed at pH 7 is about 4 times higher than at pH 4 (Held, 2000) the separation was carried out at neutral pH. As a consequence, part of the 3-tert-butylcatechol polymerized (as indicated by brownish color formation on the column) and was not eluted. This resulted in a final yield of 84% for product purification. The small amount of polymerization was deemed acceptable compared to the benefits of the reduced column volume. In this paper we have shown that the application of a modified 2-hydroxybiphenyl 3-monooxygenase, in combination with in situ product recovery and product purification over aluminum oxide, allows the efficient synthesis of 3-tert-butylcatechol. The approach described here provides a general procedure for the production of a broad variety of 3-substituted catechols by engineered HbpA.
ACKNOWLEDGEMENTS
We want to thank Carsten Boehler (ETH Zurich) for recording the NMR spectra and the Swiss National Science Foundation for financial support (grant number 5002-046098).
82 CHAPTER IV
CHAPTER IV:
HYDROXYLATION OF INDOLE BY LABORATORY EVOLVED
2-HYDROXYBIPHENYL 3-MONOOXYGENASE
ANDREAS MEYER, MICHAEL WÜRSTEN, ANDREAS SCHMID, HANS-PETER E. KOHLER,
AND BERNARD WITHOLT
JOURNAL OF BIOLOGICAL CHEMISTRY, 2002, 277(37): 34161-34167
83 INDOLE HYDROXYLATION BY LABORATORY EVOLVED HBPA
SUMMARY
Directed enzyme evolution of 2-hydroxybiphenyl 3-monooxygenase (HbpA, EC 1.14.13.44) from Pseudomonas azelaica HBP1 resulted in an enzyme
variant (HbpAind) which hydroxylates indole and indole derivatives, such as hydroxyindoles or 5-bromoindole. The wild-type protein does not catalyze these
reactions. HbpAind contains amino acid substitutions Asp222Val and Val368Ala. The activity towards indole hydroxylation was increased 18 fold in this variant.
Concomitantly, the Kd value towards indole decreased from 1.5 mM to 78 µM.
Investigation of the major reaction products of HbpAind with indole revealed hydroxylation at the carbons of the pyrrole ring of the substrate. Subsequent enzyme independent condensation and oxidation of the reaction products lead to the formation of indigo and indirubin.
The activity of the HbpAind mutant monooxygenase towards the natural substrate 2-hydroxybiphenyl was 6 times lower than that of the wild-type
enzyme. In HbpAind, there was significantly increased uncoupling of NADH oxidation from 2-hydroxybiphenyl hydroxylation, which could be attributed to the substitution Asp222Val. The position of Asp222 in HbpA, the chemical properties of this residue, and the effects of its substitution indicate that Asp222 is involved in substrate activation in HbpA.
84 CHAPTER IV
INTRODUCTION
Indole is produced from the aromatic amino acid tryptophane in tryptophanase synthesizing bacteria, such as Escherichia coli (de Moss & Moser, 1969). Enzymes that oxygenate the indole pyrrole ring are easily detectable, because the reaction products are unstable and form pigments. This observation was first made when the genes responsible for naphthalene oxidation were expressed in E. coli, which resulted in the biosynthesis of indigo (Ensley et al., 1983). Based on these results, and because of its importance as a dye, the biocatalytic production of indigo by naphthalene dioxygenase was a major goal of the biotech industry for some time (Bialy, 1997; Murdock et al., 1993). Naphthalene dioxygenase was soon found not to be the only enzyme capable of indole oxidation: several other oxygenases that accept indole as a substrate have been identified (Bhushan et al., 2000; Gillam et al., 2000; Hart et al., 1992; O'Connor et al., 1997; O'Connor & Hartmans, 1998; Panke et al., 1998). These are either enzymes similar to naphthalene dioxygenase that activate oxygen with iron centers or members of the cytochrome P450 family (Ensley & Gibson, 1983; Gillam et al., 2000). Although flavin nucleotides may be involved in electron transfer from cofactors in these proteins, none is known to be a flavoprotein oxygenase. This situation has changed with our recent finding that flavin-dependent oxygenases can be modified to accept unnatural substrates. 2-Hydroxybiphenyl 3-monooxygenase (HbpA, EC 1.14.13.44) from Pseudomonas azelaica HBP1 is a flavoprotein aromatic hydroxylase that catalyzes the hydroxylation of a variety of 2-substituted phenols to the corresponding catechols (Eppink et al., 1997; Kohler et al., 1988; Suske et al., 1997). The mechanism of HbpA has been extensively studied by spectroscopic techniques, which revealed that molecular oxygen is activated via the formation of a flavin (C4a)-hydroperoxide (Suske et al., 1999), a common intermediate in the reaction cycle of this enzyme family (van Berkel et al., 1997). HbpA has a broad substrate spectrum but does not
85 INDOLE HYDROXYLATION BY LABORATORY EVOLVED HBPA
hydroxylate indole (Held et al., 1998; Kohler et al., 1988). Recently, we changed the substrate reactivity of HbpA by directed enzyme evolution towards 2-tert-butylphenol, a substrate which is not converted by the wild type enzyme (Meyer et al., 2002a; Meyer et al., 2002b). As a side product of this work, we
also obtained a HbpA variant, which we denoted HbpAind, with activity towards the hydroxylation of indole.
In this study we report on the characterization of HbpAind with respect to its catalytic properties. While previous work on indole oxygenating enzymes mainly aimed at the biotechnological production of indigo, we were especially interested in the formation of the byproduct indirubin. Indirubin and its analogues have been identified as potent inhibitors of cyclin-dependent kinases (CDK) (Hoessel et al., 1999). The crystal structure of CDK2 in complex with indirubin derivatives showed that indirubin binds to the kinase’s ATP binding site. As a consequence, it inhibits the proliferation of a wide range of cells and belongs to a group of novel anticancer compounds that act on the cell cycle (Buolamwini, 2000).
MATERIALS AND METHODS
Chemicals, strains, and plasmids Escherichia coli JM101 (Sambrook et al., 1989) and the pUC18 plasmid (Yanish-Perron et al., 1985) were used throughout for cloning and expression of the hbpA gene. Alkaline phosphatase (EC 3.1.3.1) was purchased from Roche Molecular Biochemicals (Basel, Switzerland). Catalase (EC 1.11.1.6) from beef liver and formate dehydrogenase (EC 1.2.1.2) from Candida boidinii were obtained from Fluka AG (Buchs, Switzerland). 4-Hydroxyindole and 5-hydroxyindole were from ICN Biomedicals Inc. (Aurora, USA). Components for complex media were obtained from Difco Laboratories (Detroit, USA). All other chemicals were of purest available quality and obtained from Fluka AG (Buchs, Switzerland).
86 CHAPTER IV
Directed evolution of HbpA Directed evolution of HbpA was performed by error prone PCR based on in vitro manganese mutagenesis as described earlier (Meyer et al., 2002b). The mutant library was subsequently plated onto LB medium. Cells harboring enzymes with activity towards the hydroxylation of indole formed deep blue colonies.
The single mutant D222V (HbpAD222V) was constructed using the QuickChangeTM site directed mutagenesis kit from Stratagene (La Jolla, USA).
Protein synthesis and purification
Synthesis of wild type HbpA and HbpAind was done in recombinant E. coli using M9 mineral medium and glycerol as carbon source (Held et al., 1998). After harvesting the cells the proteins were purified according to the method described recently (Meyer et al., 2002b).
Analytical methods Determination of activity towards 2-hydroxybiphenyl. The activity of wild type
HbpA and HbpAind was determined by measuring substrate consumption and product formation with reverse phase HPLC as described elsewhere (Kohler et al., 1993a). The assay contained 0.2 µM HbpA or variant protein, 0.3 mM NADH, 0.2 mM 2-hydroxybiphenyl, and 20 mM air saturated phosphate buffer pH 7.5. Determination of in vivo indigo formation. The activity of recombinant E. coli JM101 towards the formation of indigo was determined in 250 mL shaking flasks containing 50 mL LB medium (Sambrook et al., 1989). Cultures of E. coli
JM101 harboring a pUC18 derivative encoding HbpA or HbpAind were
inoculated to an OD450 of 0.1. The cultures were incubated at 30°C and vigorously shaken. When the culture color turned olive, samples of 1.1 mL were taken. 100 µl of these were used to determine the cell dry weight (CDW) at 450 nm (Witholt, 1972). The remaining 1 mL was centrifuged and the supernatant was carefully removed. Cell associated indigo was extracted with
87 INDOLE HYDROXYLATION BY LABORATORY EVOLVED HBPA
ε -1 -1 N,N-dimethylformamide (DMF) and quantified at 610 nm ( 610 15900 l mol cm ) (O'Connor & Hartmans, 1998). Determination of in vitro indigo formation. The activity towards indole was determined using an assay with NADH regeneration by formate dehydrogenase (EC 1.2.1.2, FDH) from Candida boidinii (Fig. 1) (Kula & Wandrey, 1987). The assay contained 0.2 µM HbpA or variant, 0.25 U FDH, 160 mM sodium formate, 10 U catalase (EC 1.11.1.6) from beef liver, 0.3 mM NADH, and 2 mM indole in 1 mL 50 mM sodium phosphate buffer pH 7.5. The assay was stopped by the addition of perchloric acid, and the precipitated proteins were spun down. The indigo in the pellet and in the tube was extracted with DMF and spectrophotometrically quantified. Determination of dissociation constants. Dissociation constants between the enzymes and indole were determined by monitoring the absorption changes of the enzyme bound FAD upon binding of substrate (van Berkel et al., 1992). For
this 12 µM purified wild type HbpA and HbpAind were titrated with known concentrations of 2-hydroxybiphenyl or indole and the resulting spectra were recorded using a Varian Cary E1 UV/Vis spectrophotometer. Plotting delta absorbance at a specific wavelength allowed the calculation of the dissociation constants by weighted non-linear regression analysis (Enzfitter, Elsevier- Biosoft, UK).
O2 H2O OH O H air oxidation N
N HbpA N N H H H O
NADH + H+ NAD+
FDH CO 2 HCOOH
Fig. 1: Scheme of in vitro indigo formation assay. HbpA, 2-hydroxybiphenyl
3-monooxygenase or HbpAind; FDH, formate dehydrogenase.
88 CHAPTER IV
High Pressure Liquid Chromatography-Mass Spectroscopy (HPLC-MS) analysis. Analysis of compounds formed during in vitro indigo assays was done with reverse phase HPLC-MS (Hewlett Packard, 1100 MSD). The compounds were separated with a Hypersil ODS column (5 µm, 4.5 x 125 mm) and detected with a diode array detector and a mass spectrometer. Acidified (0.1%
formic acid) H2O (solvent A) and 50% methanol/50% acetonitrile (solvent B) were applied as mobile phase according to the following timetable: 0 to 8 min, 85A/15B, flow 1 mL min-1; gradient to 10 min, to 65A/35B, flow 2 mL min-1; to 15 min, 65A/35B, flow 2 mL min-1. Standards for isatin, 4-hydroxyindole, 5-hydroxyindole, 2-indolinone, and indole were commercially available. 3-Indoxyl was prepared by dephosphorylating 3-indoxylphosphate with alkaline phosphatase (EC 1.3.1.3) under anaerobic conditions. HPLC-MS analysis of the formed pigments was done under isocratic conditions at a flow rate of 1 mL min-1 with 70% (v/v) methanol as mobile phase for the pigments derived from indole, and 40% (v/v) methanol for the one derived from 4- and 5-hydroxyindole. Thin Layer Chromatography (TLC) analysis. Pigments were analyzed by TLC using silica gel cards and either toluene-acetone (4:1) or chloroform-acetone (97:3) as the mobile phase (Eaton & Chapman, 1995). Electron microscopy. For ultrathin sectioning, cells were fixed in 2.5% glutaraldehyde for 60 min and subsequently washed with water and embedded
in low-melting-point agarose. After fixation in 1% OsO4 for 60 min the blocks were dehydrated with ethanol and acetone and embedded in Epon-Araldit (Hess, 1966). Sections cut from the Epon-Araldit preparation were contrasted with uranyl acetate and lead citrate. Freeze fracturing was carried out following standard procedures using a Balzers BAF 300 apparatus (Balzers-Union Inc., Balzers, Liechtenstein). The specimen sandwiches were fractured at -150°C and immediately replicated with platinum-carbon. All pictures were taken with a Philips EM301 electron microscope.
89 INDOLE HYDROXYLATION BY LABORATORY EVOLVED HBPA
RESULTS
Directed evolution of HbpA We recently changed the substrate reactivity of 2-hydroxybiphenyl 3-monooxygenase (HbpA) from Pseudomonas azelaica HBP1 by directed evolution (Meyer et al., 2002b). This work led to a mutant monooxygenase with increased activity towards the hydroxylation of indole. This HbpA variant was
denoted HbpAind. E. coli JM101 cultures synthesizing HbpAind turned deep blue when grown overnight on LB medium. Electron microscopy revealed the extracellular accumulation of material, which we believe, consists of the water insoluble pigment (Fig. 2). After centrifugation the pigment was extracted from the pellet with N,N-dimethylformamide (DMF). It was authenticated as indigo by thin layer chromatography (TLC) with toluene-acetone (4:1) as the mobile phase and commercially available indigo as standard. This analysis also
revealed the presence of a major byproduct. The Rf value of this red pigment
corresponded to the Rf value determined earlier for indirubin (Eaton & Chapman, 1995). Analysis by HPLC-MS with 70% (v/v) methanol as mobile phase showed two prominent molecular ion (MH+) peaks at m/z 263 with retention times of 3.7 and 4.2 min. The UV/Vis spectra and the fragmentation patterns were compared with literature data (Eaton & Chapman, 1995; Fearon & Boggust, 1950; Laatsch & Ludwig-Köhn, 1986), which confirmed that these two compounds were indigo (3.7 min) and indirubin (4.2 min). The formation of indigo by recombinant E. coli JM101 growing on LB medium
was quantified. Cultures expressing the hbpAind gene accumulated 150 µM indigo within 8 hours, while cultures of the host synthesizing HbpA remained colorless (Fig. 3). Recombinant protein levels in both cultures were checked by SDS-PAGE and HbpA levels were determined to be in the same range of about 20% of total cell protein.
90 CHAPTER IV
Fig. 2: Electron microscopy of E. coli synthesizing HbpAind grown on LB. The arrows point to needle shaped precipitates that are believed to consist of indigo (see text). Left, ultrathin sectioning; Right, freeze fracturing.
10 0,16 Fig. 3: In vivo indigo formation by
0,14 HbpA and HbpAind. Formation of indigo 0,12 during growth of recombinant E. coli 1 0,1 JM101 on LB broth synthesizing wild 0,08 type HbpA or HbpAind. Symbols for cell 0,06 0,1 indigo [mM]
CDW [mg/mL] dry weight (CDW): , wild-type HbpA; 0,04 , HbpAind. Symbols for indigo 0,02 concentration: I, wild-type HbpA; G, 0,01 0 200 250 300 350 400 450 HbpAind. time [min]
Stability of biotechnologically produced indigo When pigments were extracted with DMF from recombinant E. coli JM101 cultures, the extract had a deep blue color. The blue color disappeared upon storage at room temperature and the solution turned red. The red pigment was analyzed by UV/Vis spectroscopy, HPLC-MS, and TLC. It was authenticated as indirubin by comparison with literature data (Eaton & Chapman, 1995; Fearon & Boggust, 1950; Laatsch & Ludwig-Köhn, 1986). Buffering the pH at a value of 7
91 INDOLE HYDROXYLATION BY LABORATORY EVOLVED HBPA
or acidification with 0.1% (v/v) 10 M hydrochloric acid stabilized the formed indigo, whereas the addition of 0.1% (v/v) 10 M sodium hydroxide or heating accelerated the disappearance of the blue color.
General properties of HbpAind
The mutant monooxygenase HbpAind differs from wild-type HbpA by two amino acids: Asp222 was substituted by valine and Val368 was substituted by
alanine. HbpAind was purified according to the procedure developed for the wild- type enzyme with a yield of about 30%. Analytical size exclusion chromatography showed that the mutant monooxygenase formed a tetramer, which is also the case for wild-type HbpA (Suske et al., 1997).
Major reaction products of indole hydroxylation by HbpAind To identify the major reaction products of indole hydroxylation, in vitro indigo formation assays were performed. After 30 min a sample was collected and immediately saturated with argon. The proteins were precipitated and separated by centrifugation. The pigments were extracted from the pellet with DMF and analyzed by TLC. They were identified as indigo and indirubin. Analysis of the aqueous phase by HPLC-MS revealed the presence of 3-hydroxyindole (indoxyl) and 2-indolinone (oxindole). When a sample was taken after 60 min assay time, isatin was also detected (Table 1).
92 CHAPTER IV
Table 1: Major reaction products of indole hydroxylation by HbpAind. The major reaction products of in vitro indole assays were authenticated by reverse HPLC-MS. The assays
contained 0.2 µM HbpAind, 0.25 U formate dehydrogenase, 160 mM sodium formate, 10 U catalase, 0.3 mM NADH, and 2 mM indole in 1 mL 50 mM sodium phosphate buffer pH 7.5.
a λ Compound Retention time Structure UV/Vis: max m/z
min nm MH+
Isatin 5.5O 242, 302 148
O N H
3-Hydroxyindole 6.1OH 228, 381 134 (indoxyl)
N H
2-Indolinone 7.6 204, 249 134 (oxindole) O N H
Indole 12.1 217, 270 118
N H
a All compounds were compared to commercially obtained standards.
93 INDOLE HYDROXYLATION BY LABORATORY EVOLVED HBPA
Substrate spectrum of HbpAind
To investigate the substrate range of the HbpAind mutant monooxygenase, in vitro assays were performed with 4-hydroxyindole, 5-hydroxyindole, and 5-bromoindole. For the variant enzyme color formation could be observed with all substrates while the control assays with the wild-type protein remained colorless. The pigment derived from 4-hydroxyindole was purple, that from 5-hydroxyindole orange, and that from 5-bromoindole pink. The polar reaction products of the assays with 4- and 5-hydroxyindole were analyzed with HPLC-MS. For this the in vitro assay was stopped by the addition of perchloric acid and the proteins were spun down. Mass peaks (MH+) at m/z 150 were detected in the supernatants from both reactions. This mass correlates with the single hydroxylated substrates. The corresponding compounds eluted between 4.5 and 6.5 minutes when using 40% (v/v) methanol as mobile phase. The main condensation products had a prominent molecular ion peak (MH+) at m/z 279 and had a retention time of 4.1 min for the assay with 4-hydroxyindole and of 3.5 min for the assay with the 5-substituted isomer. UV/Vis spectra showed the peak at 4.1 min to have a maximum at 494 nm, whereas the peak at 3.5 min had a maximum at 480 nm.
Catalytic properties of HbpA and HbpAind Specific activities towards 2-hydroxybiphenyl and indole. The in vitro activity of the purified proteins towards the natural substrate 2-hydroxybiphenyl was determined by measuring substrate consumption and product formation by
reverse phase HPLC. The kcat of HbpAind was significantly lower than that of the wild type enzyme (Table 2). Indole hydroxylation activities were determined in assays with purified proteins and NADH regeneration by formate dehydrogenase from Candida boidinii. The assay mix containing the mutant monooxygenase showed a blue color within the first 30 min, whereas the assay mix with the wild type enzyme
remained white. HbpAind formed up to 170 µM indigo, whereas hardly any indigo formation could be observed for HbpA (Fig. 4). The indole hydroxylation activity
94 CHAPTER IV
-1 of HbpAind was approximately 20 mU mg purified protein or about 18 times higher than the corresponding value for the wild type enzyme.
Table 2: Kd and kcat values of wild-type HbpA and HbpAind towards 2-hydroxybiphenyl and indole.
2-Hydroxybiphenyl Indole
a b Enzyme kcat Kd kcat/Kd kcat Kd kcat/Kd
s-1 µM s-1 M-1 s-1 µM s-1 M-1
HbpA 15.6 9.9±0.7c 1.6 × 106 5 × 10-3 1500±70 3.3
× 5 × -2 × 3 HbpAind 2.3 8.8±0.7 2.6 10 9 10 78±7 1.1 10
a Determined by measuring substrate consumption and product formation by reverse phase HPLC. Values are the average of 3 independent measurements and have a standard error <10%. b Determined by an in vitro indigo formation assay with NADH regeneration by formate dehydrogenase. Values are the average of 2 independent measurements and have a standard error <10%. c Value adapted from Suske et al. (Suske et al., 1999).
0,2 Fig. 4: In vitro indigo formation by HbpA
and HbpAind. The assay contained 0.2 µM 0,15 HbpA or HbpAind, 0.25 U formate dehydrogenase, 160 mM sodium formate, 0,1 10 U catalase, 0.3 mM NADH, 2 mM indole in 1 mL 50 mM sodium phosphate buffer pH
indigo conc. [mM] 0,05 7.5. Indigo formation was determined at I 610 nm. , wild-type HbpA; , HbpAind. 0 050100 150 200 time [min]
95 INDOLE HYDROXYLATION BY LABORATORY EVOLVED HBPA
Equilibrium binding of substrates to HbpA and HbpAind. The affinities of the enzymes towards 2-hydroxybiphenyl and indole were determined by titration of the purified proteins with known concentrations of substrate (Fig. 5, insets). Plotting the absorption difference at a specific wavelength as a function of
substrate concentration (Fig. 5) allowed the determination of the Kd values. Whereas the dissociation constants for 2-hydroxybiphenyl were in the same
range for both proteins, the Kd value for indole was 20-fold lower for HbpAind than for HbpA (Table 2). Uncoupling of NADH oxidation from 2-hydroxybiphenyl hydroxylation. Wild- type HbpA shows an uncoupling of NADH oxidation from 2-hydroxybiphenyl hydroxylation of 21% (Meyer et al., 2002b). Substitution of a single amino acid
(Val368Ala in HbpAT1) completely coupled these two reactions (Meyer et al., 2002b). In contrast, there was significant uncoupling of NADH oxidation from
hydroxylation for HbpAind, compared to HbpA and HbpAT1 (Table 3). To
investigate whether the increased uncoupling in HbpAind is an effect of the combination of the two amino acid substitutions or only due to the D222V
exchange, the single mutant D222V (HbpAD222V) was constructed by site directed mutagenesis. Uncoupling of NADH oxidation from 2-hydroxybiphenyl
hydroxylation was 3-fold higher for the HbpAD222V mutant monooxygenase than for the wild-type protein.
96 CHAPTER IV
Table 3: Uncoupling of NADH oxidation from 2-hydroxybiphenyl hydroxylation The assays were performed at 30°C in 20 mM sodium phosphate buffer (pH 7.5) containing 0.3 mM NADH and 0.2 mM 2-hydroxibiphenyl.
Specific activity Unproductive Amino acid Enzyme NADH Uncoupling substitution NADH Product oxidation oxidationa formationb
s-1 s-1 s-1 %
HbpAc - 15.6 12.3 3.3 21
c HbpAT1 V368A 16.2 15.8 0.4 3
HbpAD222V D222V 15.0 5.1 9.9 66
HbpAind D222V/V368A 5.2 2.3 2.9 56
a Determined by monitoring the NADH decrease at 340 nm. b Determined by measuring substrate consumption and product formation with reverse phase HPLC. c Values adapted from Meyer et al. (Meyer et al., 2002b).
0.016 0.01 0.014
0.008 0.012
0.008 0.01 0.01 0.006 0.006 0.004 0.005
0.002 0.008 0 0
0.004 -0.002 0.006 -0.005 Absorbance Absorbance ∆
-0.004 ∆ -0.01 Absorbance at 490 nm -0.006 Absorbance at 495 nm 0.004 ∆ ∆ -0.008 0.002 -0.015 350 400 450 500 550 350 400 450 500 550 Wavelength [nm] 0.002 Wavelength [nm]
0 0 0 1234567 0 100 200 300 400 500 600 700
Indole [mM] Indole [µM]
Fig. 5: Equilibrium binding of indole to wild-type HbpA and HbpAind. Left: Absorption changes at 490 nm during titration of 12 µM wild type HbpA with 0.3, 0.7, 1.3, 2.3, 4, and 6.7 mM indole. The inset shows the difference spectra in the presence of 0.3, 0.7, and 2.3 mM
substrate. Right: Absorption changes at 495 nm during titration of 12 µM HbpAind with 33.3, 66.7, 100, 166.7, 266.7, 433.3, and 666.7 µM indole. The inset shows the difference spectra in the presence of 33.3, 66.7, and 100 µM substrate.
97 INDOLE HYDROXYLATION BY LABORATORY EVOLVED HBPA
Comparison of HbpA with phenol 2-monooxygenase and p-hydroxy- benzoate hydroxylase The only flavoprotein aromatic hydroxylases with known three-dimensional structures are phenol 2-monooxygenase (PHHY) from Trichosporon cutaneum and p-hydroxybenzoate hydroxylase (PHBH) from Pseudomonas fluorescens (Enroth et al., 1998; Schreuder et al., 1989). Structure homology modeling of HbpA with PHHY showed that the FAD-binding and substrate-binding domains of both enzymes are structurally conserved (Meyer et al., 2002b). In this model Asp222 is located close to the bound substrate in the substrate-binding domain of HbpA. This residue corresponds to Ala267 of PHHY and Tyr201 of PHBH (Enroth et al., 1998; Gatti et al., 1994; Schreuder et al., 1994). Ala267 of PHHY is located in a β sheet close to the catalytically important residues Arg287 and Tyr289 (Enroth et al., 1998). Tyr201 of PHBH is part of a hydrogen-bond network and is involved in substrate activation (Entsch et al., 1991; Lah et al., 1994).
DISCUSSION
Hydroxylation of indole by HbpAind Indole is oxidized by different oxygenases that contain either protein-bound iron or cytochrome to activate molecular oxygen (Carredano et al., 2000; Gillam et al., 2000). No flavoprotein oxygenase has thus far been shown to accept
indole as a substrate. A 2-hydroxybiphenyl 3-monooxygenase variant (HbpAind), which we obtained during directed evolution of HbpA, showed the ability to
hydroxylate indole. The kcat/Kd of HbpAind for indole was 330-fold higher than that of wild-type HbpA, and is on the same order as the catalytic efficiency determined for an engineered fatty-acid hydroxylase, P450 BM-3 (Li et al., 2000). This P450 variant was obtained by saturation mutagenesis and is the enzyme with the highest known catalytic efficiency towards indole.
Cultures of E. coli JM101 that synthesized HbpAind during growth on LB medium had an indigo productivity of about 5 mg L-1 h-1. In comparison, a recombinant E. coli HB101 culture that expressed the naphthalene dioxygenase
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genes produced about 1 mg L-1 h-1 during growth on the same medium to a similar cell density (Ensley et al., 1983). E. coli cultures that synthesized human cytochrome P450s reached productivities of approximately 0.3 mg L-1 h-1 on
fortified TB medium (Gillam et al., 2000). Thus, the HbpAind flavoprotein recombinant is considerably (5 to 15-fold) more active in vivo in the formation of indigo from complex medium. Analysis of the pigments formed also showed the presence of the byproduct indirubin, a structural isomer of indigo that is known to be formed by indole oxidizing enzymes (Bialy, 1997; Hart et al., 1992). Indigo extracted from recombinant E. coli cell cultures showed only limited stability when stored in DMF at room temperature. This could be attributed to the pH of the solution. It is known from denim manufacturing, that at basic pHs indigo is chemically reduced to its water soluble form indigo white. In contrast, the indirubin was stable when extracted and stored in DMF.
Major reaction products of indole hydroxylation The products of indole oxidation by different mono- and dioxygenases have been investigated. Indole-epoxide has been suggested as a product for the reaction catalyzed by styrene monooxygenase (O'Connor et al., 1997), and indoxyl (3-hydroxyindole) has been identified as a hydroxylation product of cytochrome P450 enzymes (Gillam et al., 2000). Oxidation by naphthalene dioxygenase results in the formation of 2,3-dihydroxy-2,3-dihydroindole (Ensley et al., 1983). All these reaction products are unstable and spontaneously form pigments. The identification of the intermediates formed during in vitro indigo
assays with HbpAind suggests a similar route as observed for the P450 enzymes (Fig. 6). The identification of 3-hydroxyindole and 2-indolinone indicates direct hydroxylation at the carbons of the pyrrole ring of indole. In contrast to the P450 enzymes, no hydroxylation at the benzene ring of the substrate was observed (Gillam et al., 2000). The presence of isatin could have two origins: it was either produced by hydroxylation of indoxyl or oxindole, or by the decomposition of indigo and indirubin. The formation of indigo and indirubin from the enzymatic hydroxylation products of indole is spontaneous and biocatalyst independent. In
99 INDOLE HYDROXYLATION BY LABORATORY EVOLVED HBPA
short, condensation of two molecules of indoxyl followed by air oxidation leads to the production of indigo, while the condensation of indoxyl and 2-indolinone yields indirubin (Eaton & Chapman, 1995; Russell & Kaupp, 1969). In addition, indirubin can also be formed by the reaction of indoxyl with isatin (Bialy, 1997; Maugard et al., 2001). That this latter reaction does indeed take place in the
case of indirubin formation by HbpAind is supported by the fact that the ratio of indirubin to indigo increased with time when recombinant E. coli were grown on LB.
N H 1
OH O
O OH N N N N H H H H 3a 3 2 2a
O catalyzed by HbpAind O spontaneous N 4 H
O O H N
N NH N H H O O 5 6
condensation of 2 + 3 condensation of 2 + 2 or 2 + 4
Fig. 6: Proposed pathway for the formation of indigo and indirubin by HbpAind. 1) indole; 2) 3-hydroxyindole (indoxyl), 2a) 3-indolinone; 3) 2-hydroxyindole, 3a) 2-indolinone (oxindole); 4) isatin; 5) indirubin; 6) indigo.
100 CHAPTER IV
Substrate spectrum of HbpAind Indirubin inhibits cyclin-dependent kinases and therefore belongs to a group of promising anticancer compounds (Hoessel et al., 1999). Analogs such as indirubin-3’-monoxime or halogenated indirubins show even higher potency (Marko et al., 2001). This increased biological activity can be attributed to the lower hydrophobicity of the derivatives compared to indirubin. Thus the uptake of the compound is facilitated and the probability that it reaches the biological sites of action is increased (Hansch, 1969). In this context we investigated the
substrate spectrum of HbpAind. The variant showed activity with several indole derivatives, such as 4- and 5-hydroxyindole. The products of these reactions are potentially interesting because their logP values are significantly different from that of the unsubstituted indirubin and are hardly accessible by chemical means. However, analysis of the resulting pigments showed that the dihydroxyindirubin derivatives were only a minor product of the reaction of
HbpAind with hydroxyindoles. The mass and the spectral properties of the major condensation product indicates that mainly the indoxyl red derivatives were formed. Indoxyl red is known to be formed from the reaction of 3-oxo 3H-indole with indole (Capdevielle & Maumy, 1993). The proposed pathway for the
formation of the dihydroxy derivatives by HbpAind from hydroxyindole is shown in Figure 7. The electron donating hydroxyl group at the benzene ring facilitates the oxidation of the indolinone compared to the unsubstituted compound and
may explain why indoxyl red was not detected from the reaction of HbpAind with indole. Indoxyl red shares a high degree of structural similarity with indirubin. The dihydroxy derivatives have a strongly decreased hydrophobicity and their potency in inhibiting cyclin-dependent kinases is worth investigating.
101 INDOLE HYDROXYLATION BY LABORATORY EVOLVED HBPA
OH O O HbpAind oxidation HO HO HO HO N N N N H H H
1 234
O OH
1 + 4 HO N NH
5
Fig. 7: Proposed pathway for the formation of dihydroxy derivatives of indoxyl red. 1) 4- or 5-hydroxyindole, 2) 4- or 5-hydroxyindoxyl 3) 4- or 5-hydroxy-2-indolinone 4) 4- or 5-hydroxy-3-oxo-3H-indole 5) 4,4’- or 5,5’-dihydroxyindoxyl red.
Catalytic properties of HbpAind
We characterized HbpAind with respect to its catalytic properties. The variant’s activity towards indole was about 18-fold increased compared to the wild type enzyme. This increase was concomitant with an enhanced affinity of the enzyme towards this substrate. The in vitro activity of the mutant monooxygenase towards the natural substrate 2-hydroxybiphenyl was significantly decreased, while its affinity towards this substrate remained unchanged. This is mostly due to a slower flavin reduction, as indicated by the reduced NADH oxidation rate. In addition, uncoupling of NADH oxidation from
substrate hydroxylation was significantly increased in HbpAind. HbpAind evolved
from the single mutant V368A (HbpAT1) by directed enzyme evolution. In
contrast to wild-type HbpA, HbpAT1 fully couples NADH oxidation to 2-hydroxybiphenyl hydroxylation. We have suggested that this is due to the stabilization and/or improved positioning of the flavin (C4a)-hydroperoxide towards the substrate (Meyer et al., 2002b). This enhanced hydroxylation
efficiency was completely destroyed by the D222V substitution: in HbpAind the uncoupling was twice that of the wild-protein with 2-hydroxybiphenyl as
102 CHAPTER IV
substrate, while the unproductive NADH oxidation rate was similar in both
enzymes. The single mutant HbpAD222V even showed a 3-fold increased uncoupling, which was concomitant with a 3-fold increased unproductive NADH
oxidation rate. Thus, substitution D222V in HbpAind and HbpAD222V directly increases the ratio of flavin (C4a)-hydroperoxide decay to 2,3-dihydroxybiphenyl This ratio is primarily influenced by stabilization of the flavin (C4a)-hydroperoxide, solvent access to the active site, and the reactivity of the substrate towards electrophilic attack by the terminal oxygen of the peroxide (Fig. 8).
HbpA HbpAT1 2-hydroxybiphenyl + H2O2 2-hydroxybiphenyl + H2O2
R R R R O N N O O N N 21 N N O 3 N N
100% N : 104% N : N N N N H H N H O N H H O H O O O HO 79 O HO 97 EFlHOOH-S EFlHOOH-S EFl ox EFl ox
2,3-dihydroxybiphenyl + H2O 2,3-dihydroxybiphenyl + H2O
HbpAD222V HbpAind 2-hydroxybiphenyl + H2O2 2-hydroxybiphenyl + H2O2
R R R N N O O R 66 N N O N N 56 33% N N O 96% N N : N N : H N N H N H O H O H O N H HO O O 34 HO 44 O EFlHOOH-S EFl ox EFlHOOH-S EFl ox
2,3-dihydroxybiphenyl + H2O 2,3-dihydroxybiphenyl + H2O
Fig. 8: Ratio of flavin (C4a)-hydroperoxide decay to 2,3-dihydroxybiphenyl formation in the catalytic cycle of HbpA and the variants. For each of the variants the rate of formation of the flavin (C4a)-hydroperoxide is normalized compared to wild-type enzyme, assuming flavin reduction to be the rate-limiting step. EFlHOOH-S, flavin (C4a)-hydroperoxide enzyme-substrate
complex; EFlox, enzyme containing oxidized flavin.
Interestingly, Asp222 in HbpA corresponds with Tyr201 of p-hydroxybenzoate hydroxylase (PHBH) from Pseudomonas fluorescens (Gatti et al., 1994; Schreuder et al., 1994). In PHBH, Tyr201 is critically involved in the ionization of the substrate, thus affecting flavin movement and allowing efficient
103 INDOLE HYDROXYLATION BY LABORATORY EVOLVED HBPA
hydroxylation (Gatti et al., 1996; Palfey et al., 1999). Substitution Tyr201Phe results in an increased uncoupling of NADH oxidation from substrate hydroxylation up to 95% (Entsch et al., 1991). In phenol 2-monooxygenase (PHHY) from Trichosporon cutaneum, Tyr289 is hydrogen bonded with the hydroxyl group of the substrate, comparable to Tyr201 in PHBH (Enroth et al., 1998). Substitution of this residue by Phe increased the uncoupling from 10 to 34% (Xu et al., 2001). With respect to the localization of Asp222 in the HbpA model, the chemical properties of this residue, and the effects resulting from its substitution, it is likely that Asp222 plays a similar role in HbpA as do the tyrosine residues in PHBH and PHHY.
In summary, we have characterized the HbpAind mutant monooxygenase, the first flavoprotein able to hydroxylate indole. These investigations point to the importance of amino acid residue Asp222 in the catalytic cycle of HbpA. Thus, the results obtained here may serve as the basis for further elucidation of the mechanism of substrate activation in this enzyme.
ACKNOWLEDGEMENTS
We thank Dr. E. Wehrli (ETH Zurich) for performing the electron microscopy and the Swiss National Science Foundation for financial support (grant number 5002-046098).
104 CHAPTER V
CHAPTER V:
ASPARTATE 222 IN 2-HYDROXYBIPHENYL 3-MONOOXYGENASE:
A NEGATIVE CHARGE ESSENTIAL FOR EFFICIENT CATALYSIS
ANDREAS MEYER, DANIEL TANNER, ANDREAS SCHMID, HANS-PETER E. KOHLER,
WILLEM J. H. VAN BERKEL, AND BERNARD WITHOLT
105 ROLE OF ASP222 IN 2-HYDROXYBIPHENYL 3-MONOOXYGENASE
SUMMARY
Recently, we characterized a 2-hydroxybiphenyl 3-monooxygenase (EC
1.14.13.44, HbpA) variant (HbpAind) which showed hydroxylation activity with indole (Meyer et al., 2002c). In particular, the substitution Asp222Val, which significantly decreased the hydroxylation efficiency in HbpA, prompted us to
investigate the HbpAind mutant monooxygenase further. The redox potential of the FAD in HbpA was determined to be -190 mV for the free enzyme and –166 mV for the enzyme-2-hydroxybiphenyl complex. The
two substitutions D222V/V368A in HbpAind decreased the redox potential of the free enzyme to -260 mV, while the value for the enzyme-substrate complex was considerably higher at -182 mV. Asp222 was replaced by different amino acids to study the role of this residue in catalysis. Substitution by a negatively charged amino acid had no significant effect on the catalytic properties of HbpA. In contrast, substitution by an uncharged residue significantly increased the uncoupling of NADH oxidation from product formation. No residual catalytic activity was observed when Asp222 was exchanged with a positively charged residue. The results shown here provide additional evidence for the earlier suggestion that Asp222 is involved in 2-hydroxybiphenyl deprotonation, thereby activating the substrate for efficient catalysis.
106 CHAPTER V
INTRODUCTION
2-Hydroxybiphenyl 3-monooxygenase (EC 1.14.13.44, HbpA), the first enzyme involved in degradation of 2-hydroxybiphenyl in Pseudomonas azelaica HBP1 (Kohler et al., 1988), is a homotetrameric flavoenzyme that catalyzes the ortho-hydroxylation of a variety of 2-substituted phenols using NADH as electron donor (Kohler et al., 1988; Kohler et al., 1993a; Suske et al., 1997). 2-Hydroxybiphenyl 3-monooxygenase is a member of the family of flavoprotein aromatic hydroxylases (van Berkel et al., 1997; van Berkel & Müller, 1991). Due to the unique optical properties of the flavin prosthetic group, these enzymes can be conveniently studied by spectroscopic techniques (Macheroux, 1999; van Berkel et al., 1999). Such studies have led to significant understanding of the catalytic mechanism of flavoprotein aromatic hydroxylases (Arunachalam & Massey, 1994; Arunachalam et al., 1994; Entsch & Ballou, 1989; Entsch et al., 1976; Husain & Massey, 1979; Maeda-Yorita & Massey, 1993; White-Stevens & Kamin, 1972). The available evidence suggests that the reaction is an electrophilic substitution at an activated carbon atom of the substrate aromatic ring (Maeda-Yorita & Massey, 1993; Massey, 1994; Ortiz- Maldonado et al., 1999a). The required activation of oxygen is achieved by the transient stabilization of a covalent flavin (C4a)-hydroperoxide (Massey, 1994). Equation 1 shows the overall reaction scheme of flavoprotein aromatic hydroxylases, where X represents an activating group in the aromatic substrate
(e.g. -OH or -NH2) (Massey, 1994). + → + X-Ar + NAD(P)H + H + O2 X-Ar-OH + NAD(P) + H2O (Eq. 1) As can be seen from equation 1, flavoprotein aromatic hydroxylases have three substrates: the aromatic substrate to be oxygenated, NAD(P)H to reduce the enzyme bound flavin, and molecular oxygen. The presence of the aromatic substrate enables the NAD(P)H-dependent reduction of the flavin prosthetic group. Thereafter, the reduced substrate-enzyme complex readily reacts with molecular oxygen to form a flavin C(4a)-hydroperoxide. The terminal oxygen of this peroxide is then transferred to the aromatic substrate and results in a flavin
107 ROLE OF ASP222 IN 2-HYDROXYBIPHENYL 3-MONOOXYGENASE
C(4a)-hydroxide enzyme-product complex. After product release and elimination of water from the flavin C(4a)-hydroxide, the enzyme can undergo another catalytic cycle (van Berkel et al., 1997). The only enzymes in the family of flavoprotein aromatic hydroxylases with known structure are p-hydroxybenzoate hydroxylase and phenol hydroxylase (Enroth et al., 1998; Schreuder et al., 1989; Wierenga et al., 1979). In part because of this, p-hydroxybenzoate hydroxylase has become the model protein of this enzyme family and has been the ideal object of numerous structure- function studies in the past two decades (Entsch & van Berkel, 1995; Gatti et al., 1994; Schreuder et al., 1994; Schreuder et al., 1989). It has been shown that flavin movement, which is induced by substrate binding and activation, plays a crucial role during catalysis (Gatti et al., 1996; Gatti et al., 1994; Schreuder et al., 1994). In an effort to change the substrate specificity we subjected 2-hydroxybiphenyl 3-monooxygenase to directed evolution (Arnold, 1998a; Arnold & Volkov, 1999; Moore & Arnold, 1996; Stemmer, 1994) and enzyme variants with different substrate spectra and catalytic efficiencies have been
identified (Meyer et al., 2002a; Meyer et al., 2002b). One of these, HbpAind, hydroxylates indole (Meyer et al., 2002c). Initial characterization of this mutant monooxygenase suggests that Asp222 is involved in substrate activation in
HbpA, since HbpAind does not efficiently hydroxylate the natural substrate 2-hydroxybiphenyl anymore. In this study we report on the characterization of
HbpAind with respect to the function of amino acid residue Asp222.
MATERIALS AND METHODS
Chemicals, strains and plasmids Escherichia coli JM101 (Sambrook et al., 1989) and the plasmid pUC18 (Yanish-Perron et al., 1985) were used for cloning and expression of the hbpA gene. Taq DNA polymerase, restriction enzymes, and T4 DNA ligase were
108 CHAPTER V
purchased from Roche Molecular Biochemicals. All chemicals were of purest available quality and obtained from Fluka AG (Buchs, Switzerland).
Directed evolution of HbpA Construction of a random mutant library of HbpA was based on in vitro manganese mutagenesis (error prone PCR) as described recently (Meyer et al., 2002b). Subsequent in situ screening was applied to select HbpA variants with altered substrate specificity.
Site directed mutagenesis Site-directed mutants D222E, D222H, D222K, D222N, D222S, and D222V were obtained with the QuickChangeTM site directed mutagenesis kit from Stratagene (La Jolla, USA) as described in the instruction manual. The following mutagenesis primer pairs were used: 5’-CGCGGAACATCCAGTAC ATXXXGCCTTTACGATGTTCACACA-3’ where XXX stands for the altered codon (Table 1).
Table 1: Base substitutions in Mutation Original codon Mutated codon the mutagenic primers for site D222E GAC GAA directed mutagenesis of amino
222 D222H GAC CAC acid residue Asp D222K GAC AAG D222N GAC AAC D222S GAC AGC D222V GAC GTC
Preparation of cell extract Cells from a 5 mL LB culture were spun down at 5000 x g for 15 min and resuspended in 800 µL 50 mM phosphate buffer (pH 7.2). This suspension was transferred to a 1.5 mL Eppendorf tube containing 1.2 g glass beads (Ø 0.1 - 0.2 mm) and the cells were disrupted in a Retsch mill (Retsch GmbH, Hann, Germany) for 10 min at 90% power. The cell extracts were separated from the
109 ROLE OF ASP222 IN 2-HYDROXYBIPHENYL 3-MONOOXYGENASE
glass beads by centrifugation (15000 x g, 15 min) and supplemented with FAD to a final concentration of 50 µM.
Protein purification E. coli JM101 harboring either the wild type or variant hbpA gene was grown in M9 medium (Sambrook et al., 1989) with glycerol as carbon source and supplemented with MT trace element solution (Lageveen et al., 1988). 150 mg L-1 ampicillin were added to ensure plasmid maintenance. After harvesting the cells, HbpA and variants were purified as described elsewhere (Meyer et al., 2002b).
Analytical methods Standard activity assays were performed in 20 mM air saturated sodium phosphate buffer (pH 7.5, 30°C) containing 0.3 mM NADH and 0.2 mM 2- hydroxybiphenyl (Kohler et al., 1988). Specific activities were determined by measuring substrate consumption and product formation by reverse phase HPLC (Kohler et al., 1993a) using a Hypersil ODS column (5 µm, 4.5 x 125 mm) and a Hewlett Packard HP 1050 Ti HPLC coupled to a diode array detector (HP DAD 1040M). The samples were diluted 1:1 with MeOH/0.1% phosphoric acid
and separated with 60% MeOH/40% H2O (0.1% phosphoric acid) as mobile phase. NADH oxidase activities were measured by the same assay conditions by monitoring NADH consumption at 340 nm in an ATI Unicam UV4 UV/VIS spectrophotometer.
Molar absorption coefficients of HbpA and variant HbpAind were determined by unfolding the enzymes with 0.5% SDS in 20 mM sodium phosphate buffer pH 7.5 (McKean et al., 1983). Absorption spectra were taken before and after unfolding using a Hewlett Packard HP 8453 diode array spectrophotometer. Dissociation constants of enzyme-substrate complexes were determined from absorption spectral perturbations of the enzyme bound FAD during titration with known concentrations of substrate (Suske et al., 1999). Spectra were recorded at 25 °C in 20 mM Hepes, pH 7.5, using an Aminco DW-2000 double beam spectrophotometer.
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Determination of redox potentials Redox potentials of wild type HbpA and HbpA variants were determined by the xanthine/xanthine oxidase method of Massey (Massey, 1991) in 20 mM HEPES buffer (pH 7.5) at 25°C. 10 µM enzyme, 10 µM reference dye, 2.5 µM methyl viologen, and 400 µM xanthine were made anaerobic by flushing with argon. The reduction was started by the addition of a catalytic amount of xanthine oxidase. Oxidized species of HbpA and dye were monitored spectrophotometrically until reduction was complete. To reach equilibrium conditions between the oxidized and reduced form of enzyme and the dye took
1- 2 hours. Anthraquinone-2,6-disulfonate (Em = -184 mV) (Fultz & Durst, 1982)
and phenosafranine (E m = -252 mV) (Fultz & Durst, 1982) were used as reference dyes. The oxidation-redox potentials were calculated according to the method of Clark (Clark, 1960) using the following equations:
• Eh(dye) = Em(dye) + (59/ndye) log(dyeox/dyered) • Eh(E) = Em(E) + (59/nE) log(Eox/Ered)
At equilibrium, Eh(dye) = Eh(E) (Eq. 2)
Where Eh is the redox potential; Em is the midpoint redox potential; ndye is the
number of electrons accepted by the dye; nE is the number of electrons accepted by the prosthetic group of the enzyme.
By plotting log(Eox/Ered) versus log(dyeox/dyered) according to the method of Minnaert (Minnaert, 1965), the midpoint redox potentials could be determined.
111 ROLE OF ASP222 IN 2-HYDROXYBIPHENYL 3-MONOOXYGENASE
RESULTS
Properties of HbpAind, HbpAT1, and HbpAD222V
General properties. The general properties of wild-type HbpA, HbpAind,
HbpAT1, and HbpAD222V were reported recently (Meyer et al., 2002b; Meyer et al., 2002c). The amino acid substitutions in these mutant monooxygenases are summarized in Table 2. Table 2: Amino acid substitutions in HbpA mutants
Enzyme Amino Acid Position 222 368 HbpA Asp Val
HbpAT1 Asp Ala
HbpAD222V Val Val
HbpAind Val Ala
Spectral properties. The absorption spectra of HbpA and the HbpAind variant showed only small differences (Fig. 1), suggesting that the amino acid replacements cause minor changes in the vicinity of the FAD prosthetic group.
The spectral properties of HbpA and HbpAind are summarized in Table 3. No significant differences were observed in the absorption spectrum of the protein
bound FAD of the two single mutants HbpAT1 and HbpAD222V either.
0.16 Fig. 1: Absorption spectra of HbpA and
0.14 variant HbpAind. Shown are absorption 0.12 spectra of wild type HbpA (solid line) and 0.1 variant HbpAind (dotted line) in 50 mM 0.08 phosphate buffer (pH 7.5) at 25°C. Both
Absorbance 0.06 0.04 proteins are present at 15 µM and in the 0.02 oxidized form. 0 350 400 450 500 550 Wavelength [nm]
112 CHAPTER V
Table 3: Spectral properties of oxidized wild type 2-hydroxybiphenyl
3-monooxygenase and variant HbpAind. The data were taken from the spectra shown in Figure 2.
λ ε Enzyme max ox
nm mM-1cm-1 ε ε ε ε III I II I/ II HbpA 377 452 9.5 10.0 0.95
HbpAind 379 447 9.7 10.5 0.92
Catalytic properties of HbpAind, HbpAT1, and HbpAD222V Endogenous NADH oxidation. In the absence of 2-hydroxybiphenyl, HbpA has an NADH oxidase activity of 0.14 U mg-1 protein (Suske et al., 1997).
HbpAT1 showed about 30% endogenous NADH oxidation compared to the wild
type enzyme, whereas the rate for HbpAD222V was slightly increased. In the
HbpAind variant, the rate of NADH oxidation was increased more than 10-fold compared to the wild-type enzyme (Table 4). Induction of NADH oxidation by 2-hydroxybiphenyl. The addition of the aromatic substrate increases the rate of NADH oxidation of HbpA by a factor of 30 (Suske et al., 1997). 2-Hydroxybiphenyl also stimulated the rate of NADH
oxidation in the single mutants HbpAT1 and HbpAD222V. However, with the
HbpAind variant the rate of NADH oxidation was 15% lower with the enzyme- substrate complex than with the free enzyme (Table 4). Activity towards 2-hydroxybiphenyl and uncoupling of NADH oxidation from substrate hydroxylation. A summary of these already published data and the endogenous NADH oxidation rates is provided in Table 4.
113 ROLE OF ASP222 IN 2-HYDROXYBIPHENYL 3-MONOOXYGENASE
Table 4: NADH oxidase activity of wild type HbpA and variant HbpAind. The assay mix contained 10 mM sodium phosphate buffer (pH 7.5), 0.3 mM NADH and, if required, 0.2 mM 2-hydroxybiphenyl. The reaction was carried out at 30°C.
kcat
HbpA HbpAT1 HbpAD222V HbpAind
s-1 s-1 s-1 s-1 NADH oxidation1 NADH 0.6 0.2 1.0 6.2 NADH + 2-hydroxybiphenyl 15.6 16.2 15.0 5.2 Hydroxylation2 NADH + 2-hydroxybiphenyl 12.3 15.8 5.1 2.3
Uncoupling (%) 21 3 66 56
1 Determined by monitoring the NADH decrease spectrophotometrically at 340 nm. 2 Determined by measuring substrate consumption and product formation with reverse phase HPLC.
Affinity of 2-hydroxybiphenyl towards HbpAD222V. The dissociation constants
of 2-hydroxybiphenyl from HbpA and HbpAind have been reported earlier (Meyer
et al., 2002c; Suske et al., 1999). The Kd value of 2-hydroxybiphenyl towards
HbpAD222V was determined to be 5.0±1.6 µM and was in the same range as observed for the wild-type enzyme.
Redox potentials of the FAD in HbpA and HbpAind
The redox potentials of the enzyme bound FAD in HbpA and HbpAind were determined to assess the effects of the amino acid substitution in the mutant monooxygenase on the redox properties. We therefore reduced HbpA and the variant with the xanthine/xanthine oxidase system in the presence of an appropriate reference dye. Anthraquinone-2,6-disulfonate was chosen as the mediator compound for the determination of the midpoint redox potential of the flavin of the free wild-type enzyme and the enzyme-2-hydroxybiphenyl complex. The same reference dye was used for the FAD of the enzyme-substrate
complex of HbpAind, whereas phenosafranine was used for the flavin in the free
114 CHAPTER V
enzyme (Fig. 2). In all cases the reference dye and the enzyme-bound FAD were reduced simultaneously in a single two-electron reduction process. The
redox potentials were calculated by plotting log(dyeox/dyered) as a function of
log(Eox/Ered) (inset Fig. 3)(Minnaert, 1965). The midpoint redox potential of the
FAD in the free enzyme was decreased by 70 mV in the HbpAind variant whereas the difference between the redox potentials in the two enzyme-ligand complexes was only 14 mV (Table 5).
0.35 1 Fig. 2: Determination of the redox
0.8 potential of HbpA variant HbpAind. 0.3 0.6
0.4 Absorption spectra recorded at 15 min
0.25 log [ox/red] phenosafranine
0.2 0 0.2 0.4 0.6 0.8 intervals during the reduction of 10 µM 0.2 log [ox/red] HbpAind
variant HbpAind in the presence of 10 µM 0.15
Absorbance phenosafranine by the xanthine/xanthine 0.1 oxidase system. The inset shows the 0.05 log(Eox/Ered) versus log(dyeox/dyered) plot. 0 330 370 410 450 490 530 570 610 650 Wavelength [nm]
Table 5: Midpoint redox potentials of HbpA and variant HbpAind. Calculated midpoint redox potentials measured in 20 mM HEPES buffer (pH 7.5) at 25°C.
Em
HbpA HbpAind mV mV Free enzyme -190a -260b Enzyme + 2-hydroxybiphenyl -166a -182a
a Reference dye used: Anthraquinone-2,6-disulfonate (Em = -184 mV) b Reference dye used: Phenosafranine (Em = -252 mV)
115 ROLE OF ASP222 IN 2-HYDROXYBIPHENYL 3-MONOOXYGENASE
Relative activities of single Asp222 mutants
Earlier studies on the catalytic properties of HbpAind showed that substitution of Asp222 by valine significantly influenced the uncoupling of NADH oxidation to product formation (Meyer et al., 2002c). To further investigate the function of Asp222 single mutants with Glu, His, Lys, Asn, and Ser at this position were constructed by site-directed mutagenesis. The endogenous NADH oxidation rates and the specific activities of variants derived from site-directed mutagenesis experiments were compared with the corresponding rates of the wild type enzyme (Table 6). The conserved replacement of Asp222 by Glu had no significant influence on catalysis. Substitution by uncharged amino acids decreased the activity at least 50%, in part due to an increased uncoupling of NADH oxidation from substrate hydroxylation. Substitution of Asp222 by Lys and His resulted in complete loss of enzymatic activity.
Table 6: Relative activities of Asp222 variants towards 2-hydroxybiphenyl. NADH oxidase activity was determined by spectrophotometrically monitoring NADH consumption. Activities based on product formation were calculated by measuring product formation and substrate consumption by reverse phase HPLC. In both cases the experiments were performed in air-saturated phosphate buffer (20 mM, pH 7.5) at 30°C with concentrations of 2-hydroxybiphenyl and NADH of 0.2 mM.
Relative Relative activity endogenous NADH NADH Product Enzyme oxidation oxidationa formation
%%% HbpA 100 100 100 Asp222Glu 110 90 120 Asp222Ser 170 65 35 Asp222Asn 110 70 50 Asp222Lys 000 Asp222His 000 a Values corrected for endogenous NADH oxidation
116 CHAPTER V
DISCUSSION
The catalytic mechanism of flavoprotein aromatic hydroxylases has been studied extensively (van Berkel & Müller, 1991). The reaction can be divided into a reductive and an oxidative half reaction, which are also physically separated in the enzyme. In the so called ‘out’ position of the flavin, a charge- transfer complex between NADPH and FAD is created and the flavin is reduced. Substrate hydroxylation takes place while the flavin adopts the ‘in’ conformation (Gatti et al., 1994; Schreuder et al., 1994). We have recently obtained a 2-hydroxybiphenyl 3-monooxygenase variant
(HbpAind) with increased activity towards indole (Meyer et al., 2002c). Concomitant loss of the ability to effectively hydroxylate the natural substrate
2-hydroxybiphenyl prompted us to investigate HbpAind in more detail, hoping to obtain additional information about sequence-function relationships in HbpA.
Flavin environment Spectroscopic techniques have been used to investigate the region surrounding the flavin group in different flavoproteins (Macheroux, 1999; Swenson & Krey, 1994; van den Heuvel et al., 2000b). Substitutions
D222V/V368A in HbpAind changed the absorption spectrum of HbpA only marginally. This result suggests that these mutations did not cause drastic changes in the microenvironment of the protein bound FAD and therefore the active site of the protein. If there had been a direct contact between the negatively charged Asp222 and the FAD isoalloxazine ring, the substitution with an uncharged amino acid should have resulted in significant changes in the FAD absorption spectrum. The shifts of the absorbance maxima of the transition states also indicate that Asp222 does not contribute directly to the FAD environment. The red shift of transition I, the blue shift of transition II, and the higher extinction coefficients at both absorbance maxima point to a more polar
222 environment in the HbpAind variant compared to the wild-type HbpA. If Asp were located in the close vicinity of the FAD moiety, substitution by valine would be expected to result in a less polar environment.
117 ROLE OF ASP222 IN 2-HYDROXYBIPHENYL 3-MONOOXYGENASE
Redox potential The midpoint redox potential of free 2-hydroxybiphenyl 3-monooxygenase was determined to be -190 mV, which lies between the potential of phenol hydroxylase (-213 mV) (Massey, 1991) and p -hydroxybenzoate hydroxylase (-163 mV) (Entsch et al., 1991). Substrate binding does not affect the redox potential of p-hydroxybenzoate hydroxylase and only slightly decreases that of phenol hydroxylase. In contrast, the potential of the protein bound FAD in HbpA was 24 mV more positive in the presence of 2-hydroxybiphenyl. Upon HbpA titration with substrate it could be observed that the absorbance above 500 nm, characteristic for a charge transfer complex (Kosower, 1966), disappeared (Suske et al., 1999). This was attributed to the disruption of a charge transfer complex between an electron donating active site residue and the FAD moiety. The more positive redox potential is the logical consequence of blocking charge transfer upon substrate binding, which results in a reduced electron density in the flavin prosthetic group. Comparison of the redox potential of the protein-bound FAD in wild-type
HbpA and in HbpAind also yields information about the amino acid substitutions. It has been shown for flavodoxin that replacement of acidic residues by neutral amide counterparts within 13 Å of the flavin increased the redox potential by approximately 15 mV per unit of charge (Swenson & Krey, 1994). This observation also correlates with measurements of the redox potential of p-hydroxybenzoate hydroxylase, where substitution of Asn300, an amino acid residue which has direct contact with the isoalloxazine ring, by aspartic acid, resulted in a 17 mV more negative redox potential for the free enzyme (Palfey
et al., 1994b). In contrast, HbpAind has a 70 mV decreased midpoint potential compared to wild type HbpA. This observation is therefore not consistent with the removal of a negative charge near the FAD, which would facilitate the reduction of the flavin. Together with the spectral similarities of the wild-type
222 enzyme and the HbpAind variant, this clearly indicates that Asp is not located in the immediate vicinity of the flavin prosthetic group. In addition, from the
118 CHAPTER V
decreased redox potential of the FAD in HbpAind it can be concluded that the
significantly increased endogenous NADH oxidation in HbpAind is not a result of the changed electrochemical properties of the prosthetic group. The lower potential would rather make the reduction more difficult. Reconstitution of the apoenzyme of p-hydroxybenzoate hydroxylase (PHBH) with different 8-substituted flavins has revealed that the rate of flavin reduction is not dependent on the redox potential of the flavin derivative, but on blockage of the ‘out’ conformation by steric hindrance (Ortiz-Maldonado et al., 1999b). In conclusion, the increased endogenous NADH oxidation rate points to an improved accessibility for NADH to the flavin. Most probably this is effected by stabilizing the flavin ‘out’ position in the free enzyme, which is supported by the
spectral properties that indicate a more polar environment of the FAD in HbpAind compared to HbpA.
Catalytic role of Asp222 Studies on single mutants of Asp222 showed that a negative charge at this position is required for efficient catalysis. Substitution by an uncharged amino acid resulted in a significantly increased uncoupling of NADH oxidation from substrate hydroxylation. Three main factors that have an effect on the uncoupling of NAD(P)H oxidation from substrate hydroxylation are known: i) transient stabilization of the reactive flavin (C4a)-hydroperoxide; ii) solvent access to the active site; iii) reactivity of the substrate towards the flavin (C4a)- hydroperoxide. From spectral and redox properties of the protein bound FAD and the localization of Asp222 in the HbpA model, the involvement of this residue in the stabilization of the flavoperoxide is unlikely. If solvent access to the active site would be the reason for the increased uncoupling, substitution by a positively charged amino acid would not lead to complete loss of activity. As a consequence, Asp222 must be involved in activation of the substrate prior to the hydroxylation by the flavin (C4a)-hydroperoxide. For PHBH it has been shown that a proton transfer network is responsible for the activation of the aromatic substrate (Entsch et al., 1991; Gatti et al., 1996; Schreuder et al., 1994). This
119 ROLE OF ASP222 IN 2-HYDROXYBIPHENYL 3-MONOOXYGENASE
network consists of the hydroxy groups of Tyr201 and Tyr385, two water molecules, and the surface residue His72 (Gatti et al., 1996). Substitutions Tyr201Phe, Tyr385Phe, and His72Asn all increased the uncoupling of NADPH oxidation from substrate hydroxylation (Entsch et al., 1991; Frederick et al., 2001). Recently we postulated that Asp222 in HbpA is involved in activating the substrate towards electrophilic attack of the reactive flavin (C4a)-peroxide similar to the tyrosine residues in PHBH (Meyer et al., 2002c). The work presented here has given additional evidence for this earlier suggestion. First, the participation of Asp222 in the transient stabilization of the reactive flavin (C4a)-hydroperoxide could be excluded by showing that this residue is not close to the isoalloxazine ring of the prosthetic group. Second, substitution of Asp222 by a positively charged amino acid completely destroyed the catalytic activity of HbpA. Lysine most probably prevents substrate activation, whereas an alternative hydrogen acceptor can be invoked when Asp222 is replaced by an uncharged residue. Further detailed characterization of the Asp222 single mutants may yield final resolution of the participation of this amino acid in deprotonation and thus activation of the substrate prior to its hydroxylation.
120 CHAPTER VI
CHAPTER VI:
CRYSTALLIZATION AND PRELIMINARY X-RAY ANALYSIS OF NATIVE AND
SELENOMETHIONINE 2-HYDROXYBIPHENYL 3-MONOOXYGENASE
ANDREAS MEYER, DANIEL TANNER, ANDREAS SCHMID, DAVID F. SARGENT,
HANS-PETER E. KOHLER, TIMOTHY J. RICHMOND, AND BERNARD WITHOLT
121 CRYSTALLIZATION OF 2-HYDROXYBIPHENYL 3-MONOOXYGENASE
SUMMARY
2-Hydroxybiphenyl 3-monooxygenase (HbpA, E.C. 1.14.13.44) from Pseudomonas azelaica HBP1 was expressed in E. coli both as the native and SeMet-labeled protein. The two enzymes were purified to homogeneity and crystallized by the hanging-drop vapor-diffusion method. The crystals belong to the monoclinic space group C 2, with unit cell dimensions of a = 108.6, b = 196.8, c = 79.3, and ß = 97.7 for the native protein and a = 108.3, b = 196.8, c = 79.0, and ß = 97.8 for SeMet HbpA. Crystal packing considerations led to the assumption of two HbpA subunits per asymmetric unit, which corresponds
3 -1 to a VM value of 3.3 Å Da and a solvent content of 62%. The crystals were radiation sensitive and had a lifetime of only about 120 seconds when exposed to synchrotron radiation. To obtain complete data sets data was collected from 23 native and 26 derivative crystals. The high-resolution limit was 2.0 Å for native HbpA and 2.25 Å for the SeMet derivative.
122 CHAPTER VI
INTRODUCTION
Hydroxylation reactions are among the most widespread enzymatic activities occurring in all life forms from bacteria to humans. The elucidation of how enzymes that catalyze such reactions handle the three required substrates (molecular oxygen, the electron donor, and the substrate to be hydroxylated) remains a challenge (Holland & Weber, 2000). Despite our limited information base, hydroxylases have an enormous biosynthetic value due to their ability to carry out unique oxidations, usually with high regio- and enantioselectivity, that go well beyond the toolkits of classical organic synthesis (Duetz et al., 2001b). 2-Hydroxybiphenyl 3-monooxygenase (EC 1.14.13.44, HbpA) is the first enzyme of the 2-hydroxybiphenyl degradation pathway of Pseudomonas azelaica HBP1 (Kohler et al., 1988). HbpA is able to catalyze the ortho-hydroxylation of different 2-substituted phenols to the corresponding catechols using NADH as electron donor (Fig. 1) (Kohler et al., 1988; Suske et al., 1997). HbpA has a homotetrameric structure with a total mass of 256 kDa, and each subunit contains a noncovalently but tightly bound FAD (Suske et al., 1997). The catalytic mechanism has been investigated by spectroscopic techniques and enzyme kinetics have been reported in detail (Suske et al., 1999). Because of its conserved sequence motifs for FAD/NADH binding, HbpA was classified as a flavoprotein aromatic hydroxylase (Eppink et al., 1997). Of this large family, only two three-dimensional structures have thus far been resolved: those of phenol 2-monooxygenase and p-hydroxybenzoate hydroxylase (Enroth et al., 1998; Schreuder et al., 1989; Wierenga et al., 1979). However, both of these enzymes are homodimers and show only small overall homology with HbpA. 2-Hydroxybiphenyl 3-monooxygenase has proven its potency as an efficient biocatalyst in biotechnological organic syntheses. Different 3-substituted catechols were produced in in vivo and in vitro processes using this enzyme (Held et al., 1999; Held et al., 1998; Schmid et al., 2001). In addition, directed enzyme evolution was successfully used to change the substrate specificity of
123 CRYSTALLIZATION OF 2-HYDROXYBIPHENYL 3-MONOOXYGENASE
HbpA to enable the synthesis of compounds that could not be produced with the wild-type enzyme (Meyer et al., 2002a; Meyer et al., 2002b).
+ + NADH + O2 + H NAD + H2O OH OH OH
R HbpA R
R = phenyl, 2-OH-phenyl, methyl, ethyl, propyl, i -propyl, butyl, sec -butyl, fluoro, chloro, bromo, or iodo.
Figure 1: Reaction catalyzed by 2-hydroxybiphenyl 3-monooxygenase
We have started structural studies for two main reasons: i) to further investigate the catalytic mechanism of HbpA with special reference to the variants obtained by directed evolution, and ii) to rationally design novel biocatalysts for bioorganic synthesis. In this study we report on the production of native and selenomethionine HbpA crystals to reach these goals.
MATERIALS AND METHODS
Expression and purification of HbpA and SeMet HbpA Plasmid pAA1 (Meyer et al., 2002b), a pUC18 derivative harboring the hbpA gene (Yanish-Perron et al., 1985), was used for expression of HbpA and the selenomethionine labeled protein (SeMet HbpA). For synthesis of HbpA the construct was transformed into E. coli JM101 cells (Sambrook et al., 1989), whereas SeMet HbpA was produced in E. coli LE392 (Sambrook et al., 1989). The recombinant strains were grown in 3-L shaking flasks containing 1 L M9 mineral medium (Sambrook et al., 1989), supplemented with 0.1% (v/v) MT
trace element solution (Lageveen et al., 1988), 0.1 mM CaCl2, 2 mM MgSO4, 150 mg L-1 ampicillin, and 0.5 % (w/v) glucose. In addition 0.001% (w/v) thiamine was added for E. coli JM101 and 50 mg L-1 L-SeMet and 50 mg L-1 L-Trp were added for E. coli LE392 cultures. After incubation overnight at 303 K, 0.5% glycerol was added to allow hbpA expression. To ensure plasmid maintenance the addition of ampicillin was repeated and
124 CHAPTER VI
L-SeMet was again added to the E. coli LE392 culture. The E. coli JM101 cultures were harvested after 8 hours, whereas the E. coli LE392 cultures were harvested after incubation overnight. Purification of HbpA and SeMet HbpA was performed as reported recently (Meyer et al., 2002b).
Screening for crystallization conditions Initial screening of crystallization conditions was performed by sparse-matrix screens using the hanging-drop vapor-diffusion method (Cudney et al., 1994; Jancarik & Kim, 1991). The reservoir contained 1 mL precipitant solution and the drop consisted of 2 µL precipitant and 2 µL HbpA solution (10 mg mL-1). Crystallization set-ups were allowed to equilibrate at 293 K. Systematic screening around initial conditions resulted in an optimized precipitant solution
containing 1.6 M ammonium sulfate, 0.1 M NaCl2 and 0.1 M MES/NaOH pH 7.5
Diffraction experiments X-ray diffraction experiments were performed at 100 K. To this end, the crystals were transferred directly into cryoprotectant solution consisting of 30% (v/v) glycerol, 1.6 M ammonium sulfate, 0.1 M NaCl, and 0.1 M MES/NaOH pH 7.5. Both HbpA and SeMet HbpA data sets were measured at the protein crystallography beamline of the Swiss Light Source (SLS, PSI Villingen, Switzerland). Data were recorded using a marCCD detector at 0.9793 Å, the peak wavelength of the SeMet HbpA derivative for processing as SAD or SIRAS data. Integration was done with MOSFLM (Leslie, 1992) and the CCP4 program suite was used for scaling (Collaborative Computational Project, 1994).
125 CRYSTALLIZATION OF 2-HYDROXYBIPHENYL 3-MONOOXYGENASE
RESULTS AND DISCUSSION
Expression and purification of SeMet HbpA Because the catalytic activity of HbpA is completely inhibited by the presence of metals such as mercury, silver and platinum, we produced a SeMet HbpA heavy-atom derivative for phasing. HbpA contains 15 methionine residues per subunit of 64 kDa. We achieved synthesis of SeMet HbpA as 20% of the total protein in E. coli strain LE392. SeMet HbpA could be purified to homogeneity with the method developed for the native protein and showed 80% activity compared to HbpA. MALDI-TOF-MS analysis of HbpA and SeMet HbpA indicated a complete substitution of Met by SeMet. In addition, it revealed the post-translational excision of the N-terminal methionine, which is common in E. coli when, as in the case of HbpA, serine is the penultimate amino acid (Hirel et al., 1989).
Crystallization Crystals of native and SeMet HbpA were obtained within an equilibration time of 10 days (Fig. 2). They were colored bright yellow, indicating an oxidized state of the flavin prosthetic group. Crystals grew up to 0.8 mm in the longest dimension. However, it turned out that such large crystals were of inferior quality. Crystals with a size of about 100 × 200 × 400 µm showed clearer diffraction patterns, though to a lower resolution.
Figure 2: Native HbpA crystal with dimensions of 100 × 200 × 400 µm.
126 CHAPTER VI
Diffraction experiments Native crystals of HbpA diffracted to a resolution of 2.0 Å, whereas 2.25 Å was the limit for SeMet HbpA. Crystals from both protein types were isomorphous and belong to the monoclinic space group C2. Based on two
HbpA subunits per asymmetric unit the crystal packing parameter V M is 3.3 Å3 Da-1, which corresponds to a solvent content of 62% (Matthews, 1968). Although the measurements were performed at 100 K, the HbpA and SeMet HbpA crystals were radiation sensitive and showed structural damage after a total of 120 seconds exposure time. Data from 23 HbpA crystals were therefore collected over 10° each with oscillations of 0.5°. 26 SeMet HbpA crystals, each collected over 12.5°, were used for the derivative data set. The detailed statistics of the merged data are listed in Table 1.
Table 1: Data statistics for HbpA and SeMet HbpA crystals. Values in parentheses refer to the outer resolution shell. The data set for HbpA consisted of 23 different crystals and for SeMet HbpA of 26 crystals.
Native HbpA SeMet HbpA
Wavelength (Å) 0.979297 0.979297 Resolution range (Å) 20.0-2.0 (2.11-2.0) 20.0-2.25 (2.37-2.25) Spacegroup C2 C2 Unit cell a = 108.6, b = 196.8, a = 108.3, b = 196.8, parameters (Å, °) c = 79.3, ß = 97.7 c = 79.0, ß = 97.8 Completeness (%) 92.2 (87.5) 99.7 (100) Redundancy 4.9 (4.1) 6.1 (6.3)
† Rmerge (%) 9.4 (22.2) 11.0 (24.6) 〈I/σ(I)〉 overall 4.8 (3.0) 4.4 (2.8)
† Σ 〈 〉 Σ Rmerge = I - I / I, where I is the integrated intensity of a given reflection.
The localization of the selenium sites in large structures of SeMet labeled proteins is still a challenge. However, computational advances facilitate this task and have allowed the successful elucidation of a number of large selenium
127 CRYSTALLIZATION OF 2-HYDROXYBIPHENYL 3-MONOOXYGENASE
substructures (Deacon & Ealick, 1999). We are confident that the recorded data will allow the positioning of the heavy atoms in SeMet HbpA.
ACKNOWLEDGEMENTS
The authors would like to thank Clemens Schulze-Briese, Takashi Tomizaki, Claude Pradervand, and Roman Schneider from the protein crystallography beamline at the SLS for excellent assistance throughout the data collection.
128 CHAPTER VII
CHAPTER VII:
CONCLUDING REMARKS AND OUTLOOK
ANDREAS MEYER
129 CONCLUDING REMARKS AND OUTLOOK
Directed evolution of 2-hydroxybiphenyl 3-monooxygenase (HbpA) from Pseudomonas azelaica HBP1 has shown the value of protein engineering for biocatalytic organic synthesis. Mutagenesis by error prone PCR allowed HbpA to be modified for the production of 3-tert-butylcatechol, a product that can not be obtained with the wild-type enzyme. A general point concerning directed evolution experiments is how far one should go in optimizing a new enzymatic activity. With only two rounds of mutagenesis and screening, as was done to
obtain HbpAT2, it is unlikely that the limit for increasing the catalytic activity towards 2-tert-butylphenol has been achieved. Further cycles of evolution may well lead to additional improvement. How much can the activity of an oxygenase
-1 be increased? Currently, no oxygenase with a kcat higher than 100 s is known. This can be attributed to the complexity of the enzymes, the different substrates they have to handle simultaneously and, last but not least, the slow electron transfer step during catalysis, which may set an upper limit for activity for physical reasons. A reasonable goal for engineering HbpA for biocatalytic
-1 applications would be a kcat of 5 s . This is the threshold above which the biocatalytic synthesis is limited by the process conditions rather than the activity of the enzyme. Since this value is only one third of the activity of the wild-type enzyme towards the natural substrate 2-hydroxybiphenyl, it would seem to be achievable. Widely discussed is the question whether rational or random design is to be preferred in changing specific protein properties. Both approaches have been successfully applied to modify different enzyme properties. When a three-dimensional structure and basic structure-function relationships are known, rational design can lead to a faster positive result. However, the modifications are restricted to the active site of the protein and this approach is therefore quite limited. Furthermore, protein properties such as thermostability, the fundamentals of which are not yet completely understood, can hardly be engineered by rational protein design. For such alterations random techniques are more powerful. Random approaches were also the method of choice to
130 CHAPTER VII
change the substrate specificity of HbpA, since its three-dimensional structure is not known. Nevertheless, would rational design be promising for the engineering of HbpA? With the help of a structural model, preliminary attempts to rationally alter the substrate specificity of HbpA were undertaken, albeit without any success. This is likely due to inaccuracies in the HbpA model, for example improper orientation of the substrate in the active site of the protein. Thus, the final word about the feasibility of rationally designing HbpA for biotechnological application will only be possible when the three-dimensional structure of the enzyme is available. The structural studies on HbpA started in this work are aimed at solving this problem. Furthermore, the three-dimensional structure will allow a more sophisticated interpretation of the amino acid substitutions generated by the directed evolution experiments. Of special interest here is how the exchange of Val368 by alanine results in complete coupling of NADH oxidation to 2-hydroxybiphenyl hydroxylation, whereas in wild-type HbpA these two reactions are uncoupled by about 20%. Understanding this phenomenon will give deeper insight into the mechanism of flavin movement during catalysis in flavoprotein aromatic hydroxylases.
131
CHAPTER VIII
CHAPTER VIII:
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162 CURRICULUM VITAE
CURRICULUM VITAE
Name: Andreas Jörg MEYER Born: July 23, 1971 Nationality: Swiss, citizen of Frauenfeld (TG) Family Status: Single
School Education 1978-1984 Primarschule Frauenfeld, Switzerland 1984-1991 Kantonsschule Frauenfeld, Switzerland Matura Typus B, January 1991
State Service 1991-1992 Military Service
University Education 1992-1997 Eidgenössische Technische Hochschule Zürich, ETHZ Diploma of Natural Sciences, October 1997
Post-Graduate Education 1998-2002 Eidgenössische Technische Hochschule Zürich, ETHZ Institute of Biotechnology Doctoral Dissertation
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